Cellular Domains

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CELLULAR DOMAINS

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CELLULAR DOMAINS

Edited by IVAN R. NABI

A JOHN WILEY & SONS, INC. PUBLICATION

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Copyright © 2011 by Wiley-Blackwell. All rights reserved Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, (978) 750-8400, fax (978) 750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, (201) 748-6011, fax (201) 748-6008, or online at http://www.wiley.com/go/permissions. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at (800) 762-2974, outside the United States at (317) 572-3993 or fax (317) 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com. Library of Congress Cataloging-in-Publication Data: Cellular domains / edited by Ivan R. Nabi. p. ; cm. Includes bibliographical references and index. ISBN 978-0-470-59544-2 (cloth) 1. Cell membranes. I. Nabi, Ivan R. [DNLM: 1. Cell Membrane Structures. 2. Cell Physiological Phenomena. QH601.C435 2011 571.6'4–dc22

3. Cytoplasmic Structures.

QU 350]

2010042298 Printed in Singapore oBook ISBN: 9781118015759 ePDF ISBN: 9781118015735 ePub ISBN: 9781118013742 10

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This book is dedicated to my wife Hakima and children Nessim and Zachary as well as to my parents Ruth and Jim who have supported me throughout my career. It is first of all the work of the contributors, whom I thank enormously for their efforts. It is also the result of my own personal scientific journey that was shaped by my mentors, Avraham Raz and Enrique Rodriguez-Boulan, as well as by all the stimulating interactions I have enjoyed over the years with colleagues, collaborators, students and post-docs.

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CO NTENTS

PREFACE

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CHAPTER 9

ENDOSOMES

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Thierry Galvez and Marino Zerial CONTRIBUTORS PART I

xi CHAPTER 10 LYSOSOMES AND PHAGOSOMES

MEMBRANE DOMAINS

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Guillaume Goyette and Michel Desjardins CHAPTER 1 CYTOSKELETON-INDUCED MESOSCALE DOMAINS

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Ziya Kalay, Takahiro K. Fujiwara, and Akihiro Kusumi CHAPTER 2

CLATHRIN-COATED PITS

CHAPTER 11 ENDOPLASMIC RETICULUM JUNCTIONS

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Jesse T. Chao and Christopher J.R. Loewen

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PART III

CYTOSKELETAL DOMAINS

James R. Thieman and Linton M. Traub CHAPTER 3

CAVEOLAE

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Dan Tse and Radu V. Stan CHAPTER 4

LIPID RAFTS

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Leonard J. Foster CHAPTER 5

MODELING MEMBRANE

DOMAINS

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Daniel Coombs, Raibatak Das, and Jennifer S. Morrison PART II

CHAPTER 12

CHAPTER 13

CHAPTER 14

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Jody Groenendyk and Marek Michalak CHAPTER 8

THE GOLGI APPARATUS

James W. Dennis and Ivan R. Nabi

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MICROTUBULES

229

CILIA

245

Laura K. Hilton and Lynne M. Quarmby

Michael Zick and Andreas S. Reichert CHAPTER 7 THE ENDOPLASMIC RETICULUM

MICROVILLI

Geoffrey O. Wasteneys and Bettina Lechner CHAPTER 15

ORGANELLAR DOMAINS MITOCHONDRIA

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Florent Ubelmann, Sylvie Robine, and Daniel Louvard

CHAPTER 16 CHAPTER 6

THE ACTIN CYTOSKELETON

Jonathan A. Kelber and Richard L. Klemke

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INTERMEDIATE FILAMENTS

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Normand Marceau, Anne Loranger, Stéphane Gilbert, and François Bordeleau

PART IV ADHESIVE AND COMMUNICATING DOMAINS CHAPTER 17

FOCAL ADHESIONS

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Caitlin Tolbert and Keith Burridge

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CONTENTS

CHAPTER 18

THE ADHERENS JUNCTION

303

Christopher P. Toret and W. James Nelson

CHAPTER 22

NEURONAL DOMAINS

371

Jennifer S. Goldman and Timothy E. Kennedy

CHAPTER 19 SPECIALIZED INTERCELLULAR JUNCTIONS IN EPITHELIAL CELLS: THE TIGHT JUNCTION AND DESMOSOME 321

PART VI DOMAINS REGULATING GENE EXPRESSION

Keli Kolegraff, Porfirio Nava, and Asma Nusrat CHAPTER 23 CHAPTER 20

GAP JUNCTIONS

339

Jared M. Churko and Dale W. Laird

CHAPTER 24

POLARIZED CELLULAR DOMAINS PART V

CHAPTER 21

EPITHELIAL DOMAINS

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THE NUCLEAR PORE

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Richard W. Wozniak, Christopher Ptak, and John D. Aitchison 351

Nancy Philp, Liora Shoshani, Marcelino Cereijido, and Enrique Rodriguez-Boulan

CHAPTER 25

CYTOPLASMIC RNA DOMAINS

429

Henry Parker and Tom C. Hobman INDEX

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NUCLEAR DOMAINS

Dale Corkery, Kendra L. Cann, and Graham Dellaire

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PREFACE Cellular compartmentalization into and within organelles segregates biochemical reactions and increases local molecular concentrations, thereby promoting efficiency of cellular processes. Within membranes, subdomains generate lateral heterogeneity that organizes the spatial distribution of glycoprotein receptors and membrane proximal effectors. Morphologically identifiable plasma membrane domains include not only clathrin-coated pits and caveolae but also lipid rafts that form a class of membrane domains that are poorly defined morphologically. Cellular organelles, such as mitochondria, the endoplasmic reticulum, the Golgi apparatus, endosomes, and lysosomes, also define morphologically distinct domains whose functionality depends, in large part, on the establishment of “domains within domains.” Cellular organization is determined by cytoskeletal elements, including the actin and microtubule cytoskeletons, that generate cell surface microvilli and cilia, respectively, as well as intermediate filaments. Adhesive and communicating domains regulate interaction of the cell with the substrate through focal adhesions, as well as with other cells via adherens junctions, tight junctions, desmosomes, and gap junctions. The latter are particularly expressed in epithelial cells whose apical–basolateral polarization is critical to their transport function. Essentially, all cells are polarized, and the neuron represents a prime example of how cellular polarization results in the formation of functional domains. Nuclear domains control genetic regulation and transcription, and nuclear– cytoplasmic exchange and transport is mediated by the nuclear pore that delivers RNA to cytoplasmic domains that regulate RNA translation and degradation. Molecular determinants of cellular domains therefore include essentially all molecular components of the cell, including DNA, RNA, proteins, lipid, and glycans. Defining domains and understanding the molecular basis of their formation is central to understanding cellular function. Ivan R. Nabi

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CONTRIBUTORS John D. Aitchison, PhD, Institute for Systems Biology, Seattle, WA 98103-8904 François Bordeleau, Centre de recherche en cancérologie de l’Université Laval and Centre de Recherche du Centre Hospitalier de Québec (CRCHUQ), Quebec City, Quebec, Canada Keith Burridge, PhD, Department of Cell and Developmental Biology and Lineberger Cancer Center, University of North Carolina, Chapel Hill, NC 27599 Kendra L. Cann, Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Marcelino Cereijido, MD, PhD, Center for Research and Advanced Studies, Department of Physiology, Biophysics and Neurosciences, Mexico City, Mexico Jesse T. Chao, Department of Cellular and Physiological Sciences, University of British Columbia, Vancouver, British Columbia, Canada Jared M. Churko, Department of Anatomy and Cell Biology, Dental Science Building, University of Western Ontario, London, Ontario, Canada Daniel Coombs, PhD, Department of Mathematics and Institute of Applied Mathematics, University of British Columbia, Vancouver, British Columbia, Canada Dale Corkery, Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Raibatak Das, PhD, Department of Mathematics and Institute of Applied Mathematics, University of British Columbia, Vancouver, British Columbia, Canada Graham Dellaire, PhD, Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada James W. Dennis, PhD, Samuel Lunenfeld Research Institute, Mount Sinai Hospital, Toronto, Ontario, Canada Michel Desjardins, PhD, Département de pathologie et biologie cellulaire, Université de Montréal, Montreal, Quebec, Canada Leonard J. Foster, PhD, Centre for High-Throughput Biology and Department of Biochemistry & Molecular Biology, University of British Columbia, Vancouver, British Columbia, Canada Takahiro K. Fujiwara, PhD, Center for Meso-Bio Single-Molecule Imaging (CeMI) and Institute for Integrated Cell-Material Sciences (iCeMS), Kyoto, Japan Thierry Galvez, PhD, Max Planck Institute for Molecular Cell Biology and Genetics MPI-CBG, Dresden, Germany Stéphane Gilbert, PhD, Centre de Recherche en Cancérologie de l’Université Laval and Centre de Recherche du Centre Hospitalier de Québec (CRCHUQ), Quebec City, Quebec, Canada Jennifer S. Goldman, Center for Neuronal Survival, Montreal Neurological Institute, McGill University, Montreal, Quebec, Canada xi

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CONTRIBUTORS

Guillaume Goyette, PhD, Département de pathologie et biologie cellulaire, Université de Montréal, Montreal, Quebec, Canada Jody Groenendyk, PhD, Department of Biochemistry, School of Molecular and Systems Medicine, University of Alberta, Edmonton, Alberta, Canada Laura K. Hilton, Department of Molecular Biology & Biochemistry, Simon Fraser University, Burnaby, British Columbia, Canada Tom C. Hobman, PhD, Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada Ziya Kalay, PhD, Center for Meso-Bio Single-Molecule Imaging (CeMI), Institute for Integrated Cell-Material Sciences (iCeMS), Kyoto, Japan Jonathan A. Kelber, PhD, UCSD School of Medicine, Department of Pathology and Moores Cancer Center, La Jolla, CA 92093-0612 Timothy E. Kennedy, PhD, Center for Neuronal Survival, Montreal Neurological Institute, McGill University, Montreal, Quebec, Canada Richard L. Klemke, PhD, UCSD School of Medicine, Department of Pathology and Moores Cancer Center, La Jolla, CA 92093-0612 Keli Kolegraff, Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, GA 30322 Akihiro Kusumi, PhD, Center for Meso-Bio Single-Molecule Imaging (CeMI), Institute for Integrated Cell-Material Sciences (iCeMS) and Research Center for Nano Medical Engineering, Institute for Frontier Medical Sciences, Kyoto University, Kyoto, Japan Dale W. Laird, PhD, Department of Anatomy and Cell Biology, Dental Science Building, University of Western Ontario, London, Ontario, Canada Bettina Lechner, PhD, Department of Botany, University of British Columbia, Vancouver, British Columbia, Canada Christopher J.R. Loewen, PhD, Department of Cellular and Physiological Sciences, University of British Columbia, Vancouver, British Columbia, Canada Anne Loranger, PhD, Centre de recherche en cancérologie de l’Université Laval and Centre de Recherche du Centre Hospitalier de Québec (CRCHUQ), Quebec City, Quebec, Canada Daniel Louvard, PhD, CNRS, Institut Curie, Paris, France Normand Marceau, PhD, Centre de recherche en cancérologie de l’Université Laval and Centre de Recherche du Centre Hospitalier de Québec (CRCHUQ), Quebec City, Quebec, Canada Marek Michalak, PhD, Department of Biochemistry, School of Molecular and Systems Medicine, University of Alberta, Edmonton, Alberta, Canada Jennifer S. Morrison, Department of Mathematics and Institute of Applied Mathematics, University of British Columbia, Vancouver, British Columbia, Canada Ivan R. Nabi, PhD, Department of Cellular and Physiological Sciences, Life Sciences Institute, University of British Columbia, British Columbia, Canada Porfirio Nava, Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, GA 30322

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CONTRIBUTORS

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W. James Nelson, PhD, Department of Biology, Stanford University, Stanford, CA 94305 Asma Nusrat, PhD, Department of Pathology and Laboratory Medicine, Emory University School of Medicine, Atlanta, GA 30322 Henry Parker, PhD, Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada Nancy Philp, PhD, Department of Pathology, Anatomy and Cell Biology, Jefferson Medical College, Thomas Jefferson University, Philadelphia, PA 19107 Christopher Ptak, PhD, Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada Lynne M. Quarmby, PhD, Department of Molecular Biology & Biochemistry, Simon Fraser University, Burnaby, British Columbia, Canada Andreas S. Reichert, PhD, CEF Makromolekulare Komplexe, Mitochondriale Biologie, Fachbereich Medizin, Goethe-Universität Frankfurt am Main, Frankfurt am Main, Germany Sylvie Robine, PhD, CNRS, Institut Curie, Paris, France Enrique Rodriguez-Boulan, MD, Dyson Vision Research Institute, Departments of Ophthalmology and Cell Biology, Weill Medical College of Cornell University, New York, NY 10065 Liora Shoshani, PhD, Center for Research and Advanced Studies, Department of Physiology, Biophysics and Neurosciences, Mexico City, Mexico Radu V. Stan, MD, Department of Pathology, Dartmouth Medical School, One Medical Center Drive, Lebanon, NH 03756 James R. Thieman, Department of Cell Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA 15261 Caitlin Tolbert, Department of Cell and Developmental Biology, University of North Carolina, Chapel Hill, NC 27599 Christopher P. Toret, PhD, Department of Biology, Stanford University, Stanford, CA 94305 Linton M. Traub, PhD, Department of Cell Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA 15261 Dan Tse, MD, Department of Pathology, Dartmouth Medical School, One Medical Center Drive, Lebanon, NH 03756 Florent Ubelmann, PhD, CNRS, Institut Curie, Paris, France Geoffrey O. Wasteneys, PhD, Department of Botany, University of British Columbia, Vancouver, British Columbia, Canada Richard W. Wozniak, PhD, Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada Marino Zerial, PhD, Max Planck Institute for Molecular Cell Biology and Genetics MPI-CBG, Dresden, Germany Michael Zick, Adolf-Butenandt-Institut für Physiologische Chemie, Ludwig-MaximiliansUniversität München, München, Germany

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PART

I

MEMBRANE DOMAINS M

E MBR ANE BI L AYE R S form hydrophobic dividers within the aqueous environment that exists both within and without the cell. The plasma membrane surrounds the cell, segregating the intracellular milieu from the outside environment. Domain organization of the plasma membrane is based not only on lipidand protein-based interactions but also on the organization of the underlying actin cytoskeleton that impacts on molecular dynamics and function in the membrane (Chapter 1). Plasma membrane domains include not only clathrin-coated pits (Chapter 2), caveolae (Chapter 3), and less well-characterized lipid rafts (Chapters 4 and 5) but also cell-substrate adhesions (Chapter 17), cell–cell junctions (Chapters 18–20), and specialized polarized cellular domains (Chapters 13, 15, 21, and 22). Molecular transport across the plasma membrane and exchange with the extracellular milieu is the key to cellular functionality. It is mediated in large part by endocytosis via clathrin-coated pits (Chapter 2) and also, as described in later sections, by Golgi secretion (Chapter 8), exosomes (Chapter 9), transporters (Chapter 21), and gap junctions (Chapter 20). How lipids organize domains is a subject of intense investigation that has focused primarily on the role of lipid rafts (Chapters 1, 4, and 5). The underlying cytoskeleton controls and contributes to molecular dynamics in the plane of the membrane and to downstream signaling (Chapter 1; also Chapter 12), and the tools available to study and model membrane domain organization in living cells (Chapters 1 and 5) represent key elements of future research. Importantly, membrane domain organization is relevant not only to the plasma membrane but also to membranes of intracellular organelles (Chapters 6–11).

Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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CH A P T E R

1

CYTOSKELETON-INDUCED MESOSCALE DOMAINS Ziya Kalay Takahiro K. Fujiwara Akihiro Kusumi

DEFINITION The cell is much more than just a bag of protein juice. Indeed, many mechanisms exist in a living cell to keep its contents well organized. One of the most important apparatuses that the cell utilizes to organize the cytoplasm is the cytoskeleton. Therefore, the influence of the cytoskeleton on the two-dimensional fluid of the plasma membrane is an interesting subject. In this chapter, we will focus on this issue, and review the literature about the membrane domains delimited by the part of the actin-based cytoskeleton that is closely opposed to the cytoplasmic surface of the plasma membrane. This part of the cytoskeleton is referred to as the membrane skeleton. Due to its close association with the cytoplasmic surface of the plasma membrane, the membrane-skeleton meshwork directly influences the functions of the plasma membrane. As a consequence of the membrane-skeleton meshwork, the plasma membrane is effectively partitioned into mesoscale domains, or compartments, with sizes varying between 30 and 250 nm (with the exception of the larger 750-nm domain in the doubly nested compartments in normal rat kidney [NRK] cells; Fujiwara et al. 2002). We emphasize the characteristic size of these compartments, one to several hundred nanometers, that falls in the mesoscale, where collective dynamics of molecules play critical roles. At this size scale, the number of molecules in the system is insufficient for thermodynamics to hold, but is still too big to be tractable for quantum mechanics. In the plasma membrane, there are three types of major mesoscale domains (mesodomains): (1) membrane compartments delineated by the actin-based membrane skeleton; (2) raft domains, where specific proteins, glycosphingolipids, and cholesterol are concentrated; and (3) the protein oligomer domains. In this chapter, we will concentrate on the membraneskeleton-induced membrane compartments. Membrane lipids and proteins are both temporarily trapped in these membrane compartments with residency times between 1 ms and 1 second (Kusumi et al. 2005). Namely, the two key functional elements of the membrane are both influenced by the membrane skeleton, and a growing number of findings support the involvement of the membrane skeleton in many membrane processes (Gaidarov et al. 1999; Nakada et al. 2003; O’Connell et al. 2006; Lajoie et al. 2007; Chung et al. 2010; Treanor et al. 2010). In this respect, one of our main goals in this chapter is to provide an account of the various functions of these membrane-skeleton-based mesodomains. Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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CHAPTER 1

CYTOSKELETON-INDUCED MESOSCALE DOMAINS

HISTORICAL PERSPECTIVE One of the most important characteristics of the plasma membrane is that it is a twodimensional liquid. The fluid mosaic model proposed by Singer and Nicolson (1972) successfully accounted for many properties of the plasma membrane. In fact, this model is still widely believed to represent the basic structure of the plasma membrane (and intracellular membranes) of all living cells on earth. However, it fails to answer two basic questions, which have puzzled scientists for the past three decades: 1. Why do the membrane proteins and lipids diffuse faster in artificial membranes than in the cellular plasma membrane by a factor of ∼20 (ranging from 5 to 50)? (Murase et al. 2004) 2. How do molecular complexes become immobilized on the cell surface or diffuse at surprisingly lower rates, as compared with single molecules? (Kusumi et al. 2005)

Early Fluorescence Recovery After Photobleaching (FRAP) Observations Early measurements of diffusion coefficients using FRAP revealed significantly slower diffusion in the plasma membrane, as compared with that in artificially reconstituted membranes, by a factor of ∼20 (ranging from 5 to 50) (summarized by Murase et al. 2004). Some of these findings suggested that the discrepancy might be due to interactions between the cytoskeleton and the plasma membrane. In 1980, Sheetz et al. found that the diffusion coefficient of band 3 proteins in mouse erythrocyte mutants lacking the spectrin network is an order of magnitude higher than that in normal cells. In these cells, the membrane skeleton is formed by the spectrin network, and therefore, the results obtained with these mutant cells suggested that the membrane skeleton directly interferes with the diffusion of membrane proteins. For some other early examples of FRAP measurements that detected significantly reduced diffusivity in the plasma membrane (but not rotational diffusion; Tsuji et al. 1988), see the following references: Axelrod et al. (1976), Golan and Veatch (1980), Chang et al. (1981), Tsuji and Ohnishi (1986), and Tsuji et al. (1988).

Early Single-Molecule Observations: Membrane-Skeleton Fence Model for Transmembrane Proteins The advent of single-molecule imaging methods, such as single-particle tracking (SPT) and single fluorescent-molecule tracking (SFMT), allowed researchers to image and track single molecules in the plasma membrane of living cells. In 1994, by using SPT techniques, Sako and Kusumi observed that transferrin receptor and α2-macroglobulin receptor undergo a peculiar kind of motion, which is characterized by temporary confinement of the molecule in a bounded region of the membrane, with an average area of 0.25 μm2, interrupted by rare hops into adjacent, yet still temporarily confining, regions. This behavior was termed hop diffusion, as the membrane molecules seemed to be hopping between adjacent membrane compartments and diffusing freely within a compartment. Subsequent experiments showed that all of the transmembrane proteins in every cell type observed displayed hop diffusion as well. As the single-molecule imaging technique became more popular, a wealth of data started to accumulate that needed to be properly analyzed. The single-particle trajectories that exhibit hop diffusion have been analyzed by fitting appropriate models to them (see also Chapter 5 for further discussion of modeling single-particle trajectories). An early

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HISTORICAL PERSPECTIVE

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theoretical treatment by Powles et al. (1992) considered the diffusion of a particle trapped between partially permeable barriers arranged periodically in space. Here, exact mathematical expressions for some transport quantities in the system were obtained. In more recent treatments by Kenkre et al. (2008) and Kalay et al. (2008), exact formulas for the time-dependent mean square displacement and diffusion coefficient for a particle moving in a similar compartmentalized space were acquired, and the effects of disordered fence strengths and compartment sizes were predicted. The results of these works have successfully been used to deduce compartment sizes from single-particle trajectories recorded in the plasma membrane. Based on these findings, the membrane-skeleton fence model was proposed (Sako and Kusumi 1994; Kusumi and Sako 1996), in which the plasma membrane is effectively partitioned into mesoscale compartments. In this model, the membrane skeleton forms a meshwork near the cytoplasmic part of the plasma membrane with which the cytoplasmic domains of transmembrane proteins can interact. As a result of this interaction, transmembrane proteins are temporarily confined in membrane compartments induced by the membrane-skeleton meshwork. Transmembrane proteins can hop between adjacent compartments if the distance between the meshwork and the membrane becomes large enough, or if the meshwork temporarily and locally dissociates. See Figure 1.1 (top) for an illustration of these ideas. This new, compartmentalized view of the plasma membrane was further supported by an atomic force microscopy study of its cytoplasmic surface (Takeuchi et al. 1998), which produced similar estimates for the spectrin membrane-skeleton mesh size to those obtained by SPT of band 3 proteins in human erythrocyte ghosts (Tomishige et al. 1998).

Super-Speed Single-Molecule Imaging: Hop Diffusion of Phospholipids as well as Transmembrane Proteins and Anchored-Protein Picket Model Another striking discovery was made in 2002. Fujiwara et al. (2002) demonstrated that even phospholipids, which are the most basic molecular species for membrane formation, undergo hop diffusion between compartments with sizes similar to those detected by protein hop diffusion. In previous single-molecule observations, images were obtained at video (30 frames/s) or slower rates. However, the characteristic time for lipid residency within a compartment is generally much shorter than the time between two consecutive image frames employed in those observations. Therefore, the detection of membrane compartments for lipids had to wait for the development of ultrafast methods for performing SPT. Fujiwara et al. (2002) performed their measurements at a rate of 40,000 frames/s, the fastest single-molecule imaging ever made, and have increased the rate even further to 160,000 frames/s. The discovery of lipid molecules undergoing membrane-skeleton-dependent hop diffusion in the outer leaflet of the plasma membrane was very surprising, since they only reach halfway through the membrane and lack the cytoplasmic domains of transmembrane proteins. To explain the hop diffusion of phospholipids in the outer leaflet, the anchoredprotein picket model was proposed (Fujiwara et al. 2002; Kusumi et al. 2005), where lipids are envisaged to interact with the membrane skeleton indirectly through picket proteins that are attached to the membrane skeleton, as shown in Figure 1.1 (middle). Several different mechanisms for the interaction between lipids and protein pickets were proposed. First, the picket can block the passage of a lipid molecule due to volume exclusion, causing steric hindrance. Second, the lipid molecules can be packed more in the immediate vicinity of the picket than the bulk membrane, and thus the free area available for a lipid molecule

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CHAPTER 1

CYTOSKELETON-INDUCED MESOSCALE DOMAINS

Membrane-skeleton fence model Transmembrane protein

START Actin filament (membrane skeleton)

Anchored-protein picket model START

Phospholipid

Anchored-protein pickets

Actin filament (membrane skeleton)

Picket-induced slowdown effect Anchored-protein picket Phospholipid

Actin filament

Low diffusivity domain

Figure 1.1. Schematic illustrations of the membrane-skeleton fence and anchored-protein picket models, and the picket-induced slowdown effect. According to the membrane-skeleton fence model (top), transmembrane proteins with protrusions in the cytoplasm are temporarily confined in membrane-skeleton-based compartments formed by a dynamic meshwork of actin filaments. Consequently, these proteins undergo hop diffusion, as illustrated by the color-coded trajectory. Lipids in the upper leaflet of the plasma membrane can also undergo hop diffusion even though they lack cytoplasmic domains. In the anchored-protein picket model (middle), the presence of pickets, proteins that are anchored to the boundaries of membrane-skeleton-based compartments, restricts the motion of all membrane molecules including lipids. As explained in the text, the presence of an immobile protein may locally reduce the diffusivity of lipids. This so-called picket-induced slowdown effect is illustrated in the bottom figure. Here, a lipid molecule in the close proximity of a picket, which we refer to as the low diffusivity domain, diffuses at a slower rate as compared with those far from the picket, as indicated by the smaller step size in the particle’s trajectory.

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ELECTRON TOMOGRAPHY OF THE THREE-DIMENSIONAL STRUCTURE

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to move into decreases, leading to diminishing mobility around the picket (Sperotto and Mouritsen 1991; Almeida et al. 1992). Third, lipid molecules can experience hydrodynamic slowing near an immobilized picket molecule, similar to a fluid particle that moves close to the boundary of a container (Bussell et al. 1994, 1995; Dodd et al. 1995). Therefore, lipid diffusivity can be reduced in the vicinity of a picket protein for different reasons, a phenomenon we call the picket-induced slowdown effect. A schematic illustration of the low diffusivity domain due to this effect is shown in Figure 1.1 (bottom). When many such immobilized picket proteins are aligned along the membrane-skeleton fence, the membrane molecules cannot easily pass through the compartment boundaries, and thus become temporarily confined within a compartment. Namely, in the anchored-protein picket model, the entire plasma membrane is partitioned into compartments by transmembrane protein pickets, lining the membrane-skeleton fence. By performing Monte Carlo simulations that account for the reduction in free area, the picket density along the compartment boundary necessary for reproducing the observed hop diffusion of lipids was estimated to be 20–30% (Fujiwara et al. 2002). Namely, the compartment boundaries do not have to be closed off by concentrating the transmembrane protein pickets there, but when only 1/5–1/3 of the boundary is occupied by picket proteins, it would be sufficient to induce temporary trapping of lipids within a compartment. It is important to note that the effect of anchored-protein pickets is not limited to that of mere protein crowding. Several studies have indicated that if the obstacles are mobile, then they do not lead to a significant decrease in the diffusion coefficient of the rest of the mobile particles. Monte Carlo studies by Saxton (1987, 1990), which did not consider hydrodynamic effects, demonstrated that mobile obstacles do not reduce the diffusivity as much as their immobile counterparts. Later on, Bussell et al. (1994, 1995) and Dodd et al. (1995) showed that the inclusion of hydrodynamic effects did not change this conclusion, provided that the lipids can be assumed to form a continuum. This assumption would not hold in cases where the protein diameter is comparable with the diameter of lipids (we will address this point later in this chapter), and one might consider the possibility that the details of collective protein–lipid dynamics on the mesoscale may determine the properties of the system. However, since the diffusion coefficients of proteins and lipids within a compartment are similar to those measured in liposomes, reconstituted membranes, and membrane blebs (see figs. 1 and 2 in Fujiwara et al. 2002 and figure 8 in Murase et al. 2004), the immobilization of obstacles (pickets) seems to be a key factor in the picket-induced slowdown effect. In addition, the proteins do need not be continually anchored in order to function as effective pickets. If the proteins are immobilized for more than ∼10 μs at a time, then this may suffice to produce the observed hindrance of lipid diffusion.

ELECTRON TOMOGRAPHY OF THE THREE-DIMENSIONAL STRUCTURE OF THE CYTOPLASMIC SURFACE OF THE PLASMA MEMBRANE Perhaps the most direct evidence for the membrane-skeleton-based partitioning of the plasma membrane was obtained in 2006 by Morone et al., who imaged the three-dimensional structure of the cytoplasmic surface of the plasma membrane by electron tomography. Here, electron tomography was first applied to platinum-replicated samples: the threedimensional structure of the cytoplasmic surface of the plasma membrane was reconstituted from the platinum-coated membrane specimen, prepared with minimal intrusion, by using the freeze-etching technique. These images clearly demonstrated that the membrane

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CHAPTER 1

CYTOSKELETON-INDUCED MESOSCALE DOMAINS

NRK

Electron Tomography SPT

FRSK Finish

Start Finish Start

1 µm NRK cell Phospholipid (DOPE)

Figure 1.2. Electron microscopic images of the membrane skeleton of NRK (upper left) and fetal rat skin keratinocytes (FRSKs) (lower left) cells, where the scale bars in the main figures correspond to 100 nm. The inset in the lower left image highlights the 5.5-nm periodic striped pattern characteristic of actin filaments, showing that the membrane skeleton is primarily composed of actin filaments (scale bar 50 nm). Note that both of these images also contain clathrin-coated pits, which are distinguished by their lattice structure (from Morone et al. 2006). In the lower right corner, the trajectory of a gold-tagged phospholipid (DOPE) in an NRK cell, obtained by SPT at 40,000 frames/s, is displayed. The trajectory was color coded after performing a quantitative analysis that detects jumps between adjacent domains. The histograms in the upper right corner show that the size distribution of the membrane-skeleton meshes directly contacting the cytoplasmic surface of the plasma membrane, as determined by electron tomography, is very close to that of the compartments determined from the DOPE diffusion data in either NRK or FRSK cells, whereas the distributions for these two cell types are entirely different (in part from Fujiwara et al. 2002).

skeleton entirely covers the cytoplasmic surface of the plasma membrane, except for certain membrane domains, such as clathrin-coated pits (CCPs) (Chapter 2), caveolae (Chapter 3), and focal adhesions (Chapter 17), that the membrane-skeleton meshwork is primarily composed of actin filaments since almost every filament exhibited a distinct striped pattern with a 5.5-nm periodicity (see Fig. 1.2 for typical electron micrographs, although they are normal two-dimensional images), and that some of the meshwork is located as close as within a nanometer of the plasma membrane. Based on these images, the size distribution of the actin skeleton mesh on the cytoplasmic surface of the plasma membrane was obtained and found to agree well with the compartment sizes found by analyzing phospholipid hop diffusion as revealed by SPT. This result strongly supports the partitioning of the plasma membrane by membraneskeleton fences and transmembrane protein pickets lining the fence.

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ANSWERS TO THE TWO THREE-DECADE-OLD ENIGMAS The fence–picket model gives straightforward answers to the two three-decade-old questions raised in the beginning of this section. 1. Why do membrane proteins and lipids diffuse faster in artificial membranes than in the cellular plasma membrane by a factor of ∼20 (ranging from 5 to 50)? As stated in the previous paragraph, the diffusion coefficient within a compartment is not small, as compared with that in artificial membranes. However, if the diffusion coefficient is measured on much greater scales, for example, a FRAP spot size of ∼500 nm, or at slower observation rates, for example, single-molecule tracking observed at a rate of 30 frames/s or slower, then the diffusion appeared to be slow because what was observed is the apparent diffusion coefficient, which is affected by the presence of compartment boundaries. It takes time to hop from one compartment to an adjacent one, which makes the macroscopic diffusion of lipids and proteins in the plasma membrane very slow. 2. How do molecular complexes become immobilized on the cell surface or diffuse at surprisingly lower rates, as compared with single molecules? This can be explained by the “oligomerization-induced trapping” effect of the fences and pickets. Monomers of membrane molecules may hop across the intercompartment boundaries with relative ease, but upon forming oligomers or molecular complexes, the entire complex, rather than single molecules, has to hop across the picket–fence all at once, and therefore, these complexes are expected to hop across the boundaries at much slower rates. In addition, due to the avidity effect, molecular complexes are more likely to be bound to the membrane skeleton, perhaps temporarily, which also induces (temporary) immobilization or trapping of oligomers and molecular complexes. Such enhanced confinement and binding effects induced by oligomerization or molecular complex formation are collectively termed oligomerization-induced trapping (Kusumi and Sako 1996; Iino et al. 2001; Kusumi et al. 2005). One might argue that even in the absence of membrane-skeleton fences and anchored pickets lining the fences, the oligomerization of membrane proteins could greatly reduce the diffusion coefficient. However, experimental and theoretical studies clearly showed that this does not occur with transmembrane proteins that contain approximately three or more membrane-spanning α-helices (Saffman and Delbrück 1975; Peters and Cherry 1982; Vaz et al. 1982; Liu et al. 1997; Gambin et al. 2006). Based on these studies, we predict that tetramer formation from monomers (a twofold increase in radius) will only decrease the diffusion coefficient by a factor of 1.1, and even 100mers (a 10-fold increase in radius) will have a diffusion rate reduced by only a factor of less than 2 from that of monomers. Therefore, the large reductions of the diffusion coefficient, upon oligomerization or molecular complex formation, clearly indicate that the plasma membrane cannot be considered as a two-dimensional fluid continuum, and that the fence–picket model is consistent with the experimental observations and theoretical predictions. In artificial membranes without the membrane skeleton, Liu et al. (1997) and Gambin et al. (2006) reported decreases of the diffusion coefficient by a factor of ∼2 when the probe hydrodynamic diameter (in the cross section of the transmembrane domain parallel to the membrane surface) was increased by a factor of ∼2 from the original diameter of ∼0.5 nm. In this spatial scale, the probe (solute) size is comparable with the solvent

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molecular size, and the continuum fluid model is no longer correct. Therefore, the translational diffusion coefficient of the test particle strongly depends on its diameter on this particular spatial scale. However, this should not be confused with the reduction in the diffusion coefficient when single-pass transmembrane proteins, such as many receptor molecules, form oligomers in the plasma membrane. The size-dependent decrease of the diffusion coefficient observed in artificial membranes (Liu et al. 1997; Gambin et al. 2006) occurs only for test particles of ∼0.5 nm in diameter undergoing virtually simple diffusion, which exhibits a monomer diffusion coefficient of ∼10 μm2/s. In the plasma membrane, the effective diffusion coefficients of single-pass transmembrane protein monomers are generally ∼0.2 μm2/s, which are already slower by a factor of 50 than those found in artificial membranes. Oligomerization of such a single-pass transmembrane protein tends to decrease the diffusion coefficient by a factor of ∼2, but the weak resemblance of these reduction factors is merely incidental. The oligomerization-induced reduction of the diffusion coefficient in the plasma membrane was not due to changes in protein interaction with membrane lipids, as was found in artificial membranes, but rather to the compartment boundaries. Evidence for this statement comes from high-speed single-molecule tracking data. Single-pass transmembrane proteins generally exhibit hop diffusion between compartments, with a microscopic diffusion coefficient within a compartment of 5–10 μm2/s and a residency time of 20–100 ms, and oligomerization lengthened the residency time without affecting the compartment size and the microscopic diffusion coefficient within a compartment (Murase et al. 2004; Kusumi et al. 2005). This clearly indicates the necessity for being careful in interpreting long-range, slow-speed diffusion measurements, for carrying out high-speed single-molecule tracking, and for careful attention to loosely applying the results of Liu et al. (1997) and Gambin et al. (2006) to the observations made in the plasma membrane. In addition to the corralling effect of the membrane skeleton and associated transmembrane protein pickets, three major factors are frequently discussed in the literature to account for the slower diffusion of membrane molecules in the cellular plasma membrane, observed by methods with low spatiotemporal resolutions: the crowding effect of transmembrane proteins, the trapping or exclusion effect of raft domains, and the ordering effect of cholesterol. All three of these factors can lead to a decrease in molecular mobility in the plasma membrane, but even if all of these three factors are combined, a decrease in the diffusion coefficient by a factor of 20 cannot be explained (perhaps, a factor of 2 could be explained). Some of the phenomena including the immobilization of receptor complexes can only be explained by considering the effects of the membrane skeleton.

MEMBRANE-SKELETON-BASED MESODOMAINS ARE NOT DETECTED IN EVERY STUDY A number of studies failed to detect the effect of the membrane skeleton on the diffusivity of membrane molecules. Two FRAP studies (Schmidt and Nichols 2004; Frick et al. 2007) found that disruption of the cortical actin did not lead to significant changes in the diffusivity of some membrane proteins and a lipid analog. However, experiments involving drug-induced actin (de)polymerization revealed various effects, depending on the drug concentration, the treatment duration, and the cell type, and thus comparisons of the results are difficult (Vrljic et al. 2005; Umemura et al. 2008). For the intricate cell reactions to such treatments, see Suzuki et al. (2005). Furthermore, Fujiwara et al. (2002) and Murase

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et al. (2004) found that without directly observing the compartment size and the residency time within a compartment, detecting drug-induced changes will be difficult. In studies using fluorescence correlation spectroscopy (FCS) (Wawrezinieck et al. 2005; Lenne et al. 2006) and stimulated emission depletion–FCS (STED-FCS) (Eggeling et al. 2009), phospholipid analogs were observed to diffuse freely. To detect hop diffusion in FCS experiments, the focal area of the laser should be less than the characteristic compartment size. In STED-FCS (Eggeling et al. 2009), the diameter of the laser beam can be as small as 30 nm, which is comparable with the smallest compartment sizes reported in the literature (Murase et al. 2004). Therefore, a further decrease in the diameter of the laser beam seems to be necessary for determining the true nature of molecular diffusion by FCS techniques. In addition to these results obtained by measuring the signal from many molecules simultaneously, several single-molecule/particle studies also did not find hop diffusion. For instance, Wieser et al. (2007, 2008) found simple Brownian diffusion of lipids and a glycosylphosphatidylinositol (GPI)-anchored protein CD59, and Crane and Verkman (2008) reported the observation of freely diffusing aquaporin-1 water channels. In these studies, the rate at which the images were acquired was 2000 Hz or less, resulting in determination of the positions of single molecules at relatively sparse time points such that it would simply be impossible to detect fast hop diffusion among mesoscale compartments. In fact, careful attention must always be paid to the camera frame rates relative to the molecular hop frequency in each cell type when interpreting the single-molecule tracking results, since this is a key factor in the ability to detect confinement. The residency time of a molecule in a compartment may vary between one to hundreds of milliseconds, and will be shorter for lipids (Fujiwara et al. 2002; Murase et al. 2004) and longer for proteins (Tomishige et al. 1998; Tomishige and Kusumi 1999). Especially for lipids, to obtain statistically significant results, single-molecule/particle tracking must be performed at very high speeds. At a camera speed of 40,000 Hz, which was used by Fujiwara et al. to detect the hop diffusion of lipids, the time between two consecutive frames is 25 μs, thus allowing one to obtain 40 data points even for short residency times such as 1 ms (Fujiwara et al. 2002; Murase et al. 2004). Readers are referred to Murase et al. (2004) for the residency times of an unsaturated phospholipid, 1,2-dioleoyl-sn-glycero-3phosphoethanolamine (DOPE), in a number of different cells. The residency time divided by 40 essentially provides a good estimate of the inverse frame rate that is necessary to detect hop diffusion. For example, Sahl et al. (2010) reported that they failed to detect the hop diffusion of a fluorescent phospholipid analog, but instead found alternating temporary entrapment within a domain (30% of the observed duration) and simple Brownian diffusion (70%) in the plasma membrane of PtK2 cell, using an SFMT method with a time resolution of 0.5 ms. Using the same cell line, the average residency time of a phospholipid within a compartment was found to be ∼1 ms (Fujiwara and Kusumi, unpublished observations). This result indicates that, to detect the hop diffusion of phospholipids, the time resolution of the instrument must be better than 0.025 ms, suggesting that the results obtained by Sahl et al. (2010) are due to the lack of time resolution (on average, they made only two coordinate determinations during the average residency time of 1 ms): when a molecule stays in a compartment much longer than average, even the 0.5-ms observation rate would be sufficient to detect confinement, but when the molecule stays in compartments for shorter periods, a 0.5-ms resolution would be insufficient to detect confinement, thus reporting simple Brownian diffusion.

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MOLECULAR COMPOSITION OF THE MEMBRANE SKELETON As shown previously (Heuser and Kirschner 1980; Hirokawa and Heuser 1981; Hirokawa et al. 1982; Morone et al. 2006; Hanson et al. 2008), the membrane skeleton mainly consists of actin filaments (see also Chapter 12). Actin filaments grow at their fast growing ends, which interact with the membrane (Small et al. 1978; Wang 1985). Therefore, the membrane compartments should depend on the proteins involved in actin polymerization, including the Arp2/3 complex, Wiskott–Aldrich syndrome protein (WASP), suppressor of cyclic AMP receptor (SCAR)/WASP family verprolin-homologous protein (WAVE), cortactin, and the formin family of proteins (Pollard 2007; Campellone and Welch 2010), as well as those involved in actin depolymerization such as the cofilin family of proteins (Bernstein and Bamburg 2010). Actin filaments and the plasma membrane are also linked by proteins that can laterally bind to actin, including ponticulin; the ezrin/radixin/moesin family of proteins; the villin–gelsolin superfamily proteins; the epithelial protein lost in neoplasm, filamin, dystrophin and utrophin, and tropomyosin; and the myosin family of proteins (Morone et al. 2008). Furthermore, actin filaments bind to various other membrane-associated proteins and lipids with low affinities, to facilitate dynamic associations with the membrane. This easy binding to and dissociation from the membrane would be needed for the dynamic regulation of the diffusivities of various molecules in the plasma membrane. Despite such low affinities, the binding sites would be numerous, making the dynamic binding very effective, although rendering their binding undetectable in general pull-down assays. As indicated in a previous section, transmembrane proteins transiently bound to the membrane skeleton would serve well as a diffusion barrier, if the anchored durations are greater than ∼10 μs. In normal erythrocytes, the organization of the membrane skeleton has been studied extensively and is known to be different from those of other cell types. The erythrocyte membrane-skeleton mesh is composed of a network of spectrin tetramers, acting as fences, which are cross-linked by protein complexes formed by short actin filaments, adducin, band 4.1, and the transmembrane protein glycophorin C, anchoring the entire network to the plasma membrane (Goodman et al. 1988; Bennett 1990; Anong et al. 2009). Spectrin tetramers form by the tail-to-tail linkage of spectrin dimers, and the dimers and the tetramers exist in a very dynamic equilibrium. For the passage of transmembrane proteins across the spectrin fences, a spectrin tetramer dissociation gate model (SPEQ gate model) was proposed. In the model, the dissociation of a spectrin tetramer into dimers entails transient gate opening, allowing transmembrane molecules to pass through the compartment boundaries (Tomishige et al. 1998). Malfunctions of the human erythrocyte membrane skeleton often lead to serious types of anemia, indicating the important roles played by the membrane skeleton in the functions, stability, and flexibility of the plasma membrane (Bennett and Healy 2008; Kodippili et al. 2009).

FUNCTIONS OF MEMBRANE-SKELETON-INDUCED MESODOMAINS Effects of the Membrane Skeleton on Signaling B-Cell Receptor (BCR) Signal Transduction: The Membrane Skeleton May Be Involved in the Formation of Receptor Oligomers and/or Engaged-ReceptorBased Lipid Rafts Several different mechanisms for the relationship between the actin cytoskeleton and BCR signaling have been proposed. In the oligomeric BCR complex model (Reth et al. 2000;

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Schamel and Reth 2000), disruption of BCR oligomers was considered to be required for signaling, and the role of the membrane skeleton in regulating the formation and maintenance of oligomers was discussed. In another approach, the effect of lipid rafts was emphasized (see also Chapters 4 and 5). In this model, BCR is considered to enter rafts only upon antigen binding, which is accompanied by the coalescence of smaller rafts. This in turn enables BCR to react with raft-associated kinases to start signaling (Tolar et al. 2005; Gupta et al. 2006; Gupta and DeFranco 2007; Sohn et al. 2008). The membrane skeleton can participate in the regulation of signaling as it restricts the growth and coalescence of lipid rafts. Recent observations by Treanor et al. (2010) showed that the treatment of cells with actin-modulating chemicals resulted in up to a 10-fold change in the diffusivity of BCR, suggesting that the membrane skeleton can control the mobility of these molecules. Furthermore, the expression level of ezrin, which is among the proteins that can form a link between the cytoskeleton and membrane proteins, also caused a change in BCR diffusivity by a factor greater than 3. This finding is also consistent with the anchored-protein picket model, since ezrin could act as a link between the protein pickets and the membrane skeleton. Moreover, the crosslinking of BCR and the dynamics of the membrane skeleton were correlated, and BCR signaling upon antigen binding leads to significant cytoskeletal reorganization (Fleire et al. 2006; Arana et al. 2008; Lin et al. 2008). Interestingly, alteration of the actin cytoskeleton also led to B-cell signaling comparable with that elicited by BCR cross-linking. Therefore, a clear link exists between the membrane skeleton and BCR signaling, implying that membrane mesodomains can play a critical role in this process, by controlling the distribution of membrane molecules.

Epidermal Growth Factor (EGF) Receptor (EGFR) and IgE-Fc (FC portion of Immunoglobulin E) Receptor Signal Transduction: The Membrane Skeleton May Increase the Receptor Oligomerization Rate and Promote Receptor Activity Together with the Galectin Lattice Lajoie et al. (2007) raised the question of how the three mechanisms of regulating the dynamics, clustering, distribution, and function of the EGFR are coordinated: (1) Galectin-3 is located on the extracellular surface of the plasma membrane and cross-links β1,6GlcNAc-branched N-glycans on cell-surface glycoproteins, including EGFR, to form a heterogeneous lattice; (2) caveolin-1 (Cav1), a major constituent of caveolae, acts as a negative regulator of growth factor signaling (Parton and Simons 2007), and in addition, Cav1 forms noncaveolar microdomains, which are likely to contain at least 15 Cav1 molecules (Parton et al. 2006); (3) membrane-skeleton fences and pickets suppress the diffusion of EGFR, and particularly that of the signal-capable receptor oligomers. Lajoie et al. (2007) showed that the extracellular galectin-3 lattice interacts with the N-glycans on EGFR and impedes its diffusion, which predominantly protects EGFR from loss to caveolae and Cav1 microdomains where EGFR signaling is suppressed. Disruption of the actin membrane skeleton with latrunculin A increased the mobile fraction of EGFR measured by FRAP, suggesting that the galectin-bound EGFR is further stabilized by the actin membrane skeleton. It is likely that galectin-3 cross-links EGFR to other actinassociated membrane glycoproteins or pickets, thus generating actin-stabilized signaling domains. Using quantum-dot-based SPT techniques, Chung et al. (2010) measured the diffusion coefficient of EGFR as a function of time. Assuming a relationship between diffusivity and molecular size, the authors inferred that EGFRs form transient oligomers, even in the absence of ligand, with lifetimes ranging from a few to a few tens of seconds. Interestingly, the probability of finding receptor dimers was reportedly higher at the

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periphery of EGFR-overexpressing A431 and BT20 cells, in an actin-dependent manner. The abundance of dimers along the cell periphery might be a consequence of the increased collision frequency of receptors confined in the mesodomains delimited by the membrane skeleton. Previous observations provided evidence for the increased density of actin filaments near the leading edge of migrating cells (Borisy and Svitkina 2000; Pollard et al. 2000). In this respect, another function of the membrane-skeleton-induced compartments could be promoting the formation of receptor complexes, and enabling the polarized response after ligand binding that is necessary in certain cellular processes such as chemotaxis. The involvement of raft domains induced by receptor engagement in signaling, by enhancing the recruitment of Lyn kinase and reducing the collisions with phosphatases, was shown in the case of the IgE-Fc receptor, FcεRI (Field et al., 1997; Wu et al. 2004; Young et al. 2005). In addition, the involvement of membrane-skeleton-delimited compartments was found by simultaneous observations of the quantum-dot-labeled FcεRI and green fluorescent protein (GFP)-tagged actin (Andrews et al. 2008). The diffusion rate of cross-linked receptor in cells with a disrupted actin cytoskeleton was significantly higher than that in intact cells, in accordance with oligomerization-induced trapping. Immobilization of cross-linked receptors at the onset of signaling (Menon et al. 1986) also enables the cell to remember the position of the stimulus for short periods (on the order of 10 seconds) and to perform localized responses. This is essential in certain processes, such as chemotaxis. Therefore, the actin meshwork is also involved in reliably responding to local changes in the environment (Kusumi and Sako 1996; Kusumi et al. 2005). In fact, receptor redistribution and clustering are key steps in many signal transduction pathways (Petrini et al. 2004; Minguet et al. 2007; Briegel et al. 2009; Nikolaev et al. 2010). Several reports have indicated the active roles played by the cytoskeleton in inhibiting (Wang et al. 2001; Boggs and Wang 2004) or enabling (Gomez-Mouton et al. 2001; Rodgers and Zavzavadjian 2001; Baumgartner et al. 2003) the redistribution/ clustering of membrane molecules. In oligodendrocytes, stimulation of the cells induced actin depolymerization that was followed by coclustering of membrane molecules, including myelin basic protein and galactosylceramide (Boggs and Wang 2004). However, when the actin filaments were artificially stabilized, coclustering was not observed. This observation is consistent with the presence of membrane-skeleton-based mesodomains, since lateral diffusion, which is necessary for coclustering, is hindered by the presence of actinbased membrane-skeleton fences and pickets. Kv2.1 Potassium Channels: The Membrane Skeleton Is Involved in Cluster Maintenance and Distribution FRAP and quantum-dot-based imaging revealed that Kv2.1 potassium channels form dynamic clusters with sizes and spatial distributions that are influenced by the actin-based membrane skeleton (O’Connell et al. 2006). Interestingly, the individual channels diffused freely within a cluster, suggesting that the clusters might actually be channels corralled by the membrane skeleton. This proposal was further supported by the results obtained upon disruption of the membrane skeleton by latrunculin A treatment, which resulted in a 10-fold increase in the average cluster area and a decrease in the number of clusters, indicating that the smaller clusters merged during the treatment. Furthermore, the spatial distribution of Kv2.1 channels in hippocampal neurons was also affected by the disruption of the membrane skeleton (O’Connell et al. 2006). In cells with an intact cytoskeleton, the channels were restricted to the cell body. However, after the latrunculin A treatment, the channels were found in the neurites as well as in the cell body. These observations suggest that the Kv2.1 clusters are maintained in membrane-

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skeleton-based mesodomains and that an actin-based mechanism is involved in determining the spatial distribution of Kv2.1 channels.

Micron-to-Cell-Sized Diffusion Barriers Generated by the Membrane-Skeleton Fences and Associated Pickets Numerous experiments have indicated that diffusion barriers, with sizes between a micron to several tens of microns, thus producing macroscopic plasma membrane domains, might be generated by the actin-based membrane-skeleton fences and the transmembrane picket proteins lining the fence. In the following, we will discuss a few such cases. A study on fish keratocytes revealed the presence of an F-actin-dependent lipid diffusion barrier, located at the leading edge of migrating cells (Weisswange et al. 2005). In boar sperm cells, observations indicated the presence of a diffusion barrier between the equatorial segment and the postacrosome, which prevents the passage of large molecular complexes. Based on the diffusivity measurements and the topographical properties of the cell surface revealed by atomic force microscopy, the authors suggested that a high concentration of anchored transmembrane proteins in the region could be involved in forming the barriers (James et al. 2004). The plasma membrane of the neuron consists of two distinct domains: the somatodendritic and axonal domains (Kobayashi et al. 1992; Winckler et al. 1999). Single-molecule imaging of a phospholipid revealed the developmental formation, by day 10 after birth in cultured hippocampal neurons (see also Chapter 22 on neuronal domains), of a diffusion barrier that blocks the diffusion of even phospholipids in the plasma membrane, in the boundary region between the two domains, called the initial segment (Nakada et al. 2003). Moreover, the barrier was formed by highly concentrating various transmembrane proteins and membrane skeletal proteins, including actin filaments and ankyrin, which bound to each other to create very dense rows of anchored pickets. This would effectively block diffusion, even that of phospholipids, in accordance with the anchored-protein picket model. More recently, Renner et al. (2009) made a surprising discovery concerning lipid diffusion, not necessarily involving diffusion barriers, in the synaptic membranes of mature hippocampal neurons. The authors showed that the lipids ganglioside GM1 and DOPE, and the artificial fluorescent lipid probe DiIC18, undergo confined diffusion in submicron-sized regions whose extent is proportional to molecular size, being the smallest for DOPE (∼110 nm). For a review on how singlemolecule techniques have played an important role in the advancement of synaptic biology, the readers are referred to Triller and Choquet (2008).

The Role of the Membrane Skeleton in CCP Formation during Endocytosis CCP-mediated endocytosis constitutes the major route for the uptake of nutrients and signaling ligands, as well as for the entry of viruses and toxins (see also Chapter 2 on CCPs). CCPs are very closely associated with or bound by the actin filaments in the membrane skeleton, as seen in the electron microscopic images in Figure 1.2. Consistent with this observation, the mobility of CCPs visualized by GFP-tagged clathrin light chain was found to be highly restricted to submicron regions (0.5–0.8 μm in diameter) in the plasma membrane, and the disruption of actin filaments by latrunculin B led to the increased mobility of CCPs (Gaidarov et al. 1999). In a different study (Ehrlich et al. 2004), the analysis of the experimental data suggested that the CCPs are preferentially formed in the so-called active domains of ∼400 nm in diameter, which are similar in size to the membrane-skeleton-based domains. Total internal reflection fluorescence

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microscopic studies (Merrifield et al. 2002; Yarar et al. 2005) have revealed that actin polymerization is required when CCPs are undergoing invagination and scission. These results suggest the involvement of the dynamic actin meshwork in CCP formation. However, others have shown that this is not always true, depending on the cell type (Fujimoto et al. 2000) and the choice of locations on the cell surface (Saffarian et al. 2009; upper surface vs. lower surface in contact with the coverslip). Thus, CCP researchers are still vigorously discussing the possible roles of both actin polymerization (to generate force for membrane deformation) and depolymerization (to remove the dense actin meshwork that would otherwise form a steric barrier) in the regulation of CCP formation.

Regulation of Lipid Rafts by the Membrane-Skeleton-Based Compartments Lipid domains/rafts are important, and difficult to detect, constituents of the plasma membrane (see also Chapters 4 and 5 on lipid rafts). They are considered to participate in many membrane processes, including the formation of platforms for signaling molecules. In resting cells, rafts could be highly dynamic entities with a wide range of diameters. In the literature, the estimated diameters of raft-associated molecule clusters have been reported to be 5–15, 20, 50–80, 120, and 700 nm. Many different structures and processes have been proposed to describe the generation and maintenance of rafts (Kusumi et al. 2004, 2010). Among these, membrane-skeleton-induced compartmentalization can be a major regulator of raft dynamics. Most transmembrane proteins are known to exclude cholesterol from their boundary regions. Therefore, an array of anchored-protein pickets would prevent the motion and growth of rafts in their vicinity. As a result, one would expect the raft size to be limited by the typical membrane-skeleton-delimited compartment size of 30–250 nm, which is quite consistent with experimental findings.

Reaction Kinetics: Increased Collision Rate due to the Presence of Membrane-Skeleton-Based Compartments When reactants are confined to a region with a characteristic size that is comparable with their mean free path, the frequency of collisions between them will increase (see, for instance, Saxton 2002 for a discussion). However, the overall efficiency of the reaction can increase or decrease depending on the details of the compartmentalization and the properties of the reactants (Melo et al. 1992). The collision frequency of molecules in the plasma membrane has not been directly measured. Nevertheless, as mentioned earlier in this section, at least one experiment in the live-cell plasma membrane suggested a modification in the oligomerization rate of EGFR by the increased density of the actin-based membrane skeleton (Chung et al. 2010). The presence of membrane-skeleton-based compartments has two competing effects on reaction kinetics. One of them is to increase the time required for the reactants to collide for the first time (if initially they are not in the same compartment), since the long-term diffusivity is reduced by compartmentalization. The other is a substantial decrease in the first recollision time, along with a significant increase in the collision frequency, when the reactants are within the same compartment. Together, these effects can selectively control the kinetics of reactions in the membrane. For instance, at low concentrations, chemically limited reactions that require many collisions would be promoted, whereas diffusion-limited reactions would be suppressed. A detailed analysis of the modifications of reaction kinetics due to membrane-skeleton-based compartments is in progress in our laboratories.

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Artificial Regulation of Membrane Molecular Dynamics Using Fabricated Mesostructures Recently, membrane-skeleton-induced mesodomains or compartments have inspired new lines of research. Using nanofabricated materials, structures similar to membrane mesodomains have been conducted. Several recent examples of such studies can be found in Mossman et al. (2005), Tsai et al. (2008), Kam (2009), Takimoto et al. (2009), and Salaita et al. (2010). Tsai et al. (2008) constructed supported lipid bilayers on a glass substrate patterned with 50-nm-wide chromium or titanium stripes, separated by 125–250 nm. The stripes were tall enough to act as perfect barriers to lipid diffusion, but they contained periodically placed gaps that allowed the lipids to pass. Using FRAP, the authors demonstrated that the characteristics of lipid diffusion on large and small spatial/temporal scales are very different, with the short-range/time diffusivity being much higher. Although this artificial system is quite different from the cellular plasma membrane, it is a good example of a controllable nano–meso structure that can mimic biological systems. In addition, Mossman et al. (2005) used artificial structures to manipulate the geometry of immunological synapses, the interfaces between T cells and their target cells (Grakoui et al. 1999; Yokosuka and Saito 2010), which provided valuable insight into the functions of membrane compartmentalization. Mossman et al. (2005) placed T cells on a supported antigen-containing lipid membrane that was partitioned into micron-sized compartments by either an artificial meshwork or more elaborate structures. As a result, the mobility of T-cell receptors was significantly modified. When the artificial structure was a meshwork, a strong correlation between the receptor activity and position was found, such that receptors at the periphery of the interface showed prolonged signaling activity, as compared with the others. For another recent study using similar techniques, see Salaita et al. (2010), who reported the correlations between the activation of a signaling pathway related to breast cancer and the mechanical forces that act on the receptors. In the future, these artificial systems will be invaluable for studying the mechanisms of protein immobilization and the effects of pickets on other molecules, which both require further clarification.

FUTURE PERSPECTIVES Membrane-skeleton-based compartmentalization has been observed in all of the cultured mammalian cells examined thus far and in many other cell types, suggesting that it is a common feature rather than a rare occurrence (Murase et al. 2004). We should stress that although this type of plasma membrane compartmentalization might be ubiquitous, it is probably just part of a hierarchy of domain structures that organize the membrane in a coordinated fashion. As we discussed in the previous section, these compartments may contain even smaller, and possibly highly dynamic, membrane domains with various diameters, such as lipid rafts, and these compartments also coexist with CCPs (Chapter 2), caveolae (Chapter 3), cell-adhesion structures (Chapter 17), and immunological/ neuronal synaptic junctions. Similarly, these compartments reside in larger domain structures, such as the apical/basolateral domains in epithelial cells (Chapter 21) and the somatodendritic/axonal domains in neurons (Chapter 22). The actin cytoskeleton (Chapter 12) and the membrane skeleton are likely to play important roles in organizing and orchestrating all of these structures. As we highlight in this chapter, a variety of cell functions are affected by the disruption of the membrane skeleton. Among the plethora of factors that determine membrane

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function, the reduction of membrane homogeneity may play a particularly special role. This point of view encourages us to think about the evolutionary reasons behind domain formation. For instance, if plasma membrane molecules were well mixed, then various cell functions, such as the regulation of the activity of receptors that is dependent on oligomerization and the localization of receptors, could become extremely inefficient. This suggests that many structures or mesoscale domains that increase membrane heterogeneity, including the membrane-skeleton-induced compartments, might have been selected and preserved during the long course of evolution. In a reductionist point of view, the actin fences and anchored-protein pickets provide a basis for molecular interactions in the plasma membrane. When many molecules are involved, these basic interactions can lead to rich collective dynamics. The presence of membrane-skeleton-based compartments may facilitate the persistence of molecular complexes, required for many cellular functions, that otherwise would be disrupted by thermal fluctuations. The membrane compartments and partitioning thus play crucially important roles in various plasma membrane functions, ranging from signaling to endocytosis, and future research will lead to a better understanding of their role in multiple cellular processes.

ABBREVIATIONS BCR Cav1 CCP DOPE EGF EGFR FCS FRAP FRSKs

B-cell receptor caveolin-1 clathrin-coated pit 1,2-dioleoyl-sn-glycero3-phosphoethanolamine epidermal growth factor epidermal growth factor receptor fluorescence correlation spectroscopy fluorescence recovery after photobleaching fetal rat skin keratinocytes (cell line)

GFP GPI NRK SCAR SFMT SPT STED-FCS WASP WAVE

green fluorescent protein glycosylphosphatidylinositol normal rat kidney (cell line) suppressor of cyclic AMP receptor single fluorescent-molecule tracking single-particle tracking stimulated emission depletion–FCS Wiskott–Aldrich syndrome protein WASP family verprolin-homologous protein

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CH A P T E R

2

CLATHRIN-COATED PITS James R. Thieman Linton M. Traub

DEFINITION Eukaryotic cells contain abundant surface molecules that must be maintained at appropriate concentrations for normal operation and cellular homeostasis. This can be accomplished by regulating the itinerary and delivery of outward-bound molecules (termed exocytosis) as well as by the inward-bound removal of macromolecules and transmembrane proteins from the surface (endocytosis) as necessary. Clathrin-coated pits are morphologically discernible invaginations of small regions (∼200 nm) of the plasma membrane that function as endocytic cellular domains. Composed of a clathrin protein scaffold, numerous adaptors, and phosphoinositide lipids, these pits serve a variety of functions in cell metabolism and organism homeostasis including neurotransmission, nutrient uptake, signaling termination, regulation of immune surveillance, cell motility, and pathogen entry. Clathrin-coated pits are a morphologically and temporally defined intermediate of this endocytic process, which proceeds from initial adaptor and clathrin placement at sites on the plasma membrane, through simultaneous cargo capture and membrane invagination, membrane scission, vesicle uncoating, and, finally, recycling of adaptors and clathrin for further rounds of endocytosis. The resulting uncoated vesicle can then fuse with early endosomes for further intracellular sorting of membrane components. While only occupying 1–2% of the cell surface (Anderson et al. 1978), it is estimated that in fibroblasts, clathrin-coated pits can turnover nearly the equivalent of the entire plasma membrane in 1 hour (Bretscher 1982). A heterogeneous population of transmembrane proteins, termed cargo, is recognized by clathrin-coated pits. Through capture of nutrient receptors, for example, the pits can efficiently concentrate ligands orders of magnitude over freely diffusing molecules in the extracellular milieu for packaging into vesicles. Indeed, while other endocytic pathways coexist at the plasma membrane, including pinocytosis, phagocytosis, caveolae-mediated endocytosis, and other less well-defined clathrin-independent endocytic routes, the hallmark of clathrin-mediated endocytosis remains selectivity. Central to this selectivity are phylogenetically conserved peptide-based endocytic sorting signals present within receptor cargo or appended protein folds that can be read by clathrin adaptors for packaging into these trafficking vesicles.

HISTORICAL PERSPECTIVE The first ultrastructural observations of clathrin-coated pits as discrete plasma membrane specializations were by Thomas Roth and Keith Porter in 1964 (Roth and Porter 1964). Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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They were interested in how circulating proteins within female mosquito hemolymph were deposited as yolk inside developing oocytes. They noticed that shortly following a blood meal, a proliferation of “bristle-coated pits and vesicles” on the cytoplasmic surface of the mosquito oocyte plasma membrane occurs. These pits and vesicles contained three distinguishable layers by electron microscopy: an inner, extracellular/luminal layer corresponding to massed yolk protein; a middle layer of ∼75-Å corresponding to the bilayer membrane; and an outer layer ∼200 Å thick that resembled a coat of fibrous bristles in the thin sections (Fig. 2.1A). Most remarkably, they correctly predicted from static images that these coated pit domains likely operated as a shuttle: They concentrated the yolk and then budded from the surface, uncoated and fused to form the large mature yolk bodies seen in mosquito oocytes––central tenets of modern coated vesicle biogenesis models. Subsequent studies revealed that these coats are arranged into cages composed of pentagonal and hexagonal facets closely apposed around the central vesicle. Similar appearing coated vesicles are also found in a variety of vertebrate tissues. In 1975, Barbara Pearse was the first to biochemically purify and characterize the coat complex as a single protein that she named “clathrin” (Pearse 1976). A ∼190-kDa molecule, the clathrin heavy chain, is organized into three-legged trimers named triskelions with three noncovalently associated clathrin light chains of ∼25 kDa (Ungewickell and Branton 1981). Somewhat surprising was that clathrin has no inherent affinity for membranes. Less abundant “associated proteins” or “assembly polypeptides” (APs) form a layer between assembled clathrin and the plasma membrane to assist in adhering clathrin to assembling pits and vesicles. Around this time it was appreciated that extracellular nutrients and hormones had specific receptors on the cell surface for their binding. Landmark work by Joseph Goldstein and Michael Brown with the low-density lipoprotein (LDL) receptor, a surface receptor for the plasma LDL particles that transport cholesterol and esterified lipids, showed that it could cluster in the plasma membrane above clathrin-coated pits and use these zones as portals of entry into the cell. Furthermore, fibroblasts from patients suffering from familial hypercholesterolemia, a hereditary condition in which patients have elevated plasma cholesterol contributing to cardiovascular disease, were shown to have defects in the clustering of LDL receptor into clathrin-coated pits as a result of a mutation in the receptor. On pinpointing disease-causing inherited mutations within the cytosolic domain of the LDL receptor, Goldstein and Brown proposed that this receptor could be concentrated and internalized by clathrin-coated pits through interactions between its cytosolic portion and the coat components (Goldstein et al. 1979). These observations laid the groundwork for the characterization of a whole collection of endocytic sorting signals that are the basis for the selectivity of clathrin-coated pits for its cargo. When two or more different receptors were found to be present within individual coated pits, it became apparent that these structures were common entry points for diverse cell-surface proteins (Maxfield et al. 1978). Throughout the early 1980s, researchers began to characterize the assembly polypeptides. Originally described as potential contaminating proteins in coated vesicle preparations, these APs of approximately 100 and 50 kDa copurify with coated vesicles and are found sandwiched between the vesicle and the clathrin coat (Vigers et al. 1986). APs promote the polymerization of clathrin into more uniform-sized cages in vitro than clathrin alone (Zaremba and Keen 1983). Later, it was determined that these polypeptides correspond to the tetrameric adaptor complexes AP-1 and AP-2, involved in clathrin-coated vesicle formation at the trans-Golgi network (TGN) and endosomes or at the plasma membrane, respectively (see also Chapters 8 and 9 on the Golgi and endosomes, respectively). From brain-derived coated vesicles, a separate fraction of ∼100-kDa proteins would turn out to be other monomeric endocytic adaptors that bind to clathrin, and the

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Figure 2.1. Microscopic observation of clathrin-coated pits. (A) Thin-section electron micrograph of synaptosome membranes generated from synaptic terminals. The plasma membrane is visible as two dense parallel lines contiguous with the inner layer of the deeply invaginated clathrin-coated pits. The bristle-like clathrin coat is visible around the forming vesicles. (B) Freeze-etch electron micrograph of clathrin-coated pits on the adherent plasma membrane of cultured cells. The clathrin coat has been pseudocolored blue, and the position of an individual clathrin triskelion has been colored red. Notice the array of pentagons and hexagons of the clathrin lattice in both flat and invaginating pits. (C, D) HeLa cervical carcinoma cells in culture visualized by transmitted light (C) or fluorescence microscopy for exogenously expressed clathrin light chain (D). (E, F) BS-C-1 kidney cells under the same conditions as in C and D. Notice the uniform diffraction-limited clathrin structures of BS-C-1 cells in F compared with the diffractionlimited and large clathrin structures of HeLa cells in D. Scale bars represent (A) 50 nm, (B) 100 nm, and (C–F) 10 μm.

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APs and can also polymerize clathrin in vitro (Ahle and Ungewickell 1986). Then in the early 1990s, two structurally unrelated classes of tyrosine-based sorting signals and one leucine-based sorting signal were discovered within the cytosolic domains of various cellsurface receptors. Because they are autonomous and transplantable, these conserved amino acid stretches specify a receptor ’s fate for endocytosis. The 50-kDa subunit of AP-2 was shown to recognize one class of tyrosine-based sorting signal, describing a molecular mechanism for adaptor sorting during clathrin-mediated endocytosis. It was clearly evident at this time that clathrin-coated vesicles were well-ordered protein assemblies––a defined cellular domain. The cloning of cDNAs for clathrin and adaptors, starting in 1987 (Kirchhausen et al. 1987; Robinson 1989), opened up the field for mutagenesis studies and the creation of fluorescent protein fusions. This permitted the identification of additional regions of the adaptors that recognize receptor cargo and clathrin as well as the ability to perform studies of clathrin and adaptor dynamics in living cells. Concurrently, structural studies by X-ray crystallography, nuclear magnetic resonance, and cryo-electron microscopy were revealing the architecture of clathrin and adaptors and providing an exciting picture of how these proteins engage one another (Owen 2004). The past decade has seen a rapid expansion of research using sophisticated, timeresolved imaging techniques to study endocytosis. In the process, many of the original tenets of endocytosis proposed earlier were confirmed and extended (Roth 2006). Recent results suggest that the adaptors and clathrin are recruited to regions of the plasma membrane undergoing endocytosis in a regulated manner both in space and time.

MOLECULAR COMPONENTS OF CLATHRIN-COATED PITS AND VESICLES Clathrin-coated structures are discernable as flat to deeply invaginated regions of the plasma membrane that range from ∼50 to >500 nm (Heuser 1980). By thin-section electron microscopy, they appear to have a cytosol-oriented, electron-dense, fuzzy coat in longitudinal sections (Fig. 2.1A), while by freeze-etch electron microscopy, the coat displays a honeycomb-like polyhedral appearance (Fig. 2.1B) (Heuser 1980). Although clathrincoated pits only occupy a small percentage of the surface area at steady state (Fig. 2.1C–F), this dedicated domain is also visible from the extracellular surface of the plasma membrane. In freeze-etch electron micrographs, spherical ∼25-nm LDL particles are seen clustered at dimpled specializations (Heuser and Anderson 1989), and immunofluorescence analysis shows transferrin, LDL, and epidermal growth factor (EGF), as well as other extracellular ligands, clustered in discrete puncta, which are coated on the intracellular surface with clathrin. The clear concentration of cargo away from the freely diffusing molecules in the plasma membrane and the defined appearance of the assembled clathrin lattice make this sorting device a discrete membrane domain physically and functionally distinguishable from the surrounding membrane. Because the clathrin-coated membrane domain typically progresses to clathrincoated vesicles that physically detach from the plasma membrane, the protein composition of biochemically isolated coated vesicles was the starting point for molecular characterization of the chief coated pit constituents. But a diverse array of proteins are now known to accumulate at clathrin-coated pits, either transiently or throughout the entire life of these structures, as determined by methods such as biochemical purification, proteomics, light microscopy, and electron microscopy. The arrangement and complexity of the clathrincoated pit domain reflects its role as the first sorting station that surface receptors encounter on their way through the retrograde endomembrane system.

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Clathrin Clathrin, the principal protein of coated pits and vesicles, is a 1675 residue, ∼190-kDa heavy chain with one associated 25- to 29-kDa light chain. Three heavy chains combine to form a trimer known as a clathrin triskelion (Fig. 2.2A). Triskelions have a threefold rotational symmetry about their trimerization domain. The leg is made up of an extended α-helical solenoid fold with kinks at regular intervals, giving the legs an overall arched appearance. Either under acidic pH or at physiological conditions with the assistance of adaptor proteins, the triskelions engage one another in an antiparallel manner to form a spherical, polygonal array called a clathrin coat. At the free end of each clathrin triskelion leg, and projecting down into the inner layer of the clathrin-coated pit or vesicle, is the so-called clathrin terminal domain. This corresponds to the N-terminal section and performs the vital function of attaching clathrin to the membrane. Unlike the adaptor proteins that have dedicated domains for engaging lipids or receptor cargo at the plasma membrane, clathrin does not bind membranes directly. Instead, the terminal domains, composed of a seven-bladed β-propeller fold, provide binding pockets for unstructured and conserved motifs within the adaptors. The ability of the clathrin terminal domain to recognize multiple adaptor-type proteins, the presence of three terminal domains in each triskelion, and the tight packing of triskelions together (Fig. 2.1B) create a very stable scaffold that assists in bending and stabilizing as well as organizing the plasma membrane beneath the coat (Edeling et al. 2006).

The Adaptor Complexes AP-2 The APs were discovered as proteins copurifying with clathrin-coated vesicles and apparently positioned between the vesicle and the outer clathrin coat. Their established role in sorting cargo in addition to coat assembly has led them to more commonly be dubbed “adaptor protein” complexes. Four structurally analogous AP complexes are ubiquitously expressed: AP-1, AP-2, AP-3, and AP-4. Clathrin-coated vesicle formation is a general cellular phenomenon for producing transport vesicles and occurs not only during endocytosis at the plasma membrane but also at the TGN and on endosomes along the biosynthetic and degradative pathways. While AP-1, AP-3, and AP-4 can be found at the TGN and endosomes, AP-2 is exclusively found on the plasma membrane and chiefly coordinates clathrin-mediated endocytosis (Nakatsu and Ohno 2003). While clathrin-coated structures on the TGN and endosomes may morphologically resemble clathrin-coated pit domains at the surface, they are biochemically distinct, are regulated differently, and, in some cases, sort distinct cargo. AP-2 (like the other APs) is a heterotetramer composed of two large subunits, α and β2; one medium, μ2; and one small subunit, σ2 (Fig. 2.2B). The subunits combine to form a brick-like core composed largely of solenoid-folded α-helices (the α- and β2-subunit trunks), while the μ2 is mostly β-sheet with its N-terminus contributing to core structural stability and its C-terminus tucked against the core. The σ2-subunit is structurally analogous to the N-terminus portion of μ2 (Collins et al. 2002). The large subunits also contain unstructured hinge domains near their C-termini that extend from the core and end in independently folded appendage domains. This overall arrangement has implications for clathrin-coated pit and vesicle formation. The AP-2 core orients upon the plasma membrane by recognizing the negatively charged phosphate groups on the lipid phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) through the α- and μ2-subunits while positioning the appendages toward the cytosol. Attached to intrinsically unfolded linkers,

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SIDE

TOP

heavy chain

GPCR

β -arrestin EGF receptor

epsin

light chain

CD4

AP-2

90o

terminal domain

AP-2 transferrin receptor

LDL receptor

Dab2

clathrin binding

(B)

β2

PtdIns(4,5)P2

μ2

ARH

σ2

LDL receptor

hinge

α

core

clathrin coat

cargo

membrane

adaptors

appendages

Figure 2.2. Clathrin-coated pit assembly. (A) Schematic of a clathrin triskelion. Three clathrin heavy chains trimerize at the vertex of the triskelion, while three light chains bind noncovalently to the triskelion legs. The legs are curved in appearance. (B) Adaptor protein 2 (AP-2) is a heterotetramer. Large subunits α and β2 are in blue and green, respectively. Medium subunit μ2 is in magenta. The μ2 N-terminus is circular and C-terminus is elliptical. The small subunit σ2 is in orange. The large subunit C-termini appendages are shown connected to the core by flexible hinge domains. Regions dedicated to clathrin binding are indicated by yellow circles. (C) Initial placement of adaptors and clathrin on the plasma membrane prior to membrane invagination. AP-2 molecules are transiently recruited by binding PtdIns(4,5)P2 through the α-subunit (far left) and are stabilized by μ2 rearrangement (far right). This promotes receptor binding (center). Clustering of AP-2 and adaptors promotes clathrin binding through AP-2 appendages and clathrin box motifs in the AP-2 hinge and monomeric adaptors. This flat clathrin assembly will proceed to a spherical coat such as those seen in Figure 2.1A, B.

(C)

(A)

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the motional freedom of the appendages allows sampling of the nearby cytosol, increasing the frequency of appendage collision with, and binding to, one of their many interaction partners (Fig. 2.2C). Importantly, the β2-subunit appendage and a defined region of the adjacent unstructured hinge can bind bivalently to clathrin, at the leg and terminal domain, respectively (Fig. 2.2B). This multivalent interaction allows AP-2, in part, to act as a glue holding clathrin to the plasma membrane (Edeling et al. 2006). AP-2 acts not only to hold clathrin to the plasma membrane but also to recognize and retain select cargo within forming clathrin-coated vesicles. Two classes of sorting signals within the cytoplasmic portion of transmembrane proteins are recognized by AP-2: (1) YXXØ (where Y is tyrosine, X is any amino acid, and Ø represents a bulky hydrophobic residue) and (2) [DE]XXXL[LIM] (where the first position can be aspartic acid or glutamic acid and the last position can be leucine, isoleucine, or methionine). These short linear signals are recognized by the μ2 C-terminus for YXXØ or the α/σ2-subunit hemicomplex for dileucine signals (Ohno et al. 1995; Kelly et al. 2008). The YXXØ signal is found in numerous receptors, including the prototypical clathrin-coated vesicle cargo, the transferrin receptor, while the dileucine signal is often found in immunological receptors such as CD4. Functionally, this means that AP-2 can simultaneously sort at least two different kinds of receptors. This reduces competition at the plasma membrane for receptor entry (Traub 2009b). For example, a clathrin-coated pit could perform the task of iron uptake through transferrin receptor capture by AP-2 but not compromise its ability to also regulate surface expression of the CD4 receptor. Monomeric Adaptors While the tetrameric AP-2 complex is the quintessential adaptor coordinating interactions between clathrin, the plasma membrane, cargo, and numerous other endocytic proteins, many other monomeric adaptors are present within the cell. The general architecture of these proteins typically reveals a folded, modular domain at one terminus flanked by an unstructured segment typically containing short peptide-based interaction motifs, often tandemly arranged. Some of these motifs facilitate binding to the clathrin terminal domain such as the “clathrin box” (consensus sequence LØ[DEN]Ø[DEN]), while other motifs specifically engage the AP-2 appendages. The function of these monomeric adaptors is one or both of the following: (1) to bind clathrin and assist in tethering it to the membrane, and (2) to act as alternate adaptors by recognizing additional sorting signals within the cargo not decoded directly by AP-2 and connecting it to the nascent endocytic machinery. Because these coat components play an important role in cargo selection, they have been designated clathrin-associated sorting proteins (CLASPs) (Traub 2009b). β-Arrestins β-Arrestins 1 and 2 were the first proteins recognized to act as alternate adaptors by increasing the sorting potential of clathrin-coated pits beyond cargo recognized by AP-2 (Goodman et al. 1996). Following stimulation of G-protein-coupled receptors (GPCRs) with an appropriate ligand and subsequent phosphorylation of the GPCR by kinases, β-arrestins recognize the phosphorylated serine/threonine residues within the cytosolic loops of these seven-pass transmembrane receptors. β-Arrestin is usually soluble in the basal state as the largely unstructured C-terminus is bound back on the major β-arrestin fold as an inhibitory clamp. Upon receptor engagement, electrostatic associations with the receptor-attached phosphates discharge the C-terminus, revealing AP-2 appendage and clathrin box binding motifs. This interaction rapidly desensitizes signal propagation by uncoupling G proteins from the receptor. In addition, the revealed clathrin coat interaction motifs target β-arrestin and the associated desensitized receptor to preexisting clathrin-coated pits for endocytosis (Fig. 2.2C) and downregulation where

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they will either recycle or be degraded (Luttrell and Lefkowitz 2002). Hence, clathrinmediated endocytosis is important in the establishment of a refractory period in the cell in many GPCR signaling pathways. Autosomal Recessive Hypercholesterolemia (ARH) and Disabled-2 (Dab2) The ARH protein and Dab2 CLASPs cooperate in recognizing cargo containing FXNPXY sorting signals such as those commonly found in lipoprotein receptors and integrins. Mutations in this sorting signal discovered by Goldstein and Brown prevent the LDL receptor from clustering in pits (Anderson et al. 1977). ARH and Dab2 contain N-terminal phosphotyrosine-binding domains that, despite the name, bind selectively to unphosphorylated FXNPXY signals and PtdIns(4,5)P2 simultaneously. Again through unstructured AP-2 appendage and clathrin-binding motifs in ARH and Dab2, these adaptors target cargo such as the LDL receptor to clathrin-coated pits for internalization (Traub 2009b). In the case of ARH, the unstructured AP-2 motif becomes structured as it engages the AP-2 β2-appendage. Interestingly, mutations in ARH that prevent its recognition of the LDL receptor result in failure to clear LDL receptor from the surface and hypercholesterolemia with a different inheritance pattern than that studied by Goldstein and Brown (Garcia et al. 2001). ARH and/or Dab2 may also be important for integrin internalization during cell migration for redistribution to the leading edge (Teckchandani et al. 2009). AP180 AP180 was the second AP to be discovered following the heterotetrameric AP-1 and AP-2 complexes. This adaptor binds to PtdIns(4,5)P2 via the AP180 N-terminal homology (ANTH) domain. The unstructured C-terminus harbors several motifs for AP-2 appendage and clathrin binding as well, so AP180 contributes to the structural integrity of clathrin-coated pits. AP180 expression is restricted to neurons, but a ubiquitously expressed isoform CALM performs similar functions in other tissues. There is some evidence that AP180/CALM might be involved in targeting soluble N-ethylmaleimidesensititive fusion protein (NSF) attachment receptors (SNAREs), which promote the ethylmaleimide-sensititive fusion protein (NSF) attachment receptors (SNAREs), which promote the subsequent membrane fusion of uncoated transport vesicles with the appropriate acceptor compartment (Harel et al. 2008). Epsin and Eps15 Epsin and Eps15 are monomeric adaptors found at clathrincoated pits at the cell surface. Through either an epsin N-terminal homology (ENTH) domain structurally related to the ANTH domain or an Eps15 homology (EH) domain, these two proteins are able to bind PtdIns(4,5)P2, while their unstructured C-termini can engage the AP-2 appendages. Epsin also binds to clathrin and therefore stabilizes the assembly, but Eps15 does not. Both proteins contain ubiquitin-interacting motifs (UIMs) that allow them to bind receptors that have been ubiquitinated. In this respect, they are able to target cargo that becomes ubiquitinated following activation, such as the EGF receptor, to clathrin-coated pits for internalization (Traub 2009b). This increases the diversity of cargo that can be endocytosed in a clathrin-dependent fashion beyond that recognized by the AP-2 adaptor.

Regulatory Proteins and Lipids PtdIns(4,5)P2 Lipid Clathrin-coated pits are not only protein assemblies but also “lipid assemblies” as they are constructed on the plasma membrane and depend on the lipid PtdIns(4,5)P2. Phosphatidylinositol is a phospholipid with two acyl chains, a glycerol backbone, and a phosphate group connecting a six-carbon inositol ring. Variable phosphorylation of three of the other five carbons generates a total of eight phosphoinositide species, a subset

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of which has defined distributions throughout the endosomal system (Roth 2004). Phosphorylation of carbons 4 and 5 hydroxyl group yields PtdIns(4,5)P2, which is found highly concentrated on the inner leaflet of the plasma membrane. PtdIns(4,5)P2 is a vital component for assembly and progression of clathrin-coated pits and AP-2, and many monomeric adaptors bind this lipid (Haucke 2005). This bioactive lipid may be transiently enriched at sites of coat assembly; however, this may not be obvious due to the rudimentary methods currently used to visualize this lipid in cells (i.e., fluorescently labeled lipidtargeting protein domains that uniformly label the plasma membrane). Strong evidence for an actuating role of PtdIns(4,5)P2 in clathrin-coated pit assembly comes from the synaptic phenotypes in PtdIns(4,5)P2-generating PIPKIγ (Di Paolo et al. 2004) and PtdIns(4,5)P2-hydrolyzing synaptojanin (Cremona et al. 1999) gene-specific knockout animal models, and cellular perturbation of PtdIns(4,5)P2 metabolism in cultured cells (Zoncu et al. 2007). Dynamin Successful production of a clathrin-coated vesicle requires the scission of the coated pit from the plasma membrane. The GTPase dynamin can oligomerize into a ring-like collar at the base of deeply invaginated clathrin-coated pits such as those in Figure 2.1A. At this late stage of clathrin-coated pit assembly, in which only a tubular bilayer stalk connects the bud to the plasma membrane, very little further incorporation or escape of sorted cargo is expected to occur due to the reduced surface area for cargo transfer. Through conformational changes induced by GTP binding/hydrolysis and likely coincident tensile force from actin-based propulsion of the vesicle, dynamin promotes cleavage of the membrane. There is also evidence for dynamin regulating assembly of clathrin-coated pits (Mettlen et al. 2009). The precise mode of lipid destabilization and reorganization that facilitates bilayer fusion is not yet clear, but dynamin may exit the bud before final fission has occurred. Synaptojanin, Auxilin/G-Cyclin-Associated Kinase (GAK), and Hsc70 Through numerous protein–protein interactions, many of high avidity, a relatively stable assembly surrounding the bud and nascent vesicle is constructed. In order to recycle clathrin triskelions and adaptors for future rounds of clathrin-coated pit assembly and for the vesicle to sterically recognize and fuse with a target membrane, the overlying cytosolic components must be stripped from the vesicle. To accomplish this, the molecular chaperone ATPase Hsc70 is recruited to late-stage clathrin-coated pits or fully formed vesicles by the co-chaperone auxilin (expressed in neurons) or GAK (expressed ubiquitously). Through ATP hydrolysis, Hsc70 promotes the dissociation of clathrin triskelions, destabilizing the underlying adaptors and promoting uncoating (Eisenberg and Greene 2007). Hydrolysis of PtdIns(4,5)P2 by synaptojanin also plays a role in clathrin/adaptor uncoating, and AP-2 removal from vesicles requires dephosphorylation of the μ2-subunit (see below), which may require competitive removal of the μ2 targeting adaptor-associated kinase (AAK1) from AP-2 (Semerdjieva et al. 2008).

Structural Proteins Bin/Amphiphysin/Rvs161/167 (BAR)-Domain-Containing Proteins Proteins containing BAR domains are involved in membrane curvature in the endocytic/ endosomal membrane system. BAR domains are crescent-shaped protein dimers lined with positively charged residues along their concave surfaces that recognize negatively charged phospholipids. Their ability to tubulate membrane in vitro and their structure suggest they may generate or stabilize bending of target membranes (Brett and Traub 2006). Amphiphysin, endophilin, and sorting nexin 9 are BAR domain proteins that target to

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clathrin-coated pits by binding to the AP-2 appendages. These proteins are thought to assist in later stages of invagination at the neck of clathrin-coated pits where membrane curvature is highest. Many BAR proteins have Src homology-3 (SH3) domains that can directly engage the large GTPase dynamin and might recruit this late regulator to the neck of coated-pits. Filamentous Actin and Its Adaptor One striking feature of clathrin-coated pits by deep-etch cryo-electron microscopy is their frequent positioning adjacent to the cortical actin cytoskeleton. In fact, while controversy remains over the absolute requirement for actin polymerization in scission and propulsion of nascent clathrin-coated vesicles away from the plasma membrane, it is likely that filamentous actin at a minimum plays a role in the regulation of vesicle formation. Clathrin adaptors HIP1/Hip1R bind to the plasma membrane via an ANTH domain, form homoand heterodimers and bind clathrin light chain through a central coiled-coil domain, and bind to filamentous actin via an ILWEQ/THATCH domain. These proteins, conserved from yeast to human, support a role for actin attachment to clathrin-coated pits during clathrincoated pit progression (Brett and Traub 2006).

FUNCTIONAL IMPLICATIONS OF CLATHRIN-COATED PIT ORGANIZATION Constructing Clathrin-Coated Pits For years, it was presumed that either receptors destined for endocytic uptake or an unidentified, compartment-specific nucleating membrane protein were responsible for recruiting the AP-2 adaptor to regions of the plasma membrane to begin assembly of clathrin-coated pits. However, AP-2 is not recruited to endosomal membranes containing highly concentrated receptors, so it is clear that regulation of AP-2 placement involves additional factors. Instead, it is now known that AP-2 and monomeric adaptors initially bind to PtdIns(4,5) P2 through their dedicated lipid-binding modular domains. In nerve terminals, PtdIns(4,5) P2 for clathrin-mediated endocytosis is generated from phosphatidylinositol 4-phosphate (PtdIns(4)P) by type Iγ phosphatidylinositol 4-phosphate (PIP) kinase (Wenk et al. 2001). For AP-2, a patch of basic residues in the α-subunit recognize the negatively charged phosphate groups of PtdIns(4,5)P2. However, this interaction is weak, and further contacts are needed to stabilize AP-2 at the membrane. Phosphorylation of the μ2-subunit of AP-2 by AAK1 causes AP-2 to undergo a conformational change. The μ2-subunit is displaced, positioning itself parallel with the plasma membrane. Thereby, a second PtdIns(4,5)P2 recognizing patch of residues on μ2 is realigned to bind this lipid, increasing the affinity of AP-2 for the plasma membrane by roughly 100 times (Maldonado-Baez and Wendland 2006). A second consequence of this open AP-2 conformation is its receptiveness to binding YXXØ sorting signal-containing cargo binding by μ2-subunit. This could further stabilize AP-2 on the membrane. Following the initial binding of a few molecules of AP-2 to the plasma membrane, additional AP-2 and monomeric adaptors are concentrated at these sites through a mechanism termed coincidence detection (Carlton and Cullen 2005). In this scenario, the placement of a protein within the cell is determined by the close proximity of not one but two or more binding partners for simultaneous engagement. As AP-2-recognized cargo becomes concentrated in regions of the membrane along with PtdIns(4,5)P2, the likelihood of AP-2 becoming deposited is increased. Similarly for monomeric adaptors, locally increased

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concentrations of PtdIns(4,5)P2 can potentially be recognized by ENTH, ANTH, and EH domains, while the AP-2 appendage domains can be recognized by motifs presented by the CLASPs. This helps ensure that AP-2 and CLASPs/adaptors are recruited selectively to the plasma membrane relative to other organellar membranes within the cell that may contain recycling or degrading cargo or PtdIns(4,5)P2. While clathrin does not recognize membrane lipids directly, it binds AP-2 and CLASPs with high avidity. Each of the three clathrin terminal domains can engage at least one AP-2 or CLASP through short linear clathrin box motifs (Edeling et al. 2006). Thus, once an appreciable number of adaptors become concentrated on the plasma membrane, they could in theory begin to recruit clathrin triskelions. Adjacent positioning of clathrin triskelions on a planar surface would promote their self-association into a flat, hexagonal lattice (Fig. 2.1B). The appearance of additional clathrin molecules would provide positive feedback for AP-2 and monomeric adaptor recruitment and help drive assembly.

Generating Membrane Curvature Clathrin-coated pit domains must transition from an initial two-dimensional membrane assembly through high levels of curvature to a spherical end product in the course of vesicle generation. It is known that clathrin is required for inducing curvature, stabilizing membrane bending in the nascent bud, or both, as cells depleted of clathrin form discernable adaptor protein assemblies on the membrane that are entirely flat (Ungewickell and Hinrichsen 2007). The transition from a flat clathrin lattice formed mostly of hexagons to an invaginated coat (Fig. 2.1B) also requires the introduction of new pentagonal facets, but whether this provides a driving force for membrane curvature is unknown. Another possibility is that the adaptor epsin assists in curvature generation by the insertion of an amphipathic helix into the plasma membrane inner leaflet, changing relative bilayer spacing. The local expansion of the inner leaflet to the outer leaflet could promote bending toward the cytosol. Finally, BAR domain proteins with different intrinsic radii of curvature may assist at different stages of membrane bending either by generating curvature or stabilizing the existing membrane surface conformation (Ungewickell and Hinrichsen 2007).

Incorporating Cargo Beginning as early as the deposition of a cluster of adaptors on the plasma membrane and possibly occurring up until the neck of the late-stage pit becomes occluded or sealed off by dynamin, clathrin-coated pits perform their principal function of cargo capture. As the plasma membrane is roughly 50% protein by weight (Cooper 2000) and many of these proteins can diffuse within the membrane, clathrin-coated pits are confronted with the challenge of recognizing specific receptor cargo for entry into the cell within a vast sea of heterogeneous protein. It can be envisioned that clathrin-coated pits act as filters on the plasma membrane, capturing cargo that has specific sorting signals within its exposed cytosolic domain as it diffuses through and allowing nonspecific proteins to diffuse out. Endocytic adaptors are arranged in a way such that cargo recognition takes place near the plasma membrane and deep within the clathrin coat. Monomeric adaptors typically contain their modular PtdIns(4,5)P2-binding domain at their N-termini. For βarrestins, Dab2, and ARH, cargo recognition also occurs through this domain. For epsin, a region just to the C-terminus of its lipid-binding ENTH domain is responsible for recognizing ubiquitinated cargo. Similarly for AP-2, the membrane adjacent μ2-subunit can bind to YXXØ sorting signals. A binding pocket for [DE]XXXL[LIM] dileucine sorting

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signals is created by a space between the AP-2 α- and σ2-subunits. However, this site appears to be blocked by the large β2-subunit and might need a conformational change in order to recognize dileucine sorting signals. Altogether, these adaptors recognize sorting signals a short distance from the membrane (Traub 2009b). In contrast, interactions that are structural in nature and act to connect monomeric adaptors to AP-2 and adaptors to the clathrin scaffold occur at a greater distance from the membrane. These interactions converge around the AP-2 appendage domains. The appendage domains have the potential to bind numerous partners. They recognize short, linear, and, in some cases, α-helical motifs within monomeric adaptors and accessory endocytic proteins (Traub 2009b). Often, these motifs are at the C-termini of these proteins distal to membrane association domains. The unstructured segments of monomeric adaptors also often contain short, linear clathrin boxes (Maldonado-Baez and Wendland 2006). Together, these monomeric adaptors and the β2-subunit of AP-2 bind and recruit soluble clathrin triskelions to the plasma membrane for lattice assembly and invagination.

Clathrin-Coated Plaques and De Novo Forming Clathrin-Coated Pits Thus far, clathrin-coated pit assembly has been described as a process that gives rise to one clathrin-coated vesicle and in doing so removes this temporary domain from the plasma membrane. For some cells in culture, these de novo forming structures are the primary clathrin-coated pits decorating the plasma membrane in the cell and are diffraction limited (<200 nm) by light microscopy (Fig. 2.1E, F). However, in other cultured and primary cells, large (>500 nm), static and mostly flat clathrin-coated structures can also be observed on the adherent plasma membrane (Fig. 2.1C, D) (Traub 2009a). These so-called clathrin-coated plaques sequester cargo, yet the regulation and functional consequences of plaque formation versus de novo pit formation are unknown. Static freeze-etch electron microscope images suggest that clathrin-coated vesicles can bud from the periphery of these structures (Fig. 2.1C), but it has also been suggested that the entire “plaque” may internalize (Saffarian et al. 2009). How a “plaque” is kept in a flat conformation and what triggers clathrin rearrangements will be topics for future research.

Cargo-Dedicated Clathrin-Coated Pits From what we now know of how AP-2 and CLASPs recognize their cargo for incorporation into clathrin-coated pits, and the diversity of receptors containing various sorting signals encoded in their cytosolic domains, it would be expected that an individual clathrin-coated pit would contain numerous different cargo molecules. Interestingly, there is evidence for some GPCR cargos utilizing only a subset of clathrin-coated pits. In one study, exogenously expressed and ligand-bound delta opioid receptor occupied only ∼40% of clathrin-coated pits (Puthenveedu and von Zastrow 2006), while in another study, exogenously expressed purinergic receptors P2Y1 and P2Y12 localized to distinct coated pits upon ligand stimulation (Mundell et al. 2006). Typically, CLASPs have a wide distribution and colocalize with nearly all clathrin-coated pits (Keyel et al. 2006) including a mutant form of β-arrestin capable of localizing to clathrin-coated pits in the absence of activated GPCRs (Burtey et al. 2007). Thus, how certain cargo manages to be selectively incorporated into a subset of pits, if these pits are morphologically and biochemically different, and whether they are targeted to distinct endosomal compartments remains to be determined.

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ABBREVIATIONS

35

Future Perspectives Great strides have been made in the past two decades in uncovering novel endocytic proteins that contribute to clathrin-coated pit domain construction and deciphering how these proteins interact with one another. In addition, various structural studies have uncovered how endocytic protein architecture dictates unique modes of regulation of this process. While we have some understanding of the individual pieces of the puzzle, exactly how and when these proteins assemble remain a major focus in the field. It may be possible to imagine clathrin-coated pit assembly is analogous to an energy landscape model of protein folding (Onuchic and Wolynes 2004). In these models, an unfolded protein can assume many different conformations of high entropy, but as it folds, it moves down the sides of its folding funnel to its native conformation of lowest free energy and lowest entropy. There are many different pathways (or sides of the funnel) to reach the same final protein fold. Clathrin-coated pits and vesicles may form in a similar fashion in which different components could have the ability to contribute at different times. Different cell types may also use slightly different endocytic components and adaptors, yet as adaptors and clathrin triskelion begin to assemble, they limit the possible subsequent conformations of the forming coated pit and drive the process forward vectorially. Implied by this model is that a series of ordered protein–lipid and protein–protein interactions could guide the formation of a clathrin-coated pit. It has been proposed that interactions of lower affinity are progressively replaced by interactions of higher affinity thereby giving directionality to the assembly process (Mishra et al. 2004; Praefcke et al. 2004). As other fields have moved toward a systems biology holistic approach to understanding complex biological reactions, future studies into clathrin-coated pits and vesicles may also follow this trend. In order to fully understand how clathrin-coated pits assemble a comprehensive characterization of endocytic protein motifs, sorting signals, interaction dissociation constants, and endocytic protein cellular concentrations would provide valuable data that could be used to build computation-based assembly models––an approach that can now be fully realized thanks to greatly improved computing capabilities. This information could be used to validate current hypotheses on clathrin-coated pit assembly. Other exciting avenues include determining how clathrin-coated plaques form, if they are capable of budding from the plasma membrane en masse, and if they perform specialized sorting functions in comparison with de novo forming clathrin-coated pits. Also, whether cargo-specific clathrin-coated pits exist, how they would exclude specific cargo, and what benefit this early sorting has for the downstream endosomal system are now being explored.

ABBREVIATIONS APs ANTH ARH BAR CLASP Dab2 EGF EH

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associated proteins or assembly polypeptides AP180 N-terminal homology autosomal recessive hypercholesterolemia bin/amphiphysin/Rvs161/167 clathrin-associated sorting protein disabled-2 epidermal growth factor Eps15 homology

ENTH GPCR LDL NSF PtdIns(4)P PtdIns(4,5)P2 SH3 SNARE TGN

epsin/Eps15 N-terminal homology G-protein-coupled receptor low-density lipoprotein N-ethylmaleimide sensitive fusion protein phosphatidylinositol 4-phosphate phosphatidylinositol 4,5-bisphosphate Src homology-3 soluble NSF attachment receptor trans-Golgi network

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REFERENCES Ahle S, Ungewickell E. 1986. Purification and properties of a new clathrin assembly protein. EMBO J 5:3143–9. Anderson RG, Goldstein JL, Brown MS. 1977. A mutation that impairs the ability of lipoprotein receptors to localise in coated pits on the cell surface of human fibroblasts. Nature 270:695–9. Anderson RG, Vasile E, Mello RJ, Brown MS, Goldstein JL. 1978. Immunocytochemical visualization of coated pits and vesicles in human fibroblasts: relation to low density lipoprotein receptor distribution. Cell 15: 919–33. Bretscher MS. 1982. Surface uptake by fibroblasts and its consequences. Cold Spring Harb Symp Quant Biol 46(Pt 2):707–12. Brett TJ, Traub LM. 2006. Molecular structures of coat and coat-associated proteins: function follows form. Curr Opin Cell Biol 18:395–406. Burtey A, Schmid EM, Ford MG, Rappoport JZ, Scott MG, Marullo S, Simon SM, McMahon HT, Benmerah A. 2007. The conserved isoleucine-valine-phenylalanine motif couples activation state and endocytic functions of beta-arrestins. Traffic 8:914–31. Carlton JG, Cullen PJ. 2005. Coincidence detection in phosphoinositide signaling. Trends Cell Biol 15:540–7. Collins BM, McCoy AJ, Kent HM, Evans PR, Owen DJ. 2002. Molecular architecture and functional model of the endocytic AP2 complex. Cell 109:523–35. Cooper GM. 2000. The cell: a molecular approach. Washington, DC; Sunderland, MA: ASM Press; Sinauer Associates. Cremona O, Di Paolo G, Wenk MR, Luthi A, Kim WT, Takei K, Daniell L, Nemoto Y, Shears SB, Flavell RA, McCormick DA, De Camilli P. 1999. Essential role of phosphoinositide metabolism in synaptic vesicle recycling. Cell 99:179–88. Di Paolo G, Moskowitz HS, Gipson K, Wenk MR, Voronov S, Obayashi M, Flavell R, Fitzsimonds RM, Ryan TA, De Camilli P. 2004. Impaired PtdIns(4,5)P2 synthesis in nerve terminals produces defects in synaptic vesicle trafficking. Nature 431:415–22. Edeling MA, Smith C, Owen D. 2006. Life of a clathrin coat: insights from clathrin and AP structures. Nat Rev 7:32–44. Eisenberg E, Greene LE. 2007. Multiple roles of auxilin and hsc70 in clathrin-mediated endocytosis. Traffic 8: 640–6. Garcia CK, Wilund K, Arca M, Zuliani G, Fellin R, Maioli M, Calandra S, Bertolini S, Cossu F, Grishin N, Barnes R, Cohen JC, Hobbs HH. 2001. Autosomal recessive hypercholesterolemia caused by mutations in a putative LDL receptor adaptor protein. Science 292: 1394–8. Goldstein JL, Anderson RG, Brown MS. 1979. Coated pits, coated vesicles, and receptor-mediated endocytosis. Nature 279:679–85. Goodman OB, Jr., Krupnick JG, Santini F, Gurevich VV, Penn RB, Gagnon AW, Keen JH, Benovic JL.

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1996. Beta-arrestin acts as a clathrin adaptor in endocytosis of the beta2-adrenergic receptor. Nature 383: 447–50. Harel A, Wu F, Mattson MP, Morris CM, Yao PJ. 2008. Evidence for CALM in directing VAMP2 trafficking. Traffic 9:417–29. Haucke V. 2005. Phosphoinositide regulation of clathrinmediated endocytosis. Biochem Soc Trans 33:1285–9. Heuser J. 1980. Three-dimensional visualization of coated vesicle formation in fibroblasts. J Cell Biol 84:560–83. Heuser JE, Anderson RG. 1989. Hypertonic media inhibit receptor-mediated endocytosis by blocking clathrincoated pit formation. J Cell Biol 108:389–400. Kelly BT, McCoy AJ, Spate K, Miller SE, Evans PR, Honing S, Owen DJ. 2008. A structural explanation for the binding of endocytic dileucine motifs by the AP2 complex. Nature 456:976–79. Keyel PA, Mishra SK, Roth R, Heuser JE, Watkins SC, Traub LM. 2006. A single common portal for clathrinmediated endocytosis of distinct cargo governed by cargo-selective adaptors. Mol Biol Cell 17:4300–17. Kirchhausen T, Harrison SC, Chow EP, Mattaliano RJ, Ramachandran KL, Smart J, Brosius J. 1987. Clathrin heavy chain: molecular cloning and complete primary structure. Proc Natl Acad Sci U S A 84:8805–9. Luttrell LM, Lefkowitz RJ. 2002. The role of betaarrestins in the termination and transduction of G-proteincoupled receptor signals. J Cell Sci 115:455–65. Maldonado-Baez L, Wendland B. 2006. Endocytic adaptors: recruiters, coordinators and regulators. Trends Cell Biol 16:505–13. Maxfield FR, Schlessinger J, Shechter Y, Pastan I, Willingham MC. 1978. Collection of insulin, EGF and alpha2-macroglobulin in the same patches on the surface of cultured fibroblasts and common internalization. Cell 14:805–10. Mettlen M, Pucadyil T, Ramachandran R, Schmid SL. 2009. Dissecting dynamin’s role in clathrin-mediated endocytosis. Biochem Soc Trans 37:1022–6. Mishra SK, Hawryluk MJ, Brett TJ, Keyel PA, Dupin AL, Jha A, Heuser JE, Fremont DH, Traub LM. 2004. Dual engagement regulation of protein interactions with the AP-2 adaptor alpha appendage. J Biol Chem 279: 46191–203. Mundell SJ, Luo J, Benovic JL, Conley PB, Poole AW. 2006. Distinct clathrin-coated pits sort different G proteincoupled receptor cargo. Traffic 7:1420–31. Nakatsu F, Ohno H. 2003. Adaptor protein complexes as the key regulators of protein sorting in the post-Golgi network. Cell Struct Funct 28:419–29. Ohno H, Stewart J, Fournier MC, Bosshart H, Rhee I, Miyatake S, Saito T, Gallusser A, Kirchhausen T, Bonifacino JS. 1995. Interaction of tyrosine-based sorting signals with clathrin-associated proteins. Science 269:1872–5. Onuchic JN, Wolynes PG. 2004. Theory of protein folding. Curr Opin Struct Biol 14:70–5.

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Owen DJ. 2004. Linking endocytic cargo to clathrin: structural and functional insights into coated vesicle formation. Biochem Soc Trans 32:1–14. Pearse BM. 1976. Clathrin: a unique protein associated with intracellular transfer of membrane by coated vesicles. Proc Natl Acad Sci U S A 73:1255–9. Praefcke GJ, Ford MG, Schmid EM, Olesen LE, Gallop JL, Peak-Chew SY, Vallis Y, Babu MM, Mills IG, McMahon HT. 2004. Evolving nature of the AP2 alphaappendage hub during clathrin-coated vesicle endocytosis. EMBO J 23:4371–83. Puthenveedu MA, von Zastrow M. 2006. Cargo regulates clathrin-coated pit dynamics. Cell 127:113–24. Robinson MS. 1989. Cloning of cDNAs encoding two related 100-kD coated vesicle proteins (alpha-adaptins). J Cell Biol 108:833–42. Roth MG. 2004. Phosphoinositides in constitutive membrane traffic. Physiol Rev 84:699–730. Roth MG. 2006. Clathrin-mediated endocytosis before fluorescent proteins. Nat Rev 7:63–8. Roth TF, Porter KR. 1964. Yolk protein uptake in the oocyte of the mosquito Aedes aegypti L. J Cell Biol 20: 313–32. Saffarian S, Cocucci E, Kirchhausen T. 2009. Distinct dynamics of endocytic clathrin-coated pits and coated plaques. PLoS Biol 7:e1000191. Semerdjieva S, Shortt B, Maxwell E, Singh S, Fonarev P, Hansen J, Schiavo G, Grant BD, Smythe E. 2008. Coordinated regulation of AP2 uncoating from clathrin-coated vesicles by rab5 and hRME-6. J Cell Biol 183:499–511.

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Teckchandani A, Toida N, Goodchild J, Henderson C, Watts J, Wollscheid B, Cooper JA. 2009. Quantitative proteomics identifies a Dab2/integrin module regulating cell migration. J Cell Biol 186:99–111. Traub LM. 2009a. Clathrin couture: fashioning distinctive membrane coats at the cell surface. PLoS Biol 7: e1000192. Traub LM. 2009b. Tickets to ride: selecting cargo for clathrin-regulated internalization. Nat Rev 10:583–96. Ungewickell E, Branton D. 1981. Assembly units of clathrin coats. Nature 289:420–2. Ungewickell EJ, Hinrichsen L. 2007. Endocytosis: clathrin-mediated membrane budding. Curr Opin Cell Biol 19:417–25. Vigers GP, Crowther RA, Pearse BM. 1986. Location of the 100 kd-50 kd accessory proteins in clathrin coats. EMBO J 5:2079–85. Wenk MR, Pellegrini L, Klenchin VA, Di Paolo G, Chang S, Daniell L, Arioka M, Martin TF, De Camilli P. 2001. PIP kinase Igamma is the major PI(4,5) P(2) synthesizing enzyme at the synapse. Neuron 32: 79–88. Zaremba S, Keen JH. 1983. Assembly polypeptides from coated vesicles mediate reassembly of unique clathrin coats. J Cell Biol 97:1339–47. Zoncu R, Perera RM, Sebastian R, Nakatsu F, Chen H, Balla T, Ayala G, Toomre D, De Camilli PV. 2007. Loss of endocytic clathrin-coated pits upon acute depletion of phosphatidylinositol 4,5-bisphosphate. Proc Natl Acad Sci U S A 104:3793–8.

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CH A P T E R

3

CAVEOLAE Dan Tse Radu V. Stan

DEFINITION Caveolae constitute a plasma membrane microdomain abundant in vertebrate cells, with few exceptions. They are defined as spherical invaginations of the plasma membrane with a strikingly constant diameter in the range of 60–70 nm. The invaginations can exist singly or in grapelike clusters with noncaveolar plasma membrane between the invaginations. Caveolae neck (or stoma, or introit) is defined as the transition region where the membrane of the invagination continues the membrane of the plasmalemma proper. In endothelial cells, the neck contains a protein structure called a stomatal diaphragm (SD). Select members of the caveolin (Cav) and cavin families of proteins are critical for the formation of the caveolar invaginations. The Cav family comprises Cav1–3 and one invertebrate member. Cav1 and Cav3 are critical for the formation of caveolae in cells where they are expressed. Cavin family of proteins is less well described. It contains four isoforms such as cavin-1/Pol I and transcript release factor (PTRF), cavin-2/serum deprivation protein response (SDPR), cavin-3/sdr-related gene product that binds to c-kinase (SRBC), and the muscle-cell-specific cavin-4/muscle-restricted coiled-coil (MURC4). Of these, cavin-1 and cavin-4 were shown to be critical for caveolae formation. Caveolae formation also requires a certain set of lipids such as cholesterol and sphingolipids, representing a subdomain of lipid rafts (see also Chapters 4 and 5 on lipid rafts) as well as phosphatidylserine (PS), which are enriched in the caveolae membrane but present in all membranes. Although both classes of preoteins are required for caveolae formation, at this point in time, caveolins seem to define the best caveolae not each individual member of the cavin family being present in every caveola. However, caveolins can also exist on flat plasma membrane, even in cells lacking morphologically distinct caveolae; thereby, their mere presence in the absence of invaginations does not define caveolae. Thus, at this point in time, defining caveolae should rely on a double criterion of morphological invagination and imunocytochemical demonstration of Cav1 on these invaginations, achieved only by electron microscopy (EM).

HISTORICAL PERSPECTIVE The introduction of the electron microscope to the study of biological materials in the second half of the last century has dramatically expanded our understanding of the inner workings of cells. It enabled a first glimpse of the cellular architecture leading to the discovery and study of subcellular organelles and microdomains. Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Uncoated membrane invaginations in the size range of caveolae were first described in 1953 in the continuous endothelium of the heart by G.E. Palade (Palade 1953) who called them plasmalemmal vesicles. Two years later, E. Yamada described similar structures in the gall bladder epithelium, naming them caveolae intracellulares (Yamada 1955) due to their resemblance to “little caves.” Since their discovery, the presence of caveolaelike invaginations has been documented by electron microscopists in most cell types (reviewed in Parton 2003; Stan 2005b). These studies revealed the structural features of caveolae (Peters et al. 1985) as well as some of their protein (Fujimoto et al. 1992; Fujimoto 1993) and lipid residents. Functional postulates were also formulated such as transcytosis of physiologically relevant cargo across endothelial cells (Palade 1960; Simionescu 1981) or uptake of lipid—binding cholera and tetanus toxins (Montesano et al. 1982) by noncoated vesicles. In muscle cells, caveolae were associated with the developing T tubules (Ishikawa 1968) and were proposed to be involved in excitation–contraction coupling (Popescu 1974; Fry et al. 1977; Severs 1988). Caveolae were also postulated to act as stretch sensors and/or as reservoirs of membrane during cycles of contraction and relaxation (Severs 1988). These studies banked on a purely morphological definition of caveolae. In the absence of molecular markers, terms such as caveolae or plasmalemmal vesicles were used to describe morphological entities that do not always correspond to the operational definitions in use today. Moreover, several uncoated membrane invaginations in the size range of caveolae but devoid of Cav1 are found in cells (Mayor and Pagano 2007; Hansen and Nichols 2009, 2010; Howes et al. 2010; Kumari et al. 2010); therefore, care should be exercised when interpreting these early data. With the discovery of Cav1 (Glenney 1992; Glenney and Soppet 1992; Kurzchalia et al. 1992) as the first molecular marker of caveolae (Rothberg et al. 1992; Dupree et al. 1993) in the early 1990s, the study of caveolae was ushered into the molecular era. After this date, ensuing biochemical and cell biological approaches have revealed numerous features of caveolae and served as basis for hypotheses involving caveolae in a plethora of cellular functions such as vesicular trafficking, signal transduction, lipid homeostasis, and cell growth and division as well as adhesion and motility. Many excellent reviews discuss these findings (Cohen et al. 2004; Parton and Simons 2007; Hansen and Nichols 2009, 2010; Lajoie et al. 2009a). Genetic models in which critical components of caveolae such as caveolins (Drab et al. 2001; Galbiati et al. 2001; Razani et al. 2001, 2002; Park et al. 2002; Zhao et al. 2002; Cao et al. 2003; Bauer et al. 2005) and cavins (Hill et al. 2008; Liu et al. 2008; Bastiani et al. 2009; Hansen et al. 2009) were depleted have tremendously helped in defining the biogenesis, function, and regulation of caveolae. Lastly and most excitingly, caveolae and their residents have been demonstrated to play important roles in disease, such as atherosclerosis, cancer, inflammation, calcium phosphate balance, kidney stone disease, and muscular dystrophy, and to be involved in host–pathogen interactions.

DISTRIBUTION AND MORPHOLOGY Caveolae occur at different surface densities in different cell types with endothelial cells, adipocytes, and muscle cells being the richest. In adipocytes, ∼20% of the plasma membrane was estimated to be caveolar membrane (Sheldon et al. 1962; Williamson 1964). There is marked variation in caveolar surface density within a given cellular type, which is best exemplified in the case of the different types of endothelium (Simionescu et al. 1974) or mesothelia (von Ruhland et al. 2004). Variations in numbers of surface-connected

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Figure 3.1. (A) Endothelium of continuous type as seen by freeze fracture. The P face of the abluminal membrane of a heart EC shows the numerous caveolae. These are organized in linear arrays, better seen in the higher magnification images (insets). Reproduced from Simionescu et al. (1974) with permission. (B–E) Caveolae with (D–E) and without (B–C) SDs as seen by TEM (C, E) or in deep-etch specimens (B, D). (B, D) Reproduced from Bearer and Orci (1985) with permission.

caveolae are caused by differentiation status (Williamson 1964; Novikoff et al. 1980), oncogenic transformation (Koleske et al. 1995; Lee et al. 1998), shear stress (Rizzo et al. 2003), and cell adhesion (del Pozo et al. 2004). Please see Parton (2003), Stan (2005b), and Parton et al. (2006) for a more detailed analysis of the morphological aspects of caveolae. Caveolae occur in ordered, linear arrays over the entire cell body (Fig. 3.1A), suggesting a link with the underlying cytoskeleton (Simionescu et al. 1974; Tani et al. 1977; Gabella 1978; Sawada et al. 1978; Frank et al. 1980; Severs 1981; Peters et al. 1985; Izumi et al. 1988; Parton et al. 1994; Fujimoto et al. 2000). In migrating cells, caveolins and caveolae seem to preferentially distribute to the retracting edge of the cell (Isshiki et al. 1998, 2002; Parat et al. 2003; Beardsley et al. 2005). In many cell types, caveolae occur single or in chains or grapelike clusters. In the endothelium, these are attached to either front of the cell (Bruns and Palade 1968a, b; Palade and Bruns 1968; Bundgaard et al. 1979, 1983; Frokjaer-Jensen 1980, 1984, 1991; Frokjaer-Jensen et al. 1988). In other epithelia, caveolae are preferentially attached to the basolateral surface such as in Madin-Darby canine kidney (MDCK) cells and in CaCo2 cells upon de novo expression of Cav1 (Vogel et al. 1998). In muscle cells (Ishikawa 1968; Parton et al. 1997) or in freshly isolated adipocytes (Thorn et al. 2003), caveolae occur mostly as single entities rather than clusters. In adipocytes (Williamson 1964; Novikoff et al. 1980; Kanzaki and Pessin 2002; Parton et al. 2002), endothelia (Simionescu et al. 1974; Uehara and Miyoshi 1999), or epithelial cells (Parton 1994), caveolae could be attached to large, fingerlike invaginations of the plasma membrane surrounded by cortical actin, called “cav-actin” (Kanzaki and Pessin 2002), “caves” (Parton et al. 2002), or “caveolae rosettes” (Fig. 3.2B) (Novikoff et al. 1980). Rosette numbers seem to be increased in endothelial cells upon overexpression of intersectin, a protein involved in caveolae endocytosis (Predescu et al. 2003) or astrocytes under the influence of okadaic acid, a phosphatase inhibitor (Bento-Abreu et al. 2009).

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(C)

(A)

(D)

(B)

(E)

(F)

(G) (H)

(I)

Figure 3.2. Caveolar coat. Striations have been demonstrated by TEM in smooth muscle (A) and fibroblasts (F). They are readily apparent by either SEM on chromium-coated replicas of endothelial cells (B) or in platinum-carbon replicas of deeply etched specimens in fibroblasts (C, E–F). The ridges are best seen when caveolae are flattened by treatment with cholesterolbinding drugs (nystatin) (E–F). (F) Immunolocalization of Cav1 to the caveolar coat ridges. Reprints from (A) Somlyo et al. (1971), (B) Peters et al. (1985), and (C–F) Rothberg et al. (1992) with permission. B is courtesy of Rob Parton.

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In frozen, unfixed specimens caveolae surface density is reduced, suggesting that fixation might induce the formation of the invaginations (McGuire and Twietmeyer 1983; Wood et al. 1986; Noguchi et al. 1987). Moreover, there are differences between the typical morphology of caveolae in aldehyde-fixed tissues and caveolae in unfixed frozen tissues. In the latter, caveolae are slightly larger, have the aspect of cups rather than omega-shaped invaginations, and can be found with different degrees of membrane curvature (Richter et al. 2008). On their cytoplasmic face, caveolar invaginations feature a filamentous coat formed by Cav polymerization (Fernandez et al. 2002; Parton et al. 2006). The existence of the coat was first suggested by transmission electron microscopy (TEM) in smooth muscle caveolae (Somlyo et al. 1971) (Fig. 3.2A) and later confirmed by high-resolution scanning electron microscopy (SEM) in endothelial cells (Peters et al. 1985) and by rapid-freeze deep-etch techniques in endothelial cells (ECs) and fibroblasts in both unfixed and glutaraldehyde-fixed cells (Izumi et al. 1988, 1989, 1991). The coat ridges are enhanced by phalloidin and could be decorated with myosin fragments (Izumi et al. 1989). Carbonplatinum deep-etch replicas of fibroblast plasma membrane sheets showed that the coat contains Cav1-positive “spiral patterns” resistant to high-salt and high-pH extraction (Fig. 3.2E, G–I) (Rothberg et al. 1992). Cholesterol-binding drugs (i.e., nystatin) flatten caveolae and disassemble the coat (Rothberg et al. 1992) (Fig. 3.2G); the fragmentation of the coat resulted in what appeared to be ∼10-nm-diameter uniform units (Fernandez et al. 2002). TEM tomography combined with manual annotation (Richter et al. 2008) or powerful computer-assisted template matching (Lebbink et al. 2010) have recently demonstrated a spiral coat on caveolae in unfixed high-pressure freeze-substituted cells or tissues.

MOLECULAR COMPONENTS Cav Family of Proteins An important milestone in the field was achieved with the demonstration by immunocytochemistry that Cav1 is a specific component of the ridges that make the caveolar coat (Rothberg et al. 1992) (Fig. 3.2H–I). This advanced the operational definition of caveolae to plasma membrane invaginations that contain caveolins. The Cav family of proteins encompassing Cav1 (Glenney and Soppet 1992), Cav2 (Scherer et al. 1996), and Cav3 (Way and Parton 1995) is part of a gene family well conserved in amniotes (Kirkham et al. 2008). Cav1 and Cav2 have similar tissue distribution being expressed in most cell types (Kurzchalia et al. 1994; Parton et al. 1994; Scherer et al. 1997; Scheiffele et al. 1998), while Cav3 occurs primarily in striated muscle cells (Way and Parton 1995). Cav1 and Cav3 seem to coexist in different subdomains in some smooth muscle cells. Cav1 was identified as a v-Src phosphorylation substrate in Rous sarcoma virustransformed fibroblasts (Glenney 1992; Glenney and Soppet 1992) and independently cloned as VIP21, a component of the trans-Golgi-derived vesicles (Kurzchalia et al. 1992, 1994). Cav1 has two isoforms: Cav1α contains residues 1–178 and Cav1β contains residues 32–178. Cav1β is translated from a different mRNA than Cav1α (Kogo and Fujimoto 2000; Kogo et al. 2004). Cav1 is phosphorylated on serine/threonine sites by protein kinase Cα (PKCα) (Smart et al. 1995; Mineo et al. 1998; Isshiki and Anderson 1999) and on tyrosines (Tyr14 being the most important) by c-Src and possibly other nonreceptor tyrosine kinases (Glenney and Soppet 1992; Li et al. 1996a; Tiruppathi et al. 1997; Aoki et al. 1999; Nomura and Fujimoto 1999; Caselli et al. 2001; Volonte et al. 2001; Cao et al. 2002; Kim et al. 2002; Kimura et al. 2002; Labrecque et al. 2004; Nohe et al. 2005).

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Tyrosine phosphorylation seems to occur mainly on Cav1α. The two Cav1 isoforms have a different subcellular distribution (Fujimoto et al. 2000) and can be expressed in a celltype-specific manner (Kogo et al. 2004). Cav1 and Cav3 (but not Cav2) are palmitoylated on three cysteines in the C-terminal region, but the palmitoylation is not necessary for Cav targeting to caveolae (Dietzen et al. 1995). In ECs, the palmitoylation occurs posttranslationally and is irreversible, which is an important difference from other acylated proteins where palmitoylation is reversible and regulatable (Dunphy and Linder 1998; Parat and Fox 2001; Resh 2004). Based on these last data and the demonstration that palmitoylation stabilizes the Cav oligomers (Monier et al. 1996), the palmitoylation of Cav has been proposed to have a role in increasing its membrane association mediated by the hydrophobic domain (Parat and Fox 2001). Another feature of the Cav molecule is the “scaffolding domain,” a region adjacent to the hydrophobic domain used by Cavs to interact with signal transduction molecules (Sargiacomo et al. 1995; Schlegel et al. 2001). Unconventional membrane attachment domains responsible for targeting the molecule to caveolae (Schlegel et al. 2001) and Golgi (Luetterforst et al. 1999) (see also Chapter 8 on the Golgi apparatus) have been proposed to exist in the Cav molecule. However, a recent paper (Ren et al. 2004) questioned the existence of a specific targeting domain, suggesting that the disruption of the overall conformation of Cav molecule would impair caveolae targeting, raft association, and Golgi exit. Cav2 has three known isoforms (α, β, γ) that were discovered in adipocytes (Scherer et al. 1996). Cav2α is the full-length isoform, whereas 2β is an alternatively spliced isoform that has a different subcellular distribution than Cav2α (Kogo et al. 2002). The phosphorylation of Cav2 on serines 23 and 36 modulates Cav1-dependent caveolae formation (Sowa et al. 2003). Cav2 is also phosphorylated on tyrosine 19 or 27. Phospho-Cav2 (pY19) and phospho-Cav2 (pY27) display different cellular localization patterns (Wang et al. 2004). Finally, mice lacking Cav2 show severe pulmonary dysfunction but without disruption of caveolar invaginations (Razani et al. 2002). Cav3 (Way and Parton 1996) is expressed in the skeletal muscle and myocardium where it is essential for the formation of caveolae (Hagiwara et al. 2000; Galbiati et al. 2001). It was also shown to be expressed in smooth muscle cells (Song et al. 1996), astrocytes (Ikezu et al. 1998), and sinus endothelial cells (Uehara and Miyoshi 2002). Loss of Cav3 results in muscle degeneration (Hagiwara et al. 2000), cardiomyopathy (Woodman et al. 2002), and increased adipose tissue deposition (Park et al. 2002). Increased Cav3 levels are associated with muscle dystrophy, cardiomyopathy, and cardiac hypertrophy inhibition (Repetto et al. 1999; Galbiati et al. 2000; Aravamudan et al. 2003). For more detailed reviews on Cavs, see Gratton et al. (2004) and Williams and Lisanti (2004b).

Cav1 and Cav3, but Not Cav2, Are Essential for the Formation of Caveolar Invaginations Several lines of evidence demonstrate the essential role of Cav proteins in the structure of caveolae. Cav1 and Cav3 expression in cells lacking both caveolae and Cav expression induces caveolae formation (Fra et al. 1995b) and reconstitutes the caveolar coat (Chung et al. 1996). Both Cav1α and Cav1β isoforms are able to induce the formation of caveolae de novo, Cav1β being less efficient (Fujimoto et al. 2000). Expression of Cav2 alone does not lead to caveolae formation; in fact, Cav2 alone does not traffic further than Golgi, needing expression of Cav1 in order to reach plasma membrane caveolae (Scheiffele et al. 1998; Mora et al. 1999; Parolini et al. 1999).

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The most compelling evidence for the structural role of Cav1 and Cav3 in the formation of caveolae came from genetic mouse models lacking one or both of these proteins (Drab et al. 2001; Razani et al. 2001; Park et al. 2002; Zhao et al. 2002). It has been shown that upon Cav3 gene disruption, muscle cells do not have any caveolae while they are still present in other cell types (Hagiwara et al. 2000; Galbiati et al. 2001). Conversely, Cav1−/− mice lack caveolae in all cells except in muscle cells (Drab et al. 2001; Zhao et al. 2002). Loss of Cav2 does not disrupt caveolae (Razani et al. 2002), Cav2−/− mice having unaltered number of caveolae. Taken together, these data show that Cav1 and Cav3 are essential for caveolae formation, whereas Cav2 is not. However, larger (100–150 nm) invaginations of the plasma membrane and vesicle clusters are present in the endothelium of the venules in the heart and lungs of Cav1−/− mice (Drab et al. 2001; Zhao et al. 2002). Recently, these structures were shown to be vesiculovacuolar organelles (VVOs) (Chang et al. 2009). This provides genetic evidence that other smooth invaginations could exist in absence of Cav1, and by our operational definition, these invaginations are not caveolae.

Cavins Recently, it became apparent that a new family of proteins, now known as cavins, is intimately involved in caveolae biogenesis. These are cavin-1/PTRF (Hill et al. 2008; Liu and Pilch 2008; Liu et al. 2008), cavin-2/SDPR (Gustincich et al. 1999; Hansen et al. 2009), cavin-3/SRBC (McMahon et al. 2009), and the muscle-specific cavin-4/MURC (Bastiani et al. 2009). Their role in caveolae biogenesis will be discussed below (reviewed in Briand et al. 2010, Chidlow and Sessa 2010, and Hansen and Nichols 2010). In terms of common features, all cavins bind PS, which is important in their targeting to the plasma membrane. They contain consensus sites and domains such as phosphorylation motifs (phosphorylated upon insulin stimulation), leucine zipper-like domains through which they interact with other proteins, and PEST (proline, glutamic acid, serine, and threonine) domains involved in protein turnover (Rahman and Sward 2009). Cavin-1 or PTRF was cloned as a transcript release factor (Jansa et al. 1998, 2001; Jansa and Grummt 1999). It was later shown to be a major caveolar protein mainly in adipocytes (Vinten et al. 2001, 2005; Aboulaich et al. 2004) and other tissues (Voldstedlund et al. 2001, 2003). It localizes to caveolae, and its total cellular levels correlate with those of Cav1 and the number of caveolae (Voldstedlund et al. 2001, 2003; Aboulaich et al. 2004; Liu and Pilch 2008). Cavin-1 was subsequently shown to be essential for caveola formation in mammalian cells (Hill et al. 2008; Liu et al. 2008; Liu and Pilch 2008), zebra fish (Hill et al. 2008), and mice (Liu et al. 2008). Cavin-2 or SDPR (or SDR) was isolated as a PS-binding protein from platelets (Burgener et al. 1990; Gustincich et al. 1999). It was shown in vitro to bind to and be a substrate for protein kinase C (PKC) isoforms in caveolae (Izumi et al. 1997; Mineo et al. 1998). Cavin-2 levels are increased by serum deprivation but not by contact inhibition (Gustincich et al. 1999). The presence of cavin-2 in caveolae was demonstrated by immunocytochemistry (Mineo et al. 1998). Cavin-2 has a role in the formation of caveolae, as demonstrated in cells in culture (Hansen et al. 2009; McMahon et al. 2009). Cavin-2 binds directly to cavin-1 and recruits it to caveolar membranes. Overexpression of cavin-2, unlike cavin-1, induces deformation of caveolae and extensive tubulation of the plasma membrane. Cavin-2 colocalizes extensively with both B-subunit of Shiga toxin (STB) and Cav1 on the similar membrane tubes induced by STB, which is lost when caveolae are absent (Hansen et al. 2009). Cavin-3 or SRBC (Gustincich et al. 1999) or protein kinase C delta-binding protein (PRKCDBP) (Burgener et al. 1990) was shown to co-immunoprecipitate with Cav1 and

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Cav2 (Bastiani et al. 2009; McMahon et al. 2009) and to remain colocalized with Cav1 when caveolae bud off (McMahon et al. 2009). Its downregulation impairs intracellular trafficking of Cav1-containing “cavicles” but does not change the number of caveolae (McMahon et al. 2009). Cavin-4 (Bastiani et al. 2009) or MURC4 (Ogata et al. 2008; Tagawa et al. 2008) is a muscle-specific, coiled-coil-containing protein involved in muscle biogenesis. It forms complexes with other cavins and has a role in caveolae biogenesis (Bastiani et al. 2009).

CAVEOLAR LIPIDS Indications of a specific lipid composition of the caveolar membranes came from EM experiments in different cell types employing cholesterol-binding probes and showed an enrichment of cholesterol in caveolae (Simionescu et al. 1983; Fujimoto 1996; Fujimoto et al. 1997). Subsequent studies have demonstrated that Cav1 binds tightly to free cholesterol and to bind to artificial phospholipid liposomes only upon cholesterol incorporation (Murata et al. 1995; Tillmann et al. 1995; Li et al. 1996b). Moreover, cholesterol is necessary to stabilize Cav oligomers, both sterol and protein being necessary to generate and maintain caveolar structure (Monier et al. 1996). These data and the sensitivity of caveolae architecture to cholesterol depletion (Rothberg et al. 1992) and oxidation (Smart et al. 1994) have established caveolae as cholesterol-enriched membrane microdomains. Caveolae are also enriched in sphingolipids (such as sphingomyelin, ceramide, and gangliosides) that occur mostly in the exoplasmic leaflet of the membrane bilayer (Parton 1994; Smart et al. 1994; Liu and Anderson 1995; Schnitzer et al. 1995a; Fujimoto 1996; Fujimoto et al. 1997). Cav1 also binds glycosphingolipids (such as GM1 ganglioside) (Fra et al. 1995a) and fatty acids (Trigatti et al. 1999). The lipid composition of caveolae (i.e., cholesterol, sphingomyelin, glycosphingolipids, and saturated fatty acids) is similar to that of “lipid rafts,” membrane microdomains that occur in all cell types (Brown and London 2000; Simons and Toomre 2000; Shogomori and Brown 2003; Simons and Vaz 2004). Comprehensive comparison of the lipid composition of the caveolae/lipid raftenriched fractions from cells that express or not Cav1 (and caveolae), also show that both lipid rafts and caveolae are enriched in cholesterol, sphingomyelin, arachidonic acid containing ethanolamine plasmalogens, and PS with decreases in phosphatidylinositol and phosphatidylcholine (Pike et al. 2002). Interestingly, only cholesterol and not sphingomyelin was enriched in raft fractions of cells expressing Cav1 over cells that do not express it. Because Cav1 could not directly bind all the excess cholesterol present in the caveolae/ lipid raft fraction, it was postulated that it might direct the organization of a structure that facilitates cholesterol enrichment (Pike et al. 2002). It was also concluded that the extra cholesterol must either be sequestered in some fashion or restricted to the inner leaflet of the plasma membrane so that it does not alter the balance of cholesterol and sphingomyelin at the cell surface. The observation that lipid rafts and caveolae were highly enriched in PS in the cytoplasmic leaflet of the membrane (Pike et al. 2002) nicely dovetails with recent findings that all cavin proteins that are required for caveola biogenesis bind PS (Hill et al. 2008; Liu and Pilch 2008; Liu et al. 2008; Bastiani et al. 2009; Hansen et al. 2009). Lipid analysis of immunoisolated adipocyte caveolae (Ortegren et al. 2004) found the same cholesterol, sphingomyelin, and glycerophospholipid makeup. In addition, glycosphingolipid GD3 was highly enriched in caveolae, whereas GM3, GM1, and GD1a were present inside as well as outside the caveolae membrane. GD1b, GT1b, GM2, GQ1b, sulfatide, and lactosylceramide sulfate were not detected in adipocyte caveolae (Ortegren et al. 2004).

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CAVEOLAR NECKS: TRANSITION ZONE BETWEEN CAVEOLAR MEMBRANE AND PLASMALEMMA PROPER The neck of caveolae is the region where the membrane of the caveolar invagination is in continuity with that of plasmalemma proper. In aldehyde-fixed tissues, this region forms an inverted bend, which, in some cases, appears rounded, whereas in others, it appears as a sharp angle. Electron opaque material fills the cytoplasmatic aspect of the neck. The sharp angle is mostly present in endothelial cells where caveolae are provided with an SD (see below). However, electron microscopic examination of unfixed frozen tissues or highpressure freeze-substituted samples revealed differences with the fixed material. In the former conditions, caveolae have a cuplike appearance with a much larger introit and no electron opaque material around the neck (Westermann et al. 2005; Richter et al. 2008; Schlormann et al. 2010). The same is encountered in endothelial caveolae featuring SDs (Richter et al. 2008). The neck region seems to have a different composition than the rest of the caveolar body. In endothelial cells (Moldovan et al. 1995; Schnitzer et al. 1995b; Stan et al. 1999a) and adipocytes (Foti et al. 2007), the neck is readily soluble in anionic detergents. Insulin receptors seem to be concentrated at the neck of adipocyte caveolae (Foti et al. 2007). In endothelial cells, the neck contains a specialized SD (Bruns and Palade 1968a, b; Clementi and Palade 1969a, b; Stan et al. 1999a, b; Stan 2004; Stan et al. 2004) (reviewed in Stan 2005b, 2007) (Figs. 3.1D–E and 3.3). The existence of caveolae with or without SDs has been clearly demonstrated by rapid-freeze deep-etch techniques (Bearer and Orci 1985; Noguchi et al. 1987; Fig. 3.1B–E) as well as unfixed high-pressure freezesubstituted samples (Richter et al. 2008). The SD is a thin (∼5–6 nm) protein (Simionescu et al. 1981, 1982) barrier of unknown function that occurs in the caveolar neck (introit, mouth, or stoma—hence the name) or at the stomata (or communication) between fused caveolae. Similar diaphragms occur in normal endothelium on transendothelial channels (TECs), fenestrae (Clementi and Palade 1969a, b; Stan et al. 2004), or VVOs in the vascular endothelium in tumors and the endothelium of normal venules (Dvorak and Feng 2001). Recently, the SDs were postulated to be formed by the Plvap/PV1 (Fig. 3.3), type II membrane glycoprotein found in vertebrates (Stan et al. 1999a, b, 2001, 2004; Stan 2004). PV1 is a key structural component of both the SDs and fenestrated diaphragms (FDs), necessary and sufficient for diaphragm formation (Stan et al. 2004). This is based on several lines of evidence: (1) Several PV1 homodimers reside in close proximity within the same diaphragm as shown by cross-linking experiments of PV1 carried out in situ in rat lungs and kidneys (Stan 2004). (2) De novo formation of FDs and SDs correlates with PV1 expression. As expected, PV1 was found in the newly formed FDs and SDs. (3) PV1 mRNA silencing by siRNA prevented the formation of both FDs and SDs. (4) Expression of tagged PV1 in either endothelial cells or nonendothelial cells (e.g., fibroblasts, COS7, HeLa, HEK293), which lack PV1 expression and diaphragms, led to the formation of caveolar SDs (Stan et al. 2004). A model of PV1 integration in the diaphragms was recently proposed (Stan 2005b, 2007).

CAVEOLAE BIOGENESIS Immediately after synthesis in the endoplasmic reticulum (ER; see also Chapter 7), Cav1 assembles into 8S complexes that concentrated in ER exit sites, due to a DXE sequence in the N-terminal domain (Hayer et al. 2010). The coat protein II (COPII) machinery

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Caveolae

TEC

Fenestrae

(A)

Stomatal diaphragms

(B)

Fenestral diaphragms

(C)

Rim (plasma membrane) (D)

N-glycosylation Central knob

PV1 dimer

Scaffold proteins Diaphragm filbrils

Stan, figure

Rim

Figure 3.3. Working model for PV1 integration in the structure of the diaphragms. (A) Perpendicular TEM sections of a caveola provided with SD (left), TEC (middle), and fenestra (right) from specimens labeled with anti-PV1 antibodies, demonstrating the presence of PV1 in these structures. (B) En face views of SDs (left) and FDs (right), as shown by deep-etch rapidfreeze techniques, demonstrating the fibrils in the FDs and the hints of fibrils in their SD counterparts. Reprinted from Bearer and Orci (1985) with permission. (C) Schematic of the membrane insertion and features of the PV1 monomer. (D) Model of PV1 integration in the endothelial diaphragms. Left panel: PV1 dimers participate in the formation of the fibrils inserted in the rim (via PV1 N-terminus) and interweaving in the central mesh (via PV1 C-terminus). The glycan antennae (accounting for ∼15% of PV1 mass) are situated near the membrane, which would keep the protein “afloat” by preventing collapse on the plasma membrane. The sharp angle formed by the plasma membrane at the level of the rim is maintained by “scaffold proteins” that can be either transmembrane or associated with the cytosolic face of the membrane. Right panels: PV1 dimer seen from the top (up) and in a section perpendicular on the diaphragm plane (bottom).

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allows rapid transport to the Golgi complex where the Cavs lose their diffusional mobility, undergo conformational changes (Ren et al. 2004), and assemble into 70S complexes by association with cholesterol (Hayer et al. 2010). Together with green fluorescent protein– glycosylphosphatidylinositol (GFP-GPI), the newly assembled Cav complexes undergo transport to the plasma membrane in vesicular carriers distinct from those containing vesicular stomatitis virus (VSV) G protein (Hayer et al. 2010). After the arrival at the plasma membrane, PTRF/cavin-1 is recruited to the caveolar domains over a period of 25 minutes or longer (Hayer et al. 2010). PTRF/cavin-1 is present in 60S complexes that form in the absence of Cav1 (Hayer et al. 2010) possibly by interactions with other cavins (Hansen et al. 2009). Cavin-1 was postulated to act as a stabilizer of caveolae (Hill et al. 2008). Its lack results in loss of caveolae and a higher lateral mobility of Cav1, which forms a uniform smooth pattern into the membrane and is targeted for degradation via endocytosis (Hill et al. 2008). In cancer cells that lack both Cav1 and cavin-1, reintroduction of Cav1 results in membrane tubules that contain Rab8 and EHD (dynamin-like C-terminal Eps15 homology domain) proteins (Verma et al. 2010). Actomyosin-induced tension causes the tubules to disperse into small vesicles (Verma et al. 2010). Similar tubules are formed when cavin2/SDPR is overexpressed (Hansen et al. 2009). Coexpression of cavin-1 and Cav1 in these cells results in the inhibition of membrane tubulation induced by Cav1 (Verma et al. 2010). At this point in time, the essential role of Cavs in the formation of the invaginations is pretty well established. The evidence for a coat on the cytoplasmic face of caveolae is very strong and the list of the structural proteins required for caveolae formation is growing. What is unclear is the precise role of the coat in caveolae structure and the precise spatial arrangement of its molecular makeup.

CELLULAR FUNCTIONS OF CAVEOLAE Although they were implicated in many cellular functions, the precise roles of caveolae are still debated (for a review, see Parton and Simons 2007). As better, more discriminative tools became available, some caveolar roles begin to crystallize.

Endocytosis of Caveolae Early data demonstrated noncoated invaginations to participate in transcytosis across endothelial cells (Palade 1960) and endocytosis (Montesano et al. 1982). The capacity of caveolae to internalize ligands was further reinforced during the years and much was learned about the process (Parton and Simons 2007; Howes et al. 2010; Kumari et al. 2010). At the cell surface, caveolae have low lateral mobility as shown by fluorescence recovery after photobleaching (FRAP) experiments (Thomsen et al. 2002) and by cell fusion experiments (Tagawa et al. 2005). They describe kiss-and-run movements (Pelkmans and Zerial 2005) and internalize quite slowly (Thomsen et al. 2002; Pelkmans et al. 2004; Kirkham et al. 2005) in actin cytoskeleton, tyrosine kinase, and dynamin-2-dependent manner (Oh et al. 1998; Pelkmans et al. 2001, 2002, 2004; Tagawa et al. 2005). Caveolae endocytosis is triggered by cell detachment from the extracellular cell matrix (del Pozo et al. 2004, 2005), by cargo molecules such as serum albumin (Tiruppathi et al. 1997) or SV40 virus (Pelkmans et al. 2001; Tagawa et al. 2005), or treatments with phosphatase inhibitors (Parton et al. 1994; Vepa et al. 1997). Cell loading with either cholesterol or glycosphingolipids (Sharma et al. 2003; Singh et al. 2003), or removal of sialic acid

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moieties from glycoproteins and gangliosides (Singh et al. 2010), was also reported to increase caveolae internalization. After internalization caveolae carriers fuse with either the caveosomes or the Rab5positive early endosomes (see also Chapter 9) from where they can be recycled to the plasma membrane (Pelkmans et al. 2004). Very interestingly, caveolae seem to remain as a stable structure upon fusing with these compartments exchanging soluble cargo but not membrane (Pelkmans et al. 2004). It is not very clear how fission and fusion occur in the light of the apparent stability of the caveolae containers. A possibility would be a small subdomain of more fluid membrane (possibly derived from the caveolar neck) that can support these processes. The field is at an impasse with respect to specific markers to be used for internalization studies. Those thought to be the most specific include serum albumin, SV40 virus, cholera toxin B subunit as a ligand for GM1 ganglioside, lactosyl ceramide, and GPIanchored proteins. Recently, it has become clear that all these markers can be internalized via other pathways (Nichols 2003; Lajoie and Nabi 2007; Parton and Simons 2007; Howes et al. 2010; Kumari et al. 2010). GPI-anchored proteins are primarily internalized via a cdc42-regulated clathrin and Cav-independent pathway (Sabharanjak et al. 2002). Only 1 in 10 SV40 particles may enter cells via caveolae, the virus being able to induce invaginations de novo (Damm et al. 2005; Ewers et al. 2010). Cholera toxin enters cells via multiple pathways (Kirkham et al. 2005; Lajoie et al. 2009b). In endothelial cells, serum albumin is supposedly internalized in caveolae via albondin/gp60, its low affinity receptor (Schnitzer and Oh 1994; Tiruppathi et al. 1997; Minshall et al. 2000). However, it has been clear for a long time that albumin can enter cells via other pathways chiefly as fluid phase (SiflingerBirnboim et al. 1991). Lastly, fluorescent lipid analogs such as lactosylceramide can be used for identification of the carriers with limited application for studies at the cell surface. Caveolae were proposed to mediate the exchange of macromolecules across the endothelial barrier by the process of transcytosis (reviewed in Stan 2002, 2005a), and recent in situ data seem to bolster the idea (Oh et al. 2007). However, with exception of a mild hypertriglyceridemia, the blood homeostasis in Cav1−/− mice is unperturbed, especially with respect to protein levels (Drab et al. 2001). Loss of caveolae in Cav1−/− (Schubert et al. 2002) or siRNA-mediated downregulation of Cav1 (Miyawaki-Shimizu et al. 2006) resulted in an increase in albumin clearance and accumulation into the tissues (except for aorta; Schubert et al. 2001). The increased vascular permeability to albumin might be the result of the opening of endothelial intercellular junctions. These studies are contradicted by another (Rosengren et al. 2004) showing no difference in transport in the Cav1-null and wild-type mice in a peritoneal dialysis model, arguing against a significant role for caveolar transcytosis in transport across the endothelium. These Cav1 deletion models seem to raise more questions than provide answers. It is not clear whether these findings are due to a lack of significant role of caveolae in transcytosis or to compensatory mechanisms such as increased permeability due to disregulation of endothelial nitric oxide (NO) synthase and NO production (Bauer et al. 2005; Lin et al. 2007; Murata et al. 2007) or the already documented dependence of albumin endocytosis on multiple pathways that may take over in Cav1−/− mice. If the opening of junctions compensates for the lack of caveolae, it is not clear how maintenance of the blood homeostasis is achieved.

Transducing Mechanical Stimuli Endothelial caveolae numbers are increased after exposure to chronic shear stress, and caveolae are required for the ensuing signaling (Rizzo et al. 1998, 2003). These early observations were bolstered by experiments in an arterial injury model in Cav1−/− mice,

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ABBREVIATIONS

51

which showed defects in chronic flow-dependent remodeling, and in acute flow-dependent dilation (Yu et al. 2006). Expression of Cav1 in endothelium rescues these defects (Yu et al. 2006). Cav1 and caveolae were involved in cyclic strain proliferative signaling in smooth muscle cells (Sedding et al. 2005). Caveolae were hypothesized that could play a role as tension regulators by flattening out in the membrane (Sens and Turner 2004).

INVOLVEMENT OF CAVEOLAE IN DISEASE There is good evidence for a role of caveolae in the pathogenesis of human disease (Williams and Lisanti 2004a). Endothelial Cav1 was shown to be involved in the progression of atherosclerosis (Frank et al. 2004; Fernandez-Hernando et al. 2009). There is a large body of data supporting a role for Cav1 in cancer (reviewed in Goetz et al. 2008). Cav1−/− cells show increased proliferation, (Koleske et al. 1995; Lee et al. 1998). Based on data from mouse models, Cav1 promotes tumorigenesis (Williams et al. 2005) although overexpression of Cav1 can inhibit tumor progression by decreasing tumor vessel permeability (Gratton et al. 2003). Cav can have different roles in different tumors (Sunaga et al. 2004). Additionally, Cav1 was involved in kidney stone disease (Cao et al. 2003), inflammatory diseases (Chidlow et al. 2009), and infections (Shin and Abraham 2001; Harris et al. 2002; Marsh and Helenius 2006; Mercer et al. 2010). The muscle-specific Cav isoform Cav3 is linked to a number of human muscle disorders including limb girdle muscular dystrophy and rippling muscle disease (Woodman et al. 2004).

FUTURE PERSPECTIVES Caveolae are membrane invaginations containing Cavs, cavins, gycosphingolipids, and cholesterol. Their biogenesis, trafficking, and maintenance are now beginning to be understood. Their physiological roles are just emerging with certain support for mechanosensing, maintenance of cell lipid balance, cell migration and adhesion, and endocytosis. However, their physiological roles are far from being understood. They are important as informed by their participation in panoply of human diseases such as atherosclerosis and cancer, to name a few. Questions remain as to the structure of caveolae. What is the mechanism of invagination formation and precise mechanism of budding? Proteomic approaches (Souto et al. 2003; Oh et al. 2007; Massey and Schnitzer 2009) have revealed a list of proteins that reside in caveolae. With more pieces of the puzzle known, it will be easier to assemble the final picture.

ACKNOWLEDGMENTS RVS was supported by the NIH grants HL65418, HL083249, and HL092085.

ABBREVIATIONS Cav EM

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caveolin electron microscopy

ER FD

endoplasmic reticulum fenestrated diaphragm

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GFP GPI MURC4 NO PEST PKC PRKCDBP PTRF

CAVEOLAE

green fluorescent protein glycosylphosphatidylinositol muscle-restricted coiled-coil nitric oxide proline, glutamic acid, serine, and threonine protein kinase C protein kinase C delta-binding protein Pol I and transcript release factor (also cavin-1)

SD stomatal diaphragm SDR or SDPR serum deprivation protein response (also cavin-2) SEM scanning electron microscopy SRBC sdr-related gene product that binds to c-kinase (also cavin-3) TEC transendothelial channel TEM transmission electron microscopy VSV vesicular stomatitis virus VVO vesiculo-vacuolar organelle

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CH A P T E R

4

LIPID RAFTS Leonard J. Foster

DEFINITION Lipid rafts are small (∼100 nm) cholesterol- and sphingomyelin-rich subdomains of the plasma membrane that function in numerous cellular processes. Simons and Ikonen (1997) proposed the concept, envisioning the ordered lipids like a raft in the large ocean of the plasma membrane. Rafts have been contentious for many years, starting very early on when the idea was one of the first to challenge the homogenous distribution of proteins and lipids predicted by the Singer and Nicolson (1972) fluid mosaic model of membrane bilayers. Simons et al. originally identified a role for rafts in lipid trafficking between and to the apical and basolateral surfaces in polarized epithelial cells; subsequently rafts have been implicated, to one degree or another, in a wide variety of cellular processes. Membrane lipids in general can be considered a solvent for membrane proteins, and the particular solvent properties of the lipids in rafts create an environment into which a specific set of membrane proteins are thought to partition; it is these proteins that convey many of the functional roles of rafts. They are best known for their role in signal transduction, where rafts seem to act as platforms for coordinating signaling events, but by virtue of their particular biophysical properties are also integral to maintaining nonuniform lipid distribution within cells, as in the case of polarized epithelial cells. Being located on the plasmalemma, rafts mediate interactions between the cell and various external processes: They are the attachment and/or entry point for numerous pathogens and the home for the diverse family of extracellular proteins anchored to the cell via a glycophosphatidylinositol (GPI) moiety.

HISTORICAL PERSPECTIVE The near-liquid crystal state of lipids in rafts makes them stable even in nonionic detergents, and it is this property, together with their relatively low density imparted by their high lipid content, that was initially used as the basis for biochemically enriching rafts (Brown and Rose 1992). In the raft enrichment procedure (Fig. 4.1) most commonly used today, suspension or adherent cells are typically extracted gently (i.e., with no mechanical force used) in a cold solution of 1% Triton X-100. This solution is then diluted 1:1 with 90% sucrose and layered in the bottom of an ultracentrifuge tube, with 35% and 5% sucrose solutions layered on top of that. After centrifugation at 150,000 to 200,000 relative centrifugal force for 18 hours, the rafts float to the interface between 35% and 5% sucrose, where they can be visualized as a faint, light scattering band, and harvested for further analysis (see Foster 2009 for detailed protocols). The fraction extracted from the interface Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Discontinuous

Continuous

5%

5%

35%

45% 45%

Figure 4.1. Conventional and high-resolution enrichment of detergent-resistant membranes (DRMs). DRMs are typically enriched using a discontinuous sucrose and other density gradients. In this way, all detergent-insoluble cellular components with a density of between 5% and 35% sucrose will appear to comigrate. Since different detergent-insoluble components could conceivably have more subtle differences in density, a continuous or pseudo-continuous density gradient can be used to partially or completely resolve some of the components of DRMs.

between 35% and 5% sucrose in this way, however, is more accurately referred to as a detergent-resistant membrane (DRM) fraction since it is most certainly contaminated by nonraft material (see below). Thus, throughout this chapter, we use the term DRM to refer to the biochemical fraction and the term “raft” to refer to the actual liquid-ordered domain in the plasma membrane; to reinforce the point, DRMs are not equivalent to rafts, rather rafts are a subset of all DRMs. Although Brown and Rose (1992) used Triton X-100 initially—and this reagent is by far the most popular for biochemically enriching rafts—rafts are also stable in other nonionic detergents (Chamberlain 2004). The complication, however, is that the composition of DRMs isolated by different detergents can vary widely and that it is far from clear which detergent, if any, extracts “true” rafts (for a review of raft proteomics, see Foster and Chan 2007). Quantitative proteomic analysis of DRMs isolated with Triton X-100 versus Brij-96, the second most popular detergent for these experiments, reveals some striking differences in many classes of proteins (Blonder et al. 2006). Importantly, neither detergent was effective in enriching all accepted markers of rafts, and the nonraft proteins in the DRM preparations were also not consistent. Thus, the choice of detergent can have a significant impact on the results of raft biochemistry experiments. Rafts have faced a barrage of criticism, especially since the initial claims (Brown and Rose 1992) that they could be enriched using detergents. For many biophysicists, the fact that detergents were required to enrich rafts suggested that the rafts were forming in the presence of the detergent and that they did not actually exist prior to the introduction of the detergent. This doubt seems to have been finally put to rest (see “Raft Biophysics”), but the whole argument was always a bit of a Catch-22 for raft biochemistry since it is difficult to fully remove all the nonraft lipids and proteins without using detergents because they are, by definition, insoluble in water. The nonraft lipids could be solubilized in organic solvents; however, such reagents are rarely useful when working with proteins, and so other approaches for enriching rafts were explored. The main detergent-free approach has involved physical disruption of the cells, usually by sonication, followed by flotation on a sucrose or other density gradients similar to that used with detergents (Fig. 4.1; Smart et al. 1995). Besides mechanical disruption, another point of distinction with the conventional method has been the use of high pH (∼11.5) to strip off nonintegral membrane

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proteins (Song et al. 1996). These methods were originally developed to enrich caveolae (discussed elsewhere in this book), but the fractions isolated in this way contain many of the same proteins found in DRMs (Foster et al. 2003), perpetuating the controversies surrounding the distinction between rafts and caveolae. Perhaps most importantly, detergentfree methods for enriching rafts yield a membrane fraction that contains far more known contaminants than those enriched using detergents (Foster et al. 2003). Thus, since rafts have been demonstrated to be not just an artifact of detergent extraction, the use of detergent-free methods seems no longer justified given the levels of contaminants in these preparations.

Protein Composition A great many proteins have been localized to rafts with one technique or another, but a small set of proteins (Table 4.1) is acknowledged as reliable markers of lipid rafts based on a plethora of supporting data. More widely, proteins have been localized to rafts based on microscopic colocalization or equivalent buoyant density with one or more of the markers listed in Table 4.1. Until more recently, when fluorescence resonance energy transfer (FRET) and sub-diffraction imaging methods became more widespread, claims of a protein being localized to lipid rafts based on immunofluorescence colocalization must be considered dubious since rafts are typically well below the conventional resolution of visible light microscopes (see Chapter 5 for discussion of fluorescence analysis of lipid rafts). The other approach, more widely applied, has been to probe the distribution of a candidate protein in the sucrose density gradient described in Figure 4.1 versus the distribution of a known raft protein. If a protein is enriched in the 5/35% interface of the density gradient, then it can at least be claimed to enrich with rafts, although as we and others have shown, this can be misleading (Foster et al. 2003; Kierszniowska et al. 2009; Zheng et al. 2009). The final piece of evidence that is usually required to claim a protein is in rafts is a demonstration that its colocalization and/or buoyant density migration with a raft marker can be disrupted using a pharmacological antagonist of cholesterol, typically methyl β-cyclodextrin (mβCD), or one of the statin family of compounds (Keller and Simons 1998). The physical integrity of rafts has been shown to be completely dependent on cholesterol and so disruption of membrane cholesterol, either via blocking its biosynthesis (e.g., with lovastatin) or via its chelation (e.g., with mβCD), breaks up rafts and allows the other raft lipids and proteins to diffuse away and become soluble in ionic

TABLE 4.1.

Some of the Classical Markers of Lipid Rafts

Protein Flotillina Alkaline phosphatase 5′-nucleotidase Lyn Yes Complement delay accelerating factor T-cell receptor B-cell receptor Lck

Class

Anchor

Structural protein Extracellular enzyme Extracellular enzyme Tyrosine kinase Tyrosine kinase Complement receptor Receptor Receptor Tyrosine kinase

Transmembrane domain GPI GPI Palmitate and myristate Myristate GPI Transmembrane Transmembrane Palmitate and myristate

a

Also a caveolae protein.

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detergents. Thus, by microscopy, the raft marker and protein of interest should no longer colocalize after mβCD treatment, and by buoyant density both the raft marker and protein of interest should not migrate up to the 5/35% interface (Fig. 4.1). Using these lines of evidence, studies focusing on one or a few proteins have demonstrated raft localization for an estimated 200 proteins. While these studies would have been skewed toward those proteins that people thought to look for, proteomics, on the other hand, provides an excellent means to probe the composition of lipid rafts in a relatively unbiased manner—only “relatively unbiased” because current proteomic technology does not allow all the proteins in a sample to be identified and so there is a bias toward the more abundant species and those that are more hydrophilic. Nonetheless, proteomics does allow us to query the static composition of rafts and to test how the composition changes in response to stimuli. Comprehensive summaries of lipid raft proteomics literature can be found elsewhere (Foster and Chan 2007), but the major families of raftlocalized proteins are discussed below. Several other categories of proteins have been reported to be found in DRMs and therefore claimed to be in rafts (Bae et al. 2004; Man et al. 2005; McMahon et al. 2006; Kim et al. 2010), but most of these have been debunked as contaminants since their presence in DRMs is not dependent on cholesterol. The following classes of proteins are either in rafts or have been implicated in rafts. GPI-Anchored Proteins This is a functionally diverse family of proteins whose only common feature is the presence of an O- or N-linked GPI moiety that is added in the endoplasmic reticulum. There are 15 such proteins known in yeast and at least 45 in humans, comprising enzymes, receptors, and transport-type proteins, and where there is a human ortholog of a yeast GPI-anchored protein, the human protein is also known to have a GPI anchor. After they are fully processed, GPI-anchored proteins localize primarily to the exoplasmic leaflet of the plasma membrane, where many of them are targeted to rafts. They are, of course, not absolutely exclusive to the plasma membrane as they must pass through biosynthetic organelles, such as the endoplasmic reticulum and Golgi, to get to the plasma membrane. Once on the plasma membrane, GPI-anchored proteins are internalized into the endosomal system through various mechanisms; by and large, however, their primary localization is thought to be the plasma membrane. GPI-anchored proteins can be released from the membrane by phospholipases, and this approach has provided a mechanism for identifying raft-localized GPI-anchored proteins from a variety of species (Sherrier et al. 1999; Fivaz et al. 2000; Borner et al. 2003; Elortza et al. 2003; Elortza et al. 2006). In humans, some of the common raft-associated GPI-anchored proteins include CD59, Thy-1, 5′-nucleotidase, alkaline phosphatase, folate receptor, bone marrow stromal cell antigen, and decay acceleration factor. Signaling Proteins Rafts gained a great degree of popularity in the 1990s in part due to the claims that they were organizing centers for signal transduction at a time when there was an incredibly intense interest in signal transduction in general and phosphorylation cascades in particular. Interestingly, the evidence suggesting that rafts were more involved in signaling than other regions of the plasma membrane were largely anecdotal until proteomics established in an unbiased way that signal transduction proteins were enriched on rafts relative to other membranes (Foster et al. 2003). By comparing the enrichment of signaling proteins moving from a total membrane preparation through to DRMs and then to only those DRM proteins that displayed a sensitivity to MβCD, it was clear that signaling proteins are indeed more abundant on rafts than other membranes (Foster et al. 2003). While there are

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sporadic reports of many signaling pathways acting through rafts (Simons and Toomre 2000), we highlight here some of the best understood: 1. T-cell receptor (TCR) pathway. Ligation and dimerization of T-cell receptors induce raft clustering, activating Lck and Fyn kinases. This in turn leads to recruitment of Zap-70, and then GPI-anchored proteins and other signaling adaptors amplify the signal (many contributing studies—for reviews, see Janes et al. 2000; Langlet et al. 2000). 2. High-affinity immunoglobulin E (IgE) receptor pathway. Aggregation of receptors cause their translocation to lipid rafts, leading to activation of Lyn (Field et al. 1995; Sheets et al. 1999) and similar events to TCR. 3. Glial cell-derived neurotrophic factor (GDNF). Ligation of the GPI-anchored GDNF receptor-α recruits the RET tyrosine kinase to rafts (Tansey et al. 2000). 4. Hedgehog. The Hedgehog protein is posttranslationally modified with a cholesterol moiety on one end (Porter et al. 1996) and a palmitate on the other (Pepinsky et al. 1998), likely helping to localize it to lipid rafts (Rietveld et al. 1999). Binding of Hedgehog to the Patched receptor protein releases the inhibitory effects of Smoothened, leading to a signaling cascade that activates transcription. Acyl-Anchored Proteins Numerous acylated, intracellular proteins are localized to lipid rafts, particularly members of the Src family of protein tyrosine kinases such as those listed in the various signaling pathways above. With few exceptions, these acylations are saturated, unbranched lipids that can easily partition into the highly ordered lipid structure of a raft. Even Ras, perhaps the quintessential signaling protein, is found in rafts in some cases: Palmitoylation of the C-terminal CAAX motif can target it to plasma membrane rafts where it can actively transduce a signal to Raf (Roy et al. 1999). Other small GTPases in the Ras superfamily also seem to localize to rafts, but this does not apply generally to all members (e.g., Rabs 1, 2, 3, and 10 are all in DRMs, but only isoforms 2 and 3 are mβCD sensitive [Foster et al. 2003], although all Rabs are thought to be doubly geranylgeranylated). Thus, acylation per se does not determine that a protein will segregate into lipid rafts; even specific types of acylation do not guarantee lipid raft association, although, in general, saturated, unbranched lipidations will tend to localize to rafts more often than other types. Mitochondrial Proteins Prior to the first proteomic analyses of DRMs (von Haller et al. 2001; Nebl et al. 2002; Bini et al. 2003; Foster et al. 2003; Ledesma et al. 2003; Li et al. 2003; von Haller et al. 2003a, b) there was no indication that mitochondrial proteins might reside in lipid rafts, but in all DRM proteomic studies to date, the voltage-dependent anion selective channels and components of the F1/F0-ATPase have been detected. Since there was no apparent link with the other known functions of lipid rafts, the cell biology community greeted these reports with some skepticism (Magee and Parmryd 2003). As mentioned previously, one of the hallmarks of lipid raft proteins is their sensitivity to cholesterol disruption, but two early quantitative DRM proteomic studies reported conflicting results regarding the sensitivity of mitochondrial proteins in DRMs to mβCD (Bini et al. 2003; Foster et al. 2003). Other than cell type differences and some other minor variations in experimental details, the main distinction between the two studies was the point at which the amounts of protein in the untreated and mβCD-treated samples were normalized, with one study showing protein is normalized before DRM isolation (Foster et al. 2003) and the other study, after

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DRM isolation (Bini et al. 2003). Normalizing protein after pharmacological treatment is certainly justified if the agents (i.e., mβCD) are not expected to drastically alter the amount of protein in the biochemical fraction. However, in the case of mβCD, the effect on the DRM fraction is profound: fully 50+% of DRM proteins are partially or fully depleted after mβCD treatment, making it critical to equalize the amounts of protein in each treatment before the DRM enrichment. When protein amounts are normalized prior to treatment, there is no effect of mβCD on mitochondrial proteins (Foster et al. 2003; Zheng et al. 2009). Intriguingly, however, other studies have linked mitochondrial proteins to lipid rafts as well. Kim et al. reported that the F1/F0 ATPase (the mitochondrial complex) can be found on the cell surface and is even reactive to impermeant biotinylation reagents (Kim et al. 2010). More recently, they have also demonstrated that mitochondrial oxidative phosphorylation complexes can be surface exposed and that mβCD can disrupt the production of extracellular reactive oxygen species by these complexes (Kim et al. 2010). The very low resolution of the discontinuous density gradient used for most DRM preparations may also contribute to mislocalization of proteins, particularly mitochondrial proteins. In the normal method, all membranes or other complexes with densities between 35% and 5% sucrose will appear to comigrate with lipid raft proteins. To get around this limitation, we used a quantitative proteomic approach called peptide correlation profiling– stable isotope labeling by amino acids in cell culture (PCP–SILAC) to measure how closely mitochondrial proteins and lipid raft proteins comigrate on a higher resolution, pseudo-continuous sucrose density gradient (Fig. 4.1; Zheng et al. 2009). Under these conditions, we were able to clearly resolve the mitochondrial proteins from the classical lipid raft markers, reinforcing our previous claims that mitochondrial proteins cannot be localized to rafts and must instead be contaminants of the normal DRM preparation.

Lipid Composition As mentioned, cholesterol and sphingomyelin are two of the principal components of lipid rafts. Indeed, together with certain glycosphingolipids and saturated glycerophospholipids, these lipids can form a liquid-ordered state all on their own (Brown and London 1998). Beyond these broad general categories, there have been no detailed lipidomic descriptions of rafts, as of yet, presumably because of the problems involved in obtaining “pure” raft preparations through the conventional approaches (see above). However, using immunoaffinity purification, Zech et al. (2009) were able to generate highly purified membrane domains containing the TCR and then use these to investigate how the TCR-bound domains change in response to stimulation. Not surprisingly, they found that the TCR membrane domains were highly enriched in cholesterol and sphingomyelin, relative to transferrin receptor membrane domains. However, they also noted an overabundance of phosphatidylserine and phosphatidic acid in TCR membrane domains, in contrast to an underrepresentation of phosphatidylcholine and phosphatidylinositol.

RAFT BIOPHYSICS: DO THEY REALLY EXIST? A subset of the published raft studies using biophysical methods has shown that the presence of Triton can force the formation of liquid-ordered domains in synthetic and biological membranes (Heerklotz 2002; Heerklotz et al. 2003; Heffer-Lauc et al. 2005), suggesting that many of the results based on DRM preparations may be artifacts. Countering this, however, is a series of biochemical, biophysical, and cell biological experiments suggest-

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ing that sphingolipid–cholesterol rafts do exist in biological membranes before the addition of a detergent and that these domains are effectively extracted by detergents (but not that detergent extracts are composed purely of these domains). Studies employing cross-linking (Friedrichson and Kurzchalia 1998), FRET (Varma and Mayor 1998), biophysical studies in model membranes (Ahmed et al. 1997; Garner et al. 2008), and single-particle measurements of diffusion in conventional light microscopy (Pralle et al. 2000) and sub-diffraction imaging using fluorescence photoactivation localization microscopy (Hess et al. 2007) all indicate that rafts exist in biological membranes and that the detergent-based methods for extracting them are extracting the expected structures. The preponderance of evidence now strongly favors the existence of rafts and justifies most of the work that has been carried out on them over the past two or more decades (see also Chapters 1 and 5).

RAFTS IN HOST–PATHOGEN INTERACTIONS A great many parasites, viruses, and bacteria have evolved the ability to invade host cells and co-opt the host machinery to survive, replicate, and cause disease. Most of these pathogens rely on the host machinery to gain entry as well, and it is here that lipid rafts and/or caveolae are frequently engaged. Disruption of rafts through the use of mβCD, filipin, or nystatin is known to disrupt internalization of numerous bacterial pathogens, including Campylobacter jejuni (Wooldridge et al. 1996), Brucella abortus (Watarai et al. 2002), Escherichia coli (Duncan et al. 2004), and Mycobacterium kansasii (Watarai et al. 2002). For other pathogens, rafts seem to mediate signaling pathways that the bacteria interfere with in order to avoid host defenses, as is the case with Pseudomonas aeruginosa (Zhang et al. 2005). Rafts in different guises are involved in classical clathrin-mediated endocytosis (Stoddart et al. 2002; Abrami et al. 2003), providing a mechanism for some pathogens (e.g., viruses, some bacteria) and virulence factors or toxins to enter into host cells. For example, in the case of Bacillus anthracis, the anthrax toxin receptor is located in lipid rafts and is internalized on ligation with the anthrax toxin (Abrami et al. 2003). Although not yet widely recognized, Salmonella bacteria also employ host lipid rafts, if not for entry then at least for the initial adherence step (Lim et al. 2010). Salmonella are among those pathogens that can induce their own internalization via a phagocytosis-like mechanism (Brumell et al. 1999); although the functional link between rafts and the process of phagocytosis is not yet established, there is increasing evidence that rafts are involved in some way as the composition of rafts changes during phagocytosis (personal observations) and the raft components of phagosomes change with phagosome maturation (Dermine et al. 2001; Rogers and Foster 2007; see Chapter 10 for a discussion of raft domains in phagosomes). The examples mentioned above likely only represent those host– pathogen systems where investigators have considered the role of rafts or where the system is a well-established model (Zaas et al. 2005); it is likely that rafts will be found to be critical to the ability of many other pathogens to cause disease as well.

FUTURE PERSPECTIVES Lipid rafts are now assured a place in our model of the cell and of membrane structure. The biophysical arguments that rafts are simply an artifact of detergent usage died out two or three years ago, and there is now widespread agreement that they are a real function of phase separation between different classes of lipids, together with structural assistance provided by proteins. Now that the reality versus artifact discussion has concluded, the

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field can finally move on and focus on the role of rafts in cell biology. Clearly, rafts play a fundamental role in many signaling processes, as well as in intracellular trafficking. As domains/organelles go, lipid rafts are an excellent subject for “omic” approaches since their composition is relatively simple; with quantitative lipidomic and/or proteomic approaches that can control well for contaminants or functional relevance, studies of how the protein and lipid composition changes in response to various stimuli will surely open up new avenues for investigation. Partnering these discovery approaches with some of the amazing new imaging methods (Hess et al. 2007) currently available could make rafts the most exciting field in cell biology again.

ABBREVIATIONS DRM FRET GDNF GPI

detergent-resistant membrane fluorescence resonance energy transfer glial cell-derived neurotrophic factor glycophosphatidylinositol

mβCD methyl β-cyclodextrin PCP–SILAC peptide correlation profiling–stable isotope labeling by amino acids in cell culture TCR T-cell receptor

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Stoddart A, Dykstra ML, Brown BK, Song W, Pierce SK, Brodsky FM. 2002. Lipid rafts unite signaling cascades with clathrin to regulate BCR internalization. Immunity 17:451–62. Tansey MG, Baloh RH, Milbrandt J, Johnson EM, Jr. 2000. GFRalpha-mediated localization of RET to lipid rafts is required for effective downstream signaling, differentiation, and neuronal survival. Neuron 25:611–23. Varma R, Mayor S. 1998. GPI-anchored proteins are organized in submicron domains at the cell surface. Nature 394:798–801. von Haller PD, Donohoe S, Goodlett DR, Aebersold R, Watts JD. 2001. Mass spectrometric characterization of proteins extracted from Jurkat T cell detergent-resistant membrane domains. Proteomics 1:1010–21. von Haller PD, Yi E, Donohoe S, Vaughn K, Keller A, Nesvizhskii AI, Eng J, Li XJ, Goodlett DR, Aebersold R, Watts JD. 2003a. The application of new software tools to quantitative protein profiling via ICAT and tandem mass spectrometry: I. Statistically annotated data sets for peptide sequences and proteins identified via the application of ICAT and tandem mass spectrometry to proteins co-purifying with T cell lipid rafts. Mol Cell Proteomics 2:425–27. von Haller PD, Yi E, Donohoe S, Vaughn K, Keller A, Nesvizhskii AI, Eng J, Li XJ, Goodlett DR, Aebersold R, Watts JD. 2003b. The application of new software tools to quantitative protein profiling via ICAT

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MODELING MEMBRANE DOMAINS Daniel Coombs Raibatak Das Jennifer S. Morrison

DEFINITION Cell membranes are known to be heterogeneous structures, featuring distinct domains on the scale of tens to hundreds of nanometers. Membrane domains are characterized by the enrichment or exclusion of membrane constituents and proteins, and are believed to play important roles in modulating cell signaling through surface receptors. Fluorescence microscopic techniques, including fluorescence recovery after photobleaching (FRAP), fluorescence resonance energy transfer (FRET), single-particle tracking (SPT), and fluorescence correlation spectroscopy (FCS), have played important roles in shaping our understanding of membrane domains and their dynamics. These techniques have been used to study the membrane domains known as lipid rafts, which were originally defined as detergent-resistant regions that are enriched in glycosphingolipids (see also Chapter 4). We comment on our evolving understanding of these domains and show how careful data analysis and mathematical modeling naturally complement fluorescence microscopy in this context.

HISTORICAL PERSPECTIVE The Singer–Nicolson fluid mosaic model for the membrane of a cell states that the membrane of a cell can be thought of as a two-dimensional fluid, within which membrane components and transmembrane proteins diffuse freely (Singer and Nicolson 1972). However, ongoing work indicates that a better picture of the membrane may be one consisting of dynamically rearranging structures, or domains, on the scale of 50–500 nm in size. Membrane domains can be characterized according to which molecules are enriched in the region (e.g., the canonical lipid raft, enriched in cholesterol and sphingolipids), and they may influence the mobility of membrane components, lipids, and proteins by confining or hindering their motion. It is widely believed that the choreography of intracellular signaling following surface receptor engagement is affected by localization of key signaling proteins. Several well-characterized examples are found in the immunological context: FcεRI receptor signaling on mast cells following cross-linking by immunoglobulin E (IgE) and antigen, B-cell receptor (BCR) signaling following receptor cross-linking by antigen, and T-cell receptor (TCR) signaling following peptide–major histocompatibility complex (MHC) binding at the immunological synapse. Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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The definition of membrane rafts has evolved considerably in the last decade concurrently with advances in high-resolution imaging that have allowed cellular membranes to be probed with increasing spatiotemporal resolution. Until the early 2000s, rafts were defined primarily on the basis of their biochemical properties (detergent resistance, glycosphingolipid enrichment, and the preferential partitioning of glycosylphosphatidylinositol [GPI]-anchored proteins) (see Chapter 4 for a discussion of detergent-resistant raft domains) and were thought to be a distinct phase in the plasma membrane such as was observed in model membrane systems with well-defined lipid composition (Edidin 2003). With rapid advances in imaging techniques (reviewed in Lagerholm et al. 2005), and the emergence of suboptical resolution microscopy (Betzig et al. 2006), it became obvious that membrane heterogeneities occur at many length scales and have different characteristic persistence times (Hancock 2006; Hess et al. 2007). In this respect, cellular membranes differ substantially from model membranes that exhibit persistent and large-scale phase separation. As a result, a more recent definition of membrane rafts is short-lived spontaneous aggregates of variable size in resting cells, which may coalesce around activated receptors and could aggregate to form larger rafts (Pike 2006). A greater appreciation has also emerged for the role played by the membrane-associated proteins in organizing the lipid environment around them (Jacobson et al. 2007; Lingwood and Simons 2010). The recent discovery of receptor microclusters in T and B cells also supports aspects of this hypothesis, while raising new questions about the formation and function of these entities, and the role played by the actin cytoskeleton in regulating membrane organization (Bunnell 2010). The purpose of this chapter is to outline how fluorescence microscopic techniques can be used to study the dynamics of membrane domain organization, and to highlight how careful mathematical modeling and data fitting can give important insights into membrane dynamics. We will not discuss other important tools such as electron microscopy (EM) and atomic force microscopy (AFM). EM in particular has been used alongside fluorescence microscopic approaches to characterize the distribution of surface receptors and surface-associated signaling molecules with great success (e.g., Lillemeier et al. 2010). However, EM cannot be used to track dynamic reorganization of domains in response to stimuli, since cells must be fixed and the membranes removed from the cells before imaging. For this reason, EM will probably continue to be used in tandem with other techniques for studying dynamics of membrane domains. In contrast, AFM can be used with live cells and has great promise to provide resolution comparable with EM, with reasonable time resolution (Lee et al. 2007). Wider use of AFM in studies of membrane organization and signaling can be expected in the future.

THEORETICAL BASICS Fluorescence microscopic techniques for studying membrane domains are intimately connected with the theory of diffusion of molecules in the membrane. Brownian diffusion is the simplest form of random (thermal) motion and assumes that a molecule moves randomly and without any memory of its previous motion. This means that a purely Brownian particle behaves identically at all times and in all parts of the membrane. For Brownian diffusion, the expected mean square displacement (MSD) of a molecule is a linear function of time: r 2 (t ) = 4 Dt ,

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where D is defined as the diffusion coefficient of the particle, and the angle brackets indicate averaging over many particle trajectories. According to the Einstein–Stokes relationship, D is inversely proportional to the particle size and the effective viscosity of the membrane (Einstein and Fürth 1956). The existence of membrane domains breaks the assumption that the molecular motion is spatiotemporally homogeneous, because (1) the particle mobility may be altered within a domain and/or (2) domains may be defined by fence-like structures that do not permit the particle to pass. As a result, the motion is “subdiffusive” and the expected MSD grows less than linearly over time: r 2 (t ) ~ 4 Dt α , where the exponent α is less than 1. Intuitively, this can be understood by imagining a particle diffusing within a fixed cage. Over short times, the walls of the cage do not significantly affect the motion. However, over longer times, the particle’s overall displacement cannot grow beyond the size of the cage. This means that long-term diffusion is slower than short-term diffusion, or equivalently, the diffusivity of the particle can be thought of as time dependent (Saxton 2001). Similarly, a particle moving in and between cages with partially porous boundaries will show reduced long-term mobility compared with short-term mobility. At the other extreme, particles might be described as “superdiffusive” when the exponent α is greater than 1. This would be the situation for a particle undergoing directed motion (e.g., a surface protein that is actively transported along a filament of the cell cytoskeleton). If a particle moves with velocity v, superimposed on Brownian diffusion, then its MSD is given as r 2 (t ) = 4 Dt + v 2 t 2 . The term involving t2 is the square of the distance traveled due to directed motion. At long times, the MSD is dominated by this term and thus grows faster than linearly. Data from each of the microscopic techniques we describe here can be analyzed based on these basic ideas of diffusion processes. In the case of SPT, statistical comparisons can be made directly between labeled protein tracks and the diffusion model. On the other hand, when fluorescent proteins are labeled en masse (e.g., in FRAP experiments, see below), we have to use a diffusion process averaged over many particles. In the case of Brownian diffusion, this leads to a relatively simple partial differential equation that can be solved analytically or numerically (Crank 1980). For sub- or superdiffusive processes, the averaging process can be much more complex because the motion of the particle depends on its previous history of movements. Mathematical research in this area is ongoing but it is not clear how to confidently fit experimental data (e.g., from fluorescence recovery) to subdiffusive models and thus detect membrane domains.

FRAP FRAP is routinely used to study the mobility of cell-surface proteins. In this method, a fluorescent probe is conjugated to the surface population of the protein of interest. A short period of high laser intensity is used to bleach the probe in a specific region, reducing the fluorescence in that region to background levels. Motion of unbleached proteins into the bleached region causes fluorescence in the bleached region to recover. The level of fluorescence in the bleached region is monitored over time, generating a saturating recovery curve. The shape of the recovery curve is governed by the motion of the labeled protein

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and the precise details of the experimental protocol. FRAP recovery curves can be fit to mathematical models, yielding estimates of the diffusion coefficient and the fraction of labeled proteins that are mobile on the cell surface. The book chapter by Rabut and Ellenberg provides a detailed description of the technique, taking into account practical considerations for FRAP experiments (Rabut and Ellenberg 2004). The question of how FRAP recovery curves would be affected by subdiffusive particle behavior was addressed in Saxton (2001). In this study, FRAP recovery curves were simulated for three different mathematical models that yield subdiffusive motion. The main result was that, for a single experiment, determining the nature of the molecular dynamics (diffusive, subdiffusive, confined, etc.) is a very difficult task. This is shown in that paper by fitting simulated data with both anomalous and standard diffusion models and obtaining good fits in both cases. More recently, FRAP experiments have been used to study transient binding of a labeled mobile protein to a slow-moving or stationary binding partner (Sprague and McNally 2005). This is achieved by fitting the recovery curves to a reaction–diffusion model (rather than a simple diffusion model) that is parameterized by forward and reverse rate constants for a binding interaction and the diffusion coefficients of the free protein and the bound complex. Qualitatively, transient binding to an immobile partner slows recovery and changes the shape of the recovery curve. We applied this approach to study the mobility of TCRs on the surfaces of live T cells (Dushek et al. 2008). We found that the recovery of fluorescently labeled TCR was best fit by a model of TCR diffusion and transient binding to an immobile partner. The binding hindrance was removed upon treating the cells with cytoskeletal inhibitors, leading us to hypothesize that the cortical actin cytoskeleton of the cell provided (unspecified) transient binding sites. However, in other situations, this method might detect transient confinement zones within the cell membrane. In that situation, the effective on and off rates that are estimated from the data would be interpreted as rates of arriving and leaving the confinement zones. A complete comparison of the characteristic FRAP recovery curves for anomalous motion due to confinement in membrane domains versus transient binding to immobile partners would be informative. If such a study were performed, it might lead to the development of diagnostics as to which models are suitable to interpret deviations from purely diffusive behavior. However, as shown by Saxton, recovering molecular-scale information from a (relatively coarse) FRAP experiment is a near-impossible task. For this reason, we move on to discussion of other techniques that can offer greater precision.

FRET FRET microscopy is a sensitive tool to detect the close proximity between two fluorophores (Selvin 2000; Lakowicz 2006). The two fluorophores may be conjugated to the different parts of the same biomolecule, so that the FRET signal reveals conformational changes in the molecule, or they may be on two different molecules, in which case it reveals close proximity, and thus potential association. In typical FRET experiments, a donor–acceptor pair of fluorophores is chosen in a way that the emission spectrum of the donor overlaps with the excitation spectrum of the acceptor. When the donor is excited by incident light tuned to its excitation wavelength, it can transfer energy to a nearby acceptor molecule, leading to a decrease in the donor emission, while increasing the acceptor emission. The efficiency of this energy transfer process is typically measured as donor quenching in the presence of the acceptor: E = 1 − FDA / FD ,

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where FD and FDA are the donor fluorescence intensities in the absence and the presence of an acceptor, respectively. This efficiency can also be calculated by measuring the change in the lifetime of a donor in the presence of the acceptor. FRET efficiency is strongly dependent on the distance between the two fluorophores and their relative orientations, and is given as E = 1 /(1 + ( R / R0 )6 ), where R is the separation between the molecules. When R = R0, the Forster radius, the energy transfer is 50% efficient (Forster 1959). The Forster radii for a number of FRET pairs are tabulated (Lakowicz 2006) and are typically in the range of 20–60 Å. Because of the strong sixth power dependence on fluorophore separation, FRET efficiency drops off precipitously for separations exceeding R0, and in practice, FRET is only useful for measuring separations in the range of 10–100 Å.

What Do We Know about Membrane Organization from FRET Studies? A number of studies have used fluorophore density-dependent FRET measurements of GPI-anchored proteins and glycosphingolipids to examine their organization into microdomains variously called lipid rafts or membrane rafts. The principles underlying these studies are the following: For proteins (or lipids) that are organized into spatial clusters (raft) with fixed densities of component molecules, FRET efficiency is independent of the overall surface density of a raft component. In contrast, for randomly distributed molecules, FRET efficiency increases with increasing surface density, as the molecules are packed more closely. Initial studies (Kenworthy and Edidin 1998; Varma and Mayor 1998) led to conflicting reports and were followed up with a more careful examination of the membrane distributions of different GPI-anchored proteins and glycosphingolipids (Kenworthy et al. 2000; Sharma et al. 2004). The results from these studies are now reconciled within the raft hypothesis, which posits that membrane rafts are small, heterogeneous, and dynamic structures that span tens of nanometers, and that may be stabilized to form larger structures during cell signaling (Lagerholm et al. 2005; Pike 2006). Moreover, only a small fraction (5–20%) of the total amount of putative raft component is presumed to be associated with these dynamic rafts in resting cells. FRET measurements using fluorescent lipid probes also support the existence of nanometer-scale heterogeneities in the plasma membrane of mast cells (Sengupta et al. 2007).

Consequences for Cell Signaling Lipid rafts are postulated to segregate components of signal transduction. The generally accepted mechanism for the role of membrane rafts in B cells and mast cells proposes that, in resting cells, monomeric BCRs or the high-affinity IgE receptors, FcεRI, are largely excluded from lipid rafts, while the Src family kinase Lyn is preferentially distributed in rafts. Membrane phosphatases such as CD45 are also excluded from rafts, and serve to maintain the receptors in a predominantly dephosphorylated state in resting cells. Upon binding to a multivalent antigen, cross-linked receptors redistribute into Lyn-enriched lipid rafts, where they are susceptible to Lyn phosphorylation, while being shielded from phosphatases. Other signaling components such as CD19, Syk, and LAT are also enriched in rafts, either due to transient associations with the phosphorylated receptors or by virtue of their lipid anchors (for LAT), and thus rafts provide a spatial platform for downstream signal propagation (reviewed in Pizzo and Viola 2004). Some of the most detailed direct observations of these events have emerged in a series of FRET microscopy studies from the lab of Susan Pierce (reviewed in Tolar et al.

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2008). Using an elegant system of reconstituted BCRs in J558L B cells containing different cyan fluorescent protein (CFP)- or yellow fluorescent protein (YFP)-labeled BCR components, Pierce and colleagues established the stoichiometry of the BCR (1 mIg : 1 Igα-Igβ heterodimer) and demonstrated (1) antigen-induced receptor aggregation, (2) Lyndependent conformational change in the cytosolic domains of the BCR, and (3) transient associations between cross-linked receptor and Lyn that peak within 10 seconds of antigenstimulation and dissipate within minutes (Tolar et al. 2005; Sohn et al. 2006). Subsequently, these observations were recapitulated for B-cell activation with a surface-bound antigen, presented through a supported bilayer, replicating a B-cell immunological synapse (Sohn et al. 2008). Notably, high-resolution imaging of the B-cellsupported bilayer interface using total internal reflection fluorescence (TIRF) microscopy revealed the formation and migration of signaling competent BCR microclusters at the immune synapse (Depoil et al. 2008). Receptor microclusters have also been observed during the formation of a T-cell immune synapse at the interface with an antigen-containing bilayer (Bunnell et al. 2002; Campi et al. 2005). These microclusters form spontaneously at the periphery of a synapse within seconds of contact with the antigen, contain phosphorylated Src kinases, and recruit downstream signaling molecules such as Syk/Zap-70 and SLP-76 (Yokosuka et al. 2005). The molecular basis of microcluster formation and their role in immune signal transduction is still poorly understood.

SPT In contrast to FRAP and FRET, techniques that both monitor a population of molecules and thus provide an ensemble measurement, SPT is a powerful single-molecule technique that yields direct observation of molecular motion (see also Chapter 1). The protein of interest is labeled with an immunofluorescent probe or an antibody-coated bead, at a sufficiently low concentration so as to allow the visualization of single molecules or diffractionlimited clusters. Using high-speed video microscopy, the particle’s position is then tracked over time to obtain its trajectory, as the experimental output. Typical SPT experiments have a spatial resolution of tens of nanometers and a time resolution of tens of milliseconds (although experiments with microsecond resolution have been performed; Fujiwara et al. 2002; Murase et al. 2004; Umemura et al. 2008). It is also common to treat cells with pharmacological agents (such as cytochalasin D to disrupt the actin cytoskeleton) and examine changes in particle trajectories. In principle, particle trajectories contain information about the nature of molecular motion, signatures of spatiotemporal heterogeneity, and the presence of multiple subpopulations, though it is a nontrivial task to extract this information in a statistically reliable fashion.

Analysis of Particle Tracks In classical SPT analysis, particle trajectories are classified on the basis of their MSD versus time plot (see Theoretical Basics). The MSD as a function of time is calculated from the individual displacements (Qian et al. 1991; Kusumi et al. 1993), and tracks are then sorted into various modes of motion based on fitting their MSD curve to the equation MSD(t ) = 4 Dt α (see Theoretical Basics; Anderson et al. 1992; Wilson et al. 1996; Saxton and Jacobson 1997). Other MSD-based parameters, such as the relative deviation from Brownian diffusion (Kusumi et al. 1993) and the radius of gyration tensor (Rudnick and Gaspari 1987; Saxton 1993), have also been used to categorize particle trajectories.

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A series of theoretical studies by Saxton has examined the origin of anomalous diffusion due to confinement, the presence of obstacles, or binding, and proposed algorithms for detecting these through MSD measurements (Saxton 1993, 1994, 1995, 1996). An analytical treatment also exists for diffusion through an infinite array of impermeable barriers, relating the macroscopic and microscopic diffusion coefficients to the average compartment size and the average residency time within a compartment (Powles et al. 1992). The results from this study were used to test a hop diffusion model for membrane phospholipids and MHC molecules (Fujiwara et al. 2002; Umemura et al. 2008). These MSD-based classifications assume a single mode of motion within each trajectory, and as such, are suitable for identifying the global behavior of all trajectories in an experiment, or classifying trajectories into subpopulations exhibiting different characteristics. A different class of statistical analysis exists for identifying transient confinement (Simson et al. 1995; Meilhac et al. 2006), or transient directed motion (Bouzigues and Dahan 2007; Arcizet et al. 2008) within individual trajectories. Methods used to detect these transient events are typically based on a sliding average technique: MSD is calculated for short windows along the trajectory to identify deviations from Brownian diffusion within segments of an individual track. An issue with these methods is their need for a judicious choice of input parameters such as the length of the sliding window and the assumed size of confinement regions. Changes in these parameter values could alter the results significantly. We have recently developed a novel technique to identify transient changes in diffusivity within single trajectories that does not require any user-supplied parameters. Our method uses a “hidden Markov” model to identify transient periods of reduced mobility in single-particle tracks of the integrin LFA-1 and the phosphatase CD45 on the surface of T cells (Das et al. 2009; Cairo et al. 2010). The term “hidden Markov” refers to a collection of methods that exploit changes in the behavior of an observed variable to make inferences about an unobservable, or “hidden,” variable. In our approach, we use observations of particle motion to break particle trajectories up into rapidly diffusing and slowly diffusing segments. We were able to attribute reduced mobility to a binding interaction between the cell-surface receptor and the cortical actin cytoskeleton. Furthermore, our method gives estimates for the rates of transitions between the two distinct states that can be interpreted as chemical binding rates for the integrin–actin interaction. As an extension, we have also developed an algorithm to identify transient periods of directed motion within individual trajectories (Morrison 2010). As shown in Figure 5.1, our segmentation algorithm robustly detects regions within which particle diffusivity is reduced, and we believe that this type of analysis will allow SPT-based assays to be used to detect and characterize membrane domains on the cell surface. We are currently testing this application. Directed motion on the plasma membrane has been observed in SPT experiments for proteins such as E-cadherin (Kusumi et al. 1993), MHC II (Wilson et al. 1996), and gamma-aminobutyric acid (GABA) receptors on nerve growth cones (Bouzigues and Dahan 2007). Bouzigues and Dahan presented a sliding window statistical analysis that estimates velocity and diffusion coefficients in tracks where a particle switches between directed and pure diffusive modes. They found that drift occurs most often in the central region of the cell rather than in the actin-rich lamellipodia, thus defining two distinct domains for GABA receptor mobility.

Models of Plasma Membrane Domains Studied with SPT The analysis of particle trajectories with anomalous diffusion motivated the study of three main classes of membrane domain models:

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pure diffusion confined diffusion scale: 0.5 µm Figure 5.1. Detection of regions of reduced mobility in SPT data. We show a simulated diffusive particle track across a landscape where the particle mobility is reduced within diskshaped domains. The particle’s ability to enter the disks is also reduced to simulate picket fences at domain boundaries. The particle track has been segmented into free (black) and reduced mobility (gray) parts using our hidden Markov analysis (Das et al. 2009). Parameters: Dconfined = 0.01 μm2/s, Dfree = 1 μm2/s, pjump = 0.1.

• membrane picket fence model, • lipid rafts, • protein–protein interactions and clustering. The membrane-skeleton fence model was first proposed by Sako and Kusumi after performing SPT on the α2-macroglobulin receptor (Sako and Kusumi 1994). They observed short-term confined diffusion within a submicron-sized compartment and longterm hop diffusion between compartments (see also Chapter 1). Partial destruction of the cytoskeleton decreased the confined diffusion mode and increased the simple diffusion. This model is further evidenced by experiments where the residency times within corrals decrease when the cytoplasmic side of the protein is removed (Sako et al. 1998; Leitner et al. 2000). More recently, an impressive experiment shows that single FcεRI receptors on the surface of mast cells tend not to cross green fluorescent protein (GFP)-labeled cortical actin cytoskeleton barriers (Andrews et al. 2008). Hop diffusion has been observed in the trajectories of many transmembrane proteins such as E-cadherin (Sako et al. 1998), transferrin receptor (Sako and Kusumi 1994), Band-3 (Tomishige et al. 1998; Leitner et al. 2000), and MHC II (Umemura et al. 2008). The model was revised after SPT experiments with the unsaturated phospholipid 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) were performed by the Kusumi lab. Labeled DOPE was only tracked on the outer leaflet of the plasma membrane and thus had no cytoplasmic domain to directly interact with the cytoskeleton fence. Despite this, hop diffusion was observed in the trajectories (Fujiwara et al. 2002; Murase et al. 2004). This led to the hypothesis that there are transmembrane anchor protein pickets bound to the skeleton fence that block the free diffusion of lipids and proteins via steric

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hindrance and hydrodynamic friction effects. Careful biophysical modeling led to the prediction that the skeleton fence must be 20–30% covered by these picket proteins (Fujiwara et al. 2002). There are two major consequences of the picket fence model. First, it explains why diffusion coefficients measured in liposomes are much higher than in live cells. Second, oligomers—formed due to receptor cross-linking, or through other molecular associations—have a much lower probability of crossing fence barriers due to their size, explaining the discrepancy between what is predicted by the Stokes–Einstein relation and what is observed experimentally (Kusumi et al. 2005). Lipid rafts may also play a major role in modulating the mobility of membrane components. It is conjectured that lipid rafts may increase clustering of raft-philic proteins during signaling (Kusumi et al. 2005; Day and Kenworthy 2009). Because lipid rafts are so small (<10–200 nm) (Jacobson et al. 2007), SPT is an effective way to study them due to its high resolution. To investigate the importance of rafts, it is common to deplete cholesterol or sphingolipids to see how this influences the diffusivity of single molecules. Jacobson and colleagues used this protocol in their experiments with the raft-associated transmembrane proteins Thy-1 and GM1. Depletion led to a decrease in the <200-nm confinement zones and confinement times, yet it had no effect on the diffusion of a nonraftassociated protein (Sheets et al. 1997; Dietrich et al. 2002). However, interpretations of these experiments remain controversial since cholesterol depletion affects the distribution of phosphatidylinositol 4, 5-bisphosphate (PIP2) with unknown effects on the cortical actin cytoskeleton. It has also been suggested that the small size and short lifetimes of lipid domains results in inadequate detection by SPT (Destainville et al. 2008). Protein clusters are important for receptor signaling, and it is believed that cluster formation is facilitated by lipid rafts and the cytoskeleton fence. However, specialized protein–protein interactions provide the specificity required for signaling within these microdomains. Destainville and colleagues showed that under pairwise protein interaction, the small-scale individual protein diffusion coefficient (Dmicro) is inversely proportional to the concentration of proteins within a cluster and thus is proportional to the area of the cluster (Dmicro ∼ area). This relationship was observed in single-particle tracks of the μopioid receptor (a G-protein-coupled receptor) on normal rat kidney (NRK) cells (Daumas et al. 2003). After thorough statistical analysis, they concluded that confinement by lipid rafts or the cytoskeleton cannot explain the observed area dependence (Meilhac et al. 2006). Instead, they proposed that the receptors are undergoing “walking confined diffusion” described by short-term confined diffusion within a protein cluster and long-term simple diffusion of the entire cluster. Stable protein aggregates have been observed using SPT on other cell types as well—activated T cells being one example (discussed below) (Cherry et al. 1998; Douglass and Vale 2005). Their existence is also supported by numerical simulations by Destainville (2008). Although the role and biogenesis of many protein clusters remain as an open question, SPT has allowed for rapid advancement of knowledge in this area.

Consequences for T-Cell Signaling SPT has been an invaluable tool in the study of T-cell signaling. During T-cell activation, a coordinated reorganization of the cytoskeleton and signaling molecules forms a highly concentrated signaling region (the immunological synapse) containing clusters of TCRs. Using SPT, cytoskeleton fences, lipid rafts, and protein–protein interactions have all been found to play important roles in this process. A critical step is the recruitment of signaling molecules into the core of the immunological synapse. The recruitment of one particular protein, Lck (an Src family kinase), was observed through SPT—and both lipid rafts

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and the cytoskeleton fence were required (Ike et al. 2003). It is known that the actin cytoskeleton is thought to be concentrated toward the core of the synapse, leading to slower diffusion and thus longer residency time around TCR clusters. Researchers hypothesize that this effect is enhanced by Lck’s association with lipid rafts (Ike et al. 2003). Once at the center, further single-particle experiments combined with FRAP and fluorescence imaging show that Lck preferentially associates with some proteins (CD2 and LAT) and not with others (CD45). These interactions are independent of the cytoskeleton and lipid rafts (Douglass and Vale 2005). It is hypothesized that these protein–protein interactions concentrate or exclude specific proteins in order to facilitate effective signaling. This is one example of how sophisticated analysis of single-particle trajectories has led to a better understanding of the complex mechanisms governing a protein’s spatiotemporal dynamics.

FCS FCS is a technique that measures fluctuations in the intensity of a freely diffusing fluorophore within a small excitation volume (Lakowicz 2006; Mütze et al. 2009). The autocorrelation function of the fluctuations contains information about the concentration and mobility of the molecule: The higher the concentration, the larger the magnitude of the autocorrelation function at short times, and the greater the mobility, the faster the decay in autocorrelation. The cross correlation between signals from two different fluorophores are a measure of correlated motion and putative associations between them. A typical FCS setup consists of a confocal microscope augmented with sensitive avalanche photodiodes (APDs) to efficiently detect the low fluorescence intensity within the focal volume, and a correlator that can rapidly compute the correlation functions in real time.

What Does FCS Reveal about Membrane Organization? A number of technical challenges limit the application of FCS to the study of membrane organization and dynamics. As discussed in detail in Ries and Schwille (2008), these include difficulty in accurately determining the axial position of the membrane within the focal volume, leading to a systematic error in estimating the detection area, and the necessity of long acquisition times due to slow diffusion in the membrane, resulting in artifacts such as photobleaching and a loss of stability. Consequently, FCS measurements have been primarily limited to identifying differences in translational mobility of raft and nonraft markers on model membranes (Bacia et al. 2004) and have yielded limited insight into membrane organization in live cells. The problem of estimating the detection area has been somewhat circumvented by the development of “calibration-free” FCS techniques such as Z-scan FCS and two-focus FCS (reviewed in Chiantia et al. 2009). Despite these limitations, FCS measurements have revealed important details about the spatial organization of signaling in mast cells and T cells. Cross-correlation measurements between fluorescently labeled FcεRI and Lyn on live mast cells show multiple transient peaks in the association between them within minutes of antigen stimulation, followed by a decline at longer times (Larson et al. 2005). Notably, in this study, the lateral mobility of Lyn was found to decrease throughout the time course of the experiments, suggesting spatial reorganization of the membrane as a consequence of IgE signaling. A similar pattern of association was observed in T cells using FCS measurements between TCR and LAT. The two molecules show little or no association in the absence of antigen stimulation, but an appreciable correlation that persists for tens of minutes in the presence of antigen (Lillemeier et al. 2010).

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FUTURE PERSPECTIVES Fluorescence microscopy has become an invaluable tool for studying membrane domains. However, beyond the methods described here, it is likely that new experimentation and improved data analysis/modeling techniques will play important roles in the future. We draw particular attention to the rapid development of super-resolution fluorescence techniques such as stimulated emission depletion (STED) and photo-activated localization microscopy (PALM) (Hess et al. 2006; Hein et al. 2008). In both these techniques, resolutions of tens of nanometers are achieved, which will allow us to probe raft structures at a finer scale than previously. For example, the mobility of labeled membrane components has been studied at the millisecond timescale using STED by holding the (50-nm scale) region of observation fixed and observing single molecules transiting the region (Eggeling et al. 2008). However, studying raft dynamics over larger spatial scales may be challenging since the temporal resolution of STED and PALM over a larger area is currently limited (at most, tens of frames per second have been achieved; Hess et al. 2007). The most likely scenario is that super-resolution techniques will be applied in tandem with other techniques described here. The recent study of Lillemeier et al. is a case in point, combining EM, PALM, and FCS to study protein islands during T-cell activation (Lillemeier et al. 2010). The combination of high- and medium-resolution techniques provides a potent combination for the study of static and dynamic properties of membrane domains and cell signaling.

ACKNOWLEDGMENTS This work was supported by the National Science and Engineering Research Council, the Pacific Institute of Mathematical Sciences, and the Mathematics of Information Technology and

Complex Systems National Centre of Excellence. We thank Christopher Cairo, Omer Dushek, Gerda de Vries, and Vishaal Rajani for useful discussions.

ABBREVIATIONS AFM BCR CFP D EM FCS FD FDA FRAP FRET

atomic force microscopy B-cell receptor cyan fluorescent protein diffusion coefficient of the particle electron microscopy fluorescence correlation spectroscopy donor fluorescence intensities in the absence of an acceptor donor fluorescence intensities in the presence of an acceptor fluorescence recovery after photobleaching fluorescence resonance energy transfer

GABA GFP GPI MHC MSD R R0 SPT TCR TIRF YFP

gamma-aminobutyric acid green fluorescent protein glycosylphosphatidylinositol major histocompatibility complex mean square displacement separation between molecules Forster radius single-particle tracking T-cell receptor total internal reflection fluorescence yellow fluorescent protein

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Sohn HW, Tolar P, Jin T, Pierce SK. 2006. Fluorescence resonance energy transfer in living cells reveals dynamic membrane changes in the initiation of B cell signaling. Proc Natl Acad Sci U S A 103:8143–8. Sohn HW, Tolar P, Pierce SK. 2008. Membrane heterogeneities in the formation of B cell receptor-Lyn kinase microclusters and the immune synapse. J Cell Biol 182: 367–79. Sprague BL, McNally JG. 2005. FRAP analysis of binding: proper and fitting. Trends Cell Biol 15:84–91. Tolar P, Sohn HW, Pierce SK. 2005. The initiation of antigen-induced B cell antigen receptor signaling viewed in living cells by fluorescence resonance energy transfer. Nat Immunol 6:1168–76. Tolar P, Sohn HW, Pierce SK. 2008. Viewing the antigeninduced initiation of B-cell activation in living cells. Immunol Rev 221:64–76. Tomishige M, Sako Y, Kusumi A. 1998. Regulation mechanism of the lateral diffusion of band 3 in erythrocyte

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membranes by the membrane skeleton. J Cell Biol 142: 989–1000. Umemura YM, Vrljic M, Nishimura SY, Fujiwara TK, Suzuki KG, Kusumi A. 2008. Both MHC class II and its GPI-anchored form undergo hop diffusion as observed by single-molecule tracking. Biophys J 95:435–50. Varma R, Mayor S. 1998. GPI-anchored proteins are organized in submicron domains at the cell surface. Nature 394:798–801. Wilson KM, Morrison IE, Smith PR, Fernandez N, Cherry RJ. 1996. Single particle tracking of cell-surface HLA-DR molecules using R-phycoerythrin labeled monoclonal antibodies and fluorescence digital imaging. J Cell Sci 109:2101–9. Yokosuka T, Sakata-Sogawa K, Kobayashi W, Hiroshima M, Hashimoto-Tane A, Tokunaga M, Dustin ML, Saito T. 2005. Newly generated T cell receptor microclusters initiate and sustain T cell activation by recruitment of Zap70 and SLP-76. Nat Immunol 6:1253–62.

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PART

II

ORGANELLAR DOMAINS ME MBR ANE - BO UND cellular organelles represent fundamental cellular domains that are characterized by their distinctive morphology. Mitochondria, organized as an intracellular network and encircled by a dual-membrane system, fulfill various, often pivotal functions, being most well known for their role as the cellular powerhouse (Chapter 6). The endoplasmic reticulum (Chapter 7) and Golgi apparatus (Chapter 8) are key components of the secretory pathway that controls protein and lipid biosynthesis and targeting for secretion or delivery to intracellular sites (see also Chapters 18, 21, and 22). Endosomes (Chapter 9) are downstream of endocytic plasma membrane domains, such as clathrin-coated pits (Chapter 2), and function to sort internalized material for recycling or targeting to lysosomes (Chapter 10) for degradation. Lysosomal compartments also receive material for degradation via phagocytosis of large particles and autophagy of cellular material (Chapter 10). Importantly, domain organization of membrane organelles (domains within domains) is key to their function. This can be observed both at the ultrastructural level (i.e., rough vs. smooth endoplasmic reticulum and Golgi cisternae) and at the level of lipid- and protein-based domain organization, suggesting that mechanisms that regulate domain organization at the plasma membrane (Chapters 1–5) may also apply to intracellular membrane organelles. Intriguingly, it is increasingly clear that interactions between organelles, in particular endoplasmic reticulum junctions (Chapter 11), regulate organellar crosstalk and function.

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CH A P T E R

6

MITOCHONDRIA Michael Zick Andreas S. Reichert

DEFINITION On an average day the human body consumes a quantity of adenosine-5′-triphosphate (ATP) that almost equals its own weight. The vast majority, to satisfy this tremendous demand, is supplied by mitochondria, which are cytosolic organelles of endosymbiotic origin. Mitochondria harbor a sophisticated enzymatic machinery that employs molecular oxygen to complete the oxidation of fats, proteins, and sugars to carbon dioxide and water, finally transforming the released energy into ATP. As they function in an integrated network that is subjected to constant remodeling by fusion and fission events, one can consider mitochondria as the “power grid” of eukaryotic cells. Furthermore, they play pivotal roles in a diverse set of functions such as the metabolism of lipids and amino acids, the biosynthesis of iron–sulfur clusters, heme, and further prosthetic groups, ion homeostasis, cell signaling, and the regulation of apoptosis. To be able to fulfill these various functions, the mitochondrion is organized into different compartments (Fig. 6.1). This is achieved by the two membranes, inner (IM) and outer (OM) mitochondrial membranes, that surround the organelle. The IM encloses the mitochondrial matrix and is further subdivided into two distinct regions: (1) the inner boundary membrane (IBM), which is adjacent to the OM, and together with it delineates the intermembrane space (IMS); (2) the crista membrane (CM), which represents invaginations of the IM that protrude into the matrix space, thus surrounding the intracristal space (ICS). This compartmentalization appears to be dynamic, and loss of its integrity marks a key step during the course of programmed cell death.

HISTORICAL PERSPECTIVE Discovery of Mitochondria The first time that the organelles which now refer to as mitochondria were observed dates back more than one and a half centuries. Already in 1841, J. Henle reported characteristic granules within myocytes (Henle 1841). In the year 1856, the Swiss anatomist A. Kölliker performed a first careful study of the sarcoplasmic components and described an arrangement of “blasse Körnchen” (pale grains) between the myofibrils; he even suggested a correlation of these granules with the metabolism of muscle fibers (Kölliker 1856). Due to the development of improved staining techniques, in particular the introduction of Crystal Violet (Benda 1901) and Janus Green (Michaelis 1900), these organelles had been

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-40

Outer Membrane (OM)

nm

30 -2 00 nm

12

Crista Junction (CJ)

InterMembrane Space (IMS)

- TCA Cycle - Amino Acid & Lipid Metabolism - Heme Biosynthesis - etc.

Fusion / Fission

Oxidative Phosphorylation

Apoptosis Cristae

Matrix

Inner Membrane (IM) Inner Boundary Membrane (IBM) + Crista Membrane (CM) Intracristal Space (ICS)

Fe/S Cluster Biosynthesis

Protein Synthesis

Protein Import

M

IM

IMS

OM

CS

Figure 6.1. Scheme of a mitochondrion. The architecture of a mitochondrion is determined by its two membranes, which partition the organelle into several discrete compartments that host a multitude of functions crucial for eukaryotic life. M, matrix; IM, inner membrane; IMS, intermembrane space; OM, outer membrane; CS, cytosol.

observed in basically every type of eukaryotic cell until the early twentieth century. They have been referred to with a multitude of various terms, such as chondriosomes, bioblasts, and plastochondria, until F. Cavers noticed in 1914 that these structures were essentially all the same (Cavers 1914). What has become the common name originates from C. Benda, who called these organelles “mitochondria” due to their appearance as threads (Greek mitos or “μιτoσ”) and grains (Greek chondros or “χóνδρoσ”) (Benda 1898).

Origin of Mitochondria The Endosymbiotic Theory The idea that eukaryotic organelles could be derivatives of bacteria was already put forward in the late nineteenth century, however, mainly for chloroplasts (Schimper 1883; Mereschkowsky 1905). While Altmann (1894) recognized an apparent resemblance between mitochondria and bacteria, Wallin (1923) was the first to extend the idea of an endosymbiotic origin to mitochondria in 1923. Nevertheless, it took about half a century until the endosymbiotic hypothesis gained considerable attention due to the popular work of L. Margulis (Margulis, 1981; Sagan 1967). The hypothesis received substantial support by the discovery of mitochondrial DNA (mtDNA) (Nass and Nass 1963a, b), and it was generally accepted that mitochondria were the direct descendants of a bacterial endosymbiont (Gray 1992). Comparative genomic approaches imply that mitochondria are monophyletic in origin, and the closest relatives to the protomitochondrial ancestor are considered to be found within the rickettsial subdivision of the α-proteobacteria (Emelyanov 2001). It is, however, a matter of discussion whether the endosymbiotic evolution of eukaryotic cells was indeed a serial event or mitochondria originated at essentially the same time as the nuclear compartment (Gray et al. 1999; Emelyanov 2003).

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Mitochondria-Related Organelles: Anaerobic Mitochondria, Hydrogenosomes, and Mitosomes Mitochondria evolved over a period of at least 1500 million years under an enormous range of environmental conditions (van der Giezen and Tovar 2005). Thus, a diverse set of related organelles emerged, supposedly from one single ancestor, ranging from archetypal aerobic mitochondria, via various anaerobic versions and hydrogenosomes, to the most distantly evolved forms, mitosomes (Tielens et al. 2002; Hackstein et al. 2006; van der Giezen 2009). Anaerobic mitochondria, which can be found in several protists, flatworms, parasitic nematodes, invertebrates, and algae, possess a respiratory chain, as classical aerobic mitochondria do, but they do not use oxygen as a final electron acceptor. Rather they harbor alternative terminal oxidases that are capable of using either an environmental (e.g., NO3−) or an endogenously produced (e.g., fumarate) substrate as a final electron acceptor (Tielens et al. 2002). Hydrogenosomes, which can be found in a wide spectrum of anaerobic protists, represent another type of anaerobic ATP-producing relative of mitochondria. They do not possess a membrane-associated electron transport chain but employ substrate-level phosphorylation and use protons as terminal electron acceptor, thus producing hydrogen, to generate ATP (Hackstein et al. 2006; van der Giezen 2009). Another type of double-membrane enclosed organelles, which are considered the result of reductive evolution of mitochondria, are mitosomes (van der Giezen 2009). They have been identified in a range of anaerobic/microaerophilic parasitic protozoa in recent years (Tovar et al. 1999; Williams et al. 2002; Tovar et al. 2003; Goldberg et al. 2008). Mitosomes have lost the capability of producing ATP, and so far it remains largely elusive whether they have a relevance to cell physiology beyond their role in iron–sulfur cluster biogenesis (Tovar et al. 2003; Aguilera et al. 2008; Maralikova et al. 2009). Further research on the organellar descendants of mitochondria will substantially contribute to our global understanding of the evolution and cell biology of eukaryotes.

Mitochondrial Research throughout the Last Century Over its long history, mitochondrial research has revealed a large number of essential findings, primarily concerning cellular metabolism and energy conversion as the driving force of life (Fig. 6.2). Along the way, it has sustainably influenced many scientific disciplines, for example, biochemistry and cell biology. Due to their enormous breadth, the tremendous advancements leading to our current knowledge on mitochondria are beyond the scope of this contribution. However, the progress of mitochondrial research has been well documented by a series of excellent reviews throughout the years, to which the reader can refer as sources of additional information (Wallin 1923; Cowdry 1953; Crane 1961; Ernster and Schatz 1981; Rasmussen 1995; Scheffler 2001; Bechtel and Abrahamsen 2007).

THE BUILDING BLOCKS OF MITOCHONDRIA The Mitochondrial Genome Reminiscent of their origin, mitochondria of all species contain their own set of DNA. However, the size of these genomes, their architecture, as well as the number and nature of the encoded proteins (Burger et al. 2003), and even the mitochondrial genetic code itself (Knight et al. 2001), vary greatly among different organisms. The human mitochondrial

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Apoptosis Dynamics Mitochondrial Disease Metabolism Bioenergetics

Cytology 1900

1925

1950

Molecular Evolution Protein Import 1975

2000

Figure 6.2. A century of mitochondrial research. During the twentieth century mitochondrial research has experienced many highly productive periods that have significantly advanced our understanding of many aspects of mitochondria in particular, and cell biology in general. Thus, it is not surprising that a considerable number of researchers that have dedicated their interest to mitochondria have been rewarded with a Nobel Prize in Physiology or Medicine, or in Chemistry (indicated by encircled N below the timeline; http://nobelprize.org). Some key discoveries of mitochondrial biology are also specified with their corresponding references: 1Michaelis (1900); 2 Warburg (1913); 3Keilin (1925); 4Krebs and Henseleit (1932a,b, c); 5Krebs and Johnson (1937); 6 Hogeboom et al. (1947, 1948); 7Palade (1952) and Sjostrand (1953); 8Mitchell (1961); 9Nass and Nass (1963a, b) and Schatz et al. (1964); 10Sagan (1967); 11Anderson et al. (1981); 12Wallace et al. (1988a, b); and 13Liu et al. (1996).

genome, for instance, encodes 37 genes on a compact, circular DNA molecule of about 16.6 kbp in size (Fig. 6.3). The multiple copies are organized in numerous protein–DNA complexes, commonly referred to as nucleoids, which are likely to play important roles in protection, inheritance, replication, and transcription of the mitochondrial genome (Chen and Butow 2005; Malka et al. 2006; Falkenberg et al. 2007; Spelbrink 2009). Analysis of mtDNA provides highly useful information in different scientific fields. (1) Alterations of mtDNA play a crucial role in numerous human disorders (Taylor and Turnbull 2005; Brandon et al. 2006; Copeland 2008; DiMauro and Schon 2008) and in the aging process (Trifunovic et al. 2004; Trifunovic et al. 2005; Kujoth et al. 2007; Edgar et al. 2009; Greaves and Turnbull 2009; Khrapko and Vijg 2009). (2) Since the mitochondrial genome of almost 2000 distinct organisms is completely sequenced (O’Brien et al. 2009), and approximately 7000 human mtDNA sequences are available (Ingman and Gyllensten 2006; Ruiz-Pesini et al. 2007), mtDNA represents a versatile tool for evolutionary analyses and studies in population biology (Ballard and Rand 2005; Pakendorf and Stoneking 2005; Underhill and Kivisild 2007). (3) Analysis of mtDNA even plays an expanding role in the forensic sciences (Budowle et al. 2003).

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12

OH T

D Loop

SNHL DEAF

P

PL

MM E N DM LS LHON Dystonia

rR S

16

MELAS MMC DMDF LHON Dystonia

MELAS

Human mtDNA

MELAS Q ADPD

I M ND2

W

16,569 bp

MERRF LS LHON

LS A O L N C Y SNHL CO I

LS NARP PEM MILS AMDF MERRF FBSN UCN S COII

K

ATP6

I CO

II

ND 3 R 4L D N G

ATP8

D

LS

LCUN AGY S H

ND 4

ND1

UUR

ND 5

L

Cy tb

D6

NA

V

PH1 PH2 F A RN Sr

91

Figure 6.3. The human mitochondrial genome. In humans, the mitochondrial DNA encodes 37 gene products; 2 ribosomal RNAs (12S and 16S), 22 tRNAs (indicated by the cognate amino acids in one-letter code), and 13 protein subunits of the respiratory chain: NADH dehydrogenase (ND1-4, -4L, -5, and -6); cytochrome b (Cytb); cytochrome c oxidase (COI-III); ATP synthase (ATP6 and ATP8). The origins of light- and heavy-strand replication (OL and OH), the promotor of the light strand (PL), as well as the two transcription initiation sites of the heavy strand (PH1 and PH2) are depicted. The D Loop region is a non-coding, regulatory region involved in mtDNA replication. A growing number of primarily neurological disorders have been linked to mutations within the mtDNA. In fact, with a prevalence of about 10 in 100,000 they have to be considered one of the most common inherited neuromuscular disorders (Schaefer et al. 2008). Some of the disease entities are indicated next to the correspondingly affected gene product. Abbreviations: ADPD, Alzeimer ’s disease and Parkinson’s disease; AMDF, ataxia, myoclonus and deafness; DEAF, maternally inherited deafness; DM, diabetes mellitus; DMDF, diabetes mellitus and deafness; FBSN, familial bilateral striatal necrosis; LHON, Leber hereditary optic neuropathy; LS, Leigh syndrome; MELAS, mitochondrial encephalomyopathy, lactic acidosis, and stroke-like episodes; MERRF, myoclonic epilepsy and ragged red muscle fibers; MILS, maternally inherited Leigh syndrome; MM, mitochondrial myopathy; MMC, maternal myopathy and cardiomyopathy; NARP, neurogenic muscle weakness, ataxia, and retinitis pigmentosa; PEM, progressive encephalomyopathy; SNHL, sensorineural hearing loss.

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Mitochondrial Biogenesis Mitochondrial energy supply and metabolic activity require a vivid modulation in order to meet constantly changing physiological demands. Therefore, mitochondrial biogenesis is dynamically regulated, primarily via the transcriptional control of both nuclear and mitochondrial genomes (Hock and Kralli 2009). As mitochondria are nonautonomous organelles, the vast majority of its constituents need to be imported. Best characterized is the translocation of proteins across the two mitochondrial membranes; a basic overview is shown in Figure 6.4. Considerably less well explored is the delivery of phospholipids to the mitochondrial compartment (Daum and Vance 1997). The zones of close contact between mitochondria and the endoplasmic reticulum (ER; see also Chapter 11 for further discussion of ER– mitochondria interactions), also called mitochondria-associated membranes (MAM), seem to play a crucial role in the synthesis and nonvesicular trafficking of phospholipids (Gaigg et al. 1995; Hayashi et al. 2009; Wieckowski et al. 2009). The import of one further class of molecules, tRNAs, and its physiological relevance to mitochondrial translation is only just beginning to be understood (Tarassov et al. 2007; Alfonzo and Soll 2009; Duchene et al. 2009).

The Proteome of Mitochondria Mitochondria represent an extremely protein-rich compartment exhibiting a highly concentrated mixture of hundreds of enzymes in the matrix (26–56% weight/volume; Hackenbrock 1968) as well as a high protein : phospholipid ratio in the IM (>3:1 by weight; OM membrane, ∼1:1 by weight; Sperka-Gottlieb et al. 1988). Thus, in order to gain a deeper understanding of the multiple functions of this complex organelle, a tremendous effort has been made in defining the mitochondrial proteome (Da Cruz et al. 2005; Meisinger et al. 2008). Around 1500–2000 distinct proteins are estimated to be localized to the mitochondrial subcompartments: matrix (∼45%), IM (∼40%), IMS (∼5%), and OM (∼10%). Multiple databases on the protein composition of mitochondria in diverse organisms are available; a representative list can be found in Table 6.1. The problem of false positive or negative attribution of mitochondrial localization is aggravated by the fact that at least one-tenth of the mitochondrial proteins may exhibit a dual cellular localization (Sickmann et al. 2003; Regev-Rudzki and Pines 2007; Pagliarini et al. 2008). A next and crucial step will be to link the many newly discovered members of the mitochondrial inventory with their functions. Analysis of the dynamic alterations of the mitochondrial proteome under various, including pathological, conditions will hopefully expedite this endeavor. In particular, the integration of all the newly obtained information into a comprehensive mitochondrial interactome has the potential to uncover further details about the roles that mitochondria play in cellular processes and the pathogenesis of disease (Shutt and Shadel 2007; Reja et al. 2009). More specialized studies, surveying the mitochondrial phosphoproteome, provided good evidence that protein phosphorylation serves as a regulatory mechanism across a broad range of important processes in mitochondria (Lee et al. 2007; Reinders et al. 2007; Ito et al. 2009). In addition, acetylation has recently emerged as a posttranslational modification that regulates mitochondrial proteins (Huang et al. 2010); it gained enormous interest, as the main mitochondrial deacetylase (SIRT3; Lombard et al. 2007) has been implicated in modulation of mitochondrial bioenergetics (Ahn et al. 2008) and control of longevity (Bellizzi et al. 2005). A further posttranslational modification that needs to be explored is tyrosine nitration (Liu et al. 2009). Comprehensive quantitative analyses of

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Precursor with internal targeting signal

Precursor with MTS presequence

CS

++ ++

OM Proteins

93

β-Barrel Proteins ?

TOM small Tims

IM Proteins (Stop-Transfer)

Oxa1 IM Proteins (conservatively sorted or mtDNA mtRibosome encoded)

OM

TOB

Oxidized IMS Proteins S S IMS S S

MIA TIM23

IM

TIM22 Carrier Proteins

Hsp70

M

Hsp70

MPP

Matrix Proteins

Figure 6.4. Mitochondrial protein import. The major portion of mitochondrial proteins is encoded in the nuclear DNA. They are synthesized by cytosolic ribosomes and consecutively transported to their respective destination within mitochondria. Thereby they are directed by mitochondrial targeting signals. The necessary information is present either in the matrix-targeting sequence (MTS), a presequence of 10–80 amino acids that forms an amphipathic helix with one hydrophobic and one positively charged face or within an internal targeting signal instead. Several major complexes are involved in the translocation of proteins across the mitochondrial membranes (for a more detailed view see Neupert and Herrmann 2007; Chacinska et al. 2009). TOM complex: Effectively all proteins that are imported into mitochondria are translocated across the outer mitochondrial membrane via the TOM complex. Besides its function as an entry gate, the TOM complex also mediates the insertion of certain outer membrane proteins (Walther and Rapaport 2009). TOB complex (topogenesis of β-barrel proteins; also termed SAM complex): Precursors of β-barrel proteins pass through the TOM channel, form a temporary complex with small Tim proteins in the IMS, and are thereby guided to the TOB complex, which then facilitates membrane insertion and assembly of the new β-barrel protein (Paschen et al. 2005). TIM23 complex: In concerted action with the TOM complex, it translocates all matrix proteins, the majority of inner membrane proteins, and many proteins of the intermembrane space (not depicted; Popov-Celeketic et al. 2008). The TIM23 translocase is thereby driven by the electrochemical potential (Δψ) across the inner membrane. Subsequently, the matrix localized import motor, which is driven by ATP hydrolysis, takes over. One of its core components is the chaperone Hsp70, which mediates the vectorial protein movement supposedly via a Brownian ratchet mechanism (Yamano et al. 2008). Inner membrane proteins can either be arrested at the level of TIM23 while being translocated and thereupon laterally sorted (Stop-Transfer pathway); alternatively, the preprotein is initially completely translocated into the matrix and successively inserted into the inner membrane in an exportresembling manner (Conservative sorting pathway). A third class is assembled into the inner membrane via the TIM22 complex: The family of solute carrier proteins and hydrophobic TIM subunits are complexed in the IMS by small Tim proteins that guide the precursors to the TIM22 translocase into the inner membrane in a Δψ-dependent reaction (Rehling et al. 2004). Oxa1 complex: Several of the conservatively sorted inner membrane proteins are inserted by the Oxa1 complex. Additionally, this complex mediates the insertion and assists the assembly of the mtDNA-encoded subunits of the respiratory chain—at least in part cotranslationally (Ott and Herrmann 2009). MIA complex: A subset of IMS proteins contains disulfide bonds. Their import is mediated by the Mia40/Erv1 disulfide relay system. This pathway is coupled to oxidative folding of the proteins that are thereby trapped in the IMS (Deponte and Hell 2009).

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TABLE 6.1.

MITOCHONDRIA

List of Mitochondrial Proteome Databases

Database MitoP2 MitoMiner MitoCarta MitoRes

Species

Methods

Hs, Mm, Nc, Sc, At Hs, Mm, Rn, Dm, Ce, Sc, Bt, Pf Hs, Mm

MS, FT, DA MS, FT, DA MS, FT, DA DA

http://www.mitop.de: 8080/mitop2/ http://mitominer.mrcmbu.cam.ac.uk/ http://www.broadinstitute. org/pubs/MitoCarta/ http://mitores.ba.itb.cnr.it/

Elstner et al. 2008; Elstner et al. 2009 Smith and Robinson 2009 Pagliarini et al. 2008

MS, DA

http://www.mitoproteome.org/ http://bioinfo.nist.gov/ http://141.61.102.16/ormd/ http://www.biochem.oulu.fi/ proteomics/ymp.html http://www-deletion.stanford. edu/YDPM/ http://www.plantenergy.uwa. edu.au/ampdb/

Cotter et al. 2004; Guda et al. 2007 — Foster et al. 2006 Ohlmeier et al. 2004

MitoProteome

Hs, Mm, Rn, Dm, Ce, Bt, Dr, m* Hs

HMPDb ORMD YMP

Hs Mm Sc

DA MS, FT MS

YDPM

Sc

MS

AMPDb

At

MS, DA

TrypsProteome

Tb

MS

URL

http://www.trypsproteome.org/

Reference

Catalano et al. 2006

Steinmetz et al. 2002; Prokisch et al. 2004 Heazlewood et al. 2004; Heazlewood and Millar 2005 Panigrahi et al. 2009

Several databases exist that integrate the growing body of proteomic data on the composition of mitochondria in diverse organisms. Species: At, Arabidopsis thaliana; Bs, Bos taurus; Ce, Caenorhabditis elegans; Dm, Drosophila melanogaster; Dr, Danio rerio; Hs, Homo sapiens; m*, metazoa listed in UniProt Knowledgebase; Mm, Mus musculus; Nc, Neurospora crassa; Pf, Plasmodium falciparum; Rn, Ratus norvegicus; Sc, Saccharomyces cerevisiae; Tb, Trypanosoma brucei. Methods: MS, mass spectrometry of purified mitochondria; FT, subcellular localization of fluorescently tagged proteins; DA, analysis of public databases.

the posttranslational modifications of mitochondrial proteins (Distler et al. 2008) will sustain these concepts in the future (Allison et al. 2009), and help to understand their role in signaling pathways within mitochondria (Pagliarini and Dixon 2006).

Mitochondrial Membrane Lipids The role of mitochondrial membrane lipids (Table 6.2; Daum 1985), in contrast, has been rather neglected. Recent lipidomic studies, however, imply that the phospholipid composition of mitochondria has a considerable impact on their function (Kiebish et al. 2008a, b, c). Future analyses focusing on the dynamic changes in phospholipid composition of mitochondrial membranes will potentially reveal highly interesting novel insights. The characteristic mitochondrial phospholipid, diphosphatidylglycerol, better known as cardiolipin, has, nevertheless, gained considerable attention throughout the years (Schlame et al. 2000; Chicco and Sparagna 2007; Houtkooper and Vaz 2008). Preferentially localized to the mitochondrial IM, it affects energy metabolism by stabilizing respective enzyme complexes (Bogdanov et al. 2008; Claypool 2009; Klingenberg 2009; Wittig and Schagger 2009). Furthermore, it plays an important role in the structural organization of mitochondrial membranes (Schlame 2008; Lewis and McElhaney 2009; Schlame and Ren 2009) and regulates different steps of the mitochondrial apoptotic pathway (Orrenius et al. 2007; Ott et al. 2007; Kagan et al. 2009; Schug and Gottlieb 2009). Impairment of

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TABLE 6.2.

Phospholipid Composition of Mitochondria

Mammals

PC PE CL PI PS PA

95

Plants

Fungi

IM

OM

IM

OM

IM

OM

40 34 18 5 3 —

54 29 <1 13 2 —

42 38 15 5 — —

47 27 3 23 — —

38 30 20 8 2 2

45 23 6 21 3 2

Adapted from Daum and Vance (1997) with permission from Elsevier. Data taken from Daum (1985), Hovius et al. (1990), Zinser et al. (1991), de Kroon et al. (1997), and Wriessnegger et al. (2009). In contrast to phospholipids, sterols comprise only a minor fraction of mitochondrial membranes (∼1:30; Bottema and Parks 1980). The approximate levels of each individual phospholipid are indicated as percentage of the total phospholipid content. PC, phosphatidylcholine; PE, phosphatidylethanolamine; CL, cardiolipin = diphosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; PA, phosphatidic acid; IM, inner mitochondrial membrane; OM, outer mitochondrial membrane.

cardiolipin species homeostasis, caused by a deficiency in cardiolipin remodeling, thus leads to a polysymptomatic disease called Barth syndrome (Hauff and Hatch 2006; Schlame and Ren 2006).

RECENT AND PROSPECTIVE DEVELOPMENTS IN MITOCHONDRIAL RESEARCH Many of the topics that are currently fueling mitochondrial research have already been in the focus of attention in earlier times. Gathered knowledge and the continuous advancement of the experimental tools allow readdressing and extending these questions with confidence toward increasing our understanding of mitochondrial biology.

Mitochondrial Ultrastructure Organization of the IM: IBM, Cristae, and Crista Junctions A major breakthrough in our understanding of mitochondria was obtained by the pioneering work in electron microscopy of Palade and Sjöstrand in the 1950s (Palade 1952, 1953; Sjostrand 1953; Rasmussen 1995). Since then, mitochondria have been conceived as cellular compartments that are encircled by two membranes: the OM and the IM. The latter is further subdivided into two compartments: the IBM, which is apposed to the OM, and the CM, which represents membrane invaginations that protrude into the matrix compartment (Figs. 6.1 and 6.5A, B). Cristae exhibit a multitude of morphological appearances, depending on the physiological state of a cell, on the tissue, or the organism (Munn 1974; Fawcett 1981). Structural alterations, including electron-dense bodies within the matrix, are described in numerous human disorders, and dynamic changes in cristae morphology were reported to occur during apoptosis (for review see Perkins et al. 2009; Zick et al. 2009). Two major misconceptions are coined by the classical textbook presentation of mitochondria. First, mitochondria are usually represented as isolated organelles, although they rather form an interconnected, three-dimensional, tubular network in many tissues

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(A) 1959

(C)

(B) 1997

(D)

(E)

(F)

(G)

Figure 6.5. Morphology of mitochondria. (A, B) Evolving view on the ultrastructure of mitochondria. (A) Three-dimensional reconstruction of a motor end plate mitochondrion made from electron micrographs of serial thin sections at a magnification of 185,000×. Reproduced from Andersson-Cedergren (1959) with permission from Elsevier. (B) Two views of a three-dimensional reconstruction of an electron tomography of a Purkinje cell dendritic mitochondrion imaged in situ. The outer membrane is shown in dark blue, the inner boundary membrane in light blue, and the cristae membranes in yellow. Reproduced from Perkins et al. (1997) with permission from Elsevier. (C–G) Morphology of the mitochondrial network. (C) Tubular mitochondrial network in the budding yeast Saccharomyces cerevisiae. A yeast strain expressing a green fluorescent protein targeted to mitochondria was grown under nonfermentable conditions to the logarithmic growth phase. The cell wall was separately stained with calcofluor white. A surface-rendered three-dimensional stack recorded with an MMM-4Pi confocal fluorescence microscope is shown. Reproduced from Egner et al. (2002) with permission. © 2002 National Academy of Sciences, USA. (D–G) Mitochondrial morphology in HEP-G2 cells transfected with a mitochondrially targeted fluorescent protein. All images were recorded by two-photon excitation with a confocal microscope. Adapted from Plecita-Hlavata et al. (2008) with permission from Elsevier. (D, E) Cells were cultivated in 25 mM glucose. The mitochondria form a tubular reticulum with highly interconnected branches. (F, G) Cells were cultivated in 5 mM glucose before treatment. Dissipation of the mitochondrial membrane potential by the addition of 10 μM rotenone induced the fragmentation of the originally highly interconnected mitochondrial reticulum.

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(Fig. 6.5C–G). Second, cristae are still often depicted as membrane folds harboring broad openings resembling the so-called baffle model. However, in the 1990s this view was challenged by electron tomographic studies, which demonstrated that the IBM and the CM are rather connected by small pore-like openings, termed crista junctions (CJs; Figs. 6.1 and 6.5A, B; Mannella et al. 1994, 1997; Frey and Mannella 2000). Interestingly, already careful analyses of thin serial sections, performed by Andersson-Cedergren in the 1950s and by Daems and Wisse in the 1960s, suggested the existence of such narrow entry pores, termed pediculi cristae (“cristae feet”) at that time (Andersson-Cedergren 1959; Daems and Wisse 1966). The appearance of CJs is rather uniform in size and shape, exhibiting narrow tubular, ring- or slot-like structures with diameters ranging from 12 to 40 nm and lengths ranging from 30 to 200 nm (Zick et al. 2009). Compartmentalization and Dynamic Protein Distribution Based on these structural properties and the location of CJs separating the IBM from the CM, CJs were suggested to represent diffusion barriers that regulate the distribution of proteins, lipids, and metabolites between the individual mitochondrial subcompartments (Mannella et al. 1997). A major implication imposed by such a degree of organization is that entry and exit of adenosine-5′-diphosphate (ADP) or protons might be regulated by CJs. As ADP is often the limiting metabolite in oxidative phosphorylation this would represent a novel way of regulating this fundamental bioenergetic process. And indeed, several lines of evidence support that the IM is composed of two distinct subcompartments. The CM mostly contains proteins involved in oxidative phosphorylation, iron–sulfur cluster biogenesis, protein synthesis, and transport of mtDNA-encoded proteins. The IBM, however, was rather enriched in proteins involved in mitochondrial fusion and protein transport of nuclear-encoded proteins (Vogel et al. 2006; Wurm and Jakobs 2006). This protein distribution is not static but rather appears to be dynamic, depending on the physiological state. How this is regulated and what components are required to form cristae and CJs are a matter of intensive research. Molecular Players Governing Cristae and CJ Formation Even though numerous proteins affect cristae morphology, convincing evidence for a direct role has been provided only in a few cases (Zick et al. 2009). The mitochondrial F1F0-ATP synthase was shown to assemble into dimers and higher oligomeric supercomplexes via its subunits e and g (Arnold et al. 1998). Disruption of this arrangement does not compromise the enzymatic activity of the F1F0-ATP synthase but results in a gross disorganization of the mitochondrial IM, which appears concentrically stacked, mimicking an onion-like structure (Velours et al. 2009). It appears that the F1F0-ATP synthase supercomplexes act as a structural backbone mediating normal cristae appearance (Paumard et al. 2002). Thereby, dimers and oligomers of the F1F0-ATP synthase are preferentially located at regions of the membrane that exhibit a positive membrane curvature, for example, at the outer rim of a flat crista sheet, or at the tip of a crista (Strauss et al. 2008; Rabl et al. 2009). In contrast, the yeast protein Fcj1, which was also shown to be required for CJ formation, is enriched at CJs, a membrane region exhibiting a negative membrane curvature. Fcj1 genetically interacts with subunits e and g and modulates the oligomeric state of the F1F0-ATP synthase in an antagonistic manner. By that interplay between Fcj1 and the subunits e and g, IM curvature might be regulated, thereby determining cristae morphology and CJ formation (Rabl et al. 2009). Mitofilin, the mammalian ortholog of Fcj1, was also proposed to be required for normal cristae morphology and CJ formation (John et al. 2005). Other factors, such as prohibitins, were also suggested to be involved

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in CJ formation (Merkwirth and Langer 2009; Osman et al. 2009a, b), and certainly further still unknown, molecular players will have to be discovered. Apoptotic Cristae Remodeling Besides their potential role in regulation of oxidative phosphorylation, CJs gained major interest regarding their role during apoptosis. Induction of apoptosis was reported to cause opening of CJs, which in turn promoted the efficient release of cytochrome c (Scorrano et al. 2002). Thus, CJ remodeling was hypothesized as a necessary step in cytochrome c-dependent cell death pathways. Indeed, overexpression of OPA1, a large dynamin-like GTPase of the inner mitochondrial membrane, exerted an anti-apoptotic effect and prevented CJ opening during apoptosis (Cipolat et al. 2006; Frezza et al. 2006). Conversely, downregulation of OPA1 led to changes in cristae morphology and promoted cytochrome c release (Olichon et al. 2003). Due to the ability of OPA1 to form high-molecular-weight complexes, and to undergo self–self interactions, it was proposed to keep CJs in a tight or closed conformation, thereby preventing cytochrome c release. On the contrary, another study reported that upon addition of BH3-only proteins to isolated mitochondria, CJ diameters were even reduced while still cytochrome c release was promoted and OPA1 complexes were disassembled (Yamaguchi et al. 2008). This suggests that cytochrome c release can also be promoted in the absence of CJ opening. Nevertheless, OPA1 disassembly appeared to be correlated with cytochrome c release in all cases reported so far. The role of OPA1 in cristae remodeling in general is supported by the fact that Mgm1, the yeast ortholog of OPA1, was also suggested to be involved in stabilizing cristae morphology and that this also depends on its ability to form homotypic interactions (Meeusen et al. 2006). Future studies will have to address the controversial role of OPA1 in cristae remodeling during apoptosis, and to decipher how the large dynamin-like GTPases Mgm1 and OPA1 modulate cristae structure in general.

Mitochondrial Dynamics The highly dynamic behavior of mitochondria was already noticed almost a century ago (Lewis and Lewis 1914). These observations, however, fell into oblivion until the 1990s when the underlying events of fusion and fission, which constantly remodel the mitochondrial network (Fig. 6.5C–G), attracted considerable attention (Bereiter-Hahn and Voth 1994; Nunnari et al. 1997). This newly gained interest was sustained by the discovery that impairment of these processes is linked to the pathogenesis of several human neuropathies. Defects in fission or fusion of mitochondria lead to Charcot–Marie–Tooth neuropathy types 2A (Zuchner et al. 2004) and 4A (Baxter et al. 2002; Cuesta et al. 2002; Niemann et al. 2005), and autosomal dominant optic atrophy (Alexander et al. 2000; Delettre et al. 2000). Impaired mitochondrial dynamics were also associated with a pathophysiological role in neurodegenerative diseases, for example, Parkinson’s and Alzheimer ’s disease (Chen and Chan 2009; Su et al. 2010). Most of the molecular players that constitute the core machinery of mitochondrial dynamics were initially discovered and characterized in model organisms; while several key components are remarkably well conserved, the regulatory mechanisms seem to be more diverse (Westermann 2009). Fusion The fusion of mitochondria is particularly remarkable in two aspects. (1) Not only two but four lipid bilayers need to be fused in a coordinated manner. (2) In contrast to all other characterized cellular fusion events, mitochondrial fusion is not mediated by SNARE proteins. Instead, large dynamin-like GTPases that reside in the IM (Mgm1 in Saccharomyces

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cerevisiae/OPA1 in Homo sapiens) and OM (Fzo1 in S. cerevisiae/mitofusins Mfn1 and Mfn2 in H. sapiens), respectively, drive the sequential fusion of these membranes (Meeusen et al. 2004; Chan 2006; Hoppins et al. 2007; Song et al. 2009). This process is so far best characterized in yeast (Hoppins and Nunnari 2009): OM fusion requires the interaction of Fzo1 in trans, the hydrolysis of guanosine-5′-triphosphate (GTP) and a proton gradient across the IM (Meeusen et al. 2004). The in trans interaction of Mgm1 is similarly required for IM fusion, as well as an IM electrical potential is (Meeusen et al. 2004; Meeusen et al. 2006). A third protein that is essential for mitochondrial fusion in yeast is Ugo1, a modified member of the carrier protein family without any known homolog. It is distinctly required for both OM and IM fusion, and is proposed to operate at the lipid-mixing step (Hoppins et al. 2009). It is likely that further proteinaceous components of the mitochondrial fusion apparatus will be discovered. In addition, lipids play an important role in mitochondrial dynamics. Several yeast mutants defective in mitochondrial morphology were reported to exhibit altered levels of either cardiolipin, phosphatidylethanolamine, or both (Osman et al. 2009a). The mitochondrial phospholipase D (MitoPLD) was suggested to promote fusion by hydrolyzing cardiolipin after docking of two opposing outer membranes (Choi et al. 2006). Although initial concepts can be devised, we are still far from understanding the mechanistic details of how mitochondrial fusion is orchestrated. Fission Mitochondrial fission is also mediated by a large dynamin-like GTPase (Dnm1 in S. cerevisiae, DRP1 in H. sapiens), which is recruited from the cytosol to distinct patches on the outer surface of mitochondria, where it self-assembles into helical structures. This selfassembly facilitates the fission process and depends on auxiliary proteins (Fis1 and Mdv1 in S. cerevisiae), which were suggested to act as a platform on mitochondria (Lackner et al. 2009). Irrespective of the controversial discussions about the precise molecular mechanism underlying this process and its regulation, it appears that diverse modifications, such as ubiquitination, phosphorylation, sumoylation, and nitrosylation, of Dnm1/DRP1 are involved (Harder et al. 2004; Nakamura et al. 2006; Chang and Blackstone 2007; Taguchi et al. 2007; Zunino et al. 2007; Cho et al. 2009). In addition to generating individual organelles, mitochondrial fission was reported to be required for cytochrome c release during apoptosis (Frank et al. 2001; Karbowski et al. 2002; Lee et al. 2004). The proapoptotic Bcl-2 family members Bax and Bak, in turn, were suggested to promote mitochondrial fission (Autret and Martin 2009). Still, the necessity of mitochondrial fission in apoptosis remains controversial (Suen et al. 2008), as, for example, a knock-out mouse lacking the fission factor DRP1 showed no signs of impaired apoptosis (Ishihara et al. 2009). Quality Control of Mitochondria Mitochondria were shown to be strongly fragmented in numerous cases of mitochondrial dysfunction. This can be attributed mainly to the inactivation of the mitochondrial fusion machinery (Herlan et al. 2004; Duvezin-Caubet et al. 2006; Ishihara et al. 2006). Thereby dysfunctional mitochondria may become spatially separated from the intact mitochondrial network, preventing further harm, and the damaged, isolated organelles degraded in a directed manner (Herlan et al. 2004; Skulachev et al. 2004). Indeed, dysfunctional mitochondria were shown to be removed in toto by an organelle-specific autophagic process termed mitophagy (Lemasters 2005; Priault et al. 2005; Kim et al. 2007; Goldman et al. 2010). Moreover, mitophagy in mammalian cells was shown to depend on the fission factor DRP1, further supporting the role of mitochondrial dynamics in the selective removal of dysfunctional mitochondria (Twig et al. 2008). Mitophagy depends on the general

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autophagic machinery but also requires several specific factors that direct the degradation of mitochondria (Kanki and Klionsky 2010), so far studied in baker ’s yeast: Uth1, Aup1, Atg11, Atg21, Atg24, Atg32, Atg33 (Kissova et al. 2004; Tal et al. 2007; Kanki and Klionsky 2008; Okamoto et al. 2009; Kanki et al. 2009a, b); and in mammalian cells: DRP1 (Twig et al. 2008), NIX (Schweers et al. 2007; Sandoval et al. 2008; Novak et al. 2010), Parkin (Narendra et al. 2008), and PINK1 (Dagda et al. 2009). The two latter proteins are involved in Parkinson’s disease, pointing to a role of defective mitophagy in this human neuropathy (Geisler et al. 2010; Narendra et al. 2010). Certainly, other control mechanisms apply as well. Misfolding and aggregation of proteins is minimized by chaperones and numerous proteases present in the different mitochondrial subcompartments (Voos 2009). Furthermore, severely and irreversibly damaged mitochondria may initiate programmed cell death and thus be eliminated at the cellular level. In general, ensuring functionality of mitochondria occurs at different levels, and we still only begin to understand the underlying mechanisms (Tatsuta and Langer 2008).

Crosstalk between Mitochondria and the ER Next to its previously mentioned role in lipid metabolism, the intimate physical relation of mitochondria with the ER (see also Chapters 7 and 11) governs critical functions in calcium homeostasis, signaling, and regulation of the apoptotic cell death (Pizzo and Pozzan 2007; Romagnoli et al. 2007; Pinton et al. 2008; Giorgi et al. 2009; Hayashi et al. 2009; Rizzuto et al. 2009). In contrast to the initially assumed role of mitochondria as mere buffers for cytosolic calcium excesses, the stream of calcium ions into the mitochondrial matrix appears to act as an important regulator of oxidative phosphorylation (Denton 2009; Griffiths and Rutter 2009). The massive and prolonged accumulation of Ca2+ within mitochondria, however, induces the opening of a large conductance channel, the so-called permeability transition pore, in the IM (Lemasters et al. 2009), swelling of the organelle that leads to the rupture of the outer membrane, and the concomitant release of several proapoptotic factors into the cytosol, ultimately resulting in cell death (Kroemer et al. 2007). While it is a known fact that up to 20% of the mitochondrial surface is in close contact with the ER (Rizzuto et al. 1998), the macromolecular complexes that tether the two membranes at a distance of 10–25 nm (Csordas et al. 2006) are scarcely characterized. Two recent studies indicated that several proteinaceous bridges redundantly link the mitochondrial with the ER (de Brito and Scorrano 2008; Kornmann et al. 2009). Future studies will have to elucidate the organization and dynamic regulation of this structural entity, and to decipher whether the various tethers fulfill different functions within the mitochondria– ER communication.

Mitochondrial Disease Based on the intricate role that mitochondria play in cellular physiology, dysfunction of literally any process involving mitochondria results in a pathologic condition (Duchen 2004; Schapira 2006; McFarland et al. 2007). The classical mitochondrial diseases are linked to primary defects within the process of oxidative phosphorylation (Fosslien 2001; Smeitink et al. 2001; Smeitink et al. 2006; Kucharczyk et al. 2009), which can result from either mtDNA mutations (Fig. 6.3; Leonard and Schapira 2000a) or nuclear gene defects (Leonard and Schapira 2000b). However, disruptions of any other mitochondrial function (Jacobs 2003; Fadeel and Orrenius 2005; Hauff and Hatch 2006; MacKenzie and Payne 2007; Copeland 2008; Rouault and Tong 2008; Scaglia and Wong 2008; Liesa et al. 2009;

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Moczulski et al. 2009; Zutz et al. 2009), which might entail a secondary impairment of the energy metabolism (McFarland and Turnbull 2009), result in a diverse set of disorders in multiple organ systems (Finsterer 2006). Thus, the consideration given to mitochondria in regard to disease pathogenesis has tremendously widened in recent years (Wallace 2005b; McFarland et al. 2007; DiMauro and Schon 2008; Wallace and Fan 2009). As the list of mitochondria-associated diseases includes diabetes (Friederich et al. 2009; Schiff et al. 2009), Alzheimer ’s disease (Moreira et al. 2007; Reddy 2009), Parkinson’s disease (Thomas and Beal 2007; Banerjee et al. 2009), and cancer (Wallace 2005a; Rustin and Kroemer 2007; Gogvadze et al. 2008), interest in mitochondria as therapeutic targets is constantly growing (Armstrong 2007; Henchcliffe and Beal 2008; Kroemer and Pouyssegur 2008; Koene and Smeitink 2009; Zorzano et al. 2009; Moreira et al. 2010; Wallace et al. 2010). In the long run, however, a necessary prerequisite to making these efforts a success will be a more thorough understanding of the complex entanglements of mitochondria with all aspects of the biology of eukaryotic cells. If this can be achieved, the modulation of mitochondrial functions holds promise to become an effective therapeutic strategy in the amelioration of symptoms in a broad range of diseases.

ACKNOWLEDGMENTS We are very grateful to Drs. L. Plecitá-Hlavatá, P. Ježek, S. Jakobs, S.W. Hell, and G.A. Perkins for providing high-quality images of their excellent work. This work was supported by the University of Munich FöFoLe program (MZ), the Deutsche Forschungsgemeinschaft SFB 594

project B8, and the Cluster of Excellence “Macromolecular Complexes” at the Goethe University Frankfurt DFG project EXC 115 (AR). We apologize to all colleagues whose excellent research could not be mentioned due to space constraints.

ABBREVIATIONS ADP ATP CJ CM

adenosine-5′-diphosphate adenosine-5′-triphosphate crista junction crista membrane

IBM inner boundary membrane IM inner membrane MitoPLD mitochondrial phospholipase D OM outer membrane

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fission: an opportunity for drug discovery? Curr Opin Drug Discov Devel 12:597–606. Zuchner S, Mersiyanova IV, Muglia M, BissarTadmouri N, Rochelle J, Dadali EL, Zappia M, Nelis E, Patitucci A, Senderek J, Parman Y, Evgrafov O, Jonghe PD, Takahashi Y, Tsuji S, Pericak-Vance MA, Quattrone A, Battaloglu E, Polyakov AV, Timmerman V, Schroder JM, Vance JM. 2004. Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat Genet 36:449–51. Zunino R, Schauss A, Rippstein P, Andrade-Navarro M, McBride HM. 2007. The SUMO protease SENP5 is required to maintain mitochondrial morphology and function. J Cell Sci 120:1178–88. Zutz A, Gompf S, Schagger H, Tampe R. 2009. Mitochondrial ABC proteins in health and disease. Biochim Biophys Acta 1787:681–90.

FURTHER READING A valuable number of excellent monographs covering the diverse facets of mitochondrial biology in greater detail are available. We would like to suggest a few of them to the readers that became captivated by

mitochondria and intend to widen and deepen their knowledge about this versatile and fascinating organelle.

General Lane N. 2006. Power, sex, suicide: mitochondria and the meaning of life. Oxford, UK: Oxford University Press. Sczheffler IE. 2008. Mitochondria, 2nd ed. Hoboken, NJ: Wiley-Liss. Nicholls DG, Ferguson SJ. 2002. Bioenergetics 3, Amsterdam, the Netherlands; London, UK: Academic Press.

Schaffer SW, Suleiman MS. 2007. Mitochondria: the dynamic organelle. Advances in biochemistry in health and disease 2. New York: Springer. Chadwick D, Goode J. 2007. Mitochondrial biology: new perspectives. London, UK: John Wiley.

Origin of Mitochondria and Related Organelles Martin WF, Müller M. 2007. Origin of mitochondria and hydrogenosomes. Berlin, Germany; New York: Springer.

Tachezy J. 2008. Hydrogenosomes and mitosomes: mitochondria of anaerobic eukaryotes. Microbiology Monographs 9. Berlin, Germany: Springer.

Mitochondria and Disease Holt IJ. 2003. Genetics of mitochondrial diseases. Oxford, UK: Oxford University Press. Berdanier CD. 2005. Mitochondria in health and disease. Oxidative stress and disease 16. Boca Raton, FL: CRC Press. Gibson GE, Ratan RR, Beal MF; New York Academy of Sciences. 2008. Mitochondria and oxidative stress

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in neurodegenerative disorders. Boston; Oxford, UK: Blackwell. Lemasters JJ, Nieminen A-L. 2001. Mitochondria in pathogenesis. New York; London, UK: Kluwer Academic Publishers. Singh KK, Costello L. 2008. Mitochondria and cancer. New York: Springer.

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Experimental Approach to Mitochondria Pon LA, Schon EA; American Society for Cell Biology. 2007. Mitochondria, 2nd ed. Methods in Cell Biology 80. San Diego, CA: Academic Press. Leister D, Herrmann J. 2007. Mitochondria: practical protocols. Methods in Molecular Biology 372, Totowa, NJ: Humana Press.

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Stuart JA. 2009. Mitochondrial DNA: methods and protocols, 2nd ed. Methods in Molecular Biology 554. Totowa, NJ: Humana Press.

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7

THE ENDOPLASMIC RETICULUM Jody Groenendyk Marek Michalak

DEFINITION OF THE DOMAIN The endoplasmic reticulum (ER) is an intracellular membrane organelle that forms a network of tubules, vesicles, and cisternae within the cell, and is interconnected with the nuclear envelope that surrounds the nucleus. The ER is a multifunctional organelle responsible for the synthesis and facilitation of intracellular transport of membrane-associated and secreted proteins, lipid and steroid synthesis and transport, Ca2+ signaling, communication with other intracellular organelles including mitochondria and plasma membrane, and stress responses. The ER contains a number of multifunctional integral and resident molecular chaperones and folding enzymes responsible for quality control of proteins, posttranslational modifications, regulation of Ca2+ signaling, and homeostasis and control of ER stress. For example, calreticulin, calnexin, and ERp57 play a role in the quality control of newly synthesized glycoproteins. Other enzymes, including oxidoreductases, oligosaccharide transferases, glucosidases, and mannosidases, are responsible for protein modifications such as disulfide bond formation and N-linked glycosylation. ER luminal chaperones are also responsible for the regulation of the ER stress response and ERassociated degradation (ERAD). ER resident chaperones are major ER luminal Ca2+ buffers, which, together with the inositol-1,4,5-trisphosphate (InsP3) receptor/Ca2+ channel, the sarcoplasmic/ER Ca2+ ATPase (SERCA), and the stromal-interacting molecule 1 (STIM1), a sensor of ER luminal Ca2+, are critical for the regulation of Ca2+ signaling in cells. The ER is also one of the most sophisticated sensors of cellular stresses evoked by environmental or intracellular signals. The membrane has developed a network of integral membrane kinases and resident proteins to regulate ER stress responses. Therefore, the ER is able to contribute during cellular adaptation to many different challenges, which include environmental, metabolic, and intrinsic demands as well as developmental pathways.

INTRODUCTION AND HISTORICAL PERSPECTIVES The ER was first visualized using electron microscopy, by staining Purkinje cells and muscle using heavy metals or basic dyes in the late nineteenth century and has since been found to be present in nearly all cells of higher plants and animals. In 1945, using electron microscopy techniques, Keith Porter and collaborators noted the presence of a “lacelike” structure in cells grown in culture and termed this structure the ER (Porter et al. 1945;

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Palade 1956). Further examination recognized the ER as the organelle responsible for protein production by the discovery of ribosome-studded portions that corresponded to areas of high biosynthetic activity (Palade and Porter 1954; Palade 1955). Electron microscopy studies have identified the ER as a continuous, dynamic, interconnected membrane system containing microenvironments associated with specific functions and that interacts with other organelles within the cell (Sabatini et al. 1963; Song et al. 1991). The concept of the ER as an important intracellular Ca2+ store was proposed by Heilbrunn over 60 years ago (Heilbrunn and Wiercinsky 1947). Later, the concept of Ca2+ transporters (SERCA) and Ca2+-induced Ca2+ release was developed (Franzini-Armstrong 1998), followed by the discovery of the receptor-activated, second-messenger-induced intracellular Ca2+ release (Nielsen and Petersen 1972; Case and Clausen 1973). Seminal experiments a decade later by Berridge and his colleagues showed that this alternative pathway for intracellular Ca2+ release is mediated by the second messenger InsP3 (Streb et al. 1983). The ER Ca2+ release channels were molecularly characterized revealing two main families: the Ca2+-gated and InsP3-gated channels. Discoveries of ER signaling (Horton et al. 2002; BengoecheaAlonso and Ericsson 2007) and ER stress (Malhotra and Kaufman 2007) pathways further reinforced the distinctive nature of this fascinating intracellular organelle. The focus of this chapter is on the structure and function of the ER, with a major emphasis on its role as the entry site for the secretory pathway and a modulator of Ca2+ homeostasis. Sarcoplasmic reticulum (SR), a muscle-specific ER, is discussed here in context of ERdependent regulation of Ca2+ homeostasis. Many excellent reviews on the structure and function of the SR have been published (Mazzarello et al. 2003; Treves et al. 2009) including the latest commentary on the ER and SR in cardiac muscle (Michalak and Opas 2009).

STRUCTURE AND DYNAMICS OF THE ER Today, the ER is recognized as a heterogeneous and dynamic membrane system containing specific microdomains involved in protein folding, transport, and Ca2+ signaling (Papp et al. 2003). There are three main areas of the ER: the rough ER, the smooth ER, and the nuclear envelope. The rough ER is characterized by association with ribosomes, which are responsible for supporting translation of mRNA into protein and targeting the mRNA to a translocon located in the ER membrane. The smooth ER is where the ER cargo is exported and is characterized by the presence of coat protein complex (COP) II coatomer and cargo receptors such as the ER–Golgi intermediate compartment 53 (ERGIC-53) (Hobman et al. 1998). Smooth ER might also play an important role in the control of cellular Ca2+ homeostasis. The third subdomain, the nuclear envelope, is responsible for transit to the nucleus of Ca2+ and proteins involved in gene expression and will not be discussed here (see also Chapter 24 on the nuclear pore). One critical part of the ER structure is its dynamic capacity, continually moving and shaping via the action of motor proteins attached to the microtubule network (WatermanStorer and Salmon 1998; Wozniak et al. 2009) (see also Chapter 14 on microtubules). In cells that are treated with a microtubule inhibitor such as nocodazole, or by downregulation of kinesin, a motor protein, motility of the ER structures is blocked (Wozniak et al. 2009). As well, a direct interaction between ER tubules and growing microtubule plus ends via tip attachment complexes has recently been identified, with these contacts generated by the interaction of the microtubule plus-end-binding protein, EB1 with an ER resident membrane protein, STIM1 (Grigoriev et al. 2008). Another motor protein, dynein, is involved in the movement of ER tubules in the reverse direction (Lane and Allan 1999). Blockage of kinesin slows cargo exit from the ER, while loss of microtubules results in

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lateral mobility of translocon machinery (Nikonov et al. 2007; Gupta et al. 2008). Cytoskeleton-linking membrane proteins (CLIMPs) and cytoplasmic linker proteins (CLIPs) link the ER membrane to the microtubule network via tubulin-binding domains (Rickard and Kreis 1996; Klopfenstein et al. 1998). Ankyrins are directly involved in restricting specific proteins (including the ryanodine receptor [RyR], InsP3 receptor, and SERCA) to specialized regions of the ER (Tuvia et al. 1999). These discrete subdomains may be critical for coordination and optimization of global ER function and interaction with other organelles or membranes (Baumann and Walz 2001).

ER PROTEOME Following the recent wave of total genome sequencing analysis, research has turned to understanding the functions, modifications, and regulation of all proteins encoded by a genome (Au et al. 2007). Large-scale proteomics technologies and tools have made organelle-scale studies feasible and have become powerful methods for studying organelles, including the ER and their components and dynamics (Yates et al. 2005). Proteomics is the identification of the total protein complement or proteome of various organelles and cells. The basic rationale of proteomics research is to integrate the information obtained from analysis into a comprehensive proteome database that contains protein identities and functional characteristics. Subcellular fractionation followed by two-dimensional gel electrophoresis combined with peptide analyses using matrix-assisted laser desorption/ ionization mass spectroscopy and tandem mass spectrometry provides a sensitive analytical tool that identifies proteins. These techniques have been used successfully to determine the protein composition of subcellular structures including the luminal ER (Knoblach et al. 2003; Gilchrist et al. 2006), trafficking vesicles (Takamori et al. 2006), and specialized cell types such as platelets (O’Neill et al. 2002). Proteomic analysis of the ER has identified over 400 proteins associated with the ER, the majority belonging to functional categories including ribosomal proteins, translocon subunits, chaperones, secretory proteins, and glycolipid processing enzymes (Knoblach et al. 2003; Gilchrist et al. 2006; Chen et al. 2010). The ER cargo proteins are another important class of proteins identified, which move through the ER via the secretory pathway, an example being transferrin, an ironbinding protein; a number of secreted pheromone-binding proteins termed major urinary proteins (MUPs); and apolipoprotein A, which transports cholesterol from tissues to the liver (Knoblach et al. 2003). Additional classes of identified proteins are involved in other vital functions of the ER, including membrane trafficking, Ca2+ storage, and oligosaccharide biosynthesis. Table 7.1 shows a list of selected ER-associated proteins identified by proteomic techniques. An important benefit of doing organellar proteomics is the identification of novel proteins, such as two protein disulfide isomerase (PDI)-like proteins, ERp19 and ERp46 (Knoblach et al. 2003), as well as sapreticulin involved in sphingolipid degradation (Gilchrist et al. 2006). The real power of proteomics analysis of the ER will be realized when comprehensive and comparative analyses of the ER proteome are carried out in different cell types, under different physiological states, and in normal and diseased cells.

ER: A MULTIFUNCTIONAL ORGANELLE The ER is a multifunctional intracellular organelle influencing virtually every aspect of cellular function (Baumann and Walz 2001; Sitia and Braakman 2003). The ER is the entry

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TABLE 7.1. Selected Proteins Identified from Endoplasmic Reticulum Proteomic Analysis (Knoblach et al. 2003; Gilchrist et al. 2006; Chen et al. 2010)

Protein Name Spectrin alpha chain Clathrin heavy chain Protein disulfide isomerase (PDI) precursor 78-kDa glucose-regulated protein precursor glucose-regulated protein (GRP) 78 Transferrin Glutamate dehydrogenase, mitochondrial precursor Eukaryotic translation elongation factor 2 Cytochrome P450 2D2 Myosin heavy chain myr 4 ERp46 Calreticulin precursor Calnexin precursor 60S ribosomal protein L4 Endoplasmic reticulum–Golgi intermediate compartment 53 (ERGIC-53) protein precursor Carboxylesterase 3 precursor Pyruvate carboxylase, mitochondrial precursor Tubulin β2 Polymeric immunoglobulin receptor precursor Annexin A5 Endoplasmic reticulum protein 29 Golgi synaptosomal associated protein (SNAP) receptor complex member 1 Major urinary protein (MUP) precursor Low-density lipoprotein receptor-related protein-associated protein 1 Apolipoprotein A Chloride intracellular channel protein 4 N-cadherin Ras-related protein Rab-18 Signal recognition particle receptor subunit beta Endoplasmic reticulum α-1,2-mannosidase ERp19

Function Cytoskeleton Coatomer PDI family Chaperone Secretion Mitochondria Translation Mitochondria Cytoskeleton PDI family Chaperone Chaperone Translation Secretion Lipid metabolism Mitochondria Cytoskeleton Cargo Cytoskeleton PDI family Secretion Secretion Cargo Cargo Cargo Cytoskeleton Signaling Transcription Oligosaccharide PDI family

point of the secretory pathway, with the ER environment containing a high concentration of protein (∼300 mg/mL), having an oxidizing environment and high in Ca2+ concentration, similar to the outside of the cell. As one-third of newly synthesized proteins traverse the ER, homeostasis of the ER secretory pathway is mandatory and is maintained by entrance of nascent polypeptides balanced by exit of either correctly folded proteins or degradation of misfolded proteins. The ER is the site for specific protein modifications such as addition of N-linked glycosylation, disulfide bond formation, and attachment of glycosylphosphatidylinositol (GPI) anchors. If this secretory pathway is disrupted in any manner, ER stress results due to a buildup of misfolded protein, triggering a cellular response termed the unfolded protein response (UPR). In conjunction, ERAD is initiated in an attempt to recover homeostasis, and subsequently, the apoptotic pathway is activated. Not surprisingly, there are a variety of pathological conditions linked to a breakdown in ERassociated processes, including vascular diseases such as myocardial infarction and atherosclerosis (Brooks 1997, 1999; Jakob et al. 2001; Sherman and Goldberg 2001; Graf

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TABLE 7.2.

Protein

Knockout Mouse Models and Associated Diseases

Knockout Mouse Model

BiP/GRP78

GRP94 ERp57

Embryonic lethal at E3.5 • Failure of embryo to periimplant Embryonic lethal Embryonic lethal at E13.5 • Disruption in Stat3 signaling • Disruption in major histocompatability complex (MHC) Class I loading (ERp57 conditional knockout) Not done

PDI

117

Calnexin

Not embryonic lethal • Postnatal motor disorders

Calreticulin

Embryonic lethal at E14.5 • Defective cardiac development

UGGT

Embryonic lethal at E13

Diseases

References

Cancer, Alzheimer ’s disease, Parkinson’s disease, prion diseases, atherosclerosis

Luo et al. (2006); Hoshino et al. (2007)

Cancer, prion diseases, autoimmune diseases Prion diseases, Alzheimer ’s disease, autoimmune diseases

Wanderling et al. (2007) Coe et al. (pers. comm.); Garbi et al. (2006); Solda et al. (2006)

Alzheimer ’s disease, Parkinson’s disease Alzheimer ’s disease, cystic Denzel et al. (2002); fibrosis, Charcot–Marie– Kraus et al. (pers. comm.) Tooth disease Cardiac hypertrophy, Mesaeli et al. (1999) Alzheimer ’s disease, autoimmune diseases None associated Molinari et al. (2005)

et al. 2004); neurodegenerative diseases, such as Alzheimer ’s disease, Creutzfeldt–Jakob disease and Charcot–Marie–Tooth disease (Jeffery et al. 2000; Soti and Csermely 2002; Shames et al. 2003; Rutkowski and Kaufman 2004); systemic diseases, such as cystic fibrosis and familial hypercholesterolemia (Chevet et al. 1999; Sorensen et al. 2006); as well as a number of cancer conditions, including malignant melanoma and breast cancer (Li et al. 2001; Dissemond et al. 2004) (Table 7.2).

Protein Synthesis, Folding, and Quality Control Folding within the ER involves three classes of proteins termed molecular chaperones, folding enzymes, and folding sensors. Molecular chaperones reduce the rate of protein folding by transiently interacting with substrates and increasing the efficiency of folding. Folding enzymes accelerate the rate-limiting steps of the folding process, such as disulfide bond formation, while folding sensors detect misfolded protein and interact with exposed hydrophobic domains and send the unfolded protein back into the folding cycle. For example, the molecular chaperones BiP/GRP78 and GRP94 recognize exposed hydrophobic regions assisting in the folding and assembly of the nascent protein (Argon and Simen 1999; Gething 1999), while calreticulin and calnexin interact with glycoproteins via their lectin-binding ability (Helenius et al. 1997). The folding enzymes PDI and ERp57, both thiol oxidoreductases, utilize the oxidizing environment of the ER to generate disulfide linkages (Oliver et al. 1997; Molinari and Helenius 1999; Noiva 1999), with the formation of these intra- and interchain disulfide bonds, an integral part of the maturation of most secretory and membrane-bound proteins in the ER. The folding sensor, uridine diphosphate (UDP)-glucose : glycoprotein glucosyltransferase (UGGT), recognizes unfolded proteins that are glycosylated and sends them back into the quality control cycle (Taylor et al. 2003). These chaperones and folding enzymes share a specific retention and

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retrieval signals responsible for their retention in the ER. ER luminal resident proteins contain a C-terminal amino acid sequence of KDEL (Lys-Asp-Glu-Leu) or its variant (Nilsson and Warren 1994) and are retrieved by a receptor-mediated process (Michelsen et al. 2005). ER integral membrane proteins contain a KKXX (Lys-Lys-X-X) C-terminal amino acid sequence (Griffiths et al. 1994; Nilsson and Warren 1994) and utilize a COPdependent pathway for retention (Nilsson and Warren 1994). Carbohydrate modification of newly synthesized proteins plays an important role in protein folding and posttranslational modification (Helenius and Aebi 2004) (see Chapter 8 for discussion of glycosylation in the Golgi apparatus). As nascent glycoproteins are translocated into the lumen of the ER, they associate with Ca2+-dependent molecular chaperones (Helenius and Aebi 2004) responsible for appropriate folding (Molinari and Helenius 2000). The proteins utilize the quality control machinery, which includes the lectin chaperones calreticulin and calnexin, oxidoreductase ERp57, glucosidases, mannosidases, and UGGT (Helenius and Aebi 2004; Hebert and Molinari 2007) (Fig. 7.1).

Ribosome Oligosaccharide-DP

Flippase

OST

CNX

ERp57

ERp57

CRT

Glucosidase II

UGGT

α-Mann I

BiP α-Mann I PDI

ER lumen

GRP94

ERAD Transit of correctly folded protein

UPR

Figure 7.1. Endoplasmic reticulum quality control. The oligosaccharide is generated in the cytoplasm and is linked to a membrane-bound anchor, dolicholphosphate (DP), followed by flipping to the lumen of the ER, where it is linked to the N in the specific sequence, NXS/T of a nascent protein by the enzyme oligosaccharyltransferase (OST). This nascent protein then associates with the quality control apparatus, including calnexin (CNX), calreticulin (CRT), ERp57, and UDP-glucose : glycoprotein glucosyltransferase (UGGT). These enzymes and chaperones cooperate to fold nascent glycoproteins. If this protein is not able to fold after repeated cycles of quality control, it undergoes cleavage of mannose residues by the α-mannosidase I (α-mann I) and is targeted for ERAD. As well, the accumulation of misfolded protein triggers the UPR upon the interaction of BiP/GRP78 with the misfolded protein. Once the protein is folded properly, it exits the ER and transits to other locations within the cell. PDI, protein disulfide isomerase.

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Nonglycosylated proteins associate with BiP/GRP78 and GRP94 molecular chaperones and PDI folding enzyme (Ni and Lee 2007) (Fig. 7.1). BiP/GRP78 utilizes the energy of ATP to assist in folding the nascent protein and plays a role as a central protein in the lumen of the ER by performing numerous functions including buffering Ca2+, functioning as a permeability barrier via physically blocking of the translocon pore, ratcheting the nascent protein through the pore, binding hydrophobic domains and folding nascent proteins, sensing and binding misfolded proteins that are exposing hydrophobic regions, and regulating the UPR and ERAD (Ni and Lee 2007). The molecular chaperone GRP94 interacts with partially folded polypeptides after BiP/GRP78, in an ATP-dependent manner with the hydrolysis of ATP a necessary part of the folding process (Ostrovsky et al. 2009). Calnexin and calreticulin, two members of the quality control cycle, interact with partially folded polypeptides by binding immature N-linked oligosaccharides. A core carbohydrate containing two N-acetylglucosamine residues, nine mannose residues with three terminal glucose residues is assembled and transferred onto an asparagine amino acid of the nascent polypeptide chain. As soon as the oligosaccharide is added, trimming begins. Three terminal glucoses are trimmed by glucosidase I and II, and a terminal mannose is trimmed by one or more ER mannosidases. The binding of calreticulin and calnexin to their substrates is mediated, at least in part, by a lectin site that recognizes the N-linked oligosaccharide processing intermediate Glc1Man9GlcNAc2 (Leach et al. 2002). This serves to prevent aggregation, protect proteins from premature degradation, and assure the correct folding status of proteins before continuing in the intracellular trafficking pathway. There is evidence that calnexin and calreticulin also recognize the polypeptide segments of the glycoproteins as well (Swanton et al. 2003; Williams 2006). Binding of substrates to calreticulin or calnexin is terminated by removal of the terminal glucose by glucosidase II. If the protein is not correctly folded, it can be reglucosylated by UGGT, allowing reassociation with calreticulin and calnexin. Finally, a terminal mannose is trimmed via an ER-localized α-1,2-mannosidase in a time-dependent manner (Fig. 7.1). Mannose trimming allows discrimination between unfolded and terminally misfolded glycoproteins. Misfolded proteins are recognized by part of the degradation machinery, the ER degradation-enhancing 1,2-mannosidase-like protein (EDEM), a membrane protein that selectively trims mannose glycans on misfolded proteins for recognition by the ERAD lectin, osteosarcoma-9 (OS-9), which targets the misfolded protein for retrotranslocation and degradation (Lederkremer 2009; Aebi et al. 2010). This quality control cycle serves as an efficient retention method, as unfolded glycoproteins can be reglucosylated by the unfolding sensor UGGT and interact with calreticulin or calnexin again until proper folding occurs. This cycle may occur numerous times until the protein is folded correctly or, if the quality control is unable to fold the protein, these misfolded and unfolded proteins accumulate and result in a variety of signaling pathways being activated to control the ensuing ER stress, including ERAD and UPR (Fig. 7.1). An important component of protein folding is the generation and isomerization of disulfide bonds, which assist in protein folding by lowering the number of available conformations that the protein can assume, as disulfide linkages physically limit the protein. Calnexin and calreticulin also present polypeptides for disulfide bond formation to a member of the PDI family, ERp57, a folding enzyme that directly binds to calnexin and calreticulin (Oliver et al. 1999) (Fig. 7.1). ERp57 is one of many PDI-like oxidoreductases associated with the ER (Elliott et al. 1997). PDI is an abundant and versatile oxidoreductase that has oxidase, reductase, and isomerase activity, and interacts with a variety of substrates (Edman et al. 1985). PDI contains a thioredoxin active site, composed of two cysteines separated by two other amino acid residues (the CXXC motif), as well as the typical fold of thioredoxin (Ferrari and Söling 1999). The protein also works as a molecular

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chaperone by binding polypeptides and preventing protein aggregation (Freedman et al. 1994). The combination of redox and chaperone activities is what makes PDI an effective and important enzyme in oxidative protein folding (Sitia and Molteni 2004). ERp57 differs from PDI by catalyzing disulfide bond formation almost exclusively in glycoproteins in conjunction with calreticulin and calnexin (Oliver et al. 1997, 1999). Since disulfide bonds increase the stability of the native conformation, their absence or mispairing produces severely misfolded species, resulting in protein aggregation.

ERAD ERAD is a process by which misfolded ER proteins are detected and removed from the secretory pathway by ER-resident factors and directed to translocation machinery for retrotranslocation into the cytoplasm, where they undergo ubiquitin- and proteasomedependent degradation. ERAD is triggered when these misfolded proteins are targeted by the ER α-1,2-mannosidase I, which cleaves one or more mannose residues from the middle branch of the oligosaccharide (Frenkel et al. 2003). Trimming of these mannose residues effectively prevents reglucosylation by UGGT, thereby disrupting any interaction with calnexin in an attempt to refold the protein (Eriksson et al. 2004). Mannose-trimmed oligosaccharides are specifically recognized by EDEM (Spiro 2004), which accepts the protein from calnexin (Oda et al. 2003) and selectively trims further mannose glycans on the misfolded protein for recognition by the ERAD lectin, OS-9, which targets the misfolded protein for retrotranslocation and degradation (Hosokawa et al. 2001; Jakob et al. 2001; Oda et al. 2006; Aebi et al. 2009; Lederkremer 2009). The misfolded protein is unfolded and retrotranslocated into the cytoplasm, poly-ubiquitinated, and subsequently targeted to the 26S proteasome for degradation (Tsai et al. 2002). The retrotranslocation machinery, named Derlin1, in complex with cytoplasmic valosin-containing protein (VCP) also termed AAA ATPase p97 (Ye et al. 2005), and N-glycanase, which removes the oligosaccharide (Katiyar et al. 2005), appears to be distinct from the Sec61 translocon (Lilley and Ploegh 2004, 2005; Ye et al. 2004). Derlin1 transfers the misfolded polypeptide from the lumen of the ER to the cytoplasm for degradation by the proteasome, with this machinery an essential constituent of ERAD (Oda et al. 2006).

ER Ca2+ Homeostasis: Ca2+ Buffers As the ER is the major physiological source of Ca2+ within the cell, it plays an instrumental role as a signal-transducing organelle that responds to environmental cues that trigger intracellular Ca2+ signaling, as well as maintaining Ca2+ homeostasis (Berridge 2002; Michalak et al. 2002). Changes in Ca2+ homeostasis within the ER as well as Ca2+ released from the ER activates transcriptional and translational cascades that ultimately regulate chaperones responsible for protein folding, ER stress, UPR, and ERAD, as well as proteins involved in the apoptotic pathway (Berridge 2002; Breckenridge et al. 2003). Ca2+ homeostasis and signaling are maintained by controlling Ca2+ release from the ER by the InsP3 receptor (Patel et al. 1999) and RyR (Franzini-Armstrong and Protasi 1997), with the stores being refilled by SERCA (Ashby and Tepikin 2001) (Fig. 7.2). The ER Ca2+ stores not only serve as a source of easily accessible Ca2+ but are also important as a regulator of a number of ER enzymes and proteins including feedback regulation of the InsP3 receptor, the RyR, and SERCA. Localized to the lumen of the ER are proteins involved in Ca2+ signaling and buffering, affecting numerous aspects of ER function (Bergeron et al. 1994; Williams and Watts 1995; Meldolesi and Pozzan 1998; Nauseef 1999; Corbett and

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Ca

InsP3R

CSQ

CRT

PM

ERp57

BiP PDI

SERCA ERp57

GRP94

CRT

ORAI1 Stim1

ERp72 ERcalcistorin ER lumen

Figure 7.2. Endoplasmic reticulum Ca2+ homeostasis. The ER contains the greatest concentration of Ca2+ within the cell and maintains this concentration through the efforts of SERCA, an ATPase pump, and InsP3R, a Ca2+ transporter. Ca2+ is buffered in the lumen of the ER through the actions of calreticulin, PDI, BiP/GRP78, GRP94, calsequestrin (CSQ), ERp72, and ERcalcistorin. ER Ca2+ stores once released are refilled with STIM1 sensing Ca2+ concentration and triggering opening of the plasma membrane (PM) transporter, termed ORAI1, which increases cytoplasmic Ca2+ and allows refilling of ER Ca2+ stores. CRT, calreticulin; PDI, protein disulfide isomerise.

Michalak 2000; High et al. 2000; Molinari and Helenius 2000; Baumann and Walz 2001; Jakob et al. 2001). Ca2+-buffering proteins are necessary as the total Ca2+ concentration of the ER is in the micromolar to millimolar range. Stringent maintenance of low ER luminal free Ca2+ concentrations (below 300 μM) via Ca2+-buffering proteins is important for the preservation of cellular integrity. Many of the ER Ca2+-buffering proteins have a high Ca2+-binding capacity (>10 mol 2+ of Ca per mole of protein) coupled with low affinity (Kd = 1 mM or higher), while others have low capacity (1–2 mol of Ca2+ per mole of protein) but high affinity (Kd = 1 μM). Calreticulin utilizes an acidic region at its C-terminus as a high-capacity Ca2+ binding site, with 43 acidic residues in the last 82 amino acids of the protein, to bind 25 mol of Ca2+ per mole of protein with low affinity (Kd = 2 mM) (Baksh and Michalak 1991). Calreticulin is responsible for buffering approximately 50% of ER luminal Ca2+ (Nakamura et al. 2001). GRP94, like calreticulin, contains an acidic C-terminal region, which comprises its

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low-affinity Ca2+ binding site. It contains approximately 19 Ca2+ binding sites, 4 of which have high affinity (Kd = 2 μM) and low capacity (1 mol of Ca2+ per mole of protein) and 15 of which have low affinity (Kd = 600 μM) but high capacity (10 mol of Ca2+ per mole of protein) and are composed of negatively charged amino acids (Van et al. 1989). While BiP/GRP78 has a relatively low capacity for binding Ca2+ (1–2 mol of Ca2+ per mole of protein), it contributes possibly as much as 25% of the total Ca2+ buffering capacity of the ER, and elevated expression causes an appreciable increase in ER Ca2+ storage capacity (Lievremont et al. 1997) (Fig. 7.2). ER resident oxidoreductases, a PDI-like family of proteins, also contribute to Ca2+ buffering in the lumen of ER. PDI utilizes pairs of acidic residues that constitute the highcapacity Ca2+ binding site (approximately 20 mol of Ca2+ per mole of protein) with low affinity (Kd = 2–5 mM) (Lebeche et al. 1994). ERcalcistorin/PDI is a luminal 58-kDa calsequestrin-like high-capacity, low-affinity Ca2+-binding protein, binding 23 mol of Ca2+ per mole of protein with low affinity (Kd = ∼1 mM) (Lucero and Kaminer 1999). ERp72 is a luminal member of the PDI family that binds Ca2+ with high capacity (Lucero et al. 1998). Calsequestrin is a unique and muscle-specific Ca2+-buffering protein found in the lumen of the SR membrane that, in conjunction with triadin and junctin, regulates the RyR (MacLennan 1974). Calsequestrin binds approximately 50 mol of Ca2+ per mole of protein with low affinity (Kd = 1 mM) (Beard et al. 2004) (Fig. 7.2). Ca2+-buffering proteins in the ER are necessary to maintain intracellular Ca2+ homeostasis.

ER Ca2+ Homeostasis: Ca2+ Transport Ca2+ release from the ER is triggered by agonist-driven, InsP3-dependent Ca2+ release via InsP3 receptor. The resulting increase in cytoplasmic Ca2+ concentration activates numerous Ca2+-dependent proteins including calmodulin, calcineurin, and Ca2+/calmodulindependent protein kinase II (CaMKII), which then trigger a downstream signaling cascade. After the release of ER Ca2+ stores into the cytoplasm, the cell has a recovery mechanism to reduce cytoplasmic Ca2+ levels and refill ER Ca2+ stores via action of the SERCA pump. Emptying ER Ca2+ stores leads to activation of store-operated Ca2+ entry (SOCE) (Parekh and Putney 2005; Putney 2007). SOCE is an example of coordination between Ca2+ signals originating from the ER luminal store and the extracellular media (see also Chapter 11 for a discussion of SOCE). It includes ER Ca2+ sensor STIM1 and plasma membrane Ca2+ channel ORAI1 (Fig. 7.2). STIM1 is a 685-amino acid, type I transmembrane ER protein containing two luminal EF hands, a sterile alpha motif (SAM) domain as well as cytoplasmic coiled-coil domains and serine-, proline-, and lysine-rich regions (Dziadek and Johnstone 2007). STIM1 senses the decrease in ER Ca2+ stores via an EF-hand motif (Williams et al. 2001), resulting in oligomerization of STIM1 and complex formation with the plasma membrane Ca2+ channel, ORAI1 (Baba et al. 2006; Feske et al. 2006; Stathopulos et al. 2006; Vig et al. 2006; Zhang et al. 2006) (Fig. 7.2). This interaction stimulates Ca2+ influx from the extracellular milieu, leading to increased cytoplasmic Ca2+ levels and corresponding transit to refill the ER stores. Release of ER Ca2+ does not result in a uniform increase of Ca2+ in all areas of the cytoplasm, but generates areas of high concentration, closely associated with the compact region between the ER and plasma membrane where Ca2+ channels on the plasma membrane open, called Ca2+ microdomains (Parekh 2008; Barritt et al. 2009) (Fig. 7.2) (see also Chapter 11 for further discussion of ER–plasma membrane contacts). The complexity of Ca2+-signaling proteins in these microdomains may account for the specific characteristics of individual Ca2+ signals (Ambudkar et al. 2004; Parekh 2008).

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ER Stress and UPR The ER responds to an increase in unfolded protein and returns the ER to its normal physiological state by several simple adaptive mechanisms. The immediate response includes downregulation of the biosynthetic load of the ER via interruption of protein synthesis on a transcriptional and translational level, with increased clearance of misfolded or aggregated proteins from the ER by upregulating the ERAD machinery. This is followed by upregulation of the folding capacity of the ER through induction of ER resident molecular chaperones and folding enzymes as well as increasing the size of the ER (Menzel et al. 1997). The primary aim of the UPR is to limit damage to the cell by adapting the cell to the situation causing the ER stress. Three resident transmembrane proteins, in combination with the ER molecular chaperones BiP/GRP78 and calreticulin (Bertolotti et al. 2000), are responsible for the reaction to ER stress (Fig. 7.3). The UPR mechanism involves transcriptional activation of chaperones and members of the ERAD by the transcription factor, activating transcription factor 6 (ATF6) (Shen et al. 2002), in conjunction with the ER membrane kinase and endoribonuclease inositol-requiring 1 kinase (IRE1) (Cox et al. 1993), as well as translational repression of protein synthesis by the ER kinase dsRNA-activated protein kinaselike ER kinase (PERK) (Rutkowski and Kaufman 2004; Schroder and Kaufman 2005; Zhang and Kaufman 2006). Instant cessation in protein synthesis triggered by PERK reduces the translocation of nascent proteins into the ER lumen (Schroder and Kaufman 2006). IRE1 and ATF6 mediate the transcriptional activation of genes encoding proteins that increase protein folding, export, and degradation (Ye et al. 2000; Schroder and Kaufman 2006) (Fig. 7.3). As this activity requires chaperone transcription and translation, it follows the translational attenuation with a slight delay to allow recovery in protein translation (Schroder and Kaufman 2006). ATF6 is a transmembrane protein localized to the ER membrane that upon cleavage becomes a potent transcription factor. Under nonstress conditions, BiP/GRP78 interacts with ATF6, masking its Golgi localization signals (Shen et al. 2002), retaining it in the ER, but upon accumulation of unfolded nascent proteins, BiP/GRP78 is sequestered away, binding to the hydrophobic sections of these nascent proteins, resulting in the release and transit of ATF6 to the Golgi where it undergoes cleavage by site 1 and site 2 (S1P and S2P) proteases (Shen et al. 2002). This yields a soluble basic leucine zipper (bZIP) transcription factor that translocates to the nucleus and induces target genes by binding directly to the ER stress response element (ERSE) located upstream of the target gene (Yoshida et al. 1998). Several target proteins that are upregulated include BiP/GRP78, and proteins involved in ERAD, such as the EDEM receptor, and X-box-binding protein 1 (XBP1) mRNA (Mori et al. 1996). Recent evidence has recognized ATF6 as a glycoprotein, with three N-linked oligosaccharides, identified to interact with calreticulin (Hong et al. 2004) (Fig. 7.3). The ATF6 pathway provides a powerful molecular mechanism for cells to deal with increased accumulation of misfolded proteins or other stresses imposed on the ER membrane. In addition, BiP/GRP78 is sequestered away from PERK and IRE1, resulting in their homodimerization and conformational modification that is transmitted across the membrane, leading to activation of their kinase activity (Bertolotti et al. 2000; Schroder and Kaufman 2006). The transmembrane protein, PERK, upon activation, triggers the phosphorylation of the translation initiation factor eIF2α resulting in the inhibition of protein synthesis by sequestering the tRNAmet responsible for initiating the translation of nascent protein (Bertolotti et al. 2000). IRE1 is a type I ER transmembrane protein that contains

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misfolded proteins ER lumen BiP

ATF6

Caspase-12

IRE1a

PERK

TRAF2 ASK1

elF2α-P XBP1

Golgi

XBP1s

JNK pathway

Cleavage of 28S rRNA

Cleaved ATF6 Protein

synthesis ATF4

Activation of genes encoding chaperones and ERAD proteins

Apoptosis

CHOP

Figure 7.3. The unfolded protein response. Upon an accumulation of misfolded protein in the ER lumen, BiP/GRP78 is sequestered away from three transmembrane proteins: ATF6, IRE1, and PERK. ATF6 is transported to the Golgi, undergoes site-specific cleavage by S1P and S2P proteases, releasing an active transcription factor involved in the induction of ER chaperones. IRE1 dimerizes upon release from BiP/GRP78, resulting in activation of ribonuclease activity and cleavage of the mRNA for XBP1, resulting in a spliced product, XBP1s, that translates into an active transcription factor. IRE1 has been found in complex with tumor necrosis factor (TNF) receptor-associated factor 2 (TRAF2) and apoptosis signal-regulating kinase 1 (ASK1), resulting in activation of the c-Jun N-terminal kinase (JNK) pathway, eventually signaling apoptosis. Caspase 12, an ER stress-dependent caspase, forms complexes with TRAF2 to initiate apoptotic events. IRE1 ribonuclease activity is involved in inhibition of protein synthesis by cleavage of ribosomal RNA. PERK, upon loss of interaction with BiP/GRP78, dimerizes and autoactivates, resulting in the phosphorylation of eIF2α and inhibition of protein synthesis. eIF2α activates a transcription factor, ATF4, involved in the induction of apoptosis via C/EBP homologous protein (CHOP) induction.

a serine–threonine kinase module and a C-terminal endoribonuclease domain (Tirasophon et al. 1998). Activation of UPR results in IRE1 homodimerization, autophosphorylation, and activation of its endoribonuclease activity that cleaves 28S rRNA to inhibit protein synthesis as well as splice and activate XBP1 mRNA (Yoshida et al. 2001). Splicing of XBP1 mRNA produces mRNA with an altered reading frame (Yoshida et al. 2001; Calfon et al. 2002). The protein product of this new XBP1 splice variant acts as a transcription factor, binding to genes containing ERSE (Yoshida et al. 2001; Calfon et al. 2002; Lee et al. 2002; Yamamoto et al. 2004). This signal triggers downstream transcriptional upregu-

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lation of BiP/GRP78 and other chaperones to compensate for the amount of unfolded protein (Fig. 7.3).

Mouse Models of ER Protein Deficiency and Clinical Relevance of ER Dysfunction Table 7.2 demonstrates the significant difference in phenotype severity in mouse models deficient of specific ER proteins. A number of ER chaperones, including calreticulin, BiP/ GRP78, GRP94, ERp57, and UGGT are embryonic lethal due to a variety of developmental problems. Others are not embryonic lethal but lead to a number of diseases, such as autoimmune disease with malfunction of the B cells, seen in the ERp57-targeted knockout as well as neuromuscular disease seen in the calnexin-deficient mouse model (Table 7.2). A number of these proteins are linked to human disease as seen in Table 7.3. Impaired function of the ER, especially its protein folding and quality control machinery, leads to many severe pathologies including Parkinson’s disease and cystic fibrosis (Table 7.3). For example, Parkinson’s disease is a neurodegenerative disorder characterized by selective loss of neurons in the brain and results in progressive motor disturbances such as tremors, akinesia, and rigidity. Buildup of the protein parkin, a RING-type E3 ubiquitin protein ligase, is observed to suppress ER stress-induced cell death (Imai et al. 2000). Cystic fibrosis is a hereditary disease affecting the entire body, resulting in progressive disability and early death. The cystic fibrosis transmembrane conductance regulator (CFTR) protein is a highly glycosylated cAMP-activated chloride TABLE 7.3.

Selected Endoplasmic-Reticulum-Related Diseases

Disease Cystic fibrosis Diabetes mellitus Tyrosinase deficiency α1-Antichymotrypsin deficiency α1-Chymotrypsin Tay–Sachs disease Charcot–Marie–Tooth disease β-Amyloid deficiency Atherosclerosis Marfan syndrome Sitosterolemia Hypercholesterolemia Down’s syndrome Scurvy Fabry disease Polycystic kidney disease Creutzfeldt–Jakob disease Alzheimer ’s disease Breast cancer Metastatic melanoma

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Affected Protein Cystic fibrosis transmembrane receptor Insulin receptor Tyrosinase α1-Antichymotrypsin α1-Chymotrypsin β-Hexosaminidase Peripheral myelin protein β-Amyloid Apolipoprotein A Fibrillin Sterol receptor Low-density lipoprotein (LDL) receptor β-Amyloid Collagen α-Galactosidase Polycystins Prion mutation Amyloid/presenilin protein Heregulin/human epidermal growth factor receptor MHCI

Clinical Phenotype Lung disease Diabetes Pigment defect Lung and liver disease Lung disease Neurological defect Neurological disease Neurodegenerative disease Heart disease Connective tissue disorder Metabolic defect Heart/circulatory defect Neurological disorder Connective tissue disorder Circulatory disorder Kidney disease Neurodegenerative disease Neurodegenerative disease Breast cancer Skin cancer

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channel expressed at the apical membrane of epithelial cells. A mutation in the gene encoding this protein results in the autosomal recessive disease cystic fibrosis. One frequent mutation, F508del, causing misfolding of CFTR, is quickly recognized by calnexin, retained in the ER and subsequently targeted for ERAD (Pind et al. 1994; Amaral 2004). ER stress and protein folding are implicated in a number of cancer conditions that specifically trigger ER stress, including hypoxia, glucose deprivation, and acidosis, observed in tumor cells (Ma and Hendershot 2004). Alterations in the UPR are reported in gastric tumors, hepatocellular carcinoma, and breast cancer (Gazit et al. 1999; Song et al. 2001; Shuda et al. 2003; Hosoi and Ozawa 2010). A key therapeutic target appears to be promotion of protein folding. Table 7.3 demonstrates the varied phenotypes of ER-related folding diseases.

CONCLUSION The ER performs many varied functions within the cell including Ca2+ storage, protein folding, quality control, and modification. Ca2+ within the ER is necessary for normal function of the cell and is required for proper signaling that occurs during cell differentiation and growth, tissue biogenesis, and organism embryogenesis. As can be concluded from the multiple functions of the proteins in the ER, disruption of ER Ca2+ homeostasis not only results in organelle dysfunction but also has detrimental effects at cellular and systemic levels. The ability of the ER to modulate the levels of Ca2+ within the cell plays an important role in the growth and development of the organism. Consequently, the ER and contents must be essential for the growth and subsequent well-being of the organism.

ACKNOWLEDGMENTS Work in our laboratory is supported by grants from the Canadian Institutes of Health Research Heart and Stroke Foundation of Alberta, Alberta Innovates—Heath Sciences. J. Groenendyk is sup-

ported by the Canadian Institutes of Health Research, the Heart and Stroke Foundation of Canada, and the Membrane Protein and the Cardiovascular Disease Training Program.

ABBREVIATIONS ATF6 COP EDEM ER ERAD GRP InsP3 IRE1 MHC PDI PERK

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activating transcription factor 6 coat protein complex endoplasmic reticulum degradation-enhancing 1,2-mannosidase-like protein endoplasmic reticulum endoplasmic-reticulum-associated degradation glucose-regulated protein inositol-1,4,5-trisphosphate inositol-requiring 1 kinase major histocompatability complex protein disulfide isomerase dsRNA-activated protein kinase-like endoplasmic reticulum kinase

RyR ryanodine receptor SERCA sarcoplasmic/endoplasmic reticulum Ca2+ ATPase SNAP synaptosomal associated protein SOCE store-operated Ca2+ entry SR sarcoplasmic reticulum STIM1 stromal-interacting molecule 1 TNF tumor necrosis factor UDP uridine diphosphate UGGT UDP-glucose : glycoprotein transferase UPR unfolded protein response XBP1 X-box-binding protein 1

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THE GOLGI APPARATUS James W. Dennis Ivan R. Nabi

DEFINITION The Golgi apparatus is a morphologically distinct organelle composed of multiple stacks or cisternae that represents a critical element of the secretory pathway. The Golgi is localized at the site of the centrosome during interphase, and its structure (cisternae organization and number) varies between species and between cells and tissues. The cis-Golgi receives newly synthesized proteins from specialized exit sites of the endoplasmic reticulum (ER) and modifies them during their transit of the cis-, medial, and trans-Golgi stacks. Golgi modifications include terminal glycosylation of proteins and lipids delivered from the ER as well as sulfation and phosphorylation. Golgi-mediated glycosylation of proteins and lipids occurs sequentially as cargo passes through the Golgi and is critically dependent on expression of specific glycosyltransferases and their sugar-nucleotide substrates. How traffic through the Golgi occurs remains a subject of debate, and the role of vesicular traffic, cisternal maturation, and membrane partitioning remains to be clearly defined. Upon arrival to the trans-Golgi network (TGN), proteins and lipids are sorted and delivered to their target sites, including different plasma membrane domains, endosomes, and lysosomes. Sorting signals include amino acid sequences, lipid anchors, as well as Golgi modifications, including mannose-6-P phosphorylation, which targets lysosomal hydrolases for intracellular delivery from the Golgi to late endosomes and lysosomes. Golgi association with the microtubule organizing center (MTOC) and directional exocytosis together with its control of various posttranslational modifications, including, but not limited to, glycosylation, palmitoylation, and phosphorylation, contribute to its central role in the regulation of cellular signaling, secretion, mitosis, and polarity as well as cell–cell communication during development and tissue renewal.

HISTORICAL PERSPECTIVE The Golgi apparatus is named after Camillo Golgi, a professor of General Pathology and Histology at the University of Pavia in the late 1800s and early 1900s. Golgi developed a stain, the black reaction or Golgi-stain, which labeled neurons but also led to the identification of an “internal reticular apparatus.” Definitive proof of the existence of the Golgi came with the development of electron microscopy (EM) and visualization of the ultrastructure of this organelle (see Mazzarello et al. 2009 for an interesting discussion of the early history of the Golgi apparatus). Following initial EM studies by Dalton and Felix

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(1954), subsequent work led to the general characterization of Golgi ultrastructure as a stack of smooth-surfaced cisternae, often slightly curved, with one side oriented toward the rough ER (RER; see Chapter 7) and the other toward the plasma membrane or nucleus, giving rise to the distinct cis–trans polarity of the Golgi apparatus (Farquhar and Palade 1981). Three-dimensional EM analyses of the Golgi showing extensive continuities between cisternae have described the Golgi apparatus as an interconnected membrane network with a large number of associated vesicles (Rambourg et al. 1979; Rambourg and Clermont 1990; Ladinsky et al. 1999; Marsh et al. 2004). A role for the Golgi apparatus as an intermediate in the secretory pathway was initially determined using radioautography of pancreatic slices pulse labeled with radioactive amino acids to follow de novo biosynthesized protein. Initial radioactivity accumulated in the rough ER, passed transiently through the Golgi before accumulating in zymogen vacuoles (Jamieson and Palade 1967). Similarly, collagen precursors synthesized in the ER of odontoblasts passed through the Golgi prior to delivery to secretory vesicles (Weinstock and Leblond 1974). Using temperature-sensitive vesicular stomatitis virus (VSV) G protein, which accumulates in the ER at the nonpermissive temperature, the Golgi was shown to be intermediate in the route of transit of the protein to the plasma membrane (Bergmann et al. 1981). Importantly, VSV G protein was found to arrive asymmetrically at the cis face of the Golgi (Bergmann and Singer 1983) and, in a study using a 20°C temperature block that prevents protein exit from the Golgi, to leave from the trans face of the Golgi (Griffiths et al. 1985). The site of exit from the Golgi is referred to as the TGN and functions to sort proteins into vesicles for targeting to different intracellular sites (Griffiths and Simons 1986). A number of EM-based approaches, including osmium impregnation of cis cisternae (Friend and Murray 1965), localization of thiamine pyrophosphatase and alkaline phosphatase activities to trans cisternae (Goldfischer et al. 1964), and enrichment of cholesterol from cis to trans cisternae (Orci et al. 1981), provided the first evidence of the distinct molecular composition of the various stacks of the Golgi. Glycan-processing enzymes were localized to Golgi membranes (Fleischer et al. 1969; Schachter et al. 1970; Kornfeld and Kornfeld 1985) and separation of early-acting processing enzymes (α-mannosidase) from late-acting transferases (N-acetylglucosaminyltransferase II and galactosyltransferases) by differential centrifugation localized these activities to distinct Golgi regions (Dunphy et al. 1981; Dunphy and Rothman 1983). Using immunocytochemistry of EM frozen sections, the localization of galactosyltransferase (Roth and Berger 1982) and the galactose-specific lectin Ricinus communis agglutinin 1 (RCA) (Griffiths et al. 1982) confirmed the localization of galactosyltransferase activity to trans-Golgi cisternae. N-acetylglucosaminyltransferase I was localized to central Golgi stacks (Dunphy et al. 1985), while in monensin-treated cells Semliki Forest viral proteins accumulated in central Golgi cisternae (Griffiths et al. 1983). The terminal step in protein glycosylation, sialylation, was localized to the TGN, the site of protein exit from the Golgi (Roth et al. 1985). Together, these results led to the current concept of three functional Golgi compartments, cis, medial, and trans, in addition to the TGN, through which newly synthesized proteins leaving the ER pass and are sequentially modified by Golgi-associated glycan-modifying enzymes (Fig. 8.1). The Golgi matrix, an electron-dense material that surrounds and links the cisternae, was first seen by EM in plant cells (Franke et al. 1972; Staehelin et al. 1990). These proteinacious bridges were shown to form a structural scaffold that holds cisternae together in a stacked unit (Cluett and Brown 1992; Slusarewicz et al. 1994; Seemann et al. 2000). The Golgi matrix is composed of the Golgins and Golgi reassembly stacking proteins (GRASPs), such as giantin and GM130, and plays both structural and vesicle tethering

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TGN trans medial cis

Figure 8.1. Domain partitioning in the Golgi. Cargo traversing the Golgi undergoes exchange (white arrows) between processing (dark) and transport (light) domains in each Golgi stack. Residence time in processing domains will determine both the extent and the nature of glycosylation and impact inversely on the rate of transport (black arrow) though the Golgi for subsequent release in the TGN (small black arrows). Glycosyltransferase distribution between cisternae is indicated to the right. How these domains are organized and the role of vesicular transport, cisternal maturation, and intercisternal tubules in cargo transfer from one stack to the other remains to be clarified.

roles (Shorter and Warren 2002; Short et al. 2005). Golgi targeting signals were localized to the transmembrane domains of Golgi enzymes (Nilsson et al. 1991; Machamer 1993). The transmembrane domains of glycosyltransferases and other Golgi-retained proteins are shorter than plasma membrane-targeted proteins, and transmembrane domain length was shown to be a determinant of Golgi retention (Munro 1995). The reduced cholesterol content of Golgi membranes relative to the plasma membrane and the cholesterol gradient within the Golgi from cis to trans stacks (Orci et al. 1981; Coxey et al. 1993) was proposed to limit the width of Golgi membranes and therefore retain proteins with shorter TMDs in the Golgi, and selectively allow exit of proteins with longer TMDs (Bretscher and Munro 1993).

GOLGI GLYCOSYLATION Proteins synthesized in the RER have N-terminal signal peptides for insertion into the membrane. Signal peptides commonly have five to eight basic residues followed by a seven- to fifteen-residue hydrophobic core, then six polar amino acids that contain the signal peptide cleavage site. During transit into the ER lumen, proteins are glycosylated on a subset of asparagines (Asn; N-glycosylation) and serine/threonine (Ser;Thr; O-glycosylation) residues (see also Chapter 7 for further discussion of glycosylation in the ER). Several O-glycosylation pathways produce distinct structural classes of glycan modification including the mucin O-glycans, proteoglycans (sulfated heparin, chondroitin, and dermatin chains), as well as O-linked fucose, glucose, and mannose elongated by GlcNAc-Gal-SA. N-glycosylation also begins in the rough ER with transfer from the Glc3Man9GlcNAc2-pp-dolichol donor to NXS/T (X≠Pro) sites by oligosaccharyltransferase (OST). OST is found widely in all domains of life, but the N-glycan structures transferred from dolichol are truncated in many microorganisms (Banerjee et al. 2007). Mammalian OST is physically associated with the translocation channel (Sec61) complex and ribosome-associated membrane protein fraction. Bacterial OST activity is encoded in

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a single polypeptide, while the mammalian OST is a complex of nine gene products, including a catalytic peptide and other subunits that appear to enhance site recognition and allow conditional regulation of OST (Izquierdo et al. 2009). Depletion of OST or inhibition of Glc3Man9GlcNAc2-pp-dolichol synthesis with the natural product tunicamycin results in ER stress and the unfolded protein response (UPR), which inhibits protein synthesis. Mammals have two genes encoding the catalytic subunit; STT3A is primarily responsible for co-translational modification of nascent polypeptide entering the RER, while STT3B modifies sites missed by STT3A prior to protein folding (Ruiz-Canada et al. 2009). Interestingly, the bacterial system glycosylate sites located in flexible parts of folded proteins, whereas in eukaryotic cells, OST acts largely co-translationally and prior to folding (Kowarik et al. 2006). Preassembly of the donor substrate Glc3Man9GlcNAc2pp-dolichol begins on the cytosolic face of the ER and is completed in the ER lumen, with each sugar added by a specific glycosyltransferase. The Man5GlcNAc2-pp-dolichol intermediate is translocated across the membrane by a “flip-ase” activity exposing the glycan to enzyme that completes donor biosynthesis. The 14 yeast altered-in-glycosylation (ALG) genes define the pathway, and rare hypomorphic mutations in mammalian homologs result in congenital disorders of glycosylation type I (CDG). After transfer of the glycans to asparagines (Asn), two glucose (Glc) units are removed in the ER. The presence of the N-glycan promotes protein folding directly or by recruiting the ER luminal chaperones calnexin and calreticulin. The remaining Glc residue is recycled by α-glucosidase II and ER α-glucosyltransferase, which delays transit to the Golgi until folding is complete (Helenius and Aebi 2004). Glycoproteome analyzed by mass spectrometry confirms that 97% of all glycosylated Asn sites conform to NXS/T (X≠P) with >75% of unique sites modified at efficiencies of ∼98% (Zielinska et al. 2010). Recognition by OST is enhanced by neighboring aromatic amino acids, and by turns that expose motifs at the convex surface of the peptide, but OST is less efficient in the proximity of disulfide bonds, the transmembrane domain, and at the C-terminus of proteins (Jones et al. 2005). Mutagenesis of individual NXS/T sites suggest that most N-glycans are not required for protein folding, but their post-ER modification in the Golgi promotes localization at the cell surface and extracellular matrix (Elleman et al. 2000). N-glycan structural diversity in vertebrates is due to partial saturation of N-glycan branching, as well as competition between Golgi enzymes that add terminal sialic acid (SA), fucose (Fuc), and/or N-acetylgalactosamine (GalNAc) in various linkages. In the cis-Golgi, N-acetylglucosaminyltransferases I, II, III IV, and V (encoded by Mgat1, Mgat2, Mgat3, Mgat4a/b, and Mgat5) comprise the N-glycan branching pathway (Schachter 1986). These enzymes catalyze sequential reactions (Fig. 8.1) and are limited by their relative activities, uridine diphosphate N-acetylglucosamine (UDP)-GlcNAc, and transit time of glycoprotein substrates through the Golgi (Lau et al. 2007a). Glycoproteins emerge from the Golgi as a molecular distribution or population of glycoforms, which is characteristic of cells, tissues, and their environment. For example, >40 N-glycan structures have been identified on EGF receptor (EGFR) from a single source (Stroop et al. 2000). Eight NXS/T sites on EGFR are occupied with N-glycans, and when the glycoform distribution is modeled with only 14 possible N-glycans distributed randomly over each site, the potential number of glycoforms is 203,490 (Lau et al. 2007a).

TRAFFICKING TO, FROM, AND WITHIN THE GOLGI The Golgi is situated at the crossroads of traffic in the secretory pathway, between the ER and the plasma membrane. Newly synthesized cargo is delivered from the ER via COPII

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vesicles that bud and form vesicular-tubular clusters equivalent to the ER–Golgi intermediate compartment (ERGIC) that move along microtubules from ER exit sites to the cisGolgi (Barlowe et al. 1994; Cole et al. 1996; Hauri and Schweizer 1992; Kuge et al. 1994; Presley et al. 1997). Coat exchange from COPII anterograde vesicles to COPI retrograde vesicles serves a retrieval function, within ERGIC and throughout the Golgi, returning ER proteins back to the ER, in part via the KDEL retrieval motif for luminal ER proteins (Aridor and Balch 1996; Pelham 1996). Recruitment of COPI to transport vesicles is mediated by adenosine diphosphate (ADP)-ribosylation factor 1 (ARF1), whose sensitivity to the fungal metabolite Brefeldin A (BFA) is responsible for dissolution of the Golgi and its return to the ER upon BFA treatment (Lippincott-Schwartz et al. 1989; Donaldson et al. 1992a, b). Trafficking out from the Golgi is thought to occur for the most part at the TGN. Proteins are sorted into COPI vesicles for retrograde trafficking, clathrin-coated vesicles for delivery to endosomes, and TGN-to-plasma membrane carriers (TPCs) for delivery to the plasma membrane. There are at least two types of TPCs, derived from raft-like domains and apically targeted in polarized epithelial cells, and basolaterally targeted and sorted via interaction of peptide-based sorting signals through interaction with adaptor proteins (Bard and Malhotra 2006; see also Chapter 21 on epithelial domains). Selective targeting of a fluorescent ceramide analog, C6-NBD-ceramide, from the TGN to the apical surface of polarized Madin-Darby canine kidney (MDCK) epithelial cells (van Meer et al. 1987) led to the proposal of lateral separation of sphingolipids and glycerolipids into microdomains within the trans-Golgi and TGN and their segregation into distinct vesicles destined for delivery to the apical membrane (Simons and van Meer 1988). These studies contributed to the formulation of the raft hypothesis of cholesterol/sphingolipid-rich microdomains (see Chapters 1, 4, and 5 for further discussion of lipid rafts, and Chapters 13 and 21 for discussion of raft-dependent apical targeting). Apically targeted, glycosylphosphatidylinositol (GPI)-anchored proteins were shown to segregate into detergent-resistant, glycolipid-rich domains during transit through the Golgi (Brown and Rose 1992). Partitioning of Golgi enzymes and cargo between two membrane phases within Golgi cisternae has recently been proposed to mediate intraGolgi transport (Patterson et al. 2008), and caveolin-1, a raft-targeted protein, forms scaffolds that segregate from VSV G protein in Golgi stacks (Hayer et al. 2009). While trafficking to and from the Golgi has been relatively well characterized, traffic within the Golgi remains something of a black box. A debate spanning several decades concerns the relative role of vesicular traffic versus cisternal maturation in passage of cargo through the Golgi. The vesicular traffic model argues that cargo is transported anterogradely via vesicular transport from one cisternae to the next while modifying glycosyltransferases remain stably localized to their respective cisternae. The cisternal maturation model proposes that cisternae mature together with cargo from the cis to trans direction, with modifying glycosyltransferases moving retrogradely via vesicular transport. EM reconstructions show quite clearly the multiple vesicles associated with Golgi cisternae, including, but certainly not limited to, the TGN (Ladinsky et al. 1999). Indeed, an intra-Golgi vesicle fusion assay based on Golgi-dependent glycosylation led to the identification of the NSF–SNAP–SNARE fusion complex (NSF: N-ethylmaleimidesensitive factor; SNAP: soluble NSF attachment protein; SNARE: SNAP receptors; Block et al. 1988). At the same time, the ability of large cargo, such as 300-nm-long procollagen fibers, to traverse the Golgi argues that vesicle transport is not sufficient to account for all traffic through the Golgi (Bonfanti et al. 1998). However, cargo vesicle size may be variable; a protein localized to ER exit sites, TANGO1, may facilitate loading of bulky cargo, such as collagen VII, into COPII vesicles for transport to the Golgi (Saito et al. 2009). In yeast, cisternal maturation has been convincingly demonstrated using dual color fluorescent

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experiments in which cargo and resident Golgi proteins are differentially labeled (Losev et al. 2006; Matsuura-Tokita et al. 2006). A number of reviews have dealt quite extensively with the arguments for and against both anterograde vesicular transport and cisternal maturation (Rothman and Warren 1994; Glick and Malhotra 1998; Warren and Malhotra 1998; Elsner et al. 2003; Glick and Nakano 2009). In spite of the extensive debate, it is clear that neither model alone can account for the complexity of traffic through the Golgi. As mentioned previously, the tremendous numbers of possible glycoforms and the functional implications of these on cellular behavior argue that Golgi structure and organization must necessarily impact on the extent of glycosylation reactions within the Golgi. Indeed, for the case of the repeated galactosyltransferase and GlcNAc-transferase reactions that generate Mgat5-dependent polylactosamine chains, the residence time of a protein in the Golgi correlates inversely with the extent of its glycosylation (Nabi and RodriguezBoulan 1993). This suggests that a slower rate of passage through the Golgi may result in increased interaction with glycosyltransferases. Indeed, prevention of protein exit from the Golgi using a 20°C block results in extended polylactosamine glycosylation (Wang et al. 1991; Nabi and Dennis 1998). Any model for Golgi traffic must necessarily account for glycoform variability and differential rates and quantities of cargo transit through the Golgi. One way that the Golgi could selectively regulate both rates of passage and glycoform variability would be through lateral partitioning into processing domains enriched in Golgi glycosyltransferases and transport domains (Patterson et al. 2008). In this study, the rapid and exponential loss of Golgi-associated VSV G-protein fused to yellow fluorescent protein (YFP) fluorescence observed using a photobleaching approach (inverse fluorescence recovery after photobleaching [iFRAP]) did not correspond to a predicted cisternal maturation model that assumed a 25-minute Golgi transit time. To explain exponential exit of cargo from Golgi, the partitioning model proposed that cargo exits the Golgi from all cisternae (Patterson et al. 2008), a conclusion difficult to reconcile with the sequential glycosylation events through Golgi stacks, the lack of appearance of immature, endoglycosidase H (endo H)-sensitive glycoproteins at the plasma membrane, and the waves of cargo that have been observed moving through Golgi stacks (Emr et al. 2009). Indeed, cargo waves of VSV G protein have been observed to pass sequentially through the Golgi following a 15°C block and to be associated with the formation of intercisternal tubules (Trucco et al. 2004). Furthermore, Golgi transit time may be highly variable. Using acquisition of endo H resistance by newly synthesized LAMP-2 as a measure of arrival to the cis-Golgi (t1/2 = 25 minutes) and surface delivery as a measure of Golgi exit (t1/2 = 28 minutes), fully mature glycosylated LAMP-2 was shown to transit the Golgi very rapidly. Interestingly, Golgi transit rates were slower (∼15 minutes) in cells plated for only 1 day in which LAMP-2 presented extended polylactosamine chains (Nabi and Rodriguez-Boulan 1993). Rapid and differential rates of exit from the Golgi could therefore potentially be explained without invoking exit from pre-TGN structures. Golgi models that can explain the complex patterns of glycosylation observed in mammalian cells will likely include lateral microdomain organization and exchange between processing domains that mediate glycoform processing and transport domains that efficiently transport proteins through the Golgi (Fig. 8.1). Tremendous variations and heterogeneity in N-glycan branching and elongation are observed on secreted glycoproteins, suggesting that Golgi-dependent glycosylation is a stochastic process, limited by time as well as enzyme and sugar nucleotide levels (Stroop et al. 2000; Lau et al. 2007b). Residence time in processing domains will impact not only on the extent of glycosylation but also, inversely, on the rate of transit through the Golgi, as observed for LAMP-2 (Nabi and Rodriguez-Boulan 1993). Differential and local transport through the Golgi may involve a combination of vesicular traffic and cisternal maturation but may also occur via

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continuities between Golgi stacks observed by three-dimensional EM analyses (Rambourg et al. 1979; Rambourg and Clermont 1990; Ladinsky et al. 1999; Marsh et al. 2004; Trucco et al. 2004).

THE GOLGI AND CELLULAR SIGNALING AND POLARITY Golgi N-Glycan Remodeling and Surface Half-Life The number of receptors at the cell surface is an important determinant of ligand-receptor complex formation and thereby intracellular signaling. In breast cancer, levels of EGFR family members are associated with aggressive tumor behavior and reduced survival time (Slamon et al. 1987). However, a significant fraction of many receptors and solute transporters are located in endomembrane compartments that serve in part as reservoirs for mobilization to the cell surface. Endocytosis and recycling rates vary with cell type and can limit the fraction of receptors at the cell surface (Di Fiore and De Camilli, 2001; Doherty and McMahon 2009). Cancer mutations in proteins of the endocytic and recycling machinery, such as avalanche, Rab5, tsg101, and vps25, can either retain growth receptors at the cell surface or maintain their activity in endosomes (reviewed in Mosesson et al. 2008; see also Chapter 9 on endosomes). EGFR has a surface half-life of approximately 8 hours in cultured cells (Wiley et al. 2003), but acute EGF binding recruits Cbl-CIN85endophilin, resulting in rapid coated-pit endocytosis (∼5 minutes) and inactivation of the receptor by phosphatases in the early endosomes (Soubeyran et al. 2002; Offterdinger et al. 2004). Membrane remodeling and endocytosis is often hyperactivated in malignant cells, driven by Src and the small GTPases Rab5/Ras/Cdc42/Rac1, and act to reduce surface levels of receptors. Receptors are glycoproteins, and Golgi remodeling of their N-glycans determines affinities for animal lectins that can oppose loss to endocytosis. The N-glycans on EGFR and other surface glycoproteins bind galectin-3, forming a dynamic lattice or microdomain that slows lateral diffusion and loss to endocytosis (Partridge et al. 2004; Lajoie et al. 2007). The galectins are a ubiquitous family of lectins that bind to N-acetyllactosamine (Galβ1,4GlcNAc) (Cooper 2002), a sequence common to branched N-glycans on cell surface glycoproteins (Patnaik et al. 2006). The lattice also “insulates” against ligandindependent EGFR dimerization and raises the threshold for ligand activation. Receptor affinities for the lattice increase in proportion to the number of N-glycan sites (NXS/T), as well as Golgi modification of individual N-glycans (Lau et al. 2007a). The levels of high-affinity tri- and tetra-antennary N-glycans are dependent on developmental expression of Golgi enzymes but also on metabolic supply of UDP-GlcNAc to the pathway (Lau et al. 2007a). Polylactosamine (repeats of Galβ1,4GlcNAcβ1,3) further enhances affinities for galectins, and as discussed earlier, polymer length is inversely proportional to glycoprotein transit rates through the Golgi (Wang et al. 1991; Nabi and Dennis 1998). Regulation of receptor glycosylation through both selective glycosyltransferase expression and Golgi structure can therefore impact on receptor surface expression and signaling potential.

Golgi and Cell Polarity The Golgi is adjacent to the MTOC or centrosome (Fig. 8.2; see also Chapters 14, 15, and 22 for further discussion of the centrosome and Golgi in cell polarity). Both organelles can quickly realign in the direction of cell motility when a cell monolayer is wounded.

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Figure 8.2. The Golgi and microtubules. The Golgi apparatus of Cos7 cells labeled in red with antibodies to the Golgi matrix protein GM130 is localized around the MTOC from which microtubules (green) emanate. Scale bar = 20 μm. Image generated by Pascal St. Pierre.

Under these conditions, the Golgi matrix proteins GRASP65, Golgin-160, and GMAP210 are required for organelle repositioning, directed secretion, and cell polarity (Bisel et al. 2008; Yadav et al. 2009). In filamentous fungi, hyphal formation involves formin/actindependent redistribution of the Golgi complex to the growing tip where secretion efficiency is enhanced, presumably in a polarized manner (Rida et al. 2006). Posttranslational modifications of proteins and directional secretion through the Golgi complex may cooperate to maintain gradients of receptors and membrane-associated signaling proteins across the cell. For example, H-Ras, N-Ras, and K-Ras have similar biochemical activities in vitro, but in vivo their cellular locations differ due to posttranslational lipid modifications encoded by divergent sequences at the C-terminus (Rocks et al. 2006). RAS are guanine nucleotide-binding proteins that cycle between inactive GDP-bound and active GTPbound states downstream of multiple receptors and stimuli. Depalmitoylation redistributes farnesylated Ras in all membranes, and repalmitoylation retains Ras at the Golgi, where it can be redirected to the plasma membrane (PM) via the secretory pathway. Palmitoylated H-Ras and N-Ras are subject to constitutive de- and reacylation, which drives rapid exchange between the PM and the Golgi apparatus. The Golgi–PM de- and repalmitoylation cycle supports the establishment of gradients of Ras across the cell and thereby enables polarized signaling in response to growth factors (Rocks et al. 2005). RasGRP1/2, the guanine nucleotide exchange factor (GEF) associated with the T-cell receptor (TCR), also transit between the Golgi and PM, a process that influences signaling

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strength in a cooperative manner with Vav1 and PI3 kinase signaling (Ruiz et al. 2007). TCR clusters to form an immune synapse with antigen-presenting cells, followed by rapid realignment of the MTOC and Golgi for directional release of lytic factors and IL-2/IFN-γ cytokines into the synapse (Huse et al. 2008). The threshold for TCR clustering is dependent on N-glycan modifications in the Golgi that regulate affinities for galectins and slow TCR recruitment into the immune synapse (Demetriou et al. 2001). However, the inflammatory cytokine tumor necrosis factor (TNF) appear to be secreted in all directions suggesting a distinction between cell–cell contact and longer-range communication, where non-polar secretion may be appropriate (Huse et al. 2006).

Mitosis, Cell Polarity, and Fate Extracellular matrix is secreted at the basal surface of epithelial cells (see Chapter 21), where cells are sensitive to spatial gradients that can direct asymmetric force on microtubule asters, which orient the mitotic spindle and cleavage plane (Thery et al. 2007). Downstream of extracellular cues, gradients of cytosolic proteins, and differences in mother–daughter centrosomes of mammalian neocortex are instructive on orientation, asymmetric cell division, and cell fate in daughter cells (Lasorella et al. 2006; Wang et al. 2009). For T cells, contact and activation by an antigen-presenting cell induce an immune synapse where TCR and integrins cluster and activate growth signaling. This leads to growth and polarization of the T cell, followed by asymmetric cell division, where the contacting cell is fated to be an effector and the noncontact cell, a memory lineage (Chang et al. 2007). The Golgi realigns to ensure rapid assembly of the immunological synapse and, subsequently, polarized secretion of cytokines and lytic granules by the effector cell (Stinchcombe et al. 2001). Polar body formation in mouse oocytes is an extreme example of asymmetric cell division and also requires Golgi-based membrane fusion, presumably for delivering critical cues to the PM for cytoskeletal reorganization (Wang et al. 2008). One such cue may be asymmetric distribution of Numb, which is opposed by ACBD3, a protein associated with the Golgi apparatus in interphase progenitor cells but that becomes cytosolic after Golgi fragmentation during mitosis. Numb polarization in the PM distinguishes the two daughter cells (Zhou et al. 2007). In hippocampal neurons, the first neurite that develops after mitosis is nearest the centrosomes and Golgi, but perpendicular to the plane of cell division, an arrangement necessary and sufficient for neuronal polarization (de Anda et al. 2005). This suggests that pre-mitotic extracellular cues direct spindle orientation, and, in turn, post-mitotic Golgi and MTOC positions instruct polarity. The MTOC, centrosomes, and basal body are well-studied centers for cell-polarizing activities (see also Chapters 14 and 15). The Golgi is less well studied, but, as discussed here, is likely to be a critical axis of trafficking that establishes and maintains cell polarity.

Golgi and Planar Cell Polarity The atypical cadherins Fat and Dachsous (Ds) (see also Chapter 18 on the adherens junction) restrict proliferation and are required to establish planar cell polarity (PCP; Matakatsu and Blair 2006). Fat and Ds exhibit gradients of opposing polarity across the fly imaginal disk that integrate signaling across the field of cells to regulate organ size and shape. Ds on neighboring cells binds to Fat and stimulates the Hippo kinase pathway, which blocks YAP/TAZ-mediated transcription of cyclin E, Diap1, and Four-jointed (Fj) (Sopko and McNeill 2009). Mice deficient in FAT4 display misoriented cell divisions and develop cystic kidney disease linking spindle orientation to the reestablishment of polarity and

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growth control in daughter cells (Saburi et al. 2008). Graded Golgi expression of Fj across the field of cells enhances Fat activity regionally by steepening the Fat/Ds gradient. Fj shares the type II transmembrane and Golgi localization of glycosyltransferases, but remarkably the luminal domain of Fj is an atypical kinase that selectively phosphorylates domains in Fat and its counter-receptor Ds (Ishikawa et al. 2008). The kinase activity of Fj modifies Ds and Fat en route to the cell surface and regulates their interaction. Since the Golgi is a focus for polarized trafficking of other cargo to the cell surface, it is possible that Fj-dependent phosphorylation of Ds and Fat is a critical determinant of their opposing polarization across each cell to form the required smooth gradient within and between cells across the morphogenetic field. The incremental changes in Fj expression due to transcription form a discontinuous gradient from cell to cell, but the Golgi activity of Fj may convert stepwise changes into a smooth molecular gradient of Golgi-modified FAT and Ds across individual cells. It is tempting to speculate that Fj kinase activity may regulate glycosylation of FAT and Ds, and thereby impact on their cell surface half-life and recruitment, as shown for cytokine receptors into plasma membrane microdomains (Demetriou et al. 2001; Partridge et al. 2004; Chen et al. 2007; Lajoie et al. 2007; Lajoie et al. 2009).

CONCLUSION Since its discovery over a hundred years ago, the Golgi apparatus has intrigued based on its unique morphology and central role in the secretory pathway and glycosylation. While understanding Golgi function continues to be a challenge, it is becoming increasingly clear that the Golgi is a cellular hub for translational modification of cellular and secreted proteins that functions in signaling, secretion, mitosis, polarity, and cell–cell communication.

ACKNOWLEDGMENT Supported by a grant from the Canadian Institutes for Health Research (CIHR MOP-43938) to JWD and IRN.

ABBREVIATIONS ALG Asn BFA CDG Ds EGFR EM endo H ER ERGIC GalNAc GEF Glc

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altered-in-glycosylation Asparagine Brefeldin A congenital disorders of glycosylation type I Dachsous EGF receptor electron microscopy endoglycosidase H endoplasmic reticulum ER–Golgi intermediate compartment Fuc—fucose N-acetylgalactosamine guanine nucleotide exchange factor glucose

GlcNAc GRASPs Man MTOC NSF OST PCP PM RCA RER SA Ser SNAP SNARE

N-acetylglucosamine Golgi reassembly stacking proteins mannose microtubule organizing center N-ethylmaleimide-sensitive factor oligosaccharyltransferase planar cell polarity plasma membrane Ricinus communis agglutinin 1 rough endoplasmic reticulum sialic acid serine soluble NSF attachment protein soluble NSF attachment protein receptors

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REFERENCES

TGN Thr TNF TPCs

trans-Golgi network Threonine tumor necrosis factor TGN-to-plasma membrane carriers

UPR VSV VTC YFP

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unfolded protein response vesicular stomatitis virus vesicular-tubular cluster yellow fluorescent protein

REFERENCE Aridor M, Balch WE. 1996. Principles of selective transport: coat complexes hold the key. Trends Cell Biol 6:315–20. Banerjee S, Vishwanath P, Cui J, Kelleher DJ, Gilmore R, Robbins PW, Samuelson J. 2007. The evolution of N-glycan-dependent endoplasmic reticulum quality control factors for glycoprotein folding and degradation. Proc Natl Acad Sci U S A 104:11676–81. Bard F, Malhotra V. 2006. The formation of TGN-toplasma-membrane transport carriers. Annu Rev Cell Dev Biol 22:439–55. Barlowe C, Orci L, Yeung T, Hosobuchi M, Hamamoto S, Salama N, Rexach MF, Ravazzola M, Amherdt M, Schekman R. 1994. COPII: a membrane coat formed by Sec proteins that drive vesicle budding from the endoplasmic reticulum. Cell 77:895–907. Bergmann JE, Singer SJ. 1983. Immunoelectron microscopic studies of the intracellular transport of the membrane glycoprotein (G) of vesicular stomatitis virus in infected Chinese hamster ovary cells. J Cell Biol 97:1777–87. Bergmann JE, Tokuyasu KT, Singer SJ. 1981. Passage of an integral membrane protein, the vesicular stomatitis virus glycoprotein, through the Golgi apparatus en route to the plasma membrane. Proc Natl Acad Sci U S A 78:1746–50. Bisel B, Wang Y, Wei JH, Xiang Y, Tang D, MironMendoza M, Yoshimura S, Nakamura N, Seemann J. 2008. ERK regulates Golgi and centrosome orientation towards the leading edge through GRASP65. J Cell Biol 182:837–43. Block MR, Glick BS, Wilcox CA, Wieland FT, Rothman JE. 1988. Purification of an N-ethylmaleimidesensitive protein catalyzing vesicular transport. Proc Natl Acad Sci U S A 85:7852–6. Bonfanti L, Mironov JAA, Martínez-Menárguez JA, Martella O, Fusella A, Baldassarre M, Buccione R, Geuze HJ, Mironov AA, Jr., Luini A. 1998. Procollagen traverses the Golgi stack without leaving the lumen of cisternae: evidence for cisternal maturation. Cell 95:993–1003. Bretscher MS, Munro S. 1993. Cholesterol and the Golgi apparatus. Science 261:1280–1. Brown DA, Rose JK. 1992. Sorting of GPI-anchored proteins to glycolipid-enriched membrane subdomains during transport to the apical cell surface. Cell 68:533–44. Chang JT, Palanivel VR, Kinjyo I, Schambach F, Intlekofer AM, Banerjee A, Longworth SA, Vinup KE, Mrass P, Oliaro J, Killeen N, Orange JS, Russell SM, Weninger W, Reiner SL. 2007. Asymmetric T lymphocyte division in the initiation of adaptive immune responses. Science 315:1687–91.

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ENDOSOMES Thierry Galvez Marino Zerial

DEFINITION Like the Golgi complex, the endoplasmic reticulum, or lysosomes, endosomes are intracellular, membrane-bound compartments of eukaryotic cells with a specific and complex morphology. They are composed of vesicular and tubular structures of various shapes and sizes (e.g., simple vesicles, tubules, cisternae, multivesicular bodies [MVBs]) that can dynamically change over time and travel long distances within a cell (i.e., in the meter range in the case of an axon). Endosomes transport cargo molecules from mainly two sources, the cell surface and the biosynthetic pathway, toward two main destinations, the lysosomes and the cell surface. Historically, this distinction contributed to define the degradative endosomal pathway, leading to lysosomes, and the recycling endosomal pathway, returning to the plasma membrane. However, the biochemical characterization of endosomes revealed another level of complexity in the diversity of endosome classes. We know now that each endocytic route is composed of multiple endosomal populations defined by a specific combination of molecular components. On the endosomal surface, these components assemble and disassemble dynamically into membrane subdomains that may represent the actual functional units of the endosomal system responsible for essential activities such as vesicular fission and fusion, and cargo sorting. In addition to cargo sorting and transport, endosomes play a major role in cell signaling, positioning them at the heart of many important homeostatic functions. Endosomes are therefore integrators and effectors of multiple cellular functions and thereby participate in cell and tissue homeostasis and morphogenesis.

AN OVERVIEW OF THE ENDOSOMAL NETWORK Historical Considerations The first report of an intracellular compartment containing extracellular material can be traced back to the end of the nineteenth century with Metchnikoff ’s studies on phagocytosis (see also Chapter 10), but the concept of endosomes as it is understood today originates from the work of Werner Straus who studied the uptake of intravenously injected peroxidase by kidney epithelial cells (Straus 1957, 1964). Using astute staining strategies, he demonstrated that at early time points, the peroxidase was found in peripherally located,

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intermediary vacuoles prior to being delivered to lysosomes and degraded. These nonlysosomal vesicles containing endocytosed material en route to lysosomes were later named endosomes by Helenius and colleagues (1983). But endosomes were soon recognized not to be just intermediates in degradation. Steinman and colleagues showed that macrophages internalized 190% of their surface every hour, but their lysosomal area represented only 18% of the cell-surface area (Steinman et al. 1976). Since membrane protein turnover rates were known to be 2% or less per hour, it was proposed that most of the endocytosed membrane must escape degradation and be reutilized, thus revealing the existence of the recycling route to the cell surface. Similar reasoning, taking into account rates of endocytosis and protein turnover, led Brown and Goldstein to propose that the receptor for lowdensity lipoprotein (LDL) recycled to the cell surface after internalization, whereas the ligand, LDL, was degraded in lysosomes (Brown and Goldstein 1976; Goldstein et al. 1979). Although the works from Steinman, Brown, and Goldstein presented endosomes as a sorting compartment, no sorting mechanism was proposed. An important conceptual leap was made by Geuze and colleagues who used electron microscopy techniques to follow the endocytic routes of both a ligand, asialoglycoprotein, and its receptor. They observed endosomes in which ligands and receptors were spatially segregated: The receptors were enriched in thin tubules connected to vesicles containing the ligands in their lumen (Geuze et al. 1983). This observation hinted at an elegant sorting mechanism based on the geometry of endosomes. In this model, membrane receptors would thermodynamically partition into tubules where the surface-to-volume ratio is high, whereas soluble ligands would diffuse into the lumen of the vesicle. In a pioneering approach using quantitative fluorescent microscopy on populations of endosomes, Maxfield and colleagues confirmed and complemented this model, showing that recycling membrane receptors are rapidly and also iteratively removed from endosomes by geometric sorting, whereas cargo molecules destined to lysosomes were concentrated in the endosomal lumen, establishing a fundamental property of the endosomal system (Dunn et al. 1989). More than 30 years later, these key results and concepts still constitute the essence of the paradigms used to interpret today’s discoveries.

Entries, Routes, and Destinations Entering the Endocytic System Molecules entering the endosomal system have two origins: the cell surface (plasma membrane and intracellular space) and the biosynthetic pathway (see Fig. 9.1). Different types of cargos enter the cell via many different endocytic mechanisms, all involving three essential steps: (1) cargo selection, (2) membrane invagination, and (3) scission of vesicles or tubules. However, these different endocytic routes differ by the molecular machinery at play: clathrin mediated, caveolin or raft mediated, and dependent or not on dynamin for vesicle fission; for a review, see Doherty and McMahon (2009). Initially, different entry mechanisms may lead to distinct endosomal compartments: clathrin-coated vesicles (CCVs) (see Chapter 2) or tubular clathrin-independent carrier (CLIC), but remarkably, the majority of these different types of cargos will eventually reach early endosomes (EEs) in a process regulated by the small GTPase Rab5 (see below). In addition to cell-surface material, EEs receive vesicles originating from the trans-Golgi network (TGN) and transporting, for instance, mannose-6-phosphate receptors (M6PRs) (Ghosh et al. 2003). In EEs, incoming cargos are sorted toward three main destinations: lysosomes, TGN, and plasma membrane. Sorting occurs by membrane remodeling processes that belong essentially to two main categories: (1) membrane retrieval by generation of endosomal tubules and (2) biogenesis of “late,” multivesicular endosomes characterized by the accu-

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7 TGN-to-endosomes Clathrin coat Figure 9.1. Endocytic routes and Rab domains. CCP, clathrin-coated pit; CCV, clathrin-coated vesicle.

mulation of intralumenal vesicles (ILVs) and the acquisition of the capacity to fuse with lysosomes. These two types of events are nevertheless intimately coupled. Membrane Retrieval Routes Lipids and membrane proteins, which are recycled to the plasma membrane or to the TGN are retrieved from EEs by membrane tubulation and geometric sorting (Figs. 9.1 and 9.2). The machinery involved in these membrane shaping events is described below (see section on “Shaping Endosomal Membranes: Bin/Amphiphysin/Rvs [BAR]-Domain-Containing Proteins and ESCRT”). In mammalian cells, at least two routes were proposed to lead

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Figure 9.2. Spatiotemporal dynamics of the endosomal systems. Rab5 early endosomes are first detected at the cell periphery in the proximity of the plasma membrane and subsequently move toward the cell center where they convert into Rab7 endosomes and fuse with lysosomes. Early in their lifetime, Rab5 endosomes undergo multiple rounds of homotypic fusion resulting in larger endosomes, whereas most of endocytosed membrane and recycling cargos are retrieved by tubulation. As a result, cargos destined to lysosomes are concentrated in the lumen of maturing endosomes.

from EEs to the plasma membrane on the basis of differences in kinetics of cargo transport: a fast (t1/2 ≤ 2 minutes) and a slow (t1/2 ≥ 12 minutes) route. Most of the cargo molecules studied so far are able to recycle via both routes (e.g., transferrin receptor [TFR], G proteincoupled receptors [GPCRs] like β2-AR) except the lipids lactosylceramide and sphingomyelin, which appear to follow mainly the fast route (Hao and Maxfield 2000; Choudhury et al. 2004). As it will be mentioned below in the context of the endosomal compartmentalization (see section on “Rab Domains as Functional Units of the Endosomal System”), the small GTPases Rab4 and Rab35 have been implicated in fast recycling (van der Sluijs et al. 1992; Choudhury et al. 2004; Yudowski et al. 2009), whereas Rab11 is required for the slow route (Chen et al. 1998; Ren et al. 1998). Nevertheless, the identity of the compartments involved in membrane recycling remains poorly defined at the molecular level.

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Morphologically, recycling carriers form a dense network of highly motile vesicles and tubules, more densely packed in the pericentriolar region of some cell types, which define the recycling endosome (RE). The nature of this compartment is not clear: Does it constitute a stable compartment or rather a population of intermediates between EEs, biosynthetic pathway, and cell surface? Functionally, the RE is capable of elaborated sorting operations, recycling cargo to the plasma membrane but also transporting cargo to the TGN (e.g., TGN38, Shiga toxin [Rohn et al. 2000]) or from the TGN to the cell surface (e.g., TFR in some cell types [Ang et al. 2004]). Endosomal tubules are also generated at later stages of endosomal maturation, independently of the activity of Rab4 and Rab11. These tubules contain specific cargo destined to the TGN (e.g., M6PR and sortilin, wntless) and rely on different machineries, Rab9 and its effectors tail-interacting protein of 47 kD (TIP47) and RhoBTB3 on the one hand, the retromer complex on the other (Bonifacino and Rojas 2006; Espinosa et al. 2009) (see Fig. 9.3). Probably, Rab9 and the retromer belong to different routes. Formation of MVBs and Lysosomal Targeting In parallel to the intense tubulation activity, EEs undergo additional membrane remodeling, generating endosomes with specific biochemical and morphological characteristics that eventually fuse with lysosomes (see also Chapter 10). Gradually, cargo molecules destined to lysosome are packaged into 50- to 80-nm-diameter ILVs (see Fig. 9.1). Biochemically, late endosomes (LEs) acquire lysobiphosphatidic acid (LBPA), accumulate lysosome-associated membrane proteins (LAMPs), their luminal pH decreases (from ∼6.5 to ∼5.0), and the small GTPase Rab7 replaces Rab5 through a Rab conversion process (Fig. 9.2 and see section on “Rab Domains as Functional Units of the Endosomal System”) (Rink et al. 2005). Soluble molecules present in the lumen of endosomes seem to be sorted to lysosomes by default (e.g., LDL) but most membrane protein (e.g., epidermal growth factor [EGF] receptors, growth hormone receptor and C-X-C chemokine receptor 4 [CXCR4]) are actively sorted into ILVs in order to reach lysosomes. The best characterized ILV-targeting signal is ubiquitin that is recognized by the endosomal sorting complex required for transport (ESCRT) (see Fig. 9.3 and section on “Shaping Endosomal Membranes: Bin/Amphiphysin/Rvs [BAR]-Domain-Containing Proteins and ESCRT”). Noteworthy, some endosomal membrane proteins (e.g., LAMP-1 and LAMP-2) destined to lysosomes are not found in ILVs hinting at some other sorting mechanisms. In certain cell types, after back fusion of the MVBs with the plasma membrane, ILVs can be released as exosomes in the extracellular space where they may play a role in cell–cell communications (Simons and Raposo 2009).

Dynamics of the System The endosomal system is highly dynamic, its steady state resulting from constant fusion, fission, and transport of vesicles and tubules. The current model is that EEs are constantly generated (or recycled) at the proximity of the plasma membrane and subsequently move toward the cell center (Fig. 9.2). At the same time, they undergo multiple rounds of homotypic fusion, thereby growing in size. Because fusion occurs concomitantly to the membrane retrieval activity involved in recycling, the surface-to-volume ratio of endosomes decreases, and cargo molecules destined to lysosomes (e.g., LDL, EGF) are concentrated in the lumen of maturing endosomes (Dunn et al. 1989; Stoorvogel et al. 1991). Hence, the design of the system allows the concentration of specific cargo and endosomal machineries according to a spatiotemporal order imposed by the endosomal dynamics (Fig. 9.2). Remarkably, such design principles received functional support in the recent

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(A)

Rab5

(B)

Figure 9.3. Examples of functional domains on endosomes. (A) Trafficking domains (only some representative components are shown). (1) Rab5-dependent fusion machinery. (2) Membrane recycling machinery (including Rab11, Rab4, and SNX4). (3) Ubiquitinylated protein sorting into intraluminal vesicles (ILVs) by the endosomal sorting complex required for transport (ESCRT). (4) Endosome-to-Golgi retrieval machinery (including Rab9 and its effector TIP47 and the retromer). (B) Signaling domains. (5) Rab5/APPL1/AKT/GSK-3β signaling domain. (6) Transforming growth factor-β receptor type I (TGFβIR) signaling complex on early endosomes. (7) Extracellular signal-regulated kinase (ERK) scaffolding complex on late endosomes. The MP1-p14 complex is anchored to the late endosome membranes by the p18 adaptor and binds MEK1 and its substrate ERK.

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genome-wide, RNA interference (RNAi)-based survey of the endosomal system (Collinet et al. 2010). Using image-based profiling of HeLa cells containing internalized fluorescent cargos, it was possible to extract multiple descriptors of the endosomal population (e.g., size, cargo concentration, or distance from the cell center). Systematic calculations of these parameters for thousands of perturbations and subsequent analysis of their correlations indeed demonstrated that the radial position of endosomes within the cells determines the concentration of EGF (but not of transferrin, a recycling cargo) as well as the size and local density of endosomes.

MOLECULAR DOMAINS OF ENDOSOMAL MEMBRANES Rab Domains as Functional Units of the Endosomal System The Rab GTPase Code A striking feature of the endosomal network is its “Rab code.” The human genome encodes over 60 Rab GTPases among which 31 have been reported to associate with the endosomal system (Wasmeier et al. 2006; Stenmark 2009). As GTPases, Rab proteins function as molecular switches alternating between GDP-bound “off” and GTP-bound “on” states. In their GTP-bound conformation, they interact with specific sets of effectors. Remarkably, Rab GTPases and their effectors localize to specific intracellular organelles, and this feature is especially prominent in the endocytic system where specific Rab GTPases define distinct subclasses of endosomal subcompartments (e.g., EEs by Rab4 or Rab5; LEs by Rab7; REs by for instance Rab17, Rab11, or Rab35 and carriers from endosomes to TGN by Rab9; see Fig. 9.1). Soluble N-ethylmaleimide-sensitive factor [NSF] attachment protein [SNAP] receptors (SNAREs), key components of the vesicular fusion machinery, as well as phosphoinositides, are also restricted to specific populations of endosomes, but the compartmentalization of Rab GTPases and their effectors is much narrower, revealing a fundamental Rab code of the endocytic system. In addition, cargo flows from one Rab domain to the next, through molecular machinery that leads a Rab GTPase to activate the next one sequentially in the pathway (see below). At their specific location, Rab GTPases and their associated machineries control pivotal functions as diverse as budding, fusion, directional transport, and signaling. For instance, Rab5, the prototypical endocytic Rab, is essential for homotypic EE fusion and acts by recruiting membrane-tethering factors (i.e., early endosomal antigen 1 [EEA-1] and Rabenosyn-5 in complex with Vps45, a member of the Sec1 family of SNARE regulators) and by activating the phosphatidylinositol3-OH kinase Vps34, which regulates the synthesis of the endosomal phospholipid phosphatidylinositol-3-phosphate (PtdIns[3]P). This oligomeric complex associates with endosomal SNAREs (syntaxin 6 and syntaxin 13) as well as the NSF ATPase in order to form the complete fusion machinery. The Concept of Rab Domains Importantly, it has been observed by light and electron microscopy that different Rab GTPases do not occupy the endosomal surface uniformly but are rather compartmentalized into membrane subdomains. For instance, distinct Rab5, Rab4, and Rab11 domains are observed on endosomes along the recycling routes (Sonnichsen et al. 2000). In a similar manner, Rab7 and Rab9 form domains on LE membranes (Barbero et al. 2002). These domains presumably correspond to different effector machineries performing specific functions like sorting (e.g., Rab9 and its effector TIP47 for M6PR sorting [Pfeffer 2009]),

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tubulation (e.g., Rab7 and the retromer complex [Rojas et al. 2008]), or motility (e.g., Rab11A and its effector RAB11FIP2, which is linked to myosin Vb (Hales et al. 2001; Wang et al. 2008; see Fig. 9.3). Super-resolution microscopy techniques such as stimulated depletion (STED) microscopy, photoactivation localization microscopy (PALM), and stochastic optical reconstruction microscopy (STORM) (Huang et al. 2009) will certainly help define the organization and roles of these domains. At the molecular level, analysis of Rab5 effector function including the recent reconstitution of the Rab5-dependent machinery for endosomal fusion, provided several important insights into Rab domain assembly (Ohya et al. 2009). The emerging model is that Rab domains, constituted by oligomeric complexes of effectors and associated proteins (McBride et al. 1999), undergo self-assembly as a consequence of the high degree of cooperative binding between the components of the system (see Fig. 9.4). For example, Rabaptin-5 (i.e., RABEP1), a Rab5 effector, forms a complex with Rabex-5 (i.e., RABGEF1), a Rab5 exchange factor, thereby amplifying Rab5 activity and recruitment (positive feedback loop) (Horiuchi et al. 1997; Lippe et al. 2001); Rab5 stimulates Vps34p150 (i.e., VPS34-PIK3R4) (Shin et al. 2005), which synthesizes PtdIns(3)P and contributes to the recruitment of other effectors containing PtdIns(3)P-binding Fab 1, YOTB (i.e., hypothetical C. elegans protein ZK632.12), Vac 1, and EEA-1 (FYVE) domains (e.g., EEA-1, Rabenosyn-5 [i.e., ZFYVE20]); also, SNAREs contribute binding sites for the recruitment of the Rab5 effectors on the membrane (McBride et al. 1999; Ohya et al. 2009). By requiring such a high number of interactions, the system favors the formation of specific proteolipid Rab-organized domains, which may contribute to the robustness and specificity of the assembly of the membrane tethering and fusion machinery.

Figure 9.4. Cooperativity as a driving principle of Rab5 domain self-assembly. Because of multiple positive feedback loops built in the system, the endosomal recruitment of Rab5 domain components is a cooperative process. Conceptually, an elementary motif is recurrent: the recruitment of a given component (A) enhances the recruitment of another component (B), which further stabilizes (A) on endosomal membrane. (1) On endosome, Rab5-GTP recruits its effector Rabaptin-5, which is associated with the Rab5 GEF Rabex; by stimulating Rab5 exchange of GDP by GTP, Rabex enhances Rab5 recruitment on endosomes (since Rab5-GTP is more stable on endosome than Rab5-GDP): This is one feedback loop. (2) Rab5-GTP recruits its effector Vps34, which produces PtdIns(3)P on endosomes and thereby stabilizes the FYVE-domaincontaining Rab5 effectors (EEA-1 and Rabenosyn-5); in turn, EEA-1 and Rabenosyn-5 stabilize Rab5-GTP on endosomes: It is a second feedback loop. (3) Rab5-GTP recruits EEA-1; EEA-1 interacts with SNAREs, which in turn contribute to stabilizing Rab5 on endosomes (Ohya et al. 2009): It is a third feedback loop.

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Rab Domains and the Directionality of Membrane Traffic Rab domains are sequentially traversed by cargos. For instance, the recycling route is landmarked by Rab5, Rab4, and Rab11 domains, whereas cargo molecules bound to lysosomes are first found in Rab5 and then in Rab7 domains. Understanding how the specificity and directionality of transport from one Rab domain to the other are achieved is crucial. The concept of a Rab cascade where the initial Rab, Rabi, activates and recruits or stabilize the next coming Rab in the route, Rabi+1, has been proposed as a potential mechanism to ensure transport directionality (Zerial and McBride 2001). Effectors harboring distinct binding sites for two consecutive Rabs could in principle support such a Rab cascade (see Fig. 9.5A, B). For example, both Rabaptin-5 and Rabenosyn-5 bind Rab5 and Rab4 independently and can therefore recruit Rab4 to Rab5 domains (Vitale et al. 1998; de Renzis et al. 2002). Similarly, D-AKAP2, which binds both Rab4 and Rab11 could recruit Rab11 to Rab4 domains (Eggers et al. 2009). Another mechanism involves the sequential recruitment of a guanine nucleotide exchange factor (GEF) for the next coming Rab. Originally evidenced in the yeast secretory pathway (Ortiz et al. 2002), this design has been shown to drive the Rab5-to-Rab7 transition (Rink et al. 2005). In the latter example, Rab5 recruits the homotypic fusion and vacuole protein sorting (HOPS) complex, a subunit of which (Vps39) is a GEF for Rab7, therefore contributing to the nucleation of the downstream Rab7 domain. One still has to understand to what extent and how two consecutive Rab domains can mix or segregate (see Fig. 9.5C). Interestingly, a mathematical model of the Rab5-to-Rab7 transition proposed that a negative feedback loop on Rab5 is initiated by Rab7 (Del Conte-Zerial et al. 2008). At the molecular level, this feedback loop may be embodied by a Rab5-GTPase-activating protein (GAP), which is at the same time an effector for Rab7 or a Rab7-dependent inhibitor of a Rab5-GEF. Evidence for the latter has recently been obtained (Poteryaev et al. 2010). In the particular case of the Rab5-toRab7 transition, the inhibition of Rab5 is complete and Rab7 substitutes Rab5 totally in a phenomenon called Rab conversion (Rink et al. 2005). However, one can speculate that in other cases, the two domains could be maintained separately if the next coming domain is pulled away before having fully inhibited the initial domain (for instance, if it contains a motor protein), hence causing budding or tubulation.

Shaping Endosomal Membranes: Bin/Amphiphysin/Rvs (BAR)-Domain-Containing Proteins and ESCRT Endosomal sorting relies on membrane deformation processes like budding and fission of vesicles and tubules. Although membranes are only 40-Å-thin phospholipidic structures, they do not bend easily. The energy required for bending a typical bilayer is about 20 times higher than what is provided by thermal fluctuations. Therefore, generating membrane curvature requires active mechanisms and machineries ad hoc (Kozlov 2010). Two main families of proteins can induce and/or stabilize membrane curvature on endosomes: BAR-domain-containing proteins as well as membrane coats and their adapters (Zimmerberg and Kozlov 2006). BAR domains are crescent shaped; they bind membrane as dimers and are found in a large variety of proteins including the endosomal proteins APPL1&2 and the sorting nexin (SNX) family (see Fig. 9.3). Owing to their curved shape, BAR domains induce (by induced fit) or stabilize membrane curvature. In vitro, they can assemble into helical filaments that support the formation of membrane tubules in a similar way as dynamin, another membrane-shaping protein involved in the formation and scission of a vesicle neck (Shimada et al. 2007; Frost et al. 2008). The generation of endosomal tubular carriers en route to the TGN from LEs involves two BAR domain proteins, SNX1 and SNX2, components of the pentameric retromer complex (Seaman

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(A)

(B)

(C)

Figure 9.5. Directional propagation of Rab domains. (A) The first acting Rab (Rabi) possesses an effector (Eff.) with a binding site for the second acting Rab in the cascade (Rabi+1) in its GTP form. Rabi+1-GTP binding to Rabi effector stabilizes Rabi+1-GTP on endosomes and favors the nucleation of its domain. (B) The first acting Rab (Rabi) possesses an effector with a GEF activity for the second acting Rab (Rabi+1), resulting in the stable recruitment of Rabi+1-GTP on endosomes. (C) Rabi+1-GTP recruits a GAP for Rabi, thereby destabilizing Rabi domain (i.e., conversion process) or segregating the two domains (i.e., domain segregation).

et al. 1998). Similarly, SNX4 is a potential component of the tubulation machinery required for recycling from EEs to the plasma membrane (Traer et al. 2007). Coat proteins such as clathrin, and their adapters, influence curvature by acting as local molds for membranes after polymerization. In all cases, membrane bending must be coupled to force generation in order to generate and pinch off tubules. SNX4 associates with the minus-end motor dynein and the nucleation-promoting factor Wiskott–Aldrich syndrome protein and suppressor of cAMP receptor (SCAR) homolog (WASH), which activates

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Arp2/3 and actin filaments branching has been specifically found on endosomes where it associates with the retromer (see Fig. 9.3) (Gomez and Billadeau 2009). Depletion of WASH by RNAi reduces F-actin on endosomes and induces the formation of long tubules consistent with a role of actin nucleation in fission (Derivery et al. 2009; Gomez and Billadeau 2009). Interestingly, assembly of the machinery for membrane deformation may depend on Rab domains. For instance, the recruitment of the BAR-domain-containing protein adaptor protein, phosphotyrosine interaction, pleckstrin homology (PH) domain, and leucine zipper containing 1 (APPL1) on endosomes depends on Rab5 (Miaczynska et al. 2004; Zoncu et al. 2009) and endosomal recruitment of the retromer requires the sequential formation of Rab5 and Rab7 domains (Rojas et al. 2008). The ESCRT, which catalyses inward budding and the biogenesis of ILVs, is an essential machinery for membrane deformation. ESCRT is conserved from a subset of Archae to mammals and is composed of five subcomplexes ESCRT-0, -I, -II, -III, and Vps4-Vta1, as well as several associated proteins (Wollert et al. 2009b). ESCRT-0, -I, and -II interact directly with ubiquitinated cargos through various structural motifs. ESCRT-0 (composed of the two subunits Hrs and STAM) associate with clathrin to form flat endosomal domains involved in the capture and clustering of cargos. ESCRT-I and -II cooperate to deform the membrane and cause budding of the nascent ILVs (Wollert and Hurley 2010). Interestingly, the ESCRT-III subunit Snf7 can polymerize when overexpressed and form filamentous spirals (Hanson et al. 2008). As for dynamin or BAR domain proteins, these spirals are proposed to catalyze membrane tubulation and scission. Another subunit, the ATPase Vps4 acts after membrane scission to disassemble the filaments and recycle ESCRT-III proteins, a mechanism that is reminiscent of the dissociation of the SNARE complex by the ATPase NSF after vesicle fusion (Wollert et al. 2009a). Although ESCRT plays major roles in ILV formation, lipids may be instrumental as well by contributing directly to membrane deformation or by localizing protein machineries. The cone-shaped lipid LBPA has been implicated in ILV formation in vitro as well as in cells where it interacts with ESCRT-I and -III via the LBPA-binding protein Alix (Matsuo et al. 2004). Similarly, PtdIns(3,5)P2 may be important for compartmentalizing the ESCRT machinery (Whitley et al. 2003).

Compartmentalization of the Signaling Machinery In addition to their sorting functions, endosomes play important and specific roles in signal initiation, propagation, or processing (Sorkin and von Zastrow 2009). A long list of miscellaneous signaling molecules have been found to function on endosomes including activated surface receptors, kinases (e.g., PKC, AKT, c-Src, extracellular signal-regulated kinase [ERK]1&2, and mammalian target of rapamycin [mTOR]), small GTPases (e.g., H-Ras, Rheb), subunits of heterotrimeric G proteins, and transcription factors (e.g., STAT3). Hence, the modular organization of the endosomal system allows the spatial segregation of signaling molecules. Assembly of signaling domains depends on Rab domains assembly. For instance, AKT is recruited to endosomes via its interaction with the endosomal adapter and Rab5 effector APPL1 (Mitsuuchi et al. 1999; Miaczynska et al. 2004). Similarly, endosomal domains enriched in PtdIns(3)P produced by Vps34 under the control of Rab5 contribute to the recruitment of FYVE-domain-containing signaling proteins. Among many examples are SMAD anchor for receptor activation (SARA) and endofin, two pivotal components of the transforming growth factor-β (TGF-β) signaling pathway; the ubiquitin protein ligase caspase-8 and -10-associated RING protein 2 (CARP2) is also recruited on PtdIns(3)P domains where it interacts with the internalized tumor necrosis factor (TNF)-receptor complex and limit the intensity of TNF-induced

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nuclear factor “kappa-light-chain-enhancer” or activated B-cells (NF-κB) activation (Liao et al. 2008) (see Fig. 9.3). Several molecular scaffolds contribute to the assembly of signaling platforms on endosomes. For example, the lipid-raft-anchored complex formed by the proteins p18, p14, and MEK1 partner 1 (MP1) recruits the components of the mitogenactivated protein kinase cascade, bringing together MEK1 and its substrate ERK1&2 (Teis et al. 2006); the endocytic adapters for GPCRs, β-arrestins recruit Raf-1, MEK1, and ERK1&2 on EEs or REs. Interestingly, APPL1 or SARA domains are on EEs; the p18/ p14/MP1 domain assembles on late, Rab7-positive endosomes, whereas H-Ras and βarrestin scaffolds are found on Rab11 endosomes. The endosomal signaling machinery can generate very specific signaling outputs. For instance, downregulation of the Rab5 effector APPL1 in zebrafish impairs cell survival by specifically inhibiting the Akt-glycogen synthase kinase 3 beta (GSK-3β) signaling branch. This phenotype can only be rescued by endosome-localized APPL1, demonstrating the importance and the specificity of Akt signaling from endosomes (Schenck et al. 2008). Another example is the activation of the transcription factor STAT3 by the hepatocyte growth factor (HGF). First, the localization of HGF receptor c-MET to endosomes is necessary for optimal HGF-mediated stimulation of ERK1&2 and in addition, c-MET must accumulate in a perinuclear endosomal compartment in order to trigger the nuclear translocation of its effector STAT3. Interestingly, the amplitude of STAT3 translocation is lower when initiated from endosomes than when activated from the cell surface by oncostatin M (Kermorgant and Parker 2008).

ENDOSOMES AS ORGANIZERS OF CELL AND TISSUES Endosomes and Cellular Homeostasis At the crossroad of vesicular traffic among plasma membrane, biosynthetic pathway, and lysosomes, the endosomal system is inherently connected to multiple cellular activities. For instance, via the endosomal system, a cell can spatially redistribute its membrane components from one domain of its surface to the other and thereby control the local assembly of adhesion and cell junction complexes, which influences cell migration and polarity. Similarly, endosomes regulate integrated physiological functions (e.g., glycemia by controlling the number of glucose transporters at the surface of adipocytes and myocytes, water absorption by controlling the number of aquaporins in kidney, learning and memory by controlling the number of glutamate receptors in the central nervous system). In addition, endosomes form an intracellular compartment with very specific physicochemical properties (e.g., low pH, controlled volumes according to a spatiotemporal gradient, see section on “Dynamics of the System”) that are instrumental to various cell functions. For instance, with values ranging from 6.5 to 5.5, the acidic pH of endosome, contributes to the dissociation of specific ligand–receptor complexes resulting in making nutrients available (e.g., iron, LDL [Brown et al. 1983; Dautry-Varsat et al. 1983]) or in the inactivation of the receptor signaling activity. The activity of multiple proteases can be spatially restricted to endosomes because their activity is optimal within the endosomal pH range. Owing to their structural properties and role as signaling platforms, endosomes and endosomal subcompartments are important nodes of signaling networks, acting as signal initiators, modulators, integrators, or effectors of cellular signals. It is therefore not surprising that every trafficking step (e.g., coat formation and disassembly, vesicles docking, fusion and fission, actin polymerization, transport along microtubules) is tightly

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controlled as shown by the high number of protein kinase substrates among the components of the endocytic machinery (Liberali et al. 2008). Recent functional genomics studies confirmed this tight and complex control of the endosomal system by other cell functions. RNAi screens in mammalian cells coupled to image-based profiling of the endosomal system revealed that numerous signaling and metabolic pathways control endosomal organization (Pelkmans et al. 2005; Galvez et al. 2007; Collinet et al. 2010). For example, the PtdIns(3,4,5)P3-Akt-mTOR signaling module, which acts as a sensor of metabolic activity and controls cell growth, was a significant hit in all studies (noteworthy, TOR itself has been localized to endosomes both in yeast, fly, and mammals). In addition, the recent genome-wide, multiparametic profiling of the endocytic system revealed significant roles of TGF-β, Notch, and Wnt pathways in controlling endosome function and spatial distribution (Collinet et al. 2010). As systems biology approaches help in deciphering the connections between endosomal and signaling systems at the molecular level, recent studies integrating cellular functions are bringing into prominence the role of endosomes and their dynamics in cell and tissue morphogenesis. We will first illustrate these two concepts at the cellular level with the process of cytokinesis and then at the tissue level with the formation of tubes from epithelial sheets.

Endosomes and Membrane Remodeling: Cytokinesis Cytokinesis is the last stage of mitosis corresponding to the separation of the two daughter cells. After partition of sister chromatids, the cleavage furrow ingresses leaving the postmitotic cells connected by a thin bridge containing antiparallel arrays of microtubules (midbody) before its final cut or abscission. The mechanisms of abscission are largely unknown but endosomes and actin accumulate at the midbody just before abscission and seems to have a major role in this process. The small GTPases Rab11, Rab35, and ARF6 localize at the cleavage furrow and are required for abscission timing and completion (Steigemann and Gerlich 2009). Rab11 endosomes are recruited to the intercellular bridge via the Rab11 and ARF6 effector Rab11 family interacting protein 3 (FIP3)/nuclear fallout. In addition, the multifunctional protein and BIR-containing ubiquitinconjugating enzyme (BRUCE) acts as a midbody targeting and clustering factor for Rab11 and Rab8 endosomal vesicles. There, the endosomal domains devoted to recycling may contribute as membrane reservoir for the plasma membrane expansion required for abscission (Lecuit and Wieschaus 2000; Wilson et al. 2005; Boucrot and Kirchhausen 2007; Pohl and Jentsch 2008) or as local regulators of actin dynamics by ferrying RhoGEF2, an activator of RhoA and consequently of actin polymerization (Cao et al. 2008). This concept of endosomes conveying cargo, membrane, or specific machineries to the midbody is also supported by the proposed role of Rab35. The latter stimulates the assembly of PtdIns(4,5)P2 domains at the zone of abscission where it determines the local recruitment of septins, which are required for bridge stabilization and completion of abscission (Kouranti et al. 2006). Recently, the endosomal ESCRT has been directly implicated in cytokinesis as well. A component of the ESCRT-I complex, TSG101, and the associated protein Alix are present at the midbody where they interact with the centrosome protein 55 (CEP55) and are also required for the completion of abscission (Carlton and Martin-Serrano 2007). One role of ESCRT-I seems to be the recruitment of the ESCRT-III complex. The latter is essential for midbody localization of the microtubulesevering protein spastin but, because of its ability to form circular filamentous arrays, is also an attractive candidate to drive the closure of the bridge (Hanson et al. 2008).

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Shaping Tissues By transporting membranes and serving as local platform for assembly of specific molecular machineries, endosomal trafficking plays essential roles in membrane remodeling of the cell. However, it is becoming clear that the endosomal system exerts its influence at higher organizational levels as well, regulating for instance morphogenesis of tissues. This can be illustrated by the importance of endocytic trafficking during epithelial branch formation in the Drosophila trachea. Tubular organs like tracheae are instrumental for transport of gas, nutrients, and cells within an organism. They include the blood and lymph circulatory system, mammalian lungs and kidneys, and the tracheal respiratory system of insects. In the dorsal trunk of Drosophila airways, Wingless promotes the expression of the transcription factor Spalt, which inhibits cell intercalation and thereby preserves a wide diameter lumen. In the tracheal branches where Spalt is not expressed, cells intercalate and eventually convert most of their intercellular adherens junctions into autocellular junctions, resulting in single-cell-enclosed lumens. Interestingly, Rab11 and dRip11 are the key targets of Spalt in this process. In the trunk, elevated Rab11-dependent recycling mediates the surface delivery of E-cadherin complexes, promoting intercellular adhesion to oppose cell intercalation and resulting in the formation of multicellular lumens (Shaye et al. 2008). Hence, signaling molecules like Wnt/Wingless control tissue formation by regulating the activity of the endosomal system. This is reminiscent of the recent functional genomics survey that identified Wnt, TGF-β, and Notch pathways as endocytic regulators in mammalian cells (Collinet et al. 2010). Interestingly, these signaling pathways are established regulators of epithelial–mesenchymal transition (EMT) (Thiery et al. 2009), thus hinting at another link between endosome function and tissue morphogenesis. Other lines of evidence strengthen the role of endosomes in tissue patterning: For instance, in the tracheal system of Drosophila, a burst of endocytic activity is responsible for the lumen clearance (Tsarouhas et al. 2007); endosomal activity is also essential for the establishment of morphogen gradients responsible of the patterning of tissues in developing embryos (Kicheva et al. 2007; Yu et al. 2009).

FUTURE PERSPECTIVES In summary, the emerging picture of the endosomal system is that of a highly interconnected, multifunctional membrane-bound compartment. First, membrane sorting and cell signaling are so inherently intertwined that one must consider the endosomal system as a rightful signaling module. Endosomal membranes are major effectors of cell decisions (e.g., cell migration, cell division, or differentiation) by their ability to control in space and time the assembly of multiprotein complexes and the redistribution of membranes and their associated machineries. The segregation of endosomal subdomains calls for revisiting transport and signaling reactions at a higher level of spatial resolution than done so far. It will be important to explore the rules governing the assembly of signaling complexes not just on endosomes but preferably at the level of the specific membrane domains composing them. This is currently not the level of resolution that is mostly investigated. It is fascinating to consider that the organization of the endosomal system at the level of one cell within a tissue will reflect on the organization of the tissue itself, influencing the development of organisms but also pathogenesis (e.g., cancer, metastasis). The endeavor of systems biology, by implementing quantitative, global, and unbiased approaches at the cell or tissue level, will certainly continue to reveal an even more integrated system. Ideally, data sets from RNAi-based screens should be completed by large-scale quantitative proteomics

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studies of endosomes, but the existence of multiple endosomal populations together with the inherent contaminations of present subcellular fractionation techniques have been major limitations so far. At the molecular level, endosomes act as motile and multifunctional platforms for the assembly of diverse machineries. Understanding the logic of assembly and function of these nanomachines will require two main types of “bottom-up” strategies: (1) a synthetic strategy aiming at the in vitro reconstitution of the molecular machineries on endosomes as recently accomplished for the Rab-dependent fusion and the ESCRT-dependent fission machineries (Ohya et al. 2009; Wollert et al. 2009a), and (2) direct observation using the newly developed super-resolution microscopy techniques (e.g., STED, PALM, STORM) (Huang et al. 2009). These approaches, which can also be combined, will contribute to important breakthroughs in the near future. Whatever level is considered (i.e., tissue, cell, or molecular assembly), the “Holy Grail” will be to identify the design principles or “laws” that govern the structural and functional organization of the endosomal system.

ABBREVIATIONS APPL1 BAR BRUCE CARP2 CCP CCV CLIC EEs EEA-1 EGF EMT ERK ESCRT FIP3 FYVE GAP GEF GPCR GSK-3β ILV

adaptor protein, phosphotyrosine interaction, PH domain, and leucine zipper containing 1 bin/amphiphysin/Rvs baculovirus IAP repeat (BIR)-containing ubiquitin-conjugating enzyme caspase-8 and -10-associated RING protein-2 clathrin-coated pit clathrin-coated vesicle clathrin-independent carrier early endosomes early endosomal antigen-1 epidermal growth factor epithelial-mesenchymal transition extracellular signal-regulated kinase endosomal sorting complex required for transport RAB11 family interacting protein 3 Fab 1, YOTB (hypothetical C. elegans protein ZK632.12), Vac 1, and EEA1 GTPase-activating protein guanine nucleotide-exchange factor G protein-coupled receptor glycogen synthase kinase 3 beta intralumenal vesicle

LBPA lysobiphosphatidic acid LDL low-density lipoprotein LE late endosome M6PR mannose-6-phosphate receptor MP1 MEK partner 1 MVB multivesicular body NSF N-ethylmaleimide-sensitive factor PALM photoactivation localization microscopy PH pleckstrin homology RE recycling endosome RNAi RNA interference SARA SMAD anchor for receptor activation SNAREs soluble NSF attachment protein (SNAP) receptors SNX sorting nexin STED stimulated depletion STORM stochastic optical reconstruction microscopy TFR transferrin receptor TGN trans-golgi network TIP47 tail-interacting protein of 47 kD TOR target of rapamycin WASH Wiskott-Aldrich syndrome protein and SCAR homolog

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LYSOSOMES AND PHAGOSOMES Guillaume Goyette Michel Desjardins

DEFINITION Lysosomes are dense spherical organelles functionally defined as the end station of the endocytic pathway where the bulk of the internalized molecule degradation is performed. These structures, originally characterized at the biochemical level by de Duve and his colleagues (de Duve 1983), are filled with hydrolases and display a low pH in the range of 4.5–5.0. In the context of the other endocytic organelles, lysosomes have often been presented as structures showing large amounts of lysosomal membrane proteins (mainly LAMP-1, LAMP-2, and LIMP-2) and lacking small GTPases of the rab family. The presence of membrane microdomains in lysosomes is uncertain, and suggested, so far, by indirect observations reporting the association of well-known microdomain markers to this organelle. Nevertheless, the study of related organelles along the phagocytic pathway (early phagosomes, late phagosomes, and phagolysosomes) has shed light on molecular mechanisms related to microdomain structure and functions that may also apply to the endocytic pathway. Phagosomes are formed at the surface of a variety of cells, including macrophages, neutrophils, and dendritic cells, during the internalization of large particles such as dead cells, bacteria, or parasites, as well as inert particles. After their formation, phagosomes are remodeled through a complex maturation process enabling the acquisition of the microbicidal properties required for their role in the killing of microorganisms and the processing of their antigens for presentation on major histocompatibility complex (MHC) molecules. Thus, phagosomes play key roles in both innate and adaptive immunity. Coordinated interactions between these two processes are essential to mount an effective response against a variety of pathogens causing infectious diseases. Recent studies have highlighted the presence of microdomains on the membrane of phagosomes, and showed their significant role in the ability of this compartment to restrict the growth of pathogenic microorganisms. This chapter discusses the biological context that led to the identification of membrane microdomains on phagosomes, their potential role in innate immunity, and the strategies evolved by microorganisms to alter their functions.

HISTORICAL PERSPECTIVE The existence of different types of internalization processes, ranging from the specific uptake of fluids and small molecules by endocytosis, the sampling of the external milieu

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by macropinocytosis, and the engulfment of large particulate materials by phagocytosis, have maximized the ability of cells to fully interact with and sample their harsh environment (Conner and Schmid 2003). These complex pathways intersect at a key organelle of eukaryotic cells, the lysosome, where the final degradation (or storage) of the internalized molecules and the recycling of their end products occur. The lysosome has been the object of a large number of studies that historically merged the use of biochemical and morphological approaches. The pioneering work of Christian de Duve, George Palade, Alex Novikoff, Keith Porter, and several others have contributed to define the initial molecular properties of the lysosome. This early interest in this dense spherical organelle filled with hydrolases and displaying a low pH has contributed to setting the foundations for the study of cellular organelles, and the emergence of modern cell biology. De Duve and Palade, together with Albert Claude, who pioneered the use of cell fractionation for the study of cell functions, were awarded the Nobel Prize in Physiology or Medicine in 1974. The internalization of a variety of molecules at the cell surface plays a key role in cell homeostasis and the regulation of several of its functions. The control (and termination) of signal transduction through ligand–receptor internalization; nutrition via the uptake of a variety of molecules; and the initiation of a sustained immune response following the sampling of the external milieu are only a few of the cellular mechanisms regulated by internalization processes (Conner and Schmid 2003; Sorkin and von Zastrow 2009). Endocytosis, the general mechanism by which cells internalize fluids and small molecules, is characterized by the passage of ingested materials in a succession of organelles displaying complex and diverse morphological and biochemical features (see also Chapter 9 on endosomes). The textbook view of the endocytic pathway describes the successive passage of molecules destined for degradation in early endosomes, followed by their transit through late endosomes, and their transfer to lysosomes, the end station that specializes in the cleavage of molecules (and their permanent storage in the case of nondegradable materials). Lysosomes play a key role in the biogenesis of related organelles such as phagolysosomes, where cell debris and potentially harmful pathogens are killed and/or degraded, and autophagolysosomes involved in the recycling of self-components in stress conditions, as well as in the clearance of certain pathogens and the initiation of an effective immune response (Levine and Deretic 2007). Phagocytosis, the process that describes the engulfment of large particulate materials at the cell surface and leads to the formation of phagosomes, has acquired its lettres de noblesse through the seminal work of Elie Metchnikoff, who was the first scientist to link this process with immunity. Metchnikoff was awarded the Nobel Prize in Physiology or Medicine in 1908 for his work on phagocytosis, together with Paul Ehrlich, an outstanding immunologist who contributed to finding a cure for syphilis, among other accomplishments. Internalization of large particles such as dying cells or cell fragments, as well as microorganisms like bacteria and parasites, occurs by phagocytosis, a process often seen as a specialized form of endocytosis (Haas 2007; Paidassi et al. 2009). In mammals, this process plays key roles in embryogenesis, tissue homeostasis, and immunity. Phagocytosis is a receptor-mediated process initiated by the binding of a variety of ligands to a wide array of receptors at the cell surface. Engagement of phagocytic receptors triggers cascades of signaling events enabling local reorganization of cortical actin at sites of internalization (Insall and Machesky 2009), leading to the formation of phagocytic cups, the extension of pseudopodia, and the engulfment of the particle (see also Chapter 12 on the actin cytoskeleton). Early phagosomes rapidly interact with endocytic organelles, promoting their remodeling and the acquisition of the lytic properties required for the killing and/or deg-

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radation of their content (Kinchen and Ravichandran 2008). In mammals, although a wide variety of cell types are able to perform phagocytosis, the most studied are specialized cells of the immune system including macrophages, neutrophils, and dendritic cells, often referred to as “professional phagocytes” (Rabinovitch 1995). While nonprofessional phagocytes mainly internalize apoptotic cells in their immediate environment, professional phagocytes are mobile and play a key role in the clearance of bacteria and other pathogens at sites of infection. By killing potentially harmful microorganisms and initiating a sustained immune response through the processing of microbial peptides and their presentation on molecules of the MHC, phagocytes play key roles in both innate and adaptive immunity (Jutras and Desjardins 2005). Therefore, phagosomes have been the focus of great attention in the last 20 years. Surprisingly, very little is known about the presence and the potential nature of the membrane microdomains of this organelle. It should be noted that an early study reported that late endocytic organelles (and lysosomes) are unlikely to display a significant amount of membrane microdomains (raft-poor), due to the involvement of the Niemann-Pick C1 (NPC1) protein in the recycling of cholesterol from endosomes to the cell surface (Lusa et al. 2001). Nevertheless, flotillin-1, a protein shown to accumulate in membrane microdomains at the cell surface, as well as on the membrane of phagosomes (see below), has also been localized to small foci of the lysosome membrane by immunoelectron microscopy (Kokubo et al. 2003), suggesting that it may be associated to some forms of membrane microdomains. The cannabinoid type I receptor (CB1R), a seven-transmembrane domain G-protein-coupled receptor (GPCR), was shown to be present in the detergentresistant membranes (DRMs; see also Chapter 4 for discussion of DRMs) of a total cell extract, and to localize to the cell surface, as well as to endocytic organelles including lysosomes (Sarnataro et al. 2005). The direct demonstration of the presence of this protein in lysosome DRMs remains to be established. Despite the limited knowledge of the lysosome microdomains, potential mechanisms regulating the organization and the functional properties of these evasive structures can be inferred from the study of a related organelle, the phagosomes and phagolysosomes.

THE PHAGOSOME PROTEOME The refinement of mass spectrometry in the 1990s represents a key event that enabled the proteomics era, leading to new ways of studying and characterizing tissues, cells, and organelles by the identification of their constituents. Phagosomes and the Golgi complex were the first cellular organelles characterized by mass spectrometry (Bell et al. 2001; Garin et al. 2001). Proteomic analyses provided significant insights into the structural organization of phagosomes and the roles played by this organelle in the clearance of microbes and the complex cellular processes involved in antigen processing and presentation. Identification of organelle function based on the characterization of its protein constituents has always been a key approach used in cell biology. Identification of phagosomal proteins by various techniques involving Edman degradation led to the identification of a handful of proteins revealing, for example, that phagosomes are organelles that mature into phagolysosomes by the sequential acquisition of small GTPases of the rab family (Desjardins et al. 1994a, b). By applying more sensitive methods taking advantage of the separation of proteins in two-dimensional gels and their identification by mass spectrometry using matrix-assisted laser desorption/ionization (MALDI–TOF–MS), we were able to identify around 125 proteins on phagosomes isolated from J774 macrophages following the internalization of latex particles (Garin et al. 2001). Some of these proteins where

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already known as components of the endosomal/phagosomal pathway, such as hydrolases, several subunits of the vacuolar proton pump, and series of proteins involved in membrane fusion. The presence of other proteins without any known involvement in phagocytosis provided direct insight into phagosome functions. The first intriguing subset of proteins consisted of a group of proteins usually observed on the endoplasmic reticulum (ER), including calreticulin, calnexin, GRP78, endoplasmin, and Erp29 (see Chapter 7 on the ER). The presence of these proteins added to the growing evidence that intracellular organelles were rapidly recruited to the cell surface during phagocytosis to potentially provide some of the membrane needed for the formation of nascent phagosomes (see below). The second newly discovered group of proteins included flotillin-1, stomatin, and prohibitin, all shown to be present in membrane microdomains (Terashima et al. 1994; Bickel et al. 1997; Lang et al. 1998; Snyers et al. 1999). This finding had a profound effect on our interpretation of phagosome membrane organization. Rather than being an organelle where lipids and proteins are randomly distributed, the phagosome membrane displayed foci of functional specialization, enabling the spatial regulation of phagosome functions. However, the biochemical and functional nature of the microdomains present on the phagosome membrane, as well as the mechanisms linked with their assembly, remains largely unknown. Flotillin-1 (also referred to as Reggie-2) was first observed in microdomains on the plasma membrane of neurons and 3T3-L1 mouse fibroblasts differentiated into adipocytes (Bickel et al. 1997; Lang et al. 1998). This protein, maintained in evolution from bilateria to vertebrates, is ubiquitously expressed in mammalian cells. Functional studies first indicated that flotillin-1 plays a role in the regeneration of axons (Schulte et al. 1997). The role of flotillin-1 on phagosomes is still unknown. On the plasma membrane, this protein, together with flotillin-2 (Reggie-1), plays a role in the assembly of signaling platforms activating small GTPases involved in actin dynamics (Stuermer 2010). Whether these proteins play a similar role on phagosomes remains to be established. However, results indicating that flotillin-1 accumulates on maturing phagosomes suggest that functional flotillin-1-enriched platforms are unlikely to be recruited directly from the plasma membrane during phagocytosis (Dermine et al. 2001).

Microdomains on the Phagosome Membrane The phagosome, unlike most cellular organelles, is formed de novo at the cell surface following the internalization of a large particle. Studies indicate that a variety of cellular organelles can fuse with the phagocytic cup, providing some of the membrane present on early phagosomes (Jutras and Desjardins 2005). The fusion of recycling endosomes with phagocytic cups was shown to involve the v-SNARE molecule VAMP-3 (Bajno et al. 2000) and the small GTPase protein ARF6 (Niedergang et al. 2003). On the other hand, late endocytic organelles were shown to fuse and deliver membrane to forming phagosomes in a VAMP-7-dependent manner (Braun et al. 2004). Morphological analyses also indicated that the ER interacts with the membrane of phagosomes containing latex beads or the intracellular parasite Leishmania donovani before complete engulfment of this pathogen by macrophages (Gagnon et al. 2002). In that context, phagocytosis was shown to be partially inhibited following RNAi treatment for the ER SNARE Syntaxin-18 (Hatsuzawa et al. 2006). Altogether, these data clearly indicate that early phagosomes are made from a pool of membranes originating from various organelles (Fig. 10.1). It is not known whether these membranes rapidly mix to generate a more “homogeneous” structure, or whether they are kept as distinct foci on the phagosome membrane for a significant

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THE PHAGOSOME PROTEOME

ENDOSOME EEA1 Rab 5 Rab 7

V-ATPase MEMBRANE RAFT

PLASMA MEMBRANE α

Fc Receptor Toll-like Receptor

Proteases

Phagosome Cargo

Cathepsins A,B,D,L,S,Z

Lipases Sec61

Actin

β Flotillin-1 Stomatin Prohibitin

Ceramidase Lysosomal acid lipase

Calnexin Calreticulin Proteasome

G Proteins

LAMP1 LAMP2

Tap1/ Tap2

LIMP II LYSOSOME

ENDOPLASMIC RETICULUM MHC class I Tap1/Tap2 MHC class I

Figure 10.1. The phagosome membrane is composed of different microdomains. The phagosome membrane is possibly composed of membrane domains derived from different sources including the plasma membrane (yellow), endosomes (blue), lysosome (purple), and the endoplasmic reticulum (green). Furthermore, cholesterol-enriched membrane rafts form gradually on maturing phagosomes and sequester specific proteins (red), suggesting that functional properties are sequestered in different spatial foci. The mixed origin of the phagosome membrane is proposed to enable the link between the properties needed to degrade antigens in the lumen, and their handling for further processing by the proteasome. The processed antigens are then retrotranslocated either back to the phagosome lumen or to the ER for loading on MHC class I molecules.

period. Analyses of the maturation of phagosomes into phagolysosomes have initially demonstrated that various isoforms of the small GTPase molecules rab5 and rab7 are sequentially acquired (Desjardins et al. 1994b). This process coincides with the ability of phagosomes to fuse sequentially with early and late endosomes (Desjardins et al. 1997). Interestingly, different rab proteins were shown to be present on different foci of a given endosome (Sonnichsen et al. 2000), suggesting that subdomains enriched for rab5 or rab7 might also exist on the phagosome membrane. While these studies strongly supported

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the notion that phagosomes display multiple membrane microdomains, the functional significance, the composition of these structures, and whether they biochemically correspond to the structures isolated in DRMs following the solubilization of organelles in nonionic detergents remains to be elucidated. The first evidence for the presence of morphologically distinct membrane domains on phagosomes were provided by immunofluorescence labeling of the markers LAMP-1 and flotillin-1 (Dermine et al. 2001). While the labeling for LAMP-1, an abundant type-1 membrane protein present on late endocytic and phagocytic organelles, was distributed all around the phagosome membrane, the labeling for flotillin-1 was present in a punctate pattern. The distribution of these two markers to different membrane domains was confirmed by the extraction of DRM fractions from isolated phagosomes (Dermine et al. 2001). After solubilization with Triton X-100 and gradient purification, flottilin-1 was present on the top of the gradient, while LAMP-1 remained in the sample loaded at the bottom. Two-dimensional gel electrophoresis of phagosome proteins followed by massspectrometry analyses led to the initial identification of a handful of proteins specifically localizing to DRMs, including actin, subunits of the V-ATPase proton pump involved in the acidification of phagosomes, and heterotrimeric G-protein subunits. Although rudimentary, this list of proteins indicated that multiple functions were likely to take place on membrane microdomains, reinforcing the concept of spatial regulation of phagosome functions. The ganglioside GM1 was also localized to foci of the phagosome membrane and DRMs (Dermine et al. 2001). Interestingly, GM1 and flotillin-1 were not observed in the same region of the phagosome membrane, suggesting that multiple types of membrane domains are present on this organelle (Dermine et al. 2005). Membrane microdomains, referred to at the time as lipid rafts, were first identified as a platform where signaling molecules accumulated, highlighting the role of these structures in the control of signal transduction (Harder and Simons 1997). Very little is known about the role and composition of phagosome membrane microdomains. Since phagosomes display similarities with endosomes and lysosomes (Desjardins et al. 1994a), it is possible that the biogenesis and the nature of phagosome microdomains might share properties with similar structures present on endocytic organelles. Early studies on the membrane microdomains of endocytic organelles indicated that high amounts of cholesterol, sphingomyelin, and glycolipids were preferentially observed in early endosomes (Kobayashi et al. 1998; Kobayashi et al. 2001), suggesting that they might inherit these structures from the plasma membrane during endocytosis. In contrast, morphological analyses of phagosomes showed that the microdomain marker flotillin-1 accumulated late on these organelles, during phagolysosome biogenesis (Dermine et al. 2001). Large-scale analyses using proteomics have also confirmed the prevalence of microdomain markers on late phagocytic structures (Goyette et al., submitted). Lysosomal microdomains were described in the context of chaperone-mediated autophagy (CMA). CMA is a process by which cells under nutritional stress target cytosolic proteins to lysosomes for degradation (Dice 2007). This process is initiated by binding of substrate proteins to the constitutively expressed form of heat shock protein of 70 kDa through a specific amino acid sequence (Lys-Phe-Glu-Arg-Gln) which is called the KFERQ motif and is present on around 30% of the cytosolic proteins. The association of this complex with the multimeric form of LAMP-2a, a lysosomal membrane integral protein, is followed by an unfolding step enabling the translocation of the substrate in the lysosomal lumen and is degradation by proteases (Dice 2007). The formation of multimeric LAMP-2A complexes was shown to occur by the dynamic transfer of these proteins outside of cholesterol-enriched subregions of the lysosomes (Kaushik et al. 2006). Arguably, the presence of microdomains and high amounts of LAMP2 on the phagosome membrane suggests that a similar process might occur on this organelle.

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HOST–PATHOGEN INTERACTIONS Elements suggesting that phagosome microdomains are involved in key aspects of the immune properties of this organelle came from observations that three unrelated microorganisms, the parasite Leishmania donovani, the proteobacteria Brucella abortus, and the actinobacteria Mycobacterium tuberculosis target these structures in their strategy to survive in mammalian host cells. They appear to do so by interfering with the assembly and/or directly inducing the disassembly of microdomains on phagosomes (ArellanoReynoso et al. 2005; Dermine et al. 2005; Welin et al. 2008).

Brucella abortus Brucella, a gram-negative bacteria, is the causative agent of brucellosis. The survival strategy of this intracellular pathogen involves the inhibition of phagosome fusion with endocytic organelles, and the biogenesis of an ER-derived organelle where Brucella replicates (Celli and Gorvel 2004). Brucella’s ability to inhibit phagosome–lysosome fusion is linked to the presence of a polysaccharide in the envelope of the bacteria, cyclic beta1,2-glucans (CbetaG), which interacts with “lipid rafts” to disorganize these structures (Arellano-Reynoso et al. 2005). This polysaccharide displays structural similarities to cyclodextrins, which are used to extract cholesterol from membranes. As a result, Brucella mutants lacking CbetaG fail to inhibit phagosome–lysosome fusion and do not replicate in their host cells.

Leishmania donovani Studies indicated that Leishmania donovani can modulate the functional properties of phagosomes and inhibit their fusion with late endocytic organelles, a process required for the transfer of hydrolases to phagosomes and the degradation of their content (Desjardins and Descoteaux 1997). Although this process was shown to involve the parasite surface molecule lipophosphoglycan (LPG), the molecular mechanisms involved in the fusion inhibition are still poorly understood. However, it is clearly established that LPG alters several of the phagosome functional properties. The first phagosome property shown to be altered by LPG was the ability to mount an efficient oxidative response (McNeely and Turco 1990). This inhibition was later shown to be associated with the impaired recruitment and assembly of the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase complex on phagosomes (Lodge et al. 2006). LPG was also shown to affect the association of actin with phagosomes. Active depolymerization/polymerization of the actin network plays a key role in phagocytosis (Chimini and Chavrier 2000). The engagement of a variety of surface receptors at the onset of phagocytosis triggers a rapid reorganization of cortical actin filaments and enables the formation of pseudopodia and the engulfment of particles. In normal conditions, the F-actin coat surrounding nascent phagosomes is disassembled after a few minutes, exposing the membrane and favoring interaction with endocytic organelles. It was observed that F-actin is retained for a longer period around phagosomes containing wild-type Leishmania parasites than around those containing LPG-deficient mutants (Holm et al. 2001). Retention of actin, and several of its effectors (Cdc42, Arp2/3, WASP, α-actinin, myosin II et Nck) (Lodge and Descoteaux 2005), was proposed to act as a shield preventing the fusion of the Leishmania-containing phagosome with endocytic organelles and the proper maturation of this organelle (Lodge and Descoteaux 2008). The LPG-dependent accumulation of actin around phagosomes correlated with the impaired

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IFN-γ Receptor

JAK

F-Actin CapZ α-1

Arp2/3 α-actinin Myosin

STAT WASP Nck

Activation of immune functions

SNAREs IL-10 IL-12 IL-1

α-

SN

AP

Rabs

Rac1

p40PHOX p47PHOX p67PHOX

Cytosol

SHP-1

p22PHOX gp91PHOX Fc Toll-like Receptors Receptors

CD44 ACTIN DYNAMICS

MEMBRANE FUSION

SIGNALING

NADPH oxidase

Phagosome Lumen

OXYDATIVE BURST

Figure 10.2. Alteration of phagosome functions by Leishmania. Proteins implicated in phagosome functions impaired by Leishmania’s LPG have been described to associate with membrane rafts in organelles other than phagosomes. The disruption of phagosomal microdomains by LPG could explain its effect on multiple functions of this organelle (various colored boxes).

recruitment of protein kinase C (PKC)-α (Holm et al. 2001). This kinase was one of the first shown to be strongly inhibited by LPG (Descoteaux et al. 1992). Remarkably, proteins involved in all of the aforementioned functions have been observed in DRMs. These include, for example, the fusogenic molecules Syntaxin 3, Syntaxin 4, and VAMP-2 (Lang 2007), the NADPH oxidase complex (Vilhardt and van Deurs 2004), actin, and some of its effectors (Haglund et al. 2004; Chichili and Rodgers 2009), as well as several signaling molecules, including PKC (Bi et al. 2001; Dienz et al. 2003; Lin et al. 2003). The presence of these proteins in DRMs led us to propose that the disorganization of membrane microdomains by LPG contributes to the overall alteration of phagosomal functions (Dermine et al. 2005). By targeting a single structure, the parasite would alter vital properties enabling phagosome maturation and the ability to generate the microbicidal properties required for parasite killing and the degradation of its proteins for antigen presentation (Fig. 10.2). The molecular mechanisms enabling this profound effect of LPG are still poorly understood. Interestingly, LPG is a glycosylphosphatidylinositol (GPI)-anchored glycolipid shown to be recruited to parasite cell surface DRMs during metacyclogenesis, a process during which poorly infective promastigotes transform into highly infective promastigotes in the insect vector (Denny et al. 2004). This, and the fact that LPG shed from the parasite surface can insert itself into host cell membranes (Tolson et al. 1990), suggest that this molecule might selectively associate with phagosome mem-

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173

brane microdomains where it would interfere with the assembly of functional foci (Dermine et al. 2005). We have shown that LPG alters phagosome functions at the sites of its insertion, a process that requires its repetitive carbohydrate moieties. Thus, the assembly of functional foci on the phagosome membrane appears to be a key aspect of the molecular mechanisms enabling the multiple functions of this organelle.

Mycobacterium tuberculosis A recent study indicated that Mycobacterium tuberculosis, the causative agent of most cases of tuberculosis, might also alter the integrity of phagosome membrane microdomains to inhibit phagosome maturation. It was shown that lipoarabinomannan (LAM), a virulence factor that is also a GPI-anchored glycolipid, is inserted into membrane rafts via its GPI anchor, resulting in reduced phagosomal maturation (Welin et al. 2008). Interestingly, the effect of LAM on phagocytosis does not implicate the disruption of lipid microdomains, contrary to LPG or CbetaG.

Other Microorganisms The handling of a variety of microorganisms by mammalian host cells is believed to involve membrane microdomains at the cell surface. Phagocytosis of Afipia felis, a gramnegative bacteria, is impaired when cholesterol is extracted from the plasma membrane (Schneider et al. 2007). The internalization of Pseudomonas aeruginosa by alveolar macrophages was shown to occur in cholesterol-enriched regions at the cell surface and to involve Lyn kinase (Kannan et al. 2008). This study also showed that membrane microdomains and Lyn were involved in the respiratory burst in these cells. Engulfment of Neisseria gonorrhoeae in human embryonic kidney cells (HEK 293) was shown to involve the cell surface molecule CEACAM6 (carcinoembryonic antigen-related cell adhesion molecule 6), expressed at the surface of epithelial cells, and to require the integrity of membrane microdomains (Schmitter et al. 2007). Whether the mechanisms of entry of these pathogens share similarities with raft-dependent endocytosis (Lajoie and Nabi 2007) remains to be established.

FUTURE PERSPECTIVES Several questions have to be answered in order to understand the role played by membrane microdomains on lysosomes and phagosomes. The composition of these structures, as well as the molecular mechanisms involved in their assembly, is still poorly characterized. On the host–pathogen front, very little is known about the particular features of phagosomes containing microorganisms, and the potential presence of membrane microdomains. Because each microorganism interacts in a specific way with its host cells, for example, through the engagement of selected receptors, it is likely that they reside in phagosomes displaying a wide variety of composition and features. At the molecular level, the mechanisms by which Brucella and Leishmania disorganize and/or inhibit the assembly of membrane microdomains still has to be investigated. A deeper knowledge of the composition of lysosome and phagosome membrane microdomains is likely to provide valuable insights into the biology of these important structures, and a better understanding of the molecular mechanisms used by pathogens to alter their functional properties.

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ABBREVIATIONS Arp2/3 a-SNAP CbetaG CEACAM6 CMA DRM ER GPCR GPI IL-1 IL-10 IL-12 JAK LAM LAMP

actin-related protein 2/3 N-ethylmaleimide-sensitive factor attachment protein alpha cyclic beta-1,2-glucans carcinoembryonic antigen-related cell adhesion molecule 6 chaperone-mediated autophagy detergent-resistant membranes endoplasmic reticulum G-protein-coupled receptor glycosylphosphatidylinositol interleukin-1 interleukin-10 interleukin-12 Janus kinase lipoarabinomannan lysosomal-associated membrane protein

LIMP

lysosomal integral membrane protein LPG lipophosphoglycan MALDI–TOF–MS matrix-assisted laser desorption/ ionization–time-of-flight–mass spectrometer MHC major histocompatibility complex NADPH nicotinamide adenine dinucleotide phosphate PKC protein kinase C SNARE soluble NSF attachment protein STAT signal transducers and activator of transcription V-ATPase vacuolar-type H+-ATPase WASP Wiskott–Aldrich syndrome protein

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CH A P T E R

11

ENDOPLASMIC RETICULUM JUNCTIONS Jesse T. Chao Christopher J.R. Loewen

DEFINITION Membrane contact sites (MCSs) have been identified in organisms ranging from bacteria to humans suggesting they are ancient structures that play important roles in all cells (Levine 2004). In this chapter, we will focus on a subset of intracellular membrane contacts, those that are formed between the endoplasmic reticulum (ER) and other organelles. We will call these ER junctions (ERJs) (Levine 2004; Levine and Loewen 2006). ERJs function to facilitate nonvesicular exchange of lipids and calcium between organelles. An ERJ corresponds to a region in the cell where a subdomain of the ER is in close contact with another membrane-bound organelle, and in which the contact is mediated by one or more protein bridging complexes that span the gap between membranes. In most cases, the apposing membranes at ERJs do not fuse, although there may be exceptions under special circumstances (Gagnon et al. 2002; Becker et al. 2005). ERJs have been visualized by electron microscopy (EM) between ER and Golgi (Golgi-ERJ), mitochondria (MitoERJ), and plasma membrane (PM-ERJ) among others (Achleitner et al. 1999; Ladinsky et al. 1999; Pichler et al. 2001). The gap between membranes is tightly controlled and is generally ∼10 nm, but can range from ∼5 to 25 nm (Csordas et al. 2006), whereas the length of an individual junction can vary greatly. Tremendously, in budding yeast, the ER contacts the PM over 1100 times per cell, which is nearly 15 times as many as between ER and mitochondria, the second most extensive ERJ in this organism (Pichler et al. 2001). In muscle cells, the sarcoplasmic reticulum interacts extensively with PM, creating a structure known as the triad junction (Endo 2009). ERJs between ER and mitochondria (Vance 1990; Achleitner et al. 1999) and ER and PM (Pichler et al. 2001; Koziel et al. 2009) have been isolated biochemically, indicating that the structures observed by EM are not artifactual but are stable, organized structures. A list of ERJ components can be found in Table 11.1.

HISTORICAL PERSPECTIVE The triad junction of muscle cells was likely the first example of an ERJ ever described (Franzini-Armstrong 1970). Its highly organized and repetitive nature enabled it to be readily characterized by EM (Bers 2008; Endo 2009). However, for other ERJs, their

Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Transfers ceramide Binds oxysterol 25-hydroxycholesterol May transfer phosphatidylinositol and phosphatidylcholine May catalyze the transfer of phosphatidylinositol and phosphatidylcholine between membranes May transfer glycolipids Transfer PI IP3Rs Ion channels Calcium channel Detects the calcium concentration in the ER lumen; activates store-operated calcium channel activity

Mammalian Mammalian Drosophila Mammalian Mammalian Yeast Mammalian Mammalian Mammalian Mammalian

Osh1 Osh2 Osh3 Osh4 CERT OSBP RdgB

Nir2 (PTPM1)

FAPP2 Sec14 Inositol triphosphate receptor (IP3R) Transient receptor potential (TRP receptors) Orai channel Stim1

Yeast

Enoyl-CoA reductase Contacts substrates such as the insulin receptor, PKCδ, and Src either while in the PM or shortly after endocytosis May bind and transport sterols

Yeast Mammalian

Localizes FFAT motif proteins; may form PM-ERJ Forms ERMES complex to bridge Mito-ERJ

Yeast Yeast

Scs2 Mdm12 Mdm10 Mmm1 Mdm34 Tsc13 PTP1-B

Forms NVJ with Vac8

Molecular Function

Forms NVJ with Nvj1

Yeast

Species

Vac8

Nvj1

Protein

Proteins with Potential Functions at ER Junctions

*For most proteins, their localizations to ERJs are strongly inferred.

Signaling

Channels

Lipid transfer proteins

Enzymes

Structural components

Functions

TABLE 11.1.

Integral

Integral

Soluble

Integral

Integral Soluble Integral

Soluble

Integral

Integral/Soluble

PM ER

ER PM

TGN

TGN

ER and Golgi Cell cortex in bud Cytoplasm Cytoplasm TGN TGN PM

ER ER

Nuclear membrane Vacuolar membrane ER Punctate

Localization

PM-ERJ

PM-ERJ

Golgi-ERJ

Golgi-ERJ

Golgi-ERJ Golgi-ERJ Golgi-ERJ PM-ERJ

NVJ PM-ERJ

NVJ PM-ERJ

PM-ERJ (ER) Mito-ERJ (ER)

Nvj

ERJ* (Side)

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organization was not as obvious and thus they were overlooked for decades. Early on, it is likely that electron microscopists were unsure if the association of ER with other organelles in their micrographs was random or specific. The Golgi-ERJ was first seen by Novikoff, which he called GERL for Golgi-ER-lysosome (Novikoff 1976). GERL was later renamed the trans-Golgi network (TGN) (Griffiths and Simons 1986), and 3D electron tomography studies have confirmed it makes extensive contacts with smooth ER (Marsh et al. 2001; Mogelsvang et al. 2004). Junctions between ER and mitochondria were overlooked for similar reasons until more detailed ultrastructural analyses were performed (Franke and Kartenbeck 1971; Morre et al. 1971). Even physical linkages likely representing protein bridges have now been observed at Mito-ERJs (Csordas et al. 2006). Despite these detailed structural descriptions, an understanding of the function and molecular components of ERJs remained extremely limited. However, the biochemical purification of mitochondria-associated ER membrane, called MAM (Rusinol et al. 1994), and the finding that MAM was enriched in lipid-synthesizing enzymes, suggested a role for the Mito-ERJ in lipid metabolism. This was supported by earlier studies that indicated that phospholipid transfer between ER and mitochondria was a requirement for de novo phospholipid biosynthesis and was dependent on direct contact between these organelles (Voelker 1989; Vance 1990). Calcium was also thought to traffic at ERJs, but this was not directly demonstrated until Rizzuto et al. in a groundbreaking work imaged its flow in living cells between ER and mitochondria (Rizzuto et al. 1998). Genetic screens in yeast have led the way in the identification of factors that regulate ERJ structure and function (Trotter et al. 1998; Pan et al. 2000; Wu et al. 2000; Schumacher et al. 2002; Kornmann et al. 2009). However, the proteins responsible for forming and maintaining ERJs are largely unknown, except in one case. It is for the nucleus–vacuole junction (NVJ), a vacuolar ERJ in yeast, that the molecular bridge has been clearly defined (Pan et al. 2000; Roberts et al. 2003). The intent of this chapter is to introduce key concepts related to ERJ structure, assembly, and function, as well as outline many of their known molecular constituents and roles in cell physiology. This chapter does not cover the anatomy of all potential ERJs, but rather focuses on those that have at least a few defined molecular constituents. We will begin with the NVJ in yeast as it is an excellent starting point to understand the basic building blocks of an intracellular MCS. We will then cover the PM-ERJ, Golgi-ERJ, and Mito-ERJ before finishing with unresolved issues in the field.

MOLECULAR COMPOSITION OF JUNCTIONS NVJ In budding yeast, a portion of the perinuclear ER (nuclear envelope) makes a stable contact with the vacuolar membrane, the yeast equivalent of the lysosome. Because of the size and stability of the NVJ, it can be easily monitored by both light and EM, and since it is not an essential structure, it can be manipulated by reverse genetic techniques. The NVJ is assembled by the direct interaction of an integral membrane protein of the ER, called Nvj1, with a peripheral palmitoylated protein of the vacuole, called Vac8 (Fig. 11.1). Through studies on vacuole fusion, Nvj1 was identified as an interactor of Vac8 initially by yeast two-hybrid and later confirmed biochemically (Pan et al. 2000). Nvj1 was then found to localize to a subregion of the outer nuclear envelope apposed to the vacuole, and when Vac8 was deleted, Nvj1 relocalized uniformly to the perinuclear ER, suggesting that its interaction with Vac8 resrtricted Nvj1 to a contact site between nucleus and vacuole.

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(A)

Vacuole-ERJ

Mito-ERJ

PM-ERJ PM

Vacuolar membrane

Mdm34

Outer mitochondrial membrane

Mdm10

Polarisome

Vac8

?

Mdm12

Scs2

Mmm1

Nvj1

ER

Cortical ER

Perinuclear ER

(B)

FFAT

ORP

OSBP

PH

ORD

OSH1,2,3

PH

ORD

OSH4,5,6,7 ORP5,8

CRAL/TRIO SEC14, SFH1

ORD ORD

PH

TM

CRAL/TRIO FFAT

START

CERT

PITP

NIR2

GLTP

FAPP2

PH

START FFAT

PITP

PH

DDHD

GLTP

LNS2

100 aa’s

Vac8, on the other hand, localizes fairly uniformly to the vacuole membrane. The truly exciting result came when cells lacking Nvj1 were observed by EM and found to have no NVJ, indicating that the direct binding of these proteins resulted in the formation of a structural bridge between the two membrane systems. Until this point, very little was known about the NVJ other than that it excluded nuclear pore complexes (Severs et al. 1976), suggesting it was a unique membrane domain of the ER. Goldfarb and coworkers also found through overexpression studies that the size of the NVJ was governed simply by the amount of Nvj1 present to “velcro” the two membranes together (Pan et al. 2000). They later discovered stress response elements in Nvj1’s promoter that regulate its expression and also a function for the NVJ in autophagy (Roberts et al. 2003). Thus, the interaction of Vac8 with Nvj1 gives us the basic understanding that ERJs are formed through protein structural bridges between the two membrane systems, and this is a paradigm for all ERJs. Two other proteins in yeast are known to localize to the NVJ, and both have roles in lipid metabolism. Tsc13 is an integral protein of the ER that is an enoyl-coenzyme A (CoA) reductase enzyme, which is critical for the formation of very long-chain fatty acids

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Figure 11.1. (A) Structural components of ERJs. In only a few cases have the bridging complexes required for formation of ERJs been described. For the three examples shown here, the direct binding partners on the participating membranes have been identified, except for Scs2 (see below). These complexes have also been clearly demonstrated to be required for ERJ formation. The N-terminus of Vac8 is myristoylated to anchor it to vacuolar membrane and its C-terminus, which interacts with the integral ER protein NVJ1, is palmitoylated, likely to regulate this interaction (Pan et al. 2000). The ERMES complex at the mito-ERJ is formed by a type I transmembrane protein of the ER, Mmm1, interacting with Mdm10 on the outer mitochondrial membrane and a soluble protein Mdm12. Mdm34 of the outer mitochondrial membrane has not been shown to directly bind to ERMES but is required for its correct localization. The type II ER transmembrane protein, Scs2, is localized to sites of PM-ERJ formation by the polarisome (Loewen et al. 2007). Mutation of either Scs2 or the polarisome disrupts PM-ERJ formation, but a direct interaction has yet to be demonstrated. (B) LTP families. Shown are LTPs for which there is evidence for their having functions at ERJs. Not all members of each family and not all families are depicted (for a more comprehensive list of LTPs, see D’Angelo et al. 2008). ORPs contain a core OSBP-related domain (ORD) LTP domain that binds sterols (Im et al. 2005); CRAL-TRIO domain in Sec14 and Sfh1 binds PI, PC, and phosphatidylethanolamine (PE) (Schaaf et al. 2008); the START in CERT binds ceramide (Kudo et al. 2008); phosphatidylinositol transfer protein (PITP) domains bind PI (Tilley et al. 2004) and PC (Yoder et al. 2001); and glycolipid transfer protein (GLTP) domains bind glycosphingolipids (Malinina et al. 2006). LTP domain organizations are shown to scale, and LTP family names are highlighted in red while gene names are italicized. Noteworthy are the similar domain organizations of ORP, START, and GLTP family proteins. Domain abbreviations: CRAL, cellular retinaldehyde domain; TRIO, a triple functional domain encoded by the Trio gene in humans; STAR, steroidogenic acute response protein; PH, pleckstrin homology; FFAT, short conserved sequence motif “EFFDAxE” that binds to VAP/Scs2 on the ER; TM, transmembrane; DDHD, metal-binding domain with conserved sequence “DDHD”; LNS2, found in Lipin/ Ned1/Smp2 proteins.

(Kohlwein et al. 2001), and localizes to the NVJ by binding to Nvj1 (Kvam et al. 2005). Osh1 is one of seven yeast homologs of a highly conserved family of sterol transporting lipid transfer proteins (LTPs) related to oxysterol-binding protein (OSBP) (Fig. 11.1) that localizes to the NVJ also by binding to Nvj1 (Levine and Munro 2001; Kvam and Goldfarb 2004). The molecular function of Osh1 remains to be defined; however, it clearly plays a functional role at the NVJ as Osh mutants also have defects in autophagy, but no NVJ structural defects are observed. This is also true for Tsc13 (Kvam et al. 2005). A Paradigm for ERJ Formation/Localization From studies on the NVJ, a paradigm for ERJ formation and function begins to emerge in which (1) a structural bridge is formed by stable binding of proteins from opposing membranes, (2) the amount of the limiting binding partner governs the size of the junction, and (3) the protein bridge itself recruits accessory proteins/enzymes to perform functions specific for that ERJ. In the case of Tsc13, it is sequestered within a subdomain of the ER at the NVJ, similar to Nvj1 itself, while for Osh1, it is a soluble protein that is recruited peripherally to the NVJ. And it also appears that initial formation of the structural bridge simply depends on random association of the two membranes, binding of the cognate proteins (e.g., Vac8 and Nvj1) and “zippering” to create the MCS. An emerging concept for the general targeting of proteins to preexisting ERJs is that any protein with affinity for two membranes that is large enough to bridge the gap at the ERJ should be enriched there. Hence, both soluble and integral membrane proteins with multiple membrane-targeting determinants are good candidates for acting at ERJs (Levine 2004). This is a key concept that will reemerge as we discuss the proteins that likely function at ERJs.

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PM-ERJ Triad Junction A key feature of this highly specialized PM-ERJ is that it enables direct contact between the dihydropyridine receptors in the transverse tubules of the PM and the ryanodine receptors in the sarcoplasmic reticulum, which results in channel opening and calcium release in response to depolarization of the PM. However, this interaction is not thought to comprise the structural bridge. Instead, another class of ER proteins called junctophilins likely perform this role, although their binding partners on the PM have yet to be identified (Takeshima et al. 2000) and awaits further study. Store-Operated Calcium Entry (SOCE) A similar mechanism to excitation contraction coupling at the triad junction in muscle underlies SOCE in virtually all cell types. In SOCE, inositol triphosphate receptors (IP3Rs) of the ER contact transient receptor potential (TRP) receptors on the PM indirectly via a cytosolic protein called homer (Yuan et al. 2003), allowing replenishment of ER calcium stores. Calcium traffic has now been visualized between PM and ER (Fahrner et al. 2009). It is critical that this occurs at PM-ERJs so that the incoming calcium does not flood the cytoplasm. In lymphocytes, an incredible mechanism is emerging on coupling the calcium status of the ER to opening of Orai calcium channels in the PM in SOCE (Fahrner et al. 2009). An ER-embedded calcium sensor protein, Stim1, detects the calcium concentration in the ER lumen using its EF-hand domain (helix-loop-helix structural domain found in a large family of calcium-binding proteins; this structure allows calcium to be embedded in its core via interaction with negatively charged amino acids). When calcium drops below a certain threshold, Stim1 oligomerizes, which allows its cytoplasmic domain to directly bind to the Orai channel of the PM. The unexpected part is that Stim1 does not traffic to the PM through the secretory pathway; instead, it remains in the ER but relocalizes to ER that is in close proximity to PM (PM-ERJ) where it promotes tetramerization and activation of the Orai channel (Penna et al. 2008). In fact, ER calcium depletion increases the number of PM-ERJs per cell, but not their size, whereas overexpression of Stim1 increases the size, but not the number (Wu et al. 2006). A recent study found that phosphoinositides (PIPs) on the PM might help localize Stim1 to ER in contact with PM by binding to Stim’s C-terminus (Ercan et al. 2009), and this sorting signal appears to be conserved in all eukaryotes. So, based on our model ERJ, the NVJ, these data suggest that the Stim1–Orai interaction does not comprise the core structural bridge, but that once formed at a PM-ERJ by the aid of a general targeting determinant for the PM in Stim1, the Stim1–Orai interaction modulates the size and or stability of that ERJ. Cortical ER (cER) in Yeast In budding yeast, incredibly, the ER contacts the PM over 1100 times per cell as quantified by EM of serial sections (Pichler et al. 2001) and is often referred to as cER because of its obvious location at the periphery of the cell. cER is formed at the tip of newly forming buds during each round of the cell cycle and is therefore easily studied in this genetically tractable organism (Du et al. 2004). A seminal work that characterized the structure and dynamics of cER in living yeast and identified the first mutants in the pathway clearly showed that cER in yeast is remarkably similar to metazoan ER (Prinz et al. 2000). cER consists of ER tubules connected by three-way junctions that are capable of dynamically rearranging on the actin cytoskeleton. However, neither actin nor microtubules were required to maintain cER once formed at the periphery of the cell, indicating that direct physical contacts with the PM likely hold it in place. Mutations in genes that are involved in ER to Golgi traffic, such as coat protein complex I (COPI), and mutations in the signal

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recognition particles (SRP) receptor altered general ER morphology, but the specific roles for these components in PM-ERJs are unclear. Additional visual screens have identified mutants with delayed inheritance of ER to the bud (Prinz et al. 2000; Du et al. 2004, 2006; Estrada de Martin et al. 2005; De Craene et al. 2006), but cER was not quantified by EM in these studies. As a result, it was difficult to determine their effects on PM-ERJ formation. A new class of proteins responsible for tubulating the ER, called reticulons, clearly play a role in structuring cER (Voeltz et al. 2006; Hu et al. 2008); however, it remains attached to the PM in these mutants, suggesting that reticulons are not part of the structural bridge between ER and PM. Another conserved integral ER protein, called Scs2 or vacuolar membrane-associated protein (VAMP)-associated protein (VAP) in humans, is a good candidate for being part of a structural bridge (Loewen et al. 2007) (Fig. 11.1). A role for Scs2 in cER was discovered because of its unusual localization; it is segregated within the ER to sites of polarized growth, including the bud tip, the site of cER formation. Loss of SCS2 resulted in a substantial decrease in PM-ERJ formation without causing defects in ER tubules or nuclear envelope structure. Furthermore, when Scs2 was released from the ER by deleting its C-terminal transmembrane domain, it retained its polarized localization, suggesting it has targeting determinants also on the PM (Loewen et al. 2007). Thus, binding of Scs2 directly to the PM could provide a simple mechanism that facilitates PM-ERJ formation in yeast. This study also found a link between PM-ERJ formation and the nuclear cycle in that defective PM-ERJ formation activated a cell cycle checkpoint, thus uncovering an important function for ERJs in cell cycle regulation. Sterol Traffic and the OSBP-Related Protein (ORPs) A role for PM-ERJs in sterol traffic has been implied for almost 30 years (DeGrella and Simoni 1982), and studies on cultured cells treated with brefeldin-A (Urbani and Simoni 1990) and on Sec temperature-sensitive mutants in yeast (Baumann et al. 2005; Schnabl et al. 2005) demonstrated that the bulk of sterol moves from ER to PM independently of the secretory pathway, implying traffic at ERJs. A highly conserved family of LTPs has now been found to play a role in this transport in yeast (Im et al. 2005). This family includes 16 ORPs in humans and seven OSBP homolog (OSH) proteins in yeast (Fig. 11.1) (Olkkonen and Levine 2004). The feature shared by all these proteins is a highly conserved domain at the C-terminus, called ORD for OSBP-related domain, suspected to be a lipid-binding/transfer domain. Yeast Osh4 was found to be capable of transporting sterol between donor and acceptor liposomes in vitro, suggesting that it may function as an LTP (Im et al. 2005). The structure of yeast Osh4 was solved in complex with cholesterol and ergosterol (Im et al. 2005) and tellingly the lipid was buried in a deep hydrophobic pocket, which was accessible to the cytoplasm via a channel that even included a lid. Further work demonstrated that Osh proteins, especially Osh4, are responsible for a significant proportion of sterol transport in vivo in yeast (Raychaudhuri et al. 2006), consistent with them acting as LTPs in vivo. Is there evidence these proteins function at PM-ERJs? Unfortunately, the bridging complexes between ER and PM remain ill-defined, so it has not yet been possible to break apart the structure and assay for Osh-dependent lipid traffic. Even though the Δscs2 yeast mutant has close to a 70% reduction in PM-ERJs, this mutant does not have reduced rates of sterol transfer between PM and ER (Prinz, unpublished data), implying that this transfer must be very robust if cER and PM-ERJs are indeed important. But a recent study has now shown that Osh proteins in yeast very likely localize to PM-ERJs and that simultaneous targeting to both membranes is a requirement for lipid transfer (Schulz et al. 2009).

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ERJ Targeting: The Two Phenylalanines in an Acidic Tract (FFAT) Motif A unique property of the ORP family of proteins is that they contain, in addition to a sterol transfer domain, multiple membrane-targeting domains that may allow them to localize to ERJs (Olkkonen and Levine 2004). What are known as the “long” forms contain a pleckstrin homology (PH) domain near their N-terminus, while the “short” ORPs lack this domain. The PH domain allows ORPs to associate with different membranes based on their selectivity for binding different PIPs (Olkkonen and Levine 2004). An important advance was the discovery that OSBP and the long ORPs, such as Osh1/2/3, contain a highly conserved short motif for targeting the ER, called the FFAT motif (Loewen et al. 2003). The FFAT motif contains a core element, “EFFDAxE,” that is flanked by acidic amino acids, and it enables direct binding to VAP/Scs2. At first, this discovery appeared paradoxical as how could these proteins be localized to the ER and the PM simultaneously? One explanation was that they could only have affinity for one membrane at a time. But the data showed that the PH domain and FFAT motif synergized to specify Osh protein subcelluar localization; that is, there was integration of the targeting signals (Loewen et al. 2003). This strongly suggested that the long ORPs were interacting with both membranes simultaneously, and hence were localizing to ERJs. For Osh1, the localization is most clear, as the NVJ is an easily identifiable ERJ. For Osh2 and -3, there is now additional evidence that their suspected localization to PM-ERJs (Loewen et al. 2003) is indeed true (Schulz et al. 2009). But what about the short ORPs that lack PH domains and FFAT motifs? Even these have now been found to target ER and PM simultaneously, and this is required for their sterol transfer activity both in vitro and in vivo (Schulz et al. 2009). Finally, there are large ORPs in metazoans (ORP5, -8) that contain, in addition to PH and ORDs, an additional C-terminal transmembrane domain that localizes them to the ER (Loewen, unpublished data) and suspiciously lack FFAT motifs. If these proteins are truly LTPs, that they are firmly anchored in the ER indicates they must function at ERJs. Other Activities It appears that PM-ERJs also play a role in cell-surface receptor dephosphorylation, as the ER-embedded tyrosine phosphatase, PTP-1B, contacts substrates such as the insulin receptor, PKCδ, and Src either while in the PM or shortly after endocytosis (Anderie et al. 2007; Yudushkin et al. 2007). ER-bound PTP-1B also plays a role at focal adhesions where it is important for their assembly or maintenance, but its exact substrates are unknown (Hernandez et al. 2006). The association of ER with septins on the PM in yeast is required to create a diffusion barrier for ER proteins between bud and mother (Luedeke et al. 2005), and PM-ERJs in turn seem to be important also for septin assembly (Loewen et al. 2007), stressing the functional interplay between these structures. Also, a Drosophila protein called Rdgb likely functions at PM-ERJs in photoreceptors where it is required for traffic of phosphoinositol (PI) between these organelles (Milligan et al. 1997) and is discussed in more detail in the next section.

Golgi-ERJ The TGN makes extensive contacts with smooth ER, which have been studied in detail by 3D electron tomography (Marsh et al. 2001; Mogelsvang et al. 2004). Astoundingly, in this study, the ER could be seen moving in between the cisternae of the Golgi stack while remaining continuous with the bulk ER. However, the structural bridging complexes have yet to be discovered. Nevertheless, the functions for Golgi-ERJs have been recently uncovered—in lipid traffic—with the discovery of multiple classes of LTPs localizing to the TGN and functioning in sphingolipid synthesis.

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Ceramide Transfer Protein (CERT) and Sphingomyelin Synthesis Ceramide, the precursor of sphingomyelin, is synthesized in the ER and must be transported to the lumenal leaflet of the TGN, where the sphingomyelin synthase is located, to be converted to sphingomyelin. Approximately half of the ceramide is trafficked in vesicles by the secretory pathway, while the other half is mediated by an LTP called CERT (ceramide transporter) and is nonvesicular in nature (Hanada et al. 2003, 2009). Similar to the ORP family, CERT has an N-terminal PH domain that targets PIPs on the Golgi, a FFAT motif for targeting the ER, and a C-terminal steroidogenic acute regulatory (STAR)related domain (START), which binds ceramide and buries it in a deep hydrophobic pocket in the protein structure (Kudo et al. 2008) (Fig. 11.1). Thus, again we find an LTP with domains that target two organelles, one of which is the ER. Does CERT therefore function at Golgi-ERJs? The answer again is very likely yes. Although CERT localizes to the TGN and is not seen localizing to ER (Kawano et al. 2006), mutation of either the PH domain or FFAT motif in CERT abolishes its ceramide transfer activity in vivo. Given the extensive associations between the TGN and ER, CERT likely functions at Golgi-ERJs. Highresolution cryo-EM or tomography should show us if this is true. OSBP OSBP was discovered as the major target of the oxysterol 25-hydroxycholesterol in cells, which is a potent regulator of transcription factor SCAP-SREBP (SCAP, sterol regulatory element-binding protein [SREBP] cleavage activating protein) and sterol metabolism (Ridgway et al. 1992). However, it was not until the discovery of CERT that OSBP’s function in regulating sphingomyelin synthesis was uncovered (Perry and Ridgway 2006). Like CERT, OSBP too very likely localizes to Golgi-ERJs because it has a PH domain that localizes to the TGN and a FFAT motif for ER targeting. Although OSBP generally localizes to the TGN and is not seen on the ER, a PH-domain mutant, W174A, localizes to the ER (Wyles et al. 2002). This suggests that the wild-type protein is also in contact with the ER, but the combination of Golgi and ER targeting restricts its localization to Golgi-ERJs. In further support, both the PH domain and FFAT motif are required for OSBP to stimulate CERT in the presence of oxysterols (Perry and Ridgway 2006). OSBP has now been shown to act as cholesterol LTP in vitro (Ngo and Ridgway 2009). Therefore, OSBP appears to be the molecular link between sterol metabolism and sphingolipid production, acting both as sensor (of oxysterols) and LTP (for cholesterol). ORP9 may also have a similar role, although it does not bind oxysterols and therefore likely participates solely in trafficking cholesterol between ER and Golgi (Ngo and Ridgway 2009). Indeed, knockdown of ORP9 did not prevent stimulation of sphingolipid synthesis by oxysterols, as is the case for OSBP (Ngo and Ridgway 2009). But ORP9 knockdown did affect Golgi structure and protein secretion, implying that it may also be involved in the formation or maintenance of Golgi-ERJs. Phosphatidylinositol-Four-Phosphate Adapter Proteins (FAPPs) and Glycosphingolipid Synthesis LTPs may also be involved in glycosphingolipid metabolism at the TGN. Two recent papers showed conclusively that the protein FAPP2, which contains a glycolipid transfer protein (GLTP) domain (Fig. 11.1) and localizes to the TGN is required for the production of complex glycosphingolipids (D’Angelo et al. 2007; Halter et al. 2007). However, these two groups came to slightly different conclusions about the route taken by glycosphingolipids, but they agreed that it is mediated by nonvesicular traffic. D’Angelo et al. demonstrated that FAPP2 can transfer glucosylceramide (GlcCer) in vitro, and this is dependent on the GLTP domain, indicating that it is a bona fide LTP. But these authors suggested that FAPP2 acts on the cytoplasmic face of cis-Golgi (where the GlcCer synthase enzyme

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was wrongly thought to be) and transfers GlcCer to the TGN, where the GlcCer is flipped to the luminal leaflet to be made into complex glycosphingolipids. Evidently, their model does not involve an ER step. On the other hand, Halter et al. showed that GlcCer synthase is enriched in the TGN and that GlcCer does not flip from the cytosolic to lumenal leaflet at the TGN, which is a problem for the D’Angelo model. Halter et al. then found, surprisingly, that the GlcCer made at the TGN is trafficked by FAPP2 back to the ER to be flipped to the luminal leaflet before being trafficked in vesicles to cis-Golgi for conversion into complex glycosphingolipids. It thus appears that FAPP2 does indeed function at GolgiERJs, but it traffics GlcCer from the TGN back to the ER. Interestingly, in flies, GlcCer synthase is located in the ER, and flies do not have a FAPP2 homolog, suggesting that transporting GlcCer back to the ER truly is FAPP2’s role (Levine 2007). Curiously, FAPP2 does not have a FFAT motif for ER targeting, indicating it must have an as yet unidentified lipid/protein receptor. Other Potential LTPs Yet another class of LTPs, called Nir (Lev et al. 1999), has roles in structuring the Golgi and regulating secretion, likely through affecting Golgi diacylglycerol (DAG) levels (Litvak et al. 2005; Peretti et al. 2008). Nir proteins are highly conserved (called RdgB for retinal degeneration B in Drosophila) and contain a phosphatidylinositol transfer protein (PITP) lipid transfer domain present in mammalian PITP (D’Angelo et al. 2008) (Fig. 11.1). This domain in RdgB can transfer PI in vitro, and in flies, RdgB transfers PI from ER to PM to replenish rapidly depleting PI(4,5)P2 levels in photoreceptors due to activation of phospholipase C during phototransduction (Vihtelic et al. 1993; Milligan et al. 1997). Nir proteins also have a FFAT motif, and in the case of RdgB, it localizes to ER directly apposed to the photoreceptor PM (PM-ERJ). Nir2 and Nir3 localize to the Golgi, and it awaits confirmation that this is the Golgi-ERJ. However, their interaction with VAP in the ER is required for function (Peretti et al. 2008), suggesting that Nir proteins transfer lipids from ER to TGN or vice versa. Reconstitution of their LTP activity in vitro also still awaits. Yeast has an analogous family of PI transfer proteins, called Sfh for Sec14 homolog, that contain the Sec14 PI/phosphatidylcholine (PC) lipid transfer domain (Peretti et al. 2008) (Fig. 11.1). Sec14 has been shown to transfer these lipids in vitro and is essential for Golgi structure and secretion in vivo (Bankaitis et al. 1990). And once again, the structure revealed a hydrophobic pocket for burying the lipid and a lid to seal it shut, although this structure incorporated two detergent molecules in place of lipid (Sha et al. 1998). However, recent structural analysis of Sec14 bound to PI and PC combined with structure-based mutagenesis suggests that Sec14 function in vivo is more complicated than it simply acting as an LTP (Schaaf et al. 2008). The new data suggest that Sec14 presents lipids to PI kinases in cis and is therefore a key regulator of PI metabolism instead. It awaits to be determined if Sec14-mediated stimulation of PI kinase activity can be measured in vitro.

Mito-ERJ Mito-ERJs, the ER component of which is called MAM, have been studied extensively as they are readily observed by EM (Franke and Kartenbeck 1971; Morre et al. 1971; Csordas et al. 2006) and have been isolated biochemically (Voelker 1989; Vance 1990; Achleitner et al. 1999). MAMs have been proposed to have multiple functions. For example, calcium traffic between ER and mitochondria occurs at Mito-ERJs (Rizzuto et al. 1998), which uses a similar arrangement as calcium coupling at PM-ERJs and is extensively reviewed

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elsewhere (Hayashi et al. 2009). Also, MAMs are enriched in lipid-synthesizing enzymes and are likely the site of lipid translocation between ER and mitochondria (Vance 2003). Mitochondria are capable of synthesizing some, but not all, of the phospholipids they require. Since mitochondria are off the secretory pathway, they must exchange lipids with the ER by nonvesicular traffic, which has been reconstituted in vitro and depends on protein components on the respective membranes (Achleitner et al. 1999). Furthermore, genetic screens in yeast have identified an E3 ubiquitin ligase, Met30 that is required for phospholipid traffic between ER and mitochondria, suggesting that ubiquitination plays a role at Mito-ERJs (Schumacher et al. 2002; Choi et al. 2006). However, a mechanism remained elusive for many years until the discovery of a first bridging complex called ER–mitochondria encounter structure (ERMES) (see below). And suspiciously, the ERMES subunit Mdm34 is a target of another ubiquitin ligase, Mdm30 (Ota et al. 2008). The First Structural Bridge: ERMES To identify a structural bridge for Mito-ERJs, Kornmann et al. designed a synthetic bridging molecule called “chimera” that contained a mitochondrial and an ER membranetargeting signal separated by a linker that included green fluorescent protein (GFP). They then took advantage of the power of yeast genetics and performed a mutagenic screen looking for mutants in which their synthetic bridge became essential. After screening 100,000 colonies, they found two mutants harboring mutations in the same gene, MDM12. Mdm12 forms a complex with three other proteins, Mmm1, Mdm10, and Mdm34, that localize to punctae in the cell and are required for respiratory growth and mitochondrial morphology (Berger et al. 1997; Boldogh et al. 2003; Youngman et al. 2004), and which they have now renamed “ERMES” (Fig. 11.1). Although these genes are not directly conserved to mammals, Mmm1 and Mdm12 contain synaptoptagmin-like mitochondrial proteins (SMP) domains, which are highly conserved and may function similarly in vertebrates. Chimera expression in the deletion mutants rescued the respiratory defects and largely the mitochondrial morphology defects. Interestingly, when the mitochondrial subunits Mdm10 and -34 were deleted, Mdm12 and Mmm1 relocalized from punctate to general ER, showing that their interaction with mitochondria restricted their localization within the ER, implying that these subunits interact across the Mito-ERJ and comprise a structural bridging complex. Fascinatingly, Mito-ERJs were visualized in a living cell for the first time. Unexpectedly, there are far fewer “dots” than expected based on earlier EM studies on mitochondrial ER contact sites, by this study from 2 to 10 per cell. It is unclear if the ERMES dots represent all Mito-ERJs in the cell or if each dot is of similar composition. Lastly, Kornmann et al. found that disruption of ERMES led to a decrease in the kinetics of conversion of phosphatidylserine (PS) into PC via the mitochondrial PS decarboxylation pathway, which depends on nonvesicular traffic (Choi et al. 2006), but phospholipid levels in mitochondria were normal, except for a modest decrease in cardiolipin. This is not surprising given the nature of their original chimera screen as it would preclude identification of the lipid transporters involved in the traffic, and since they did not measure for the loss of ER–mitochondria association, it is not clear if ERMES is solely responsible for Mito-ERJ formation. Their lipid transfer data argue it is not, or that the transporters can operate without Mito-ERJs or with severely disabled junctions in the ERMES mutants. Even though the lipid transporters that act at Mito-ERJs are still unknown, knowing the identity of one structural bridge not only marks a major advance in the field but will also enable further definition of the roles for Mito-ERJs. The ERMES bridge shares some similar characteristics with the Nvj1–Vac8 bridge. Both involve heterotypic interactions between two membrane-bound proteins in each membrane, and these interactions restrict their localizations to the ERJ. For ERMES, a

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minor difference is the presence of a soluble protein, Mdm12, that links the membraneembedded proteins, which may reflect a need for greater regulation at the Mito-ERJ than the vacuole-ERJ. Indeed, that the ERMES bridge is regulated posttranslationally by Mdm30 suggests that, unlike the Vac8–Nvj1 bridge, which is regulated at the transcriptional level, ERMES regulation is even more dynamic. Other Components Autocrine motility factor receptor (AMFR)/GP78 (also known as GP78) is an integral ubiquitin ligase of the ER that participates in ER-associated protein degradation (ERAD). Studies using an antibody to AMFR/GP78, 3F3A, have identified a subdomain of smooth ER that is associated with mitochondria (Wang et al. 2000; Goetz et al. 2007; Fairbank et al. 2009). Specifically, the antibody 3F3A labels smooth ER that is in contact with mitochondria by EM and by confocal microscopy and shows extensive colocalization with mitochondrial HSP70 (Wang et al. 2000). Other studies using 3F3A labeling and structural EM showed that free cytosolic calcium regulates the extent of association between ER and mitochondria, and dissociation with increased calcium (∼100 nM) is rapid and reversible, indicating that it is a functional coupling that is physiologically relevant. Overexpression of AMFR/GP78 saturates colocalization with mitochondria (Goetz et al. 2007), implying that this interaction is specific. However, 3F3A-labeled ER does not colocalize with reticulons (Goetz et al. 2007), suggesting that the 3F3A ER domain is either not tubular in nature, or that reticulons are specifically excluded from these regions. This unique localization of AMFR/GP78 also suggests an intriguing role for Mito-ERJs in retrotranslocation and the ERAD pathway. Nevertheless, several key questions still need to be answered. For instance, does knocking down AMFR/GP78 decreases Mito-ER contacts? What is the receptor for AMFR/GP78 at Mito-ERJs? Another protein, PACS-2, localizes both to ER and mitochondria and targets Bid to mitochondria in the presence of apoptotic stimuli to promote apoptosis (Simmen et al. 2005). Interestingly, knockdown of PACS-2 disrupts mitochondrial morphology and association with ER through caspase activation of BAP31, a proapoptotic factor that activates a dynamin-related protein Drp1/Dlp1 that drives mitochondrial fission. However, it is not known if PACS-2 localizes to Mito-ERJs or what its receptors are, but it is exciting to speculate that another role for Mito-ERJs might be in regulating cell death. In fact, PACS-2 is mutated in over 40% of colorectal cancers (Simmen et al. 2005), and it is required for killing of tumor cells (Aslan et al. 2009). In another study, a synthetic linker protein was engineered that contained both an ER and mitochondrial targeting signal separated by an intervening red fluorescent protein (RFP), and remarkably it was found to alter Mito-ER coupling both structurally and functionally (Csordas et al. 2006). Expression of this ∼5-nm linker decreased the average mito-ER gap size from 24 to 6 nm and increased the length of interface over fourfold. This led to increased rates of calcium transfer and calcium overloading of mitochondria, similar to what the authors observed under apoptosis-inducing conditions, suggesting that the physical dimensions of the junction regulate transport activity.

UNRESOLVED ISSUES It has yet to be directly proven whether LTPs transfer lipids at ERJs in vivo and if they do so by shuttling a short diffusible distance or by bridging the small intermembrane gap at ERJs. Perhaps by modulating ERJ gap width in vivo and measuring rates of lipid transfer, we will be able to define the appropriate mechanism. Alternatively, one could artifi-

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cially tether an LTP to both membranes and measure the rates of traffic. Another outstanding issue is the feasibility of bulk traffic of lipids by LTPs. Thus far, the measured rates of LTP-mediated lipid transfer in vitro are orders of magnitude slower than the rates of nonvesicular traffic in vivo. Hopefully, it will be possible to account for the differences in rates by reconstituting an ERJ in vitro. Also, there is accumulating evidence that LTPs have roles other than in lipid transfer (Beh et al. 2009). For instance, OSBP might act as a sterol sensor or lipid-presenting protein because OSBP has been found to bind to two different phosphatase complexes and regulate extracellular-regulated kinase (ERK) signaling, and this is dependent on sterol (Wang et al. 2005). This may also be the case for the canonical LTP, Sec14, given new insight from structural and functional data (Schaaf et al. 2008) that it acts as a lipid-presenting protein for PI kinases in membranes in cis. Such a role may also be true for mammalian PITP family proteins, as Nir2 seems to play a similar role as yeast Sec14 in regulating exit from the TGN. Finally, Osh proteins in yeast appear to regulate cell polarity and vesicular transport through Cdc42 and Rho (Kozminski et al. 2006), but direct interactions between Oshs and the polarization machinery have yet to be demonstrated. Defining the biochemical activities of these proteins in vitro will undoubtedly help to define the precise roles for LTPs in signaling, cell polarity, and secretion.

CONCLUSIONS If this chapter is your first introduction to interorganelle membrane contacts, I hope that you now can appreciate the interconnectedness of membrane domains within cells and the impact these domains have on cell physiology. Although we have made significant progress in the past 10 years, our understanding of the fine organization of ERJs and the cellular process they contribute to remains extremely limited. This begins with the lack of identification of the structural building blocks of ERJs, and then of course how these molecules recruit other factors. By identifying the proteins that localize to ERJs and uncovering their functions, we will gain a better understanding of the diverse roles that these highly conserved ubiquitous cellular domains perform. Given that ERJs predate the evolution of the eukaryotic secretory pathway and that they are minimally involved in trafficking ions and lipids (and even possibly proteins [Schnabl et al. 2005]), they may represent the primordial cellular trafficking domain.

ABBREVIATIONS AMFR cER CERT COPI DAG ER ERAD ERJ ERK ERMES FAPP FFAT GlcCer

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autocrine motility factor receptor cortical ER ceramide transfer protein coat protein complex I diacylglycerol endoplasmic reticulum ER-associated protein degradation ER junction extracellular-regulated kinase ER–mitochondria encounter structure phosphatidylinositol-four-phosphate adapter protein two phenylalanines in an acidic tract (EFFDAxE) glucosylceramide

GLTP IP3R LTP MAM MCS Mito NVJ ORD ORP OSBP OSH PC PE PI PM

glycolipid transfer protein inositol triphosphate receptor lipid transfer protein mitochondria-associated membrane membrane contact site mitochondria nucleus–vacuole junction OSBP-related domain OSBP-related protein oxysterol-binding protein OSBP homolog phosphatidylcholine phosphatidylethanolamine phosphoinositol plasma membrane

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phosphatidylserine red fluorescent protein SREBP cleavage activating protein synaptoptagmin-like mitochondrial proteins store-operated calcium entry sterol regulatory element-binding protein

SRP START TGN VAP

signal recognition particles steroidogenic acute regulatory (STAR)-related domain trans-Golgi network vacuolar membrane-associated protein (VAMP)-associated protein

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CYTOSKELETAL DOMAINS C E L L UL AR ARC HI T E C TU R E is structurally supported by its cytoskeletal components: actin filaments (Chapter 12), microtubules (Chapter 14), and intermediate filaments (Chapter 16). The actin cytoskeleton plays a critical role in the formation of cellular protrusions and associated signaling and focal adhesion dynamics that regulate cell migration (Chapters 12, 17, and 22). At the plasma membrane, it regulates membrane domain organization (Chapter 1), but is also critically involved in the formation of microvilli (Chapter 13), specialized projections found on the apical surface of polarized epithelial cells (Chapter 21), as well as in the stabilization of adherens (Chapter 18) and tight (Chapter 19) junctions in epithelial cells. Microtubules (Chapter 14) derive from the microtubule organizing center (or centrosome) that corresponds to the basal body from which cilia derive (Chapter 15) and play a critical role in cellular polarization (Chapters 8, 21, and 22). Microtubules are critical regulators of intracellular membrane trafficking and regulate the distribution and dynamic exchange of various intracellular organelles including mitochondria (Chapter 6), the endoplasmic reticulum (Chapter 7), the Golgi apparatus (Chapter 8), and endosomes (Chapter 9). Intermediate filaments (Chapter 16) are specialized, depending on cell type, with epithelial keratin filaments stabilizing desmosomal junctions (Chapter 19) and microvilli (Chapter 13). The cytoskeleton therefore controls the dynamics and interactions of cell surface domains and intracellular organelles and plays a key role in determining cell organization and structure.

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THE ACTIN CYTOSKELETON Jonathan A. Kelber Richard L. Klemke

DEFINITION Actin is one of the most abundant proteins in the cell and is critical for survival of all organisms. It is necessary for many fundamental cellular processes including cell migration, axon pathfinding, differentiation, and proliferation (Richards and Cavalier-Smith 2005). On the other hand, improper regulation of actin contributes to numerous human diseases including developmental disorders, inflammation, and cancer (Richards and Cavalier-Smith 2005). Actin is a highly abundant protein in cells and is highly conserved throughout all eukaryotes (Erickson 2007). F-actin is composed of individual β-actin and γ-actin subunits polymerized into helical double-stranded filaments that are bundled into fibrous networks of cables to provide mechanical support, force generation, and transport of intracellular cargo. The F-actin cytoskeleton can be grouped into the following subcellular compartments: the membrane-associated cytoskeleton (including ruffles, filopodia, spikes, pseudopodia, and lamellapodia), the cortical actin rim, and internal stress fibers and cables (Pollard and Cooper 2009; Prasain and Stevens 2009; Fig. 12.1A). The membrane cytoskeleton and stress fibers are made up of shorter F-actin structures, while the cortical actin rim is composed of long F-actin fibers bundled together (Heimann et al. 1999). The membrane actin cytoskeleton possesses a thickness of only a few nanometers and primarily regulates membrane architecture, allowing the cell membrane to adopt a dynamic nature. The cortical actin rim lies adjacent to and beneath the membrane actin cytoskeleton. This structure is composed of dense actin fibers and is modulated by actinbinding and cross-linking proteins (Prasain and Stevens 2009). Finally, stress fibers are actomyosin contractile bundles capable of producing force (Hall et al. 1993; Hotulainen and Lappalainen 2006; Stricker et al. 2010). These three actin subdomain structures have diverse roles in controlling cell shape, membrane protrusion dynamics, and cell motility, and must be coordinately integrated to achieve the overall function of the cell (Stricker et al. 2010). Additionally, the actin cytoskeleton domain is a highly dynamic scaffold upon which mechanosensing and signal transduction events are spatially and temporally integrated in response to a diverse array of extracellular stimuli. Interestingly, little is known about how these dynamic events are globally integrated to modulate cell shape, motility, and overall cell physiology. Here, we provide an overview of the actin cytoskeleton domain and its regulatory components, and using informatics and extensive literature searches, we also present a contemporary view of the actin cytoskeleton interactome and its spatial compartmentalization within the cell interior.

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Figure 12.1. F-actin cytoskeleton network. (A) Fluorescence microscopy image of the F-actin network in a mouse embryonic fibroblast spreading on fibronectin (F-actin stained with Phalloidin-Alexa546) showing the membrane actin, cortical actin rim, and stress fiber structures. Scale Bar: 15 μm. (B) IPA-derived network for F-actin identifies 193 proteins that have been functionally or biochemically linked to F-actin. This network was constructed to include both direct (binding) (solid line) and indirect (regulatory) (dashed line) F-actin network partners.

HISTORICAL PERSPECTIVES In the middle of the twentieth century, actin and myosin were discovered to comprise an intricately organized force-producing center that makes up more than half of the total protein in muscle tissue (see Geeves et al. 2005; Geeves and Holmes 2005; Pollard and Cooper 2009). It was not until two decades later that actin was discovered in other cell and tissue types, demonstrating that the actin filaments found in muscle were a specific example of a more general cellular network (Hatano and Oosawa 1966; Adelman and Taylor 1969). At the molecular level, an atomic model of the actin filament was first generated in 1990 (Holmes et al. 1990). Subsequently, molecular components of the membrane cytoskeleton (see also Chapter 1) were initially identified in erythrocytes (De Matteis and Morrow 2000), and later resolved in nonerythroid cells (Wu et al. 2001). Actin, spectrin, and spectrin-binding proteins are the primary components of the membrane actin cytoskeleton (Pradhan et al. 2001; Baines 2009). Importantly, membrane proteins link to the spectrin-based membrane cytoskeleton directly and indirectly through adaptor proteins. Through the interactions between the membrane actin cytoskeleton and focal adhesion complexes (see also Chapter 17), cells are able to connect sites of cell–substrate contact to the cortical actin rim. While the membrane actin cytoskeleton and the cortical actin rim are distinguishable subcellular structures (Fig. 12.1A), several multiprotein complexes allow these actin subdomains to interact, which facilitates the regulation of membrane dynamics in concert with specific cellular functions (Prasain and Stevens 2009). Stress fibers consist of short F-actins with repeating units of myosin-II motor proteins (Cramer et al. 1997) which are cross-linked by α-actinin and other actin-binding partners (Hotulainen and Lappalainen 2006). These actomyosin contractile bundles are integral during centripetal cellular tension generation, and for regulating cell–extracellular substrate interactions

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(Small et al. 1999; Dudek and Garcia 2001). While the membrane actin cytoskeleton and the cortical actin rim are close to the cell membrane, these stress fibers exist throughout the cell (Fig. 12.1A). In a unique balancing act, the stress fibers generate a centripetal force that offsets the centrifugal force generated by the cortical actin rim. In addition to playing an integral role in regulating cell morphology and motility, stress fibers facilitate intracellular transport of signaling molecules throughout the cell in order to spatially and temporally orchestrate signaling events. As discussed below, the actin cytoskeletal domain represents a collection of highly dynamic structures that contribute to the morphology and structure of the cell, as well as function, to receive and transduce important signals in response to extracellular cues.

MOLECULAR COMPOSITION OF THE DOMAIN Components of the Actin Cytoskeleton Network The actin cytoskeleton represents a large cellular scaffold with which numerous cytoplasmic and membrane proteins assemble and interact. While extensive research over the years and the advent of functional genomics and proteomics has significantly increased the number of known actin cytoskeletal components and interacting partners, our understanding of how these components are functionally integrated and spatially organized in the cell is quite limited. Nonetheless, further understanding the actin regulatory network as a whole can help identify functional protein units that operate together in a defined space within the cell and help identify where and how certain multicomponent interactions occur. Simple linear pathways fail to describe modular interactions and provide little insight into possible feedback mechanisms, drug interactions, or cellular consequences of silencing specific genes. To catalogue the known actin regulatory components and begin to understand their complex interactions, we utilized the Ingenuity software program and Knowledge Database (IPA). Ingenuity is a bioinformatics resource that facilitates the mapping of signaling networks and interactomes involving a range of biomolecules as they relate to normal physiological and pathophysiological functions (http://www. ingenuity.com) (Gehlenborg et al. 2010; O’Donoghue et al. 2010). By using this comprehensive informatics program, it is possible to model, analyze, and understand the complex integration of biological systems. IPA utilizes the Ingenuity Knowledge Base, a large repository of biological interactions and functional annotations created from millions of individually modeled relationships between proteins, genes, complexes, cells, tissues, drugs, and diseases. These modeled relationships include links to original articles and are manually reviewed biweekly for accuracy. The Ingenuity Knowledge Base includes information from a wide range of published biomedical literature, textbooks, reviews, internally curated knowledge (such as pathways), and a variety of established informatics sources and databases (e.g., EntrezGene, RefSeq, online mendelian inheritance in man [OMIM] disease associations, KEGG, GWAS, LIGAND, and BIND). Using the IPA software program and F-actin as the hub of our network, we have assembled an F-actin network of 193 interaction partners and a diverse array of regulatory components (Fig. 12.1B) consisting of cytokines/growth factors, ion chanels, kinases, membrane receptors, and transcription factors. This network was constructed to include both direct (binding) (solid line) and indirect (regulatory) (dashed line) F-actin network partners. For example, α-actinin is catalogued as a direct binding partner of F-actin based on its role in bridging the actin cytoskeleton to focal adhesion complexes, while the extracellular cytokine epidermal growth factor (EGF) is catalogued as an indirect regulatory

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molecule of F-actin based on its ability to directly activate signaling pathways that impinge on F-actin remodeling (e.g., small GTPases) (Feldner and Brandt 2002; Mogilner and Keren 2009). In contrast, IPA does not classify FAK as a member of the F-actin cytoskeleton since it does not directly bind to actin or directly/indirectly regulate F-actin remodeling. This differs from other focal adhesion proteins such as vinculin or paxillin that have been reported to bind to F-actin directly (Martin 1998; Tsubouchi et al. 2002; Nolz et al. 2007). For further information on focal adhesion complexes, see Chapter 17 and these reviews (Martin et al. 2002; Mitra and Schlaepfer 2006; Zaidel-Bar et al. 2007). We also mapped the known interactions between these components and their spatial organization in the cell, creating a systems view of the actin cytoskeleton interactome (Fig. 12.2). Over 80% of the known F-actin-interacting proteins (162 components in the F-actin network, Fig. 12.1B) are functionally or biochemically linked with one another in this interactome. As an example, F-actin nucleation, polymerization, length, turnover, branching and cross-linking are intricately regulated by numerous proteins such as profilin, cofilin, formins, Wiskott–Aldrich syndrome family protein (WASP)/Scar, gelsolin, and Arp2/3 (Pollard and Cooper 2009). As expected, these proteins are identified as key cytoplasmic F-actin interaction and regulatory hubs that are highlighted in the F-actin interactome. However, while these signaling nodes are primarily cytoplasmic, they are functionally connected to proteins that reside in the plasma membrane and nucleus.

Components Regulating Actin Polymerization and Branching This F-actin interactome reveals some novel and unique information about actin polymerization/depolymerization and branching mechanisms. In fact, we have identified 13 regulators of these processes by highlighting the direct interacting and regulatory partners of Arp2/3 proteins in this interactome (Fig. 12.2, yellow). Actin monomers (G-actin) can form long, stable filaments (F-actin); however, early polymerization events exhibit lower stability than late polymerization events, once long actin filaments have been formed (Pollard 2007). The formation of actin filaments coincide with the binding and hydrolyzing of adenosine triphosphate (ATP) to adenosine diphosphate (ADP), which then primes F-actin for depolymerization during cellular events that depend on the dynamic rearrangement of the actin cytoskeleton (Pollard 2007). Actin filaments are polar due to the directional assembly of actin monomers. Importantly, this facilitates preferential filament growth in one direction, allowing for cells to regulate the direction of actin polymerization to control cell motility and morphology as well as intracellular transport of signaling molecules. This process is primarily regulated by formins and α-catenin, where formins bind to the barbed end of the filament to facilitate elongation (Fig. 12.3A; Kobielak et al. 2004). As previously reported (Pfaendtner et al. 2010), cofilin is a key cytoplasmic modulator of F-actin depolymerization that functions to offset the role of formins and α-catenin (Fig. 12.2). Also, several mechanisms exist for the formation of new filaments including branch formation on the side of an existing filament, cleavage of existing filaments to form two new ends, or filament polymerization from the pool of actin monomers (Pollard 2007). The Arp2/3 complex is an example of a key component of the F-actin interactome that is well described and plays a central function in actin filament formation. The Arp2/3 complex contains seven subunits (Arp2/3 and ARPC1-5), two of which (2 and 3) have similar structures and sequences to that of actin. The complex becomes activated via its interactions with a WASP family protein sequence known as WCA and an actin filament (Robinson et al. 2001; Kiselar et al. 2007; Nolen and Pollard 2007). When active, Arp2 occupies a filament-like conformation next to Arp3. The first actin subunit, bound to the

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Figure 12.2. F-actin cytoskeleton interactome. The F-actin interactome demonstrates that 162 of the proteins from the F-actin cytoskeleton network are functionally and/or biochemically connected with one another. Proteins shaded yellow refer to those involved in actin polymerization, depolymerization, and branching functions. Proteins shaded in blue refer to those involved in the regulation of myosin–actin force generation. Green shading refers to proteins found in both the blue and yellow populations. Finally, proteins shaded in red indicate small GTPases and their interaction partners, a group of molecules well-documented for their role in mediating actin reorganization. To generate this interactome, the F-actin network in Figure 12.1B was further analyzed by connecting only direct interactions that have been documented in the Ingenuity Knowledge Base and limited to activation/inhibition, protein–protein interactions, regulation of expression, and phosphorylation events. Interactions were characterized as binding (line) or direct modification (line with arrow) of a target molecule.

WCA region of N-WASP, binds at the barbed end of Arp2. Ultimately, these interactions facilitate the association of an actin filament “pointed end” (Fig. 12.3A) with a centrally located actin monomer in its partnering filament. While important details still need to be elucidated regarding the mechanism of Arp2/3-mediated branched actin filament

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Figure 12.3. F-actin formation and phosphorylation within the cell. (A) The Arp2/3 complex and formins work together with actin monomers (G-actin) to generate actin filaments (F-actin). Arp2/3 complexes are stable assemblies of seven subunits that are believed to regulate actin branching, while formins are dimers that sit on growing ends of actin filaments and mediate the elongation of linear actin networks. In collaboration with profilin, these molecules recruit actin monomers to facilitate actin filament assembly. Actin filaments are polarized and the “barbed ends” preferentially bind actin monomers to facilitate elongation of the filaments. (B) Fluorescence image of the F-actin cytoskeleton (red) and phosphotyrosine proteins (green) in a mouse embryonic fibroblast spreading on fibronectin. Note the strong phosphotyrosine staining at the end of many of the stress fibers terminating at focal adhesions, which are rich in actin-binding and phosphotyrosine proteins. F-actin stained with Phalloidin-Alexa546 (red) and phosphotyrosine proteins stained with anti-phosphotyrosine (4G10, Millipore, Billerica, MA) + antimouse-Alexa488 (green). Scale Bar: 15 μm.

generation (Insall and Machesky 2009), it is likely that the Arp2/3 complex makes new filaments by side-branching. Furthermore, it appears that the Arp2/3 subunits in the branching structure form the initiation seed for the nascent daughter filament. Molecular modifiers (e.g., kinases) and other compounds (e.g., ATP) are also likely to be important components involved in regulating the conformational changes that activate the Arp2/3 complex. LeClaire et al. (2008) have shown that the Arp2/3 complex undergoes tyrosine and threonine phosphorylation and that these posttranslational modifications are required for proper actin nucleation and lamellapodia formation. However, little is known about the kinases or phosphatases that regulate these crucial events in the cell. Consequently, the F-actin interactome demonstrates that there are no direct kinases connected to the Arp2/3 complex (Fig. 12.2).

Components Regulating Actin Cytoskeleton Plasticity The actin cytoskeleton is regulated by a complex interplay of signals that target the transcription of actin and its binding partners and effectors. These signals also control posttranslational modifications of actin-associated proteins, which in turn modulate effector protein levels, protein–protein interactions, and actin polymerization/depolymerization events. Here we will primarily focus on the biochemical regulation of the actin cytoskeleton as it relates to actin polymerization/depolymerization and effector modulation through

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phosphorylation. For more detailed information on transcriptional regulation and subcellular localization of actin, see these reviews (Schleicher and Jockusch 2008; Zheng et al. 2009). The inherent plasticity of the actin cytoskeleton stems from the ability of actin to polymerize and depolymerize in response to extracellular cues. The ability of cells to sense their extracellular environment and respond mechanically is critical for mediating cell shape change, migration, and cytokinesis. The mechanosensing function of the actin cytoskeleton is primarily mediated by integrins, G-protein-coupled receptors, and tyrosine kinase receptors at the membrane surface (Fig. 12.2). These receptors relay a cascade of signals that eventually impinge on the Rho family of small GTPases (Cdc42, Rac, and RhoA), which temporally and spatially modulate the required changes in the actin cytoskeleton (Hall et al. 1993; Ridley et al. 2003; Burridge and Wennerberg 2004; Raftopoulou and Hall 2004; Myers and Casanova 2008; Insall and Machesky 2009; Jiang et al. 2009; Mogilner and Keren 2009). Using the F-actin interactome, we have identified 21 small GTPase binding partners, targets, or regulators (Fig. 12.2, red) that control actin cytoskeletal dynamics. In addition to regulating actin plasticity, these signaling pathways also regulate cellular functions that depend upon the contractility of the actin–myosin system (Chodniewicz and Klemke 2004b; Clark et al. 2007; Nambiar et al. 2010; Pollard 2010). For example, our group has previously demonstrated that ERK targets myosin light-chain kinase (MLCK) in order to stimulate actin–myosin contractility (Klemke et al. 1997; Cheresh et al. 1999). The MAPK and myosin signaling pathways are identified as integral regulators of cytoskeletal contractility in the F-actin interactome (Fig. 12.2) and we have identified 17 components of this module (blue). The unique ability of the actin cytoskeleton to remodel as well as mediate myosin contractility provides sensitive and powerful machinery capable of transducing force throughout the cell (Stricker et al. 2010).

Kinase and Phosphoprotein Components Regulating the Actin Cytoskeleton Many processes that require the spatial and temporal organization and regulation of the actin cytoskeleton occur via reversible protein phosphorylation on serine, threonine, or tyrosine amino-acid residues. These phosphorylation events represent a major mechanism utilized by cells to coordinate the localization, activation, stability, and assembly of macromolecular signaling complexes that interact with the actin cytoskeleton (Linding et al. 2007). However, aberrant regulation of protein localization and protein phosphorylation can also contribute to numerous pathologies—this is especially prominent in cancer (Brugge 1993; Aranda et al. 2006; Mitra and Schlaepfer 2006; Wang et al. 2010). Therefore, it is imperative to understand these events that are critical regulatory elements in the biology of the actin cytoskeleton. With this goal in mind, we have generated an F-actin phosphorylation interactome with molecules that are involved in phosphorylation events during actin cytoskeleton regulation (e.g., phosphoproteins, kinases, and phosphatases) (Fig. 12.4). Our analysis has uncovered 81 phospho-related proteins that are functionally linked within the F-actin network. This represents approximately 40% of the complete F-actin network (Fig. 12.1B), indicating that phosphorylation is a major mechanism that regulates this domain. This can also be seen in the overlay image of phosphotyrosine and actin cytoskeleton immunofluorescence staining of fibroblast cells (Fig. 12.3B). Within this subnetwork (Fig. 12.4), we have identified 13 kinases (green) and 2 phosphatases (red) that have previously been reported to regulate actin cytoskeletal dynamics, including p21-activated kinase (PAK),

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Figure 12.4. F-actin phosphorylation interactome. The F-actin phosphorylation interactome demonstrates that 81 of the proteins from the F-actin cytoskeleton network are functionally and/or biochemically connected with one another via phosphorylation events. Proteins shaded green are kinases and proteins shaded in red are phosphatases. To generate this interactome, the F-actin network in Figure 12.1B was further analyzed by connecting both direct and indirect interactions that have been documented in the Ingenuity Knowledge Base and limited phosphorylation events. Interactions were characterized as binding (line) or directional modification (line with arrow) of a target molecule.

LIMK, Rho-associated kinase (ROCK), Abl1/2, and protein kinase A/C (PKA/C; Howe 2004). Interestingly, the F-actin phosphorylation interactome identifies 10 extracellular factors as also being linked to the phospho-regulation of the actin cytoskeleton. However, this is not surprising as many of these factors indirectly activate signaling pathways that regulate the F-actin domain via phosphorylation events (these indirect regulatory events are denoted by dashed lines).

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FUNCTIONAL IMPLICATIONS AND ROLES FOR DOMAIN ORGANIZATION Spatial Organization of the Actin Domain in Polarized Cells A hallmark of polarized cells is their ability to compartmentalize cellular contents and partition key physiological processes into subregions within the cell interior. This process leads to the segregation of subnetworks of multiprotein complexes to different poles of the cell. The polarized signaling response can then direct a division of labor in the cell that dictates its specific functions and morphological attributes. In many cases, cell polarity is determined by its interconnection with neighboring cells and exposure to gradients of chemokines and extracellular matrix proteins, which provide directional cues. This is most apparent in the establishment of epithelial basolateral polarity, cell migration polarity, and growth cone navigation during brain development (see also Chapters 21 and 22). As discussed below, polarization of the actin cytoskeleton domain plays a prominent role in all these processes. The actin cytoskeleton and associated regulatory proteins are distinctly polarized in migrating cells to form a leading front and trailing rear compartment. At the front of the cell, actin polymerization and assembly drive membrane protrusion leading to the formation of filopodia and the lamellipodium. At the sides and rear of the cell, cortical actin provides cell rigidity, whereas stress fibers work in concert with myosin to facilitate strong force and contraction events that pull up the trailing tail from the underlying substratum. The repeated cycle of membrane extension at the front and tail retraction at the back facilitate cell translocation (Ridley et al. 2003; Chodniewicz and Klemke 2004a; VicenteManzanares et al. 2005). The super family of small GTPases, kinases, and membrane receptors spatially signal to the actin polyermization and actomyosin contractile machinery to induce polarization and motility (Insall and Machesky 2004; Sanz-Moreno and Marshall 2009). The spatial intricacies of small GTPase signaling are best illustrated by the Rho family members Rac, RhoA, and Cdc42, which have been extensively studied in recent years in relation to mesenchymal mode of cell migration (Hall et al. 1993; Small et al. 1999; Ridley et al. 2003; Insall and Machesky 2009). These proteins are regulated by a complex network of upstream signaling pathways via GEFs and GAPs that couple GTPase activity to actin nucleation and actin cytoskeletal remodeling events (Figs. 12.1 and 12.2, red). Recent work aimed at understanding their spatial signaling in migrating cells has revealed complex and overlapping activities of these proteins. Previously, it was thought that the structural output of GTPase activity was linear with Rac mediating lamellipodia formation, Cdc42 mediating filopodia formation, and RhoA mediating stress fiber formation. This led to the belief that RhoA activity in migrating cells predominated at the rear of the cell to mediate tail retraction, while Cdc42 and Rac controlled lamellipodia and filopodia formation at the front (Ridley et al. 2003; Burridge and Wennerberg 2004; Raftopoulou and Hall 2004). However, the development of methods to selectively isolate the front and back of polarized cells for biochemical analyses and the advent of förster resonance energy transfer (FRET)-based biosensors to monitor spatial GTPase activity in migrating cells have challenged this belief (Cho and Klemke 2002; Brahmbhatt and Klemke 2003; Pertz et al. 2006; Pertz et al. 2008; Machacek et al. 2009). Both of these methods revealed a significant level of RhoA activity in the lamellipodium. Subcellular imaging of RhoA biosensor activity showed that it was concentrated in a sharp band directly at the edge of membrane protrusions, and was in peripheral ruffles and pinocytic vesicles. It was also observed sporadically in retracting tails, and was low in the cell

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body. However, it is notable that in contrast to randomly migrating cells, growth factorinduced membrane protrusions have low RhoA activity, potentially because growth factors strongly activate Rac, which has previously been shown to antagonize RhoA activity (Sander et al. 1999; Nimnual et al. 2003). While additional work is necessary to clarify the interrelationship of Rho family proteins and their spatial organization in migrating cells, this body of work highlights the functional compartmentalization of actin dynamics in polarized migrating cells and its complex spatial regulation. Finally, recent work has uncovered novel modes of cell migration that do not involve actin-mediated membrane protrusion, which is characteristic of mesenchymal movement (Friedl and Gilmour 2009; Friedl and Wolf 2010).This amoeboid-like migration mode is characterized by a blebbing-like protrusive process driven by internal changes in hydrostatic pressure and ROCK contraction. While the actual mechanism of bleb formation and the directional cues that drive amoeboid-like migration have not yet been fully defined, the cortical actin rim and membrane actin cytoskeletal structures are likely altered under these conditions due to changes in hydrostatic pressure. Glandular epithelia in organs consist of cells that are organized around a central lumen. Such cells exhibit an asymmetric distribution of proteins along the apical, lateral, and basal surfaces (Suzuki and Ohno 2006; Aranda et al. 2008). Direct association of the actin cytoskeleton with cell adhesion proteins in cell–cell junctions plays a central role in the assembly and stabilization of barrier function and tissue integrity (see Chapters 18 and 19 on adherens and tight junctions). The resulting apical–basal polarity of the actin cytoskeleton and cell–cell junctions are essential for proper organ function and is intimately regulated by Rho family proteins (Drubin and Nelson 1996; Samarin and Nusrat 2009; Terry et al. 2010). On the other hand, during pathogenesis, disruption of the polarity apparatus by oncogene expression leads to loss of epithelial cell–cell contacts, tumor invasion, and cancer progression to a more malignant state. For example, ErbB2 overexpression in normal mammary epithelial cells directly uncouples the Par6–Par3–aPKC–CdC42 polarity protein complex by displacing Par3 and Cdc42 from the Par6–aPKC complex (Aranda et al. 2006). In the absence of cell–cell contacts, the Par6–aPKC complex exists independently of Par3. However, when epithelial cells come into contact, GTP-bound Cdc42 and Par3 are recruited to form an active Par complex (Par3–Par6–aPKC–GTP– Cdc42) (Suzuki and Ohno 2006). This polarity complex maintains the stability of both the actin and microtubule cytoskeleton in order to maintain epithelial cell polarity. Thus, one mechanism by which ErbB2 contributes to cancer progression is by disrupting normal epithelial polarity and by driving destabilization of the actin cytoskeleton and the loss of cell–cell contacts leading to a more invasive phenotype. Interestingly, an emerging picture indicates that epithelial polarity proteins and cell–cell junction proteins like occludins and zona occludins one (ZO-1) can also control the actin cytoskeleton and microtubule networks to direct polarized cell migration (Tsukita et al. 2008; Franke 2009; Franke et al. 2009; Tuomi et al. 2009). This has important implications, as metastatic cancer cells may be able to hijack the polarity and junctional complex machinery to invade tissue and disseminate to distant organs. If so, novel therapeutics might be designed to block this process and the spread of cancer.

Neuritogenesis and Growth Cone Dynamics The development of a properly connected and functional nervous system depends on the ability of neurons to guide their axons along appropriate routes in the embryo to locate a suitable synaptic partner, a process known as pathfinding. Axon guidance relies on the pathfinding abilities of the growth cone, which is a highly dynamic and motile structure

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at the tip of growing neurites and axons that respond to molecular guidance cues in the extracellular environment (Arimura and Kaibuchi 2007). These factors include chemokines, growth factors, and extracellular matrix proteins that attract or repel growth cones to guide their directional migration to their target cell within the developing nervous system (Geraldo and Gordon-Weeks 2009). The guidance molecules activate intracellular signaling pathways that primarily converge on the growth cone cytoskeleton (Zhou and Snider 2006), which consists of microtubules and actin filaments (F-actin). The dynamic behavior of these filaments has been extensively studied (Kalil and Dent 2005; Lowery and Van Vactor, 2009). Here we will focus on the role of the actin cytoskeleton in growth cone dynamics (see also Chapter 22 on neuronal domains). For a review on the role of the microtubule cytoskeleton, see Conde and Caceres (2009). As in migrating cells, the small GTPase family of effectors are central regulators of the dynamic actin cycles that control neurite protrusion and growth cone steering (Jones et al. 2004; Pertz et al. 2008). This includes peripheral F-actin assembly, retrograde flow of F-actin, and the depolymerization and recycling of F-actin. The process of actin polymerization at the leading edge during growth cone protrusion is controlled by multiple regulators, including the Arp2/3 complex and the formins (Korobova and Svitkina 2008). Additional molecules including ENA/VASP play a role in this regulatory process as well. In this context, the Arp2/3 complex functions to regulate Rac1 and Cdc42 (Goley and Welch 2006). Notably, inhibition of the Arp2/3 complex in neurons prevents lamellipodia and filopodia protrusions, which coincides with increased RhoA activity. However, work remains to determine the complete role of RhoA during growth cone motility and guidance (Korobova and Svitkina 2008). The ENA/VASP proteins enhance F-actin elongation by multiple methods, such as attracting actin subunits for further F-actin elongation (Drees and Gertler 2008). While ENA/VASP proteins are primarily regulated through kinases downstream of guidance cue signaling (Drees and Gertler 2008), the ENA regulator Ableson tryrosine kinase can function through Rac and Rho (Jones et al. 2004; Sini et al. 2004) which suggests that crosstalk may exist between these proteins during the dynamic rearrangement of the actin cytoskeleton as the growth cone migrates and is guided by extracellular cues.

Contemporary Methods to Study Actin Cytoskeleton Signaling One of the most exciting technical breakthroughs in the last two decades has been the advent of GFP and FRET-based technologies for studying protein–protein interactions and protein activity in living cells with high spatial and temporal resolution (Heim et al. 1994; Heim et al. 1995; Pertz and Hahn 2004; Hodgson et al. 2008; Machacek et al. 2009; Hodgson et al. 2010) (see also Chapters 1 and 5 on the use of these techniques to study membrane domains). There are several excellent, in-depth reviews available on this large topic in the literature (Giepmans et al. 2006; Tsien 2006, 2009; Hodgson et al. 2010). Here, we will provide only brief highlights of recent work in relation to the actin cytoskeleton. This approach has proven ideal for studying the dynamic actin cytoskeleton and its binding partners (e.g., small GTPases) in relation to various cellular processes such as membrane protrusion and/or retraction (Ting et al. 2001; Pertz and Hahn 2004; Pertz et al. 2006, 2008; Hodgson et al. 2008). In addition to small GTPase biosensors, N-WASP biosensors have enabled the spatiotemporal analysis of N-WASP function during actin polymerization at the leading edge of lamellapodia and invadopodia in carcinoma cells (Lorenz et al. 2004a, b; Parsons et al. 2005). More recently, fluorescently tagged Lifeact constructs have been developed to visualize the remodeling of the actin cytoskeleton (Riedl et al. 2008).

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Together with total internal reflection fluorescence (TIRF) microscopy techniques, these reagents have enabled researchers to investigate the membrane–cytoskeleton interface in real time to observe dynamic changes in the molecular composition and regulation of the actin cytoskeleton. Also, high throughput screening studies using these fluorescent technologies to monitor cytoskeletal changes in cell models of disease are underway (Brugge 1993; Clark and Brugge 1995; Bakal et al. 2007; Simpson et al. 2008; Winograd-Katz et al. 2009). This approach will likely provide important insight into the progression of these diseases and provide specific therapeutic reagents that elicit their effects on the cytoskeleton. The work presented here provides an initial glimpse of the actin cytoskeleton interactome; however, there remains a substantial need for a more complete inventory of the actin-associated proteins and their integration at a systems level. To achieve this goal it will be necessary to fully define the actin cytoskeleton proteome and phosphoproteome. In this regard, large genomic and proteomic screens that target modifiers of the actin cytoskeleton have been limited (Brugge 1993; Clark and Brugge 1995; Bakal et al. 2007; Simpson et al. 2008; Winograd-Katz et al. 2009). Specifically, biochemical methods for direct extraction of the actin cytoskeleton and its associated proteins that are compatible with current mass spectrometry-based proteomics and phosphoproteomics techniques are very limited (Ramsby and Makowski 1999; Borner et al. 2009). Once these methods are resolved and the actin cytoskeleton proteome is defined, powerful informatics programs and computer models can be used to understand these protein–protein interactions, signal transduction events, and kinase/phosphatase activation networks. To begin to address these limitations in our laboratory, we have combined quantitative large-scale proteomic and phosphoproteomic analyses with subcellular fractionation of actin-rich protrusions (pseudopodia and neurites) (Cho and Klemke 2000; Brahmbhatt and Klemke 2003; Cho and Klemke, 2002; Wang et al. 2007; Pertz et al. 2008; Wang et al. 2010). Using these methods along with enrichment procedures for phosphoproteins, thousands of phosphorylation sites and phosphoproteins have been annotated and assigned to signaling pathways that control cytoskeletal remodeling in these structures. This approach also allowed us to uncover low abundant and novel phosphoproteins that regulate the pseudopod and neurite cytoskeletons including the novel proteins LASP-1 and the atypical kinase PEAK1 (Lin et al. 2004; Wang et al. 2010). LASP-1 is a novel focal adhesion and actin-binding phosphoprotein necessary for cell survival and migration. Interestingly, Abl kinase activation by apoptotic agents specifically promotes phosphorylation of LASP-1 at tyrosine 171. This phosphorytion event blocks LASP-1 localization to focal adhesions leading to the induction of cell death. Recent research has further linked LASP-1 to brain development and cancer progression (Grunewald and Butt 2008). PEAK1 is a large 190-kD nonreceptor tyrosine kinase that localizes to actin-rich protrusions, focal adhesions, and stress fibers in adherent cells, and is a member of the Src/CAS/Crk and ERK signaling pathways (Wang et al. 2010). It is regulated by growth factor receptors and Src kinase activation and is necessary for proper cell spreading, migration, and cancer cell proliferation. PEAK1 regulates ERK activity in cancer cells and may also be a good biomarker for colon cancer as it is overexpressed in approximately 80% of patients with this disease. In related work Shankar et al. (2010), used a combination of genomic, proteomic, and pseudopodia purification methods to discover a panel of cytoskeleton proteins that mediate cancer cell invasion. Interestingly, knockdown of several of these proteins including AHNAK, septin-9, eIF4E, and S100A11 in metastatic cells reduced cytoskeleton dynamics and induced mesenchymal–epithelial transition (MET). Together these studies provide an important new source of biomarkers that may predict metastatic potential of cancer cells. These studies also highlight the power of using con-

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temporary “-omics” approaches for identifying actin cytoskeletal-associated proteins in normal and diseased cells.

FUTURE PERSPECTIVES In summary, the actin cytoskeleton is more than a simple support system that provides cell shape. It is a highly dynamic structural and signaling scaffold on which mechanosensing and signal transduction events are spatially and temporally integrated in response to a diverse array of extracellular stimuli. Numerous important cellular processes depend on the actin cytoskeleton including cell migration, axon pathfinding, differentiation, and proliferation. Although progress has been made in understanding actin polymerization and many important F-actin modulators have been identified, there is still a central need to better define the actin cytoskeleton proteome, understand how the nuts and bolts of this domain are assembled into a functional cytoskeletal network, and how it is integrated with the other cellular domains that comprise a complete functioning cell.

ABBREVIATIONS ABL ADP ARP ARPC ATP BIND CAS EGF ENA/VASP ERK FAK FRET GAP GEF GFP

abelson kinase adenosine diphosphate actin-related protein ARP complex adenosine triphosphate biomolecular interaction network database Crk-associate substrate epidermal growth factor enabled/vasodilator-stimulated phosphoprotein extracellular-regulated kinase focal adhesion kinase förster resonance energy transfer GTPase activating protein GTPase exchange factor green fluorescent protein

GWAS IPA KEGG LASP-1 LIMK MAPK MET MLCK OMIM PAK PEAK1 PKA/C ROCK TIRF WASP ZO-1

genome-wide association studies ingenuity pathway analysis Kyoto encyclopedia of genes and genomes LIM and SH3 domain protein one LIM kinase mitogen-activated protein kinase mesenchymal–epithelial transition myosin light-chain kinase online mendelian inheritance in man p21-activated kinase pseudopodium-enriched atypical kinase one protein kinase A/C Rho-associated kinase total internal resonance fluorescence wiskott-aldrich syndrome family protein zona occludins one

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MICROVILLI Florent Ubelmann Sylvie Robine Daniel Louvard

DEFINITION Microvilli consist of fingerlike membrane protrusions supported by organized bundles of actin filaments that are present on most cell types irrespective of their origin. Few epithelial cell types show ordered and homogeneous arrays of microvilli (or microvilli-like structures) that cover the entire cell apex. Two major examples of cellular domains made of ordered microvilli include the brush border and the stereocilia (see Fig. 13.1B, D). Brush borders are observed on the intestinal and proximal kidney epithelia, and are constituted by thousands of 1- to 2-μm-long tightly packed microvilli, homogeneous in length and width, well adapted to the intense absorptive processes performed by these epithelia. Moreover, these highly specialized domains display a molecular composition suitable for their physiological functions such as enrichment in digestive enzymes and ion exchangers in intestinal microvilli. On the inner ear cell apexes, stereocilia are microvilli-like structures ranging from 1 to 120 μm organized in a “staircase,” allowing mechanosensory transduction elicited by sound pressure. It is worth stressing that stereocilia, named after cilia, which are microtubule-based cellular protrusions, are also supported by bundles of actin filaments. They share numerous structural characteristics with intestinal microvilli and possibly derive from common rudimentary microvilli. Within each intestinal microvillus, actin polymerization at the growing extremities of 20–30 oriented actin filaments organized in a dense meshwork provides sufficient mechanical force and resistance to maintain the membrane protrusion. The exceptional homogeneity of microvilli-based domains implies the existence of precise regulatory mechanisms to control assembly and dynamics of the brush border. Indeed, the actin filaments within microvilli are tightly bundled together by actin-bundling proteins and are laterally attached to the membrane by myosin bridges and proteins of the ezrin–radixin–moesin (ERM) family. In addition, the microvillar actin bundles extend deeply inside a subapical zone, the so-called terminal web, where each microfilaments bundle is referred as the rootlet of the microvillus. The rootlets are interconnected by a network made of spectrin proteins and are connected to an underlying cytokeratin meshwork, thus providing stabilization to the overall brush border.

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Figure 13.1. Comparison of the different molecular components involved in the shape regulation of intestinal microvilli (right) and stereocilia (left). (B, D) General aspect of brush border domains made of (B) microvilli found on intestinal cell apexes and (D) stereocilia found on inner ear cell apexes. (C) Different proteins organize the parallel actin bundles that support microvilli structures. (A) Highlight on the tip of microvilli structures where major regulation of actin polymerization is ensured by protein complexes identified in stereocilia. Such complexes remain to be found on intestinal microvilli.

HISTORICAL PERSPECTIVE Discovery and Identification of the Brush Border Domain Microvilli were originally identified as the individual components of the intestinal brush border at the apex of enterocytes, the major intestinal cell type. This peculiar cell surface was first described by Henle in 1841 and referred as “free border” with no apparent structure. In the following years, several studies made by Gruby and Delafon in 1843 and Funke in 1855 depicted the striated aspect of the intestinal brush border observed by light microscopy. Two major interpretations were then elaborated: Gruby and Delafon and later, Zimmerman and Hindenhain in 1898 and 1899, respectively, proposed that striations might represent fine protoplasmic filaments within the “free border” describing them as “analogous to cilia.” In contrast, Funke in 1855 suggested that striations might be “pore-canals” that could be involved in nutrient absorption and transport (Baker 1942). Granger and

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Baker revealed the ultrastructure of the brush border by electron microscopy in 1950, showing the exquisite regular array of microvilli covering the apical pole of intestinal cells and thus ended the controversy (Granger and Baker 1950).

Ultrastructural Organization The brush border covering each human intestinal cell, as described by Brown et al. in 1962, consists of about 1700 microvilli measuring 1–2 μm in length and 0.1 μm in diameter, increasing 30- to 40-fold the intestinal surface (Brown 1962). Millington and Finean detected, within intestinal microvilli, filamentous structures extending in the terminal web and further suggested that these microvillar filamentous cores could be essential to support the membrane protrusions (Millington and Finean 1962). First proposal that actin filaments could be the central component of the microvillar filamentous core was made in 1969 by Ishikawa and colleagues who succeeded to decorate extracted filamentous cores by heavy meromyosin, which forms characteristic arrowhead complexes with actin filaments in situ (Ishikawa et al. 1969). The formal demonstration was done in 1971 by Tilney and Mooseker who characterized the major microvillar molecular component as indiscernible to smooth muscle actin on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) (Tilney and Mooseker 1971). This work was based on a reliable procedure for isolation of intact hamster brush borders that is still in use nowadays (Miller and Crane 1961). A detailed analysis of actin microfilament organization, by Tilney and Mooseker in 1975, showed that the microvillar actin core is composed of actin filaments identically polarized toward the microvillus tip. They also described the lateral membrane attachment of the actin cores along the microvilli through regular cross bridges (Mooseker and Tilney 1975). Hirokawa and colleagues in the 1980s, using the quick-freeze deep-etch technique followed by electron microscopy, showed an impressive view of the intertwining of the different cytoskeletal meshworks supporting the brush border (Hirokawa and Heuser 1981). In 1982, Pollard and Mooseker provided a new insight on the morphogenesis of these structures. Indeed, they realized an in vitro polymerization assay using purified filamentous actin (F-actin) extracted from microvilli and showed that the actin monomer addition takes place exclusively at the microvillus tip, suggesting that actin polymerization could drive the membrane protrusion (Pollard and Mooseker 1981). However, considering the homogeneity in length of each microvillus that constitutes the brush border, actin polymerization and organization have to be precisely regulated. Besides actin, the three other major molecular components were detected as being three actin-binding proteins that modulate actin assembly and organization: villin (95 kDa), plastin-1 (formerly I-fimbrin), and myosin-1a (Bretscher and Weber 1979, 1980; Matsudaira and Burgess 1979; Howe and Mooseker 1983).

MOLECULAR DETERMINANTS INVOLVED IN THE CONTROL OF THE MICROVILLI SHAPE In order to give a better understanding on the microvillus morphogenesis, we will briefly recall some essential biochemical properties of actin, the protein that drives the membrane protrusion (see also Chapter 12 on the actin cytoskeleton). The polymerization of actin monomers establishes polarized actin microfilaments showing different association/ dissociation rates at both extremities of each protomer. Indeed, the addition of monomeric actin is favored at the so-called barbed end, resulting in actin assembly, whereas it is

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unfavorable at the so-called pointed end where actin disassembles. This results in a dynamic equilibrium between polymerization and depolymerization leading to the “treadmilling” phenomenon that maintains the microfilament to a constant length. However, the “treadmilling” is highly dependent on free actin monomer concentration: In case of an excess or a lack of free actin monomers, the equilibrium between the biochemical reactions leading to monomer addition/removal is disturbed, leading to a growing or shrinking actin filament (Pollard and Borisy 2003).

Actin: The Major Microvillar Component Actin assembly occurs at the microvillus tip, providing force for the membrane deformation, whereas disassembly takes place at the base of the actin bundle (Mooseker et al. 1982; Stidwill et al. 1984). The F-actin core supporting the microvillar structure is therefore constantly renewed by the “treadmilling” process, which has to be tightly regulated to ensure a constant length to the mature microvillus. One level of regulation is reached through the free actin monomers (G-actin) availability; indeed, the G-actin/F-actin ratio remains constant in adult intestinal cells. However, during chick development, from day 1 to 3 after hatching, the nascent intestinal microvilli undergo a fast extension. This increase in length is concomitant to an increase in the G-actin/F-actin ratio, suggesting that an imbalance of the “treadmilling” process could drive the lengthening of microvilli (Stidwill and Burgess 1986). Actin biochemistry per se is therefore an important player in microvilli length regulation. Cellular domains made of microvilli exhibit an impressive homogeneity of organization that requires different actin-binding proteins to precisely control the shape of the domains. These factors possess different actin-related functions that regulate actin assembly or disassembly, F-actin cross-linking, and actin dynamics to allow the creation of accurate meshworks. Indeed, proteins that sequester actin monomers, sever existing actin filaments, or cap barbed ends to block actin polymerization, allow a precise control of the microfilaments’ length. In addition, the actin filaments can be crosslinked together by actin-bundling proteins to form stiff actin bundles. In microvilli, the complete regulation network is far from being completely elucidated; we will thus attempt to describe the different factors identified so far, which are involved in the control of microvillar shape.

Regulation of Actin Polymerization at the Tips of Intestinal Microvilli At the microvillus tip, where actin polymerizes, actin assembly is subject to major modulation by various actin-binding proteins that regulate the addition of monomers. Actincapping proteins seem to play a prominent role in the microvilli length regulation by blocking actin polymerization at the barbed ends. CapZ, identified in skeletal muscle, has been detected at the tips of intestinal microvilli (see Fig. 13.1A) (Schafer et al. 1992). Recently, the capping protein eps8 has been identified at the tips of intestinal microvilli in Caenorhabditis elegans (Croce et al. 2004). In absence of eps8, worms present irregular and longer microvilli, indicating a lack of proper termination of actin filament elongation. On the other hand, eps8 knockout mice present regular and shorter microvilli, a difference that likely reflects the presence of additional mechanisms involved in microvillar length regulation in mammals. Additionally, nutritional defects have been reported for these mice, although no clear explanation is provided for this observation (Tocchetti et al. 2010). The contribution of myosin motors to transport or anchor these tip-restricted factors has been described in stereocilia (Boeda et al. 2002; Belyantseva et al. 2005). No such

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data are available for microvilli; however, similar mechanisms likely occur at the microvillus tip where myosin V has been detected (Heintzelman et al. 1994).

Regulation of Actin Polymerization at the Tips of Stereocilia In stereocilia, recently published data furthermore stress the role of proteins regulating actin assembly at the tips of microvilli-like structures and provide accurate details on the mechanisms involved in microvillar length regulation. The “staircase” organization of stereocilia is of particular interest because it implies a differential regulation of the actin bundles length. A tip concentration of twinfilin-2, a barbed-end capping protein, has been reported for the shorter stereocilia row only (Peng et al. 2009). Interaction of twinfilin-2 with myosin IIIA is essential to its proper localization. The precise function of this interaction is still unclear, but a role in twinfilin-2 transport and/or anchorage at the tip is very likely (Rzadzinska et al. 2009). Functionally, in a renal cell line, twinfilin-2 transfection induces a restriction in microvilli length (Peng et al. 2009). A second complex composed of myosin XVa and whirlin has been identified at the tips of stereocilia, independently of their length (Rzadzinska et al. 2009). The function of myosin XVa and whirlin remains obscure; however, lack of either one or the other components of this complex results in the formation of very short stereocilia without disturbing the staircase organization (Probst et al. 1998; Mburu et al. 2003), suggesting a positive role of this complex in actin polymerization. In summary, the myosin XVa/whirlin complex promotes actin assembly, whereas the myosin IIIa/twinfilin-2 complex inhibits actin assembly (see Fig. 13.1A). The interplay between these determinants could determine the length gradation of the stereocilia staircase organization (Rzadzinska et al. 2009). Such regulatory complexes are probably also present at the tips of microvilli and their identification is of prime importance in order to understand better the regulation of these structures. One potential candidate would be a member of the formin protein family, which has been shown to be present at the tips of filopodia, which are migratory structures related to microvilli. At the filopodia tips, formin could nucleate actin to form unbranched microfilaments and protect the resulting barbed ends from capping proteins to increase F-actin length (Chhabra and Higgs 2007).

Microvilli F-Actin Parallel Bundles: United We Stand F-actin parallel bundles function as frameworks to support and stabilize membrane protrusions and thus determine their dimensions. A set of actin-bundling proteins that crosslink actin filaments ensures the straightness and stiffness of the actin framework that allows and maintains the membrane protrusion (Fath and Burgess 1995; Bartles 2000). These proteins exhibit multiple actin-binding motifs and, for some of them, present specific domains regulating their activity in correlation to external stimuli, for example, calcium concentration. The parallel bundles of intestinal microvilli are organized by three actin-bundling proteins: villin, plastin-1, and espin (see Fig. 13.1C) (Bartles 2000). Villin expression is restricted to the kidney proximal tubule and intestinal epithelial cells where it concentrates in the brush border and represents the more abundant microvillar component besides actin (Maunoury et al. 1988). Villin bundles actin filaments together through its homodimerization (George et al. 2007). In addition to its bundling activity, villin exhibits actin-severing activity, actin nucleation, and capping activities; all of these activities are regulated by external calcium concentration (Glenney et al. 1981b). Plastin-1 is a member of the plastin family. This isoform is found in intestinal/kidney microvilli and in stereocilia (Bretscher and Weber 1980; Tilney et al. 1992). Plastin proteins bundle actin filaments together, through two actin-binding sites, in a calcium-dependent manner

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(Glenney et al. 1981a). Espin, the third actin bundler is present in stereocilia and intestinal microvilli (Zheng et al. 2000). Intestinal espin is an espin small splicing variant, based on the C-terminal sequence that contains two actin-binding motifs. The F-actin bundling activity of espin does not depend on calcium concentration (Bartles et al. 1998). During embryogenesis, the actin-bundling proteins accumulate in microvilli in a stepwise manner. Villin apically concentrates concomitantly with the apparition of the first poorly organized microvilli, followed by apical concentration of plastin-1 a few days later (Shibayama et al. 1987). Espin concentrates apically later in development, on a core bundle that is largely organized and thus could “lock” its organization (Bartles et al. 1998). In cell culture, overexpression of villin, espin, or the T isoform of plastin increases microvilli length (Friederich et al. 1989; Arpin et al. 1994; Loomis et al. 2003). The microvillar lengthening is likely attributable to the bundling activities of these proteins as transfection of the actin cross-linking motif of espin is sufficient to drive microvilli elongation (Loomis et al. 2003). Conversely, villin knockdown in cell culture causes the disruption of the brush border in a colonic cell line (Costa de Beauregard et al. 1995). Surprisingly, the intestinal microvilli of villin knockout mice do not present any morphological defect (Pinson et al. 1998), neither do the Jerker mice, which do not express any espin isoform (Zheng et al. 2000) (see Fig. 13.2). On the other hand, although plastin-1 knockout mice still develop an organized brush border, it contains shorter microvilli and presents a strong alteration of the underlying terminal web (Grimm-Gunter et al. 2009) (see Fig. 13.2). As

Villin –/–

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Figure 13.2. Impact of the absence of major microvillar proteins on the intestinal brush border morphogenesis assessed by knockout studies. Villin–/– and espin–/– microvilli do not present any morphological defect as compared with wild-type microvilli. Plastin 1–/– microvilli are shorter; note the absence of rootlets. In myosin-1a–/–, microvilli are irregular, shorter, and loosely packed. On ezrin–/– neonates, microvilli are strongly misorganized, shorter, and misoriented. Myosin1a–/– and ezrin–/– micrographs were very kindly provided by Dr. Mark Mooseker and Dr. Andrea McClatchey, respectively.

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none of the three knockout mice models exhibit strong defect of the intestinal brush border morphogenesis, some redundancy between these actin-bundling proteins must necessarily exist.

The Microvillar F-Actin Parallel Bundle Is Connected to the Plasma Membrane and to the Terminal Web Different proteins link the microvilli actin core to the plasma membrane or to the subapical terminal web and thus anchor the F-actin bundle to the cell. Within each microvillus, two proteins have been shown to connect the actin filaments to the plasma membrane. Myosin1a labels the distinctive regular “bridges” between the F-actin core and the membrane seen by electron microscopy (Howe and Mooseker 1983). This plus-end-directed myosin motor could stabilize the microvillus, probably through the development of membrane tension by the connection with the microvillus actin core (Mooseker and Cheney 1995; Nambiar et al. 2009). Indeed, myosin-1a knockout mice show poorly packed microvilli that present variable length, shorter diameter, and membrane extrusions (see Fig. 13.2). These defects have been linked to the loss of the lateral bridges and the associated membrane tension that would result in decreased membrane stiffness (Tyska et al. 2005). However, in cell culture, the great majority of the total brush border myosin-1a pool (80%) is mobile based on fluorescence recovery after photobleaching (FRAP) experiments. This result shows that most of the myosin-1a in microvilli does not function as a stable linker and highlights the myosin-1a dynamics in microvilli (Tyska and Mooseker 2002). Ezrin, a membrane-cytoskeleton linker connects the microvillar actin core to the plasma membrane in intestinal cells. Ezrin knockout mice die soon after birth; however, the altered physiological functions leading to death remain to be determined. Ezrin knockout neonates display intestinal cells with strong alterations of the brush border, which contains shorter, poorly organized, and misoriented microvilli (see Fig. 13.2). Moreover, these animals show a thicker and undulating terminal web suggesting that ezrin participates in its organization (Saotome et al. 2004). The terminal web consists of an extremely dense intertwining of actin microfilaments interconnected by proteins of the spectrin family and by myosin-IIa, which confers contractile properties to the brush border. Intermediate filaments (see also Chapter 16) underlie this acto-myosin meshwork and participate in the establishment of the enterocyte brush border as cytokeratin 8 knockout mice present around 20% of enterocytes with an extensive atrophy of their microvilli (Ameen et al. 2001). The cohesion of the apical pole could be further ensured by the interconnection between the microvillar F-actin bundles and the terminal web as suggested by the molecular interaction between plastin-1 and cytokeratin-19. The shorter microvilli observed in plastin-1 knockout animals (see Fig. 13.2) would therefore be the consequence of the loss of this stabilizing interaction (Grimm-Gunter et al. 2009).

INTESTINAL MICROVILLI FORM A DOMAIN SPECIALIZED IN ABSORPTION Brush borders on intestinal cells allow a dramatic cell-surface increase and thus endow the intestinal epithelium with an extensive absorptive capacity. Indeed, a broad variety of membrane-bound proteins, such as enzymes, transporters, and ion channels, are inserted in the plasma membrane of the intestinal microvilli (see also Chapter 21 for further discussion of polarized transport in epithelial cells).

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The Particular Lipid Composition of Microvillar Membranes Facilitates the Insertion of Digestive Enzymes The majority of digestive enzymes present on intestinal microvilli are glycosylated and lipid raft associated (Danielsen 1995). The microvillar localization of raft-associated enzymes is dependent on lipid microdomains, which have been proposed to function as a major sorting platform during apical exocytosis (Simons and Ikonen 1997). Lipid rafts have been operationally defined as detergent-insoluble membrane fractions at low temperature and called detergent-resistant membranes (DRMs) (Brown and Rose 1992) (see also Chapter 4 on DRMs). Interestingly, the intestinal brush border is particularly rich in lipid rafts as determined by this biochemical approach (Danielsen 1995). Lipid rafts can create morphologically distinct membrane microdomains such as caveolae (Chapter 3) (Anderson 1998). Such morphologically distinguishable microdomains are not found on microvilli. However, raft and nonraft domains appear heterogeneously distributed on microvillar extracts arguing in favor of the existence of stable lipid rafts in microvilli (Hansen et al. 2001). Unlike conventional lipid rafts, particularly enriched in sphingolipids and cholesterol, intestinal microvillar lipid rafts are cholesterol independent and mainly composed of glycolipids (Christiansen and Carlsen 1981; Hansen et al. 2001). Proper insertion of the digestive enzymes in the brush border is obviously essential for their physiological role. The type II integral membrane protein sucrase–isomaltase, which is involved in the terminal digestion of sucrose and starch, is probably the best described lipid-raft-associated microvillar enzyme. Congenital sucrase–isomaltase deficiency, a condition causing diarrhea and abdominal pain due to saccharide malabsorption, has been associated with mutations in the sucrase–isomaltase gene leading to an impairment of its apical targeting (Naim et al. 1988). Proper association between apical proteins and lipid rafts usually requires additional posttranslational modifications. Indeed, O-glycosylation of sucrase–isomaltase is essential for its association with intestinal lipid rafts, which could be mediated by a lectin, probably galectin-4, as it co-immunoprecipitates with sucrase– isomaltase (Alfalah et al. 1999; Danielsen and van Deurs 1997). Indeed, galectin-4 could act as a stabilizer/organizer of microvillar rafts by virtue of its ability to form lattices containing glycolipids and glycoproteins (Braccia et al. 2003) (see also Chapters 8 and 21 for further discussion of rafts and apical targeting). However, it is important to point out that other apical sorting mechanisms do exist as apical localization of other digestive enzymes, such as lactase-phlorizin hydrolase, are lipid raft independent (Danielsen 1995).

The Cytoskeleton Participates in the Retention of Absorptive Factors at the Membrane The microvillar cytoskeleton is involved in the retention of membrane-bound proteins at the plasma membrane. Indeed, myosin-1a, which connects the microvillar actin bundle to the membrane is essential for proper brush border localization of sucrase–isomaltase (Tyska et al. 2005). Moreover, transfection experiments with a nonfunctional myosin-1a lead to the redistribution of sucrase–isomaltase to the cytoplasm (Tyska and Mooseker 2004). These results suggest that myosin-1a, probably through its ability to link the microvillus actin core to the membrane, could stabilize sucrase–isomaltase molecules in lipid rafts and thus extend their lifetime in the brush border membrane. Another nonexclusive hypothesis would be that myosin-1a powers the apical movement of Golgi-derived vesicles containing sucrase–isomaltase (Jacob et al. 2003; Tyska and Mooseker 2004). It is also worth noticing the role of ezrin, a membrane-cytoskeleton linker, which is involved in the transport and retention at the microvillar membrane of different ion channels (Na-Pi cotransporter 2, Na+/H+ exchanger 3, or cystic fibrosis transductance regulator).

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Indeed, their retention depends on the formation of protein scaffolds mediated by adaptor proteins (such as the Na+/H+ exchanger regulatory factors), which are connected to the microvillar actin bundle by ezrin (Cha et al. 2006).

Glucose Transporter 2 (GLUT2) Targeting at the Membrane in Response to an Increase in Glucose Intake Brush border disaccharidases (sucrase–isomaltase and lactase-phlorizin hydrolase) are responsible for the enzymatic digestion of polysaccharides into monosaccharides that are transported inside the cells by glucose transporters. The apical glucose/sodium symporter SGLT1 is permanently present at the brush border and is associated with lipid microdomains in renal proximal tubular cells (Runembert et al. 2002). Following an increase in luminal monosaccharide availability, GLUT2 is specifically targeted from the terminal web to the brush border membrane to avoid SGLT1 saturation. Kellet et al. proposed that GLUT2 transport toward the brush border is regulated by an increase in intracellular Ca2+ concentration that could activate the terminal web-resident phosphokinase PKCβII, which in turn may promote GLUT2 microvillar insertion via acto-myosin contractions of the terminal web (Kellett et al. 2008). This dynamic process allows the enterocytes to adapt their absorptive capacity to changes in dietary intakes. The participation of specific plasma membrane lipid composition, cytoskeleton, and scaffolding proteins therefore allows the particular enrichment of intestinal microvilli in factors involved in absorption.

Microvilli Generate Vesicles in the Intestinal Lumen Besides increasing surface contact with the lumen, intestinal microvilli seem to further participate actively in the digestive process through the release of vesicles in the lumen. Indeed, McConnell and colleagues were able to characterize a population of unilamellar vesicles that shed from microvilli to be released in the intestinal lumen. Interestingly, myosin-1a knockout mice present striking perturbation of vesicle production, suggesting that myosin-1a could power the sliding of microvillar membrane along actin bundles leading to the vesicle shedding. These microvillar vesicles are enriched in lipid-raft-resident digestive enzymes and are oriented right side out, thus allowing the deployment of catalytic activity, provided by the digestive enzymes, in the lumen to facilitate nutrient processing. Moreover, the vesicles seem to be especially enriched in intestinal alkaline phosphatase (McConnell et al. 2009). The physiological role of this enzyme remains obscure; however, its involvement in the detoxification of bacterial lipopolysaccharide has been shown (Beumer et al. 2003). The luminally released vesicles may thus be important to minimize the proinflammatory impact of this prominent bacterial component. Secreted microvillar vesicles thus represent a newly described physiological function of microvilli that brings new insight into how the brush border dynamically participates in the digestive process. Moreover, it raises numerous questions on the regulation of these highly specialized structures and on their putative role concerning the small intestine protection.

PERSPECTIVES ON THE BRUSH BORDER DYNAMICS Intestinal Microvilli Display High Plasticity Microvilli appear to be very stable structures at steady state. However, following intestinal stresses from various origins, including mechanical stress (Tilney and Cardell 1970),

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(A)

WT

villin–/–

(B)

EPEC myosin-1a

espB

Figure 13.3. The brush border is subject to major remodeling following Ca2+ stress and enteropathogenic E. coli (EPEC) infection. (A) Following Ca2+ stress, microvilli are subject to deep morphological alteration. On wild-type (WT) animals, microvilli are subject to disruption following Ca2+ stress. Villin, probably via its severing activity (scissors), is a major actor of Ca2+-induced microvillar breakdown as its absence impairs microvilli disruption following Ca2+ stress. (B) Microvilli “attaching and effacing” by EPEC. Following EPEC attachment to the brush border, microvilli remodeling leads to the formation of “pedestals,” allowing further infection. Microvilli remodeling involves espB, a factor secreted by EPEC, which binds to the actin-binding motif of myosin-1a leading to the disruption of the actin–myosin interaction.

calcium stress (Lange et al. 1997; Ferrary et al. 1999), or during cell migration (Nusrat et al. 1992), microvilli are subject to deep alterations and even to complete disruption. Microvilli are thus effectors of the epithelial plasticity; they can adapt their molecular composition and overall organization when exposed to changes in the intestinal microenvironment. Villin seems to be involved in the microvillar morphology adaptation following calcium stress, and its role has been nicely documented. Indeed, villin knockout mice do not show microvillar morphology breakdown following various Ca2+ stresses (see Fig. 13.3A). Moreover, these animals present a reduced ability to recover from chemically induced intestinal wounds, suggesting that villin, likely through its severing activity, is a key player in actin dynamics and a major factor of actin cytoskeleton reorganization and motility (Ferrary et al. 1999). During epithelium restitution that follows injury, cells undergo an epithelial-to-mesenchymal transition to acquire a motile fibroblastic-like morphology. Villin, through its severing activity, would thus be involved in the microvilli breakdown preceding intestinal cell migration. This hypothesis is strengthened by the fact that ectopic expression of villin increases migratory capacity of epithelial cells through

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villin phosphorylation that leads to an enhancement of its severing activity (Tomar et al. 2004). Moreover, this process would increase the concentration of actin monomers available for the formation of migratory structures.

Microvilli and Pathogens Protection Ensured by the Intestinal Epithelium The brush border of intestinal cells presents a huge surface area in contact with the outside world and thus constitutes a potential threat due to external pathogens. In this part, we will focus on how the intestinal brush border creates a barrier against pathogens and how pathogens can still penetrate it. A thick (up to several hundred micrometers) mucus coating on microvilli creates a stable gel layer acting as a physical barrier. Mucus is, in majority, composed of heavily glycosylated mucins secreted by the goblet cells. Besides providing a physical barrier, mucins are of particular importance for epithelial integrity as they harbor oligosaccharide chains, which are ligands for numerous bacteria and thus form a meshwork that can “trap” pathogens (Thornton and Sheehan 2004). Another level of protection is provided by the overall organization of the brush border, made of tightly packed microvilli that hinder close contacts of the microorganisms with the cell body. Pathogens that get in close contact with the microvillus cannot be endocytosed as the stiff microvillus actin bundle prevents membrane invagination. Microvilli interdigitations at their base would therefore be the only place where endocytosis could occur (Hansen et al. 2009). Enteropathogenic Escherichia coli (EPEC) and Shigella flexneri Exploits Actin Dynamics to Bypass the Physical Barrier Pathogens have developed several strategies to subvert the host cytoskeletal components in order to bypass mucosal protection. One of the major causes of infantile diarrhea worldwide is the infection by EPEC. Infection process involves the attachment of E. coli to the intestinal epithelium followed by an important cytoskeletal rearrangement of the brush border, leading to the destruction of microvilli and formation of membrane “pedestals” (see Fig. 13.3B) (Rosenshine et al. 1996). After EPEC attachment, the type III secretion system allows the secretion, into microvilli, of factors responsible for the subsequent infection. EspB, one of these effectors is involved in the microvillar effacement. EspB binds to the actin-binding sites of several myosins including myosin II and myosin I, and therefore inhibits the actin–myosin interaction due to competition (see Fig. 13.3B). Interestingly, EPEC mutants lacking the myosin-binding region on espB do not fail to attach at the cell membrane but microvilli destruction and further infection is impaired (Iizumi et al. 2007). This result therefore suggests that loss of interaction between myosin and actin is involved in microvillar effacement, which would not be required for pathogen attachment but rather for subsequent infection. Moreover, villin participation in the microvillar effacement through cytoskeletal rearrangements driven by its actin severing activity has been proposed but never demonstrated (Goosney et al. 1999). Further bacterial activity and spreading inside the intestinal epithelium has been nicely described for another pathogen, S. flexneri, which subverts the host cytoskeletal components for its propagation. S. flexneri is an enteropathogen causing dysentery, a worldwide major health issue. S. flexneri is believed to invade intestinal epithelium through M cells, a particular intestinal cell type that presents “depressed” microvilli (Fujimura et al. 1990; Sansonetti et al. 1996). However, it is not known whether the brush border alteration on this cell type is important for the bacterial uptake. Once internalized, S. flexneri is propelled through an actin comet and propagates into adjacent enterocytes (Bernardini et al. 1989). S. flexneri recruits

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cytoskeletal components of the host cell in order to form its actin comet. Villin is recruited at the S. flexneri actin comet where its severing and capping activities enhance actin dynamics and therefore provide higher propelling force (Revenu et al. 2007). On this aspect, a dramatic reduction of infected enterocytes are detected in villin knockout mice demonstrating that S. flexneri exploits the ability of villin to restructure the actin cytoskeleton in order to enter and spread throughout the epithelium (Athman et al. 2005).

Pathologies Associated with Microvilli Defects Several pathologies involving microvilli disruption have been described. In particular, strong defects of the intestinal brush border have been reported in intestinal biopsies of patients suffering from ulcerative colitis or celiac disease, a condition causing gluten intolerance. However, these conditions involve a massive inflammatory response; the microvillar defects observed might thus represent a consequence of the inflammatory response. Conversely, microvillus inclusion disease, which is a rare genetic disorder of the small intestine leading to severe microvillar atrophy and requires lifelong parental nutrition or intestinal transplantation, is directly linked to microvilli breakdown. Besides a partial or complete atrophy of apical microvilli, large cytoplasmic vesicles containing microvilli radiating from the inner face of the vesicles toward the vesicle lumen are observed and called microvillar inclusion bodies (Cutz et al. 1989). The molecular bases of the disorder remain unclear and contradictory, although a defective apical trafficking pathway has been reported (Ameen and Salas 2000). A general failure in protein apical insertion could thus impair microvillar biogenesis. In a recent work, an alternative hypothesis was proposed, as the authors were not able to detect any defect in apical trafficking. They proposed that microvillar inclusion bodies could rather result from autophagocytosis of the apical membrane (Reinshagen et al. 2002). Genetic association studies allowed the identification of mutations on the myosin Vb gene in patients suffering from microvillus inclusion disease, highlighting a critical role for this myosin in microvilli maintenance, although its precise role remains unknown (Muller et al. 2008). Additionally, a strong reduction of rab8 expression, a GTPase essential for trafficking, has been determined on enterocytes from a microvillus inclusion disease-suffering patient albeit no mutation in the rab8 gene has been detected. Interestingly, rab8 knockout mice present severe microvilli atrophy and large cytoplasmic vesicles containing microvilli, phenotypes similar in that to the ones observed in patients suffering from microvillus inclusion disease (Sato et al. 2007). Research on microvillus inclusion disease thus appears to be of primary interest as it provides new insights into microvilli biogenesis and maintenance.

GENERAL CONCLUSIONS Despite their obvious function to expand surface contact with the external milieu, microvilli are specialized cellular structures whose study has addressed some fundamental questions in biology. The rate of growth and the length of each individual microvilli are highly dependent on the regulation of actin microfilaments dynamics by a wide variety of actin-binding proteins. Hence, the detailed analysis of the molecular mechanisms that regulate the shape, length, and dynamics of microvilli provides pertinent models to unravel key questions such as how cells measure and count. Progress in mouse genetics and the possibility to knockout gene expression in tissue and temporal specific manner during development or during adult life has permitted to decipher some functions of key microvilli regulators in an in vivo context. The information gathered over the last decade, as illus-

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trated in this chapter, has led the authors to conclude that the functions of microvilli cannot be determined by the simple analysis of the knockout of one single gene. To our surprise, the invalidation of a single gene such as villin and others later on, demonstrated that the lack of one actin-binding protein does not perturb dramatically the organization and therefore function of microvilli (see Fig. 13.2). However, invalidation of a single gene may have dramatic effects on the homeostasis of the tissue when particular stressful conditions are applied. Thus, such an experimental approach can reveal the importance of a given gene in the dynamics of microvilli and its role in nonphysiological or pathological states. As a result, future studies will have to take into account these parameters to understand the involvement of gene networks in physiopathological processes and their contributions to acquired or inherited diseases. Beyond the structural aspect, the study of microvillar structures has stressed the importance of cell signaling integration in such specialized domains. For instance, the role of actin microfilaments as a platform to recruit signaling molecules is now well documented (see also Chapter 1). Indeed, actin microfilaments shaping microvilli can provide localized binding sites. The concentration of signaling complexes in a particular cellular domain contributes to the dynamics and specificity of signal transduction mechanisms during either physiological or physiopathological processes. The composition of the proteins associated with microvilli and stereocilia has now reached an advanced stage (see Fig. 13.1). However, it remains to perform further functional studies and analysis to integrate those molecular machineries at the cell level and, later on, at the tissue level to better understand the highly dynamic processes occurring both during embryonic development and throughout adult life.

ACKNOWLEDGMENTS This work was supported by funds from Curie Institute Foundation and Centre National de la Recherche Scientifique. F.U. is supported by a

PhD fellowship from the Ministère de l’Enseignement Supérieur et de la Recherche.

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CH A P T E R

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MICROTUBULES Geoffrey O. Wasteneys Bettina Lechner

DEFINITION Microtubules are essential eukaryotic structures with central roles in many cellular processes. Recognizable as hollow cylinders 25 nm in diameter but several micrometers in length, microtubules are the largest of the cytoskeletal filaments found in eukaryotic cells and also the most rigid. They are often bundled into tracks or cross-linked into complex cellular machinery that controls cell shape, cell motility, chromosome and organelle movement, and, when modified structurally into axonemal structures, the beating of flagella and cilia. These tracks and machines undergo rapid remodeling to coordinate pivotal events in the cell cycle or in response to extracellular cues. Rapid remodeling can occur because microtubules are composed of heterodimers of globular subunit proteins, called α- and β-tubulins. The key to microtubule assembly and disassembly is that their globular tubulin subunits are GTPases, with the process of GTP hydrolysis enabling eventual polymer disassembly. The inherent polarity of the heterodimeric tubulin subunits is repeated throughout the length of the microtubule, resulting in a structural polarity that is reflected in the different assembly properties at each end of the microtubule. Thirteen (generally) protofilaments in turn interact laterally to form a hollow cylinder approximately 25 nm in diameter. One consequence of the polar arrangement of tubulins in microtubules is that subunit exchange occurs much more rapidly at the β-tubulin-exposed end of the microtubule. Consequently, microtubules tend to grow and shrink predominantly from this end, which is termed the plus end. In contrast, the α-tubulin-exposed minus end undergoes subunit exchange at a relatively slow rate and frequently not at all when capped by microtubule nucleating complexes, which also often serve to orient microtubule arrays in cells. In addition to these tubulin building blocks, some highly conserved microtubule-associated proteins (MAPs) are essential for catalyzing microtubule polymerization and depolymerization at physiologically relevant rates. Other structural MAPs play more specialized roles in cross-linking microtubules into higher order structures such as mitotic spindles, phragmoplasts, axonemes, and centrioles. Some microtubule-specific motor proteins are utilized to move cargo along microtubule tracks, while others are specialized to utilize their ATPase activity to build, modify, and bend cellular machines.

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HISTORICAL PERSPECTIVE Microtubules are such an obvious cellular domain, yet it was not until the development of glutaraldehyde fixation for the preparation of cells for transmission electron microscopy that the true tubular nature of these structures was finally revealed. The term microtubule came into broad usage after two seminal publications in 1963 documented 180-Å diameter “microtubules” in hydra (Slautterback 1963) and in 250-Å diameter “microtubules” in the cortex of plant cells (Ledbetter and Porter 1963). Although tubular elements had been reported previously that in retrospect were probably microtubules, these two articles in 1963 made a major conceptual leap forward, not only in demonstrating the ubiquity of microtubules in eukaryotic organisms but also in recognizing that these cytoplasmic tubules were structurally identical to mitotic spindle fibers, to components of centrioles, and to previously described tubules from ciliated and flagellated protozoa (Dustin 1978). In 1964, Ledbetter and Porter published images depicting the 13 protofilament substructure of microtubules, described as “slender filamentous subunits” from the root tips of two diverse plant species (Ledbetter and Porter 1964). Their measurements indicated that these circular-profiled subunits formed 70-Å thick walls, which would appear as a halo around a 180-Å core when negatively stained, a finding that helps to explain the original 70-Å disparity between microtubule diameters in hydra described by Slautterback (1963) and in the plant cell cortex by Ledbetter and Porter (1963). Ledbetter and Porter also noted the similarity between the cytoplasmic microtubules and the fibrils of the mitotic spindle as well as the filaments comprising the 9 + 2 complex (Ledbetter and Porter 1964) previously described in cilia and flagella (Fawcett and Porter 1954). A more serious disparity between Slautterback’s work and that of Ledbetter and Porter was in their initial interpretation of the nature of microtubules. Slautterback, perhaps influenced by Palay’s interpretation that neurotubules observed in dendrites and axons were smooth endoplasmic reticulum (Palay 1956), but also because of their association with Golgi bodies in nematocysts, assumed microtubules to be composed of phospholipid– protein membranes and speculated that they would be involved in water or electrolyte transport in ion-secreting cells (Slautterback 1963). Ledbetter and Porter were more on the mark in their prediction that microtubules comprised protein and played some function in tension/contraction generation. In 1962, it was already fairly clear that spindle fibers were proteinaceous elements (Roth and Daniels 1962). In that same year, an insightful prediction was made by Paul Green (Green 1962). He noted that the drug colchicine, already known to dissolve spindle fibers, also caused growing plant and algal cells to swell rather than elongate, and from this he postulated that proteins of a spindle fiber nature would be active in the control of cellulose deposition at the surface of plant cells. By verifying the existence of elements in the plant cortex that not only resembled mitotic spindle fibers but also were oriented in the same direction as cellulose microfibrils in the cell wall, Ledbetter and Porter (1963) opened the way for exploring the mechanisms that govern mitosis in all eukaryotic cells, and the determination of cell shape in the plant kingdom. The drug colchicine, an autumn crocus-derived alkaloid that has been used for centuries in the treatment of the inflammatory disease gout, played an important role in the history of microtubule biology. The importance of spindle fibers in tethering and controlling the separation of chromosomes was obvious to early cytologists, but the composition of these fibers remained a mystery. The ability of spindle fibers to refract polarized light (birefringence) was described as early as 1937 (Schmidt 1937), suggesting that spindle fibers were structurally highly ordered. The loss of birefringence when cells

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were subjected to high hydrostatic pressure indicated that spindle fiber assembly was a highly dynamic process (Marsland 1938). Colchicine, whose antimitotic effects had first been documented in the late nineteenth century (Pernice 1889), was also found to dissolve mitotic spindles and to arrest mitosis in metaphase (Dustin 1938). This so-called stathmokinetic effect, which in the absence of cytokinesis leads to the amplification of chromosome numbers, was exploited both by plant breeders for the production of larger, more productive polyploid crop plants, and by cytogeneticists for the counting of chromosomes. In the 1960s, Taylor and his colleagues used tritiated colchicine to identify colchicine-binding proteins. They observed a strong correlation between the abundance of microtubules in various cell types with the reversible binding of colchicine, and this led Taylor to speculate that colchicine was binding to a structural protein of the spindle (Taylor 1965). This led to the purification under nondenaturing conditions of a 120-kDa colchicine-binding protein (Borisy and Taylor 1967; Shelanski and Taylor 1967) that later was determined to be a dimer of two very similar subunits (Shelanski and Taylor 1968). By 1968, the name tubulin was adopted for colchicine-binding protein (Adelman et al. 1968; Mohri 1968). This noble flurry of activity by a few researchers in the early 1960s led to the recognition that diverse cellular structures were built from microtubules and that the basic building blocks for microtubules were globular proteins. It has been followed by waves of discovery that corresponded to the advent, refinement, and uptake of new technologies. Electron microscopy remained the dominant technique for microtubule research until the late 1970s, when immunofluorescence microscopy made it possible to observe entire cellular arrays of microtubules in a wide variety of cells in many organisms. Another decade saw the introduction of live cell imaging, first by the microinjection of fluorescently tagged brain tubulin, which was quickly supplanted by genetically engineered fluorescent fusion proteins, such as green fluorescent protein (GFP)-tagged tubulins or MAPs that now enable researchers to follow the construction and dismantling of microtubule machines throughout the cell cycle. In parallel, the capacity to acquire and analyze images has benefited from the evolution of microscopes from simple epi-fluorescence through to the point scanning and spinning disk confocal systems in common use today.

MOLECULAR COMPOSITION OF MICROTUBULES, STRUCTURAL PROPERTIES, AND ASSEMBLY OF MICROTUBULES Unlike many cellular domains, which often persist for the lifetime of a cell or longer, microtubules are highly labile (see Fig. 14.1). This is an important property, given their tendency to rapidly reorganize into distinct arrays as cells progress through the cell cycle or to remodel in response to developmental or environmental cues. Tubulins are GTPases, proteins that commonly function as signaling switches because of the conformational changes that occur when the bound GTP undergoes hydrolysis. Although both α- and βtubulins bind GTP, the α-tubulin-bound GTP is occluded by dimer formation and remains unhydrolyzed. In contrast, hydrolysis of the β-tubulin-bound GTP takes place soon after polymerization. This hydrolysis, thought to be activated by the docking of an incoming α-tubulin to the outermost β-tubulin, causes the tubulin dimer to undergo a conformational change that effectively spring loads the linear protofilaments that make up the microtubule for rapid disassembly when required. Tubulin heterodimers assemble into linear protofilaments in a strictly αβαβ order, giving the microtubule an inherent structural polarity.

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Figure 14.1. Microtubules are dynamic structures. (A) Once nucleated, each microtubule can undergo several cycles of growth and shrinkage with periods of pause in between. GTP tubulin is indicated in brighter green. The GTP cap prevents the microtubule from depolymerization. (B) Various proteins (shown in orange) can bind to the microtubule or free tubulin and alter microtubule dynamics, cross-link adjacent microtubules or use the filaments as tracks for intracellular transport. (For details see Table 14.1 and text.)

Tubulins Comprise a Multigene Family of Proteins All eukaryotic organisms require at least one α- and one β-tubulin-encoding gene but multiple copies are common, ranging from 1 to 2 of each type of tubulin gene in some unicellular taxa to more than 6 of each in complex multicellular taxa such as vertebrates (Wade 2009) and vascular plants (Kopczak et al. 1992; Snustad et al. 1992). The proteins encoded by these different genes vary mainly in the negatively charged C-terminal domain of the protein, the so-called E-hook, and generally show 60% conservation among species. When multiple gene copies exist, they can either be coexpressed at one stage of development to generate isotypes of divergent function or may function in specific developmental modules or in response to different environmental conditions. The γ-tubulin subfamily proteins neither dimerize with α- or β-tubulins nor do they contribute to the main structure of microtubules. They are instead involved in microtubule nucleation and stabilization. They have 30% identity with α- and β-tubulins and are usually located in microtubule organizing centers (MTOCs) or in dispersed nucleating complexes. Additional tubulin subfamilies have been identified but they are not common to all eukaryotes. Tubulins such as δ-, ε-, ζ-, and η-tubulins appear to be involved in the duplication of centrioles and basal bodies (Wade 2009). Prokaryotes were previously thought to lack a cytoskeleton. Surprisingly, two tubulin orthologs, FtsZ of most prokaryotes and the plastid symbionts of plant cells, and the BtubA and BtubB proteins of Prosthecobacter dejongeii, have been identified in recent years. Together with eukaryotic tubulins, these proteins form a distinct family of GTPases. The sequence identity between the bacterial proteins and tubulins is rather low: 17% for FtsZ

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and 35% for BtubA/B compared with eukaryotic tubulin, but the protein structural domains remain similar. FtsZ and tubulins are likely to have evolved from a common ancestor and to have diverged at an early stage of evolution, as indicated by phylogenetic analysis. In contrast, BtubA and BtubB are more similar to tubulins than to FtsZ, which suggests that they evolved as a result of horizontal gene transfer from a eukaryotic host (Shih and Rothfield 2006).

Microtubule Dynamics and Its Regulation by Associated Proteins Microtubules undergo cycles of rapid growth and shrinkage, sometimes interspersed with short pauses, in which there is no net gain or loss of subunits. This behavior, illustrated in Figure 14.1A, is called dynamic instability, a process first described in 1984 by Mitchison and Kirschner, who demonstrated that individual microtubules can still undergo bouts of growth while the overall polymer mass is declining or that they can undergo rapid shrinkage under conditions that generally promote growth (Mitchison and Kirschner 1984). The transition from growth to rapid shrinkage is known as catastrophe and from shrinkage to growth is termed rescue. Dynamic instability is dependent on the nucleotide bound to the E-site in β-tubulin. During polymerization at the microtubule plus end, the GTP at this site is hydrolyzed slowly to GDP. Because GTP hydrolysis is not activated until after the α subunit of an incoming tubulin dimer interacts with the exposed β subunits, a so-called cap of GTP-bound subunits is retained at the growing plus end as long as the rate of incoming subunits exceeds the rate of hydrolysis. The GTP cap in turn stabilizes the microtubule against catastrophe. GDP-bound tubulin has a curved shape and is thought to promote disassembly, once the GTP cap is lost. Microtubules have distinct polymerization rates at their two ends. The plus end, at which β-tubulin is exposed to the cytoplasm, grows 10 times faster than the α-tubulinexposed minus end. The dynamic behavior of microtubules is not only regulated by GTP hydrolysis rates, but also by MAPs. Among these MAPs, some of which are illustrated in Figure 14.1B and listed in Table 14.1, are nucleating, stabilizing, and destabilizing factors, as well as microtubule severing and motor proteins (Conde and Caceres 2009; Wade 2009). Initiation of new microtubules in vivo requires nucleating proteins, which also control the number, the polarity, the timing of assembly, and the subcellular location of microtubules (Table 14.1). Nucleation complexes are required because the available cytoplasmic tubulin concentrations are generally below the critical concentration required for assembly, which is defined as the concentration at which microtubule self-assembly occurs in vitro. Nucleation usually takes place at MTOCs, which vary greatly in different organisms (see below). Nevertheless, all MTOCs have one protein in common: γ-tubulin, which is a principal component of the γ-tubulin-ring-complex (γ-TuRC). This 2-MDa complex forms an open, ring-shaped template for correct microtubule assembly (Aldaz et al. 2005; Conde and Caceres 2009). In contrast to the nucleating properties of MTOCs, tubulin-sequestering proteins stimulate catastrophes and destabilize microtubules by altering the pool of tubulin subunits. Stathmin, also called oncoprotein 18, binds two tubulin dimers and sequesters tubulins from the available pool to render tubulin assembly incompetent without reducing the overall stores of tubulin in cells. Stathmin’s binding to tubulin is an important mechanism to stimulate microtubule depolymerization, and its affinity for tubulin is in turn inhibited by phosphorylation (Manna et al. 2009). Stathmin plays a major role in cell– cycle progression and is highly expressed in many types of cancer cells (Rana et al. 2008; Wade 2009).

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MICROTUBULES

Categories of Common Microtubule-Associated Proteins

Protein family or group

Group

Example

Function

Nucleation

γ-TuRC

γ-Tubulin

Initiation of microtubule assembly

Destabilizing

Stathmin Microtubulesevering AAA ATPase

Stathmin Katanin

Sequesters tubulin

Spastin MCAK (Kinesin13)

Bends tubulin at plus end of microtubule, internal motor domain

MAP2 MAP4 Tau

In neurons, dendrites Other tissues In neurons, axons

EB1

EB1

XMAP215 CLIP

TOGp, MOR1 CLIP-170

CLASP

CLASP1

Sheet closure, tracks to plus end and recruits other +TIPs Promotes dynamics Interaction with dynein–dynactin, vesicles, and kinetochores Associates with CLIP, interaction with Golgi and cell cortex

Kinesin

KHC (Kinesin-1)

Kinesin Stabilizing

+TIPS

Transport

MAP2/Tau

Microtubule stabilization and bundling

Dynein

Cytoplasmic dynein

Plus-end directed, processive, N-terminal motor Minus-end directed, nonprocessive, C-terminal motore Minus-end directed transport, processive

Dam1-Ring TAC

Dam1 STIM1

Attachment to kinetochore Elongation of ER tubes by microtubule growth

Ncd (Kinesin-14)

Attachment

Generates internal breaks in the microtubule lattice

Other microtubule-destabilizing factors are microtubule-severing proteins, such as katanin and spastin, members of the AAA ATPase (ATPase associated with various cellular activities) protein family (Roll-Mecak and McNally 2010; Table 14.1). These microtubulestimulated ATPases sever microtubules by generating internal breaks. Both form hexameric rings only in the presence of ATP and are monomeric when ADP-bound (Roll-Mecak and McNally 2010). Katanin has two subunits: the p60 (60 kDa) subunit, which binds and severs microtubules, and the p80 subunit, which targets the complex to the centrosome. Katanin can release microtubules from the γ-TuRC, which are then transported as a short polymer, for example, into neurites (Conde and Caceres 2009). Spastin is the most commonly mutated gene in hereditary spastic paraplegia (HSP), a neurodegenerative disease (see Section on Microtubules and Human Disease). It is thought to target and partially unfold tubulin by binding the tubulin C-terminus through its positively charged ring pore. This will destabilize tubulin–tubulin interactions within the microtubule lattice and lead to microtubule breakdown (Salinas et al. 2007). After initiation of polymerization, structural MAPs such as members of the MAP2/ Tau family are known to stabilize microtubules, by binding along the microtubule lattice and alternating their dynamic behavior (Dehmelt and Halpain 2005). Unlike severing or motor proteins (see below), these are not enzymatically active. In vertebrates, all three

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members, MAP2, MAP4, and Tau, have alternative splice isoforms. Common features are the conserved C-terminal domain with microtubule-binding repeats and N-terminal projection domains of varying sizes. The size of the projection domain appears to determine the spacing between adjacent microtubules organized in bundles. The binding of these MAPs to microtubules can be regulated by phosphorylation and can inhibit transport along the microtubules. MAP2 and Tau are mainly expressed in neurons, whereas MAP4 is abundant in other tissues. These structural MAPs are important for neuritic differentiation and maintenance. MAP2 is mostly found in dendrites, where microtubules have mixed orientation. Tau is enriched in axons, where microtubules are in uniform orientation. Its binding also protects microtubules from severing by katanin. Furthermore, Tau is implicated in Alzheimer ’s disease and other dementias (Conde and Caceres 2009). A diverse group of microtubule plus end-binding proteins are called plus-end tracking proteins (+TIPs; Slep and Vale 2007). The dominant +TIP families are microtubule end binding protein 1 (EB1), cytoplasmic linker protein of 170 kDa (CLIP-170), CLIPassociated protein (CLASP), and XMAP215. These families are ubiquitous among eukaryotes and regulate the dynamics of the fast-growing plus end of microtubules. EB1 is a small, highly conserved protein that is thought to promote sheet closure of a growing microtubule, and tracks processively the growing plus end. EB1 promotes assembly of microtubules with 13 protofilaments and binds to microtubules via its N-terminal calponin homology domain (CH-domain). It has a coiled–coil dimerization domain, which provides a binding surface for EB1-interacting proteins. The acidic tail is responsible for autoinhibition, by binding to the CH-domain (Wade 2009). Members of the XMAP215 family promote rapid assembly and disassembly and inhibit pause. The highly conserved XMAP215/Dis1 family is the only known group of MAPs with orthologs in all kingdoms (Gard et al. 2004). These proteins are essential for survival, are generally encoded by single-copy genes, and are in most cases extremely large (>200 kDa). Growth and shrinkage rates of microtubules can be accelerated 10-fold by the addition of XMAP215 in vitro (Brouhard et al. 2008) or can be dramatically reduced in vivo in temperature-sensitive loss of function mutants of MOR1 (Kawamura and Wasteneys 2008), the plant homolog of XMAP215 (see Fig. 14.2B) . The most conserved feature are the N-terminal TOG domains. Family members have up to five TOG domains, where each may be sufficient to bind tubulin (Gard et al. 2004).

Microtubules Serve as Tracks for Intracellular Transport One of the most important functions of microtubules is to serve as tracks for intracellular transport. Many membranous organelles, chromosomes, and protein complexes are transported along microtubules to their final destination. The driving force for this intracellular transport is provided by molecular motors, which convert chemical energy into mechanical work. The two molecular motors associated with the microtubules are kinesins and dyneins. Both have a globular motor domain, also referred to as the “head” domain, but they are structurally and evolutionarily unrelated classes of proteins. The catalytic head domain possesses two properties crucial for molecular motors: a site for ATP hydrolysis and a nucleotide-dependent binding site for the cytoskeletal track. In many cases, the globular head domain is connected to an extended stalk, which dimerizes via a coiled–coil structure to yield a double-headed molecular motor. Members of the Kinesin-1 family (formerly conventional kinesins) are involved in intracellular long-distance transport, and especially important in axons and dendrites (Hirokawa and Takemura 2004; Hirokawa and Takemura 2005). Each ATP hydrolysis event is coupled to an 8-nm step toward the plus end of the microtubule (Hua et al. 1997; Schnitzer and Block 1997). The characteristic processive

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(A)

(B)

MOR1

microtubules

merge; blue=DNA

Figure 14.2. The microtubule cytoskeleton is rearranged during the cell cycle. To visualize microtubules under a fluorescence microscope, two approaches can be used. Cells either express recombinant tubulin or a MAP fused to a fluorescent reporter protein, such as GFP, or fixed cells can be stained with antibodies against tubulin. (A) A hypocotyl cell from Arabidopsis thaliana expressing GFP tubulin during interphase. The cortical array of microtubules is located below the plasma membrane and encircles the whole cell (picture courtesy of J.C. Ambrose). (B) Antibody-stained root tip cells of Arabidopsis in preprophase. They are stained with antibodies against the XMAP215 homolog MOR1 (red) and tubulin (green). Nuclei are stained with DAPI (blue). Microtubules concentrate in the preprophase band, which determines the cell division plane. Scale Bar: 5 μm. (Images from Kawamura et al. 2006, http://www.plantphysiol.org, Copyright American Society of Plant Biologists).

hand-over-hand type motility requires coordinated activity of two coupled motor heads (Hancock and Howard 1998; Hackney et al. 2003). Cytoplasmic dynein also moves in a processive manner, but, in contrast to the Kinesin-1 family proteins, it moves toward the minus end of the microtubule. While dyneins are strictly minus end-directed, kinesins can be plus or minus end-directed, depending on whether the motor domain is at the N (plus end-directed) or C-terminal (minus end-directed). The Kinesin-13 class is unconventional in that they contain an internal motor domain, and rather than transporting cargo these kinesins utilize their ATPase activity to promote microtubule catastrophes. They bend tubulins at the microtubule ends and, as a consequence, initiate microtubule depolymerization. Kinesin-13s, such as MCAK or KIF2A, are dimeric proteins and move along the microtubule via one-dimensional diffusion. Once bound, they remain attached to the microtubule by electrostatic interaction with the negatively charged E-hook of tubulin and diffuse along the microtubule lattice ran-

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domly to reach either the plus or the minus end. This motion is based on diffusion and is not dependent on hydrolysis of ATP (Helenius et al. 2006). Kinesin-13s either deconstruct microtubules to allow rearrangement of the network or couple the depolymerization to cargo movement, for example, chromatid separation in the meiotic and mitotic spindle (Moores and Milligan 2006). A highly sophisticated form of intracellular transport is axonal transport (see also Chapter 22 on neuronal domains). In the human body, axons can be up to 1 m in length; numerous proteins, mRNA, or even whole organelles need to be brought from the neuronal cell body to the synapse. Insufficient supply to the synapse leads to neurodegeneration, resulting in severe neuronal diseases. Transport in axons occurs in anterograde and in retrograde direction, and has two major components of distinct velocity, termed fast and slow axonal transport (Lasek 1967). Anterograde transport toward the cell periphery, which can be either fast or slow, is driven by various kinesin motors, while fast retrograde transport is driven by cytoplasmic dynein. Anterograde transport is responsible for supplying the synapse with structural components, whereas retrograde transport collects metabolites and redundant membranes for recycling in the cell body and delivers chemical messages. It is thought that vesicular cargoes are mainly delivered by fast axonal transport, while cytoskeletal and cytosolic proteins are delivered by slow axonal transport.

FUNCTIONAL IMPLICATIONS OF MICROTUBULE SPATIAL ORGANIZATION Microtubules as Cellular Machines Interphase For organisms whose cells are in isotonic solution, microtubules can support cell shape as a result of their rigid nature as long and hollow tubes, which enable them to resist deforming forces. In cultured animal cells, the microtubule network extends in a radial pattern to generate a round and flattened cell shape. In contrast, microtubules are oriented parallel to the long axis in columnar epithelial cells. In plant cells, microtubules work in a completely different, indirect manner to maintain cell shape through their influence on the formation of the cell wall. Because of the enormous hydrostatic turgor pressures that exist in plant cells, microtubules do not contribute to the actual forces that drive expansion (Wasteneys and Ambrose 2009). Plant interphase microtubules are dispersed in a selforganizing array that is located just beneath the plasma membrane (see Fig. 14.2A). Recently, these cortical microtubules have been shown to mark plasma membrane domains in which enzyme complexes synthesize cellulose microfibrils. The orientation of cellulose microfibrils plays an important role in determining the growth characteristics and thus the shape of plant cells. Cellulose microfibrils and the matrix polysaccharides they interact with form a tensile network, like hoops of a barrel, to resist lateral expansion and to promote elongation. Consequently, the isotropic turgor pressure is able to drive cell elongation. The microtubule cytoskeleton is also a formidable force-generating apparatus that moves whole cells. Single-celled organisms or motile cells of multicellular organisms use cilia and flagella to move in aqueous solutions (see also Chapter 15 on cilia). Microtubules determine cellular polarity in part through the positioning of motorassociated organelles (see also Chapters 7 and 8 on the ER and Golgi, respectively). The Golgi apparatus of animal cells is kept close to the cell center through the activity of molecular motors and an intact microtubule array (Barr and Egerer 2005). When microtubules are disassembled, the Golgi becomes fragmented and dispersed throughout the

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cell. The distribution and organization of the ER also depends on motile and static interactions with the cytoskeleton. The ER is continually moving and changing shape. The movement is driven by motor proteins. ER tubules slide along microtubules toward the periphery in animal cells. The tubules interact with plus-end trackers via TAC (tip attachment complex). This complex contains EB1 on the microtubule side and the ER resident protein stromal interacting module 1 (STIM1). ER movement can be driven by either kinesin-1 or dynein. Static attachment of the ER through 63-kDa cytoskeleton-linking protein (CLIMP63) stabilizes the extended network in vertebrate cells (Bola and Allan 2009). In plant cells, most organelle transport is dependent on myosin for movement along actin filament bundle networks (Wasteneys and Galway 2003), and many organelle systems, such as Golgi, are highly dispersed rather than centrally organized (Boevink et al. 1998). It has been suggested that myosin, which moves organelles more quickly than microtubule motors, is well suited to the transport of organelles through the plant cytoplasm, which is typically vast and thinly spread around the cell periphery (Wasteneys 2002). Nuclear and Cell Division In all eukaryotes, chromosome segregation is accomplished by the mitotic spindle. This micro-machine is composed of microtubules arranged in a bipolar manner through rapid dis- and reassembly during the transition from interphase to mitosis. In many archaeae, algae, and fungi, the nuclear envelope remains intact during mitosis, and spindle pole bodies serve as microtubule organizing centers, through which microtubules can penetrate and be captured by kinetochores. Open mitosis in animal and plant cells involves the assembly of a spindle that usually surrounds the chromosomes prior to nuclear envelope breakdown to ensure efficient chromosome capture. In animal cells, astral microtubules radiate from each of two daughter centrosomes during early prophase. Centrosomes consist of two arrays of nine triplet microtubules oriented at right angles to each other, called centrioles, which are in turn embedded in pericentriolar material. Pericentriolar material includes factors that promote microtubule assembly such as gamma tubulin ring complexes. The separation of centrosomes from one another toward the opposite ends of the nucleus is driven by the combination of microtubule polymerization and the activity of motor proteins, which interconnect and drive apart the oppositely oriented microtubules from each centrosome. As the number and length of microtubules in the spindle increase, the centrosomes eventually form the poles of the bipolar spindle. In centrosome-free animal cells, such as oocytes, chromatin itself stimulates microtubule nucleation, resulting in a jumble of microtubules that, through the action of motor proteins, self-organize into bipolar spindle after nuclear envelope breakdown. Spindle formation in plant cells is also self–organized but, as with animal cells, occurs prior to nuclear envelope breakdown. Before prophase begins, cortical microtubules concentrate in a band that marks the eventual site of cell plate attachment (see Fig. 14.2B). Microtubules from this preprophase band also extend toward the nucleus and appear to provide a framework that assists polarity establishment of the prophase spindle, which assembles and self-organizes through kinesin motor protein activity (cytoplasmic dyneins are absent in plant cells). The diverse mechanisms used for spindle assembly in different eukaryotic kingdoms, and indeed in those animal cells that are centrosome-free, illustrate how centrosomes are in fact nonessential for spindle formation, and underscore the importance of self-organization mechanisms (Wasteneys 2002). Microtubule nucleation in flagellated unicellular organisms and multicellular organisms that possess flagella (e.g., animals) is most often (but not exclusively) concentrated around centrosomes. Centrosomes are absent in yeast cells, but spindle pole bodies define analogous central organizing centers. The lack of centriole-based organizing centers in plant cells seems to be correlated with the

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loss of flagellated sperm. In most seed-bearing plants, sperm cells are carried to the ovule inside specialized tip-growing pollen tubes. Plant cells and acentrosomal animal cells achieve accurate mitoses so it is obvious that centrosomes are not essential for spindle pole formation. Instead, it is instructive to consider that the purpose of centrosomes acting as spindle poles is to ensure their transmission to both daughter cells. This tight coupling of the centrosome and nuclear division cycles has been illustrated in classic experiments on sea urchin eggs (Lingle et al. 2005). If fertilized by two sperm or when cells in metaphase are briefly treated with mercaptoethanol (Paweletz et al. 1984), sea urchin eggs form four spindle poles, and four daughter cells are produced. In the subsequent cell cycle, the daughter cells mainly form monopolar spindles because the centrosome duplication cycle results in just one rather than two centriole pairs. Thus, confining microtubule spindle nucleation to the centriolar apparatus ensures that each daughter cell will inherit one centrosome. A spindle pole in mitosis will become a basal body during interphase, from which flagella will arise. An analogous situation exists in some nonvascular plants that have cells with one single large chloroplast. Since each daughter cell should receive a chloroplast in order to maintain autotrophy, microtubule organizing centers are associated with plastids and with spindle pole separation during anaphase. Once the spindle is built in prophase, chromosome separation follows similar mechanisms in all taxa. During prometaphase, microtubules are captured by the contact to a kinetochore and stabilized. Kinetochores are stably associated with the plus end of up to 20–30 microtubules (in mammals) and form spindle fibers. The unattached kinetochore captures a microtubule from the opposite pole. Subsequently, chromosomes are moved to the spindle equator, the metaphase plate. The spindle incorporates the following types of microtubules. (1) Astral microtubules reach outward to position the spindle and determine the plane of cytokinesis. (2) Chromosomal microtubules extend between kinetochores and centrosomes. They move chromosomes during anaphase. (3) Polar microtubules reach from the centrosome past the chromosomes and overlap with the ones from the other pole. They form a structural basket. In anaphase, the sister chromatids are split apart and move toward the poles. Poleward movement is driven by shortening of plus and minus ends, which is catalyzed by microtubule-depolymerizing kinesin-13s. In yeast, the Dam1 complex forms a ring around the microtubule plus end and is attached to the kinetochore (Howard and Hyman 2007). While microtubules depolymerize, the Dam1 ring and subsequently the chromosomes are pushed backward to the poles. Similar mechanisms probably exist in other organisms. During telophase, daughter cells return to the interphase condition. The mitotic spindle disassembles and, where open mitosis has occurred, the nuclear envelope reforms. Cytokinesis usually follows and varies considerably between plant and other kingdoms, but involvement of microtubules is common in all taxa (Otegui et al. 2005). In animal cells, cytokinesis begins with indentation of the cell by a cleavage furrow that deepens through myosin-dependent actin filament sliding that occurs at a site specified by the remnants of the spindle, which now forms the cytoplasmic bridge between the two cells called a midbody. Plant cells, because of their enormous turgor pressures, must instead construct an extracellular wall between the two daughter nuclei. Wall formation starts with the formation of a membranous cell plate that often forms in the cell center and expands outward to meet the existing lateral walls in a process that is dependent on bipolar microtubule arrays called phragmoplasts. Like the midbody of animal cells, phragmoplasts arise in late anaphase from remnants of the spindle. Their close association with Golgi-derived vesicles suggests a microtubule-dependent role in Golgi trafficking, which results in the expansion of the cell plate until it separates the cytoplasm into two daughter cells at the division plane that had been marked in G2 by the microtubule preprophase band.

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Microtubules in Agriculture and Medicine Chemical Inhibitors of Microtubules as Herbicides, Fungicides, and Drugs Microtubules are the primary target of many effective agents used in agriculture either as herbicides or fungicides. Many industrially synthesized chemicals, such as the dinitroaniline herbicide oryzalin and the phosphoric amide herbicide amiprophos-methyl, target tubulins (Murthy et al. 1994), and this has led to their use as effective herbicides that kill germinating weed seedlings before they can compete with perennial crops. One of the big attractions of these herbicides is that they have a high affinity for plant tubulin but virtually no affinity for vertebrate tubulin (Murthy et al. 1994), suggesting that they can be used as low-toxicity options in agricultural practice. Weeds species, however, readily develop resistance to oryzalin along with cross-resistance to the chemically unrelated amiprophos-methyl through single nucleotide substitutions in α-tubulin-encoding genes (Anthony et al. 1998; Anthony and Hussey 1999). Such setbacks to the agro industry, however, have had interesting benefits for understanding the nature of cytoskeletal dynamics. Like plants, many important protozoan parasites that cause human disease are susceptible to dinitroaniline herbicides. And just as in plants, these parasites can achieve resistance through single point mutations. By identifying numerous point mutations that lead to oryzalin resistance in Toxoplasma—a parasite of concern for fetal mortality and birth defects—Morrissette et al. (2004) mapped oryzalin’s binding site and determined that this compound specifically disrupts the contacts between M and N loops on α-tubulin, leading to microtubule disassembly. Whether these or related compounds will one day be useful therapeutic agents remains unclear, but the quest to understand the nature of oryzalin resistance in weed species and parasites has uncovered important information on the mechanism of microtubule polymer formation. It is reminiscent of the quest to identify colchicine-binding proteins four decades earlier that led to the discovery of tubulin (Borisy and Taylor 1967; Shelanski and Taylor 1968). Microtubule Dynamics and Human Disease Microtubule function has important implications in both the cause and treatment of many human diseases including cancer, neurological disorders, polycystic kidney disease, and infertility. Mutations in human spastin, a microtubule-severing protein, cause autosomaldominant forms of HSP. The disorder is characterized by progressive spastic weakness of the lower extremities. It is called “dying-back neuropathy” because synapses of the motoneurons degrade first (Casari and Rugarli 2001; Gould and Brady 2004). There are more than 30 HSP loci known, and some of these genes are clearly associated with microtubule functions. Another example is the neuronal kinesin-1. The first in vitro studies have shown that the mutations—located in the motor domain—reduce the gross cargo flux, leading to deficient supply to the synapse and therefore degradation (Ebbing et al. 2008). The importance of precise assembly dynamics is underlined by the effective anticancer properties of drugs such as taxol, which hyperstabilizes microtubules, and vinblastine, which causes microtubule disassembly (Hadfield et al. 2003). Unfortunately, microtubule dynamics are frequently altered in cancer cells, and this can reduce the effectiveness of therapies. For example, increased microtubule dynamics is associated with resistance of lung cancer cells to taxol (Goncalves et al. 2001). Understanding the complexity of microtubule dynamics is therefore an important goal for cancer research. The differential activity of cyclin-dependent kinases alters microtubule dynamic properties as cells progress through the cell cycle. These kinases generally affect microtubule dynamics by phosphorylating MAPs. A general role for MAPs in regulating the

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dynamic and structural properties of microtubule arrays has been known for decades, but we still know little about how these proteins work, and some MAPs have only recently been discovered. One cancer-critical gene is the colonic, hepatic tumor overexpressed gene (ch-TOG). This gene was initially cloned from a human brain tumor cDNA library and was subsequently demonstrated to be overexpressed in neoplastic liver and colon tissues (Charrasse et al. 1995). Ch-TOG, which is a homolog of XMAP215 and MOR1, occurs in humans as a single-copy gene (Charrasse et al. 1998). Much ongoing research into the development of anticancer agents, as well as the understanding of the nature of the disease itself, is focused on the regulation of microtubule dynamics. The sequestration properties of the protein stathmin, for example, are currently intensively being investigated (Rana et al. 2008).

Future Perspectives with Uncertainties and Controversies Posttranslational Modification of Tubulins and Interplay with Microtubule Associated Proteins The microtubule cytoskeleton is also influenced by diverse and reversible posttranslational modifications (PTMs), such as acetylation, polyglycylation, and polyglutamylation, tyrosination, and phosphorylation (Westermann and Weber 2003; Hammond et al. 2008). Most of these modifications affect the carboxy-terminal domains of α- and β-tubulins, which are located on the outside of the microtubule where it is well positioned to influence interactions with other proteins. An exception is acetylation, which takes place on lysine 40 of α-tubulin inside the tube. Most PTMs occur on microtubules rather than on unpolymerized tubulin, and it has long been known that stable microtubules, as compared with dynamic microtubules, accumulate more modifications. PTMs are thought to influence the binding of motor proteins to direct intracellular traffic in a certain direction or to mark the microtubule for certain events, such as severing by katanin or spastin. Furthermore, PTMs influence the higher order of microtubule assembly, such as into axonemes, centrioles, or basal bodies. Many enzymes that catalyze the PTMs are unknown and their influence on cellular function is poorly understood. Nevertheless, PTMs are known to influence microtubule stability and structure as well as the recruitment of MAPs. For many decades now, microtubules have been a major focus in several fields of biological research, and yet there remain many uncertainties about their basic biology. These uncertainties constrain the development of important applications in medicine and agriculture. As highlighted in this chapter, many applications-driven research projects, such as the quest to identify the target of colchicine, the development of taxol as an anticancer therapy, and the basis for the development of resistance in plants and human parasites to the herbicide oryzalin, have generated important insights into the basic mechanisms by which microtubules form and carry out their ubiquitous and essential functions in eukaryotic cells. Future research efforts to both understand the basic biology of microtubules and develop applications relevant to human endeavours will no doubt continue to yield insight into these cylindrical cellular compartments that have the magical ability to suddenly vanish and reappear in new places.

ABBREVIATIONS AAA ATPase ATPase associated with various cellular activities CH-domain calponin homology domain

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CLASP CLIMP63 CLIP-170

CLIP-associated protein 63-kDa cytoskeleton-linking protein cytoplasmic linker protein of 170 kDa

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MICROTUBULES

microtubule end binding protein 1 green fluorescent protein hereditary spastic paraplegia microtubule-associated proteins microtubule organizing centers

STIM1 TAC +TIPs γ-TuRC

stromal interacting molecule tip attachment complex plus-end tracking proteins γ-tubulin-ring-complex

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Pernice B. 1889. Sulla cariocinesi delle cellule epiteliali e dell’ endotelio dei vasi della mucosa dello stomaco et dell’ intestino, nelle studio della gastroenterite sperimentale (nell’avelenamento per colchico). Sicilia Med 1: 265–79. Rana S, Maples PB, Senzer N, Nemunaitis J. 2008. Stathmin 1: a novel therapeutic target for anticancer activity. Expert Rev Anticancer Ther 8:1461–70. Roll-Mecak A, McNally FJ. 2010. Microtubule-severing enzymes. Curr Opin Cell Biol 22:96–103. Roth LE, Daniels EW. 1962. Electron microscopic studies of mitosis in amebae. II. The giant ameba Pelomyxa carolinensis. J Cell Biol 12:57–78. Salinas S, Carazo-Salas RE, Proukakis C, Schiavo G, Warner TT. 2007. Spastin and microtubules: functions in health and disease. J Neurosci Res 85:2778–82. Schmidt WJ. 1937. Die Doppelbrechung von Karyoplasma, Zytoplasma und Metaplasma. Berlin: Gebr. Bernträger. Schnitzer MJ, Block SM. 1997. Kinesin hydrolyses one ATP per 8-nm step. Nature 388:386–90. Shelanski ML, Taylor EW. 1967. Isolation of a protein subunit from microtubules. J Cell Biol 34:549–54. Shelanski ML, Taylor EW. 1968. Properties of the protein subunit of central-pair and outer-doublet microtubules of sea urchin flagella. J Cell Biol 38:304–15. Shih YL, Rothfield L. 2006. The bacterial cytoskeleton. Microbiol Mol Biol Rev 70:729–54. Slautterback DB. 1963. Cytoplasmic Microtubules. I. Hydra. J Cell Biol 18:367–88. Slep KC, Vale RD. 2007. Structural basis of microtubule plus end tracking by XMAP215, CLIP-170, and EB1. Mol Cell 27:976–91. Snustad DP, Haas NA, Kopczak SD, Silflow CD. 1992. The small genome of Arabidopsis contains at least nine expressed beta-tubulin genes. Plant Cell 4:549–56. Taylor EW. 1965. The mechanism of colchicine inhibition of mitosis. I. Kinetics of inhibition and the binding of h3-colchicine. J Cell Biol 25(SUPPL):145–60. Wade RH. 2009. On and around microtubules: an overview. Mol Biotechnol 43:177–91. Wasteneys GO. 2002. Microtubule organization in the green kingdom: chaos or self-order? J Cell Sci 115: 1345–54. Wasteneys GO, Ambrose JC. 2009. Spatial organization of plant cortical microtubules: close encounters of the 2D kind. Trends Cell Biol 19:62–71. Wasteneys GO, Galway ME. 2003. Remodeling the cytoskeleton for growth and form: an overview with some new views. Annu Rev Plant Biol 54:691–722. Westermann S, Weber K. 2003. Post-translational modifications regulate microtubule function. Nat Rev Mol Cell Biol 4:938–47.

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CILIA Laura K. Hilton Lynne M. Quarmby

DEFINITION Cilia are membrane-covered, microtubule-based structures that project from the surface of cells. The earliest eukaryotic cells were ciliated, and most extant lineages retain cilia. The primordial cilium may have functioned both as a processing center for signal transduction and as a device of motility (Satir et al. 2008; Quarmby and Leroux 2010). Some cilia have retained both functions whereas others have become highly specialized. The fundamental structure of the cilium is conserved, both at the level of proteins and ultrastructurally. Cilia are organized into four defined zones: the basal body (BB), the transition zone (TZ), the cilium proper, and the tip (see Fig. 15.1). Each of these regions may be more or less elaborated in different cell types. The ciliary membrane is continuous with, but distinct from, the plasma membrane. There is selectivity in both the lipids and proteins that are directed to the ciliary membrane. The mechanisms driving this selectivity are not known and are active areas of investigation. It is likely that there are unique domains within the ciliary membrane, as has been shown for a calcium permeant channel in Chlamydomonas (Fujiu et al. 2009). Underlying the membrane, providing structural support and a scaffold for assembly and function, is the axoneme. The core of the canonical axoneme is formed by nine outer doublet microtubules that are continuous with the A and B microtubules of the ninefold BB ABC triplet. Many elaborations of the axoneme, including the dynein arms, the central pair, and the radial spokes, participate in the generation and regulation of motility. The term “flagella” is commonly used in reference to some cilia—such as sperm tails and the cilia of some protists, such as Chlamydomonas. Whereas in eukaryotes “cilia” and “flagella” are synonyms, the term “flagella” is also used to describe an entirely different structure found in prokaryotes. In contrast to the eukaryotic organelle, the prokaryotic flagellum is not membrane bound and is comprised of a rigid helical protein structure that rotates like a propeller. Its composition, assembly, and mode of motility are completely different from cilia; that both are called “flagella” is an accident of history. Prokaryotic flagella are not addressed further in this chapter.

HISTORICAL PERSPECTIVE Motile cilia were likely first observed in the late fifteenth century, with the development of microscopy. The motile function of these cilia would have been obvious as early naturalists watched the protists of pond water swim about. In the twentieth century, Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Figure 15.1. Anatomy of a cilium. (A) A ciliary basal body (BB; pink), transition zone (TZ; green), and axoneme (blue) are shown, with cross sections through each region shown on either side. Adapted from O’Toole et al. (2003). (1) A ring of amorphous material at the proximal end of the BB. (2) The proximal end of the BB has a “cartwheel” structure at its center with nine triplet microtubule (MT) “blades” surrounding it. The A, B, and C tubules are indicated. (3) Transition fibers that anchor the BB to the plasma membrane are shown as triangular projections from the blades. An electron tomography (ET) image of this section of the BB is shown adjacent (from O’Toole et al. 2003, with permission). (4) Transition fibers from the distal end of the BB continue through the proximal end of the TZ. (5) Stellate fibers within the MT ring of the TZ and the termination of the transition fibers in pores surrounding the TZ. An ET image of this section of the TZ is shown adjacent; asterisks indicate the pores in the surrounding ciliary membrane (from O’Toole et al. 2003, with permission). (6) The arrangement of nine outer doublet MTs and the central pair are shown, with the A and B tubules indicated. A transmission electron microscopy (TEM) image of an axoneme is shown adjacent, with central pair and outer doublet projections visible. Reprinted from Sakato and King (2004) with permission from Elsevier. (7) The recently identified proximal site of severing where the TZ severs from the BB before mitosis. (8) The site of flagellar autotomy (SOFA) where the axoneme severs during the deflagellation stress response. (B) A simplified representation of the 96-nm repeat pattern in which inner (IDA) and outer dynein arms (ODA) and radial spokes (RSs) are arrayed. (C) Cryo-ET image and volume rendering of 160 averaged sections through two adjacent outer doublet MTs. N, nexin link/DRC; IL, dynein tail complex. From Nicastro et al. 2006. Reprint with permission from AAAS.

researchers used primarily sea urchin sperm to begin teasing apart the molecular mechanisms of ciliary motility, revealing the microtubule core, the role of the ciliary dyneins, and the sliding model for bending (see section on “Motility”; Summers and Gibbons 1971; Brokaw 1972). The unicellular biflagellated green alga Chlamydomonas became the predominant model organism for studying cilia as the facile haploid genetics and ease of preparing large quantities of isolated flagella allowed the correlation of mutant motility (paralyzed, altered waveform) with genes, proteins, and ultrastructure (Witman et al. 1978; Omoto et al. 1996). Although known for more than a century, the nonmotile primary cilium has until relatively recently been more of an enigma. The primary cilium was largely neglected and thought by many to be vestigial. In 2000, that all changed when it was reported that a Chlamydomonas flagellar assembly (FLA) mutant carried a defect in a gene known to be associated with polycystic kidney disease (PKD) in mice (Pazour et al. 2000). This discovery revived interest in the primary cilium, and at this writing, the field of ciliary research is in an explosive phase.

MOLECULAR COMPOSITION Cilia are among the most diverse of subcellular organelles (see Fig. 15.2). In the human body, motile cilia drive fluid flow over epithelial surfaces in the respiratory tract, ventricles of the brain, and, for those of us who have them, fallopian tubes. The sperm tail, or flagellum, is also an elaborated motile cilium. The molecular machinery that detects light in our eyes is organized within the outer segments of our rod and cone cells—these outer segments are highly derived cilia. Similarly, long flaccid cilia in our olfactory tissues are the site of reception of odorants. In addition, almost every cell in the human body, including neurons, kidney, liver, and skin cells, expresses a tiny cilium known as the primary cilium. Similarly, distinctive cilia can be found in protists, eukaryotic gametes of all sorts, and in the model organisms that have contributed most to our understanding of cilia: sea urchin, Chlamydomonas, Caenorhabditis elegans, and zebra fish. In spite of this diversity, the fundamental components of cilia are highly conserved, as is the machinery for building a cilium. A “typical cilium” is comprised of more than

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(A)

(C)

(B)

(D)

(E)

Figure 15.2. Cilia are diverse. Scanning electron micrograph of chick (A) olfactory and (B) respiratory cilia. From Breipohl and Fernandez 1977, with kind permission from Springer Science+Business Media). (C) Scanning electron microscopy (SEM) of Chlamydomonas reinhardtii cells (Dartmouth EM facility). (D) SEM of mouse renal collecting duct cells with primary cilia on the apical surface (Pazour et al. 2000 © The Rockefeller University Press. The Journal of Cell Biology, 2000;151:709–718). (E) TEM of mouse rod photoreceptor cells demonstrating the connecting cilium between inner and outer segments. Inset: cross section of the connecting cilium axoneme just above the BB. From Watanabe et al. 1999. With permission from Oxford University Press.

600 proteins. Listed in Table 15.1 are the key structural components, proteins involved in ciliary assembly, and many proteins that are at the forefront of ciliary biology for their roles in human disease. The molecular composition of cilia has been determined in several ways. Many of the proteins listed in Table 15.1 are known from proteomic studies of isolated flagella. All ciliated cells will shed their flagella in response to chemical or mechanical stress in a process known as deflagellation or deciliation (Quarmby 2004). This allows for relatively easy preparation of large quantities of isolated cilia, which can then be analyzed by mass spectrometry. This feature of ciliated cells has been used to obtain proteomes of Chlamydomonas flagella, rat olfactory cilia, and mouse retina photoreceptor cilia (Pazour et al. 2005; Liu et al. 2007; Mayer et al. 2009). The proteome of isolated Chlamydomonas centrioles has also advanced our understanding of how cilia are built and maintained (Keller et al. 2005). These proteomes have been invaluable for identifying components of cilia, but extensive functional characterization has been performed for only a fraction of them (Table 15.1). These functional characterizations have largely been the result of identifying genetic mutations affecting cilia in model organisms, and linking human disease genes to cilia. Comparative genomics and expression studies have also revealed a host of additional cilia-related proteins, many of which are involved in ciliary assembly but are not themselves components of the cilium (Avidor-Reiss et al. 2004; Li et al. 2004; Baron et al. 2007). Despite these advances, we are still far from having a complete picture of what goes into building a cilium.

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TABLE 15.1.

Components of Basal Bodies and Cilia

Protein/Complex Basal body (BB) proteins γ-Tubulin δ-Tubulin Centrin

Ninein

SAS-6

RGRIP1 (retinitis pigmentosa GTPase regulator [RPGR]interacting protein) OFD1

NPHP1/NPHP4

CEP290/NPHP6

RPGRIP1L (RPGRIP1like) MKS1

TZ proteins ε-Tubulin

p210

249

Description

Reference

Participates in nucleating centriolar microtubules Required for BB assembly Family of calmodulin-like proteins; component of distal fibers that contract in response to calcium signals and may participate in deflagellation; required for centriole duplication Associates with the mother centriole and participates in anchoring the distal end to the membrane Component of the cartwheel structure at the center of Chlamydomonas BBs; required for the establishment of ninefold symmetry of the BB; required for centrosome assembly in other species Localizes to connecting cilia of photoreceptors and to BB of primary cilia; interacts with nephronophthisis (NPHP)4, -5, -6, and RPGR

Stearms and Kirschner (1994)

Mutations are causative for oral–facial–digital syndrome type 1, with malformations of the face and digits and cystic kidneys; participates in assembling distal appendages and stabilizing centriolar MTs; required for ciliogenesis Localize to transition zone (TZ) (NPHP1) and BBs (NPHP4); regulates entry of intraflagellar transport (IFT) components and cargo into the cilium Localizes to BBs and to connecting cilium of photoreceptors; interacts with pericentriolar material-1 (PCM1); required for ciliogenesis and ciliary targeting of Rab8; mutations are associated with various ciliopathies (MKS, Joubert syndrome [JBTS], NPHP, etc.). Interacts with NPHP4; mutations are causative for JBTS. Required for ciliogenesis in embryogenesis; mutations cause plieotropic developmental abnormalities associated with cilia/Hh signaling. Necessary for assembly of centriolar MTs and anchoring BBs at the plasma membrane Component of Y-shaped connectors that anchor TZ to ciliary membrane

Dutcher and Trabuco (1998) Wright et al. (1985); Salisbury et al. (1987, 2002)

Mogensen et al. (2000)

Nakazawa et al. (2007)

Hong et al. (2001); Roepman et al. (2005); Otto et al. (2005); Chang et al. (2006) Ferrante et al. (2001, 2006); Singla et al. (2010)

Fliegauf et al. (2006); Mollet et al. (2005); Jauregui et al. (2008) Chang et al. (2006); Sayer et al. (2006); Frank et al. (2008); Kim et al. (2008b)

Arts et al. (2007) Weatherbee et al. (2009)

Dupuis-Williams et al. (2002); Dutcher et al. (2002); Chang et al. (2003) Lechtreck et al. (1999)

(continued)

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TABLE 15.1.

(continued)

Protein/Complex RPGR (RP GTPase regulator)

BBS3/ARL6

FA1/FA2

Axoneme α/β-Tubulin

Monomeric inner dynein arm (IDA)

IDA I1/f

Outer dynein arm

Radial spoke complex

Nexin–dynein regulatory complex

Central pair apparatus

C1a

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Description

Reference

Localizes to connecting cilia of photoreceptors, to TZ of motile cilia, and to BB of primary cilia; putative guanine nucleotide exchange factor; interacts with RPGRIP1 Small GTPase, GTP-, and GDP-locked forms stimulate ciliogenesis; localizes to the distal end of BB; modulates canonical Wnt signaling Chlamydomonas-specific proteins essential for Ca2+-induced deflagellation; localize to the distal end of the TZ where axonemal severing occurs; FA2 also localizes to this site when ectopically expressed in mammalian ciliated cells.

Hong et al. (2003); Shu et al. (2005); Gerner et al. (2010)

Compose ciliary microtubules; α-tubulin is acetylated; both can be glutamylated/ glycosylated. Six species (a–e, g) arranged along the inner surface of the A tubule of outer doublet MTs closest to the axonemal center; single dynein heavy chain (DHC) associated with actin and centrin or p28 light chain (LC). Inner and outer dynein arms are responsible for flagellar motility. Similar arrangement to monomeric IDA; two DHCs, three tryptophan aspartate-repeat (WD repeat) intermediate chains (IC), five different LCs Arranged on the outer surface of the A tubule of outer doublet MTs; composed of 26 subunits: 3 DHCs, 2 WD-repeat ICs, 18 LCs, three docking complex (DC) proteins Thin stalk with bulbous head pointing from outer doublets in toward central pair; mutations cause paralyzed flagella (PF); 23 known protein components with unknown stoichiometry Joins each outer doublet to neighboring doublets and suppresses dynein function when central pair apparatus is defective; mutations restore motility to paralyzed central pair/radial spoke mutants; seven known protein components with unknown stoichiometry Essential for flagellar motility in many cell types including Chlamydomonas, Tetrahymena, Drosophila sperm, mammalian sperm, and respiratory epithelia. Many Chlamydomonas PF mutants disrupt central pair formation altogether. PF6 (mutations disrupt assembly of C1a projection), four accessory proteins, calmodulin

Wiens et al. (2010)

Finst et al. (2000); Mahjoub et al. (2002, 2004)

Gaertig and Wloga (2008)

Reviewed by King and Kamiya (2009)

Reviewed by King and Kamiya (2009)

Reviewed by King and Kamiya (2009)

Yang and Smith (2009)

Heuser et al. (2009); reviewed in Wirschell et al. (2009)

Reviewed in Mitchell (2004, 2009)

Wargo et al. (2005)

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TABLE 15.1.

(continued)

Protein/Complex C1b

C1c C1d PF16 Bridge

Diagonal link C2a C2b

C2c

IFT proteins Kinesin-II/osmolarity mutant 3 (OSM-3)

Cytoplasmic dynein 1b

IFT complex A

IFT complex B

BBSome

251

Description

Reference

CPC1 (adenylate cyclase domain that may not be functional; defects disrupt C1b and C2b assembly), two accessory proteins, HSP70, enolase (causes reduced beat frequency due to reduced glycolytic ATP production within the cilium) Identified structurally; little biochemical/genetic data available Identified structurally; little biochemical/genetic data available Armadillo repeat protein; localizes to C1 tubule Connects C1 tubule to C2 tubule; PF20 (WD-repeat protein), PF16 (armadillo repeat protein) Identified structurally; little biochemical/genetic data available Identified structurally; little biochemical/genetic data available Hydin (hydrocephalus inducing) knockdown abrogates C2b formation in Chlamydomonas; mouse ortholog impairs ciliary motility contributing to hydrocephaly when defective. KLP1 (kinesin-like protein) knockdown eliminates EM density at C2c and disrupts C2b formation; also reduces motility; may act as a conformational switch, interacting with radial spokes to regulate axonemal dynein activity

Mitchell et al. (2005)

Anterograde motors; associate with complex B and transport cargo from the cell body into the cilium; OSM-3 is a secondary motor found in some species such as C. elegans. Retrograde motor; associates with complex A and transports cargo from the distal tip of the cilium to the cell body. IFT144, IFT140, IFT139, IFT122, IFT121, IFT43; mutations in IFT144 and IFT136 disrupt retrograde IFT. At least 17 polypeptides; core complex: IFT88, 2 × IFT81, 2 × IFT 74/72, IFT52, IFT46, IFT27, IFT25, IFT22; mutations in IFT88, IFT52, IFT46 disrupt anterograde IFT. Consists of one subunit each of BBS1, BBS2, BBS4, BBS5, BBS7, BBS8, BBS9; regulates assembly of IFT complexes; stabilizes interactions between complexes A and B in IFT trains; associates with Rab8, PCM-1

Kozminski et al. (1995); Scholey (2008); Snow et al. (2004)

Mitchell and Sale (1999) Mitchell and Sale (1999) Smith and Lefebvre (1996) Smith and Lefebvre (1997)

Mitchell and Sale (1999) Reviewed in Mitchell (2009) Lechtreck and Witman (2007); Lechtreck et al. (2008) Yokoyama et al. (2004)

Pazour et al. (1998); Scholey (2008) Cole (2009); Iomini et al. (2009) Cole (2009); Lucker et al. (2010)

Nachury et al. (2007); Ou et al. (2005); Pan et al. (2006)

(continued)

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TABLE 15.1.

(continued)

Protein/Complex Other cilia proteins (disease genes, etc.) End-binding 1 (EB1)

CNK2

LF4

PKG2

GSK3

Polycystin-1 (PC1)

Polycystin-2 (PC2)

Fibrocystin

NEK8/NPHP9

NPHP3

Inversin/NPHP2

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Description

Binds to plus ends of MTs; localizes to BB in mammalian cells, and to BB and ciliary tip in Chlamydomonas; required at the BB for assembly of mammalian primary cilia Chlamydomonas NIMA-related kinase; localizes along the length of the cilium; overexpression causes short cilia and small cell size, while knockdown causes long cilia and large cells. Chlamydomonas microtubule-associated protein (MAP) kinase; localizes to cilia; null mutations cause long flagella. Chlamydomonas cyclic guanosine monophosphate (GMP) dependent kinase 2; null mutations cause short, unequal length, or no flagella. Localizes to Chlamydomonas flagella in a phosphorylation-dependent fashion; inhibition/knockdown causes cells to become aflagellate. Most common affected gene in ADPKD; encodes a large membrane protein; participates in multiple cell proliferation and apoptosis signaling pathways including mammalian target of rapamycin (mTOR), Gα12/c-Jun N-terminal kinase (JNK) (apoptosis), cAMP/Ca2+ signaling, canonical Wnt signaling; PC2 interacts with and regulates PC1 at the cilium. Affected in ADPKD; mechanosensitive (flow-induced) Ca2+ channel; signaling pathways include nitric oxide, tumor necrosis factor alpha (TNF-α). Affected in ARPKD; affects flow-induced Ca2+ signaling, possibly through its interaction with PC2 NIMA-related kinase; associated with human NPHP type 9 and jck mice. Disease-causing mutations disrupt ciliary localization of NEK8; may affect ciliary localization of PC1/2; localizes to the Inv compartment at the proximal region of the primary cilium (see inversin) Large, 1330-amino acid protein associated with NPHP type 3 and Senior–Loken syndrome; localizes to the Inv compartment at the proximal region of the primary cilium (see inversin) Causes situs inversus and cystic kidneys in mouse models, NPHP type 2 in humans; contains ankyrin repeats; localizes to mouse node cilia and to a distinctive proximal compartment of primary cilia called the Inv compartment

Reference

Pedersen et al. (2003); Schroder et al. (2007)

Bradley and Quarmby (2005)

Berman et al. (2003)

Rasi et al. (2009)

Wilson and Lefebvre (2004)

Dere et al. (2010); Yu et al. (2010); Besschetnova et al. (2010); Lal et al. (2008)

Nauli et al. (2003); AbouAlaiwi et al. (2009); Li et al. (2008b) Rohatgi et al. (2008); Kim et al. (2008b); Wang et al. (2007) Otto et al. (2008); Trapp et al. (2008); Sohara et al. (2008); Shiba et al. (2010)

Omran et al. (2000); Olbrich et al. (2003); Shiba et al. (2010) Morgan et al. (1998); Otto et al. (2003); Watanabe et al. (2003); Shiba et al. (2009, 2010)

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TABLE 15.1.

(continued)

Protein/Complex PTCH1

Description Mammalian Sonic Hedgehog (Shh) receptor; localizes to cilia in the absence of Shh signals; exits the cilium in the presence of Shh Mammailan Shh effector; enters the cilium in the presence of Shh signals to release inhibition of downstream Shh signals Mammalian Shh effectors; become enriched in the cilium in the presence of Shh signals; ciliary localization helps regulate processing into the GLI3 repressor or maintenance of the GLI2 activator Defective in autosomal dominant RP, causing defects in the axoneme of connecting and outer segment (OS) cilia; shares homology with the MAP domain of doublecortin Small GTPase involved in exocytic vesicle docking at base of cilium; GTPase activity and associated guanine nucleotide exchange factor (GEF) Rabin8 are essential for ciliogenesis.

SMO

GLI2/3

RP1

Rab8

253

Reference Rohatgi et al. (2007)

Corbit et al. (2005)

Kim et al. (2009); Wen et al. (2010)

Liu et al. (2002); Yamashita et al. (2009)

Nachury et al. (2007)

BB BBs are the interphase form of centrioles. Centrioles come in pairs of orthogonally oriented short cylinders of nine triplet microtubules: The ABC triplet is composed of one complete microtubule (the A tubule, made of 13 protofilaments) and two partial microtubules (B and C tubules with 11 protofilaments each) that “piggyback” on the A tubule (see Fig. 15.1). With each cell cycle, centrioles undergo replication: A new “daughter” centriole forms next to each original “mother” centriole. At prophase, the original pair separates, each mother taking its daughter along into one of the daughter cells. The microtubuledriven separation of centrioles during cell division facilitates efficient separation of daughter cell components. During interphase, centrioles dock to the plasma membrane and elaborate into the BBs from which cilia are built. In addition to providing the foundation for cilia, BBs serve as microtubule organizing centers (MTOCs), directly impacting the structure of the cytoplasmic cytoskeleton (see also Chapter 14 on microtubules). BBs have a structural polarity that has been examined most thoroughly in Chlamydomonas (see Fig. 15.1; Ringo 1967; Johnson and Porter 1968; Cavalier-Smith 1974; Holmes and Dutcher 1989; O’Toole et al. 2003). In Chlamydomonas, a highly stereotyped system of fibers emanating from the BBs serves to position the two flagella, the nucleus, and the eyespot. While the specifics of this fiber system may be unique to the green algae, it is generally the case that the establishment of apical–basal and planar cell polarity (PCP) is tightly linked to BB placement in metazoans (Marshall 2010). In cells that possess only a single cilium, the mother centriole invariably serves as the BB of the cilium. With few exceptions, ciliated cells lose their cilia prior to entry into mitosis, presumably to provide flexibility and precision in centriole separation and positioning during cell division (Parker et al. 2010). However, in some organisms, such as C. elegans and Drosophila, only terminally differentiated cells are ciliated. This means

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that once centrioles become BBs, they never return to serve as centrioles again. In these cases, it is not uncommon to find that BBs are degenerate and comprised of only singlet or doublet microtubules, and become less distinguished from the microtubules of the ciliary axoneme (Marshall 2008). A key elaboration of the BB is the transitional fibers (known as distal appendages in metazoans) that facilitate docking of the BB to the cell membrane (Weiss et al. 1977). All cilia have these fibers, which are thought to play important roles in the docking and sorting of protein complexes destined for the cilium, although their exact protein composition remains elusive (Dutcher 2009). Many protein components of the BB have been identified in mutants with defective ciliary phenotypes. These include centrosome-specific tubulin isoforms δ- and ε-tubulin, the C. elegans spindle assembly protein 6 (SAS-6), and the human nephronophthisis (NPHP) proteins NPHP1 and NPHP4, which are associated with an extremely heterogeneic and plieotropic group of human genetic disorders of varying severity (see Table 15.1). Some of these proteins contribute directly to BB assembly, while others are required at the BB to regulate ciliogenesis. These mutants demonstrate the importance of the BB in building and maintaining normal cilia.

TZ The transition from the ABC triplet microtubules to the AB doublets occurs at the proximal end of the TZ. Assembly of the TZ on the BB is a key step in the formation of cilia, but to date, only two genes have been implicated specifically in this process (UNI1 and UNI2; Piasecki and Silflow 2009). Intriguingly, the junction between BB and TZ has recently been identified as the locus of a severing event that appears to serve to release the BB from the TZ for reentry into the division cycle (Parker et al. 2010). In some cilia, such as the flagella of green algae, the TZ contains highly stereotyped and distinctive electron-dense structures (Ringo 1967; O’Toole et al. 2003). In other cells, such as the primary cilia of mammalian cells, the TZ is difficult to identify. However, immunofluorescence localization of proteins involved in building and maintaining cilia has revealed specific proteins that localize to this region and thereby serve to define it, even in cells where no distinctive TZ can be identified ultrastructurally (see Table 15.1). At the distal end of the TZ is a break point that has become known as the site of flagellar autotomy (SOFA) (Mahjoub et al. 2004). Almost all eukaryotic cells shed their cilia in response to stress, and some do so as a prelude to cell division (Quarmby 2004). It is likely that a katanin-like microtubule-severing protein mediates severing of the outer doublet microtubules at the SOFA, but the evidence to date is circumstantial. A genetic screen in Chlamydomonas identified two proteins involved in breaking the axoneme at this site, FA1 and FA2 (Finst et al. 1998). FA1 appears to be a scaffolding protein and FA2 is a never in mitosis gene A (NIMA)-related kinase; both of these proteins localize to the SOFA in Chlamydomonas. Although the functionally orthologous proteins have yet to be identified in mammals, exogeneously expressed Chlamydomonas FA2 in mouse kidney cells reveals that it localizes to the analogous site on the mammalian primary cilium (Mahjoub et al. 2004).

The Cilium Deflagellation, the stress-induced ciliary shedding response, has been used for decades by ciliary researchers to provide an abundant source of cilia for biochemical and structural studies (Witman 1986). After a culture of cells has been induced to deflagellate, the flagella

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are isolated by differential centrifugation. Whole flagella can then be separated into the core axoneme and a detergent-soluble membrane-matrix fraction. The axoneme most often consists of nine doublet microtubules that run the length of the cilium. In some cases, such as in C. elegans sensory cilia, Chlamydomonas gametes, and mammalian olfactory cilia, the doublet microtubules transition to singlets toward the distal end of the cilium (Silverman and Leroux 2009). With few exceptions, motile cilia have a central pair of singlet microtubules that participate in regulating motility—these cilia are designated 9 + 2. Most nonmotile cilia, such as mammalian primary cilia, olfactory cilia, and photoreceptor outer segments, lack this central pair and are designated 9 + 0. The axoneme of a nonmotile 9 + 0 axoneme is relatively streamlined; the nine outer doublet microtubules are composed of α- and β-tubulin with various posttranslational modifications, including acetylated α-tubulin, and glutamylation and glycosylation of both tubulin isoforms. The ninefold symmetry established by the BBs continues throughout the length of the axoneme. Examined in cross section, a number of important distinguishing features of the motile 9 + 2 cilium are readily apparent (Fig. 15.1). The outer doublets are decorated with rows of inner and outer dyneins, radial spoke complexes, and interdoublet connections known as the nexin links. Recent tomographic studies have revealed that the nexin-link is the dynein regulatory complex, which plays a major role in regulating motility (Heuser et al. 2009). Ten or more heavy chain dyneins and a multitude of associated proteins form the outer arm dyneins and three classes of inner arm dynein. Different species of dynein are restricted to specific domains along the length of the flagella (Yagi et al. 2009). Further spatial organization is found in the distinct identities of each of the nine outer doublet microtubules (Wargo et al. 2005). Each singlet microtubule of the central pair has distinct projections, with the C1 tubule having longer projections than the C2 tubule. Table 15.1 describes the molecular composition of these structures. The “membrane-matrix” fraction of the cilium includes the lipids and proteins of the ciliary membrane, the intraflagellar transport (IFT) particles involved in ciliary assembly, and the as yet poorly defined soluble fraction of the compartment. IFT particles are made of two protein complexes (see Table 15.1) that span the space between the outer doublet microtubules and the ciliary membrane. The lipid and protein composition of the ciliary membrane is distinct from the plasma membrane. Predominant among ciliary membrane proteins are signaling proteins such as receptors and ion channels (Dunlap 1977; Berbari et al. 2009). The distinctive lipid composition of ciliary membranes is an active area of investigation.

The Ciliary Tip Electron microscopy (EM) has demonstrated that in the motile flagella of Chlamydomonas and Tetrahymena, the basic structure of the ciliary tip includes a plate and ball structure that anchors the central pair to the ciliary membrane, and a plug inserted into the A tubule of each outer doublet (Dentler 1980). The function and composition of these structures is not yet defined. However, the flagellar tip must perform three essential functions necessary for maintaining the axoneme: regulating the turnover of flagellar subunits, loading and unloading IFT cargo, and regulating the activity of microtubule motors that transport IFT cargo along the axoneme (Sloboda 2005). The plus-end-binding protein EB1 localizes to the flagellar tip in Chlamydomonas and has an essential role in the assembly of primary cilia (Pedersen et al. 2003; Schroder et al. 2007). It is not yet clear, however, whether this is due to the known microtubule-stabilizing functions of EB1 or some other, yet unknown function at the flagellar tip.

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The Machinery of Ciliary Assembly A major distinguishing feature of cilia is that they are synthesized de novo in each cell, and sometimes more than once in the lifetime of a single cell due to stimuli that may cause the cilium to be shed or resorbed for a period of time. A complete molecular machine known as IFT is dedicated to building and maintaining the cilium. After a cell has completed mitosis, the first step in creating a new cilium is to dock the BB at the plasma membrane (Marshall 2008). The migration of the BB to the surface of the cell involves intracellular polarity cues, as the cilium forms at the apical surface of the cell. The TZ and its associated fibers are then assembled, and this structure anchors the cilium in its permanent position in the plasma membrane. The cilium is assembled via the addition of ciliary components to the distal tip. IFT complexes A and B, assembled from distinct subunits, transport cargo from the cell body toward the tip of the axoneme in IFT “trains” containing multiple A and B complexes (Fig. 15.3). Anterograde transport (toward the plus end of the ciliary microtubules and the distal tip of the cilium) is mediated by kinesin-II, which is thought to associate with complex B, while retrograde transport is mediated by cytoplasmic dynein in association with complex A (Cole 2009). Mutations in complex B subunits obliterate ciliogenesis (Pazour et al. 2000; Brazelton et al. 2001), while mutations in complex A subunits cause accumulation of material at the ciliary tip (Iomini et al. 2009). In addition to carrying the building blocks of the ciliary axoneme, there is growing evidence that IFT trains also participate in the transport of membrane proteins along the axoneme (Emmer et al. 2010). At least one IFT protein (IFT20) is involved not only in the ciliary IFT trains but also in vesicular traffic (Follit et al. 2006). IFT20 is thought to facilitate the trafficking of vesicles carrying ciliary membrane proteins from the Golgi to the base of the cilia. These vesicles then fuse with the plasma membrane near the transitional fibers, delivering membrane proteins and lipids to the IFT particles for transport into the cilium. Baldari and Rosenbaum (2010) proposed that axonemal components might also be targeted to the cilium via association with cilia-destined vesicles (see also Chapters 8, 18, 21, and 22 for further discussion on polarized trafficking). Anterograde and retrograde IFT remain active throughout the lifetime of a cilium. Thus, cilia are dynamic structures and the steady-state length is a consequence of a balance between the length-dependent rate of assembly and the length-independent rate of disassembly; that is, cilia grow until these two processes are in balance (Marshall et al. 2005). Several proteins have been implicated in controlling this balance point, including both ciliary and cell body proteins, among them at least five distinct kinases (see Table 15.1). It is likely that diverse pathways regulate ciliary length, mediating diverse physiological changes related to cell cycle, cell polarization, cell migration, and fluid flow. We have barely begun to understand the mechanisms and physiological significance of ciliary length control.

FUNCTIONAL IMPLICATIONS OF DOMAIN ORGANIZATION Sensory Antennae Cilia evolved as sensory organelles (Satir et al. 2008). While all cilia likely perform at least rudimentary signaling functions, some are highly specialized for sensory functions. The outer segment of retina photoreceptor cells is a highly derived cilium. Primary cilia lining renal tubules sense flow. Olfactory cilia are the site of localization of olfactory

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Figure 15.3. Intraflagellar transport (IFT). (1) IFT components accumulate at the basal body and are assembled on the transition fibers. Membrane proteins arrive at the base of the cilium from the Golgi via vesicular transport pathways. (2) IFT particles transport cargo along the ciliary axoneme in IFT trains. In certain species, such as C. elegans, osmolarity mutant 3 (OSM-3) supplements kinesin-II-driven anterograde transport. (3) In these species, OSM-3 is the sole molecular motor responsible for driving anterograde transport along the distal singular MTs. (4) At the distal tip of the cilium, IFT complexes disassemble, and structural components may be added at the tip of the axoneme. As ciliary components are turned over, IFT complexes are reassembled for transport back to the cell body. (5) IFT trains transport ciliary components using cytoplasmic dynein as the exclusive molecular motor for retrograde transport. (6) Retrograde IFT complexes are disassembled at the BB, and membrane components are degraded using the endosomal recycling pathway. (7) TEM of Chlamydomonas flagella showing three IFT trains. Black arrowheads indicate the longer anterograde trains, and the white arrowhead indicates a shorter retrograde train. © Pigino et al., 2009. Originally published in the Journal of Cell Biology 187:135–148.

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receptors. C. elegans possess cilia only on the termini of sensory nerves, where they are exposed to the external environment and participate in chemosensory functions. The cilia of respiratory epithelia, whose function is to sweep mucus and debris out of the airways, possess bitter taste receptors. When bitter compounds stimulate these receptors, ciliary beat frequency increases, a response which may help remove noxious substances from the airway more rapidly (Shah et al. 2009). In Chlamydomonas gametes, receptor proteins known as agglutinins localize to flagella and trigger signal transduction events involved in mating (Quarmby 1994; Wang and Snell 2003). A common feature of sensory cilia is that receptor molecules are concentrated within the cilium (Fig. 15.4). This can be observed in photoreceptor cells, where the lightdetecting machinery is condensed in the cell’s outer segment; in olfactory cilia, where the G-protein-coupled receptors and associated G proteins activate cyclic adenosine monophosphate (cAMP) signaling and neuron depolarization from within the cilium; and in neuronal cilia, where particular somatostatin and serotonin receptors localize exclusively to primary cilia (Whitfield 2004; Berbari et al. 2009). Thus, a number of normal signaling functions are entirely dependent on the presence of a healthy cilium. A growing number of human diseases are attributed to sensory dysfunction of cilia. For example, the products of cystic kidney disease genes, such as polycystin-1 and -2 (PC1 and PC2; autosomal dominant PKD), fibrocystin (autosomal recessive PKD), and various NPHP proteins, localize to cilia (Table 15.1). In response to flow, PC1 and PC2 initiate downstream signaling events that include Ca2+ influx and transcriptional changes. Cilia are also critical for developmental and homeostatic Hedgehog (Hh) and Wnt signaling. In mammals, the Hh receptor PTCH1 and all the downstream Hh effectors localize to cilia, and proper localization of these components is essential for normal Hh signaling (Goetz and Anderson 2010). For Wnt signaling, the presence or absence of a cilium determines whether canonical or noncanonical Wnt signaling is activated (Lai et al. 2009). The roles of these pathways in tumorigenesis also strongly implicate the cilium as an important organelle in cancer. NPHP, the most severe form of juvenile cystic kidney disease, has joined a rapidly expanding group of heterogeneic ciliopathies associated with early development and physiological homeostasis: Bardet–Beidl syndrome (BBS), Meckel syndrome (MKS), Joubert syndrome (JBTS), NPHP, and isolated and syndromic forms of retinitis pigmentosa (RP) (Table 15.1). These disorders are highly pleiotropic, and symptoms can include retinal degeneration, cysts of the kidney and liver, facial malformations, neural tube defects, mental retardation, polydactyly, and obesity. While the disorders are clinically distinct, some genes contribute pathogenic alleles to multiple disorders. For example, nonsense mutations in centrosomal protein (CEP290) cause MKS, the most severe of these disorders, while hypomorphic mutations can cause RP, NPHP, BBS, or JBTS (Chang et al. 2006; Sayer et al. 2006; Frank et al. 2008; Leitch et al. 2008). Many of the abnormalities in these disorders have connections to Wnt, Hh, and PCP signals, all of which have cilia-dependent functions in development (Zaghloul and Katsanis 2009). Given that defects in this group of genes can cause defects in ciliary structure, the phenotype for any given mutation may depend on the severity of the ciliary defect, the tissue affected by ciliary defects, the time during development that these defects arise, and the cilia-dependent signaling pathways affected.

Motility Although all cilia are sensory, some are also motile. Ciliary beating drives the swimming behavior of unicellular organisms, such as Paramecium, Tetrahymena, and Chlamydomonas,

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Figure 15.4. Ciliary functions. (A) Ciliary waveform versus flagellar waveform. In ciliary waveform, an asymmetric wave provides a power stroke and a recovery stroke. In unicellular organisms such as Chlamydomonas, apical flagella pull the cell forward. This same waveform is used by epithelial cells to draw fluids across a surface. In flagellar waveform, cells such as human sperm, swim with their flagella behind them, generating motility through a whiplike action (adapted from Smith et al. 2009, with permission). (B) Signal transduction in cilia. Sensory receptors are localized along the surface of the cilium. In response to an external signal, such as light, chemicals, or flow, the sensory receptors initiate a signaling cascade to induce downstream effects on the cell. (C) During interphase, the centrosome is located adjacent to the plasma membrane of the cell, and the mother centriole becomes the BB that nucleates a cilium. Before cell division, the cilium is resorbed. The centrioles then duplicate and participate in cytokinesis. After cell division, new cilia are built from the older of the two centrioles.

and colonial and multicellular protists such as Volvox and Planaria. The gametes of multicellular organisms, for example, mammalian sperm, can be propelled by cilia (often known as flagella in this context). In complex tissues, motile cilia can drive the flow of fluid over a surface. In mammals, this includes respiratory epithelia, ependymal cells lining the brain cavities, and the epithelial cells that drive fluid flow in oviducts. Whether propelling a cell over distance or moving fluids over a surface, the molecular machinery that drives underlying ciliary beat is conserved. Commensurate with the diversity of motile cilia in the human body, a number of disease states derive directly from the loss of ciliary motility. Among the first ciliopathies

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defined is Kartagener ’s syndrome, also known as primary ciliary dyskinesia (PCD). This disorder is caused by loss of ciliary motility, often due to mutations in axonemal dynein subunits (Leigh et al. 2009). Predominant symptoms include chronic bronchitis due to inefficient clearance of mucus from the airway. Because motile cilia in the embryonic node are essential for establishing morphogen gradients that determine left–right asymmetry, approximately 50% of PCD patients have the placement of their organs reversed. The motile cilia of ventricular ependymal cells help maintain circulation of cerebrospinal fluid (CSF), without which CSF builds up in the brain causing hydrocephalus. The force that drives ciliary beating is generated by the ciliary dyneins. The dyneins are molecular motors that use ATP hydrolysis to drive large conformational changes that result in a series of steps along a microtubule track. Dynein walks toward the plus end of microtubules, which in cilia is the distal tip of the cilium. This means that the A tubule, to which the motors are bound, would be carried along the adjacent B tubule toward the tip; that is, the microtubule doublets would slide over one another. In the intact cilium, the doublets are cross-linked to one another by other proteins, and additionally, they are anchored by the BB. If all ciliary dyneins were active at the same time, the cilium would be under substantial tension, but no movement would be generated. Precise temporal and spatial regulation of dynein activity is required to generate ciliary motility. In order for a bend to form, only the dyneins on one side of the axoneme can be active. Furthermore, a simple bend will not generate propulsive force. In order to be effective, the cilium must produce an oscillatory wave. Two different types of wave are commonly observed in motile cilia. The ciliary waveform is an asymmetric, breaststroke-type beat, with a power stroke and a recovery stroke (e.g., respiratory epithelia and Chlamydomonas) that pull a cell forward (see Fig. 15.4). The flagellar waveform is a symmetric whiplike wave that pushes a cell from behind. In order to generate either of these productive waves, dynein activity must be tightly controlled and coordinated, both around the circumference and along the length of the cilium. The question of precisely how this coordination is accomplished remains an active area of current research. Although we do not yet fully understand how the generation of an oscillatory wave is accomplished, there is likely a complex interplay of forces at work. Ultimately, changes in interdoublet spacing distorted by stress may drive the propagation of the wave of motor activity/inactivity (Mitchison and Mitchison 2010). There is evidence that the dynein molecules themselves are tuned oscillators that respond to the loading conditions induced by bending (King and Kamiya 2009). Intensive research has revealed that the inner and outer dyneins may be differentially regulated and at least two systems of mechanosensory transduction feed into the generation of effective ciliary waveforms (King and Kamiya 2009). Ultimately, these transduction pathways must be modulating dynein activity. Much of what we know about the regulation of dynein activity derives from microtubule sliding assays. Mild proteolytic digestion of isolated axonemes frees the links that hold the doublets together. In the presence of ATP and ADP, the axoneme will telescope out, sometimes approaching the full ninefold increase in length (Summers and Gibbons 1971). Using mutant strains of Chlamydomonas, the sliding assay described above, and biochemical and EM-based structural studies, it has been shown that a regulatory enzymatic cascade originates at the central pair, which rotates and may thus serve as a distributor. This signal is thought to pass from the central pair to the radial spokes to inner arms (Smith 2002), activating and inactivating dyneins sequentially around the circumference and from base to tip (Mitchison and Mitchison 2010). The specific placement of different dyneins may also contribute to the generation of effective waves. For example, the three classes of inner arm dyneins, each of which comes in a variety of flavors, dock to specific sites in a 96-nm repeat pattern (Piperno et al. 1990).

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261

An additional complexity is that some dyneins are found only in the proximal realms of the cilium (Yagi et al. 2009). Beyond the striking accomplishment of generating a productive waveform, cilia can modulate beat frequency or undergo a waveform switch in response to signals from the environment. Calcium and redox potential are important relay signals that regulate various components of the motile apparatus. For example, in response to bright light, opening of a voltage-gated calcium channel that is restricted to the distal region of the flagella causes a switch from ciliary to flagellar waveform in Chlamydomonas. In spite of tremendous progress and a fabulously detailed understanding of the motile apparatus, there remains a great deal that we do not understand about the generation and regulation of ciliary motility.

Cell Proliferation and Differentiation The spectrum of developmental and physiological disorders that arise as a consequence of ciliary dysfunction are indicative of the fundamental cellular processes controlled by cilia. As we have seen above, motile cilia are required for proper organ placement, respiratory and cognitive function, and fertility; it is through the sensory functions of cilia that we see and smell the world around us. Signaling pathways localized to cilia guide development and homeostasis of our tissues, and they play important roles in our physiology. All of these functions are directly related to the motile or sensory activity of cilia. But there are additional consequences of the organization of this cellular domain. In order to function effectively, the cells in a complex tissue must be properly oriented with respect to one another. This is especially apparent in ciliated tissues where cilia must beat in the same direction in order to move fluids over an epithelial surface. Distinct from apical–basal polarity, this type of orientation is called PCP. The signaling pathways that establish and maintain PCP are an intensive focus of current research. Although as yet poorly understood, it appears that a feedback loop exists between fluid flow generated by ciliary beat and PCP signaling (Marshall 2010). Independent of its function as a sensory and sometimes motile organelle, the mere formation of a cilium impacts cell proliferation and differentiation. There is a cycle of ciliary resorption and reassembly that is tightly linked to the cell cycle in ways that we have barely begun to understand (Fig. 15.4). Releasing the BB from the TZ appears to be an important prerequisite for mitosis, and proteins that are involved in IFT during interphase may do double duty as participants in cytokinesis during cell division (Baldari and Rosenbaum 2010; Parker et al. 2010). This is a particularly rich area for future study.

ABBREVIATIONS BB BBS CEP290 CSF EM ET FA FLA Hh JBTS MKS MT MTOC

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basal body Bartdet–Beidl syndrome centrosomal protein cerebrospinal fluid electron microscopy electron tomography flagellar autotomy flagellar assembly Hedgehog Joubert syndrome Meckel syndrome microtubule microtubule organizing center

NIMA NPHP OSM-3 PC PCD PCP PKD PTCH RP SAS-6 SOFA TEM TZ

never in mitosis gene A nephronophthisis osmolarity mutant 3 polycystin primary ciliary dyskinesia planar cell polarity polycystic kidney disease patched retinitis pigmentosa spindle assembly protein 6 site of flagellar autotomy transmission electron microscopy transition zone

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INTERMEDIATE FILAMENTS Normand Marceau Anne Loranger Stéphane Gilbert François Bordeleau

DEFINITION Intermediate filaments (IFs), in concert with actin microfilaments (MFs; Chapter 12) and microtubules (MTs; Chapter 14), form the cytoskeleton in all nucleated eukaryotic cells. Each of these fibrillar networks exhibits rather unique structural and functional characteristics (Coulombe and Omary 2002), the exclusive IF features including an absence of polarity, a relative insolubility in nonionic detergents, and a wide subcellular dispersion between the surface membrane and the nucleus. Moreover, while both actins and tubulins are ubiquitous and encoded by few genes, the expression of IF proteins is encoded by more than 60 different genes (Hesse et al. 2001; Omary et al. 2009), classified into several types that are to a large extent characteristic of the tissue in which they are expressed. Furthermore, in contrast to actins and tubulins, which exhibit a highly conserved constitution, the structure of IF proteins is most diverse, except for conserved subdomains within a central domain. In fact, this structural heterogeneity of IF proteins is provided by the flanking amino and carboxy terminal domains (Coulombe and Omary 2002), which contain sites that can undergo posttranslational modifications, such as phosphorylation (Omary et al. 2006), meaning that both domains are important from regulatory and functional standpoints. Of particular note, these terminal domains are largely responsible for spatiotemporal IF organization as well as IF interactions with associated proteins (Coulombe and Wong 2004; Green et al. 2005). In line with these unique structural features, new functions have emerged recently for keratin IFs, namely in modulating cell adhesion/migration and providing resistance to mechanical stress and death-receptor-stimulated apoptosis, through interplay with surface membrane microdomains.

HISTORICAL PERSPECTIVE As stated in a recent review by Oshima (2007) on this subject, “the history of IFs is remarkably characterized by divergent beginnings, a shared common discovery period and a divergent future in terms of functions.” For instance, the first piece of evidence on IF structure was derived from X-ray diffraction data on hair keratins that reflected the alpha helical model by Pauling, and the prediction of the alpha helical coiled-coil structure

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by Crick (1952). However, the presence of IFs in tissues became real upon imaging by electron microscopy in the 1960s, particularly through the work of Holtzer et al. on developing muscle cells (Ishikawa et al. 1968); of particular note, these investigators were the ones who named them IFs because their diameter was found to be intermediate between MFs and myosin. The recognition of neurofilaments in neuronal axons then followed, based again on the microscopic evidence for abundant filaments of size intermediate between MFs and MTs (Huneeus and Davison 1970). Together, these structural observations turned out to be the first clear-cut demonstration by which IFs became recognized as a cytoskeletal network in eukaryotic cells. The next relevant breakthrough came from extensive immunochemical analyses performed on multicellular tissues, through the use of cell-type-specific IF antibodies, by Weber, Franke, Osborn, Sun, and Liem and their respective collaborators in the late 1970s and early 1980 (see Oshima 2007 for review). As a result of such cell-typing analyses, IF proteins were recognized as useful differentiation and cell-type markers at the postnatal stage. Thereafter, complementary information on IF composition came from studies of early embryonic development, namely through the generation of monoclonal antibodies against two cytoskeletal proteins of teratocarcinoma cells by Brulet et al. (1980), and protein purification work by Oshima of endo A and endo B from mouse embryonal carcinoma cells that turned out to be the antigens recognized by the above mentioned antibodies (Oshima et al. 1983); interestingly, endo A and B were subsequently identified by RNA and gene cloning as keratins 8 and 18, respectively (Oshima et al. 1996). Still, the similarity of different IF proteins was first highlighted in 1981 following the discovery of an antigenic epitope common to all IF proteins and recognized by a single monoclonal antibody (Pruss et al. 1981). At the same time, broad analyses using protein sequencing plus mRNA cloning/sequencing of cytoplasmic IFs revealed a conserved central coiled-coil domain, with the common antigenic epitope being localized at the C-terminus. Subsequent cDNA sequence and exon gene structure analyses led to key findings on the evolutionary emergence of IF proteins. For instance, nuclear lamins were identified as members of the IF superfamily, but were found to contain extended helical segments absent in cytoplasmic IFs (McKeon et al. 1986). However, these segments were discovered in IFs of invertebrates, such as Caenorhabditis elegans (Karabinos et al. 2001), thus suggesting an evolutionary path from invertebrates to nuclear lamins, to cytoplasmic IFs in mammalian cells (Hesse et al. 2001; Oshima 2007). Much of the lagging period in our knowledge of IF functions has been largely due to the fact that, in contrast to actin MFs and MTs, there was no chemical agent capable of specifically disrupting IF networks. So, most of our understanding on IF functional activities came from the development of gene targeting methods to genetically knockout the expression of IF genes in mice (Oshima 2007); the first IF gene to be inactivated was keratin 8 (K8) by Baribault and Oshima (Baribault et al. 1993). While variable numbers of K8-deficient mice were found to die at the embryonic stage, subsequent work on the survivors revealed an increased sensitivity of K8-deficient hepatocytes to mechanical stress, in support of the original hypothesis on IF mechanical function (Loranger et al. 1997). However, further analyses of these animals pointed to a hypersensitivity of the liver (hepatocytes) to toxic stresses, including those that led to cell death (Marceau et al. 2001a). On these grounds, IFs were found to exert both mechanical and nonmechanical functions, and from a historical perspective the original expectations that IFs would play comparable roles in different cell types, analogous to the well-known cytoskeletal functions of MFs and MTs, had to be re-revisited. For instance, there is accumulating evidence indicating that keratin IF and other IF networks are involved in cell signaling regulation (Gilbert et al. 2008; Bordeleau et al. 2010).

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Finally, an important IF research area emerged following the discovery made by Coulombe and Fuchs, in the early 1990s, of mutations in keratin genes that directly cause human skin diseases (Coulombe et al. 1991). Subsequent genomic analyses revealed that IF-associated diseases, designated as IF-pathies, in humans involve a broad range of tissues, in line with the cell-type expression of the IF genes (Omary 2009). On the whole, these analyses demonstrated that IF gene mutations either cause or predispose their carriers to more than 80 human diseases, including several degenerative conditions and cancer.

KEY IF PROTEIN FEATURES IFs as a Multiple Gene Family of Proteins IF proteins are classified into six types, which reflect key differentiation features of the cells of origin (Table 16.1). The first four types are present in the cytoplasm and are recognized as acid (type I) and basic (type II) keratins present in all epithelial cells; vimentin, desmin, glial fibrillary acidic protein (GFAP), and peripherin (type III) in mesenchymal, muscle, glial, and neuronal cells respectively; and α-internexin and neurofilaments (type IV) in neurons and cells of the peripheral neuroendocrine system. The type V is provided by the lamins, which form a meshwork of proteins at the inner layer of the nuclear membrane, as well as intranuclear structures of which the molecular organization is poorly understood (Weber et al. 1989; Broers et al. 2006). The two type VI proteins, filensin and phakinin, are found exclusively in the lens (Table 16.1). TABLE 16.1.

Classes of IF Proteins

Type

Name

No. of Gene(s)

Distribution

I (acid)

Human type I epithelial keratins Human type I hair keratins Nonhuman type I epithelial and hair keratins

17 11 2

Epithelia Epithelia Epithelia

II (basic)

Human type II epithelial keratins Human type II hair keratins

20 6

Epithelia Epithelia

III

Vimentin GFAP Desmin Peripherin

1 1 1 1

Heterogenous Astrocytes/glia Muscle PNS Neuron

IV

NF-L NF-M NF-H Synemin α-Internexin Nestin Syncoilin

1 1 1 1 1 1 1

CNS Neuron CNS Neuron CNS Neuron Muscle CNS Neuron Heterogenous Muscle

V

Lamin A/C Lamin B1 Lamin B2

1 1 1

Ubiquitous/nucleus Ubiquitous/nucleus Ubiquitous/nucleus

VI

Filensin Phakinin

1 1

Lens Lens

PNS, peripheral nervous system; CNS, central nervous system.

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Keratins, the IF proteins of epithelial cells, actually constitute the largest family of cytoskeletal proteins and are grouped into type I (K9–K42) and type II (K1–K8 and K71–K86) subfamilies (Schweizer et al. 2006). Keratin IFs are obligate heteropolymers that include at least one type I keratin and one type II keratin, and are coordinately expressed as specific pairs in a cell lineage and differentiation manner. Notably, the K8/ K18 pair constitutes the first cytoplasmic IF genes expressed in the blastocytes at the early embryonic stage. As the main cell lineages emerge, these keratin genes are downregulated and lineage-specific IF genes, for example, vimentin in the mesenchyme including the endothelium, are expressed. In turn, as mesenchymal progenitors differentiate into more specialized cells of the same lineage, the vimentin gene is downregulated and the specific type III IF genes are turned on, for example, desmin in muscle cells, GFAP in glial cells, and peripherin in peripheral neurons (Oshima 2007). The neuroepithelial progenitor cells express the type IV gene nestin, and as they progress along the neuronal lineage, vimentin is transiently expressed. When the neurons become fully differentiated at the late fetal period, the vimentin is downregulated and the neurofilament genes are turned on. Similarly, the emergence of the periderm is associated with the maintenance of blastocytic keratins, that is, K8/K18, and the emergence of epidermal keratinocytes is characterized by their disappearance and the expression of basal cell keratins, for example, K5/K14. In postnatal epidermis, the ratio and spectrum of keratins vary during keratinocyte terminal differentiation (e.g., K5/K14 --> K1/K10) (Coulombe and Omary 2002). In simple epithelia, all cells contain the embryonic K8/K18 pair, and some of them express 2–3 other keratins in addition (Oshima et al. 1996; Omary and Ku 1997). This is the case for biliary ductular cells in the liver and ductal cells in the pancreas, which express K7, K8, K18, and K19 (Marceau and Loranger 1995; Toivola et al. 2000). Intriguingly, hepatocytes maintain solely the blastocytic K8/K18 pair even after birth, suggesting that these differentiated liver parenchymal cells that express multiple specialized functions nevertheless maintain key embryonic phenotypes. Finally, other tissues are characterized by a mixed expression of stratified and simple epithelial keratins; this is indeed the case for the mammary glands, where the basal cells express K5 and K14, and the luminal cells contain K7, K8, K18, and K19 (Oshima et al. 1996). So, in this case, it appears that the presence of more than one keratin pair in a particular epithelium reflects the degree of its cellular complexity.

IF Protein Domains and Assembly Individual IF proteins, such as K8 and K18 (Fig. 16.1A), consist of a central α-helical (rod) domain flanked by N-terminal and C-terminal globular domains (Coulombe and Omary 2002; Herrmann and Aebi 2004). The rod domain constitutes a major driving force during the assembly of IF proteins. It consists of four segments (IA, IB, IIA, and IIB) that are separated by three linkers (L1, L1–2, L2), which act as modulators of IF assembly. The beginning and end portions of the rod domain are conserved among all IF proteins, and point mutations in these rod portions lead to aberrant IF organization and dynamics (McLean and Lane 1995; Omary and Ku 1997), which in turn result in dramatic loss of cellular integrity and severe pathologies. While type III proteins such as vimentin and desmin can assemble as homopolymers or heteropolymers, keratins assemble as obligate heteropolymers of individual type I and type II proteins (Coulombe and Wong 2004). Of particular note, while IF assembly in vitro is a spontaneous energy-independent process that does not necessitate associated proteins (Herrmann and Aebi 2004), recent imaging analysis of living cells has revealed that K8/K18 IF formation in simple epithelial cells only takes place through the involvement of IF particle precursors that originate from

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Figure 16.1. (A) Main structural features of K8 and K18, identification of phosphoSerine (Ser) residues and their associated kinases, and localization of mutations in transgenic mice and of plectin binding domain (BD). (B) Schematic representation of plectin indicating the binding domain (BD) positions for actin, microtubule-associated protein 1 and 2 (MAP1/2), IF, and RACK1.

actin-anchoring focal adhesions (FAs; see Chapter 17), a process that appears to be independent of the cell differentiation status (Windoffer et al. 2006).

IF Protein Domains and Cytoskeletal Interplay Notably, the head and tail domains contribute to most of the structural heterogeneity of IF proteins (Coulombe and Omary 2002) and are largely responsible for the spatiotemporal cytoskeletal organization as well as interactions with various associated proteins (Coulombe and Wong 2004; Green et al. 2005). This is relevant to the maintenance of overall cellular functional integrity, particularly in polarized epithelial cells, where a tightly controlled interplay among IFs, MFs, and MTs is required. Yet these networks are unable to interact directly with each other, which means that the interactions must take place through a cytolinker, such as plectin (Fig. 16.1B), a member of the plakin family, which also includes desmoplakins (Wiche 1998b; Fuchs and Yang 1999). The key molecular and cellular features of plectin are the following. (1) This is the largest (>500 kDa) member of the plakin family, and its activity depends on homodimer formation (Wiche 1998b; Rezniczek et al. 2004). (2) It possesses a binding site for IF proteins (including K8/K18) at the C-terminal domain, and others at the N-terminal domain for MT-associated proteins and actin (Wiche 1998a, b), which makes it a pivotal player in the integration of regulatory cytoskeletal events (Svitkina et al. 1996). (3) Based on sequence differences in 5′ DNA regions, several plectin isoforms have been identified in tissues (Rezniczek et al. 2003). (4) These small alternative N-terminal sequences profoundly affect their cytoplasmic localization (Green et al. 2005); for instance, one isoform is exclusively targeted to mitochondria (see Chapter 6), providing an IF connection to these organelles, whereas another one is concentrated to FAs, a surface membrane domain further described in the next section (see also Chapter 17 on FAs). In Madin-Darby canine kidney (MDCK) cells, plectin has been found to regu-

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Figure 16.2. Immunolocalization of keratin 8, actin, tubulin, plectin, and desmoplakin in mouse hepatocytes in situ and in a monolayer culture at 24 hours post-seeding. Note that the individual proteins are predominantly localized at the surface membrane. Reproduced from Marceau et al. (2001a).

late both the structural arrangement of K8/K18 bundles at the cell periphery and their association with the submembrane actin–MFs (Eger et al. 1997), perhaps linked with the formation of the membrane domains. In hepatocytes, both K8/K18 IFs and plectin are largely present at the cell periphery (Fig. 16.2) and a loss of K8/K18 alters its content, which implies a coordinated targeting of K8/K18 and plectin to surface membrane domains.

K8/K18 IFs and Cell Signaling The head and tail domains of K8 and K18 contain serine (Ser) sites with motifs that are recognized by signaling enzymes, for instance, the mitogen-activated protein (MAP) kinase cascades and protein kinase C (PKC) (Omary et al. 2006; See Fig. 16.1A). The motifs at three K8 Ser sites, for instance, Ser74, are recognized by MAP kinases p38 and JNK, and Ser432 by ERK1/2 (Omary et al. 2006). For K18, two phosphoSer sites, Ser34 and Ser53, have been documented, which are targets for CDK1 (cyclin-dependent kinase 1) and Raf-1, respectively (Omary et al. 2006; Fig. 16.1). Of note, IFs are among the most abundant cellular phosphoproteins, and several lines of work have indicated that the phosphorylation and dephosphorylation events are essential in the regulation of IF dynamics by modulating the intrinsic features of IFs, namely their solubility, conformation, and arrangement. In addition, these phosphorylation-regulated events dictate IF subcellular compartmentalization, levels and turnover, and binding with associated proteins (Omary et al. 2006). However, conversely, K8/K18 IFs can act as regulators of trafficking/signaling, a prominent functional trait that requires the involvement of scaffold/adaptor proteins. For instance, plectin possesses a binding site for receptor for activated C kinase 1 (RACK1), identified as an anchoring protein for PKC (Osmanagic-Myers and Wiche 2004). Of note, the plectin binding site for RACK1 is located next to that for IFs at the tail domain (Fig.

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16.1B), which can then facilitate IF protein–RACK1 interactions. Such findings suggest that keratins and plectin can work jointly as a signal-integrating platform in epithelial cells. This platform would be able to assimilate cues generated by signals triggered at the surface membrane, and to govern the transduction of these signals in a proper spatiotemporal sequence within the cells.

KERATIN INTERPLAY WITH SURFACE MEMBRANE DOMAINS Keratin IFs and Desmosomes One key function that is common to all keratin IFs, for example, the K5/K14 pair in keratinocytes and K8/K18 IFs in hepatocytes, is their capacity to maintain surface membrane integrity under mechanical stress largely through interactions with desmosomes, a polarity-dependent portion of the surface membrane involved in cell–cell adhesion (Fig. 16.2 and Fig. 16.3A; see also Chapter 19 on desmosomes). Keratin IFs exhibit unique viscoelastic properties that render them resistant to cell deformation and other mechanical stresses, and since desmosomes link keratin IFs of one cell to those of its neighbors, together keratin IFs and desmosomes form an integrated and mechanically resilient network across the epithelium (Green et al. 2005). The adhesive property of desmosomes is provided by proteins of the cadherin family, desmogleins/desmocollins (Marceau et al. 2001b), and the other main building elements are plakoglobin and desmoplakins (Marceau et al. 2001b). Together, these proteins are assembled according to a hierarchy of protein– protein interactions, with desmoplakins being responsible for anchoring the keratin IFs to the desmosomes. By using the K8-null mouse model generated via a targeted mutation in the germ line where the lack of K8 leads to the absence of K18, we assessed the K8/K18–desmosomal interaction in K8/Kl8-lacking and wild-type hepatocytes in vivo and in monolayer culture (Loranger et al. 2006). The results revealed that in wild-type hepatocytes, both K8/K18 IFs and desmoplakin are largely lined at the surface membrane and that the loss of K8/ K18 leads to alterations in the content and deposition of desmoplakin, but not of desmoglein in situ; of note, a reinsertion of wild-type K8 into K8-null hepatocytes rescues desmoplakin targeting, via the head domain. Moreover, since keratins are IF proteins whose dynamics and reorganization are phosphorylation-dependent, we addressed the role of Ser phosphorylation in the K8/K18–desmoplakin interaction and found that the desmoplakin/plakoglobin rescue can be modulated via phosphorylation of a specific residue (Ser24) present in the K8 head domain (Loranger et al. 2006). Such an observation points to a key role for K8 in modulating desmoplakin deposition at desmosomal microdomains, most likely as a step for the coordinate involvement of IFs–desmosomes in providing an efficient mechanically resilient network across a simple epithelial cell layer. This also provides direct evidence for the importance of K8/K18 IFs in the modulation of simple epithelial cell polarity.

Keratin IFs and Hemidesmosomes or FAs In stratified squamous epithelial cells such as keratinocytes, plectin is targeted to typical polarity-dependent microdomains involved in cell-extracellular matrix (ECM) adhesion known as hemidesmosomes (Fuchs et al., 1999; Wiche 1998b). There, plectin acts as a linker between keratin IFs and a specific set of integrins, α6β4, which in turn constitute

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Figure 16.3. Schematic representation of (A) a desmosome; (B) a focal adhesion complex; (C) a hemidesmosome; and (D) FasR compartmentalization.

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transmembrane receptors for ECM elements such as laminin, which is mainly produced by the dermis (Fig. 16.3B). However, in simple epithelial cells such as hepatocytes, there are no hemidesmosomes and the integrin-mediated transmembrane junctions to ECM are provided by FAs (Fig. 16.3C), that is, sites equivalent to the focal contacts seen in monolayer culture between adherent fibroblastic cells and ECM elements such as fibronectin (Schoenwaelder and Burridge 1999). A variety of actin-binding proteins have been identified (e.g., vinculin, talin, actinin) at the cytoplasmic side of FAs, along with signaling proteins such as FA kinase (FAK), integrin-linked kinase (ILK), paxillin, PKC, and c-Src (Schoenwaelder and Burridge 1999; Fig. 16.3C; see also Chapters 12 and 17). These proteins are assembled at integrin/FA sites according to a hierarchy of interactions that seems to vary with cell type (Burridge et al. 1997; Danen et al. 1998). Actin also combines with myosin-II to make the actomyosin network (e.g., stress fibers), which governs actinbinding assembly at integrin sites as well as cell contractility and cell shape (Burridge et al. 1997). Notably, our recent work has identified the novel PKCδ as a mediator of the K8/K18 modulation of hepatic cell adhesion and migration. Actually, a K8/K18 loss results in a time-course modulation of the RACK-1, β1-integrin, plectin, PKC, and c-Src complex formation, and a reduced FAK time-residency at FAs. On the whole, these results uncover a key regulatory function for K8/K18 IFs in the adhesion and migration of simple epithelial cells, through a dynamic interplay taking place at FA microdomains. Remarkably, cell migration is critically dependent on the ability of cells to sense and adapt to mechanical stresses applied on ECM–integrin–actin cytoskeleton connections at the surface membrane. In these terms, integrins behave as transmembrane mechanical sensors and actin MF-linked transducers at FAs (Ingber 2006). In addition, microdisplacement of the surface membrane resulting from the cellular response to mechanical stress applied on integrins at FAs has been demonstrated to reflect a localized stiffness (Matthews et al. 2004). With this background in mind, we have recently assessed the involvement of K8/K18 IFs in the simple epithelial cell response to a tangent mechanical stress (Bordeleau et al. 2008). To that end, we used monolayer cultures of K8/K18-lacking liver cells and their normal counterparts. The stress was then generated with a laser tweezers-mediated force applied on fibronectin-coated microbeads attached to integrins α5β1; in this laser setup, the displacement reflected the cell reaction to the photonic force acting on the bead. Overall, the results revealed a significant contribution of K8/K18 IFs to the cellular mechanical stress response at FA microdomains in simple epithelial cells.

Keratin IFs and Fas Receptor Conventional apoptosis is initiated through stimulation of death receptors by their respective ligands. The central execution machinery is evolutionarily conserved and is mostly based on caspase signaling (Kumar 1999; Zheng and Flavell 1999). Remarkably, two death-signaling pathways can be activated depending on cell type, namely mitochondriaindependent type I cells and mitochondria-dependent type II cells (Park and Peter 2008). In type I cells, Fas stimulation leads to its internalization and the recruitment of Fasassociated protein with death domain (FADD) allowing the recruitment of other deathinducing signaling complex (DISC) components (Park and Peter 2008) (Fig. 16.3D). Then, the DISC triggers procaspase-8 autoproteolytic activation, which is followed by a rapid cleavage of executioner procaspase and cleavage of various downstream substrates (Kumar 1999; Zheng and Flavell 1999). Type II cells produce little active caspase 8 at the DISC, and this correlates with minimal Fas internalization (Park and Peter 2008). This, however, is sufficient for the cleavage of the Bcl-2 family member Bid, causing its translocation to mitochondria, where it induces a loss of the mitochondrial transmembrane potential (ΔψM)

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and a release of death-promoting factors such as cytochrome c (Park and Peter 2008). In the cytoplasm, cytochrome c binds to Apaf1, leading to procaspase-9 recruitment to form the apoptosome, which, in turn, activates executioner procaspases and leads to cell death. With this background in mind, we addressed the K8/K18 IF modulation of Fas-induced apoptosis in mouse hepatocytes and found that K8-null hepatocytes are more sensitive than wild-type hepatocytes to FasR stimulation (Gilbert et al. 2001), with a type II to type I switch in FasR-activated death signaling as result of the K8/K18 IF loss (Gilbert et al. 2008). In line with recent reports by others (Chaigne-Delalande et al. 2008; Park and Peter 2008), these observations provided the first evidence indicating that the difference in type I and type II FasR-activated signaling is largely linked to the extent of DISC formation. Actually, it appears that ligand-dependent conformational alterations and multimerization of homotrimeric FasR, known as signaling protein oligomeric transduction structures (SPOTS; Siegel et al. 2004), constitute the first steps required to recruit the proteins essential to forming the DISC. This early FasR structural step triggers DISC formation, in association with procaspase-8 activation. Thereafter, in type I cells, these segregated FasR structures become “caps,” which in turn colocalize with lipid rafts, and internalize to amplify the procaspase-8 activation (Schutze et al. 2008). Of note, in type II cells, while lipid rafts appear to be involved, the FasR internalization is not apparent (Fig. 16.3D). In line with these propositions, our unpublished results indicate that the K8/K18 IF loss leading to the switch from type II to type I death signaling in hepatocytes alters the initial FasR activation through perturbation of lipid raft constituent distribution and organization. This integrative interplay, which takes place at FasR microdomains, adds an extended dimension to the multifunctional traits of K8/K18 IFs at surface microdomains in simple epithelial cells.

Keratin IFs and Epithelial Cell Asymmetry The sorting mechanisms responsible for the targeting of surface membrane proteins to apical and basolateral domains in single-layered (simple) epithelial cells, such as those of the intestine, have been extensively studied (Oriolo et al. 2007; see also Chapter 21 on epithelial domains as well as Chapter 13 on microvilli). Central to the acquisition of such cell asymmetry is the cytoskeleton and its associated proteins. For instance, a recent study has revealed an involvement of ezrin as a linker between cortical actin and membrane proteins at the apical domain, through local PKC-dependent activation. In these cells, keratin IFs are also cortical, either apically or apico-lateral, and previous work in K8-null enterocytes where K7, a type II keratin redundant to K8, is not expressed, has shown a loss of apical membrane polarity. Although such a phenotype is striking, the underlying mechanistic explanations remain unclear. One possibility is that this alteration occurs through an effect on the subcellular localization of microtubule organizing centers (MTOCs) and the polarized dynamics of MTs in these intestinal epithelial cells (Oriolo et al. 2007; see also Chapters 8, 14, 15, and 22 on the MTOC and cell polarity).

RELEVANCE TO HUMAN DISEASES The discovery that keratin gene mutations in keratinocytes cause several diseases has provided definitive evidence for the structural functions of epidermal keratins. This is the case for epidermolysis bullosa simplex (EBS), where a K14-point mutation induces a profound disorganization of K5/K14 IFs in the basal keratinocytes, which in turn leads to

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cytolysis and blistering of the skin (Coulombe and Wong 2004). There is also ample evidence indicating that K8/K18 instead constitutes key susceptibility genes for end-stage liver diseases in humans (Ku et al. 2003a). Omary’s group was the first to detect K8 and K18 mutations in the human genome, which exhibit heterozygocity (Ku et al. 1997). Recent analyses from the same group have led to the identification of 10 additional heterozygote mutations (e.g., K18 deletion, K8 frameshift, missense K8/K18 alterations, and several new polymorphisms), although the mutation frequency varies with population and stage of liver disease (Ku et al. 2003a; Hesse et al. 2004). Overexpression of a K8 or K18 mutant in transgenic mouse liver predisposes to Fas-induced apoptosis (Ku et al. 2003b). Actually, mutations in the genes encoding K8, K18, and K19 appear to serve as liverdisease modifiers (Omary et al. 2009). Notably, disease-causing or disease-associated mutations have been described in most IF genes of the six classes (Oshima 2007), which means that the resulting IF-associated diseases (IF-pathies) involve a broad range of tissues and reflect the tissue and cell-type expression of IF genes (Omary 2009).

UNCERTAINTIES AND CONTROVERSIES Obviously, much of the experimental evidence accumulated so far on the multifunctional features of keratin IFs related to surface microdomains has come from work performed on simple epithelial cells, particularly hepatocytes and hepatoma cells. As stated in a previous section, the advantage of using these hepatic cell types is that they contain only K8 and K18, so that a knockout (or knockdown) of K8 leads to epithelial cells lacking IFs. On this ground, the contribution of K8/K18 IFs to the different cellular activities can be directly assessed by comparing wild-type with K8-null cells. However, the presence of additional keratins, which likely reflects some functional redundancy, in other simple cell types such as those lining the intestine or forming the ductular network in liver and pancreas (Marceau and Loranger 1995; Toivola et al. 2000) raises relevant issues with regard to the relative contribution of the different keratin pairs to simple epithelial cell polarity, and their interplay with membrane microdomains in these cell models. By extension, similar uncertainties on the possible interplay between keratins and membrane microdomains can be raised with regard to postnatal epidermis, where the keratin pairs vary from K5/K14 to K1/K10 during keratinocyte terminal differentiation (Coulombe and Omary 2002), and to other highly specialized tissues such as the mammary glands, where the basal cells express the stratified keratin pair K5/K14 and the luminal cells contain the simple keratins K7, K8, K18, and K19 (Oshima et al. 1996). Controversial data over the last 20 years claim that a K8 ectoplasmic domain can be detected on the cell surface of some cells, including hepatocytes, hepatoma cells and breast cancer cells (Gonias et al. 2001). While relevant doubts have been raised about the reliability of the assays originally used (Riopel et al. 1993), this controversial issue has been revisited lately by using a monoclonal antibody that specifically recognizes an epitope sequence of the ectoplasmic domain unique to K8 (Obermajer et al. 2009). In addition, the antibody has been shown to compete with the urokinase-type plasminogen activator (uPA), thus suggesting that the ectoplasmic domain functions as a plasminogen activator receptor (uPAR; Obermajer et al. 2009). These observations can be linked to reports suggesting that cell surface uPA, bound to a uPAR, activates plasminogen to plasmin, which in turn promotes ECM degradation, tumor progression, and metastasis. If such findings can be confirmed, it would mean that the complex made of K8, uPA, and plasminogen acts at a surface membrane domain as a modulator of tumor cell adhesion/ migration.

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FUTURE PERSPECTIVES So far, the relevance of the link between IFs and surface microdomains has been derived from work performed mainly on polarized epithelial cells. Still, a polarized nonepithelial cell type of particular interest is the endothelial cell. In this cell type forming a monolayer lining the inner face of blood vessels, key microdomains are provided by the integrinforming FAs and the V-cadherin-containing adherent junctions (Czirok et al. 2008). Notably, recent work has revealed that vimentin IFs associate with FAs in endothelial cells, and their recruitment at this surface domain is tightly regulated and modulates the strength of cell adhesion to the ECM substratum (Bhattacharya et al. 2009). Actually, this mirrors what is known about the function of the keratin IF linkage to integrins at hemidesmosomes in epidermal keratinocytes during skin formation. In the case of endothelial cells, it would be of great interest to determine the mechanisms by which the vimentin-dependent modulation of cell adhesion strength occurs during angiogenesis, where precise control of adhesion appears to be central to blood vessel development and maturation. An additional issue of interest here concerns the adhesive interactions of leukocytes with endothelial cells during their migration across the endothelium (diapedesis), that is, migration via a transcellular route, where vimentin IFs of both endothelial cells and leukocytes form a dynamic anchoring structure at the site of contact between the two cell types (Nieminen et al. 2006). Among the issues that need to be addressed is the role of vimentin IFs in the formation of the anchoring and/or organizing structures for adhesion molecules on the surface of both cell types. In addition, a similar phenomenon appears to take place when invasive carcinoma cells, for example, colon cancer cells surviving in the blood circulation, also cross the vessel wall through transcellular diapedesis to invade the surrounding tissue, thus mimicking leukocyte extravasation (Tremblay et al. 2008). Since colon cancer cells express keratins, and endothelial cells express vimentin, the possible involvement of these two distinctive IFs in mediating the interplay between the cell types at the site of contact constitutes a challenging and interesting question. An intriguing issue with regard to the functions of IFs in surface microdomain dynamics concerns the relative contribution of keratins, essentially K8/K18 and K19, versus vimentin in epithelial–mesenchymal transition (EMT) during normal embryo development and carcinoma cell invasion and metastasis (Eastham et al. 2007). Although the loss of cell surface E-cadherin is considered a prerequisite for EMT, many other cellular events are required to impart increased cell migration and invasion, for instance, a proper spatiotemporal expression, interaction, and arrangement of relevant intracellular and extracellular factors (Eastham et al. 2007). In line with the evidence indicating a likely involvement of the cytoskeleton in these EMT events, it is worth noting that under both embryonic and cancerous tissue remodeling conditions, the cell IF status changes from an K8/K18-rich network connected to desmosomes to a vimentin-rich network connected to FAs (Kokkinos et al. 2007). So, considering that patient mortality following the metastatic spread of carcinoma cells is significantly increased, and these account for at least 80% of all cancers, an in-depth investigation of the keratin versus vimentin contribution to surface microdomain fate in these invasive cells would help to better understand cancer metastasis and to potentially design novel therapeutic approaches for inhibiting this fatal complication of cancer.

ACKNOWLEDGMENTS The original work from our laboratory was supported by grants from the Canadian Institute of Health Research, Cancer Research Society, and National Sciences and Engineering Research Council of Canada.

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ABBREVIATIONS CDK1 DISC ECM FA FADD FAK GFAP ILK Ks

cyclin-dependent kinase 1 death-inducing signaling complex extracellular matrix focal adhesion Fas-associated protein with death domain focal adhesion kinase glial fibrillary acidic protein integrin-linked kinase keratins

IF IF-pathies MF MT MTOC PKC RACK1 SPOTS

intermediate filament IF-associated diseases microfilament microtubule microtubule organizing centers protein kinase C receptor for activated C kinase 1 signaling protein oligomeric transduction structures

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PART

IV

ADHESIVE AND COMMUNICATING DOMAINS J

UNC T I O NS AND ADH E SIO N S are cellular domains that regulate the interaction of the cell with its environment and also provide the structural attachments for the cytoskeleton-based cytoarchitecture. Focal adhesions (Chapter 17) mediate integrin-dependent adhesive interactions of the cell with the underlying substrate or extracellular matrix and are intimately associated with actin stress fibers (Chapter 12). Focal adhesions are signaling domains whose turnover is critical to cell migration (Chapters 12 and 17) and also movement of neuronal growth cones (Chapter 22). Cadherin-based adherens junctions (Chapter 18) together with tight junctions (Chapter 19) are actin-associated cell–cell junctions that represent key structures for the development of polarized epithelial domains (Chapter 21). Other epithelial junctions include the desmosomes (Chapter 19), also cadherin based, that are stabilized by interaction with keratin intermediate filaments (Chapter 16). Cadherin-based interactions generate polarity cues that lead to the formation of the exocyst (Chapter 18) and the targeted microtubule– Golgi trafficking that is critical to the establishment of polarized cellular domains in various cell types, including epithelial and neuronal cells (Chapters 8, 14, 21, and 22). The gap junction is distinct from adhesive junctions in that it forms communicating channels that enable exchange of small molecules between adjacent cells (Chapter 20).

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FOCAL ADHESIONS Caitlin Tolbert Keith Burridge

DEFINITION For 100 years, tissue culture has provided a powerful tool for studying the structure and behavior of cells. In order to grow in culture, it is necessary for many cells to adhere to the underlying substratum. Adhesion is usually mediated by cell adhesion molecules of the integrin family binding to extracellular matrix (ECM) molecules, such as fibronectin (Fn) or vitronectin (Vn), that adsorb onto the plastic or glass substratum. For many cells in culture, the integrins mediating adhesion are not uniformly distributed over the cell surface but rather are concentrated in discrete domains, known as focal adhesions (FAs) and focal complexes (FCXs). These domains serve multiple functions including providing sites of adhesion to the surface on which the cells are growing, anchoring bundles of actin filaments and transmitting tension generated within these bundles to the substratum. In addition, these domains contain many signaling proteins that participate in signaling pathways that affect the growth and behavior of cells. “Focal adhesion,” “focal contact,” and “adhesion plaque” are all terms that have been used to describe the prominent adhesions made by cells like fibroblasts to ECMcoated substrata as discussed in the historical perspective section. We will use the term FA for these large, relatively stable adhesions, which are characterized by being dependent on the activity of the GTPase RhoA (see below). Smaller, less stable adhesive structures have also been identified and are referred to as “focal complexes” (Nobes and Hall 1995) (Fig. 17.1). Typically, these are regulated by Rac1 and Cdc42 activity, and they mature into FAs in a RhoA-dependent manner. In this brief review, we will discuss the formation of FAs and FCXs, some of their key constituents, the signaling pathways that emanate from these structures, and their dynamics. However, for more details, the reader is advised to go to other recent reviews (Webb et al. 2003; Romer et al. 2006; Zaidel-Bar et al. 2007; Dubash et al. 2009).

HISTORICAL PERSPECTIVE The structures that we recognize today as FAs were probably first observed by Abercrombie and his colleagues who were using electron microscopy to study fibroblasts growing in culture (Abercrombie et al. 1971). They identified dense plaques on the ventral surface of these cells that came into close apposition with the underlying substratum. Bundles of filaments could be seen inserting into these structures, and it was speculated

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(A)

(B)

(C)

(D)

Figure 17.1. Types of ECM adhesions. (A) Focal adhesions with (B) stress fibers and (C) focal complexes with (D) small actin filaments were visualized by staining rat embryo fibroblasts with paxillin (green) and actin (red). (C, D) Cells expressing constitutively active Rac1 and treated with Y27632, an inhibitor of Rho kinase. Scale bar 5 μm.

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that the filaments might be part of the contractile system of the cell and that the plaques were likely involved in providing adhesion. These structures became known as adhesion plaques. Shortly after this, immunofluorescence microscopy began to be applied to look at the cytoskeletal organization of cultured cells. Suddenly, by staining with an antibody against actin, stress fibers in thousands of cells could be visualized on a single coverslip, whereas previously electron microscopy had limited analysis to a few cells or even to a few regions of cells (Lazarides and Weber 1974). At the time, this represented a significant breakthrough. There was a rush to visualize many of the components that earlier had been identified in striated muscle but which had not been considered in nonmuscle cells. In this context, antibodies were generated against α-actinin, a protein of muscle Z-lines, and staining of fibroblasts showed that α-actinin is distributed along stress fibers with a periodic pattern (Lazarides and Burridge 1975). However, the initial localization of α-actinin also revealed that it was concentrated at the ends of stress fibers in regions referred to as “patches.” In this location, it was suggested that α-actinin might participate in linking the actin filaments of stress fibers to the plasma membrane (PM) (Lazarides and Burridge 1975). The authors speculated that these patches where α-actinin is enriched might correspond to the plaques seen by Abercrombie and coworkers, and subsequent work did indeed confirm this. At about the same time, the light microscopy technique of interference reflection was used to visualize the areas of closest contact between cells and the substratum. In two parallel studies, it was confirmed that adhesion plaques are areas where cells come very close to the substratum (Abercrombie and Dunn 1975), and these areas of tightest adhesion were also given the name focal contacts (Izzard and Lochner 1976). The discovery of vinculin, which, unlike α-actinin, is concentrated in the adhesions and not seen along the length of stress fibers, gave the field considerable momentum (Geiger 1979; Burridge and Feramisco 1980). There was a search for proteins that would span the membrane at these sites of adhesion that could serve to link the ECM on the outside with the cytoskeleton at the cytoplasmic face. Research from several different groups in the early to mid-1980s led to the discovery of integrins and showed they are major constituents of FAs (reviewed in Hynes 1992). Indeed, FAs can be considered as sites where integrins are clustered to form a scaffold on which many other proteins assemble. Not only are they sites of adhesion where the cytoskeleton is linked across the PM to the ECM, but they are major regions of signal transduction. Many different signaling pathways as well as signaling components have been identified in FAs and are further described in this chapter.

ASSEMBLY OF FCXS AND FAS Two different mechanisms for FA assembly have been described. Hotulainen and Lappalainen studied the formation of both stress fibers and FAs in motile cells (Hotulainen and Lappalainen 2006). FAs were observed to form close to the edge of cells and give rise to “dorsal stress fibers,” bundles of actin filaments polymerizing in an mDia-dependent fashion (see also Chapter 12 on the actin cytoskeleton). Individual dorsal stress fibers growing by polymerization extended from FAs back toward the cell body. It was observed that these dorsal stress fibers would link up either with other dorsal stress fibers growing from the other side of the cell or with “arcs,” rearward moving bundles of actin filaments that form parallel to the leading edge of cells. The fusion of these structures was observed to generate the more typically described ventral stress fibers, which are anchored at each end by FAs (Hotulainen and Lappalainen 2006). This mode of assembly of stress fibers

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and FAs, driven by actin polymerization, is probably the common mechanism as cells migrate or as they spread on an ECM matrix. A different model for studying FA assembly has used cells that are not migrating but which have been serum starved to decrease RhoA activity and disassemble FAs (Ridley and Hall 1992). Under these conditions, activation of RhoA by agents such as lysophosphatidic acid (LPA), or expression of constitutively active RhoA mutants, stimulates the rapid assembly of FAs and stress fibers. The mechanism for this assembly was shown to depend less on actin polymerization (Machesky and Hall 1997) (although some polymerization occurs) and more to be due to RhoA-driven myosin II contractility (ChrzanowskaWodnicka and Burridge 1996). FAs and FCXs are essential domains of the PM composed of tightly clustered integrins, thereby forming a scaffold to which many other structural and signaling proteins are recruited. The key question when considering assembly of these domains is how the clustering is induced. ECM is essential for the assembly of FAs and FCXs (Hotchin and Hall 1995), and it can contribute to small-scale clustering because of its multivalent nature. However, in order to form the larger clusters that make up FAs, the clustering of integrins is driven by forces within cells derived from external stimuli that activate RhoA. Integrin clustering has been more effectively studied in the model system using serum-starved quiescent cells stimulated by agents that activate RhoA. Here, the clustering was shown to be induced by the contractile activity of actomyosin (Chrzanowska-Wodnicka and Burridge 1996). Little work has been performed on FCX formation, but we suspect that a major factor inducing the clustering of integrins is the bundling of actin filaments. Possibly, the main role of Rac and Cdc42 is to promote actin polymerization at sites where integrins are engaging ECM. Since integrins engaged to their ECM ligands are tethered to actin filaments, when the filaments are brought together by cross-linking proteins, this will cluster their associated integrins, giving rise to the FCXs. However, FCXs were also shown to be dependent on myosin activity, although not downstream of RhoA (Rottner et al. 1999). There is a complex interplay between ECM adhesion and Rho activity. Initial adhesion or clustering decreases RhoA activity (Ren et al. 1999; Arthur et al. 2000), but longer adhesion activates RhoA (Cox et al. 2001). This biphasic transition of decreased RhoA activity followed by increased activity likely occurs to allow initial cell spreading prior to formation of mature FAs. Following RhoA activation, downstream effectors of RhoA, specifically Rho kinase (ROCK), control myosin-dependent contractility. ROCK causes an increase in myosin light chain (MLC) phosphorylation by directly phosphorylating MLC and by inactivating the MLC phosphatase. MLC phosphorylation activates myosin’s ATPase activity, generating force on actin filaments. Additionally, this phosphorylation promotes the assembly of myosin II bipolar filaments. The multivalent myosin filaments are efficient at cross-linking and thus bundling actin filaments, giving rise to stress fibers (Chrzanowska-Wodnicka and Burridge 1996; Pellegrin and Mellor 2007).The tension generated within stress fibers, as well as the bundling, leads to increased integrin clustering and thus formation of larger FAs (Chrzanowska-Wodnicka and Burridge 1996; Dubash et al. 2009). As RhoA activity increases over time, this is paralleled by the maturation of the FCXs into the larger aggregates, the FAs. The coupling of tension from the cytoskeleton and ROCK activity provides a positive feedback loop mechanism in order to sustain and amplify ROCK activity, which creates more cytoskeletal tension and continuing the feedback mechanism (Bhadriraju et al. 2007). The recruitment of cytoskeletal proteins, including talin and vinculin, establishes linkage between integrins and actin (Fig. 17.2). FCXs can either mature to an FA or be turned over (Geiger et al. 2001; Galbraith et al. 2002). A characteristic of FCXs maturing

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β α

-Integrin Activation --Talin --Kindlin

-Tension in ECM -Ligand Engagement of Integrins

Vinculin Arp2/3

F-actin

FAK

Src

Focal Complex

Talin

Initial Actin Formation

Figure 17.2. A model of integrin-mediated FA assembly. PM, plasma membrane.

e.g.

Outside-In Signaling Event

e.g.

Inside-Out Signaling Event

Syn4

Paxillin Tensin Zyxin

VASP α-Actinin Myosin II

Focal Adhesion

ECM

PM

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into FAs is the recruitment of vinculin to these sites of adhesion in response to tension generated by myosin II (Galbraith et al. 2002). However, in order to establish the link between integrins and actin, talin is necessary and is the initial protein recruited to adhesions (Lee et al. 2007). It has been shown that talin is recruited to integrin clusters, even in the absence of F-actin, in the presence of phosphatidylinositol 4,5-biphosphate (PIP2) (Puklin-Faucher and Sheetz 2009). Following integrin clustering, scaffold proteins (such as paxillin) are recruited in order to relocalize multiple substrates including focal adhesion kinase (FAK). Furthermore, kinases (such as FAK or Src) are activated and further transduce the adhesion signal. Multiple proteins found at FAs are tyrosine phosphorylated and FAs display the highest level of tyrosine phosphorylation in the cell. Indeed, staining with antibodies against phosphotyrosine effectively reveals FAs by immunofluorescence. Not surprisingly, tyrosine phosphorylation is a key regulator of many FA proteins. For instance, FAK and Src family kinases (SFKs) phosphorylate several proteins, including paxillin and Crk-associated substrate (p130Cas). The phosphorylation sites on these adapter proteins, in turn, create sites for binding Crk, an adapter that couples to Rac1 activation via the DOCK180/ELMO exchange factor complex (Kiyokawa et al. 1998; Ren et al. 2000; Brugnera et al. 2002; Chodniewicz and Klemke 2004). FAK and Src can directly contribute to the biphasic pattern of RhoA activity following cell adhesion to Fn. For example, both Src and FAK have been implicated in the inactivation of RhoA by promoting p190RhoGAP activity, a RhoA-specific GTPase-activating protein (GAP) (Arthur et al. 2000; Ren et al. 2000; Bass et al. 2008). However, there are several RhoA guanine nucleotide exchange factors (GEFs) that are activated by tyrosine phosphorylation, and specifically, FAK has been shown to be involved (Dubash et al. 2007; Pellegrin and Mellor 2007; Lim et al. 2008).

FA COMPLEXITY A mature FA is a complex structure with over 100 proteins having been identified in these regions. Some of these proteins play a predominantly structural role, whereas most are involved in signaling cascades. Most FA proteins interact with multiple binding partners, and many are also sensitive to conformational changes and phosphorylation states that affect their activity. While there are many proteins found in FAs, here we will highlight a few that play a significant role in this domain (see Table 17.1). For recent works demonstrating FA complexity, see Kanchanawong et al. (2010).

Integrins Integrins are heterodimers composed of α- and β-subunits that undergo allosteric changes upon ligand binding (Hynes 2002). The particular combination of the α- and β-subunits dictates ligand specificity, and to date, there are 18 α-subunits and 8 β-subunits that can generate 24 combinations in humans (Humphries et al. 2006). Much of the work on integrin structure and affinity regulation has been performed on the major integrin found in platelets, αIIbβ3, or on the β2 integrins found in leukocytes. These integrins are involved respectively in platelet aggregation during blood clot formation and in the recruitment of leukocytes out of the circulation to the endothelium during inflammation. In both cases, it is particularly important that these integrins be maintained in the low affinity state until appropriate signals activate the integrins promoting their high affinity state. Structural studies have revealed that these integrins adopt a bent-over conformation in the low affinity state that straightens out upon activation. There has been considerable interest on how

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Structural Structural

Structural

Adaptor/structural Structural

Structural

Structural

Structural Protease

Lipid

Serine/threonine kinase Tyrosine kinase

Tyrosine kinase

α-Actinin Filamin

Kindlin

Palladin Talin

Tensin

VASP

Vinculin Calpain

PIP2

PKC FAK

Src

+ + +

+ +

+ +/−

+ + +

* +

+

−

+

*

*

+

+ + +/−

*

+

+ + +/− −

Migration

* −

+

+

FA Turnover

+ −

+

+ +

FA Assembly

Proliferation, ROS generation* Mechanotransduction, proliferation, activating Rho GTPases, anti-anoikis Mechanotransduction, proliferation

Cytoskeletal reorganization

Promote actin polymerization, membrane rigidity Mechanotransduction, apoptosis Perturbs Rho GTPases

Actin bundling, mechanotransduction* Inhibit integrin activation, membrane stability, proliferation Differentiation*, proliferation*, regulate integrin affinity, cell survival* Differentiation, podosomes, tissue remodeling Mechanotransduction, regulate integrin affinity Actin capping

Activating Rho GTPases Proliferation Activating Rho GTPases Differentiation, mechanotransduction

Other Roles

Carisey and Ballestrem (2010) Kulkarni et al. (2002); Perrin and Huttenlocher (2002) Ling et al. (2006); Sheetz et al. (2006) Wu et al. (2008) Mitra et al. (2005); Wozniak et al. (2004) Playford and Schaller (2004); Wozniak et al. (2004)

Le Clainche and Carlier (2008); Lo (2004, 2006) Galler et al. (2006)

Goicoechea et al. (2008) Wozniak et al. (2004)

Wozniak et al. (2004) Wozniak et al. (2004) Wozniak et al. (2004) Hirata et al. (2008); Zaidel-Bar et al. (2004) Le Clainche and Carlier (2008) Harburger and Calderwood (2009); Sheetz et al. (2006) Lo (2006); Meves et al. (2009)

References

PKC, protein kinase C.

*, role is implicated but has not been demonstrated.

+, a positive regulator; –, a negative regulator; +/−, plays both a positive and negative role;

Shown here are cytoplasmic components known to play a role in FAs and their role in this dynamic domain. Highlighted are papers or reviews that provide greater detail on their function that is presented.

Adaptor Adaptor Adaptor Adaptor

Type

Cytosolic Composition of Focal Adhesions

Crk P130Cas Paxillin Zyxin

TABLE 17.1.

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these changes in the extracellular domains are induced by events occurring within the cytoplasm. In the resting state, the single transmembrane domains of the α- and β-subunits as well as their cytoplasmic domains appear to be closely associated, and activation induces a separation of these chains (Qin et al. 2004). Activation of integrins is mediated by binding of proteins such as talin to the integrin cytoplasmic domains (Calderwood 2004). Activation of the integrins involved in FCXs and FAs, specifically integrins α5β1 and αvβ3, is also regulated, although this has been less well characterized than for the platelet and leukocyte integrins. Activation as a result of binding to proteins to the cytoplasmic domains is known as “inside-out” signaling, whereas the signals transmitted via the integrins to the cell in response to their binding to extracellular ligands is known as “outside-in” signaling (Fig. 17.2). These signaling cascades have been reviewed (Hynes 2002; Liddington and Ginsberg 2002; Qin et al. 2004).

Syndecan-4 (Syn4) Integrins are not the only transmembrane proteins in FAs. Syn4, a heparan sulfate proteoglycan, has been found in the FAs of many cells and plays an important role in FA maturation (Woods and Couchman 2001). Syn4 binds to the heparin-binding domains (HBD) on Fn. The significance of Syn4 to FA assembly was demonstrated by multiple studies in which plating cells on the cell-binding domain (CBD) of Fn (the major integrin-binding site) were insufficient for FA assembly. Additionally, the presence of an HBD in Fn was required (Izzard et al. 1986; Woods et al. 1986). The discovery that Syn4 is present in the FAs of some cells and that it binds this HBD on Fn thrust Syn4 into the FA limelight. Members of the syndecan family share conserved sequences in the transmembrane and cytoplasmic domains while sulfated glycosaminoglycan chains attach to the extracellular region that enables binding to ECM ligands (Woods and Couchman 2001; Okina et al. 2009). While there are several syndecans in each cell type, Syn4 is the only member consistently present in FAs in mammals (Woods and Couchman 2001). Although integrins have a primary role in cell adhesion and spreading, Syn4 plays more of a role in FA maturation (Bloom et al. 1999). Recently, it has been shown that Syn4 aids in forming stable FAs by contributing to the recruitment of vinculin (Bloom et al. 1999; Bass et al. 2007a). Although vinculin has not been shown to directly bind Syn4, this indirect interaction may be mediated by the direct interaction of Syn4 with α-actinin, an actin-binding protein that contributes to actin bundling (Greene et al. 2003). Further evidence for the role of Syn4 in FAs comes from overexpression studies that yield cells with an increase in FA formation and limited cell motility (Couchman 2003). A possible explanation for the role of Syn4 in promoting FA assembly is that Syn4 may enhance RhoA activation and contribute to FA assembly when cells are plated on Fn CBDs that are spaced at too low a density to activate RhoA (Wang et al. 2005). The C-terminal tail of Syn4’s cytoplasmic domain contains a PDZ-domain (post-synaptic density 95, discs-large, zonula occludens-1) consensus binding sequence. One PDZdomain-containing protein that binds to this sequence is synectin, which has been shown to bind to the RhoA GEF Syx1 (Liu and Horowitz 2006), suggesting a direct pathway by which Syn4 may stimulate RhoA activation and evidence for RhoA activation downstream of Syn4 has been presented previously (Dovas et al. 2006). However, other experiments indicate that Syn4 regulates Rac1 activity and that cells lacking Syn4 are not able to activate Rac1 when plated on Fn (Bass et al. 2007b). Mice that are deficient in Syn4 survive but have defects in wound healing and angiogenesis (Ishiguro et al. 2000; Echtermeyer et al. 2001). Interestingly, cells that are deficient in Syn4 are still able to form FAs when plated on full-length Fn, but unlike wild-type fibroblasts when plated on the Fn CBD and

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supplemented with soluble HBD, they do not develop FAs. It is clear that more work needs to be performed on Syn4 to clarify its various functions and activities in FAs.

Cytoskeletal Proteins Talin Talin is a large structural protein with multiple functions within FAs. Binding of the talin head to β-integrin cytoplasmic domains activates the integrin, inducing a conformational change such that the integrin goes from a low-affinity to a high-affinity ligand-binding state. Talin is also a key link between integrins and the actin cytoskeleton, bridging between the integrin cytoplasmic domains and actin directly, and also indirectly via other actin-binding proteins such as vinculin (Critchley 2009). Talin consists of a globular N-terminal head and a flexible C-terminal rod domain that is made up of a series of helical bundles, with the final helical region generating a dimerization domain. How much of talin is monomeric and how much dimeric within FAs and FCXs remains to be determined. As a monomer, talin exists in a folded conformation so that the C-terminus binds to the head, blocking the integrin-binding site. This inactive state is relieved by binding PIP2 (Goksoy et al. 2008). The head region is enriched in basic residues that can interact with negatively charged residues such as the head group of PIP2 and regions of FAK as well as integrin subunits αIIb, β1, and β3 (Isenberg and Goldmann 1998; Nayal et al. 2004). The rod domain of talin binds actin and vinculin (Critchley 2009). Due to its multiple binding sites for its binding partners (integrins, actin, and vinculin) and its dimerization ability, talin is an effective cross-linker and is likely to be important for integrin clustering and stabilization of FCXs (Fig. 17.2). With the vinculin-binding sites on talin, evidence has been presented that some of these are normally buried within the molecule but open upon application of force as would occur during FA formation (Ziegler et al. 2008; del Rio et al. 2009). It is well established that mechanical force contributes to the growth and maturation of FCXs into FAs (Riveline et al. 2001). In part, this may be due to the tension leading to more integrins being drawn into the structure, but the exposure of additional binding sites, such as those on talin for vinculin is almost certainly an important part of this tensioninduced maturation (del Rio et al. 2009). Talin’s importance in FA assembly is shown through studies in which talin null cells display impaired FA assembly and fail to recruit vinculin in response to tension (Zhang et al. 2008). Talin binds one isoform of PIP kinase, PIPK1γ90, targeting this enzyme to FAs (Ling et al. 2003). The product of PIPK1γ90 is PIP2. PIP2 has many effects expected to enhance FCX and FA assembly. For example, it induces a conformational change in talin, exposing the integrin-binding site on the talin head so that it will activate integrins (Goksoy et al. 2008). It also induces a change in conformation of vinculin, exposing binding sites for multiple partners (Johnson and Craig 1995; Gilmore and Burridge 1996). PIP2 also promotes the activities of many actin-binding proteins (Ling et al. 2006; Sheetz et al. 2006). Consequently, the recruitment of PIPK1γ90 to sites of adhesion and the resulting local synthesis of PIP2 is likely to enhance the assembly of these structures. Interestingly, PIPK1γ90 binds to the head of talin and competes with the integrin β cytoplasmic domain for binding. Overexpression of this kinase disrupts FAs, probably due to the displacement of talin from integrins. However, under normal conditions, we suspect that the stoichiometry of PIPK1γ90 relative to talin is low. This should allow some talin molecules to bind to integrins while others bind PIPK1γ90. Given the potential for talin to dimerize it is, of course, conceivable that a talin dimer could simultaneously bind to both molecules. Regardless, the interaction of talin with PIPK1γ90 and the resulting local synthesis of PIP2 is probably one of several ways that talin contributes to the assembly of FCXs and FAs.

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Vinculin Vinculin is a ubiquitously expressed structural adaptor protein that when active not only binds actin but also transmits tension. According to mouse knockout studies, vinculin null cells adhere but have difficulties spreading and migrate more rapidly. The vinculin null cells are resistant to apoptosis (Xu et al. 1998). In addition, vinculin is downregulated in many tumors leading to smaller and fewer FAs, while overexpression has been found to suppress tumorigenicity (Rodriguez Fernandez et al. 1992). The full role of vinculin remains elusive. However, insights into the role of vinculin are emerging from structural studies that reveal differences in regulation depending on vinculin’s binding partners. Vinculin activation is dependent on its structural conformation. When inactive, vinculin is auto-inhibited by intramolecular interactions of the N-terminal head domain and C-terminal tail domain. Upon release of the auto-inhibition, vinculin adopts an open conformation. In this state, it acts as a scaffold, binding a number of proteins including talin and α-actinin to the head region; Arp 2/3, vinexin, and vasodilator stimulated phosphoprotein (VASP) to the hinge region; and F-actin, paxillin, and PIP2 to the tail region (Carisey and Ballestrem 2010). Interactions with talin or α-actinin have been shown to aid in the activation of vinculin due to their high binding affinity that can release vinculin’s auto-inhibition (Chen et al. 2006). However, the mechanism for releasing the autoinhibition is debated because there appear to be multiple mechanisms. One involves binding of PIP2 to the vinculin tail domain, influencing vinculin head tail associations and along with interactions of the head domain with talin, exposes the binding site for F-actin (Gilmore and Burridge 1996; Saunders et al. 2006).

Signaling Mediators FAK One of the major events triggered by integrin adhesion is tyrosine phosphorylation of multiple signaling proteins in FCXs and FAs. Integrins have short cytoplasmic domains that lack catalytic activity so this tyrosine phosphorylation is mediated by kinases that become activated in response to integrin engagement and clustering. The central kinase in this response is the FAK, which partners with members of the Src family of tyrosine kinases (SFKs) to generate many of the tyrosine phosphorylation events. FAK specifically localizes to FAs via interactions with talin or paxillin and plays a role in numerous cell processes including cell migration and proliferation. The structure of FAK implies that it acts as both a scaffold and a kinase, with its kinase domain found within the central region of the protein. FAK contains a protein 4.1, ezrin, radixin, and moesin homology (FERM) domain at the N-terminus, which serves as a binding site for growth factor receptors and can promote nuclear translocation of FAK during cellular stress, a proline-rich domain, and focal adhesion targeting (FAT) domain (Tomar and Schlaepfer 2009). In the resting state, FAK exists in an auto-inhibited conformation in which the N-terminal FERM domain interacts with the central kinase domain inhibiting its activity (Lietha et al. 2007). Activation can occur via binding of PIP2 to a site in the FERM domain, or in response to mechanical tension (Cai et al. 2008). During activation, Y397 becomes autophosphorylated, and this provides a site for SFKs to bind via their Src homology 2 (SH2) domains. SFKs then phosphorylate a series of additional tyrosines within FAK, increasing FAK activity and generating additional binding sites for other proteins. These sites include Y576 and Y577 within the kinase activation loop, and Y861 and Y925 within the C-terminus. The latter phosphorylation site provides a binding site for Grb2, which activates the Ras/mitogenactivated protein (MAP) kinase signaling pathway. FAK is elevated in many malignant cells, and the increased expression contributes toward cell migration and proliferation (Owen et al. 1999). FAK null cells have excessive adhesions, which suggest that FAK is

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FA COMPLEXITY

Active

Inactive Rho

GDP

GTP

Rho

295

Inactive GTP

GEF

Rho

GAP

GDP

Pi

GDP

Effectors

Rho

GDP

GDI

Sequestered Figure 17.3. The Rho GTPase cycle. Rho proteins cycle between a GTP-bound active state and a GDP-bound inactive state. This cycle is controlled by three types of regulatory proteins: GEFs, GAPs, and GDIs.

not necessary for FA assembly but is needed for FA disassembly (Tomar and Schlaepfer 2009). Not only does FAK contribute to FA turnover and cell migration, but it is critical in promoting anchorage-dependent cell survival. For many cells, integrin-mediated attachment and signaling is necessary to prevent “anoikis,” the form of apoptosis induced by loss of adhesion (Chiarugi and Giannoni 2008). Rho GTPases Rho GTPases are the major regulators of assembly and maturation of FCXs into FAs. Rho GTPases are part of the Ras superfamily and are composed of at least 20 members. These regulators are found to play many roles in the cell including actin and microtubule remodeling, cell motility, cell division, vesicular transport, and transcriptional regulation as well as other processes (Etienne-Manneville and Hall 2002; Jaffe and Hall 2005). These GTPases act as mechanical switches that cycle between an active and inactive state depending on whether they are bound to GTP or GDP, respectively (Fig. 17.3). When inactive, GEFs are able to bind to Rho GTPases and stimulate the displacement of GDP, which is quickly replaced by GTP since the ratio of GTP to GDP in the cytosol is 10:1. Once active, Rho GTPases are able to signal to their downstream effectors until a GAP stimulates the intrinsic GTPase activity causing the hydrolysis of GTP to GDP. When inactive, guanine nucleotide dissociation inhibitors (GDIs) are able to sequester Rho GTPases to the cytosol by binding to their C-terminus and prevent GTPase activation by blocking GDP displacement (Dubash et al. 2009). The best characterized Rho GTPases that play a role in cell adhesion and migration are Cdc42, Rac1, and RhoA, all of which have distinct roles in FA dynamics as described above. GEFs and GAPs are the primary regulators of Rho GTPases. Members of the major GEF family contain Dbl homology (DH) and pleckstrin homology (PH) domains, which are the primary domains that allow GDP/GTP exchange and recruitment to the plasma

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membrane, respectively (Rossman and Sondek 2005). Approximately 70 Dbl-related GEF family members exist, with some having a narrow specificity for the GTPases on which they act, but others acting on multiple Rho proteins (Rossman and Sondek 2005). Similarly, there are approximately 80 known GAPs that serve to negatively regulate Rho GTPases (Peck et al. 2002). If these GEFs and GAPs are not properly expressed or regulated themselves, then it leads to either the constant activation or constant inactivation of certain Rho GTPases. This misregulation of Rho GTPases can lead to abnormal development or diseases including cancer and neurological disorders (Peck et al. 2002). A novel mechanism for Rho GTPase activation was discovered by Campbell and colleagues through in vitro studies (Heo and Campbell 2005). They discovered that there is a conserved redox-sensitive cysteine within the nucleotide-binding site of Rho GTPases. Oxidation of this cysteine by reactive oxygen species (ROS) displaced the bound nucleotide. Upon reduction of the cysteine, nucleotide exchange was observed in the absence of a GEF. Exploring whether this may also occur in cells responding to ROS, activation of RhoA was noted in the presence of physiological levels of hydrogen peroxide. Consistent with the activation occurring via redox modification of the critical cysteines, activation was blocked when two cysteines within the nucleotide-binding site were mutated to alanines (Aghajanian et al. 2009). How important this mode of Rho protein activation may be remains to be determined, but it is striking that ROS are generated in many physiological as well as pathological situations.

MECHANISMS OF FA AND FCX DISASSEMBLY FA disassembly has been much less studied than assembly, but disassembly is a critical process for cell migration. Several pathways have been identified, and it seems likely that there are multiple ways by which an FA or FCX can be disassembled depending on the situation and the signal. Inhibiting RhoA activity rapidly leads to FA disassembly (Ridley and Hall 1992) and dispersal of the clustered integrins (Chrzanowska-Wodnicka and Burridge 1996). Another mechanism that contributes to turnover is signaling from FAK and Src. As previously mentioned, FAK and Src can inhibit RhoA activity through p190RhoGAP (Ren et al. 2000). Additionally, RhoA and mDia are able to localize Src to FAs in order to contribute to disassembly (Huveneers and Danen 2009). Cells lacking FAK or Src reveal decreased FA disassembly as judged by paxillin dynamics, although FA assembly was unaffected (Webb et al. 2004). This suggests that the phosphorylation of FA substrates is required in order for proper disassembly to occur. A different mechanism for disassembly involves calpain, a calcium-dependent protease (Perrin and Huttenlocher 2002; Franco and Huttenlocher 2005). A number of FA proteins are cleaved by calpain during disassembly. Although multiple calpain substrates have been identified in FAs, most attention has focused on talin. In an elegant study, a calpain-resistant version of talin was engineered. This resulted in decreased FA disassembly and decreased cell migration (Franco et al. 2004). One of the results of talin cleavage by calpain is that it generates the talin head domain, which is known to activate integrins. Investigating the fate of the talin head, it was found that this domain binds Smurf1, an E3 ubiquitin ligase that promotes its ubiquitination and proteasome-mediated degradation (Huang et al. 2009). It has been known for some time that growing microtubules (see Chapter 14) target FAs for disassembly (Kaverina et al. 1998). Ezratty and coworkers took advantage of this to develop a system where regrowth of microtubules following washout of a microtubule inhibitor resulted in the essentially synchronous disassembly of FAs (Ezratty et al. 2005).

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Strikingly, they found that this disassembly occurred even in the presence of constitutively active RhoA, indicating that it was not due to RhoA inhibition. Exploring the mechanism for disassembly, they uncovered a mechanism in which dynamin is involved, suggestive of endocytosis. Continuing this work, they and others provided evidence for a clathrinmediated endocytic mechanism of FA disassembly (Chao and Kunz 2009; Ezratty et al. 2009). However, the internalization process may be cell type specific as β1 integrins in epithelial cells can be internalized by lipid rafts, cholesterol-rich domains that contain caveolins, proteins involved in clathrin-independent endocytosis (Vassilieva et al. 2008).

IMAGING ADHESION DYNAMICS For many years, FAs were viewed as essentially static domains. Certainly compared with many regions of the cell surface, they are relatively stable. However, FAs can move over the substratum and components exchange between the adhesion and the soluble cytoplasmic pool, revealing that they are more dynamic than originally anticipated. Using a green fluorescent protein (GFP)-tagged integrin, Smilenov and colleagues showed that FAs in stationary cells move centripetally in a myosin-dependent fashion. Unexpectedly, the FAs of motile cells were less mobile with respect to the substrate on which the cells were migrating (Smilenov et al. 1999). Movement of FAs has also been observed when cytoplasmic components have been tagged with fluorescent markers and followed over time (Zamir et al. 2000). Taking advantage of the ease with which components can be visualized after expressing them as fusion proteins with GFP or other fluorescent markers, an increasing number of studies are applying a variety of biophysical techniques to examine the dynamics of FA proteins. For example, fluorescence recovery after photobleaching (FRAP) allows the dynamics of a fluorescent protein to be measured both in an adhesion and in other areas of the cell (Webb et al. 2003). Other techniques that are gaining popularity are fluctuation correlation spectroscopy (FCS) and image correlation spectroscopy (ICS). These fluorescent methods are being used to develop models that describe adhesion dynamics (see Chapter 5 for a discussion of the use of fluorescent approaches to study membrane domain dynamics). FCS uses fluctuations in fluorescence intensity to correlate protein concentration and to reflect molecular interactions (Webb et al. 2003). Similarly, ICS uses changes in fluorescence intensity within a specific area of an image utilizing a scanning confocal microscope. These types of imaging techniques have been used to monitor linkage efficiency between integrins and F-actin and to show that not only does this vary between cell types and depend on the substrate density, but that the linkage efficiency increases during FA maturation (Brown et al. 2006). Correlation spectroscopy has also been used to monitor paxillin incorporation and removal as an indicator of adhesion dynamics (Digman et al. 2008). This revealed that FAs are heterogeneous and that as large FAs slide across a substratum, there is a “treadmilling” effect, with paxillin monomers being added on one side but paxillin in aggregates dissociating from the other side. Another similar method is raster-scan image correlation spectroscopy (RICS), which analyzes diffusion and binding dynamics of proteins within a single image instead of single points on an image (Digman et al. 2005). This method can be used to measure diffusion of protein complexes and estimate the fraction of interacting proteins, as well as determine the spatial and temporal distribution of these complexes. In one study, vinculin–paxillin and FAK– paxillin complexes were monitored using two-color fluorescence. Away from adhesions, complexes were not detected, but large aggregates were seen to detach from disassembling FAs and then rapidly dissociate (Digman et al. 2009).

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FUTURE PERSPECTIVES In this chapter, we have briefly covered some of the more extensively studied proteins and mechanisms that contribute toward the assembly and disassembly of FA domains. A general theme in FA dynamics is that there are mechanisms that act in positive feedback loops promoting assembly. Increasingly, there is evidence that many of the components respond to force by adopting conformations that enhance their interactions or that expose cryptic binding sites. These include integrins themselves, talin, p130cas, vinculin, and FAK. The ultrastructural organization of FAs has been refractory to conventional electron microscopy, probably because of the density of the interacting components. However, the application of biophysical techniques to visualize the dynamics of individual FA components is rapidly increasing our ability to understand many of the interactions that occur within these domains. The next several years promise to be an exciting era in FA research.

ABBREVIATIONS CBD DH ECM FA FAK FAT FCS FCX FERM Fn FRAP GAP GDIs GEF GFP HBD

cell-binding domain Dbl homology extracellular matrix focal adhesion focal adhesion kinase focal adhesion targeting fluctuation correlation spectroscopy focal complex protein 4.1, ezrin, radixin, and moesin homology fibronectin fluorescence recovery after photobleaching GTPase-activating protein guanine nucleotide dissociation inhibitors guanine nucleotide exchange factor green fluorescent protein heparin-binding domain

ICS LPA MAP p130Cas PDZ PH PIP2 PM RICS ROCK ROS SFK SH2 Syn4 VASP Vn

image correlation spectroscopy lysophosphatidic acid mitogen-activated protein Crk-associated substrate post-synaptic density 95, discs-large, zonula occludens-1 pleckstrin homology phosphatidylinositol 4,5-biphosphate plasma membrane raster-scan image correlation spectroscopy Rho kinase reactive oxygen species Src family kinase Src homology 2 syndecan-4 vasodilator stimulated protein vitronectin

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THE ADHERENS JUNCTION Christopher P. Toret W. James Nelson

DEFINITION The adherens junction (AJ), a proteinaceous domain of the plasma membrane, is located at sites of cell–cell contact and integrates individual cells into a cohesive tissue monolayer. The primary function of the AJ, which plays a central role in development, morphogenesis, and wound healing, and is often defective in cancer and metastasis (Berx and van Roy 2009), is to form calcium-dependent adhesions between adjacent cells (Meng and Takeichi 2009; Shapiro and Weis 2009). The AJ is found in all metazoans (King 2004) and in many different cell types (Meng and Takeichi 2009), although this domain has been most thoroughly studied in the context of epithelial cells. The AJ of epithelial cells is located in a specialized region of the plasma membrane at the apex of the lateral plasma membrane that demarcates the boundary between the apical and basolateral plasma membranes in polarized epithelial cells (Meng and Takeichi 2009; see also Chapter 21). The AJ is part of a tripartite, intercellular junctional complex that includes the tight junction (apical to the AJ) and desmosomes (basal to the AJ) (Farquhar and Palade 1963; see also Chapter 19). The central component of the AJ is the transmembrane protein cadherin, which is clustered at sites of cell–cell contact and mediates extracellular attachment across approximately 20 nm of extracellular space via calcium-dependent trans-dimerization of cadherins on opposing cells (Pokutta and Weis 2007; Zhang et al. 2009). The AJ, through cadherins, interacts with cytoplasmic proteins and the actin cytoskeleton. The cytoplasmic tail of cadherin interacts with proteins that regulate cadherin turnover, cell signaling, and linkage to the actin cytoskeleton (Hartsock and Nelson 2008). In epithelial cells, a circumferential belt of bundled F-actin underlies the AJ and is thought to play in important role in maintaining cell tension and cell shape (Rauzi et al. 2008; Cavey and Lecuit 2009; Fernandez-Gonzalez et al. 2009), although its formation and connection to the adhesion machinery is not fully understood. The integration of extracellular adhesion and intracellular regulation of the actin cytoskeleton by cadherins at the AJ are important in driving epithelial cell migration during development (Cavey and Lecuit 2009). The AJ is now considered a dynamic membrane domain that is in constant flux depending on the cellular context.

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HISTORICAL PERSPECTIVE The importance of cell–cell adhesion in animal and tissue morphogenesis emerged from the fields of embryology and zoology during the early 1900s when scientists sought to understand the relationship between intrinsic cellular properties and global changes in embryos during morphogenesis (Okada 1996). For example, in 1907 Wilson showed that disassociated sponge cells were capable of re-aggregating and forming miniature sponges (Wilson 1907). Later, cell–cell adhesion and cell sorting properties were uncovered in seminal studies on a number of animal species (Holtfreter 1939; Trinkaus and Groves 1955; Moscona 1962; Steinberg 1963; Steinberg and Gilbert 2004). This early research launched studies of the disaggregation and re-aggregation properties of animal cells. Farquhar and Palade’s electron microscopy studies of cell–cell junctions revealed dense plaques on the plasma membrane referred to as the zonula adherens at the interface of two cells (Farquhar and Palade 1963). This membrane domain, which corresponds to the AJ, was bordered by the tight junction and desmosomes and defined the tripartite junctional complex that links together epithelial cells (see Chapter 19). Intracellular filamentous structures were associated with these plaques that corresponded to the actin belt that underlies the AJ, and intermediate filaments associated with desmosomes. By the 1960s it had become apparent that there was a vital role for Ca2+ in cell–cell adhesion and, after some initial confusion, a trypsin-sensitive factor was found to be required for cell–cell adhesion (Okada 1996). Work by Takeichi identified a 150 kDa proteinaceous factor that mediated Ca2+-dependent adhesion (Takeichi 1977). This protein was later refined to 124 kDa and given the name “cadherin” for its Ca2+-dependent role in cell–cell adhesion (Yoshida and Takeichi 1982). Soon thereafter, other groups independently identified cadherin, although they had given it different names including uvomorulin, L-CAM, C-CAM, A-CAM, and Cell-CAM (Kemler et al. 1977; Damsky et al. 1983; Edelman et al. 1983; Cunningham et al. 1984; Vestweber and Kemler 1984; Volk and Geiger 1984; for a historical overview see Franke 2009). The newly identified cadherin adhesion protein was subsequently localized to the zonula adherens (Boller et al. 1985). Together, these studies laid the foundation for structural and functional studies of the plasma membrane domain now recognized as the AJ. After the discovery of cadherin, many related adhesion proteins were identified. These cadherin-related proteins were all characterized by the presence of extracellular cadherin repeat (EC) domains. Together, these proteins constitute the cadherin superfamily that comprises the “classical” cadherins (E-, N-, P-, VE-, and R-cadherin), type II cadherins, protocadherins, and other outlying members (Wheelock and Johnson 2003; Perez and Nelson 2004; Halbleib and Nelson 2006). The classical cadherins have five extracellular repeats, each of which contains a calcium-binding EF-hand domain (Ozawa et al. 1990), and a highly conserved cytoplasmic tail domain. Analysis of the “classical” cadherins revealed that different cadherin proteins were present in different cell types: E-cadherin in epithelia, N-cadherin in neurons, VE-cadherin in vascular endothelium, and P-cadherin in the placenta (Halbleib and Nelson 2006; Franke 2009). Initially, the cadherins were viewed as a means of defining tissue and cell type, although it is now recognized that some cells can switch cadherin type, for example, during wound healing (Hinz et al. 2004) and neurulation (Nakagawa and Takeichi 1995). Although cadherin is the major component of the AJ, subsequent work by Takai and colleagues identified the nectin family of adhesion proteins as AJ adhesion proteins (Mandai et al. 1997; Takahashi et al. 1999). Like cadherins, nectins are plasma membrane proteins that form trans-dimers across the AJ but, unlike cadherins, are members of the Ig superfamily and are Ca2+-independent adhesion proteins (Takai et al. 2008b). Nectins

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have been proposed to play an important role during initial stages of cell–cell contact that later mature into cadherin-based contacts (Takai et al. 2008a; Takai et al. 2008b).

PERIPHERAL MEMBRANE COMPONENTS OF THE AJ In epithelial cells, the AJ mediates linkages between extracellular contacts through the tripartite junctional complex and intracellular signaling and cytoskeletal machineries (Fig. 18.1). This linkage occurs through the cytoplasmic domains of nectin and cadherin, which bind to cytoplasmic proteins that regulate cell signaling, protein trafficking, and cytoskeletal organization (Table 18.1). Using the cytoplasmic tail of E-cadherin as a probe, two interacting cytoplasmic proteins were identified and named α-catenin (102 kDa) and β-catenin (88 kDa) (Ozawa et al. 1989; Nagafuchi et al. 1991). Shortly thereafter, other cadherin-interacting cytoplasmic proteins were identified, including plakoglobin (80 kDa) and p120-catenin (Cowin et al. 1986; Hatzfeld 1999; McCrea and Gu 2010). These proteins regulate a wide range of cellular activities and thus connect the AJ to a broad spectrum of downstream cellular responses to cell–cell adhesion. cDNA sequencing of β-catenin and plakoglobin revealed that they were orthologs of the Drosophila protein, armadillo (McCrea et al. 1991; Peifer et al. 1991), which is involved in the Wnt/wingless signaling pathway and the regulation of cell proliferation (Nelson and Nusse 2004). These proteins have characteristic “arm” repeats that form a general protein–protein interaction domain (McCrea et al. 1991; Hulsken et al. 1994). That β-catenin and armadillo (and plakoglobin) are structurally related indicates that they have dual functions in the cell (Nelson and Nusse 2004): At the AJ, β-catenin links α-catenin Polarized epithelial cells

Adherens junction

Apical membranes

Tri-junctional complex

Basolateral membranes

Nectins Afadin

Tight junctions

Actin structures (Formin, Myosin, vinculin)

Adherens junctions

EPLIN

Desmosomes

+

Microtubules PLEKHA7 (Vesicle trafficking Nezha CLIP170, dynein)

_

Ubiquitin

α-catenin β-catenin p120-catenin

Hakai

Ubiquitin Endocytosis, degradation, and recycling Clathrin-mediated and Rab11-dependent

2+

Ca

2+

Lateral targeting patch (Secretion of adherens junction and basolateral proteins)

Ca

E-cadherin

Figure 18.1. Schematic diagram depicting major protein–protein interactions of the adherens junction domain that links together epithelial cells. See text for details.

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306

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p120, Nezha

E-cadherin, RhoA, PLEKHA7

Plekha7/1122

Central armadillo domain; N-terminus binds PLEKHA7

Afadin/Par3

Nectin tail/(differs for nectin type and isoform) p120/971

Actin binding, α-catenin binding

5 extracellular EC domains; Cytoplasmic tail(intrinsically unstructured by nuclear magnetic resonance analysis) 3 extracellular Ig domains

β-catenin, p120

E-cadherin tail/ 151

EPLIN/600

Central armadillo domain 151–671

Cadherin cytoplasmic domain, α-catenin

β-catenin/781

WW, PH, coiled-coil domains

Two regions of predicted disorder flanking a central LIM domain

57–264 β-catenin binding +dimerization; 385–632 M domain; 678–906 actin-binding

β-catenin, F-actin, afadin, ZO-1, EPLIN

αE-catenin/906

Domain organization

Interactions at adherens junctions

Cytoplasmic Machinery of the Adherens Junction

Protein/length (amino acid)

TABLE 18.1.

None

LIM domain

C-terminal consensus E/A-X-Y-V binds afadin None

Armadillo domain ± E-cadherin cytoplasmic domain; α-catenin binding region 118–149 bound to α-catenin 57–264 Bound to β-catenin (last 100 amino acids)

57–264 fusion to β-catenin 118–149; 82–264; M domain

Known structures

C: 85 X: 52 Z: 57 DM: 40 CE: 40 M: 76 C: 58 X: 45 Z: 36 M: 90 C: 70 Z: 52

C: 71 X: 84 Z: 82 DM: 36 CE: 31

C: 97 X: 93 Z: 90 DM: 61 CE: 38 C: 99 X: 97 Z: 97 DM: 67 CE: 34

Ortholog% identical to human (full-length)

Connect p120, Nezha

Regulator of cadherin trafficking and actin

Adhesion protein

Adhesion protein

Link classical cadherins and α-catenin

Actin regulator

Function at adherens junctions

307

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N-term microtubule binding domain, C-terminal dimerization domain

CH, WW, RasGAP, RasGAP-C-term

FHA, Ras-bnding domain, PDZ domain

APC, β-catenin

Microtubules, EB1, β-catenin, Axin, ASEF

APC

APC, KIF3

APC

β-catenin

Actin, α-catenin, Ras

AMER1/1135

APC/2843

Asef/690

Kap3/ 793

EB1/268

IQGAP1/1657

Afadin/1816

Armadillo domain 297–400; 540–662

SH3, DH, PH

N-terminal coiled-coil dimerization, armadillo

Central region homology to pyruvate dehydrogenase complex dihydrolipoamide acetyltransferase; C-terminal DUF domain No known domains

Minus-end microtubule, Plekha7

Nezha/1276

Domain organization

Interactions at adherens junctions

Protein/length (amino acid)

FHA, Ras-binding domain, PDZ domain

GAP domain, RGC domain

N-terminal domain alone, C-terminal domain complex with p150Glued

None

Autoinhibited structure 66–540;

Dimerization domain, 129–250 coiled-coil, complexes with β-catenin and Axin

None

N-term 3–141

Known structures

C: 89 X: 74 Z: 66 DM: 53 CE: 32 C: 84 X: 65 Z: 75 C: 96 X: 93 Z: 84 DM: 56 M: 96 C: 95 X: 91 Z: 88 DM: 70 CE: 56 C: 92 X: 85 Z: 83 DM: 35 X: 78 Z: 73 DM: 32 CE: 34

M: 78 X: 46 Z: 35

M: 89 X: 54 Z: 63

Ortholog% identical to human (full-length)

(continued)

Link to actin

RasGAP

Microtubule binding

Link APC to kinesin

GEF

Connection of cadherin complex to microtubule complex

Anchorage of MT to cadherin in adherens junction

Function at adherens junctions

308

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2 WD40 β-propellers

L27, 3 PDZ, SH3, GUK

240 residue coiled-coil domain at N-term (possibly BAR domain) Coiled-coil domain at N-term (possibly BAR domain); GAP SH3, PH, DBL, Coiled-coil trimerization

ARF-GAP, ankryin repeats, Coiled-coil dimerization, C-terminal paxillin binding domain

Lgl, β-PIX, APC, ZO2

Par6, syntaxin 4, aPKC, exocyst?

APC, CASK, Lin7, K+ channel

Rich1, Pals1, Patj, MPP7, MUPP1, Par3

Angiomotin, aPKC, Par3, endocytic adaptors Scrb, Git-1, Aip4 (Ub E3 ligase), PAK2

β-PIX

Scribble/1630

Lgl/1015

Dlg1 (SAP97)/904

Angiomotin (AMOT)/676

Rich1/881

Git-1/761

β-PIX (RhoGEF7)/803

2 ASD domains unique to Shroom family LRR, 2 LAP-specific domains, 4 PDZ

Actin binding

Domain organization

Shroom1/847

Interactions at adherens junctions C-terminal C2 domain (synaptotagmin-like)

(continued )

Bitesize/1102

Protein/length (amino acid)

TABLE 18.1.

Coiled-coil dimerization

Coiled-coil trimerization, SH3 (several ligand complexes), DBL

None

All 3 PDZ domains, including complexes with K+ channel peptide None

None

PDZ 1,3,4

None

None

Known structures

M: 92 Z: 74 DM: 35 CE: 30 C: 85 X: 63 Z: 60

M: 84 Z: 67

M: 59 X: 30 M: 86 Z: 58 DM: 38 M: 90 X: 68 Z: 57 DM: 36 CE: 20 X: 82 Z: 74 DM: 50 CE: 45 M: 81 X: 61

(Only insect)

Ortholog% identical to human (full-length)

ArfGAP

RhoGEF

Cdc42 RhoGAP

Scaffold

Scaffold

Scaffold

Scaffold

Needed for AJ stability (flies); organizes F-actin through moesin Actin binding

Function at adherens junctions

309

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RalA, exo84

Sec3, Sec5, Sec8

Sec3, Sec6, Exo70, IQGAP1

Sec5, Sec15, Exo84

Sec10, Rab3, Rab8, Rab11, Rab27

Sec8,Tc10

Sec5, Sec10, RalA

Sec5/924

Sec6/745

Sec8/974

Sec10/708

Sec15/804

Exo70/684

Exo84/725

At least two helical domains unique to superfamily; PH domain (Ral binding)

At least two helical domains unique to superfamily; homologous to known exocyst helical domains 4 all-helical domains, unique to this superfamily

Homologous to known exocyst helical domains

Ral-binding IPT (β-barrel) domain, remainder homology to known exocyst helical domains At least two helical domains unique to superfamily, remainder homologous to known exocyst helical domains Homologous to known exocyst helical domains

Domain organization

PH domain bound to RalA

Complete structure except first 82 amino acids (mouse, 94% identical)

None

None

None

none

Ral-binding domain, complex with RalA

Known structures

C: 85 X: 79 Z: 74 DM: 38 CE: 32 X: 87 Z: 80 DM: 34 CE: 27 C: 95 X: 89 Z: 83 DM: 43 CE: 36 C: 89 Z: 82 DM: 41 CE: 38 C: 81 X: 72 Z: 73 DM: 30 CE: 24 C: 81 X: 73 Z: 73 DM: 27 CE: 25

Z: 76 DM: 33 CE: 26

Ortholog% identical to human (full-length)

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Membrane trafficking/ exocyst subunit

Function at adherens junctions

Human AJ proteins listed with relevant interactions, domains, and functions. Sequence identities are from NCBI BLAST (M = Mus musculus [mouse], C = Gallus gallus [chicken], X = Xenopus laevis or tropicalis (clawed frog, whichever species was in database; similar numbers if both), Z = Danio rerio [zebrafish], DM = D. melanogaster [fruit fly], CE = C. Elegans [nematode]). Further details can be found in Takai et al. (2008b), Meng and Takeichi (2009), Nelson (2009), and Green et al. (2010).

Interactions at adherens junctions

Protein/length (amino acid)

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to cadherin, while a cytosolic pool of β-catenin regulates gene transcription in the nucleus in response to Wnt/wingless signaling. The ability of the AJ to bind and sequester β-catenin implies a role for the AJ in Wnt/wingless signaling regulation (Nusse 1999; Nelson and Nusse 2004; Heuberger and Birchmeier 2010). There is some evidence that cadherinbound β-catenin can be directly recruited to the nucleus under certain circumstances (Brembeck et al. 2004; Kam and Quaranta 2009). The arm-repeat protein, plakoglobin, was found to associate with classical cadherins at AJs, but its association with desomosomal cadherins has emerged as a central focus for plakoglobin studies (Green et al. 2010). Plakoglobin has also been implicated in counteracting canonical Wnt signaling, but its role is not as well understood (Schmidt and Koch 2007). p120-Catenin, another of the arm-repeat proteins, binds to the juxtamembrane domain of cadherins at a conserved octapeptide sequence (YDEEGGE) (Ferber et al. 2002). Initial studies suggested a role for p120 in regulating Rho GTPases (Noren et al. 2000), which, in turn, regulate actin cytoskeletal dynamics (Jaffe and Hall 2005). However, it is unclear how cadherin association with p120-catenin influences Rho GTPase signaling (Wildenberg et al. 2006; Reynolds 2007; Hartsock and Nelson 2008). p120-Catenin plays a clearer role in regulating E-cadherin stability and endocytosis at the plasma membrane by antagonizing the ubiquitination of E-cadherin via the E3 ligase, Hakai, which results in cadherin endocytosis (Fujita et al. 2002; Davis et al. 2003; Ishiyama et al. 2010). The other major cytoplasmic component of the cadherin complex is α-catenin (Rimm et al. 1995; Kobielak and Fuchs 2004), which has an emerging role apart from the AJ (Weis and Nelson 2006). α-Catenin, is not an arm-family protein, but has sequence homology and domain organization similar to those of the actin-binding protein vinculin (Herrenknecht et al. 1991; Nagafuchi et al. 1991) and binds F-actin (Rimm et al. 1995; Kobielak and Fuchs 2004). Three cell-type-specific isoforms of α-catenin are present in vertebrates (αE-catenin, αN-catenin, and αT-catenin), while only one is present in less complex metazoans (Kwiatkowski, pers. comm.). Initially, the actin-binding property of α-catenin was quickly recognized as a potential direct link to the actin belt at the AJ (Herrenknecht et al. 1991; Brabletz 2004; Pokutta and Weis 2007), but subsequent characterization has challenged this model. Mammalian αE-catenin forms a monomer and homodimer (Drees et al. 2005; Yamada et al. 2005; Benjamin et al. 2010). Association of monomer with β-catenin weakens the affinity of αE-catenin for F-actin, whereas αEcatenin homodimer binds strongly to F-actin (Drees et al. 2005; Yamada et al. 2005). An indirect link between actin and α-catenin may occur through the interaction of α-catenin with other actin-binding proteins including afadin (Pokutta et al. 2002) and EPLIN (Abe and Takeichi 2008), although much less is known about the mechanisms of these protein– protein interactions (Weis and Nelson 2006). Other studies have proposed that a link between actin and α-catenin may occur under mechanical tension similar to vinculin at focal adhesions (see Chapter 17), but this remains to be demonstrated biochemically (Yonemura et al. 2010). It is interesting to note that vinculin binds β-catenin at AJs (Drees et al. 2005; Peng et al. 2010), but auto-inhibition of vinculin binding to actin is not relieved (Drees et al. 2005). Recently, α-catenin was found to play a role in regulating actin dynamics and cell migration independently of cell–cell adhesion (Benjamin et al. 2010), indicating that its functions in actin regulation are broader than that immediately adjacent to the AJ. The cytoplasmic domain of nectins interacts with the cytoplasmic protein afadin (Irie et al. 2004). Afadin binds actin (Mandai et al. 1997), and is important for AJ formation in mammals (Ikeda et al. 1999). However, studies on the Drosophila homolog, canoe, suggest that is not required for AJ formation, but instead has an important role in linking

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apical cell constriction with the actin belt (Sawyer et al. 2009). Afadin also binds α-catenin (Pokutta et al. 2002), indicating that it provides a link between nectins and cadherins at the AJ. A binding domain for the small GTPase Rap1 is present in afadin (Miyata et al. 2009), which might prevent endocytosis of nonengaged cadherins (Hoshino et al. 2005), implying that nectins could have a positive role in stabilizing cadherin at the AJ.

METAZOAN DEVELOPMENT AND ORGANIZATION THROUGH THE AJ The AJ domain evolved with the appearance of primitive metazoans approximately 600 million years ago during the Precambrian period (Baum et al. 2008). The emergence of a polarized sheet of cells, an epithelium, in these early multicellular organisms provided a boundary to the outside environment, the ability to compartmentalize an internal compartment, and vectorial transport of ions and solutes between the outside and inside compartments (Magie and Martindale 2008). The AJ is absent in other multicellular organisms including plants and fungi (King et al. 2003). These distinguishing characteristics highlight the importance of the AJ in the evolutionary success of metazoans. During development, as a metazoan builds itself from a single cell to a complex multicellular organism, many cellular rearrangements occur. These organizational changes require intricate control of the cellular links between cells, and modulation of AJ function(s) to create the delicate balance between intracellular cohesion and cellular migration and rearrangements. As cells transition from a migratory phase to an adhesive cluster of cells, and then to a polarized epithelium, the axis of polarity changes from front–back to apical– basolateral, the plasma membrane differentiates, and the cytoskeleton is reorganized (Nelson 2009; see also Chapters 8 and 21 for discussions of epithelial cell polarization). These processes are initiated by formation of the AJ, and highlight the requirements for highly dynamic and complex functions of the AJ. AJ organization in different members of the animal kingdom is not always conserved. In mammalian epithelial cells, the tripartite junctional complex is stereotypically organized with the tight junction apical to the AJ, and desmosomes basal to the AJ (Meng and Takeichi 2009). In Drosophila, the junctions are organized differently: The AJ is the most apical junction, followed by the septate junction (analgous to the tight junction), but desmosomal complexes are lacking all together. Despite this organizational variation, the AJ organization and associated signaling and cytoskeletal machineries appear largely conserved (Tepass et al. 2001). Differences in AJ organization also occur between cell types. AJ structures exist in many cell types, including neurons and endothelial cells, but often have cadherin and catenin homologs different from those found in epithelia (Halbleib and Nelson 2006; Dejana et al. 2008; Suzuki and Takeichi 2008). Significantly, expression of tissue-specific cadherins plays an important role in cell sorting during development (Nose et al. 1988; Steinberg and Foty 1997; Foty and Steinberg 2004; Foty and Steinberg, 2005; Patel et al. 2006).

THE AJ AND EPITHELIAL CELL POLARITY Given the vital role of the AJ in cell and tissue rearrangements it is not surprising that a strong link between the AJ and the cell polarity machinery has emerged. Polarity protein com-

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plexes, Crumbs, PAR (partitioning defective), and Scribble (Bryant and Mostov 2008), are essential determinants of cell polarity (Roh and Margolis 2003), although it remains unclear precisely how constituent proteins function to create apical and basolateral plasma membrane domains. Polarity protein complexes appear to have scaffolding domains and regulate the cytoskeleton through Rho GTPases (Iden and Collard 2008), which can have global effects on membrane and cytoskeleton dynamics and protein trafficking (Nelson 2009). In the Drosophila embryo, the AJ does not form properly in mutants of the Crumbs complex, which comprises Crumbs, PALS1, and PATJ (Grawe et al. 1996; Muller and Wieschaus 1996; Tepass 1996). Note that the Crumbs complex localizes to the apical membrane, and many apical membrane markers are also lost in crumb mutants; whether this is a result of a lack of AJ formation or loss of Crumbs complex activity is unclear (Tanentzapf et al. 2000). The PAR complex (comprising atypical protein kinase C [aPKC]– Par6–Par3) localizes to the subapical region in Drosophila epithelial cells (Roh and Margolis 2003). Drosophila mutants of bazooka (Par3) fail to form an AJ and lose cell polarity, which is similar to the phenotype of armadillo mutants, suggesting that the bazooka/Par3 complex has a role in AJ assembly (Muller and Wieschaus 1996). The Scribble complex, which localizes on the lateral membrane below the AJ (Martin-Belmonte and Mostov 2008), may also have a link to AJ turnover through regulation of endocytic and exocytic pathways via Rho GTPases (Tepass et al. 2001; Nelson 2009; Shivas et al. 2010). In mammalian epithelial cells, these polarity protein complexes have a similar role in determining plasma membrane identity. The PAR complex localizes at the AJ and is directly linked to the AJ by an interaction between PAR3 and nectin (Takekuni et al. 2003). The relationship between the AJ and the polarity complexes appears to be intimately linked in both Drosophila and mammals; however, the precise mechanisms that coordinate the two processes remain poorly understood.

PROTEIN TRAFFICKING AND POLARITY AT THE AJ Both E-cadherin and nectin are transmembrane proteins and, therefore, must traffic through the secretory pathway to the plasma membrane to engage in cell–cell contacts. Little is known about nectin trafficking, but there appear to be several stages at which cadherin trafficking is regulated. Binding of β-catenin to the tail of E-cadherin facilitates E-cadherin exit from the endoplasmic reticulum and transit through the secretory pathway (Chen et al. 1999). The juxtamembrane domain of E-cadherin has a dileucine motif, a common cargo sorting motif for membrane proteins (Miranda et al. 2001), and binds ankyrin G (Nelson et al. 1990; Kizhatil et al. 2007), both of which are essential for polarized sorting and trafficking to the basolateral plasma membrane of polarized epithelial cells, and E-cadherin endocytosis (Miyashita and Ozawa 2007a). Finally, newly synthesized E-cadherin has an N-terminal prosequence that inhibits trans-dimerization and cell–cell adhesion, and is proteolytically cleaved prior to arrival at the plasma membrane (Ozawa and Kemler 1990). Many cellular trafficking events are coordinated by early membrane contact and initiation of the AJ (Nejsum and Nelson 2007). Nectin is believed to play an important adhesive role in the linking together of two initially contacting cells, which is subsequently followed by E-cadherin clustering and trans-dimerization (Takai and Nakanishi 2003). Studies have found that the small GTPase Rac is transiently recruited to initial sites of cell contact and is important for contact formation and regulating the actin cytoskeleton (Braga et al. 1997; Yamada and Nelson 2007; Perez et al. 2008). Delivery of membrane

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proteins to the cell surface requires the exocyst complex and SNAREs. The exocyst is linked to E-cadherin–nectin complexes (Yeaman et al. 2004) and rapidly localizes to sites of E-cadherin clustering at membrane contacts (Grindstaff et al. 1998). To form structurally and functionally different apical and basolateral plasma membranes, cells must maintain tight regulation and a delicate balance between the secretion of membrane proteins (exocytosis), the removal of membrane proteins (endocytosis), and protein trafficking between apical and basal membranes (recycling/transcytosis; see also Chapter 21). Toward this goal, the AJ plays a critical role in localizing the exocytic machinery. After recruitment of the initial exocyst components to the early cell–cell contact site, the basolateral-targeting SNARES are then recruited to the AJ (Hsu et al. 2004; Nejsum and Nelson 2009). The exocytic machinery, termed the lateral-targeting patch, provides specificity for the docking and fusion of secretory basolateral membrane vesicles. By using two aquaporins, AQP5 and AQP3, which sort to the apical and basolateral membranes, respectively, it was demonstrated that only AQP3 sorted to the forming AJ upon initial cell–cell adhesion (Nejsum and Nelson 2009). These studies revealed that the AJ plays a major role in directing the sorting of basolateral membrane cargo and the establishment of cell polarity.

MAINTENANCE AND REMODELING THE AJ Cell migration is important in embryogenesis but must occur in the context of cell–cell adhesion (Lecuit and Wieschaus 2002; Green et al. 2010). To achieve a dynamic site of cell–cell adhesion, cadherins at the plasma membrane are turned over by a balance between endocytosis and exocytosis. As noted in the last section, E-cadherin is directly targeted to the basolateral plasma membrane from the Golgi complex (see Chapter 8). E-cadherin is internalized via different pathways including clathrin- and caveolae-mediated endocytosis, lipid raft-mediated internalization, and macropinocytosis (Bryant and Stow 2004; Delva and Kowalczyk 2009). Recent photobleaching studies indicated that at mature AJs E-cadherin turnover is due to clathrin-mediated endocytosis (de Beco et al. 2009). Questions still remain about the role of other forms of endocytosis at the AJ. A study in Drosophila found a role for an Arp2/3-dependent endocytic pathway in AJ stability where the deletion of Cdc42–Par6–aPKC pathway destabilized AJ and similar phenotypes were observed when Arp2/3-dependent endocytosis was blocked (Georgiou et al. 2008; Leibfried et al. 2008). These studies indicate a role for Cdc42 in regulating cadherin endocytosis. Following endocytosis, E-cadherin may be recycled to the plasma membrane, or degraded. Drosophila DE-cadherin accumulates in Rab11-positive membranes in exocyst mutants, and mammalian E-cadherin delivery to the cell surface is dependent on Rab11, both suggesting that E-cadherin trafficking is regulated by the recycling pathway (Langevin et al. 2005; Lock and Stow 2005). Another recent study indicated a role for inactivation of Rab7 and the degradation of E-cadherin (Frasa et al. 2010). Many questions remain, however, about how cadherin stability is regulated at the cell surface, and how cadherin sorting decisions are made within the recycling and degradation pathways. The turnover of E-cadherin at the plasma membrane is clearly regulated by p120catenin. Binding of p120-catenin masks the dileucine signal in the cytoplasmic tail of cadherin, which is thought consequentially to protect E-cadherin from the endocytic machinery and degradation (Miyashita and Ozawa 2007b; Ishiyama et al. 2010). Furthermore, nectin-associated afadin increases the p120-cadherin complex in a Rap1dependent manner, which also stabilizes the AJ (Hoshino et al. 2005).

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Ubiquitination is a signal for endocytosis and degradation (Leon and HaguenauerTsapis 2009; Sorokin et al. 2009). The E3-ligase Hakai binds E-cadherin in a Src-dependent manner, and over-expression of Hakai destabilizes and increases ubiquitination of E-cadherin (Fujita et al. 2002), but questions remain about whether ubiquitylated E-cadherin is ultimately degraded via the endocytic–lysosomal pathway or the proteosome.

AJ REGULATION OF THE CYTOSKELETON Both the actin and microtubule cytoskeletons have functions at the AJ (see also Chapters 12 and 14). Actin is essential for AJ formation, but its precise role in the process remains to be established and likely changes as the AJ develops (Yonemura et al. 1995; WatabeUchida et al. 1998). In mature columnar epithelial cells, actin forms a submembranous belt of bundled actin filaments that circumscribes the AJ. The actin belt is capable of myosin-driven constriction, which plays an important role in shaping epithelial sheets during development (Cavey and Lecuit 2009; Fernandez-Gonzalez et al. 2009). Thus, the connection with the actin network at AJs may be complex with both lateral links along actin filaments and links at actin filament ends. Actin structures at the AJ differ in various cell types. Moreover, the actin organization differs between immature and mature AJs (Yamada and Nelson 2007), which suggests that the actin cytoskeleton is remodeled during AJ formation. Within the actin network, a basal to apical flow of actin filaments with cadherins at cell junctions was observed in some cell lines (Kametani and Takeichi 2007), although the biological significance of this activity is not clear. To identify how actin functions at the AJ, many studies have focused on understanding the association between the two structures, with most attention being given to αcatenin and afadin. However, other actin-binding proteins also localize to the AJ including myosin, vinculin, formin, and EPLIN (Geiger et al. 1985; Kobielak et al. 2004; Abe and Takeichi 2008; Cavey and Lecuit 2009). Recently, EPLIN was shown to bind α-catenin when in complex with β-catenin and cadherin. In cells lacking EPLIN, the actin-belt is reorganized into radial filaments at cadherins, suggesting EPLIN plays a role in actin-belt formation, but not attachment to cadherin (Abe and Takeichi 2008). Similar to other membrane processes that are linked to the actin cytoskeleton (focal adhesions, lamellipodial extensions, filopodia formation, cytokinesis, and endocytosis) (see also Chapters 9, 12, 14, and 17), more detailed biochemical analysis is needed to understand how each protein regulates the actin cytoskeleton alone and in conjunction with one another to fully understand their relationship with actin at the AJ. Microtubules do not form specialized structures at the AJ, but appear to have an essential role at the AJ (see also Chapter 14). Microtubules extend toward the AJ in a CLIP170-dependent manner (Stehbens et al. 2006), and a dynein–β-catenin link has been implicated in the attachment of microtubules to the AJ (Ligon et al. 2001). Drugs that inhibit microtubule polymerization interrupt AJ dynamics, suggesting that microtubules play a role in assembly and disassembly of the AJ (Waterman-Storer et al. 2000; Ivanov et al. 2006). The main role of AJ-directed microtubules may be associated with vesicle trafficking and delivery of AJ proteins and polarized membrane cargo (Nejsum and Nelson 2009). `The cadherin stabilizing protein p120-catenin may play an important role in linking microtubules to the AJ. p120-Catenin has been demonstrated to interact with microtubules via its C-terminus (Chen et al. 2003; Liu et al. 2007). The amino-terminus of p120-catenin also interacts indirectly with microtubules through the PLEKHA7-Nezha complex.

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315

PLEKHA7 localizes to and is essential for AJ formation (Meng et al. 2008). Nezha similarly localizes to the AJ but uniquely binds to microtubule minus ends and implicates a novel role for microtubules in AJ formation. Furthermore, the minus-end directed microtubule motor protein KIFC3 may be involved in trafficking to the tight junction. Loss of PLEKHA7, Nezha, KIFC3, or microtubules has similar defects on tight junction formation (Meng and Takeichi 2009). Taken together with the earlier CLIP170 data these results indicate that both plus and minus ends are localized to the AJ.

FUTURE PERSPECTIVES Despite recent advances in understanding the structural and functional organization of the AJ domain, many questions remain unanswered and new questions are being raised. For example, similar AJ structures are formed between static epithelial cells in tissues, between migrating mammalian cells during gastrulation, and during dynamic de novo formation of an epithelium in Drosophila cellularization; how is the AJ modified for these different functions in these different cellular contexts. While dynamics of AJ organization has been extensively studied during assembly following cell–cell adhesion, little is known about the dynamics of the AJ during cell movements within tissues. Dynamic organization of the AJ could be regulated by changing the balance between exocytosis and endocytosis of the cadherin–nectin complex. However, it remains unclear how individual cells coordinate protein trafficking, signaling, and polarity protein machinery via the AJ to globally create the different epithelial movements that drive tissue development and organization. Alternatively, or in combination, AJ dynamics could be regulated by changes in protein interactions at the cell surface. As highlighted in this review, several cytoplasmic proteins interact with cadherins and nectins at the AJ, but we know little about the molecular details of most of these protein–protein interactions, and it is possible that many more await discovery. In addition, despite the presence of known actin-binding proteins that interact with cadherins and nectins, it is not understood how they interact with, or locally regulate the actin cytoskeleton. Indeed, linkages between membrane proteins and cytoplasmic proteins that interact with the cytoskeleton are no longer envisioned as static linkages, and must be investigated in the context of mechanical strain on actin and microtubule cytoskeletons that occurs at the AJ.

ACKNOWLEDGMENTS Work for the Nelson laboratory is supported by NIH GM35527, and CT is currently supported by a

National Research Award T32 CA09151 from the National Cancer Institute.

ABBREVIATIONS aPKC CAM Cdc42 Ig domain EF-hand domain EPLIN KIF3C

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atypical protein kinase C cell adhesion molecule cell division control protein 42 imunoglobulin domain Eps15 homology domain epithelial protein lost in neoplasm kinesin family member 3C

PALS1 PAR PATJ PLEKHA7 Rap1

protein associated with Lin Seven 1 partitioning defective PALS1-associated tight junction protein pleckstrin homology domain containing, family A member 7 Ras-Proximate-1

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SPECIALIZED INTERCELLULAR JUNCTIONS IN EPITHELIAL CELLS: THE TIGHT JUNCTION AND DESMOSOME Keli Kolegraff Porfirio Nava Asma Nusrat

TIGHT JUNCTIONS Definition of the Tight Junction The tight junction is a cell–cell adhesive junction that brings together adjacent cell membranes to create a tight seal in the paracellular space. By creating selective pores within the intercellular space, the tight junction regulates the movement of ions and small molecules between epithelial cells (gate function). In addition, the junction maintains cell polarity by separating apical and basolateral plasma membrane domains within a given cell and restricting the mixing of membrane protein and lipids (fence function). The tight junction is composed of the transmembrane proteins, including the tetraspanin proteins claudins and occludin, and members of the junctional adhesion molecule (JAM) family. Plaque proteins associate with the cytoplasmic domain of the transmembrane proteins and include membrane-associated guanylate kinase (MAGUK) family members like the zonula occludens (ZO) proteins. Numerous other cytoplasmic proteins localize to the tight junction, and these proteins include transcription factors such as ZONAB and cell polarity proteins including Par3/6 and Scribble. Functionally, the formation of tight junctions between neighboring epithelial cells allows the separation of distinct body compartments and is essential for the regulation of tissue permeability. Dysregulation of tight junction proteins has been implicated in numerous pathophysiological processes affecting epithelia, including inflammation and tumorigenesis.

Kisses in the Dark: The Original Description of the ZO In the early 1960s, electrophysiological studies established that epithelial tissues could function as an electrical barrier and separate charge across the cell layers, thereby creating and defining distinct body compartments (Diamond 1962a, b, 1964; Diamond and Tormey 1966a, b; Tormey and Diamond 1967; Wright and Diamond 1968). This observation led

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Figure 19.1. Ultrastructure of cell–cell junctions in polarized intestinal epithelial cells. (A) Transmission electron microscopy of two adjacent intestinal epithelial cells (T84). At the level of the tight junction (white arrow), the intercellular space is obliterated by the close apposition of the cell membranes. The adherens junction (black arrow) is located below the tight junction (TJ). Numerous desmosomes (black arrowheads) can be observed along the lateral border of the cell membranes. (Image courtesy of Dr. James L. Madara.) (B) Freeze fracture replica of the mouse colon, demonstrating the anastomosing tight junction strands that run along the circumference of the epithelial cells. PF, P face; EF, E face; MV, microvilli. (Image courtesy of Dr. Hartwig Wolburg, Institute of Pathology, University Hospital Tübingen, Germany.)

to the notion that sites of intercellular contact must exist to impede the flow of ions and fluid across the epithelium. Examination of cells using light microscopy hinted at the presence of such intracellular bridges (Bizzozero 1864, 1870), but it was not until the 1960s that this intercellular “seal” was characterized (Farquhar and Palade 1963). Using electron microscopy (EM) to analyze polarized epithelial tissues, Farquhar and Palade described a point of contact between adjacent epithelial cells that appeared to result from a fusion of the outer leaflet of the plasma membrane from each cell (see Fig. 19.1A). This close apposition of cell membranes occurred at the apical or luminal domain of the lateral cell membrane, and tracer studies demonstrated that macromolecules and dyes could not penetrate this paracellular region (Miller 1960; Kaye and Pappas 1962; Kaye et al. 1962). Thus, this restrictive or occluding zone of the plasma membrane was referred to as the “zonula occludens” or tight junction. Other electron microscopists took a more romantic approach in their description and referred to these characteristic zones as “kisses” between neighboring epithelial cells (Diamond 1974). Initial studies supported the role of the tight junction as a paracellular “gate,” which restricted the passage of ions and macromolecules between individual epithelial cells. Interestingly, it was shown that not all tight-junction-containing tissues exhibit characteristics of a “tight” seal. Using a glass micropipette as a sensitive probe to monitor regions of electrical current, it was found that the paracellular space is a region of high conductance and different epithelial tissues have characteristic rates of conductance (Fromter 1972). Thus, epithelial tissues could be classified as “tight” or “leaky” based on these conductance parameters. Early electron microscopic studies could not distinguish the basis for this difference in ion selectivity and tissue permeability, and it was not clear what possible tight junction structure–function correlation existed in these different tissue types (Wright and Diamond 1968; Misfeldt et al. 1976).

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Nature of the Gate: Are Tight Junction Strands Composed of Lipid, Protein, or Both? During the 1970s and 1980s, technological advances in EM led to the development of the freeze fracture technique, in which frozen tissues are broken apart to allow the visualization of structural detail in the fracture plane by EM. Using this technique, the tight junction appears as parallel fibrils or strands running along the circumference of the epithelial cell with the strands interconnected by shorter branches of fibrils (Chalcroft and Bullivant 1970); see Figure 19.1B. These strands are embedded in the inner leaflet of the plasma membrane, also referred to as the protoplasmic (P) face. Complementary furrows or grooves are evident in the corresponding exoplasmic leaflet (E face). The first hint at a possible tight junction structure–function correlation came from the work of Claude and Goodenough in the 1970s, who demonstrated that “tight” epithelial tissues with reduced conductance had more parallel strands than their “leaky” counterparts (Claude and Goodenough 1973). Further work demonstrated that fragmentation or disruption of these strands correlated with changes in tissue permeability, as was observed during the absorption of glucose in the small intestine or in pathological settings such as inflammation or exposure to toxic stimuli (Sonoda et al. 1999; Fujita et al. 2000; Laukoetter et al. 2008). Therefore, the ability of a tissue to regulate permeability seemed to lie in the formation of these strands. However, it was not clear how these strands were formed and how they were able to act as an intercellular seal. Perhaps more pressing at the time was the looming question in the tight junction field: What is the molecular composition of these anastomosing fibrils? Early studies examining the composition of the tight junction fibrils pointed toward a lipid-based composition of the strands and several tight junction models proposed that the exoplasmic leaflets of neighboring cells were fused into a single membrane at the ZO, forming a cylindrical pore that controlled the passage of ions and macromolecules (Kachar and Reese 1982; Pinto da Silva and Kachar 1982). However, these models fell out of favor as the tight junction strands were shown to be sensitive to protein fixatives such as gluteraldehdye (Staehelin 1973) and were not extractable by detergents (Meller and ElGammal 1983). Furthermore, strand formation was shown to be disrupted by protein synthesis inhibitors, supporting a protein-based composition of the strands (Sang et al. 1980). In 1986, Stevenson and Goodenough identified a large cytoplasmic protein that localized to the strands of the ZO and appropriately named it “ZO-1” (Stevenson et al. 1986). However, it was not until the 1990s that the first integral membrane proteins of the tight junction were described. Using modern biological techniques, Furuse and Tsukita identified two tetraspan proteins that localize to the tight junction strands, naming them occludin, from the Latin “occludure” or “to occlude” and claudin, from the Latin “claudure” or “to close.” Further studies from the same group demonstrated that expression of the claudin proteins in fibroblasts allowed the formation of tight-junction-like strands and that claudin is the obligatory integral membrane of tight junctions (Furuse et al. 1993, 1998a, b, 1999); see Figure 19.2A. Tsukita’s group and others went on to show that charged extracellular loops of the claudin protein allow homotypic and heterotypic interactions among claudin family members (Tsukita and Furuse 2002; Colegio et al. 2003). These claudin interactions create a charged aqueous pore in the extracellular space (see Fig. 19.2B), whose electrophysiological characteristics depend on the claudin composition, as 24 claudin isoforms have been described to date. Our studies have revealed that the tight junction proteins localize in raft-like compartments at the cell membrane (Nusrat et al. 2000). These microstructures play an important role in the spatial organization of the tight junction and contribute to the regulation of paracellular permeability in epithelial cells.

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Actin–myosin ring

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Figure 19.2. Molecular composition of the tight junction and desmosome. (A) Schematic diagram depicting the transmembrane, plaque, and cytoskeletal proteins of each junction. (B) Current model of how the tight junction functions as a paracellular gate. The extracellular loops of tight junction proteins from adjacent cells interact to create aqueous pores in the paracellular space (red cylinders). The electrophysiological characteristics of these pores vary depending on the claudin composition.

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A Second Function for the ZO: The Tight Junction as a Membrane Fence between Apical and Basolateral “Pastures” An important characteristic of epithelial cells is the asymmetric localization of membrane components into at least two specialized regions: the apical surface, which faces the organ lumen, and the basolateral surface, which is in contact with the underlying connective tissue. The first evidence of such “polarized” organization of the epithelium came from electron microscopic studies of the small intestine, which demonstrated the presence of small fingerlike projections (microvilli) that were present only on the apical surface of the cells (Bizzozero 1870; Berridge and Oschman 1969). During the 1980s, a great deal of effort in the field of cell biology was focused on understanding the fundamental process of epithelial cell polarity and how proteins are targeted to and retained by specific plasma membrane domains. Because epithelial cells were known to segregate proteins and lipids to apical and basolateral surfaces, it was proposed that cells maintain this polarity through the actions of a fence-like structure within the plasma membrane (Pisam and Ripoche 1976; Sang et al. 1980). The work of Dragsten et al. using fluorescent membrane probes demonstrated that the tight junction restricts the lateral diffusion and mixing of apical or basolateral membrane components (Dragsten et al. 1981, 1982a, b; Rigos et al. 1983). Interestingly, their work showed that the tight junction only restricts the movement of lipids and proteins in the outer leaflet of the plasma membrane bilayer, as labeled probes added to the inner leaflet were not restricted in their diffusion. Thus, the evidence suggested that while the tight junction does not establish epithelial cell polarity per se, this structure was important for the maintenance of the distinct plasma membrane domains.

Molecular Composition of the Tight Junction After the discovery of ZO-1 in 1986, the number of tight-junction-associated proteins has grown substantially (see Fig. 19.2). Currently, the protein components of the tight junction can be broadly grouped as transmembrane or cytosolic/scaffolding proteins. The major components are discussed here; for more comprehensive reviews, see Gonzalez-Mariscal et al. (2003) and Gonzalez-Mariscal and Nava (2005). Tetraspan Proteins of the Tight Junction Occludin, tricellulin, and the claudin family members are integral membrane proteins that localize in the tight junction strands (Furuse et al. 1999; Ikenouchi et al. 2005). These proteins have four transmembrane domains and are believed to mediate cell–cell contact through their extracellular loops. The claudin family of proteins consists of at least 24 isoforms in humans and mice, and studies in claudin-deficient fibroblasts have demonstrated the homotypic and/or heterotypic interactions of the different claudin isoforms (Tsukita and Furuse 1999, 2000a, b, 2002; Asano et al. 2003; Daugherty et al. 2007). In addition, claudin knockout mouse models have further supported the unique role of different claudin family members in tight-junction-containing tissues, as the phenotype varies greatly depending on the deleted isoform (Kitajiri et al. 2004; Furuse and Tsukita 2006). In addition to the tetraspan integral membrane proteins, single-pass transmembrane proteins of the immunoglobulin superfamily, JAM, and Coxsackie and adenovirus receptor (CAR) have also been shown to localize to the tight junction strands (Bergelson et al. 1997; Martin-Padura et al. 1998; Roelvink et al. 1998) and to regulate barrier permeability (Liu et al. 2000; Mandell et al. 2004).

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Cytoplasmic/Scaffolding Proteins of the Tight Junction While expression of claudin family members is required for strand formation, it is the expression and localization of the ZO proteins that control where strand assembly occurs (Siliciano and Goodenough 1988). The ZO proteins are MAGUK homolog family members that contain several PSD-95/Discs large/ZO-1 (PDZ) domains, SH3 domains, and one guanylate kinase-like domain (Anderson et al. 1988; Gumbiner et al. 1991). The ZO proteins interact with the cytoplasmic tails of claudins, occludin, and JAM proteins through PDZ-dependent and -independent mechanisms (Itoh et al. 1999). Importantly, the ZO proteins link the tight junction structure to the apical actin cytoskeleton, directly or indirectly through associations with additional proteins, including afadin/AF6 (Miyoshi and Takai 2005). Importantly, it has been shown that modulation of this tight-junction-associated, circumferential actin–myosin “belt” can control epithelial permeability, by changing the paracellular pores formed by the tight junction strands.

Pathophysiology of the Tight Junction Since their initial description, tight junctions have been intimately linked with pathological conditions and diseases, and an alteration in their structure corresponds with changes in tissue permeability in both physiological and pathological settings. To date, numerous studies have examined the role of tight junction defects in disease, and human genetic disorders and mouse models have further elucidated the roles of individual tight junction proteins in normal tissue function and homeostasis. The tight junction was once thought of as a static, rigid structure that merely formed a selective seal between adjacent cells, but it is now known that this specialized zone is a dynamic, complex structure that can quickly be modulated in the context of physiology or disease. As early as the 1970s, it was reported that tight junction structure was altered during inflammation (Sosula 1975) and infection (Barker 1975). Soon it became evident that many pathogens and pathogen products could actually target the tight junction structure to cause increased tissue permeability and enhance their invasion through the body’s epithelial barriers (Laukoetter et al. 2007; Schulzke et al. 2009). Recently, tight junction proteins have been identified as receptors for a number of viruses, including CAR (Bergelson et al. 1997; Roelvink et al. 1998), reovirus (JAM) (Barton et al. 2001), and most recently, hepatitis C virus (Beard and Warner 2007; Evans et al. 2007) and HIV (claudins) (Andras et al. 2003; Zheng et al. 2005). Furthermore, bacterial toxins such as Clostridium perfringens enterotoxin have been shown to bind specifically to and degrade claudins to disrupt the intestinal epithelial barrier (Sonoda et al. 1999; Fujita et al. 2000). Interestingly, perturbation of tight junction function is also a hallmark of several inflammatory diseases, such as Crohn’s disease and ulcerative colitis (Laukoetter et al. 2008). In these disorders, inflammatory cytokines induce the downregulation and internalization of tight junction proteins or target components of the actin–myosin machinery, leading to a compromised epithelial barrier, enhanced exposure to luminal antigens, and further inflammation. A number of congenital syndromes have been described for patients with mutations in claudin family members. Interestingly, mutations in different claudin isoforms can have drastically different phenotypes. For instance, mutations in claudin-11 and claudin-14 have been shown to cause some forms of hearing loss (Wilcox et al. 2001; Kitajiri et al. 2004) due to sensory epithelial defects, whereas claudin-16 (paracellin) mutations have been detected in some patients with hypomagnesemia and hypercalciuria caused by kidney malfunction (Simon et al. 1999). Deletion of claudin-5 underlies the phenotype of the velo-cardio-facial syndrome, which affects multiple organ systems including the heart and kidneys (Morita et al. 1999).

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The tight junction has also been shown to be altered in cancers, and several tight junction proteins have been shown to contribute to tumorigenesis (Dhawan et al. 2005; Resnick et al. 2005; Darido et al. 2008). In 1977, it was reported that, in contrast to their normal cell counterparts, prostatic cancer cells possess a loosely organized network of fibrils and many individual, short strands that are disorganized and likely dysfunctional (Sinha et al. 1977). Given that a characteristic property of epithelial cancers is the loss of cell polarity, a breakdown in tight junction structure contributes to this defect in normal cellular asymmetry. Interestingly, the tumorigenic potential of several viral oncoproteins has been shown to correlate with their ability to sequester and/or inactivate tight junction proteins (Latorre et al. 2005; Gonzalez-Mariscal et al. 2009).

DESMOSOMES Definition of a Desmosome Desmosomes or “maculae adherentes” are cell–cell adhesive junctions that were originally identified in the epidermis using EM. While desmosomes are found in a number of tissue types, they are enriched in tissues that must withstand extensive mechanical stress, such as the skin or heart. The core protein components of the desmosome consist of the transmembrane cadherin family member glycoproteins desmocollin and desmoglein; the cytoplasmic plaque proteins of the plakin family, such as desmoplakin; and armadillo family members including plakoglobin and the plakophilins. The extracellular domains of the desmosomal cadherins mediate adhesion between neighboring cells, while their intracellular domains are linked to the intermediate filament cytoskeleton via the plaque proteins. The intimate association of the desmosomal plaque with the intermediate filament network reinforces the cell–cell contact sites and adds overall structural stability to the tissue. While the desmosome has always been described as a junction specialized for maintaining tissue integrity, recent evidence from animal models and human diseases demonstrate that desmosomal proteins likely have important signaling roles in proliferation, differentiation, apoptosis, and overall tissue morphogenesis, in addition to their structural role in forming desmosomes.

Discovery of the Desmosome The existence of a link or “node” between neighboring epithelial cells was first described in 1864 by the Italian medical doctor Bizzozero, who examined sections of epidermis using a light microscope (Bizzozero 1864, 1870; Calkins and Setzer 2007). The original descriptions declared these novel structures “desmosomes,” which derives from the Greek words “desmo,” meaning link or bond, and “soma,” or body (Schaffer 1920). With the advent of EM, it soon became evident that there were specialized structures linking adjacent cells in stratified epithelial tissues. Several descriptions reported the existence of symmetrical electron-dense plaques that were present in membranes of adjacent epithelial cells (Porter et al. 1945; Kelly 1966); see Figure 19.1A. These plaques appeared to also have a less defined substructure in the intercellular space. In addition, it was evident that densely packed, cytoplasmic tonofilaments converged on these plaques (Porter 1956; Odland 1958). The plaques were circular or oval in shape, and multiple plaques could be detected along the membranes of adjacent cells, leading to their description as “spot welds” in epithelial tissues. Interestingly, it was noted that these structures were almost always shared between neighboring cells; that is, the plaque was formed between cells in contact,

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and each cell membrane had an identical, aligned electron-dense plaque region with associated tonofilaments (Overton 1962; Arnn and Staehelin 1981). Occasionally, a “half” plaque would be detected, but these were usually cytoplasmic bodies suggesting that they were traveling to, or returning from, the cell membrane. Additionally, structures along the basal surface of epithelial cells were described as “hemi-desmosomes” (Kelly 1966), even though they were formed between the cell and the underlying extracellular matrix and therefore only had one-half of a circular desmosomal-like plaque. Through careful examination, electron microscopists identified three major regions of the desmosomal plaque: the midline, an electron-dense plaque consisting of an outer dense region and an inner, less dense region, and the associated tonofilaments that contacted the plaque at the inner region (Porter 1956; Odland 1958; Farquhar and Palade 1963). Around the same time, the cytoskeletal filaments were being carefully studied using the same techniques. Three major classes of filaments were known to exist in the cells: microfilaments with the smallest diameters (5–6 nm), intermediate filaments (7–10 nm), and microtubules with the largest diameters (20–25 nm). Based on these morphological criteria, it was clear that the desmosomal tonofilaments belonged to the intermediate filament class (Drochmans et al. 1978). The desmosome was first described as a structure enriched in the stratified epithelial tissues of the epidermis, and the work of many investigators demonstrated that desmosomes are also formed in other epithelial tissues, namely the simple or single-layered epithelial tissues of the intestine and kidney. In their careful characterization of junctional complexes in epithelial cells, Farquhar and Palade noted that desmosomes or “maculae adherentes” (adherent spots) were one of three cytoskeletal-associated plaques in polarized, simple epithelial tissues such as the mucosal linings of the intestine and kidney, the other junctions being the zonulae occludens (tight junctions) and the zonulae adherens (adherens junctions) (Farquhar and Palade 1963). The desmosome was distinguished from the other cell–cell junctions based on its association with the intermediate filament network in contrast to the microfilament network, which associates with the tight and adherens junctions, and because it is the most basally located of the three junction types. Around the time of the seminal Farquhar and Palade publication in 1963, structures with desmosomal morphology were also described in many other nonepithelial tissue types, including in meninges, between follicular dendritic cells of lymph nodes, and in the intercalating disks of cardiomyocytes (Fawcett and Selby 1958; Gusek 1962; Swartzendruber 1965). Importantly, it was also noted that desmosomal structures were highly enriched, and often larger, in tissues that routinely experience high mechanical stress, such as the heart and skin. While it seemed plausible that these junctions were specialized for resistance to mechanical stresses and in reinforcing tissue integrity, there was no direct evidence demonstrating this function of desmosomes.

The Desmosome as Intercellular Adhesive Cement The desmosome was a junction defined based on morphological criteria as observed using an electron microscope. Little was known about the exact role of this structure in the tissue and whether or not this junction could “malfunction” had yet to be determined. However, in the medical field, the existence of blistering skin diseases had already been described, in which patients suffer from a fragile epidermis that would separate easily from the underlying dermis (Waschke 2008). This class of diseases was referred to as “pemphigus,” and histological examination of skin biopsies demonstrated splitting of the epidermis just above the basal cell layer and detachment of individual epithelial cells from their neighbors (acantholysis). In 1964, a possible disease mechanism was proposed, as Beutner et al.

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described the presence of anti-epithelial antibodies coating the surface of keratinocytes isolated from patients with pemphigus (Beutner and Jordon 1964). These antibodies were not found in healthy skin samples, strongly suggesting that the antikeratinocyte antibodies were at least partly responsible for the disease. A series of studies using EM later demonstrated that the acantholytic keratinocytes have altered desmosomal junctions, detachment of tonofilaments, or both. Notably, Hashimoto and Lever described the appearance of abnormal granules on the surface of acantholytic keratinocytes and also observed a disruption of the amorphous intercellular substance that seemed to bridge adjacent cells (Hashimoto and Lever 1967a, b). They hypothesized that the granules were responsible for transporting the intercellular adhesive “cement” to the cell surface and that somehow this substance was abnormal in patients with pemphigus or that a lytic material was acting on the intercellular substance to break it down. Other EM studies of a different form of pemphigus, pemphigus foliaceus, also demonstrated breakdown of the desmosome, especially loss of the cementing substance, as well as loss of the tonofilament–plaque association at the membrane (Arnn and Staehelin 1981). Together with the findings that keratinocytes are coated with antibodies, it was proposed that the blistering disease pemphigus was caused by an “anti-intercellular cement” antibody that disrupted the function of desmosomes and made the skin fragile. Thus, pemphigus was the first group of diseases that were linked to desmosomal dysfunction. Later, through advances in molecular cloning and genetic studies, mutations in multiple desmosomal proteins were shown to underlie the pathogenesis of diseases affecting not only the skin, but hair and heart as well.

The Molecular Structure of the Desmosome Advances in biomedical technology of the 1980s and 1990s allowed the identification of the protein components of the desmosome (see Fig. 19.2A). Scientists exploited the highly insoluble nature of the desmosome to enrich and isolate relatively pure desmosomal fractions. Initial biochemical characterization of these fractions identified a mixture of glycosylated and nonglycosylated proteins that comprise the desmosomal plaque (Drochmans et al. 1978; Franke et al. 1981a, b). We now know that these proteins include the transmembrane glycoproteins desmocollin and desmoglein, as well as the nonglycosylated cytoplasmic plaque proteins desmoplakin, plakoglobin, and the plakophilins (Getsios et al. 2004; Holthöfer et al. 2007; Garrod and Chidgey 2008). Additional proteins have been described as being localized to desmosomes or interacting with known desmosomal proteins, but have received less attention: desmocalmin, desmoyokin, erbin, pinin, p120 catenin, p0071 (Garrod and Chidgey 2008). The Desmosomal Cadherins Desmocollin and desmoglein are calcium-dependent cell–cell adhesion proteins that belong to the cadherin superfamily. There are four desmoglein isoforms and three desmocollin isoforms that are expressed in a tissue-specific and differentiation-dependent manner (Dusek et al. 2007). The extracellular domain of these desmosomal cadherins consists of cadherin repeat domains (EC1–5) that are linked by flexible calcium-binding regions. As has been described for other cadherin family members, the adhesive site has been proposed to be in the EC1 domain of the desmosomal cadherins, and blocking peptides against this region inhibit homophilic and heterophilic interactions of desmocollin and desmoglein (Runswick et al. 2001). While the adhesive function of the desmosomal cadherins usually depends on the presence of calcium, Garrod et al. described calcium-independent “superadhesive” desmosomes that no longer require the presence of calcium to be maintained (Garrod and Kimura 2008). The membrane proximal EC5 domain (or EA domain) is less

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conserved between the proteins and may allow interactions between distinct cadherin family members (Getsios et al. 2004). The membrane spanning (TM) domain is followed by a cytoplasmic juxtamembrane domain (intracellular anchor [IA]), which has been proposed to mediate interaction with plakophilins (Bonne et al. 1999; Hatzfeld et al. 2000; Chen et al. 2002), and the intracellular cadherin sequence (ICS), which has been shown to bind plakoglobin (Wahl et al. 2000; Andl and Stanley 2001; Gaudry et al. 2001). In addition to the domains described above, the desmogleins have additional unique cytoplasmic segments, the repeating unit domains (RUDs), and the desmoglein terminal domain (DTD). The function of these domains remains to be determined. Numerous studies have examined the regulation of the assembly and disassembly of the desmosomal junction and have found that phosphorylation of the cadherin cytoplasmic tail is an important contributor to this process (Calautti et al. 1998; Garrod et al. 2002). Phosphorylation also regulates the interaction of the desmosomal cadherins with the plaque proteins, including plakoglobin (Gaudry et al. 2001). Proteolytic cleavage of the desmosomal cadherins, especially desmogleins, can occur in the extracellular domain (Bech-Serra et al. 2006; Cirillo et al. 2007a; Klessner et al. 2009) or intracellular domain (Dusek et al. 2006; Cirillo et al. 2007b; Nava et al. 2007) and regulates processes such as junction disassembly and apoptosis. In agreement with these findings, at least one human disease, the staphylococcal scalded skin syndrome (SSSS), is caused by cleavage of desmoglein-1, an isoform expressed in the upper layers of the skin. Cleavage of this protein by the bacterial toxin induces massive exfoliation of the upper layers of the epidermis and is often fatal if not treated appropriately (Amagai et al. 2000; Hanakawa and Stanley 2004; Nishifuji et al. 2008). Plakin Family Members The desmoplakin isoforms I and II are the major plakin family members of the desmosome, and immunolocalization of desmoplakin often serves as a reliable marker to identify the desmosomal junction. The desmoplakins are large (210–230 kDa) proteins that are currently thought to be the major plaque proteins that connects the cytoplasmic tail of the desmosomal cadherins to the intermediate filament network. Desmoplakins also interact with armadillo family members, thereby serving as a general scaffold for the association of a number of proteins with the desmosomal junction (Franke et al. 1982; Mueller and Franke 1983). Multiple binding domains allow numerous protein–protein interactions with desmoplakin, including a globular head domain formed by two spectrin repeats, an Src homology domain, and a central coiled-coil domain that mediates desmoplakin dimerization. Other plakin family members that have been localized to the desmosome include envoplakin, periplakin, and spectrin (Garrod and Chidgey 2008). Armadillo Family Members The 42-amino-acid-long “arm” repeat domains were first identified in the polarity gene armadillo of the fruit fly Drosophila melanogaster (Klymkowsky 1999). These domains were subsequently shown to be present in a number of signaling proteins, including betacatenin and other catenin family members. The desmosome contains several members of the armadillo family, including plakoglobin (gamma-catenin) and plakophilins 1–3, and these proteins have been shown to interact with the desmosomal cadherins and/or desmoplakin (Hatzfeld 1999; Garrod and Chidgey 2008). Interestingly, plakoglobin localizes to both the desmosome and the adherens junction, and may allow “crosstalk” between the different junction types. Also, pointing to nondesmosomal signaling roles, the plakophilins demonstrate junctional and nuclear pools in epithelial cells and may integrate signals from

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the cell membrane with changes in cell behavior and tissue morphogenesis (Klymkowsky 1999; Hatzfeld et al. 2000; Bonne et al. 2003). Intermediate Filaments All desmosomes interact with the intermediate filament network, although the cytoskeletal proteins that comprise this network differ depending on the tissue type (Jamora and Fuchs 2002; Owens and Lane 2003). In epithelial cells, the intermediate filaments consist of cytokeratin isoforms, which are differentially expressed based on the type of epithelia (simple vs. stratified) and the differentiation of the cells. For instance, keratin 8 and keratin 18 (K8/K18) are the primary keratins expressed in simple epithelia, K5/K14 in the basal cells of stratified epithelia, and K1/K10 in the suprabasal layers of the epidermis (see Chapter 16). In contrast, in nonepithelial tissues, desmosomes interact with the intermediate filament protein desmin (cardiomyocytes) or vimentin (follicular dendritic cells, meninges).

Tissue Expression of the Desmosomal Cadherin and Desmoplakin Isoforms While numerous tissues are known to form desmosomes, the constituent protein isoforms of desmosomes are not always the same. Currently, it is thought that the obligatory proteins of the desmosome are desmocollin, desmoglein, plakoglobin, and desmoplakin, although the isoforms expressed differ depending on the tissue type. In general, desmoglein-2 and desmocollin-2 are the ubiquitously expressed desmosomal cadherins and are present in all tissues that have desmosomes (Franke et al. 1981a; Cowin et al. 1985). In the skin, these proteins are expressed in the basal layer of the epidermis but are replaced in the suprabasal layer by desmoglein-3 and desmocollin-3 isoforms. In the uppermost layers of the epidermis, desmocollin-1 and desmoglein-1 replace desmoglein-3 and desmocollin-3, and the expression of these proteins is always restricted to the terminally differentiated upper layers of the epidermis and other stratified epithelia. Desmoglein-4 expression is restricted to the upper layers of the skin and hair follicles (Bazzi et al. 2006, 2009; Mahoney et al. 2006). Desmoplakins I and II have been identified in both simple and stratified epithelia, although some confusion remains on whether variant II exists in all epithelia (Franke et al. 1982; Mueller and Franke 1983; Garrod et al. 2002; Garrod and Kimura 2008).

Alterations in Desmosome Function: Lessons from Human Diseases and Knockout Mouse Studies Since the description of the defective “intercellular cement” between acantholytic keratinocytes in patients with pemphigus, alterations in desmosomes and desmosomal proteins have been shown to contribute to disease pathogenesis of a number of conditions affecting the skin, hair, and heart. Inflammatory/Infectious Etiologies In addition to the autoimmune blistering skin diseases pemphigus vulgaris (anti-Dsg3 autoantibodies) and pemphigus foliaceus (anti-Dsg1 autoantibodies), another type of skin disease called the SSSS, leads to disease through the ability of the Staphylococcus aureus exfoliative toxins A and B to specifically cleave desmoglein-1, inducing blister formation and massive skin exfoliation (Amagai et al. 2000; Hanakawa and Stanley 2004; Nishifuji et al. 2008).

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Inherited Desmosomal Diseases Given their widespread distribution, it is not surprising that inherited mutations in desmosomal proteins have been shown to affect mainly the heart, skin, and hair. For instance, numerous mutations in desmosomal proteins have been linked to arrhythmogenic right ventricular cardiomyopathy (ARVC), a heart disease in which patients experience fibrofatty replacement of cardiac muscle tissue, resulting in abnormal electrical conductance and contraction of the muscle. Mutations in desmoglein-2, desmocollin-2, desmoplakin, and plakophilin-2 have all been linked to ARVC. Interestingly, heart disease was the only manifestation of the particular mutations described in these patients, likely due to the serious and often fatal presentation of this disease. Other desmosomal protein mutations have shown a mix of skin, hair, and heart involvement, including a deletion mutation in plakoglobin, which results in Naxos disease: ARVC-like cardiomyopathy, woolly hair, and palmoplantar keratoderma. Mutations in a given gene do not always lead to the same phenotype; for instance, a second type of mutation in desmoplakin leads to striate palmoplantar keratoderma without cardiac involvement (Bazzi and Christiano 2007). Desmosomes and Cancer A number of studies have observed changes in the protein and/or mRNA of desmosomal proteins in tumor samples; however, it is not currently understood what contribution these changes may have on the development, progression, or metastasis of the tumor (Kurzen et al. 2003; Khan et al. 2006; Schmitt et al. 2007; Funakoshi et al. 2008). Knockout Mouse Models While the important role of desmosomes in the skin and heart are further supported by studies using knockout mice, many of the observed phenotypes also suggest that desmosomal proteins have essential functions beyond cell–cell adhesion (Bazzi and Christiano 2007; Green and Simpson 2007). Notably, embryos lacking the ubiquitously expressed desmoplakin or desmoglein-2 exhibit embryonic lethal phenotypes. Surprisingly, desmocollin-3 knockout embryos are also not viable and die before the appearance of desmosomes in the developing embryo (Den et al. 2006). These findings suggest that desmosomal proteins function early in embryonic development perhaps playing a role in embryonic adhesion or in proliferation of the embryonic stem cells (Eshkind et al. 2002).

FUTURE PERSPECTIVES Epithelial tissues provide an important barrier between the external environment and the body’s internal compartments. Proper function and integrity of these tissues are essential, and breakdown in epithelial function underlies the pathogenesis of a number of human diseases. Specialized adhesive contacts between epithelial cells allow the regulation of epithelial permeability (tight junctions) and reinforce cell–cell contacts to increase tissue strength (desmosomes). Furthermore, proteins of these junctions also contribute to regulation of the homeostatic processes of the epithelium, including cell proliferation, differentiation, and apoptosis; however, the mechanisms by which this occurs are not well understood. Additional studies are needed to address several important questions, which apply to both tight junctions and desmosomes: How are these junctions assembled and disassembled? Can assembly or disassembly be modulated as a therapeutic strategy, for instance, to minimize and/or reverse the effects of inflammation, infection, or autoantibodies, or to improve drug delivery through epithelial linings such as the intestine and skin? In addition, and even less understood, is the role of junction proteins in the process of

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tumorigenesis and whether or not these proteins are able to protect against the development and/or progression of cancers. Future studies should further explore the contribution of these proteins to the regulation of basic cellular processes.

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GAP JUNCTIONS Jared M. Churko Dale W. Laird

DEFINITION Gap junctions are protein-enriched aggregates formed between apposing cells that facilitate intercellular communication by allowing the passage of small molecules and metabolites from one cell to another. Virtually all cells found in solid tissues express connexins and assemble gap junctions. Gap junctions are composed of intercellular channels assembled from one or more of the 21 connexin (Cx) subunits named after the molecular weight of each family member (e.g., Cx43 has a molecular weight of 43 kD) (Sohl and Willecke 2004). Six connexins oligomerize within a cell and are transported to the plasma membrane in a configuration known as a connexon (Fig. 20.1). Connexons at the plasma membrane can open and close to exchange molecules with the extracellular environment in a state known as a hemichannel. When one connexon docks with a connexon from an adjacent cell, a gap junction channel is formed. These channels laterally diffuse and aggregate into a semi-crystalline state known as the mature gap junction or a gap junction plaque where they open and close to facilitate intercellular communication (Herve et al. 2007; Goodenough and Paul 2009). The mature gap junction itself consists of tightly aggregated connexin channels, which are thought to extrude all other integral membrane proteins. However, gap junctions are highly dynamic subject to remodeling and rapid turnover. Nucleation, growth, remodeling, internalization, and turnover of the gap junction domain are all critical for establishing the appropriate physiological levels of intercellular communication in all tissues and organs.

HISTORICAL PERSPECTIVE The term gap junction comes from historic electron microscopy studies where a space (seen by an electron opaque region caused by heavy metal infiltration) was observed in a junction-rich region between two closely apposed cardiac myocytes (Revel and Karnovsky 1967). Sometime after this initial observation, low-resolution electron microscopy and X-ray crystallography imaging revealed the first three-dimensional structure of the gap junction channel (Caspar et al. 1977; Makowski et al. 1977). These channels were composed of hexagonal arrays with a central pore embedded into two opposing lipid bilayers. The size of the central pore could be deformed with the addition of the divalent cation chelator EGTA (Unwin and Ennis 1984), and later analysis of isolated Cx26 gap junctions determined that this pore diameter can decrease from 15 Å to 6 Å after the addition of

Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Gap Junction

Hemichannel

Connexosome Golgi Apparatus Proteosome

ERAD

Lysosome

Endoplasmic Reticulum

Nucleus

Figure 20.1. An overview of the life cycle of connexins. Connexins are co-translationally inserted into the endoplasmic reticulum (ER). Misfolded, ER-retained connexins are subjected to quality control mechanisms and exported to proteasomes for degradation via the ER-associated degradation (ERAD) pathway. Properly folded connexins are transported to the Golgi apparatus, where six connexins oligomerize to form a connexon prior to delivery to the plasma membrane, where they may function as hemichannels or dock with connexons from an adjacent cell to form a gap junction channel. Gap junction channels aggregate to form a mature gap junction also commonly referred to as a gap junction plaque. Gap junctions are internalized into specialized double-membrane structures called connexosomes (also called annular gap junctions) prior to delivery to lysosomes for degradation.

calcium (Muller et al. 2002). These structural changes in the pore diameter suggested that gap junction channels can be manipulated to form two functional states (the open and closed state). While higher resolution of the gap junction channel was obtained from X-ray diffraction analysis of Cx43 with a portion of the C-terminal tail truncated (Unger et al. 1999), our understanding of the general arrangement of connexins within the connexon has not substantially changed. Confirmation of the structural organization of the connexon in split gap junctions and other gap junction preparations was acquired with the advent of

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atomic force microscopic imaging where resolution limits of 10 Å were obtained (Hoh et al. 1991; Hoh et al. 1993; Muller et al. 2002; Oshima et al. 2007).

MOLECULAR COMPOSITION OF THE GAP JUNCTION DOMAIN Humans express 21 different members of the connexin family, and the connexin topology in the membrane is essentially the same for all connexin family members (Sohl and Willecke 2003). Four transmembrane domains subdivide the primary connexin structure into a defined N-terminal, C-terminal, two extracellular, and one intracellular domain. These domains have classically been defined as having unique functional roles and can promote the opening and closing of the gap junction channel. Briefly, the extracellular loops facilitate the binding of one connexon to a connexon of an adjacent cell (Herve et al. 2007). The N-terminus contributes to voltage gating (Purnick et al. 2000), while the C-terminus and the distal end of the intracellular loop can regulate the channel by sensing the intracellular pH (Ek et al. 1994; Ek-Vitorin et al. 1996). In addition, high-resolution imaging of connexons composed of Cx26 without an N-terminus suggests that the N-terminal domain can physically plug the gap junction channel (Oshima et al. 2007; Maeda et al. 2009). Phosphorylation of the C-terminal has also been shown to promote the opening and closing of the gap junction channel as well as signaling the gap junction to be both assembled and degraded (Solan and Lampe 2009). With members of the connexin family being expressed throughout multiple cell types and tissues, it is no surprise that multiple connexins expressed in a single cell may interact with each other. Early immunolocalization studies localized two different connexin family members (Cx26 and Cx32) to the same gap junction plaque (Zhang and Nicholson 1994), and these connexins were further found to intermix within the same connexon (Sosinsky 1995; Fig. 20.2). Various combinations of connexins within the gap junction channel can now be classified into distinct groupings (Laird 2006; Goodenough and Paul 2009). Connexons containing only one type of connexin are termed homomeric channels, while mixed connexons of two or more different connexins are termed heteromeric (Fig. 20.2). If a connexon from one cell is docked with a connexon of a different type from an apposing cell, the resulting channel is heterotypic as opposed to homotypic channels, where the same connexon is found in both cells (Laird 2006). Although a wide variety of gap junction channels can be formed by these combinations, not all connexins have the ability to oligomerize with each other. For example, Cx40 and Cx43 (He et al. 1999), Cx43 and Cx45 (Martinez et al. 2002), and Cx26 and Cx32 have all been shown to co-oligomerize, but Cx26 cannot oligomerize with Cx43 (Gemel et al. 2004). Restrictions among which connexins can intermix serve to compartmentalize intercellular communication between cells of different origins or functions. The various connexin permutations within a connexon may also play an important role in many normal and disease cellular environments by selectively regulating the transjunctional molecules that are exchanged. Since gap junction channels composed of only one connexin may be specialized for the passage of a specific solute and another connexin may be specialized for the passage of a different solute, varying the ratio of each connexin subunit within a single channel may promote, or even inhibit, the passage of solutes containing intermediate properties of the two solutes. For example, oocytes expressing heteromeric gap junctions composed of both Cx43 and Cx40 are more sensitive to acidification-induced uncoupling than their homomeric channel equivalents (Gu et al. 2000).

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Homotypic Homomeric

Homotypic Homomeric

Cx43

Heterotypic Homomeric

Cx26

Heterotypic Heteromeric

Cx32

Figure 20.2. Molecular combinations of connexins within a connexon and a gap junction. Connexin family members are often thought to self-assemble to form homomeric, homotypic gap junction channels. Some connexin family members do not intermix within the same channel but may coexist in the same gap junction plaque as is the case for Cx32 and Cx43. In cases where Cx26 and Cx32 are coexpressed, they may form homomeric/heterotypic channels or intermix within the same connexon and even form heteromeric/heterotypic gap junction channels.

G F P -C x4 3 P re b le a ch

B le a ch

2 5 se c

5 0 se c

1 0 0 se c

1 5 0 se c

2 0 0 se c

3 0 0 se c

Figure 20.3. Mobility of Cx43 connexons within a gap junction plaque. A gap junction plaque composed of a GFP-tagged Cx43 mutant was subjected to fluorescent recovery after photobleaching and imaged every 25 seconds after photobleaching. Lateral migration of the GFP-tagged Cx43 mutant into the bleached area suggests that the connexons within the plaque are mobile. Bar = 2 μm.

Once connexons are transported to the plasma membrane, gap junction channels formed from docked connexons aggregate into dense concentrations of tens to hundreds of gap junction channels (Zampighi et al. 1989). Gap junctions are unique and represent a functional domain distinct from any other junctional complex. Given that the average turnover time for connexins and gap junctions is 1–5 hours, gap junctions are in a constant state of change and dynamically respond to the physiological demands of the cell (Laird et al. 1991; Beardslee et al. 1998). Earlier electron microscopy studies suggest that gap junctions mature from a loosely packed state, termed a formation plaque, to a more densely packed semi-crystalline structure (Johnson et al. 1974). Fluorescent recovery after photobleaching of Cx43-containing plaques also suggests that not all gap junctions possess the same fluidity in the gap junction plaques (Simek et al. 2009) suggestive of immature states of assembly. By measuring the recovery of green fluorescent protein (GFP)-tagged Cx43 in a photobleached region of gap junction plaques (as identified by their punctate fluorescence), the mobility of Cx43 within the gap junction plaque can be classified into two distinct mobile states (Fig. 20.3). Highly mobile GFP-tagged Cx43 may represent imma-

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ture nascent plaques, while low mobility Cx43 may represent mature densely packed plaques. In addition, the binding of scaffolding proteins to connexins within the gap junction plaque may regulate the surface dynamics of connexons within a gap junction plaque (Laird 2010). For example, the binding of ZO-1 to the distal C-terminal tail of Cx43 has been reported to regulate the gap junction plaque size, and this interaction may further regulate the turnover or stabilization of gap junctions at the plasma membrane (Maass et al. 2007). Distinct properties of gap junctions can also be attributed to the lipid composition present in the plaque environment. Gap junction plaques are resistant to most detergents, and connexons can be seen in preparations used to isolate lipid rafts (alkaline carbonate extraction) (Locke et al. 2005). Thus, the lipid environment may regulate distinct gap junction assembly stages as part of the dynamic life cycle of connexins. Lipid–connexin interactions may also be dependent on which connexin is expressed or the structural and folding state of the connexin (hemichannel or gap junction state). For example, Cx32 hemichannels specifically interact with L-α-phosphatiylcholine, while Cx26 hemichannels prefer to interact with L-α-phosphatiylserine. In a fully assembled gap junction, however, Cx26 and Cx32 channels lose most of their selectivity for specific lipids (Locke and Harris 2009). Gap junction formation and function is also linked to the cholesterol content in subdomains of the plasma membrane. Early studies have shown that gap junction plaques are rich in cholesterol (Hertzberg and Gilula 1979), and cholesterol extraction was found to have a profound impact on the structural integrity of the plaques (Henderson et al. 1979). The importance of cholesterol within gap junction plaques is also supported by the fact that cells supplemented with 20 μM cholesterol can increase their gap junction plaque content by sixfold (Meyer et al. 1990). This increase was also accompanied by an increase in intercellular dye transfer indicating that more functional channels had formed. Thus, a specific plasma membrane cholesterol concentration may be required for optimal gap junction activity. In liposome studies, the formation of gap junction channels was found to be biphasic and dependent on cholesterol concentration (Locke and Harris 2009).

GAP JUNCTION GROWTH, INTERNALIZATION, AND RENEWAL Once gap junction plaques are formed at the cell surface they remain in a dynamic state subject to remodeling, growth, and renewal. Two mechanisms are proposed to explain how connexins are added to the gap junction plaque. One mechanism supports the lateral diffusion of connexons to the periphery of an existing gap junction plaque where they dock with connexons from an adjacent cell, while other evidence supports a more directed targeting of connexons to the gap junction plaque where interconnexon docking occurs. The first mechanism is supported best by a FlAsH/ReAsH pulse-chase study where older connexins were labeled with one color and younger connexins were labeled with a second color. Visualization of newly formed gap junction channels was clearly localized along the periphery of the gap junction plaque, while older connexins were found within the interior of the gap junction plaque (Gaietta et al. 2002). This overall mechanism is supported by a number of other studies that would argue for a lateral diffusion of connexons at the cell surface to sites of nucleated gap junctions (Jordan et al. 1999; Windoffer et al. 2000; Lauf et al. 2002). Evidence for the second proposed mechanisms is limited to one study where rapid time-lapse imaging suggested that yellow fluorescent protein (YFP)-tagged Cx43 was targeted to gap junction plaques along microtubule cables, where

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they anchored to adherens junctions in close apposition to gap junctions (Shaw et al. 2007). The maximum size of a gap junction is governed by properties that remain poorly understood. Some cells have very small gap junctions, while other cells have gap junctions that are more than 2 μm in diameter (Zampighi et al. 1989). Undefined processes regulate the removal of older connexons from the gap junction plaque and entire gap junctions can be removed in a single process. Evidence supporting selective removal of older components (more central regions) of the gap junction plaque have been documented (Gaietta et al. 2002) suggesting that the plaque is subject to distinct regulatory mechanisms. This event involves the vesicular invagination of part or even the entire gap junction plaque to form a double-membrane structure called an annular junction or more recently defined as a connexosome (Larsen et al. 1979; Naus et al. 1993; Jordan et al. 2001; Laird 2006). These connexosomes are unique in that they contain not only connexons from the cell that produced them but also remnants of the plasma membrane and connexons produced by the neighboring cell. Internalized connexosomes are typically delivered to lysosomes, where they are degraded (Naus et al. 1993; Jordan et al. 2001; Leithe et al. 2009). This unusual pathway of gap junction internalization is relatively unprecedented and not observed for other junctional complexes. However, there is circumstantial evidence that suggests more classical pathways of canonical endocytosis of disassembled gap junction plaques may also exist, but our understanding of alternate pathways remains rudimentary.

SIGNIFICANCE OF THE GAP JUNCTION DOMAIN Since gap junction channels can function in isolation, why are they typically found in tightly packed clusters? This is an intriguing question that may be rooted in several possibilities. First, some evidence would suggest that gap junction channels acquire a more stable state when clustered as opposed to being distributed throughout the plasma membranes (Bukauskas et al. 2000). Given that the extracellular loops of adjacent connexins must tightly seal to prevent transjunctional and small molecule leakage to the extracellular surface, the need to have many closely associated channels could exponentially increase the stability of this interaction. Second, a localized concentration of gap junction channels within a gap junction plaque may also provide an efficient mechanism by which gap junctions can be cleared from the plasma membrane. For example, during mitosis, gap junctions are rapidly removed from the plasma membrane and intercellular communication with neighboring cells is abolished (Stein et al. 1992; Xie et al. 1997). Localizing gap junction channels to distinct regions of the plasma membrane allows for the rapid removal of numerous channels quickly in one process through the formation of connexosomes. Third, gap junction channels may work cooperatively when clustered into gap junction plaques. In HeLa cells expressing Cx30, electrophysiological recording determined that single channels are inactivated slower at a specified transjunctional voltage when compared to multichannel combinations (Valiunas et al. 1999; Valiunas and Weingart 2001). Finally, gap junction plaques are frequently found in close approximation with adherens and tight junctions, and these structural junctions may provide a microenvironment more amenable to both gap junction formation and stability. In addition, connexins and molecular components of both tight and adherens junctions (Chapters 18 and 19) share common binding and scaffolding partners, creating a framework for molecular cross-talk among these junctional complexes (Laird 2010). Furthermore, evidence supports the concept that N-cadherin-based cell adhesion precedes Cx43 gap junction formation in rat cardiomyocytes (Hertig et al. 1996; Kostin et al. 1999) and cardiomyoctes from N-cadherin null mice

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have reduced levels of Cx43 (Luo and Radice 2003). In the end, the need for gap junction channels to be in tightly packed aggregates may be due to any one or a combination of these reasons that currently extend beyond our full understanding.

FUTURE DIRECTIONS In recent years, the gap junction community has become increasingly aware that connexins neither exist nor function in isolation. The emergence of the gap junction proteome that exceeds over 40 molecules highlights the complexity of these specialized junctions (Laird 2010). The gap junction proteome consists of molecules that posttranslationally modify connexins, scaffold connexins to cytoskeletal elements, regulate connexin trafficking and assembly, and link connexins to growth control and other regulatory mechanisms. Without doubt, the gap junction proteome is far from complete and has focused primarily on only the Cx43 member of the connexin family, leaving many important discoveries to be found. A second area in gap junction biology that has come to the forefront has been the increasing understanding that the function of connexins extends beyond regulatory mechanisms associated solely with cell–cell communication. Considerable evidence now suggests that connexins can function in growth control by mechanisms that are connexin dependent but intercellular communication independent (Ionta et al. 2009). Likewise, the molecular cross-talk of how connexins may regulate their binding partners is as valid as the focus on how binding partners regulate gap junctions. Importantly, some of these intercellular communication-independent mechanisms may require connexins to be spatially localized to gap junction plaques. A third area of interest is related to the plethora of diseases that are now linked to germ-line mutations in the genes that encode connexins. These diseases range from rare diseases such as oculodentodigital dysplasia associated with mutant Cx43 expression to high-frequency diseases such as neurosensory hearing loss caused by the expression of mutant Cx26. Intriguingly, the majority of the diseases associated with connexins are linked to single missense mutations (Laird 2008; Paznekas et al. 2009), and most of the mutants investigated to date travel to the cell surface and assemble into the gap junction plaque but fail to function in intercellular communication (McLachlan et al. 2005; Gong et al. 2006; McLachlan et al. 2008; Chtchetinin et al. 2009). Thus, the nature of the disease is linked to malfunctions in connexin assembly into functional gap junctions and not to the delivery of the mutant connexins to the correct plasma membrane microdomain. Lastly, with the increase in success of obtaining high-resolution structural information (Maeda et al. 2009), it will be important to fully understand the regulatory aspects associated with pore formation, channel opening and closing, the regulatory properties of the pore, the pore lining residues, and the transjunctional molecules that truly pass through gap junction channels. With regard to transjunctional passage of molecules in a physiological setting, this remains one of the most pressing questions in gap junction biology. In the absence of this knowledge, our understanding of connexin biology will remain incomplete.

ACKNOWLEDGMENTS The authors would like to thank Jamie Simek for contributing the data presented in Figure 20.3. This work was supported by Canadian Institutes

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PO L ARI T Y commonly refers to the formation of distinct plasma membrane domains. The best studied examples of polarized cells are the apicobasal polarity of the epithelial cell (Chapter 21) and axonal–dendritic polarity of the neuron (Chapter 22). The apical and basolateral domains of epithelial cells present distinct protein and lipid compositions that are critical to the vectorial transport functions of this cell type; epithelial polarity is also dependent on the formation of epithelial junctions, such as adherens junctions, tight junctions, and desmosomes, that also contribute to the barrier function of this cell type (Chapters 18, 19, and 21). Apical specializations such as microvilli (Chapter 13) and cilia (Chapter 15) provide another example of “domains within domains.” Similarly, neurons present not only somatic, dendritic, and axonal domains but also a variety of specialized axonal, presynaptic, and postsynaptic domains (Chapter 22). Indeed, it is important to recognize that essentially every cell, even the so-called nonpolarized cells, exhibits polarized cellular domains, such as the yeast bud (Chapter 11) and the pseudopodia (lamellipodia and filopodia) of migrating fibroblasts (Chapter 12) that are critical to cellular function and specialization. The formation of these domains is critically dependent on the establishment of stabilized cell–cell and cell–substrate contacts (Chapters 17–19), the cytoskeleton (Chapters 12, 14, and 16), and polarized trafficking and recycling from the Golgi apparatus and endosomes (Chapters 8 and 9).

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EPITHELIAL DOMAINS Nancy Philp Liora Shoshani Marcelino Cereijido Enrique Rodriguez-Boulan

DEFINITION Epithelial cells display distinct apical (exposed to the external environment) and basolateral (exposed to the internal medium) plasma membrane (PM) domains, separated by tight junctions. These PM domains are populated by over 1000 gene products (solute transporters, pumps, channels, hormone receptors, etc.) that represent over 5% of the human genome and are highly polarized to one or the other domain. The polarity of individual PM proteins is highly variable among different epithelia; for example, Na,KATPase is basolaterally localized in most epithelia but is apically localized in retinal pigment epithelium (RPE) and choroid plexus (CP). The variable polarity phenotype of over 100 different epithelial cell types present in metazoa allows them to contribute specific vectorial functions in absorption and secretion required by their host tissues or organs, for example, tegument, mucosal membranes, and digestive, hepatic, respiratory, urinary, reproductive, endocrine, and exocrine organs. Here, we review the mechanisms responsible for the variable functional polarity of different epithelia, with a focus on the variable polarity of transporter proteins, which remains poorly understood. To this end, we discuss the mechanisms responsible for the apical/basolateral localization of Na,KATPase and lactate transporters, as model systems for future studies on the sorting of other transporters.

APICAL–BASOLATERAL POLARIZATION IN EPITHELIAL CELLS The Central Role of Na,K-ATPase in Transporting Epithelia The internal medium of metazoa displays ∼140 mM Na+, a fossil record of the Na+ levels in the primitive sea at the time of their appearance on earth about 700 million years ago (Cereijido et al. 2004). Physiologists have wondered for over a hundred years how amphibians thrive in a pond where Na+ levels are 10–100× lower than in the internal medium. Already in the 1850s, Émile Du Bois Raymond had shown that frog skin can generate an

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electrical potential, that in 1905, Galeotti attributed to a higher Na+ permeability in the inward than in the outward direction (Fig. 21.1A). Although this concept appeared to violate the second law of thermodynamics, it was proven to be true by Koefoed-Johnsen and Ussing (Koefoed-Johnsen and Ussing 1958), a few years after sodium radioisotopes were developed as a subproduct or research on the atomic bomb (Rotunno et al. 1970; Cereijido et al. 2001) (Fig. 21.1A). The key molecule in this transport process turned out to be Na,K-ATPase, first purified by Skou (Jorgensen and Skou 1969), which extrudes three Na+ ions in exchange for two K+ ions, using the energy provided by the hydrolysis of ATP (Skou 1982). Key aspects of the Koefoed-Johnsen and Ussing model are (Fig. 21.1B) the following: (1) Na,K-ATPase is basolaterally localized and its activity generates low intracellular Na+ levels ([Na]i), which, (2) together with the high Na+ permeability and low K+ permeability of the apical membrane, allow the entry of NaCl down a favorable electrochemical gradient; and (3) the basolateral membrane has high permeability (but low conductance) for K+, which contributes to a transepithelial electrical potential positive on the blood side and high permeability to Cl−; both factors are necessary for the net transport of NaCl into the internal medium. The Koefoed-Johnsen and Ussing model was extremely successful as a blueprint to explain epithelial transport and spawned half a century of studies that have elucidated major mechanisms underlying polarized ion transport and confirmed the central role of the Na,K-ATPase in ion and solute transport across epithelia (Fig. 21.1C). These studies showed that epithelial cells employ the Na+ gradient generated by the electrogenic function of the Na+ pump to transport sugars, amino acids, lactate, protons, lactate, Ca2+, and so on, across the apical membrane through a wide variety of Na+/glucose, Na+/amino acid, Na+/lactate, Na+/Ca2+ cotransporters, and Na+/H+ and Na+/HCO3− exchangers (Schultz and Curran 1970; Frizzell et al. 1979; Broer 2008) (Fig. 21.1C). Furthermore, the transport of Na+ creates the osmotic force necessary for the vectorial movement of fluid, another key function of epithelia, which is facilitated by a family of water channels, the aquaporins (AQPs) (Fig. 21.1C) (Diamond and Bossert 1967; Nielsen et al. 2002; Verkman 2009). Hence, it is clear that sodium transport across epithelia is important not only because of the intrinsic biological value of this ion, but also because its vectorial movements establish electrochemical and osmotic gradients across different poles of the PM that provide the driving force for the transport of a wide variety of essential nutrients.

Variable Polarity of PM Proteins in Different Epithelia Epithelial cells carry out a vast number of functions essential for the organism through a variety of nutrient, hormone, and immune receptors; enzymes; and active and passive transporters for ions, solutes, organic molecules, peptides, and vitamins. It is clear now that ∼5% of the genome (∼1000 genes) encodes transporter proteins or related proteins, including solute carriers (SLCs), ion and water channels, and ATP-dependent transporters, for example, the ATP-binding cassette (ABC) family and the ion pumps (e.g., Na+, H+, Cu++, Fe++ ATPases) (Hediger et al. 2004). In epithelial cells, these proteins are expressed in a polarized fashion at apical or basolateral PM domains, separated by tight junctions (see Chapter 19). Different epithelial cells vary dramatically the localization of individual receptors and transporters in order to perform specific functions required by the host organ. Figure 21.2 illustrates this variation in three epithelia. The first one is the kidney’s proximal tubule (PT), an absorptive epithelium that shares many organizational features with the small intestine. The human kidneys filter the blood by producing daily over 180 L of ultrafiltrate. Two-thirds of this volume is reabsorbed by the PT in a constitutive fashion. To this end,

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Figure 21.1. Central role of Na, K-ATPase in the functional asymmetry of epithelia. (A) In the second half of the nineteenth century, Du Bois Raymond observed that the frog skin establishes an electrical potential difference between its outer and inner side. In 1905, Galeotti proposed that this potential (red) can be accounted for by a higher Na+ permeability in the pond-to-blood than in the blood-to-pond direction (left to right in the figure). His interpretation was rejected on the basis that if the frog skin were mounted in a doughnut-shaped chamber with saline solution, it would originate a net transport and constitute a perpetuum mobile in violation of the laws of thermodynamics (blue arrows). (B) Half a century later, the use of radioisotopic tracers of Na+ demonstrated that, in fact, the inward flux of this ion is 20-odd times higher than the outward one. In 1958, Koefoed-Johnsen and Ussing proposed a model for the frog skin, where the pump (an Na+,K+-ATPase) was polarized to the basal domain of the epithelial cell. This model acted as blueprint for most transporting epithelia. (C) Research in the past five decades has identified hundreds of transporters, including pumps, channels, and solute transporters that are localized in a polarized fashion in epithelial cells. The Na,K-ATPase remains the central protagonist of transepithelial ion and solute transport, as it generates the critical Na+ gradients necessary for the function of these transporters. CLC, chloride channel; NBE, sodium bicarbonate exchanger; NDC, Na-dependent cotransporters of ... ; CFTR, cystic fibrosis transmembrane conductance regulator; NBC, sodium bicarbonate cotransporter.

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(A)

(B) Lumen Na+ K+ Cl− CO3H− Lactate

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Blood side Na+= 140 mM Cl− = 115 mM CO3H− = 27 mM Lactate = 1 mM Figure 21.2. Variable configuration of transporter polarity in secretory and absorptive epithelia. Different epithelial cells vary the localization of Na,K-ATPase, Na+, K+, Cl−, bicarbonate, and water transporters and channels in order to perform functions specific for each organ and tissue. (A) The kidney proximal tubule (PT) absorbs daily 120 L of glomerular ultrafiltrate by localizing Na,K-ATPase basolaterally, the Na/proton exchanger NHE3 apically, and a complement of Cl− and K+ channels either apically or basolaterally. The Na+ gradient generated by these transport activities drives fluid absorption. (B) The choroid plexus (CP) utilizes apical Na,K-ATPase and a different combination of Na+, Cl−, K+, and bicarbonate channels to secrete choroidal spinal fluid into the brain ventricle. Chloride and bicarbonate gradients drive fluid transport in this case. (C) The retinal pigment epithelium (RPE) localizes Na,K-ATPase apically, like CP. The Na+ pump secretes Na+ into and removes K+ from the subretinal space, which is necessary for the dark current of the photoreceptors. However, net transport of fluid occurs in the apical to basal direction driven by Cl− and CO3− gradients (see text). NBCE, sodium chloride bicarbonate exchanger.

the PT organizes its polarity along the classical Koefoed-Johnsen and Ussing model. The Na,K-ATPase is localized basolaterally, and the apical uptake of Na from the ultrafiltrate is carried out by an Na/proton exchanger (NHE3). The strategic localization of several K+, Cl−, bicarbonate, and water channels completes the basic configuration to transport fluid in the apical to basal direction. A large number of Na+-dependent solute transporters at the apical surface utilize the favorable Na+ gradient to transport amino acids, glucose, vitamins, peptides, and so on, into the PT cell. A complementary set of Na+-independent transporters at the basolateral membrane (e.g., Glut 2 for glucose) moves them into the blood. More distal regions of the nephron and the collecting tubules and ducts use the same basic configuration, with a variable complement of Na+ transporters, for example, NHE (thick ascending limb), the thiazide-sensitive Na+/Cl− cotransporter NCC (thick ascending limb), and the amiloride-sensitive epithelial sodium transporter (ENaC) (collecting duct). In the collecting duct, α and β cells display V-type proton ATPase at apical or basolateral domains, respectively, to transport protons and bicarbonate in opposite directions, ready to respond to variations in the pH of the internal medium of the organism (Brown et al. 2009).

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The choroid plexus (CP) epithelium (Fig. 21.2B) is a secretory epithelium, so called because it transports a net amount of fluid in the basal to apical direction, that is, from the blood toward the ventricle, to produce 500 mL per day of cerebrospinal fluid (CSF), necessary for the normal function of the brain. The CSF’s ion and nutrient composition is very similar to the blood. In order to transport fluid apically, CP localizes Na,K-ATPase apically and coordinates its Na pumping activity with strategically localized bicarbonate, Cl− and Na+ transporters and with the activity of carbonic anhydrases that promote the intracellular formation of bicarbonate (Jacobs et al. 2008). As in the kidney PT, Na+ and bicarbonate gradients constitute the driving force for the movement of fluid, which is facilitated by AQP1 (Wolburg and Paulus 2010). The RPE (Fig. 21.2C), like CP, is a neuroepithelium. However, unlike most other epithelia, the apical PM of RPE is not free, but is in contact with 30–50 photoreceptors (rods and cones), separated by the subretinal space (SRS). A major task of the RPE is to generate the correct ionic environment for the function of the photoreceptors, controlling tightly the subretinal K+ and Na+ levels in order to generate the dark current, essential for vision. Like the CP, the RPE localizes Na,K-ATPase at the apical surface; however, unlike CP and like the kidney PT, the RPE transports a net amount of fluid in the apical to basal direction, creating a negative pressure that helps attach the neural retina to the RPE (Adijanto et al. 2009). As in both CP and PT, this task requires coordinating Na,K-ATPase activity with those of bicarbonate and Cl− transporters. Diurnal variations in the activity of the retina result in large changes in the levels of Na+, K+, and CO2 in the SRS, which are balanced by changes in the activity of the various Na+, K+, Cl−, and bicarbonate transporters (Fig. 21.2C). Fluid transport in RPE is driven by the net transport of Cl− and Na+ in the apical to basal direction and may be facilitated by AQP1 (Strauss 2005), although this issue is still controversial (S. Miller, pers. comm.). Lactate, produced in large amounts by the retina and a major food source for photoreceptors, contributes to fluid transport into RPE (swelling) but not to net transepithelial fluid transport (Philp et al. 1998; Hamann et al. 2003; Adijanto et al. 2009; Miller, pers. comm.). Lactate levels in the SRS are 3–10× higher than in the blood (Strauss 2005) and is transported efficiently by proton-coupled transporters at the apical and basolateral membrane, respectively, H+/Lac− cotransporter (MCT) 1 and MCT3 (Philp et al. 1998). During the past two decades, expression cloning has resulted in the identification of hundreds of apical and basolateral pumps, channels, and transporters in epithelia (Fig. 21.1C). However, the mechanisms utilized by the cell to localize these transporters at the correct PM domain and to vary their localization in different epithelial cells remain largely unknown. On the other hand, studies with model PM proteins consisting of viral envelope glycoproteins, nutrient receptors, and a few transporters in model epithelial cell lines have begun to elucidate the sorting compartments, intracellular trafficking routes, and some of the sorting mechanisms utilized by epithelial cells to segregate apical and basolateral PM proteins. These studies have revealed that epithelial cells utilize, rather than a simple binary mechanism to sort apical and basolateral proteins, a dizzying repertoire of apical and basolateral targeting mechanisms. This multiplicity of sorting mechanisms is ontologically explained by the requirement of different epithelia for maximal flexibility in localizing individual transporters apically or basolaterally to perform their tissue- or organ-specific functions. Here, we will briefly review the apical–basolateral sorting mechanisms emerging from studies of model proteins in the prototype epithelial cell line Madin-Darby canine kidney (MDCK). Then, we will discuss recent insights on the very different mechanisms employed by epithelial cells to sort the Na,K-ATPase and proton/monocarboxylate transporters, as models for future studies on the sorting of other oligomeric epithelial transporters and channels.

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MECHANISMS TO LOCALIZE PM PROTEINS APICALLY OR BASOLATERALLY General Strategies Epithelial cells use three major strategies to localize transporters to apical or basolateral domains: 1. Expression of differently spliced isoforms. This strategy is used to change the localization of the Na+, K+, 2Cl− cotransporter (NKCC) 1 and AQPs (Brown and Nielsen 2008; Carmosino et al. 2008). NKCC1 expression in the basolateral membrane of most epithelia is directed by a dileucine motif found in the C-terminal cytoplasmic tail. The dileucine motif is encoded by optionally spliced exon 21. In most epithelia, NKCC1 is expressed with exon 21 but not in neuroectoderm-derived epithelia, thus explaining the apical distribution of NKCC1 in CP and RPE. 2. Expression of different combinations of subunits. Most transporters are either homoor hetero-oligomers. For example, the family of P-type ATPases, to which Na,KATPase belongs, as well as many amino acid, sugar, and lactate transporters, are heterodimers of a single-pass highly glycosylated subunit that acts as a chaperone for intracellular transport, and a multispan transmembrane subunit that constitutes the functional transporter. As we will discuss later in the review, epithelial cells may change the apical/basolateral localization of Na,K-ATPase by expressing differently glycosylated β-subunits or may change the localization of different lactate transporters depending on sorting signals present in the α- or β-subunits. 3. Differential expression of targeting machinery. As the molecular details of the apical and basolateral trafficking machineries are elucidated, their variations in different epithelial cells are starting to be uncovered. Recent studies have shown that lack of expression of the epithelial specific clathrin adaptor AP1B in certain epithelia, for example, RPE and kidney PT, results in apical expression of a subset of PM proteins that are basolaterally expressed in AP1B-positive epithelia (Diaz et al. 2009; Schreiner et al. 2010).

Trafficking Machinery Involved in Apical–Basolateral Localization The study of the trafficking machinery and the cell biological bases of epithelial polarity became possible with the introduction in the 1970s of the MDCK model (Cereijido et al. 1978) and the subsequent development of a large number of epithelial cell lines and primary cultures from various epithelial origins. This is an ongoing process, as new techniques are developed to facilitate the establishment of cell cultures that preserve, as close as possible, the phenotype of the original tissue. The MDCK model allowed for the first time the reconstitution of an epithelium from its individual cells. A typical protocol involves harvesting the cells with trypsin ethylenediaminetetraacetic acid (EDTA), followed by plating them on a permeable substratum (Fig. 21.3A). Cell dissociation promotes the rounding up of the cells, with loss of the surface polarity of their pumps and channels. After the cells are plated at confluency, they reform their tight junctions and progressively reconstitute their trafficking mechanisms and surface polarity over the following 12–15 hours (Fig. 21.3A). Using this type of approach and ouabain binding to study the repolarization of the Na,K-ATPase in MDCK cells, it was possible to show that after rapid reformation of the tight junctions, the Na+ pump remains depolarized by trapping of a fraction

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Figure 21.3. Mechanisms responsible for epithelial polarity. (A) Contribution of the MDCK model. The introduction of the MDCK model in 1978 (Cereijido et al. 1978) allowed for the first time the study of the reformation of an epithelium from its individual cells. When the cells are plated at confluency on a permeable substratum, they reform their tight junctions in 10–12 hours, resulting in the generation of transepithelial electrical resistance (TER). (B) Polarized viral budding in MDCK cells. Influenza virus and vesicular stomatitis virus (VSV) bud asymmetrically from apical and basolateral PM domains of MDCK cells, driven by their envelope glycoproteins influenza HA and VSVG (Rodriguez-Boulan and Sabatini 1978; Rodriguez-Boulan and Pendergast 1980). (C) Studies with virus-infected MDCK cells identified for the first time apical and basolateral trafficking routes and the key sorting role of the Golgi complex and the trans-Golgi network (TGN). Subsequent studies with viral envelope glycoproteins and single-span transmembrane receptors, such as low-density lipoprotein (LDL), polymeric IgA receptor, and transferrin receptor elucidated apical and basolateral recycling routes and various apical and basolateral sorting signals and mechanisms involved in apical delivery (e.g., lipid rafts) and basolateral delivery (e.g., clathrin and clathrin adaptors). Very little is known about the sorting signals and mechanisms directing oligomeric ion and water transporters to apical or basolateral PM domains. ARE, apical recycling endosomes; ASE, apical sorting endosomes; BSE, basolateral sorting endosomes; CRE, common recycling endosomes; TJ, tight junction.

of the surface population in the apical domain, but recovers its basolateral polarity progressively after internalization and recycling (Contreras et al. 1989). A similar situation is observed for K+ channels expressed by MDCK cells, Maxi K, Kv1.6, and Kv1.7, although in this case, restoring the polarized distribution of the channels requires de novo protein synthesis (Garcia-Villegas et al. 2007). The study of the intracellular routes and sorting compartments involved in apical– basolateral PM protein sorting in MDCK cells was initially made possible by the use of enveloped RNA viruses, for example, influenza and vesicular stomatitis virus (VSV) (Rodriguez-Boulan and Sabatini 1978; Rodriguez-Boulan and Pendergast 1980). The envelope proteins of these viruses are expressed in large amounts in infected cells and are sorted by epithelial cells using the same compartments and mechanisms used for their own PM proteins. These studies showed that apical and basolateral PM proteins are synthesized in the endoplasmic reticulum (ER) and sorted in the trans-Golgi network (TGN) into different carrier vesicles for delivery to their corresponding PM domains (Fig. 21.3B) (see also Chapter 8). Studies in the 1980s also defined the endocytic and recycling routes of

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MDCK cells and the major endosomal compartments, highlighting the major role of the common recycling endosome (CRE) in sorting PM proteins in the recycling route (Fig. 21.3C) (see also Chapter 9). Studies in the last decade have shown that TGN and endosomes cooperate in the sorting of PM proteins in both biosynthetic and recycling routes. Indeed, many newly synthesized PM proteins follow a transendosomal route from the TGN to the PM, with the final sorting decision made at the level of different endosomal compartments (Gonzalez and Rodriguez-Boulan 2009; Weisz and Rodriguez-Boulan 2009). With the introduction of DNA cloning and transfection techniques in the 1980s, it became possible to study the sorting signals of viral glycoproteins, glycosylphosphatidylinositol (GPI)-anchored proteins and a variety of model single-span PM receptors (Mostov et al. 1992; Rodriguez-Boulan et al. 2005) (Fig. 21.3C). Typically, apical sorting signals are complex and may be found in either the luminal, transmembrane, or cytoplasmic domains of the PM protein (Weisz and Rodriguez-Boulan 2009). The first apical sorting signal identified was GPI, which mediates attachment of GPI-anchored proteins to the PM (Lisanti et al. 1988). Since GPI has affinity for PM microdomains enriched in glycosphingolipids and cholesterol, termed “lipid rafts,” experiments showing that GPIanchoring promoted apical targeting of chimeric proteins (Brown et al. 1989; Lisanti et al. 1989) provided the first experimental support for the lipid raft hypothesis for apical sorting (Simons and van Meer 1988). Subsequent experiments demonstrated that other apical proteins, for example, influenza hemagglutinin, interacted with lipid rafts (Skibbens et al. 1989). The lipid raft concept is still evolving (see Chapters 4, 5, and 8): Current ideas indicate that they are formed in the Golgi complex as very dynamic structures that require aggregation into larger rafts via a variety of proteins (e.g., the tetraspanin MAL1 and certain lectins) to act as sorting platforms (Puertollano et al. 1999; Fullekrug and Simons 2004; Weisz and Rodriguez-Boulan 2009). Lipid raft association is not a universal mechanism for apical trafficking, as many apical proteins are not associated with lipid rafts; some apical proteins are sorted through direct interaction with microtubule motors, for example, rhodopsin (Sung and Tai 2000). In contrast with apical sorting signals, basolateral sorting signals are short peptide sequences most often found within the cytoplasmic domain of the protein (Gonzalez and Rodriguez-Boulan 2009). Some basolateral sorting signals resemble endocytic signals, for example, variations of the canonical endocytic dileucine, YXXΦ, and NPXY motifs (Bonifacino and Traub 2003). Other basolateral signals are unrelated to endocytic signals, for example, the tyrosine motifs in low-density lipoprotein receptor (LDLR) (Matter et al. 1994) and VSVG protein (Thomas et al. 1993), the glycine–aspartic acid–asparagine– serine (GDNS) motif of transferrin receptor (TfR) (Odorizzi and Trowbridge 1997), the multicomponent basolateral signal of polymeric immunoglobulin (Ig) receptor (Reich et al. 1996), the monoleucine motifs found in CD147 (Deora et al. 2004) and stem cell factor (Wehrle-Haller and Imhof 2001), the EXEXΦΦ motif found in the M3 muscarinic receptor (Iverson et al. 2005), the PXXP motif in the epidermal growth factor (EGF) receptor (He et al. 2002), and the PDZ-binding domains in syndecan-1 (Maday et al. 2008). Recently, it has become clear that clathrin plays a fundamental role in basolateral sorting, comparable to its key role in endocytosis from the PM (Deborde et al. 2008) (see Chapter 2 on clathrin-coated pits). Clathrin does not interact directly with endocytic or basolateral proteins but, rather, through a variety of clathrin adaptors (Bonifacino and Traub 2003). To date, only one clathrin adaptor has been shown to be involved in basolateral protein sorting: the epithelial-specific AP1B (Ohno et al. 1999; Folsch et al. 2001). The family of heterotetrameric adaptor proteins (APs) displays organelle-specific distributions; AP1B sorts basolateral PM proteins at recycling endosomes in both the biosynthetic and recycling routes (Gan et al. 2002; Cancino et al. 2007; Gravotta et al. 2007). The ubiquitous,

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highly related AP1A adaptor, which differs from AP1B in the medium subunit (μ1A instead of μ1B), mediates basolateral PM protein sorting proximally to AP1B, probably at the level of the TGN (Gravotta et al., manuscript in revision). It appears that, although clathrin and clathrin adaptors regulate the localization of many basolateral proteins, there are alternative sorting adaptors involved in basolateral PM localization. To date, these include Naked, involved in basolateral sorting of some EGF receptors (Li et al. 2004) and ankyrin G, which participates in cooperation with β2-spectrin in basolateral sorting of E-cadherin (Kizhatil et al. 2007). Unlike the PM proteins discussed above, PM transporters and channels are often multimeric. This single fact has complicated the analysis of their intracellular sorting mechanisms, which to date remain largely unexplored. Here, we discuss the mechanisms involved in the sorting of two relatively well-characterized transporters, Na,K-ATPase and lactate transporters, studied in our laboratories. These proteins are sorted by very different mechanisms, thus providing interesting models for future studies.

MECHANISMS INVOLVED IN THE SORTING OF NA,K-ATPASE Intracellular Sorting versus Retention at the PM by the Ankyrin–Spectrin Cytoskeleton Early studies demonstrated that the Na+ pump, comprised of α- and β-subunits, is sorted in the TGN and delivered directly to the basolateral membrane without significant appearance at the apical surface in some strains of MDCK cells (Caplan et al. 1986; Zurzolo and Rodriguez-Boulan 1993; Gottardi and Caplan 1993a). As Na,K-ATPase and H-ATPase are highly homologous ion pumps, yet in LLC-PK1 cells the former is polarized to the basolateral domain and the latter to the apical plasma membrane, studies were carried out to study the polarized expression of chimeric constructs of the α-subunit of H-ATPase and Na,K-ATPase in LLC-PK1 cells. These studies identified apical sorting information within the fourth transmembrane domain of the α-subunit of H-ATPase that is sufficient to redirect the normally basolateral Na,K-ATPase to the apical surface of these cells (Gottardi and Caplan 1993b). However, it remains unclear whether basolateral sorting information exists in the fourth transmembrane domain of the α-subunit of Na,K-ATPase. Studies by Nelson and coworkers highlighted an alternative role of the α-subunit in the basolateral localization of the pump: anchoring via the ankyrin–spectrin (fodrin) cytoskeleton. Early work by this group showed that in some MDCK cell lines, newly synthesized Na,K-ATPase is delivered to the PM without polarity, but nonetheless acquires a mostly lateral distribution through specific interactions of the α-subunit with ankyrin (Hammerton et al. 1991). The lateral localization of the ankyrin–spectrin cytoskeleton is dependent on the expression of E-cadherin (see Chapter 18 for further discussion of the role of E-cadherin in epithelial polarization). Ectopic expression of E-cadherin in nonpolarized fibroblasts induced the assembly of a membrane skeleton at cell–cell contact sites and leads to a restricted localization of Na,K-ATPase to these sites; however, when a truncated E-cadherin form lacking the catenin-binding domain was expressed, neither fodrin nor Na,K-ATPase was localized to cell–cell contacts (McNeill et al. 1990). Moreover, overexpression of ankyrin-binding and actin-binding domains of β-spectrin results in highly abnormal cells lacking polarized distribution of the Na,K-ATPase. A role of the ankyrin–spectrin cytoskeleton in the polarized distribution of Na,K-ATPase is also supported by observations that show that in CP and RPE, the ankyrin/fodrin cytoskeleton is

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Figure 21.4. Mechanisms involved in the polarization of Na,K-ATPase. (A) Three main mechanisms have been postulated to be involved in the polarized distribution of Na,K-ATPase in epithelia. (1) Intracellular sorting at the level of the Golgi apparatus. The interaction of α- and β-subunits of Na,K-ATPase takes place in the endoplasmic reticulum and is required for exit from this organelle. Biochemical assays indicate that in some strains of MDCK cells (derived from the kidney) and Fisher rat thyroid (FRT) cells, Na,K-ATPase is sorted into a vectorial route to the basolateral surface that does not involve passage through recycling endosomes, observed for other basolateral proteins such as VSVG protein and transferrin receptor. (2) Trapping by the spectrin–fodrin cytoskeleton. In some MDCK strains, Na,K-ATPase is delivered without polarity to both apical and basolateral membranes; nonetheless, Na,K-ATPase is still basolaterally localized in these cells, which was attributed to interaction of the α-subunit with the ankyrin–spectrin cytoskeleton, assembled at the lateral membrane under the control of the epithelial cell adhesion molecule E-cadherin (Hammerton et al. 1991). (3) Homotypic adhesion between β-subunits. More recently, evidence has accumulated supporting a key role of the β-subunit in the basolateral localization of Na+,K+-ATPase. The β-subunit has homotypic adhesion properties. Shoshani et al. (2005) showed that overexpression of the β1-subunit cause concentration of Na,K-ATPase at the lateral membrane of MDCK cells (Shoshani et al. 2005). This model depends strongly on the affinity of a β1-subunit for homotypic β1-subunit located in another epithelial cell placed on the opposite side of the intercelullar space. Thus, Padilla-Benavides et al. attached β1-subunit to beads and demonstrated that it is able to selectively bind to and retain the soluble extracellular segment of β1 (Padilla-Benavides et al. 2010). (B) Molecular representation of Na+,K+-ATPase of two neighboring epithelial cells interacting via their β1-subunits (yellow) based on the recently reported crystal structure (Shinoda et al. 2009) and on recent FRET studies by Shoshani, Cereijido, and coworkers (Padilla-Benavides et al. 2010).

predominantly apical, like Na,K-ATPase (Philp and Nachmias 1985; Gundersen et al. 1991). Furthermore, overexpression of E-cadherin in an RPE cell line that lacks E-cadherin promote redistribution of Na,K-ATPase to the lateral domain (Marrs et al. 1995). A recent study focused on identifying the trafficking itinerary of the newly synthesized Na+ pump, using the SNAP tag system (New England Biolabs, Inc.) (Farr et al. 2009). This study showed that newly synthesized Na,K-ATPase does not transit through recycling endosomes. This observation is consistent with the lack of requirement of the Na,K-ATPase’s basal distribution for the clathrin adaptor AP1B, which localizes to recycling endosomes (Cancino et al. 2007; Gravotta et al. 2007). Basolateral localization of Na,K-ATPase is also independent of clathrin, suggesting that other clathrin adaptors do not play a role in this sorting process (Deborde et al. 2008).

Role of the β-Subunit Three isoforms of the Na,K-ATPase β-subunit have been identified: β1, β2, and β3. Analysis of the 3D structure of the pump, which was recently crystallized (Morth et al. 2007; Shinoda et al. 2009), reveals that part of the extracellular domain of the β-subunit interacts with the α-subunit, while the C-terminal lobe has an Ig-like structure that resembles cell–cell adhesion molecules (Bab-Dinitz et al. 2009). Indeed, the β2-subunit was originally identified as an adhesion molecule, AMOG, in glial cells (Gloor et al. 1990). The β2-subunit has about twice the number of N-glycans as the β1-subunit, and studies by Vagin and coworkers demonstrated a role of these glycans in determining apical versus basolateral distribution of Na,K-ATPase (Vagin et al. 2005, 2006, 2007, 2008; Tokhtaeva et al. 2010). Which subunit dominates the sorting of the heterodimer is not well understood and may be different depending on the cell types. Interestingly, in RPE, where the Na+,K+-ATPase is polarized to the apical membrane, the β1 isoform is expressed (Wetzel et al. 1999). Recent work by two of us indicates that the adhesive properties of the β1-subunit play a paramount role in basolateral localization of the pump (Shoshani et al. 2005). In

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(A)

(B)

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1 α

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MDCK cells, the β1-subunit localizes at the lateral but not the basal or apical membrane domains. Transfection of the dog β1-subunit into CHO cells results in the production of Ca-independent cell aggregates, consistent with an adhesive role of this protein and with a previously described intrinsic glycan-binding capacity that could be involved in neural cell interactions (Kitamura et al. 2005). Furthermore, our most recent work using pulldown, co-immunoprecipitation, and fluorescence resonance energy transfer (FRET) assays has shown that (1) β-subunits of neighboring cells are sufficiently close to allow direct β–β interaction across the intercellular space in vitro and in vivo; (2) that β–β interaction is species specific, as it is observed between rat/rat and dog/dog β1-subunits, but not between rat/dog ones; and (3) that in cultured monolayers of MDCK cells, the extracellular domains of β1-subunits on neighboring cells are sufficiently close as to allow energy transference in FRET assays (Padilla-Benavides et al. 2010). On these bases, we have proposed the model illustrated in Figure 21.4B where Na+,K+-ATPase is anchored to the cell borders facing the intercellular space, by virtue of the β–β interaction between two neighboring cells.

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PROTON-COUPLED MONOCARBOXYLATE TRANSPORTERS Lactate is both an end product of glycolysis and a substrate for oxidative phosphorylation (see Chapter 6 on mitochondria). At physiological pH (7.4), lactic acid exists predominantly in its ionic form and is largely membrane impermeable. Lactate is transported into and out of cells by H+-coupled lactate transporters, which are members of the SLC16 gene family (Halestrap and Meredith 2004). Fourteen members of this family have been identified to date, but only MCT1, MCT3, and MCT4 have been shown to be proton-coupled lactate transporters. MCTs share structural similarities with other solute transporters: They have 12 membrane-spanning domains, and both the amino and carboxy termini are cytoplasmic. MCT1–MCT4 are heteromeric transporters like Na,K-ATPase and are comprised of a nonglycosylated catalytic α-subunit (MCT) and a highly glycosylated β-subunit (Philp et al. 2003a; Poole et al. 1996). CD147 is the accessory subunit for MCT1, MCT3, and MCT4, while embigin is the accessory subunit of MCT2 (Halestrap and Meredith 2004; Wilson et al. 2005). Both CD147 and embigin are members of the Ig superfamily and are highly glycosylated single-pass membrane proteins. The extracellular domain of CD147 is the longest region and has two Ig-like domains of the C2 type. The transmembrane domain is about 24 amino acids in length, contains a leucine zipper and a charged glutamic acid residue, and is highly conserved in all species examined including Drosophila. In vivo and in vitro studies have shown that maturation and cell-surface expression of MCT1, MCT3, MCT4, and CD147 require the expression of both subunits, and, in the absence of one subunit, the other one is targeted for degradation (Philp et al. 2003a; Deora et al. 2005; Gallagher et al. 2007). MCT1 is the most ubiquitous isoform and is found in most cells including erythrocytes and muscle. Under normoxic conditions, the MCT1 transporter supplies cells with lactate, pyruvate, and ketone bodies to use as an energy source. Under hypoxic conditions, MCT1 is upregulated in cells to release lactate produced through glycolysis. For l-lactate, MCT1 has a Km of 500 μM (Dimmer et al. 2000). MCT3 was cloned from an embryonic chicken library screened with an RPE-specific monoclonal antibody (Philp et al. 1995). It is expressed preferentially in RPE cells, where it is restricted to the basolateral membrane. MCT3 has a lower affinity for lactate than MCT1; at physiological pH, we determined the Km to be 8.3 mM (Grollman et al. 2000). MCT4 shares the greatest sequence homology with MCT3. It has low substrate affinity; the Km for lactate has been determined to be 30 mM (Dimmer et al. 2000). MCT4 is expressed in cells that have few mitochondria and are dependent on glycolysis for the production of ATP. MCT4 is expressed in embryonic tissues, but as development progresses and the embryo becomes more reliant on oxidative metabolism, there is a downregulation of MCT4 and an upregulation of MCT1.

Polarized Distribution of MCTs in Different Epithelia MCTs display variable polarity in different epithelia. MCT1 (SLC16A1) is polarized to the basolateral membrane of intestinal epithelium (Garcia et al. 1995; Koho et al. 2005) but is apical in RPE (Philp et al. 1998; Daniele et al. 2008). In contrast, MCT3 (SLC16A8) and MCT4 (SLC16A3) are localized basolaterally in all epithelia, including RPE (MCT3), thyrocytes (MCT4) (Fanelli et al. 2003), cultured RPE (MCT4) (Philp et al. 2003b), and small intestine (MCT4) (Gill et al. 2005). The retina is one of the most metabolically active tissues in the body producing large quantities of lactate and CO2, which are released into the SRS. Lactate levels in the SRS are

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between 4 and 13 mM, while lactate in the blood is 1 mM. The RPE tightly regulates the pH and osmolarity of the SRS by transporting excess lactate out of the retina to the choroidal vessels. Early physiological studies showed that RPE expresses a proton-coupled lactate transporter (1:1, H+ : Lac−) at the apical membrane in various native and cultured RPE preparations (bovine, porcine, frog, and human) (Kenyon et al. 1994; la Cour et al. 1994; Lin et al. 1994). This apical membrane H/Lac cotransporter in RPE was identified as MCT1, the first member in the family of monocarboxylate transporters (Philp et al. 1998). At the RPE basolateral membrane, lactate is transported out of the cell via MCTs (MCT3 and MCT4) (Philp et al. 2001, 2003b) and a Cl−/Lac anion exchanger (AE2) (Kenyon et al. 1994). In kidney PT and salivary gland, the reuptake of lactate from glomerular filtrate and saliva is facilitated by Na+-coupled lactate transporters in the apical membrane since the lactate concentrations in both cases is about 1 mM. Na+-dependent lactate transporters are utilized by tissues that absorb lactate against a concentration gradient into the blood, for example, kidney, small intestine, and salivary glands. In these tissues, two members of the SLC5 family, SLC5A8 (SMCT1) and SLC5A12 (SMCT2), are expressed at the apical surface, and their activity is complemented by proton-coupled monocarboxylate transporters localized at the basolateral membrane, MCT1 and/or MCT2 (Halestrap and Meredith 2004; Frank et al. 2008).

Variable Polarity of MCTs in Different Epithelia Is Determined by Tissue-Specific Variations in the Trafficking Machinery The distinct polarity of MCT1 in different tissues raised a crucial question: Is the tissuespecific polarized localization of MCT/CD147 heterocomplexes determined by the MCTs or by CD147? Initial insight into the variable sorting of MCTs in different epithelia was provided by the identification in our laboratory of a basolateral sorting signal in CD147 consisting of a single leucine (L252) near an acidic patch in its cytoplasmic tail (Deora et al. 2004). Expression of mutant rat CD147-L252A, but not wild-type CD147, in MDCK cells led to redirection of endogenous MCT1 to the apical membrane, while endogenous MCT4 or transfected MCT3 remained basolateral (Fig. 21.5). The sorting of MCT1/ CD147-L252A to the apical membrane in MDCK cells mimicked the distribution of this transporter in RPE cells. These findings led us to propose a model whereby MCT1 does not harbor any endogenous basolateral signal; hence, its trafficking to the PM is regulated by the basolateral signal of CD147. More recent work (Castorino et al. 2011) has identified strong basolateral signals in MCT3 and MCT4 that control their basolateral distribution in epithelia (Fig. 21.5). Our findings suggest a novel paradigm for the polarized sorting of heterodimeric transporters whereby the dominant sorting information can be found in the nonglycosylated transporting subunit or the glycosylated accessory subunit, according to the MCT (Fig. 21.5). Our results show that the flexible phenotype of MCT1 is due to the fact that it does not have a basolateral sorting sequence and its trafficking is determined by the basolateral signal of CD147. In PT, intestine, salivary gland, and MDCK cells, where the basolateral sorting signal of CD147 is recognized by the trafficking machinery of the cells, the transporter is targeted to the basolateral membrane. However, in RPE cells, the sorting machinery does not recognize the basolateral signal of CD147 and MCT1/CD147 are polarized apically, driven by a cryptic apical sorting signal. Our findings suggest that it is not only the array of transporters but also the array of sorting machinery expressed in a tissue-specific manner that facilitates the polarized expression of proteins to fulfill the physiological needs of the host tissue or organ.

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MDCK

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MCT3 Recessive apical signal

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L252A

Figure 21.5. Polarity of lactate transporters is determined by sorting signals in either the α- or β-subunits. Like many ion transporters, including Na,K-ATPase, monocarboxylate transporters are composed of a single-span highly glycosylated β-subunit (also called CD-147, basigin, emmprin, etc.) and a nonglycosylated, multispan α-subunit (MCT) that is responsible for the transporting properties of the heterodimer. CD147 has a strong basolateral signal in its cytoplasmic domain, based on a single leucine and a nearby acidic patch, which is recognized by MDCK cells but not by retinal pigment epithelium (RPE). The α-subunit of MCT1 has no sorting information; hence, the heterodimer is targeted basolaterally in MDCK cells and apically in RPE cells. On the other hand, the α-subunit of MCT3 (expressed constitutively in RPE cells) and MCT4 (expressed in rat thyroid cells and enterocytes in small intestine) have strong basolateral signals that target the heterodimer basolaterally in all epithelial cells, even if the basolateral signal of CD147 is inactivated by mutation to alanine.

VARIATIONS IN THE SORTING MACHINERY AMONG EPITHELIA The Na,K-ATPase expressed basolaterally by most epithelia and apically by RPE and CP appears to be composed of the same α- and β1-subunits. Similarly, MCT1 and CD147, expressed basolaterally in MDCK cells and PT and apically in RPE, are structurally identical. Hence, the variable polarity of these transporters cannot be attributed to the display of different sorting signals in different epithelia. Rather, their variable polarity is likely a result of variations in components of the sorting machinery in different epithelia. Ironically, RPE cells differ from most other epithelial cells in that they do not express the basolateral sorting adaptor AP1B; yet neither Na,K-ATPase nor MCT1/CD147 are AP1B cargos. Furthermore, Na,K-ATPase is not dependent on clathrin for its basolateral localization, indicating that other clathrin adaptors (see Chapter 2) are not likely to be involved in its basolateral targeting. Other emerging basolateral sorting adaptors, for example, Naked and ankyrin G, need to be tested for their participation in the sorting of the pump. On the other

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hand, our previously our published work (Deborde et al. 2008; Castorino et al. 2011) indicate that the basolateral polarity of MCT1/CD147 in MDCK cells is dependent on clathrin; hence, we can expect that clathrin adaptors other than AP1B control their basolateral localization. The identity of these clathrin adaptors is still unknown. Not surprisingly, since basolateral sorting signals are usually dominant over apical sorting signals and many basolateral proteins contain cryptical apical signals, variable expression of a component of the basolateral sorting machinery contributes to the most striking variations in epithelial polarity observed to date. RPE and PT do not express the clathrin adaptor AP1B, leading to apical or depolarized localization of a plethora of cognate basolateral proteins in these epithelia (Diaz et al. 2009; Schreiner et al. 2010). These proteins include adhesive PM proteins such as Coxsackie adenovirus receptor (CAR), junctional adhesion molecule C (JAMC), and neural cell adhesion molecule (NCAM); hormone receptors such as parathyroid hormone receptor (PTHR); and nutrient receptors such as LDLR and TfR. Clearly, lack of expression of AP1B must contribute in a significant way to the normal physiology of these epithelia. However, it is not yet known whether the absence of AP1B in PT and RPE results from suppression of μ1B expression as they differentiate or, alternatively, reflects the absence of AP1B in the parental epithelia from which they derive (respectively, neural ectoderm or early nephric blastema). Our observations predict that abnormal expression of AP1B in RPE and PT, perhaps as a consequence of certain diseases, may lead to pathological dysfunction.

FUTURE PERSPECTIVES While notable progress has been made in the characterization of apical and basolateral sorting signals in single-span monomeric PM proteins, the characterization of the sorting signals involved in polarized distribution of membrane transporters, which are usually oligomeric, lags considerably behind. In a few cases (e.g., AQPs and NKCC1), expression of different splicing isoforms may lead to their variable apical/basolateral localization. In other cases, for example, Na,K-ATPase and MCT1/CD147, their variable polarity may be attributed to variations in the expression of sorting machinery. Our current knowledge of the sorting machinery is very basic. For most basolateral and apical sorting signals, the corresponding “sorters” have not been identified. These areas of polarity research can expect considerable progress in the near future.

ABBREVIATIONS AE AP AQP ARE ASE BSE CAR CFTR CLC CP CRE

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anion exchanger adaptor protein aquaporin apical recycling endosomes apical sorting endosomes basolateral sorting endosomes Coxsackie adenovirus receptor cystic fibrosis transmembrane conductance regulator chloride channel choroid plexus common recycling endosome

CSF ENaC JAMC LDLR MCT NBC NBCE NBE NCAM NCC NDC NHE

cerebrospinal fluid epithelial sodium transporter junctional adhesion molecule C low-density lipoprotein receptor H+/Lac− cotransporter sodium bicarbonate cotransporter sodium chloride bicarbonate exchanger sodium bicarbonate exchanger neural cell adhesion molecule Na+/Cl− exchanger Na-dependent cotransporters of ... Na/proton exchanger

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Na+, K+, 2Cl− cotransporter plasma membrane proximal tubule parathyroid hormone receptor retinal pigment epithelium

SRS TER TfR TGN TJ

subretinal space transepithelial electrical resistance transferrin receptor trans-Golgi network tight junction

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22

NEURONAL DOMAINS Jennifer S. Goldman Timothy E. Kennedy

DEFINITION Neurons are heterogeneous and complex cells, specialized for intercellular communication at synapses. Mature neurons are functionally polarized into somatic, dendritic, and axonal compartments. While dendrites and soma receive and integrate synaptic input, axons generate and transmit neuronal output as action potentials. Within each compartment, unique subcellular domains support neuronal function. Specialized domains include the axon initial segment, nodes of Ranvier, presynaptic terminals, and postsynaptic specializations such as the postsynaptic density (PSD) and dendritic spines. The action potential is generated at the axon initial segment and is propagated over long distances, repeatedly regenerated at nodes of Ranvier that are regularly spaced along myelinated axons. The electrical depolarization of the action potential triggers the release of a chemical neurotransmitter from presynaptic terminals. The chemical transmitter diffuses a short distance across the synaptic cleft and is received by postsynaptic specializations. Whether or not the postsynaptic neuron will reach the threshold to fire an action potential is determined by the summation of excitatory and inhibitory postsynaptic events by the dendrite. To form functional neural networks during development, neurons elaborate dynamic structures that are important for appropriate neuronal migration, initial polarization, axon and dendrite guidance, process branching, and synapse formation. These include the leading and trailing edges of migratory neurons, and the growth cones of elongating axons and dendrites. In this chapter we summarize key discoveries in the history of neuronal cell biology, provide an overview of our current understanding of the structure and significance of specialized neuronal domains, and discuss examples of neurological disorders that result from malformation or degeneration of these domains in neurons.

HISTORICAL PERSPECTIVE Our current understanding of the basic biology and physiology of neurons is largely derived from an explosion in theory and experimentation that has taken place between the late eighteenth century and today (Bennett 1999; Cowan et al. 2003). In 1791, Luigi Galvani serendipitously produced twitching in isolated frog’s legs through contact of an attached nerve with mixed metals generating current (Galvani 1791), providing the first clue that the nervous system was responsive to electricity. In the 1840s, Emile Du Bois-Reymond

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discovered that nerves themselves conduct electricity (Du Bois-Reymond 1849), Hermann von Helmholz measured the velocity of conduction (Helmholtz 1850), and Julius Bernstein theorized that the origin of conductance lay in the distribution of charged ions (Bernstein 1868). Subsequently, experimental results began to hint at a chemical component of neuronal activity (Bernard 1878; Langley 1905; Loewi 1921; Brown et al. 1936; Dale et al. 1936). An intense debate began as to whether the nature of the neuronal signal was chemical or electrical, “soup or spark” (Eccles 1945; Cook 1986). In the late 1930s, methods were devised to record the activity of resting and active neurons (Cole and Curtis 1939; Hodgkin, 1939; Hodgkin and Huxley 1945). Hodgkin and Huxley went on to prove that the movement of sodium and potassium ions across the neuronal plasma membrane resulted in voltage changes across the membrane and that this is essential for neuronal signaling (Hodgkin and Huxley 1952). In 1954, with the advent of appropriate technology to record intracellularly (Graham and Gerard 1946), the chemical theory was proven by Eccles and colleagues, who were ironically some of the strongest proponents of the spark hypothesis (Brock et al. 1952). We now know that neurons use both chemical and electrical means to communicate and that specialized domains within neurons are required for this function. The low intrinsic contrast and high density of neural tissue had long impeded scientists from visualizing individual neurons. In the late nineteenth century, Camillo Golgi (see also Chapter 8 on the Golgi apparatus) developed a method to selectively impregnate a small proportion of neurons with potassium chromate and silver salts (Golgi 1873). Despite the prevalent theory that all tissues were composed of cells (Schwann 1839), Golgi believed that axons and dendrites formed a kind of reticulum, whereby neurons were physically interconnected (Mazzarello 1999). Using Golgi’s method, Santiago Ramón y Cajal developed the beginnings of what is today called the neuron doctrine, the theory that neurons are discrete cellular units, channeling a unidirectional flow of information. Cajal suggested that between neurons was a “protoplasmic kiss,” where neurons could transfer information. He was also the first to describe the presence of specialized structural domains in dendrites and axons, and to observe their development (Cajal 1909, 1933). It would be another several decades before the “neuron versus reticulum” debate would be indisputably resolved with the advent of electron microscopy and the direct visualization of the synaptic cleft between neurons (Palay 1956). In the 1960s, V. Whittaker and colleagues developed biochemical methods to isolate synapses and their component parts (Gray and Whittaker 1962; Whittaker et al. 1963). The 1970s and 1980s saw the identification of various classes of neurotransmitter, including amino acids, biogenic amines, peptides, and gases, and the identification and cloning of genes encoding ion channels and neurotransmitter receptors (Cowan et al. 2003).

NEURONAL DOMAINS The polarization of cellular compartments is the basis for the unidirectional flow of information through neurons, from dendrites, to soma, to axons, and transferred between neurons at synapses. Dendritic postsynaptic domains are closely juxtaposed with the presynaptic specializations of other neurons, to respond to the release of neurotransmitter in the synaptic cleft. The binding of chemical neurotransmitters to receptors at postsynaptic specializations increases (depolarizes) or decreases (hyperpolarizes) the membrane voltage of the local area of dendrite. The dendrites integrate depolarizing and hyperpolarizing synaptic inputs, and propagate these changes over the area of the dendritic tree, toward the soma and the axon. Due to the heterogeneity of dendrites (Fig. 22.1), different neurons perform very different computations.

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If the membrane voltage at the axon initial segment increases over the threshold for firing an action potential, the initial segment generates a spike in the membrane voltage. The action potential propagates down the axon and is regenerated along myelinated axons at nodes of Ranvier. When the action potential reaches a presynaptic specialization, neurotransmitter is released into the synaptic cleft and diffuses across the extracellular space to postsynaptic specializations. While each neuron usually produces only one type of neurotransmitter, most neurons have a multiplicity of specialized postsynaptic sites that are receptive to different types of neurotransmitters from other neurons. Among the great diversity of neurons found in nervous systems, primary sensory neurons are unique in that they may not contain dendrites. Rather than receiving input from other neurons, sensory neurons are responsive to environmental stimulation through domains specialized to detect chemical (e.g., odor and taste molecules), mechanical (e.g., depression of the skin or membrane movement in the ear), electromagnetic (e.g., visible light), electrical (e.g., navigation by electric fish), or thermal (e.g., heat) stimuli. Like other neurons, primary sensory neurons convey information to other neurons via an axon. Due to the diversity of specialized domains present in primary sensory neurons, they will not be described in further detail here; however the following citations are provided for review (Bullock 1982; Roper 1989; Buck 1996; Garcia-Anoveros and Corey 1997; Dhaka et al. 2006). All neurons contain subdomains defined by interactions between membrane, extracellular matrix, and cytoskeleton that are specialized to subserve unique functions (Fig. 22.2). A unifying principle of most neuronal domains is the concentration of specific types of transmembrane ion channels for rapid changes in electrical potential. In a resting state, neurons maintain low intracellular concentrations of Na+, Cl−, and Ca2+, and high intracellular K+ (Table 22.1). Whereas some ion channels are open in a resting state, others open in response to discrete events, including sensory stimulation, neurotransmitter signaling, intracellular signaling, or changes in membrane voltage. Ion channel opening allows diffusion of particular ions through the transient, ion-selective pore, producing changes in membrane voltage (Kandel et al. 2000; Purves et al. 2008).

Soma The neuronal cell body is much like that of other cells, containing the nucleus and organelles (see Chapters 6–10, 23) for synthesizing and sorting a great diversity of proteins and lipids, but in some cases accounts for as little as 10% of the volume of the entire neuron. Some mRNA made in the soma is transported to dendrites or to the growth cone for local, activity-induced synthesis (Wang et al. 2010), but the great majority of protein is made in the cell body and sorted into pathways bound for the axon or the dendrites, contributing to the uniqueness of the domains present in each compartment. The cell body also receives some of the most persuasive synaptic inputs since they occur so close to the axon initial segment (Kandel et al. 2000; Purves et al. 2008).

Axonal Domains The axon is a thin extension from the cell soma, which may be relatively short if the neuron communicates only with its neighbors, or more than a meter long to communicate with neurons relatively far away. The axon has a unique molecular composition specialized to generate action potentials and to propagate them to the distal presynaptic terminals, where they signal the release of neurotransmitter molecules.

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Cortical Pyramidal Neuron

LGN Relay Neuron

Cerebellar Purkinje Neuron

Cortical Basket Neuron

Spinal Motor Neuron

Retinal Bipolar Neuron

Figure 22.1. Drawings of vertebrate neurons, adapted from Cajal (1909). Pictured are a pyramidal neuron from the cerebral cortex, a relay neuron from the lateral geniculate nucleus (LGN), a Purkinje neuron from the cerebellum, a basket neuron from the cerebral cortex, a motor neuron from the ventral horn of the spinal cord, and a bipolar neuron from the retina.

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Figure 22.2. Schematic of a neuron, showing an incoming axon forming en passant and terminal synapses with the dendrites of the cell pictured. Note dendritic branches emerging from the cell soma, and dendritic spines. Note also axonal specializations, the axon hillock and myelin-associated domains, the node of Ranvier, and the internode. A specialized extracellular matrix surrounds the cell.

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TABLE 22.1. Ion Distribution between the Extracellular and Intracellular Compartments Generates an Electrical Potential Difference across the Plasma Membrane

Ionic species Potassium (K+) Sodium (Na+) Calcium (Ca2+) Chloride (Cl−)

Extracellular

Intracellular

Direction of ion flux at rest

20 440 10 560

400 50 0.0001 40–150

Out In In In

Concentration gradients of potassium (K+), sodium (Na+), calcium (Ca2+), and chloride (Cl−) ions are generated and actively maintained by transmembrane ion transporters during activity and at rest. Ion channels open in response to cellular signaling, revealing a selectivity pore, which discriminates between ion species. Opening of an ion channel results in diffusion of specific ions through the channel’s selectivity pore, to areas of lower concentration. Values shown are those reported for the membrane of the squid giant axon (Hodgkin and Huxley 1945; Purves et al. 2008).

Axon Initial Segment (AIS) The AIS is often the section of axon closest to the cell body. It maintains the unique environment of the axon by forming a physical barrier to diffusion, and functions to generate the neuronal output, the action potential. The membrane of the AIS contains a high concentration of voltage-activated Na+ channels, which sense changes in plasma membrane voltage. The resting membrane potential of the neuron is typically about −65 mV with respect to the extracellular space, but when increased over a threshold value, Na+ channels in the AIS undergo a conformational change, resulting in the inward flux of Na+, a dramatic rise in the membrane voltage, and the generation of an action potential. The plasma membrane of the AIS also contains voltage-activated K+ channels, which contribute to the falling phase of the action potential (Kandel et al. 2000; Purves et al. 2008). The AIS is endowed with a rich repertoire of intracellular proteins that regulate and stabilize ion channels in the membrane (Fig. 22.3A). Na+ and K+ channels are anchored

Figure 22.3. Schematics of the molecular composition of specialized membrane domains in neurons. The membrane at the axon hillock (A) is densely packed with voltage-gated Na+ and K+ channels, adhesion molecules (Caspr2, Tag1, NrCAM, and NF-186), scaffolding molecules (PSD93 and ankyrinG), and actin cytoskeleton. Myelin associated domains (B), the node of Ranvier, paranode, juxtaparanode, and internode each contain a functional complement of specialized proteins. At the node of Ranvier, known proteins are similar to those comprising the axon hillock. Adjacent to the node of Ranvier, the paranode marks the beginning of the myelinated segment of an axon. The paranode domain is enriched with adhesion molecules, which tether the axon to the myelinating glial cell (MGC), and prevent lateral diffusion between the node of Ranvier and the juxtaparanode. The juxtaparanode contains K+ channels, whereas the axonal membrane along the internode is not known to display a high degree of specialization. A glutamate synapse (C) is composed of the juxtaposed membrane specializations of two cells, representing presynaptic and postsynaptic neurons, separated by the synaptic cleft. The pre-synaptic domain is pictured on the upper portion, and contains neurofilament cytoskeleton (actin), scaffolding and catalytic proteins (CASK, Liprin-a, ELKS, RIM, Piccolo and Bassoon, SNAP25, Syntaxin, and Synapsin), synaptic vesicles and associated proteins (VAMP and synaptotagmin), calcium channels, transmembrane adhesion molecules, and organelles (mitochondria and endosomes). The postsynaptic domain is pictured on the lower portion, and contains glutamate receptors and associated subunits (AMPA and TARP, NMDA, and mGluR), scaffolding molecules (PSD95, Homer, Shank, and GKAP), catalytic proteins (CAMKII and actin regulatory proteins), adhesion molecules, and organelles (spine apparatus and ribosome).

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through interactions with ankyrin-G, an intracellular scaffolding molecule that plays a key role in organizing the AIS. Ankyrin-G also associates with βIV spectrin, which interacts with actin filaments, linking ion channels to the underlying cytoskeleton (see Chapters 1 and 12). The transmembrane adhesion molecules, NrCAM and Neurofascin-186, bind the extracellular matrix of the AIS and intracellular ankyrin-G, tying down the AIS on both sides of the membrane. Other scaffolding, signaling, and adhesion molecules that include PSD-93, iFGFs, Tag1, and Caspr2 are also clustered in the AIS (Ogawa and Rasband 2008; Grubb and Burrone 2010). Nodes of Ranvier, Paranodes, Juxtaparanodes, and Internodes Nodes of Ranvier are regularly spaced interruptions in myelin along the axon. Myelin is a dense, lipid insulation formed by myelinating glial cells (MGCs). MGCs encapsulate segments of the axon by wrapping their membranes around many times, substantially reducing dissipation of the action potential. Nodes of Ranvier are functionally and molecularly similar to the AIS, containing high densities of Na+ channels important for regenerating the action potential as it travels down the axon. Nodes of Ranvier are also enriched with cell adhesion molecules such as NrCAM and Neurofascin-186, the cytoskeletal adaptor ankyrin-G, and the actin-binding protein βIV spectrin. As at the axon hillock, these molecules serve to regulate the membrane localization, stability, and gating properties of Na+ channels, and the domain architecture of the node (Poliak and Peles 2003; Susuki and Rasband 2008; Thaxton and Bhat, 2009). Adjacent to the node of Ranvier is a domain called the paranode, a region of noncompact myelin, where the MGC forms a specialized adhesive junction with the axonal membrane that keeps the MGC closely associated with the axon and prevents diffusion of proteins between the membranes of the node of Ranvier and the juxtaparanode. The axonal membrane at the paranode does not contain ion channels, but is enriched in adhesion molecules that are required for the tight association between the MGC and the axon. Adjacent to the paranode is the juxtaparanode, which contains K+ channels of unknown function, adhesion molecules, including Caspr2 and Tag1, and the intracellular scaffolding molecule PSD93. Finally, between juxtaparanodes are domains of compact myelin, called internodes. The axonal domain associated with the internode is not known to be particularly specialized (Poliak and Peles, 2003). Presynapse The presynaptic specialization contains the molecular machinery to convert the electrical action potential into the release of chemical neurotransmitter (Fig. 22.3C). Conventionally, the presynaptic specialization is established by the axon, although dendritic and somatic presynaptic release sites can occur. Presynapses may be terminal (occurring at the end of a branch) or en passant (occurring along the length of the axon) (Fig. 22.2). The membrane specialization of the presynapse, organized to release neurotransmitter, is called the active zone (Kandel et al. 2000; Purves et al. 2008). In electron microscope reconstructions, the active zone appears highly structured, characterized by clusters of neurotransmittercontaining synaptic vesicles aligned along the presynaptic membrane, tethered to a dense intracellular protein cytomatrix (presynaptic grid) rich in filamentous actin, myosin, spectrin, and β-catenin. This cytomatrix is important for anchoring the active zone, and displays varying morphologies in different synapses and different species. Some examples include concave, but simple, morphology at the Caenorhabditis elegans neuromuscular synapse (NMJ), ribs and beams at the frog NMJ, T-shaped at the NMJ of Drosophila melanogaster, perpendicular to the presynaptic membrane in electroreceptor synapses of the skate, and rows of pyramids in the human hippocampus (Zhai and Bellen 2004). The presynaptic

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active zone is precisely aligned with postsynaptic neurotransmitter receptor clusters, enabling fast and reliable synaptic transmission (Cowan et al. 2003). The active zone membrane contains target-SNARE (t-SNARE; syntaxin and SNAP25) proteins, which interact with vesicular SNARE (v-SNAREs; synaptobrevin/VAMP) on synaptic vesicles. Coiled-coil protein domain interactions between t- and v- snares drive synaptic vesicle fusion with the active zone membrane. To achieve regulated fusion of synaptic vesicles during an action potential, the presynapse is enriched with voltageactivated N, P/Q, and R-type Ca2+ channels, all requiring strong depolarization to open. When an action potential invades the presynaptic terminal, synaptic vesicle associated, calcium-sensitive synaptotagmin proteins plunge into the active zone membrane in response to the brief rise in intracellular Ca2+, causing the regulated fusion of synaptic vesicle and active zone membrane, thereby releasing neurotransmitter into the synaptic cleft (Schoch and Gundelfinger, 2006). In addition to the readily releasable pool of synaptic vesicles at the active zone, a reserve pool of synaptic vesicles is tethered to the actin cytoskeleton, away from the active zone membrane. Synapsin proteins constitutively interact with actin and with synaptic vesicles in a phosphorylation-state dependent manner. Influx of Ca2+ during periods of high frequency action potential firing leads to the activation of kinases, which phosphorylate synapsins allowing reserve synaptic vesicles to join the readily releasable pool at the active zone (Cowan et al., 2003; Schoch and Gundelfinger, 2006).

Synaptic Cleft The synaptic cleft is an approximately 20-nm extracellular space between the pre- and postsynaptic membranes. Depending on the type of synapse, specific enzymes important for the degradation and reuptake of neurotransmitters are present in the cleft. The synaptic cleft is bridged by a complement of synapse-specific adhesion molecules present in the pre- and postsynaptic membranes, often tethered intracellularly to the actin cytoskeleton. Many members of the cadherin and Ig family of adhesion molecules, including sidekicks, nectins, SynCAMs, integrins, neurexins and neuroligins, eph and ephrinB, and netrin-Gs and netrin-G ligands bind homo- or heterophilically to bridge the gap between pre- and postsynaptic membranes. Synaptic adhesion molecules regulate the position, molecular composition, and functionality of both domains (Kim and Ko 2006; Dalva et al. 2007; Han and Kim 2008; Chua et al. 2010).

Dendritic Domains The word dendrite is derived from the Greek word dendron, meaning tree. The dendrites are often highly branched, underlying their ability to integrate the input of often thousands of excitatory, inhibitory, and modulatory synapses. Dendrites are the locus of most synaptic input to the neuron, displaying exquisite domain specialization at each postsynaptic site. Current generated at active synapses is integrated along the length of the dendrite and propagates toward the AIS using both passive and active dendritic properties. Postsynapse The postsynapse is the neuronal domain specialized to sense the release of chemical neurotransmitter from the presynaptic neuron. Postsynaptic specializations are quite diverse, depending on the type neurotransmitter used to signal at that synapse; however, all mature postsynapses are closely aligned with the presynaptic active zone across the synaptic cleft. Postsynapses contain concentrated clusters of neurotransmitter receptors, ion channels, and signaling molecules tethered intracellularly to scaffolding molecules and the actin cytoskeleton (Cowan et al. 2003).

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Among the various types of synapses, most is known about synapses using the neurotransmitter glutamate due to the facility with which they can be biochemically isolated and their abundance in the brain. Also referred to as an asymmetric synapse or Gray’s type 1 synapse (Gray 1969), the hallmark of an excitatory glutamatergic synapse is the PSD. The PSD is an electron-dense structure, situated just beneath the postsynaptic membrane, with an average diameter of 300–400 nm. The PSD is densely packed with layers of protein, anchored on one side to the intracellular face of the plasma membrane and on the other to the actin cytoskeleton. The first layer of the PSD is enriched with glutamate neurotransmitter receptors. Alpha-amino-3-hydroxy-5-methylisoxazole-4proprionic acid (AMPA) and N-methyl-D-aspartate (NMDA) ionotropic glutamate receptors are glutamate-activated ion channels responsible for the fast increase in membrane voltage that results from glutamate neurotransmission. Metabotropic glutamate receptors, mGluRs, are G-protein-coupled glutamate receptors, which activate intracellular signaling cascades in response to glutamate binding. G-protein signaling impinges on ion channels, opening or closing them, resulting in a slow change in the membrane voltage. Both AMPA and NMDA receptors are anchored to the PSD by a very abundant scaffolding molecule called PSD-95. While NMDA receptors bind directly to PSD-95, AMPA receptors are tethered through auxiliary subunits called transmembrane AMPAr regulatory proteins (TARPs). At the PSD, PSD-95 proteins are situated an average of 12 nm from the cytoplasmic face of the postsynaptic membrane (Kim and Ko 2006; Okabe 2007). The next layer of the PSD contains a dense network of multidomain scaffolding proteins, including Homer, Shank, and GKAP, actin and actin-regulatory molecules, mRNA trafficking and protein synthesis machinery, and a great variety of synaptic signaling molecules (Kim and Ko 2006; Okabe 2007). Calcium/calmodulin protein kinase II (CAMKII) is one of the most abundant proteins in the PSD, and exists as holoenzymes of 12–14 subunits. CAMKII holoenzymes translocate into the PSD in response to Ca2+ influx through NMDA receptors. Activated CAMKII will phosphorylate AMPA receptors, increasing the amount of current they transmit on glutamate binding. In addition, active CAMKII enhances the number and stability of AMPA receptors in the PSD, thereby enhancing the effectiveness of the synapse (Kim and Ko 2006; Okabe 2007). In addition to glutamatergic excitatory postsynapses, specialized postsynaptic domains exist for inhibitory and modulatory neurotransmitter signaling. Significantly less is known about the molecular composition of central nervous system synapses specialized for GABA, glycine, acetylcholine, and biogenic amines (serotonin, dopamine, adrenaline, and noradrenaline). Like glutamatergic synapses, other types of postsynapses contain dense clusters of both ionotropic and metabotropic receptors specifically activated by their respective neurotransmitter ligands (Maley et al. 1990; Arancibia-Carcamo and Moss 2006; Bockaert et al. 2006). Unlike glutamatergic synapses, other types of postsynaptic specializations do not contain a electron-opaque postsynaptic density, and are therefore classified as symmetrical synapses (Gray 1969; Cowan et al. 2003). Whereas glutamatergic postsynapses usually occur on dendrites or dendritic spines, it is quite common to find inhibitory postsynapses on the cell body, axon initial segment, and the necks of dendritic spines. Modulatory biogenic amine postsynapses often occur on dendrites and dendritic spine necks and are commonly also associated with the axon, where they efficiently regulate neuronal output (Cowan et al. 2003). Dendritic Spine More than 90% of glutamatergic synapses in the vertebrate nervous system are made onto dendritic spines, fine protrusions from the parent dendrite, that are thought to isolate and

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concentrate signaling molecules at the postsynapse. The classical dendritic spine consists of a constricted neck, and a broad head, containing the PSD, actin, polyribosomes, and smooth endoplasmic reticulum (spine apparatus). The volume of a dendritic spine often is very well correlated with the size of the PSD, AMPA, and NMDA receptor number, and synaptic strength (Nimchinsky et al. 2002; Tashiro and Yuste 2003). Mature dendritic spines display diverse morphologies but can be categorized into mushroom, thin, branched, or stubby subtypes based on their morphology. Mushroom spines are the most stable, most enriched in F-actin, most likely to contain polyribosomes and spine apparatus (Nimchinsky et al. 2002; Tashiro et al. 2003). Animals reared in enriched environments display increased dendritic spine density (Schrott 1997), whereas sensory-deprived animals have a significantly reduced spine density (Bryan and Riesen 1989). Two-photon time-lapse imaging studies carried out in vivo indicate that while most dendritic spines are quite stable, spines may appear, disappear, or change shape over the course of minutes, hours, or days (Grutzendler et al. 2002; Trachtenberg et al. 2002; Holtmaat et al. 2005). New spines are more likely to persist in the presence of learning, whereas old spines may be lost to preserve the overall density of dendritic spines in adult animals (Xu et al. 2009; Yang et al. 2009). Dendritic Geometry and Synaptic Integration Dendritic geometry has important implications for neuronal computation. Synapses occurring on smaller dendrites produce larger amplitude synaptic events with faster time courses, whereas synapses occurring on larger dendrites tend to produce smaller synaptic events with longer time courses. Dendritic branches furthest from the cell body tend to be smallest in diameter, whereas primary order dendrites, closest to the cell body, tend to be larger. Balancing these differences, spatial and temporal decay of the amplitude of synaptic current occurs as it propagates toward the cell body. In addition, the effect of a synaptic potential depends on the pattern of activation and proximity of other synergistic or antagonistic synapses (Gulledge et al. 2005; London and Hausser 2005; Wen and Chklovskii 2008). Thus, geometric properties of dendrites fundamentally contribute to the integration of synaptic information and different geometries perform very different computations. Although it was initially believed that dendrites were mere passive “cable-like” conductors for synaptic activity, it is now appreciated that dendrites contain a heterogeneous collection of voltage-activated Na+, Ca2+, and K+ ion channels capable of actively amplifying membrane voltage changes, and even generating action potentials when sufficiently depolarized (Gulledge et al. 2005; London and Hausser 2005). Active dendritic conductances also allow for back-propagation of action potentials generated at the axon hillock into the dendrites, a phenomenon thought to contribute to spike-timing-dependent plasticity, whereby coincident activation of a postsynapse with a back-propagating action potential increases the strength of that synapse, whereas uncorrelated activation of a synapse with a back-propagating action potential decreases the strength of that synapse (Caporale and Dan 2008).

Cytoskeleton and Neuronal Domains The neuronal cytoskeleton makes a profound contribution to the architecture of neuronal domains. It consists of microtubules (a polymer of tubulin), microfilaments (a polymer of actin), and neurofilaments (polymers of neuronal intermediate filament proteins). Whereas neurofilaments are relatively stable cytoskeletal elements, influencing the radial growth of neuronal processes during development, microtubules and microfilaments may be quite dynamic. The motility of cells and growth cones, the development of polarized

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compartments within the neuron, and the branching of axons and dendrites all fundamentally depend on microtubule and microfilament remodeling (Lambert de Rouvroit and Goffinet 2001; Pollard and Borisy 2003; Geraldo and Gordon-Weeks 2009; Valiente and Marin 2010). Cytoskeletal remodeling is regulated by a complex interplay of microfilament and microtubule regulatory proteins (Georges et al. 2008; see also Chapters 12 and 14 on the actin and microtubule cytoskeletons). In mature neurons, microtubules are polarized within compartments to form scaffolds for long-distance, vesicular transport of domain-specific proteins. All axonal microtubules are plus-end directed, meaning that the growing end of the microtubule is pointed toward the distal end of the axon, whereas dendritic microtubules have mixed polarity. Dendritic and axonal microtubules also differ in the composition of microtubule-associated proteins (MAPs). Whereas Map2 associates with dendritic microtubules, Tau specifically associates with axonal microtubules. The motor proteins kinesin and dynein carry proteincontaining cargo vesicles from the nucleus specifically toward the plus and minus ends of microtubules, respectively (Fig. 22.4A; Conde and Caceres 2009). In contrast to microtubules, which form bundles along the longitudinal axis of axons and dendrites, cortical filamentous actin (microfilament cytoskeleton) is enriched just under the membrane in a dense mesh, and is involved in local trafficking of domain-specific proteins and their stabilization (see Chapter 1). This submembranous actin serves to dock and restrain protein and lipid constituents found in all neuronal domains, including pre- and postsynaptic specializations, the axon hillock, and the adhesive contacts that generate the force underlying cell and process motility (Kusumi and Sako 1996; Letourneau 2009). Additionally, submembrane cortical actin plays a key role at the end of the secretory pathway in neurons; myosin motors walk along actin polymers to the periphery, where cargo is unloaded and secretory vesicles fuse with the membrane (DePina and Langford 1999).

Extracellular Matrix; Perineuronal Nets The extracellular matrix, or perineuronal net, is a neuron- and domain-specific collection of heparin, keratin, and chondroitin sulfate proteoglycans, tenascin, fibronectin, and hyaluronan organized into lattice-like structures, secreted by neurons and glial cells (Karetko and Skangiel-Kramska 2009). In the developing nervous system, the extracellular matrix is linked through adhesion molecules in the plasma membrane to the intracellular cytoskeleton, generating traction for motility. Many guidance cues and growth factors are linked to the extracellular matrix, providing directional and positional information for motile cells (Rhiner and Hengartner 2006). In the adult central nervous system (brain and spinal cord), perineuronal nets tightly envelop cell bodies, dendrites, axons, and synapses to stabilize the extracellular milieu, potentially limiting neuronal plasticity, synapse formation, and axonal regeneration. In the adult brain, dissolution of perineuronal nets restores heightened plasticity associated with early development (Karetko and Skangiel-Kramska 2009).

NEURONAL DOMAINS DURING DEVELOPMENT Neurons are generated from undifferentiated precursor cells in proliferative pro-neural epithelia during embryonic development and migrate to their appropriate positions in the brain, spinal cord, and ganglia. Axons and dendrites then grow into target regions, where they branch and form synapses. During each of these developmental steps, transient domains exist, appropriate to each developmental stage.

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dendrite

+

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+

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+

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axon

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(B) Filopodium

Lamellipodium

microtubules cortical actin actin bundle actin arc

transport vesicle, kinesin motor transport vesicle, dynein motor transport vesicle, myosin motor map2

tau

Figure 22.4. Cytoskeletal specializations contribute to axon/dendrite polarity and the morphology and motility of the growth cone. (A) Both axons and dendrites contain oriented microtubules along which kinesin motors transport cargo toward the plus end and dynein motors transport cargo toward the minus end. Both axons and dendrites display sub-plasmalemmal networks of cortical actin along which myosin motors transport cargo to the periphery. In axons, all microtubules are oriented with the plus end facing away from the soma, whereas microtubules in dendrites have mixed polarity. Also, axons contain the distinct microtubule-associated protein Tau, whereas dendrites contain the microtubule-associated protein Map2. (B) The growth cone is a transient axonal domain that integrates extracellularly distributed guidance cues to lead the axon to its postsynaptic target during development. Whereas the central domain of the growth cone is highly enriched with microtubules, the peripheral domain contains filamentous actin organized into networks (cortical actin), bundles, and arcs. Cortical actin supports membrane ruffling underlying the formation and motility of lamellipodia, and bundles of actin filaments support the protrusion of filopodia.

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Specialized Domains during Migration and the Establishment of Polarity Some neurons migrate through the extracellular matrix relatively unaccompanied, other neurons migrate in chains, while still others migrate along the fibers of radial glia. In all cases, specialized domains organize the adhesion, intracellular signaling, cytoskeletal reorganization, and the direction of neuronal migration (Lambert de Rouvroit and Goffinet 2001; Pollard and Borisy 2003; Valiente and Marin 2010). The leading edge is a dynamic structure oriented in the direction of migration, which may be relatively simple or branched in different cell types and in distinct microenvironments. The force to extend the membrane of the leading edge is generated by a dynamic, growing network of branched submembranous filamentous actin, and is regulated by actin regulatory proteins that include Arp2/3, profilin, cofilin, WASp, and the Rho GTPases, Cdc42, Rac, and RhoA (Jaffe and Hall 2005; see also Chapter 12). A longitudinal system of microtubules links the leading edge to the centrosome, stabilizing the leading edge and directing the trafficking of lipid and protein components. The leading edge adheres to the substrate through interactions with integrins and other adhesion molecules (see also Chapter 17), and is polarized with regard to extracellularly distributed guidance cues that include netrins, slits, semaphorins, wnts, and ephrins, giving topographical information to the motile cell (Bloch-Gallego et al. 2005; Maness and Schachner 2007; Schmid and Maness 2008; Chedotal 2010). In migrating neuronal precursors, the soma translocates behind the leading edge by a process called nucleokinesis, dependent on contractile actin–myosin forces (Lambert de Rouvroit and Goffinet 2001; Pollard and Borisy 2003; Valiente and Marin 2010). While the leading edge is specialized to adhere to and follow extracellular cues toward the neuron’s ultimate destination, the trailing edge is specialized to de-adhere, and follow the leading edge and the soma. In contrast to the net growth of actin filaments in the leading edge, the trailing edge displays high levels of RhoA activity and filamentous actin disassembly (Lambert de Rouvroit and Goffinet 2001; Kurokawa et al. 2005). For many neuron types, polarity initially arises while neurons are migrating into target tissues, and a distinct relationship exists between the leading/trailing edge and the future axons and dendrites; however, the mechanisms determining neuronal polarity can be quite complicated and cell-type specific. Because polarization is difficult to visualize in vivo, most studies have been carried out in cell culture. In some cases, the plane of axo-dendritic polarization can be predicted by apico-basal polarity of progenitor cells and the plane of cell division. During mitosis, the centrosome and Golgi apparatus are situated near the membrane opposite of the cleavage furrows between dividing cells. The centrosome organizes the polarity of microtubules and of the Golgi apparatus (see also Chapters 8, 14, and 15 for further discussion of the centrosome, Golgi, and cell polarization). During initial neurite extension, the first protrusion forms adjacent to the centrosome and Golgi and grows to become the longest process and future axon. The subsequent protrusions to emerge will become dendrites (de Anda et al. 2005). Within the nascent axon, microtubules begin to be stabilized while actin is destabilized (Witte and Bradke 2008). The axon begins to concentrate the partitioning-defective protein (PAR) complex, consisting of Par3, Par6, and atypical protein kinase C (PKC). Plus-end directed, axon-specific microtubule-based kinesin motors, initially present in all nascent processes, become excluded from all but the future axon. Eventually, the axon begins to concentrate proteins enriched in the mature axon, including PI3 kinase, CRMP-2, and GAP43, and exclude polyribosomes (Horton and Ehlers 2003; Barnes and Polleux 2009). The AIS plays an important role in the generation and maintenance of axonal polarity, demarcating the boundary between the axon and soma, and preventing the diffusion of membrane and cytosolic proteins between the two compartments (Grubb and Burrone 2010).

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Cell extrinsic cues also contribute to the generation of neuronal polarity. Secreted morphogens and neurotrophins, bone morphogenic proteins (BMPs), BDNF, NT-3, and semaphorins play complex roles, initiating signaling events that lead to directed outgrowth and the initial specification of the axon (Horton and Ehlers 2003).

Axon and Dendrite Guidance, Branching, and Synapse Formation One of the most daunting challenges of development requires axons to grow over long distances to innervate target structures. During axon guidance, the axon is endowed with a motile sensory structure at its leading edge, called the growth cone. Intracellularly, the growth cone is enriched with cytoskeletal and signaling molecules (Moore and Kennedy 2006). The central portion of the growth cone is enriched with microtubules and microtubule-binding proteins, serving to stabilize the growing axon and providing a substrate for protein trafficking (Fig. 22.4B). In contrast, the peripheral portion of a growth cone is enriched with actin, which dynamically cycles between monomeric and polymeric forms to generate motile filopodia (finger-like structures) and lamellipodia (flat-ruffling structures) (Geraldo and Gordon-Weeks 2009).The growth cone membrane is enriched with receptors for guidance cues, secreted and cell-surface ligands that provide spatial signals capable of attracting, repelling, fasciculating, or collapsing growth cones. Canonical guidance cues include the ephrins, semaphorins, slits, and netrins. Downstream signaling in response to these cues impinges on the regulation of Rho GTPases and reorganizes the actin cytoskeleton (O’Donnell et al. 2009; Bashaw and Klein 2010). Lipid raft-like membrane specializations have been implicated in the function of several guidance receptors and adhesion molecules (Kamiguchi 2006). The response of axon growth cones to guidance cues is complex and may be altered as an axon progresses along its trajectory. Such response-switching allows a growth cone to be initially attracted to a cue present at an intermediate target, but then ignore or be repelled by the same cue in order to continue toward a final target (Kamiguchi 2006; Moore and Kennedy 2006; Geraldo and Gordon-Weeks 2009; Lowery and Van Vactor, 2009; Hong and Nishiyama 2010). Dendrites also extend and are endowed with growth cones that may be directed to grow into target structures by guidance cues. Within target regions, dendrites sprout filopodia, which either disappear quickly or elongate into new dendritic branches, often sprouting filopodia of their own. Additionally, dendritic spines are thought to initially appear as thin, motile dendritic filopodia, which actively search for compatible presynaptic partners. If successful, dendritic filopodia may mature into rounder, more stable protrusions (Nimchinsky et al. 2002). Key mechanisms thought to decide the fate of a new dendritic filopodium include Ca2+ and neurotransmitter signaling, cell adhesion molecules, and membrane tension (Heiman and Shaham 2010; Jan and Jan 2010). Within targets, axons may also branch through protrusion and elongation of filopodia (interstitial branching), or by bifurcation of the migrating axonal growth cone (Kollins and Davenport 2006). A growing axon releases neurotransmitters even before reaching its target. As well, before being innervated by an axon, dendrites contain “hot spots” or clusters of neurotransmitter receptors. Both axonal and dendritic filopodia play active roles in initiating axodendritic contact, probing the environment for suitable synaptic candidates. At the site of contact, signaling through guidance, adhesion, and neurotransmitter molecules locally reorganizes the architecture of the membrane and cytoskeleton, leading to the recruitment and organization of pre- and postsynaptic proteins (Zhen and Jin 2004; Craig et al. 2006; Cai and Sheng 2009; Owald and Sigrist 2009; Shen and Cowan 2010).

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NEURONAL DOMAINS AND MENTAL DISORDERS As neuronal domains are specialized for the integration and processing of information, developmental and degenerative disorders of neuronal domains interrupt cognitive ability and impair normal behavior.

Dendritic and Synaptic Disorders Many forms of mental retardation (MR) are associated with anomalous development of dendrites, dendritic spines, and synapses (Kaufmann and Moser 2000; Newey et al. 2005). Fragile X results from the expansion of a trinucleotide CGG repeat, and hypermethylation of the FMR1 gene in the 5’ untranslated region, resulting in the underexpression of fragile X MR protein (FMRP). Normally, FMRP binds mRNA and inhibits local translation of dendritic mRNAs in response to metabotropic glutamate receptor signaling (see also Chapter 25 on cytoplasmic RNA domains). Absence of FMRP in fragile X patients results in dysregulation of dendritic mRNAs important for development, function, and plasticity of postsynaptic specializations, and the maturation of spine morphology (Irwin et al. 2001; Newey et al. 2005; Bassell and Warren 2008). Down’s syndrome results from triplication of genetic material present on the 21st chromosome, and overexpression of genes present at this locus. In cortical neurons of newborn children with Down’s syndrome, the dendrites display an overelaboration, and the density of dendritic spines is increased with respect to normal children. In normal development, dendrites become more elaborate and the density of dendritic spines increases, tending to display mushroom, thin, short, or stubby geometries. In contrast, cortical neurons from patients with Down’s syndrome display dendritic atrophy, and a great reduction in the density of mature dendritic spines (Marin-Padilla 1972; MarinPadilla 1976; Takashima et al. 1981; Becker et al. 1986). Although the etiology of autism is less well understood, polymorphisms in genes encoding proteins regulating mTor-dependent mRNA translation (Chapter 25), as well as genes encoding synaptic adhesion molecules such as Neuroligins, Neurexins, and the scaffolding molecule Shank, have all been associated with the development of autism spectrum disorders. Phenotypically, cortical areas involved in the production of language and social behaviors show an increase in cell and synaptic growth, and a shift in the balance of excitatory to inhibitory synapses (Bourgeron 2009). Alzheimer ’s disease (AD) is a degenerative disorder characterized at late stages by cortical atrophy, dysfunction, and loss of cortical synapses, senile plaques, neurofibrillary tangles, loss of basal forebrain cholinergic neurons and their innervation of the cortex, as well as severe cognitive decline. Senile plaques are abnormal aggregates of amyloid β peptide (Aβ), which is known to potently inhibit synaptic transmission and synaptic plasticity, and is toxic to cells at high concentrations. Neurofibrillary tangles are intracellular occlusions, consisting mainly of the microtubule associated protein, Tau, in an abnormally hyperphosphorylated form, aggregating into straight or paired filaments (Morfini et al. 2009; Jurgensen and Ferreira 2010; Perl 2010).

Axonal Trafficking Disorders AD also presents pathology in axonal trafficking along microtubules; pathological Tau filaments actively inhibit kinesin-mediated, microtubule-based axonal transport of protein, resulting in dystrophic neurites. Dysfunctional axonal transport is also associated with other degenerative diseases: amyotrophic lateral sclerosis (ALS), hereditary spastic paraplegias (HSPs), spinal muscular atrophies, Huntington’s disease, and Parkinson’s disease,

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all of which result in progressive decline in neuronal function, neuronal degeneration, and neurogenic deficits specific to the population of neurons lost (Morfini et al. 2009).

FUTURE PERSPECTIVES Throughout the history of neuroscience research, theory has become proof as quickly as novel methods can be devised. Today, technical innovations are taking place more rapidly than ever, providing unique insight into long-standing questions regarding neuronal structures and the functions they subserve. Today, we are witnessing especially rapid advancements in genetics and optics, with utility for making specific inquiries at the submicron scale. Until recently, it was only possible to assess neuronal activity with electrophysiology, ultimately recording the activity of a neuron for at most a few hours. The identification of genetically encoded fluorescent proteins, fluorescent calcium sensors, and voltage sensors, advances in confocal and two-photon fluorescence microscopy, and the advent of methods to visualize neurons in situ have enabled the study of live cells in living organisms doing the things they actually do in life. Additionally, the discovery and utilization of light-activated ion channels to artificially depolarize or hyperpolarize the neuronal membrane, selectively activating or inactivating specific neurons—in intact and behaving animals—is contributing greatly to our understanding of the neuronal basis of behavior. The identification of cell-type-specific promoter sequences has led to the ability to drive the expression of any of these genetically encoded tools in specific types of neurons. Contemporary insights derived from the development of such techniques indicate that specialized subcellular domains are essential elements of the neuronal computational networks that underlie nervous system function.

ACKNOWLEDGMENTS We thank Wayne Sossin, Ed Ruthazer, and Ajit Singh Dhaunchak for comments on the text. This work was supported by the Canadian Institutes of Health Research (CIHR). JSG was supported by a Jeanne-TimminsCostello Fellowship. TEK holds an FRSQ Chercheur Nationaux Award and is a Killam Foundation Scholar.

ABBREVIATIONS AD AIS ALS AMPA

Alzheimer ’s disease axon initial segment amyotrophic lateral sclerosis alpha-amino-3-hydroxy-5-methylisoxazole-4proprionic acid Arp2/3 actin-related complex (containing actin related proteins ARP2 and ARP3) Aβ amyloid β peptide BDNF brain derived neurotrophic factor BMPs bone morphogenic proteins CAMKII calcium/calmodulin protein kinase II Caspr contactin associated protein Cdc42 cell division control protein 42 CRMP collapsing response mediator protein FMRP fragile X mental retardation protein GAP-43 growth associated protein 43

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GKAP HSPs iFGF MGC MR NMDA NMJ NrCAM NT-3 PAR PI3K PKC PSD PSD-93 PSD-95 RhoA

guanylate kinase-associated protein hereditary spastic paraplegias intracellular fibroblast growth factor myelinating glial cells mental retardation N-methyl-D-aspartate neuromuscular junction Ng-CAM (cell adhesion molecule) related neurotrophin 3 partitioning-defective protein complex phosphatidylinositol 3-kinases protein kinase C postsynaptic density postsynaptic density protein, 93 kilodalton postsynaptic density protein, 95 kilodalton Ras homolog gene family, member A

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SHANK

SH3 and multiple ankyrin repeat domains protein SNAP-25 soluble NSF attachment protein SNARE SNAP receptor Tag1 transiently expressed axonal surface glycoprotein 1

TARPs VAMP WASp Wnts

transmembrane AMPAr regulatory proteins vesicle associated membrane protein Wiskott-Aldrich syndrome protein Wingless and Int proteins

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VI

DOMAINS REGULATING GENE EXPRESSION G E NE T I C RE GUL AT I O N defines the molecular composition of the cell and determines the expression of the proteins and lipids that generate the various cellular domains described previously. Domains that regulate DNA and RNA organization are not membrane bound and function in the nucleus to regulate DNA transcription and RNA production (Chapter 23) and in the cytoplasm to regulate RNA translation, trafficking, and stability (Chapter 25) in various cell types including neurons (Chapter 22). Exchange of material, including messenger RNA, between the nucleus and cytoplasm is regulated by the nuclear pore, a highly complex structure that represents a true example of a molecular machine (Chapter 24). Domain organization of genetic material therefore regulates cell differentiation and cell fate by regulating nuclear DNA expression and RNA trafficking and translation in the cytoplasm.

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NUCLEAR DOMAINS Dale Corkery Kendra L. Cann Graham Dellaire

DEFINITION The nucleus is the warehouse for the vast majority of cellular DNA in eukaryotes. As such, its responsibilities include DNA replication and cell cycle regulation; DNA repair; RNA transcription from RNA polymerases (pols) I, II, and III; and RNA splicing. The relationship between structure and function pervades biology, and the nucleus is no exception, as it is organized around the regulation and the biochemistry of these processes. At the most fundamental level, the nucleus is arranged around the DNA, which is compacted and distributed as chromatin in chromosomes (Dundr and Misteli 2001). Interacting with the chromatin are numerous proteinaceous nuclear substructures/domains (Fig. 23.1), the largest of which are the nucleoli. The other subnuclear organelles include the promyelocytic leukemia nuclear bodies (PML NBs); nuclear speckles (splicing factor compartments, interchromatin granule clusters [IGCs]); Cajal bodies (coiled bodies); Oct1, PTF (proximal sequence element-binding transcription factor) transcription (OPT) domains; gems; Polycomb group (Pc-G) bodies; Src associated in mitosis 68 kDa protein (Sam68) nuclear bodies; and the perinucleolar compartment (PNC) (Dundr and Misteli 2001; Spector 2001; Dellaire et al. 2003). All of these subnuclear structures lack membranes. However, they each contain a defining set of marker proteins (Fig 23.1 and Table 23.1) (Dellaire and Bazett-Jones 2007), they can be morphologically identified by light and/or electron microscopy, and some of them can be biochemically purified (Dundr and Misteli 2001). The nuclear domains can be further subdivided into facultative domains (Sam68 and Polycomb group bodies) that are cell type and activity related, and constitutive domains (nucleolus, PNC, Cajal bodies, PML NBs), which are present in all cell types. This chapter will focus on the physical and functional characteristics of these subnuclear compartments.

CHROMATIN AND THE INTERCHROMATIN DOMAIN SPACE Eukaryotic chromatin is organized within the nucleus as arrays of nucleosomes; a hypothesis first proposed by Roger Kornberg (Kornberg 1974). Each nucleosome is composed of two copies of the histone proteins H2A, H2B, H3, and H4 assembled into an octamer. Each octamer has 145–147 base pairs of DNA wrapped around it to form the nucleosome core. Repeating nucleosome cores are further assembled into higher-order structures that

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Figure 23.1. Overview of nuclear domains. Illustration of a nucleus and the various constitutive and facultative (*) nuclear domains. TABLE 23.1. Summary of Commonly Used Molecular Markers for the Identification of Nuclear Domains by Light Microscopy

Nuclear Domain Heterochromatin Promyelocytic leukemia (PML) body Polycomb (Pc-G) body Nucleolus Chromosome territory Nuclear splicing speckle Sam68/SLM nuclear body Perinucleolar compartment (PNC) Cajal body Gem

Molecular Markers Heterochromatin protein 1 (HP1) alpha, trimethylated H3K9 and H4K20 PML protein, Daxx, Sp100 Bmi1 Nucleolin, fibrillarin Fluorescent in situ hybridization (FISH) of individual chromosomes SC35, also known as splicing factor, arginine/serine-rich 2 (SRFS2) Src substrate associated during mitosis 68 (Sam68), Sam68-like mammalian proteins 1 and 2 (SLM-1, SLM-2) Polypyrimidine tract-binding protein 1 (PTBP1), also known as PTB p80 coilin Survival of motor neurons (SMNs), gemin2

are stabilized by linker histone H1. Packaging DNA around the nucleosome allows for a 30- to 40-fold compaction of linear DNA (Luger et al. 1997). The most obvious form of structural genome organization in the nuclei of mammals and other eukaryotes is the compartmentalization into heterochromatin and euchromatin. The distinction between these two forms of chromatin was first made by the German botanist Emil Heitz while visualizing nuclear chromosomal regions in moss in 1928 (reviewed in Fedorova and Zink 2008). He observed regions that did not undergo postmitotic decon-

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densation and termed these parts of the chromosomes heterochromatin. In contrast, fractions of the chromosome that decondense and spread out diffusely in the interphase nucleus are referred to as euchromatin. Heitz proposed that heterochromatin reflected a functionally inactive state of the genome. Heterochromatin is composed mainly of repetitive DNA sequences and is classified into two groups: constitutive heterochromatin and facultative heterochromatin. Constitutive heterochromatin contains hypoacetylated nucleosomes enriched for histone H3 methylated at lysine 9 (meH3K9) and remains condensed throughout an organism’s lifespan. An example of constitutive heterochromatin would be the large bands of pericentric satellites present next to the centromeres of human chromosomes. In contrast, facultative heterochromatin is enriched for histone H3 methylated at lysine 27 (meH3K27) and is assembled when needed to permanently silence genes (Grewal and Jia 2007). An example of facultative heterochromatin would be the mammalian inactive X, in which one female X chromosome is silenced early in development (Brockdorff 2002). The organization of chromosomes within the nucleus was realized in the 1980s using in situ hybridization. Using this technique, it was demonstrated that the DNA of individual chromosomes was not distributed over the whole nucleus during interphase, but remained confined to a smaller subvolume, or the so-called chromosome territory (CT) (Schardin et al. 1985; Lichter et al. 1988). It has been demonstrated that CTs display a radial organization that correlates with gene density (Croft et al. 1999). Gene-poor chromosomes are located more peripherally, while gene-rich chromosomes tend to occupy more interior positions (Habermann et al. 2001). The observation that chromosomes were confined to territories led to the interchromosome domain compartment (ICD) model (Cremer et al. 1993, 1995). This model speculated that a three-dimensional channel (or so-called ICD space) would surround and separate individual CTs, thereby confining regulatory factors and transcription factors to a single ICD space (Cremer et al. 1993, 1995). Since this discovery, several different versions of the ICD model have been proposed, but the idea that the territorial organization of a chromosome plays an important role in gene regulation is central to all models (Fedorova and Zink 2008). While clear-cut evidence supporting any version of the ICD model is currently lacking, there are several studies in support of gene regulation through territorial organization of chromosomes. For example, it has been shown that some loci loop out of their respective CT, and this change in localization correlates with transcriptional activity (Mahy et al. 2002a, b; Williams et al. 2002; Chambeyron and Bickmore 2004; Chambeyron et al. 2005). However, it should be noted that localization within a territory does not preclude gene transcription, nor is localization at the periphery of a CT sufficient to increase gene transcription (Morey et al. 2007, 2009; Noordermeer et al. 2008). As we will see in the following sections, many of the “constitutive” nuclear compartments found in all mammalian nuclei are found in the ICD and make contacts with chromatin. Furthermore, the functional state of chromatin with respect to transcription and replication also play an important role in the structure of several nuclear compartments.

NUCLEOLUS The nucleolus is the most prominent subcompartment of the eukaryotic nucleus (see Fig. 23.2A) and is the site for ribosomal RNA (rRNA) transcription and ribosome biogenesis (reviewed in Hernandez-Verdun 2006a; Raska et al. 2006; Sirri et al. 2008). Mammalian nuclei usually contain between one and four nucleoli, which together can take up as much

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(A)

DAPI

Nucleolin

Overlay

Nucleoli

(B)

PML

PML NBs

(C)

Coilin

Cajal Bodies

(D)

SC35

Nuclear Speckles

Figure 23.2. Protein components and in situ structures of subnuclear bodies. Normal human diploid fibroblasts (NHDFs) were stained with an antibody to nucleolin (red) to mark the nucleoli (A), an antibody to PML (green) to mark the PML NBs (B), an antibody to coilin (red) to visualize to the Cajal bodies (arrowheads) (C), or an antibody to SC35 (red) to mark the nuclear speckles (D). In all images, the cells have been counterstained with DAPI (4′,6-diamidino-2phenylindole) (blue) to visualize the DNA.

as one-third of the nuclear volume (Carmo-Fonseca et al. 2000). The nucleoli form around the ribosomal DNA (rDNA) gene repeats, which are found in tandem arrays at several chromosomal loci termed the nucleolar organizing regions (NORs). Furthermore, by electron microscopy, the nucleolus consists of three morphologically distinct components: the fibrillar centers (FCs), the dense fibrillar component (DFC), and the granular component

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(GC). The RNA pol I-mediated rRNA transcription is carried out at the interface between the FC and the DFC, and the newly made transcripts radiate out into the DFC and eventually progress into the GC. Along the way, the precursor rRNA is complexed with ribosomal and nonribosomal proteins, cleaved, processed, and matured into pre-60S and pre-40S subunits that are eventually exported to the cytoplasm (reviewed in Raska et al. 2006). The very existence of the nucleoli depends on ongoing rRNA transcription and the secondary rRNA processing events, and because of this, they represent the canonical example of the relationship between structure and function within the nucleus (Dundr and Misteli 2001). The first observations of the nucleolus came from Fontana when he noticed the presence of an ovoid body visible in the nucleus more than two centuries ago (reviewed in Sirri et al. 2008). Since then, our understanding of the nucleolus has evolved in parallel to advancements in imaging technology. Through the use of light microscopy in the nineteenth century, numerous cytologists described the variability in nucleolar morphology with great precision (reviewed in Sirri et al. 2008). The presence of RNAs in the nucleolus was first described in the 1950s, followed by identification of ribosomal genes (rDNAs) in the NOR in the 1960s through the use of in situ hybridization techniques (Caspersson 1950; Perry 1962; Ritossa and Spiegelman 1965). Around this same period, techniques were developed to allow for the mass isolation of nucleoli. This led to the biochemical characterization of nucleolar components and the suggestion that the nucleolus was the site of ribosome biogenesis. Next, as described above, the functional organization of the nucleolus was deciphered between 1980 and 2000 with the invention of electron microscopy (reviewed in Sirri et al. 2008). Finally, the development of advanced microscopy techniques, including fluorescence microscopy and live-cell imaging (Handwerger and Gall 2006; Hernandez-Verdun 2006a, b; Trinkle-Mulcahy and Lamond 2007), and largescale proteomic studies using mass spectrometry (Andersen et al. 2002; Scherl et al. 2002) have allowed much more extensive analyses of nucleolar dynamics, structure, and biochemical composition. This has helped usher in a new era of our understanding of the nucleolus (Raska et al. 2006; Boisvert et al. 2007; Sirri et al. 2008), as it is now clear that the function of the nucleolus extends far beyond ribosome biogenesis. In fact, only approximately 30% of the over 700 human proteins identified in the nucleolus have known functions in ribosome biogenesis (Boisvert et al. 2007). Other nucleolar proteins have functions that include messenger RNA export; the processing and maturation of nonnucleolar RNA and ribonucleoproteins; and the regulation of the cell cycle, DNA replication, DNA repair, cell senescence, tumor suppression, and the cell stress response (Maggi and Weber 2005; Raska et al. 2006; Boisvert et al. 2007; Dellaire and Bazett-Jones 2007; Montanaro et al. 2008; Sirri et al. 2008). Because ribosome biogenesis is required to support cell growth through the production of protein, it is perhaps not surprising that nucleolar function is tightly coregulated with cell cycle control. Growth factors and oncoproteins that promote proliferation, including c-Myc, epidermal growth factor, and insulin-like growth factor, also upregulate rRNA transcription, increasing the size and number of nucleoli (Raska et al. 2006; Derenzini et al. 2009). On the other hand, negative cell cycle regulators such as p53 and retinoblastoma protein (pRB) inhibit rRNA transcription (Voit et al. 1997; Zhai and Comai 2000). Finally, to ensure that cells do not enter the cell cycle without adequate protein production and growth, normal ribosome biogenesis is required for the expression of the critical cell cycle regulator cyclin E (Volarevic et al. 2000). These important links between nucleolar activity and cell proliferation form the biochemical basis for the use of nucleolar size as an important prognostic indicator in cancer pathology (reviewed in Pich et al. 2000; Derenzini et al. 2009).

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As mentioned above, the maintenance of nucleolar structure is intimately coupled to ongoing RNA pol I transcription, which when inhibited leads to a rapid and dramatic reorganization of this nuclear organelle (reviewed in Olson et al. 2000). Although yeast nucleoli remain intact during mitosis, the nucleoli of metazoans are highly dynamic and undergo rounds of disassembly and reassembly during the cell cycle (Carmo-Fonseca et al. 2000; Olson et al. 2000). Upon entry into mitosis, an ordered disassembly of first the GC and then the DFC of the nucleolus occurs. At this time, certain nucleolar components (e.g., upstream binding factor [UBF], a key regulator of RNA pol I transcription) remain attached to chromosomal NORs. A number of nucleolar proteins, including pre-rRNA processing components, associate with mitotic chromosomes during prophase but eventually localize to extrachromosomal nucleolus-derived foci (NDF) (e.g., nucleolin) during the progression from anaphase to telophase. During late telophase, NDF disappear and some processing components become associated with prenucleolar bodies (PNBs) (e.g., fibrillarin), which appear prior to the initiation of RNA pol I transcription. Once rRNA transcription begins, PNBs (or their components) are recruited to NORs, resulting in the formation of the nucleolus (reviewed in Carmo-Fonseca et al. 2000; Olson et al. 2000; Boisvert et al. 2007; Sirri et al. 2008). Beyond its role in rRNA biogenesis, the nucleolus appears to act as a storage site or reservoir for a number of proteins that do not have roles in rRNA metabolism (CarmoFonseca et al. 2000; Olson et al. 2000; Visintin and Amon 2000; Politz et al. 2002). In fact, the nucleolar sequestration of proteins appears to be a common means of cell cycle regulation among eukaryotes. For example, the tumor suppressor alternative reading frame product of the CDKN2A locus (ARF), whose expression is upregulated by oncogenic proteins such as Ras, Myc, and E1A, is a nucleolar protein (Tao and Levine 1999). ARF is an important activator of the p53 tumor suppressor, which regulates cell cycle checkpoints, cell senescence, and apoptosis. Normally, p53 is targeted for degradation by its negative regulator murine double minute 2 (MDM2), an E3 ubiquitin ligase (Iwakuma and Lozano 2003; Moll and Petrenko 2003). However, following DNA damage by ionizing radiation or mitomycin C, or when ARF is upregulated in response to oncogenic signals, ARF can sequester MDM2 in the nucleolus, preventing its interaction with p53 (Weber et al. 1999; Khan et al. 2004; Dias et al. 2006; Saporita et al. 2007). Therefore, nucleolar anchoring of MDM2 by ARF provides one means of regulating the cell cycle. Similarly, in Saccharomyces cerevisiae, the exit from mitosis is regulated by the nucleolar sequestration of the protein phosphatase Cdc14p, which is released from the nucleolus to promote both the degradation of the cyclin subunit B-type cyclin (Clb) and the accumulation of protein kinase inhibitor during late anaphase (Visintin et al. 1999). However, this mechanism for controlling mitotic exit is specific for lower eukaryotes like yeast, because as discussed above, nucleoli are dispersed during mitosis in metazoan cells. The nucleolar sequestration of MDM2 by ARF is only the beginning of the involvement of the nucleolus in p53 regulation. Cellular stresses that cause RNA pol I inhibition induce nucleolar disruption, initiating a release of proteins that can regulate the function of p53, which may ultimately lead to the activation of cell cycle checkpoint or apoptotic pathways (Rubbi and Milner 2003; Olson 2004; Mayer and Grummt 2005; Gjerset 2006). The list of stresses known to inhibit RNA pol I includes hypoxia, heat shock, transcriptional inhibitors, topoisomerase I inhibitors, and some DNA damaging agents (e.g., UV irradiation and cisplatin) (Rubbi and Milner 2003). Besides ARF, other nucleolar proteins that can regulate p53 include nucleostemin, nucleophosmin, nucleolin, and the ribosomal proteins (RPs) RPL5, RPL11 RPL23, and RPS7, through interactions that can involve p53, MDM2, ARF, and each other (Mayer and

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Grummt 2005; Takagi et al. 2005; Gjerset 2006; Saxena et al. 2006; Derenzini et al. 2009; Zhang and Lu 2009). Through its association with cell proliferation and tumor suppression, the nucleolus plays a critical role in cancer cell biology (reviewed in Maggi and Weber 2005; Raska et al. 2006; Boisvert et al. 2007; Montanaro et al. 2008; Sirri et al. 2008; Zhang and Lu 2009). However, the importance of this subnuclear structure in human health and disease is not limited to cancer. Nucleolar proteins whose genes are known to be mutated in human genetic diseases include TCOF in Treacher Collins (Marsh et al. 1998), ataxin7 in a form of spinocerebellar ataxia (Kaytor et al. 1999), and DKC1 in dyskeratosis congenita (Ruggero and Pandolfi 2003; Montanaro et al. 2008). The protein product of DKC1 is dyskerin, which functions in small nucleolar ribonucleoproteins (snoRNPs) and telomerase as a pseudouridine synthase (Ruggero and Pandolfi 2003; Montanaro et al. 2008). Finally, nucleolar proteins are common auto-antigens in patients with hepatocellular carcinoma (HCC), gastrointestinal, lung, and ovarian cancers (Imai et al. 1992).

PML NUCLEAR BODIES The PML protein (also called TRIM19, MYL, RNF71, or PP8675) is a tumor suppressor that localizes to punctate nuclear structures, or bodies (Dellaire and Bazett-Jones 2004; Bernardi and Pandolfi 2007). PML NBs, previously known as ND10, PML oncogenic domains (PODs), and Kr bodies, vary in size from 0.2 to 1.0 μm in diameter and can range in number from 1 to 30 bodies per nucleus (see Fig. 23.2B), depending on cell type, cell cycle phase, and differentiation stage (Dellaire and Bazett-Jones 2004). PML NBs have been found juxtaposed to other nuclear structures, such as nuclear gems and Cajal bodies, though the significance of this association is unknown (Dellaire and Bazett-Jones 2004). PML NBs are primarily protein based, with the main structural protein being the PML protein. However, they are dynamic structures, with over 75 different proteins as potential NB components (Dellaire et al. 2003). Other proteins that localize to these structures include Sp100, small ubiquitin-like modifier (SUMO), UBC9, HAUSP, Daxx, and cAMP responsive element binding protein (CREB) (CBP) (Maul et al. 2000; Borden 2002). Under normal conditions, neither chromatin nor RNA is found within the central core of these bodies, but newly synthesized RNA is associated with and extensive chromatin contacts are made at the periphery of these structures (Boisvert et al. 2000). However, in telomere negative cells that use the alternative lengthening of telomere pathway, during viral infection and following genotoxic stress, single-strand DNA may be found within PML NBs (Jul-Larsen et al. 2004; Boe et al. 2006; Grudic et al. 2007). The formation of the PML NBs is regulated by PML dimerization and SUMO-dependent multimerization (Shen et al. 2006). The dimerization is mediated by PML’s N-terminal RBCC motif, a tripartite motif that contains a RING zinc finger (R), two beta-box zinc fingers (B), and a coiled-coil domain (CC). Next, PML is sumoylated on three lysine residues by the SUMOconjugating enzyme UBC9 (Duprez et al. 1999). These sumoylated residues then interact with SUMO-interacting motif (SIM) domains on other PML proteins, seeding the multimerization (Shen et al. 2006). These sumoylation events are necessary for PML NB formation, as a PML mutant incapable of being sumoylated fails to recruit additional body components, such as Sp100 and Daxx (Ishov et al. 1999; Zhong et al. 2000a). The interaction between SIM domains and SUMO appears to be a common mechanism for the recruitment and retention of PML NB-associated proteins, including Daxx (Lin et al. 2006; Shen et al. 2006).

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PML NBs are also thought to function in gene transcription, proteasomal degradation, the antiviral response, apoptosis (programmed cell death), cell senescence, DNA repair, and tumor suppression (Dellaire and Bazett-Jones 2004; Bernardi and Pandolfi 2007). However, the biochemical functions of PML NBs in these pathways have not been completely elucidated. A few models have been proposed to explain how PML NBs exert their biological function (Zhong et al. 2000b; Negorev and Maul 2001; Borden 2002; Dellaire and Bazett-Jones 2004). The first model proposes that PML NBs operate as nuclear storage sites for the accumulation of proteins, both in normal conditions for sequestering proteins until they are required, as well as in pathological conditions where they isolate misfolded or foreign proteins. However, this view has expanded to include PML NBs being “catalytic surfaces” where proteins accumulate for posttranslational modification, assembly of protein complexes, and the regulation of protein turnover through modulation by ubiquitin-mediated proteasomal degradation (Dellaire and Bazett-Jones 2004; Takahashi et al. 2004; Bernardi and Pandolfi 2007). In fact, there is evidence that the PML protein itself can function as an E3 ligase for sumoylation (Quimby et al. 2006). Furthermore, because the PML NBs make extensive contacts with chromatin at their peripheries (Boisvert et al. 2000), they are very sensitive to changes in the cellular environment, especially changes that affect chromatin condensation (Eskiw et al. 2004; Dellaire et al. 2006b). PML NBs are also disrupted following various forms of DNA damage, including pyrimidine dimers induced by UV irradiation or damage induced by topoisomerase II inhibitors (Kurki et al. 2003), and DNA doublestrand breaks (DSBs) induced by ionizing radiation (Carbone et al. 2002; Dellaire et al. 2006a; Varadaraj et al. 2007). Therefore, the current model of PML NB function is that they are dynamic sensors of the cellular environment that regulate downstream pathways via the posttranslational modification of effector proteins (Dellaire and Bazett-Jones 2004; Bernardi and Pandolfi 2007). One of the most well-known targets of PML NB function is the critical tumor suppressor p53, which regulates the cell cycle checkpoints, apoptosis, and cell senescence (Kinzler and Vogelstein 1997; Levine 1997; Rodier et al. 2007). p53 is stabilized following cellular stress, allowing it to transactivate its transcriptional targets. This stabilization can occur at PML NBs through phosphorylation by the kinases casein kinase 1 (CK1) (AlsheichBartok et al. 2008), cell cycle checkpoint kinase 2 (Chk2) (Louria-Hayon et al. 2003), or homeodomain-interacting protein kinase 2 (HIPK2) (D’Orazi et al. 2002); through acetylation by the acetyltransferases CBP (Ferbeyre et al. 2000) or Tat interacting protein 60 (TIP60) (Wu et al. 2009); and potentially through de-ubiquitination by ubiquitin-specific protease 7 (USP7) (Everett et al. 1997; Li et al. 2002). The PML gene was originally identified as the fusion partner of the retinoic acid receptor-alpha (RARA) gene in a t(15;17) translocation in over 98% of acute promyelocytic leukemias (APLs) (de The et al. 1991; Kakizuka et al. 1991; Wang and Chen 2008). This initial link between PML and the oncogenesis of APL began its long-standing association with cancer biology (Dellaire and Bazett-Jones 2004; Bernardi and Pandolfi 2007). In APL, the PML-RARA oncoprotein disrupts the formation of the PML NBs (Dyck et al. 1994; Koken et al. 1994), and treatment of APL cells with all-trans retinoic acid results in the reformation of these structures (Koken et al. 1994). In fact, the presence or absence of PML NBs is used as a biomarker during immunohistochemical analysis for APL diagnosis and to monitor treatment efficacy, remission status, and the onset of relapse (Dyck et al. 1995). Furthermore, PML protein expression has been found to be reduced or eliminated in a significant proportion of a variety of other tumor types, including lymphomas, and lung, colon, breast, prostate, central nervous system (CNS), and germ cell cancers (Gurrieri et al. 2004).

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In addition to their function as tumor suppressors, PML NBs have another important connection to human health; they are also associated with the antiviral response (Nisole et al. 2005; Everett and Chelbi-Alix 2007). PML NBs are disrupted by herpes virus infection, due to expression of the immediate-early gene products that appear to specifically target the PML protein (e.g., IE1 of human cytomegalovirus [HCMV] and ICP0 of herpes simplex virus [HSV]) (Maul 1998). In addition, several nuclear-replicating DNA viruses (e.g., herpes viruses, adenoviruses, polyomaviruses, and papovaviruses) have been reported to have their parental genomes preferentially associated with PML NBs, and their initial sites of transcription and their DNA replication centers are often juxtaposed to these bodies or their remnants (Everett 2001).

CAJAL BODIES AND NUCLEAR GEMS Cajal bodies are highly dynamic structures of 0.2–1.0 μm in diameter and the average plant or animal cell contains 1–10 of these bodies (see Fig. 23.2C). A major component of Cajal bodies is the protein p80 coilin, which appears to be essential for body integrity and function, and serves as a marker for these structures. Cajal bodies are believed to be involved in small nuclear ribonucleoprotein (snRNP) biogenesis, guided by small Cajal-bodyspecific RNAs (scaRNAs) (Darzacq et al. 2002), and in trafficking of snoRNPs and snRNPs, which appear to move through the Cajal body en route to nucleoli or splicing speckles (Sleeman and Lamond 1999). For example, the spliceosomal U1, U2, U4/U6, and U5 snRNPs localize to these bodies, as well as the U7 snRNP, which is involved in histone 3′-end processing. The U3 and U8 snoRNPs, involved in pre-rRNA processing, also localize to Cajal bodies (Gall 2000). In addition, several gene loci, including histone, U1, U2, and the U3 genes appear to associate preferentially with these structures (Matera 1999). Cajal bodies lacking the coilin protein (Bauer and Gall 1997), also known as “residual Cajal bodies” (Tucker et al. 2001), are unable to recruit snRNPs. Surprisingly, neither coilin nor Cajal bodies appear to be essential for splicing as evident from the viability of coilin knockout mice (Tucker et al. 2001), consistent with the view that Cajal bodies increase efficiency of snRNP assembly by concentrating enzymes and substrates in one region of the nucleoplasm (Rippe 2007). The Cajal body was first described by Ramon y Cajal (Ramon y Cajal, 1903) while studying neuronal cell nuclei. They were first described as nucleolar accessory bodies due to their frequent appearance as a nucleolar cap in neurons (Hardin et al. 1969). They were given the name “coiled body” after ultrastructural studies revealed the bodies were composed of a tangle of coiled, electron-dense threads, approximately 0.5 μm in diameter (Monneron and Bernhard 1969; Malatesta et al. 2004). It was not until nearly 100 years after their first description that they were given the name Cajal body, in honor of Ramon y Cajal (Gall et al. 1999). Nuclear gems, also referred to as Gemini of coiled bodies, are often found paired or juxtaposed to Cajal bodies in mammalian nuclei and have been characterized by their association with the proteins’ survival of motor neurons (SMNs) and gemin2 (Matera 1999). Interestingly, a study of multiple tissues indicated that many mammalian cell types lack both gems and Cajal bodies (e.g., cardiac and smooth muscle, blood vessels, stomach, and spleen) (Young et al. 2000). It has since been suggested that Cajal body abundance is determined by RNA processing rates and snRNP levels in the nucleus (Sleeman et al. 2001; Cioce and Lamond 2005; Lemm et al. 2006). Cajal bodies and nuclear gems have both been implicated in disease through their association with the spinal muscular atrophy disease gene product, SMN (Carvalho et al.

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1999). SMN and an associated protein SIP1 form a complex that plays an essential role in cytoplasmic snRNP biogenesis (Fischer et al. 1997), which suggests that defects in spliceosomal snRNP assembly may be central to the development of spinal muscular atrophy.

NUCLEAR SPLICING SPECKLES Nuclear splicing speckles are thought to be storage sites for pre-mRNA splicing factors. From here, splicing factors can be recruited to RNA pol II transcription sites and nascent transcripts in the nucleoplasm. It is currently thought that varying the concentration of antagonistic splicing factors in the nucleoplasm may control alternative pre-mRNA splicing. Thus, speckles may function as reservoirs of splicing factors. In addition, speckles have been identified in a wide variety of metazoan species, underscoring the functional significance of this compartment (reviewed in Maniatis and Tasic 2002). The term “speckles” was first introduced by J. Swanson Beck in 1961 after observing rat-liver sections immunolabeled with serum from individuals with autoimmune disorders (Beck 1961). Unknown to Beck, these structures had been identified by Hewson Swift two years earlier and named interchromatin particles (Swift 1959). Swift observed a nonrandom distribution of particles localized to “clouds” within the nucleus. Cytochemical analysis of said “clouds” revealed the presence of RNA. The first connection with premRNA splicing came from examination of the distribution of snRNPs using antibodies specific to these splicing factors (Perraud et al. 1979; Lerner et al. 1981; Spector et al. 1983). The punctate staining of splicing factors observed by immunofluorescence microscopy (see Fig. 23.2D) is now referred to as splicing speckles. Electron microscopy analysis suggests that speckles may correspond to two distinct structural elements: IGCs and perichromatin fibrils (PFs). The IGCs have an apparent diameter of 0.8–1.8 μm and are composed of smaller 20- to 25-nm particles (Monneron and Bernhard 1969). IGCs, unlike PFs, do not appear to incorporate (3H)-UTP or Br-UTP and do not colocalize with DNA, suggesting that IGCs cannot be correlated with transcription sites or nascent transcripts (Fakan and Bernhard 1971; Wansink et al. 1993). PFs are structures found at the periphery of IGCs and have a diameter of 5–8 nm. PFs are thought to contain nascent transcripts, as they are sensitive to RNase treatment and are labeled rapidly with Br-UTP and (3H)-UTP (Cmarko et al. 1999). One possible relationship between IGCs and PFs is that splicing factors may be recruited from their depots in IGCs to nascent transcripts present in PFs. IGCs contain polyadenylated (poly A) RNA; however, it is unlikely that IGCs are sites of pre-mRNA splicing. Experiments employing laser scanning confocal microscopy and very high dilutions of primary antibodies against known speckle components have revealed that speckles may be much more heterogeneous and perhaps dynamic than previously thought. Mintz and Spector (2000) demonstrated that speckles are composed of numerous, smaller subdomains (subspeckles) ranging in size from 0.2 to 0.5 μm in diameter and are often arranged in loops. The loops of subspeckles are dependent on RNA pol II transcription and thus may be involved in targeting splicing factors to sites of transcription or RNA processing. Biochemical purification and characterization of IGCs has determined that at least 75 proteins are enriched in speckles, with the majority being RNA processing factors (Mintz et al. 1999). Interestingly, this approach also identified several novel gene products that may help to elucidate connections between RNA processing and other regulatory or metabolic pathways. The most extensively characterized occupants of speckles are the serine-argenine-rich (SR) protein family of essential pre-mRNA splicing factors (Fu 1995).

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SR proteins are a highly related, evolutionarily conserved family of nuclear phosphoproteins involved in both constitutive and alternative splicing. The primary structure of SR proteins consists of one or two amino-terminal RNA recognition motifs (RRMs) and a carboxyl terminal domain (CTD) that is enriched in arginine-serine dipeptides (RS domain). Sequence-specific RNA binding is conveyed by the RRMs, while the RS domain facilitates protein–protein interactions and contributes to their proper subcellular localization (Caceres et al. 1997). The modular domain structure of SR proteins reflects their roles as adapter molecules, mediating interactions between the pre-mRNA and the assembling spliceosome (Graveley 2000). Speckles are dynamic structures that respond to the levels of RNA pol II transcription in the nucleus. Fusion proteins between the green fluorescent protein (GFP) and the SR protein SF2/ASF has been a powerful tool in elucidating the functional relationship between speckles and pre-mRNA splicing. A variety of studies suggest that SR proteins are released from their storage sites in speckles and targeted to transcription sites in the nucleoplasm. Recruitment of SR proteins to transcription sites requires both the RS domain of SR proteins as well as the CTD of pol II (Misteli et al. 1997; Misteli and Spector 1999). The extent of serine phosphorylation within the RS domain is critical for regulating SR protein activities in vitro and in vivo. Accordingly, several kinases have been described that can induce the dissociation of SR proteins from speckles, thereby increasing the nucleoplasmic concentration (Gui et al. 1994; Colwill et al. 1996). The emerging picture of pre-mRNA splicing in vivo involves release of SR proteins from speckles by one or more kinase activities (Misteli et al. 1997, 1998). Once free, SR proteins are targeted to nascent transcripts via interactions with the CTD of RNA pol II, where spliceosome assembly begins. At the biochemical level, the higher phosphorylation state of liberated SR protein favors specific interactions with consensus sequences within the pre-mRNA as well as protein–protein interactions with the U1 snRNP (i.e., U1 70K) and perhaps U2AF35 (Xiao and Manley 1997, 1998). Once the spliceosome has assembled, dephosphorylation of SR proteins is necessary for catalysis and retargeting of SR proteins to speckles (Misteli 1999). Although SR proteins accumulate in the nucleus, a subset appears to shuttle continuously and rapidly between the nucleus and the cytoplasm, possibly escorting the spliced mRNA to its final destination in the cytoplasm (Caceres et al. 1998; Huang and Steitz 2001). The nucleocytoplasmic shuttling activity of SR proteins may be regulated by their state of phosphorylation and could prove to be a key regulatory target for the cell. Splice site selection in vivo is sensitive to the nuclear concentration of SR proteins and their antagonists; therefore, modulation of SR protein localization may be an effective way for the cell to control patterns of pre-mRNA splicing. More recently, SR protein function has also been expanded to the regulation of mRNA translation in the cytoplasm (Sanford et al. 2004; Michlewski et al. 2008). Nuclear splicing speckles have been intimately linked to human disease. One such disease is osteogenesis imperfecta type I (OI), which encompasses a group of inherited diseases characterized by bone fragility. OI is caused by a mutation in one of the two genes encoding collagen, COL1A1 and COL1A2 (reviewed in Roughly et al. 2003). One particular mutation causes improper splicing and retention of intron 26 of COL1A1 pre-mRNA resulting in decreased collagen production (Stover et al. 1993; Johnson et al. 2000). The addition of intron 26 within the transcript results in its sequestration in splicing speckles, ultimately rendering it unavailable for translation in the cytoplasm (Johnson et al. 2000). Another disease intimately linked to splicing speckles is myotonic dystrophy (DM) type 1. DM type 1 is characterized by skeletal muscle myotonia and degeneration and is caused by toxic CUG repeats in the 3′ untranslated region of DM protein kinase (DMPK) mRNA

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(reviewed by Cho and Tapscott 2007). Under normal conditions, DMPK will be transcribed at the periphery, escorted into splicing speckles, and escorted out of the nucleus as part of a mature mRNP. However, CUG repeat expansion prevents RNA entry into the speckles causing the messages to accumulate in foci at the periphery (Holt et al. 2007; Smith et al. 2007). These foci will sequester factors, including muscle-blind protein family 1 (MBNL1), preventing their normal nuclear activities, which includes the splicing of muscle-related transcripts (Dansithong et al. 2005).

POLYCOMB BODIES Polycomb or Pc-G bodies were first identified in the transformed human cell line U2-OS (Alkema et al. 1997). Staining these cells with antibodies against the mammalian Polycomb group protein Bmi1 revealed a punctate pattern reminiscent of the speckled distribution of PML nuclear bodies (Alkema et al. 1997). Pc-G bodies have subsequently been identified in a number of transformed mice and human cell lines (Voncken et al. 1999). The subcellular distribution of three Pc-G group proteins has been studied in Drosophila embryos (Buchenau et al. 1998). The punctate staining pattern seen previously in mammalian cells was not reproduced here, and a more diffuse nuclear staining was observed. Furthermore, primary mammalian cells lack Pc-G bodies, unless passaged for prolonged periods in tissue culture (Voncken et al. 1999) Although Pc-G bodies may well be a curiosity of the in vitro environment, the Pc-G proteins are of clear importance from flies to mammals (Gould 1997). The Pc-G genes of Drosophila, such as posterior sex combs (Psc) and suppressor two of zeste are required to maintain the correct patterns of Hox gene transcription. Loss of Pc-G function in flies leads to misexpression of Hox genes, causing the homeotic transformation of embryo parts. The role of Pc-G proteins in transcriptional repression may occur via discrete DNA elements. In support of this, Pc-G bodies have been shown to localize near pericentromeric heterochromatin (Saurin et al. 1998). The effects of Pc-G proteins in flies are broadly antagonistic to those of the trithorax group (trxG) genes. One such gene encodes the DNAdependent ATPase/helicase Brahma, which is related to a component of the SWI/SNF chromatin remodeling complex in budding yeast (Gould 1997). Mammalian homologs of the Pc-G genes have been identified, with conserved sequences and apparently conserved functions (Gould 1997). The homologs of Psc in mice, Bmi1 and Mel18, have several motifs in common with Psc, notably a RING finger, which is implicated in target specificity. Disruption of the genes encoding Mel18 and Bmi1 in mice gives rise to phenotypes consistent with a role for the mammalian Pc-G proteins in regulating Hox expression. Several Hox genes are misexpressed in both mutants, and the mice show vertebral transformations that have been interpreted as homeotic phenotypes. A third mouse Pc-G protein, M33, which shares the chromodomain of Psc, can partially rescue Drosophila Pc-G loss of function, suggesting that this may be a true homolog (Muller et al. 1995). Loss of M33 in mice also results in homeotic-like transformations. Another mammalian Pc-G gene, eed, is homologous to the Drosophila Pc-G gene extra sex combs (Core et al. 1997). Loss of eed function interferes with gastrulation in mice but may have roles in Hox regulation later in embryogenesis (Faust et al. 1995).

PNC The PNC was first discovered by performing immunofluorescence on cells using an antibody recognizing the polypyrimidine tract-binding protein PTB/hnRNP1 (Ghetti et al.

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1992). The PNC is found mostly in transformed cells and is rarely found in primary cells. This nuclear compartment is of irregular shape with a very variable size ranging from 0.25 to 1 μm in diameter. Analysis using electron microscopy revealed that this structure is composed of thick, 80–180 nm in diameter fibers, which are in direct contact with the nucleolus (Huang et al. 1997). Live imaging of cells expressing a GFP-tagged PTB revealed that the PNC is a dynamic structure moving along the periphery of the nucleolus over time (Huang et al. 1997). Several proteins have been described as being highly enriched within the PNC compared with their normal cellular localization. Most of them are RNA-binding proteins such as PTB/hnRNP 1 (Ghetti et al. 1992), CUG-binding protein (CUG-BP/hNab50) (Timchenko et al. 1996), the KH (hnRNP K homology domain)-type splicing regulatory protein KSRP (which is involved in splicing regulation) (Huang 2000), RAVER1 (Huttelmaier et al. 2001), RAVER2 (Kleinhenz et al. 2005), ROD1, and nucleolin (Kopp and Huang 2005). Also located in the PNC are small RNAs transcribed by RNA pol III, such as RNase MRP RNA, hY RNA, RNase P RNA (Matera et al. 1995; Lee et al. 1996), ALU, and SRP(7SL) (Wang et al. 2003). The function of the PNC remains largely unknown, but the presence of hnRNP proteins and splicing factors suggests a role for this compartment in RNA processing. Moreover, the presence of RNA transcripts that are not derived from RNA pol I and the fact that RNA pol II inhibitors affect the structural integrity of the PNC demonstrates that this structure is likely involved in either transcription or accumulation of newly synthesized RNA for further processing or retention (Huang et al. 1998). However, the exact function of the PNC requires further investigation. There also exists the potential for the PNC to function as a pan-cancer marker. It has been shown that PNC prevalence is 0% in normal breast tissue, and increases in parallel with clinical progression of disease reaching nearly 100% in distant metastasis (Kamath et al. 2005). This correlation between PNC prevalence and malignancy has been observed in a range of cancers, with results suggesting that PNCs form at advanced stages of transformation and are prominent in cells of high metastatic capacity (Pettaway et al. 1996; Norton et al. 2008). In this context, the PNC has potential as a prognostic marker for solid tissue tumors.

SAM68/SAM68-LIKE MAMMALIAN (SLM) NUCLEAR BODIES Sam68/SLM nuclear bodies (SNBs) are nuclear structures first visualized in HeLa cells using antibodies directed against Sam68 (Chen et al. 1999). Sam68 is an RNA-binding protein, and its activity is regulated by tyrosine kinases (Derry et al. 2000). SNBs are distinct and separate from other nuclear structures. SNBs are generally of spherical or ovoidal shape and measure ∼0.5–1 μm in diameter. HeLa cells usually have two to three SNBs per cell nucleus, frequently located in close proximity to the nucleolus, which is reminiscent of the PNC (Huang 2000). SNBs are fibrous structures that contain both proteins and nucleic acids. The nucleic acid is most likely RNA because of its fibrous and decondensed nature as visualized by electron microscopy. In addition to Sam68, two related Sam68 family members, SLM-1 and SLM-2 (also known as T-Star and Etoile), have been shown to localize in SNBs (Chen et al. 1999). Other proteins found to localize to SNBs include the alternative splicing factor YT521-B (Hartmann et al. 1999), the soluble tyrosine kinase breast tumor kinase/Src-related intestinal kinase (BRK/SIK) (Derry et al. 2000), and hnRNP A1-associated protein (HAP)

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(Denegri et al. 2001). Interestingly, the SNBs are able to recruit some splicing factors of the SR family upon stress induction (Denegri et al. 2001). The function of SNBs is unknown. SNBs have only been found in highly transformed and poorly differentiated cell lines, such as HeLa and BT-20, suggesting that SNBs may be a marker for these types of cancer (Chen et al. 1999). Many SLM proteins share SH2 and SH3 domains, suggesting that this family of proteins may serve as adaptor proteins for cellular protein kinases. In addition, their ability to bind RNA and the presence of splicing factors in SNBs suggests that they may play a role in coupling signal transduction pathways to pre-mRNA processing. Further support for a role in RNA metabolism comes from the evidence that rat SLM-2 can interact via a yeast two-hybrid system with several splicing factors (SRp30c, YT521-B, and SAF B) and can regulate the alternative splicing of a CD44 mini gene (Stross et al. 2001). SNBs have been implicated in cancer, although evidence supporting a specific role remains contradictory. For example, overexpression of Sam68 has been shown to induce G1 growth arrest while causing the downregulation of cyclin D1 (Taylor et al. 2004), suggesting that Sam68 may function as a tumor suppressor. On the other hand, Sam68 knockdown in prostate cancer cells is known to reduce proliferation and delay cell cycle progression, but cause an accumulation of cyclin D1 (Busa et al. 2007), suggesting that Sam68 may also function as a proto-oncogene. Mechanistically, there is evidence suggesting that Sam68 is the first splicing factor to promote CyclinD1b splicing in prostate cancer cells (Paronetto et al. 2010). Human cyclin D1 can be expressed as two isoforms, D1a and D1b, derived by alternative splicing of RNA. While both isoforms are frequently upregulated in human cancers, cyclin D1b displays a relatively higher oncogenic potential and a more pronounced nuclear localization (Lu et al. 2003; Solomon et al. 2003). In addition to cancer, SNBs have also been implicated in the neurodegenerative disorder fragile X-associated tremor/ataxia syndrome (FXTAS). FXTAS is characterized by gait ataxia, impaired executive cognitive functioning, and action tremors (Jacquemont et al. 2003), and has been proposed to be caused by titration of RNA-binding proteins by expanded CGG repeats (Sellier et al. 2010). Among the sequestered RNAbinding proteins is Sam68, which thereby loses its splicing regulatory function (Sellier et al. 2010).

SUMMARY Our current understanding of nuclear substructures would not have been possible without the development of advanced microscopy techniques, including fluorescence microscopy, live-cell imaging, and electron spectroscopic imaging (Dellaire et al. 2004; Olson and Dundr 2005; Handwerger and Gall 2006; Hernandez-Verdun 2006b; Trinkle-Mulcahy and Lamond 2007). We can now visualize and analyze protein interactions, nuclear trafficking, and nuclear body response to stimuli in vivo. Furthermore, we can map the nuclear and cellular localizations of almost any protein, RNA, or gene. These types of analyses led to the identification of many of the nuclear bodies described above and helped establish new functions for well-known nuclear bodies such as the nucleolus. Undoubtedly, the structures described above merely represent the beginning of our understanding of the nuclear landscape and how it is interconnected. Furthermore, an expanding list of human diseases and syndromes exhibit changes in nuclear morphology. As such, the biologies of the subnuclear compartments are providing novel insights into disease pathology and also potential therapeutic targets.

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ACKNOWLEDGMENTS G.D. is a Canadian Institutes of Health Research (CIHR) New Investigator and the Cameron Research Scientist in Cancer Biology of the Dalhousie Medical Research Foundation. D.C. is a recipient of a Dalhousie Cancer Research Training Program (CRTP) studentship from the Canadian Breast Cancer Foundation-Atlantic. K.C. is the recipient of a Nova Scotia Health

Research Foundation (NSHRF) Postdoctoral fellowship, as well as a Killam Postdoctoral Fellow. This work was supported by operating grants to G.D. from the NSHRF (MED-Project-2007-3348) and the CIHR (MOP-84260). The authors also acknowledge Christina Ridley for aid with Figure 23.1.

ABBREVIATIONS APL ARF CBP Chk2 CK1 Clb CNS CREB CT CTD DFC DM DMPK DSB FCs FXTAS GC GFP HCC HIPK2 ICD IGC MBNL1 MDM2 NDF NOR OI

acute promyelocytic leukemia alternative reading frame product of the CDKN2A locus CREB-binding protein cell cycle checkpoint kinase 2 casein kinase 1 B-type cyclin central nervous system cAMP (cyclic adenosine monophosphate) responsive element binding protein chromosome territory carboxyl terminal domain dense fibrillar component myotonic dystrophy DM protein kinase DNA double-strand break fibrillar centers fragile X-associated tremor/ataxia syndrome granular component green fluorescent protein hepatocellular carcinoma homeodomain-interacting protein kinase 2 interchromosome domain compartment interchromatin granule cluster muscle-blind protein family 1 murine double minute 2 nucleolus-derived foci nucleolar organizing region osteogenesis imperfecta type I

OPT PF PML PML NB PNB PNC POD Pol II pRB Psc PTB PTF RARA rDNA RPs RRM rRNA Sam68 SIM SLM SMNs SNBs snoRNP snRNP SR SUMO TIP60 USP7

Oct1, PTF transcription perichromatin fibril promyelocytic leukemia promyelocytic leukemia nuclear body prenucleolar body perinucleolar compartment PML oncogenic domain polymerase II retinoblastoma protein posterior sex combs polypyrimidine-tract-binding protein proximal sequence element-binding transcription factor retinoic acid receptor-alpha ribosomal DNA ribosomal proteins RNA recognition motif ribosomal RNA Src associated in mitosis 68 kDa protein SUMO-interacting motif Sam68-like mammalian survival of motor neurons Sam68/SLM nuclear bodies small nucleolar ribonucleoprotein small nuclear ribonucleoprotein serine-argenine-rich small ubiquitin-like modifier Tat interacting protein 60 ubiquitin-specific protease 7

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Mayer C, Grummt I. 2005. Cellular stress and nucleolar function. Cell Cycle 4:1036–8. Michlewski G, Sanford JR, Caceres JF. 2008. The splicing factor SF2/ASF regulates translation initiation by enhancing phosphorylation of 4E-BP1. Mol Cell 30: 179–89. Mintz PJ, Spector DL. 2000. Compartmentalization of RNA processing factors within nuclear speckles. J Struct Biol 129:241–51. Mintz PJ, Patterson SD, Neuwald AF, Spahr CS, Spector DL. 1999. Purification and biochemical characterization of interchromatin granule clusters. EMBO J 18:4308–20. Misteli T. 1999. RNA splicing: what has phosphorylation got to do with it? Curr Biol 9:198–200. Misteli T, Spector DL. 1999. RNA polymerase II targets pre-mRNA splicing factors to transcription sites in vivo. Mol Cell 3:697–705. Misteli T, Caceres JF, Spector DL. 1997. The dynamics of a pre-mRNA splicing factor in living cells. Nature 387:523–7. Misteli T, Caceres JF, Clement JQ, Krainer AR, Wilkinson MF, Spector DL. 1998. Serine phosphorylation of SR proteins is required for their recruitment to sites of transcription in vivo. J Cell Biol 143:297–307. Moll UM, Petrenko O. 2003. The MDM2-p53 interaction. Mol Cancer Res 1:1001–8. Monneron A, Bernhard W. 1969. Fine structural organization of the interphase nucleus in some interphase cells. J Ultrastruct Res 27:266–88. Montanaro L, Trere D, Derenzini M. 2008. Nucleolus, ribosomes, and cancer. Am J Pathol 173:301–10. Morey C, Da Silva NR, Perry P, Bickmore WA. 2007. Nuclear reorganisation and chromatin decondensation are conserved, but distinct, mechanisms linked to Hox gene activation. Development 134:909–19. Morey C, Kress C, Bickmore WA. 2009. Lack of bystander activation shows that localization exterior to chromosome territories is not sufficient to up-regulate gene expression. Genome Res 19:1184–94. Muller J, Gaunt S, Lawrence PA. 1995. Function of the polycomb protein is conserved in mice and flies. Development 121:2847–52. Negorev D, Maul GG. 2001. Cellular proteins localized at and interacting within ND10/PML nuclear bodies/ PODs suggest functions of a nuclear depot. Oncogene 20:7234–42. Nisole S, Stoye JP, Saib A. 2005. TRIM family proteins: retroviral restriction and antiviral defence. Nat Rev Microbiol 3:799–808. Noordermeer D, Branco MR, Splinter E, Klous P, van Ijcken W, Swagemakers S, Koutsourakis M, van der Spek P, Pombo A, de Laat W. 2008. Transcription and chromatin organization of a housekeeping gene cluster containing an integrated beta-globin locus control region. PLoS Genet 4:e1000016. Norton JT, Pollock CB, Wang C, Schink JC, Kim JJ, Huang S. 2008. Perinucleolar compartment prevalence is a phenotypic pancancer marker of malignancy. Cancer 113:861–9.

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THE NUCLEAR PORE Richard W. Wozniak Christopher Ptak John D. Aitchison

DEFINITION Eukaryotic cells segregate the majority of their genetic material within an enclosed doublemembrane system termed the nuclear envelope (NE; see Fig. 24.1). This membrane defines the periphery of the nucleus and consists of several distinct domains. The outer nuclear membrane is continuous with the endoplasmic reticulum (ER) membrane (see Chapter 7) and shares many functional features, including attached ribosomes and other components of the protein translocation machinery. This membrane continuity also establishes direct access of the lumenal contents of the NE to that of the ER. Paralleling the outer nuclear membrane, and separated by the NE lumen, is the inner nuclear membrane. This membrane lies directly adjacent to the underlying chromatin and contains a unique set of membraneassociated proteins that facilitate this interaction and influence chromatin structure and gene expression, thereby linking the structural organization of the NE with the chromatin (see Chapter 23). Some of these NE membrane proteins interact with the lumenal segments of outer membrane proteins, creating protein bridges that extend across the NE from the chromatin to cytoplasmic filaments that include actin fibers and microtubules (see Chapters 12 and 14). Beyond these interactions, direct continuity between the inner and outer nuclear membranes exists at numerous sites along the nuclear surface where a connecting cylindrical membrane, termed the pore membrane domain, extends across the nuclear envelope. These transcisternal pores (∼90 nm in diameter) are occupied by elaborate macromolecular structures termed nuclear pore complexes (NPCs) that project into the cytoplasm and the nucleus. These structures control the transport of all molecules across the nuclear envelope. This critical nexus has long been the focus of studies investigating the role of NPC in transport and, more recently, its function in dictating chromatin structure. In the following sections we will discuss the discovery and molecular characterization of the NPC and its role in regulating nuclear transport.

HISTORICAL PERSPECTIVE In 1682, the microscopist Thonius Philips van Leeuwenhoek first described a predominant intracellular structure in the red blood cells of fish which, a century and a half later, was termed the nucleus by the botanist Robert Brown following his studies of orchid leaves (Brown 1833). Many years after this, Hartog J. Hamburger was able to physically manipulate nuclei with microneedles, and in 1904 he reported the idea that the nucleus was Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

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Figure 24.1. Schematic diagram of the nuclear envelope (NE). Shown is a cartoon of the NE in cross-section. The NE is composed a double membrane that includes a cytoplasmic facing outer nuclear membrane (ONM) and a nucleoplasmic facing inner nuclear membrane (INM). The ONM is contiguous with the endoplasmic reticulum (ER). The INM and ONM are separated by a lumen termed the perinuclear space, which itself is connected to the ER lumen at ER–ONM junctions. Both the ONM and ER surfaces may be studded with ribosomes functioning in co-translational integration or translocation of proteins into and across these membranes. Among these proteins are those that are specifically localized to the ONM and INM (depicted in purple, gray, and pink). The INM-localized proteins may also interact with lamins or chromatin, while the ONM-localized proteins may contact cytoskeletal structures, such as actin fibers or microtubules. ONM KASH (Klarsicht, ANC-1, Syne Homology) domain proteins can also interact with the INM SUN (Sad1p, UNC-84) domain proteins creating a bridge across the perinuclear space and forming a physical link between the lamins and the cytoskeleton (reviewed in Burke and Roux 2009). The lamins themselves form a peripheral meshwork along the nucleoplasmic face of the INM, where, in association with other proteins, extensive contacts are made with heterochromatin. The lamina is absent at sites along the INM, where the INM and ONM are fused to form nuclear pores with a diameter of ∼100 nm. The connecting membrane forming the wall of the pore is referred to as the pore membrane. Positioned within the pore is the nuclear pore complex (NPC), integral membrane proteins, known as Poms (shown in olive), which assist in connecting NPCs to the pore membrane. Unlike the lamins, NPCs are not associated with heterochromatin but rather generally lie directly adjacent to the less condensed euchromatin. Detailed structural models of the NPC are shown in Figures 24.2 and 24.3.

“membrane” bound. However, like other organelles, the structure of the NE was only revealed with the advent of electron microscopy and its application to the analysis of cell structure, which began in the late 1940s and early 1950s. During this period the general structural features of the NE began to emerge, as did the term nuclear envelope (Anderson 1953). The presence of discontinuities was observed in cross-sections of the NE, as was a connecting membrane between the inner and outer nuclear membranes (the pore membrane) (Bahr and Beermann 1954; Gall 1954; Watson 1955). These early studies also revealed densely staining material within the pores that appeared as ring-like structures in en face views of the nuclear surface. The term pore complex was coined to describe these structures. In the years that followed, the structural complexity of the NPC was progressively revealed. Wischnitzer (1958) and Gall (1967) reported that the NPC was not merely a ring-shaped structure, but rather it exhibited octagonal symmetry about an axis perpendicular to the NE and through the NPC. As models emerged in the early 1970s, they depicted the eightfold symmetrical structures of the NPC as eight particles or granules organized to form rings on the cytoplasmic and nucleoplasmic sides of the pore that,

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Figure 24.2. Model of the nuclear pore complex (NPC). The NPC is a large proteinaceous structure that extends across the nuclear envelope at points where the inner and outer nuclear membranes are fused, creating points of transit between the nucleoplasm and cytoplasm. Shown is a model of one NPC within the nuclear envelope with the outer and inner membranes indicated. The complete NPC model (left) and a cross-sectional view (right) are shown. Major structural elements are indicated. FG-Nups are located along the filaments, the central channel, and the nuclear basket. Non-FG-Nups form the spoke and ring structures.

together, sandwiched additional eightfold symmetrical structures within the center of the pore (Franke 1970; Roberts and Northcote 1970; Fabergé 1973; Hoeijmakers et al. 1974). In 1982, a further breakthrough came from the studies of Unwin and Milligan, who used Fourier averaging methods of negatively stained Xenopus oocyte NEs to develop a 9-nm resolution map of the NPC (Unwin and Milligan 1982). The seminal features of their model were parallel cytoplasmic and nucleoplasmic rings surrounding eight spoke-like structures that radiated from the pore membrane and surrounded the central channel of the NPC. As imaging methods improved, filaments emanating from the NPC into the cytoplasm and the nucleoplasm became evident (see Fig. 24.2). The advent of molecular cloning, structural biology, proteomics, and computational biology has subsequently defined the molecular constituents and their positions in the NPC (see below), revealing a remarkable structure tuned by evolution to govern transport and an increasing number of additional functions associated with its central position linking the nucleus and the cytoplasm (reviewed in Brohawn et al. 2009; Elad et al. 2009; Strambio-De-Castillia et al. 2010).

MOLECULAR ORGANIZATION OF THE NPC Based on the analysis of many eukaryotes as widely diverse as humans and trypanosomes, it is believed that the overall structural organization of the NPC appears conserved throughout the Eukaryota domain (Mans et al. 2004; DeGrasse et al. 2009). The molecular analysis of NPC structure has been largely conducted in vertebrates and yeast. Electron microscopy data and, in yeast, more extensive analysis of isolated structures, estimated the mass of the NPC to lie within a range of 50–100 million Daltons (Reichelt et al. 1990; Rout and Blobel 1993; Yang et al. 1998; Rout et al. 2000; Alber et al. 2007b). The majority of the structure is well conserved (Yang et al. 1998) and some NPC proteins (termed nucleoporins or Nups) can function across phyla from yeast to mammals (Aitchison et al. 1995; Grandi et al. 1997; Watkins et al. 1998). Over the last two decades, biochemical and genetic approaches have led to the identification of most, if not all, Nups in vertebrates and yeast (primarily Saccharomyces cerevisiae) (reviewed in Suntharalingam and Wente 2003;

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Strambio-De-Castillia et al. 2010) culminating with mass spectrometric (MS) analyses of enriched fractions of yeast and rat NPCs that suggest NPCs are composed of ∼30 Nups (Rout et al. 2000; Cronshaw et al. 2002). Immunoelectron microscopy analysis has revealed that most of the Nups are positioned on both faces of the NPC, suggesting the nucleoplasmic and cytoplasmic rings were composed of similar Nups, which reflects the morphological symmetry of the core NPC structure. Beyond the core, symmetry is lost; the nuclear filaments form a basket-like structure, and the cytoplasmic filaments appear disordered. Accordingly, some Nups localize to only one side of the NPC and are thus believed to be an exclusive component of either the cytoplasmic filaments or the nuclear basket (Rout et al. 2000). At a superficial level, Nups can be divided into three subgroups that include: (1) integral pore membrane Nups, also termed Poms, (2) scaffold/structural Nups, and (3) phenylalanine–glycine (FG)-repeat-rich Nups, or FG-Nups. The first group consists of integral membrane proteins positioned within the pore membrane, of which three have been identified in yeast (Pom34p, Pom152p, and Ndc1p) and three in vertebrates (gp210, NDC1 and POM121) (Wozniak and Blobel 1992; Hallberg et al. 1993; Wozniak et al. 1994; Chial et al. 1998; Rout et al. 2000; Mansfeld et al. 2006; Stavru et al. 2006). Aminoacid sequence comparisons have also shown that, in spite of the functional importance of these proteins (see below), they are not well conserved, with the exception of Ndc1 (Mansfeld et al. 2006; Stavru et al. 2006). This lack of conservation reflects the observation that, in some organisms, no single Pom appears to be absolutely required for NPC assembly and maintenance. This also illustrates redundancies inherent within NPCs that contribute to the robustness of its structural integrity (Mans et al. 2004; Madrid et al. 2006; Mansfeld et al. 2006; Stavru et al. 2006; Onischenko et al. 2009). Each Pom has been predicted to possess at least one transmembrane segment that places additional domains on either side of the pore membrane (Mans et al. 2004). With domains on both sides of the membrane, these proteins contribute to structural features of the NPC within the pore, including the rings and spokes, as well as to a ring within the lumen of the NE adjacent to the lumenal face of the pore membrane (see Fig. 24.3). The functions of these membrane proteins are not well understood; however, it has long been proposed that they play a role in NPC assembly and anchoring of the structure to the membrane. The former idea is supported by various studies in both yeast and vertebrate systems showing that depletion of specific membrane proteins blocks the assembly of NPCs (Antonin et al. 2005; Mansfeld et al. 2006; Stavru et al. 2006; Madrid et al. 2006; Onischenko et al. 2009; Doucet et al. 2010). It is also important to consider that certain pore membrane proteins may also perform additional duties necessary for normal NE structure and function. For example, the yeast spindle pole body is inserted into the NE such that it spans the inner and outer NE membrane in a manner analogous to that of the NPC. The membrane protein Ndc1p is associated with both NPCs and spindle pole bodies (Chial et al. 1998), and is thus suspected to play a similar biogenic and structural role that contributes to their common topology across the NE. Indeed, certain mutant forms of Ndc1p inhibit spindle pole body duplication (Winey et al. 1993). In vertebrates, where the NE disassembles and then reassembles during the course of mitosis, the pore membrane protein Pom121 appears to function in an NE assembly checkpoint, which prevents the formation of an intact NE if key steps in NPC assembly are not completed (Antonin et al. 2005). Within NPCs, binding partners of the Poms include members of the second broad grouping of Nups, namely those that form the core scaffold of the NPC and coat the pore membrane. These Nups appear to form the eightfold symmetrical framework of the NPC, with each of the eight repeated units containing one, or more often, several copies of each

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Figure 24.3. Nuclear pore complex structural model. Shown is a map of the positions of various Nups within substructures of the yeast NPC (adapted from Alber et al. 2007b). In the top half of the figure, proposed substructures of the NPC (outer, inner, and membrane rings, linker Nups, and FG-Nups) formed by complexes of Nups are shown. In the bottom half of the figure, the positions of individual Nups within one of the eight spokes of the NPC is shown. Membrane proteins (Pom152, Pom34, and Ndc1) contribute to attaching the NPC core scaffold, consisting of the Nup170 and Nup84 subcomplexes, as well as the linker Nups Nic96 and Nup82, to the membrane. The NPC core scaffold forms the framework on which the FG-Nups (green) are attached.

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scaffold Nup. Within each of these repetitive units, the Nups are in turn organized into distinct subcomplexes (reviewed in Suntharalingam and Wente 2003; Schwartz 2005; Lim and Fahrenkrog 2006; Strambio-De-Castillia et al. 2010). The identification of these subcomplexes derives from studies conducted by various groups analyzing nearest neighbor associations between individual Nups following partial NPC disassembly (reviewed in Suntharalingam and Wente 2003). In many cases, these results are also supported by genetic analyses (Nehrbass et al. 1996; Zabel et al. 1996; Marelli et al. 1998; Siniossoglou et al. 2000; Seo et al. 2009). While these approaches have proven fruitful in identifying subcomplexes and assisting in the characterization of their functions, they have not led to a detailed architectural picture of how these subcomplexes are molded together to form the NPC. Recently, through an elegant biochemical and computational study (Alber et al. 2007a, b), Alber and colleagues have proposed a structure for the yeast NPC that provides a comprehensive view of the arrangement of subcomplexes in the NPC (see Fig. 24.3). Their architectural model of the NPC was developed using a computational approach that explores the placement of each of the 456 individual Nups into a single macromolecular ensemble. The practically infinite number of ways in which individual Nups could be assembled was limited by the large amount of experimental data acquired over many years. These included: NPC size and symmetry; a Nup inventory including their stoichiometry, size, and shape; a low precision localization map of each Nup within the NPC that was established by immunoelectron microscopy; and a large Nup–Nup protein interaction dataset established by affinity purification and mass spectrometry. The final ensemble of solutions that simultaneously satisfied all of these restraints converged to a structure of the NPC that positioned each Nup with a precision of ∼5–7 nm (see Fig. 24.3). Among the predictions of the Alber model is the placement of two NPC subcomplexes, the Nup170- and Nup84-containing complexes, adjacent to the surface of the pore membrane. This architecture is consistent with several observations that supported a physical and functional link between the Nup170p-containing complex and pore membrane proteins (Makio et al. 2009; Onischenko et al. 2009). Moreover, the close association of these Nups with the membrane was consistent with the hypothesis that the pore membrane curvature required for NPC assembly occurs through a mechanism similar to those used by vesicle coating complexes such as COPII and clathrin (see Chapter 2). This idea was spawned from structural predications based on sequence analysis of the scaffold Nups that has been validated and extended through an increasing number of crystallography studies. These structural studies revealed that components of the Nup170- and Nup84-containing subcomplexes consist of either a β-propeller, an α-solenoid, or a combination of both folds arranged as an N-terminal β-propeller and a C-terminal α-solenoid domain (Berke et al. 2004; Devos et al. 2004; Schwartz 2005; Devos et al. 2006; Brohawn et al. 2008; Whittle and Schwartz 2009). Similar structural domains and their organization into larger macromolecular assemblies have also been identified within the vesicular coat protein complexes that include clathrin and the Sec31-Sec13 heterodimer (Fotin et al. 2004; Brohawn et al. 2008; Brohawn and Schwartz 2009). These similarities, with respect to structure as well as membrane coating and curvature-inducing functions, coupled with phylogenetic analyses, have given rise to the concept that NPCs, COPs, and clathrin coats arose through the evolutionary divergence of a protocoatomer found in the last common eukaryotic ancestor (Devos et al. 2004; Mans et al. 2004; reviewed in Field and Dacks 2009). Multiple copies of the Nup170- and Nup84-containing subcomplexes coat the walls of the pore membrane and are major components of the eight repetitive, spoke-like structures that form the framework of the NPC (see Fig. 24.3). Each spoke itself is thought to be symmetrical relative to a plane parallel to the NE and passing through the center of the spoke with Nup84 and Nup170 subcomplexes on each side of the plane (Rout et al. 2000;

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Alber et al. 2007b). Recent crystallographic studies on the components of the Nup84 subcomplex have provided structural data leading to two distinct models of how they contribute to the spokes and architecture of the central channel (reviewed in Brohawn et al. 2009). In the lattice model, two sets of eight Nup84 subcomplexes form an upper and lower ring that sandwich, and so are linked by, two sets of eight Nup170 subcomplexes (also referred to as the Nic96 subcomplexes) (Alber et al. 2007b; Brohawn et al. 2008; Brohawn and Schwartz 2009). In the concentric ring model, 32 copies of the Nup84 subcomplex are predicted to form four eight-membered rings that are stacked one upon another and positioned along the surface of the pore membrane. These rings are proposed to form a porous “fence” and allow interdigitating Nups, including, for example, those of the Nup170 subcomplex, and Poms to interact through the fence (Hsia et al. 2007; Debler et al. 2008; Seo et al. 2009). Regardless of the specific model, the scaffold possesses two faces: a pore membrane face that supports membrane curvature and links the scaffold to the membrane through interactions with the Poms (Onischenko et al. 2009) and a channel face that lines the central channel of NPC and functions as a support for the centrally located FG-Nups (Rout et al. 2000). These Nups, of which there are 13 in yeast and 9 counterparts in humans, are rich in FG dipeptide repeats and, like the scaffold Nups, are present within the NPC in multiple copies. Ultimately, their localization within the central channel of the NPC positions them to play a direct role in the movement of macromolecules through the NPC (reviewed in Terry and Wente 2009). Each of the FG-Nups contains two structurally distinct regions. One domain is defined by the presence of the FG-repeat containing peptides. Depending on the Nup, between 4 and 48 repeats are distributed within a region, varying in length from ∼150 to 700 amino-acid residues (Strawn et al. 2004). Importantly, the FG-repeats and variably sized spacer regions between them form a domain that appears flexible and does not fold into secondary structures, such as extended α-helices or β-sheets, when examined using in vitro assays. In other words, they appear to adopt a natively unfolded structure (Denning et al. 2002; Denning et al. 2003; Dokudovskaya et al. 2006; Lim et al. 2006). This is in contrast to adjacent, often C-terminal, regions that generally lack FG-repeats and are predicted to contain coiled coil motifs. These more structured regions are thought to anchor the FG-Nups to the scaffold Nups, thus positioning the FG-repeat-containing regions to extend into the central channel of the NPC (see Devos et al. 2006). The FG-Nups also contribute to the cytoplasmic and nuclear filaments that extend from the core into the cytoplasm and the nucleoplasm (reviewed in Terry and Wente 2009). Understanding the structural organization of the FG-Nups within the context of the NPC remains one of the major unsolved issues in the field, and the emerging models are hotly debated. As these Nups occupy the central channel of the NPC, understanding their structure and organization within this space is essential to developing models for the mechanisms of transport through the NPC. Two preferred models have emerged in recent years with a key difference being the nature of the interactions between individual FGNups. With upwards of 128 FG-Nups per pore, FG-Nups are predicted to be tightly packed within the confines of the NPC (reviewed in Terry and Wente 2009). In one model, their arrangement is proposed to mimic “polymer brushes” with limited intermolecular interactions between individual FG-Nups (Lim et al. 2006). Polymer brushes have been studied for many years by chemists and material scientists (Milner 1991). Generally speaking, these structures are characterized by flexible (e.g., unfolded) polymers that extend from a surface with little interactions between adjacent molecules, much like bristles on a brush. Characteristics of the extended polymers, including their shape and the degree of extension of the polymer, depend on various factors including their composition and the degree of their compaction along the support surface. In the polymer brush model, FG-Nups would

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extend from the scaffold structures with the FG-repeat-containing regions toward the central axis of the NPC (Rout et al. 2000; 2003; Lim et al. 2006). A second model suggests a high degree of intermolecular interactions between FGNups within the central channel. In this “hydrogel” model, the FG-Nups are proposed to associate with one another through extensive hydrophobic interactions mediated by their FG-repeat peptides (Ribbeck and Görlich 2001, 2002). This densely packed and interconnected network of FG-Nups is proposed to form a structure akin to a gel. In vitro the FG-repeat-containing region of one yeast Nup, Nsp1p, can, in fact, form a gelatinous material. Within the NPC the hydrogel would occupy the central channel and create a hydrophobic barrier between the cytoplasm and the nucleoplasm (Frey et al. 2006; Frey and Görlich 2007).

TRANSPORT THROUGH THE NPC Whether in the form of a polymer brush or a hydrogel, the presence of the FG-Nups within the central channel of the NPC prevents the passage of macromolecules greater than ∼9 nm in diameter, while allowing metabolites and other small molecules to pass through the NPC by simple diffusion (reviewed in Terry and Wente 2009). For most proteins and ribonucleoprotein particles (see Chapter 25), travel through the NPC requires interactions with soluble nuclear transport factors (NTFs). The NTFs are proteins that recognize nuclear localization signals (NLS) or nuclear export signals (NES) on these molecules that generally consist of a short peptide on the surface of the protein. Once bound to a cargo molecule the NTF functions to escort the complex through the NPC (reviewed in Wozniak et al. 1998; Marelli et al. 2001; Pemberton and Paschal 2005). Several NTFs are directly involved in mRNA export and the import of the small GTPase Ran. Their functions have been reviewed elsewhere (Stewart 2000; Carmody and Wente 2009). However, most NTFs are members of a group of structurally related proteins termed β-karyopherins, referred to here simply as Kaps, which are further subdivided into two groups: those involved in nuclear import (importins) and those functioning to export molecules from the nucleus (exportins). There are ∼20 Kaps in metazoan cells and 14 in yeast. Each Kap appears to bind a different set of cargos; however, there is some redundancy between members of certain cargo sets as reflected by the observation that some macromolecules are imported by more than one Kap (reviewed in Weis 2002). On the basis of crystallographic analysis of individual Kaps and secondary structure predictions, it is believed that Kaps are composed of a series of ∼20 HEAT (Huntington, Elongation Factor 3, PR65/A, TOR) repeats that are connected together to give the proteins a superhelical structure (reviewed in Cook et al. 2007). Each HEAT repeat is ∼40–50 amino-acid residues in length and is composed of two antiparallel α-helices separated by a turn (Andrade et al. 2001). While the overall structure of the Kaps is similar, their sequence similarity is low. This is especially true beyond the first 200 amino-acid residues where the variability of their sequences likely reflects their ability to recognize different transport signals. The greatest degree of sequence similarity between Kaps lies within their first 150–175 amino-acid residues (Conti et al. 2006; Quan et al. 2008). This region contains a binding site for the GTP-bound form of Ran (RanGTP; see Chook and Blobel 1999; Vetter et al. 1999; Matsuura and Stewart 2004; Lee et al. 2005), which plays a key role in regulating the binding of Kaps to their cargos (see below). In addition to binding signal-containing cargos and RanGTP, Kaps also contain multiple binding sites within their HEAT repeats for the FG-repeat-containing regions of this group of Nups. The exact number of FG-binding sites in any given Kap is still not

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clearly defined. One of the most intensely studied Kaps is importin β/Kap β1 (or Kap95 in yeast). It contains at least two identified FG-Nups binding sites (Bayliss et al. 2000). This may, however, only be the tip of the iceberg. For example, molecular dynamic simulations suggest that another Kap, the exportin Cse1, may have as many as 14 Nup binding sites (Isgro and Schulten 2007). It is now clearly established that the interactions of the various Kaps with the ∼128 FG-Nups, and their ∼3500 FG-repeats peptides, present in the NPC is essential for the transport of the Kaps and their cargos through the central channel of the NPC (Rout et al. 2000; Cronshaw et al. 2002). However, the nature of these interactions and the means by which they facilitate transport is unclear. Much of our current knowledge has come from in vitro binding assays. These studies have led to the conclusion that the interactions of Kaps with individual FG-Nups are of a relatively low affinity (Bayliss et al. 1999; BenEfraim and Gerace 2001; Bayliss et al. 2002; Pyhtila and Rexach 2003). This may allow Kaps to rapidly move between different FG-Nups during transport through the NPC (Macara 2001; Rout et al. 2003; Timney et al. 2006). Also complicating the picture are data supporting the conclusion that different Kaps preferentially bind different subsets of FG-Nups. This idea has led to the proposal that specific Kaps may follow a distinct pathway through the NPC (Marelli et al. 1998; Makhnevych et al. 2003; Strawn et al. 2004; Fiserova et al. 2010). Furthermore, at least some Kaps also bind to unique, non-FGcontaining regions within FG-Nups. In one case, this interaction is cell cycle regulated and functions to prevent translocation of the interacting Kap and inhibits the import of its cargos. Two distinct models (and variations thereof) dominate our thinking around the mechanisms by which the interactions of Kaps (and other NTFs) with FG-Nups facilitate their movement through the central channel of the NPC. Both models rely on the assumption that the in vivo associations of NTFs with the FG-Nups are mediated by low-affinity interactions, as detected using in vitro assays. This property would allow the NTFs to partition into the FG-Nup-rich environment of the NPC yet allow them to move between FG-repeats via rapid dissociation–association cycles. Both models also have the feature of a virtual gate. That is, the physical properties of FG-Nups prevent macromolecules having a diameter larger than ∼9 nm to move through NPCs. This impediment is overcome through interactions between NTFs and the FG-Nups. It is believed that these interactions allow NTF–cargo complexes to transit through NPCs virtually unimpeded, at rates near diffusion, and without the need of a mechanical gate or an exogenous energy source. The key difference between these models lies in the two proposed molecular states of the FGNups within the NPC—either the hydrogel or the polymer brush. As discussed above, a fundamental property of the hydrogel proposal is that the intermolecular interactions of the FG-Nups lead to numerous hydrophobic interactions among FG domains, thereby forming a gel that occludes the channel (Ribbeck and Görlich 2001, 2002). In a transport model based on this configuration, the ability of the NTFs to bind FG-repeats is proposed to disrupt their intermolecular interactions, effectively causing localized melting of the hydrogel and partitioning of the NTF into the NPC channel. Since the FG–NTF interactions are weak, and thus transient, cycles of dissociation–association would allow the NTF–cargo complex to move from one FG-repeat to another until it exits the hydrogel. Those molecules that fail to bind the FG-repeats (i.e., most cellular proteins) would be unable to disrupt or penetrate the mesh and would be prevented from moving through the NPC (Frey et al. 2006; Frey and Görlich 2007, 2009). The second model considers nuclear transport in the context of a polymer brush wherein the FG-Nups are dynamic and dominated more by their entropy than by their interactions with one another. In this state, Brownian motion of the unfolded

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FG-repeat-containing regions is predicted to exclude macromolecules from the central channel, as their passage through the NPC would cause crowding and a loss of entropy that would be energetically unfavorable. However, the weak binding of NTFs to FGrepeats would offset the entropic cost of crowding, thus allowing NTFs to move through the dense and dynamic FG-Nup-rich environment of the NPC central channel (Rout et al. 2000; reviewed in Rout et al. 2003). When evaluating the plausibility of these two general models, it is important to keep in mind that, while distinctly different in their predictions for structural organization of FG-Nups and the mechanism NTF transport, elements of both models may contribute to the in vivo situation. For example, those FG-Nups that lie within the central regions of the transport channel are likely to be more tightly packed and restricted in their movement, a condition that could foster the formation of a gel matrix. However, those FG-Nups lying adjacent to the entrance of the central channel or contributing to the cytoplasmic or nucleoplasmic filaments are likely to be less tightly packed, which favors entropic considerations and hence a polymer brush-like structure (Patel et al. 2007). Furthermore, an alternate model suggests that the FG-Nup polymer brushes collapse on NTF binding into a more globule structure sustained by the presence of both the NTF and intermolecular FG-Nup interactions (Lim et al. 2006; 2007a; 2007b). Thus, it seems possible, and perhaps, likely, that features from each of these various models may reflect the in vivo situation. Irrespective of their specific features, all of the models predict essentially no net cost of energy for the NTFs to enter the NPC, and once there, they can just as easily exit, either into the cytoplasm or into the nucleoplasm. As discussed below, directionality appears to stem from disruption of NTF–cargo interactions mediated by RanGTP, or additional interactions in the target compartment.

ESTABLISHING DIRECTIONALITY FOR NUCLEAR TRANSPORT The models for nuclear transport outlined above predict a mechanism for the selective permeability of the NPC by NTFs and their attached cargos. Importantly, the NPC is not believed to impart directionality to the transport process. In other words, whether the NTF is an importin or exportin, it is predicted to move indiscriminately in either direction through the NPC. Rather, the unidirectional movement of cargos into or out of the nucleus is established by the small GTPase Ran and molecules that establish a gradient of RanGTP across the NE (reviewed in Görlich and Kutay 1999; Wente and Rout 2010). RanGTP is believed to be the primary energy source required to establish the concentration gradient of cargos across the NE. Ran exists in two states in the cell; in the nucleus it is present largely in its GTP-bound form due to the presence of a chromatin-associated Ran guanine nucleotide exchange factor (RanGEF). RanGEF binds RanGDP, thereby promoting the exchange of Ran-bound GDP for GTP (present in higher concentrations in the cell). By contrast, RanGTP present in the cytoplasm is rapidly converted to RanGDP by the presence of a Ran GTPase-activating protein (RanGAP) in the cytoplasm or attached to the cytoplasmic face of the NPC (depending on the organism). RanGAP binds to Ran and greatly stimulates its intrinsic GTPase activity. Together, the Ran effector molecules produce high concentrations of RanGTP in the nucleoplasm and low concentrations in the cytoplasm, creating a potential energy gradient across the NE. It is the high concentration of nuclear RanGTP that contributes to the directionality of transport. Once an importincargo complex crosses the NPC, it binds to RanGTP. Importantly, this binding triggers the release of the cargo in the nucleus. While the importin bound to RanGTP can return to the

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cytoplasm, the cargo is left behind in the nucleoplasm. Binding to RanGTP has a distinctly different effect on exportins. In this case, RanGTP binding to an exportin stimulates a cooperative interaction between the exportin and a NES-containing export cargo. This complex is stable until reaching the cytoplasm, where it encounters the RanGAP that stimulates Ran to hydrolyze GTP, leading to disassembly of the export complex.

SUMMARY AND FUTURE PERSPECTIVES The last 20 years have seen a blossoming in our understanding of the mechanisms that control the movement of macromolecules into and out of the nucleus. From the identification of targeting signals and their varied receptors (NTFs) to the structural and functional characterization of the NPC, we now have a solid understanding of the basic principles that govern nuclear transport. As this field moves forward to understand the molecular mechanisms of transport, many questions remain to be answered. Prominent among these is an understanding of the molecular basis for the movement of NTFs through the NPC, and the dynamic changes that occur in this structure during the transport process. Future work is likely to uncover additional functions for the NPC and its component parts that extend beyond nuclear transport. For example, recent studies point to a role for the NPC in regulating chromatin structure and gene expression (reviewed in Van de Vosse et al. 2011). In addition, Nups have been suggested to play a role in the spindle assembly checkpoint and, in some organisms, they are capable of interacting with kinetochores during mitosis (reviewed in Wozniak et al. 2010). As these studies mature, undoubtedly, new chapters will emerge on the functions of the nuclear pore.

ABBREVIATIONS ER FG GAP GDP GEF GTP Kap MS

endoplasmic reticulum phenylalanine–glycine GTPase-activating protein guanosine diphosphate guanine nucleotide exchange factor guanosine triphosphate karyopherin mass spectroscopy

NE NES NLS NPC NTF Nup Pom

nuclear envelope nuclear export signal nuclear localization signal nuclear pore complex nuclear transport factor nucleoporin pore membrane nucleoporin

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Schwartz TU. 2005. Modularity within the architecture of the nuclear pore complex. Curr Opin Struct Biol 15: 221–6. Seo HS, Ma Y, Debler EW, Wacker D, Kutik S, Blobel G, Hoelz A. 2009. Structural and functional analysis of Nup120p suggests ring formation of the Nup84 complex. Proc Natl Acad Sci U S A 106:14281–6. Siniossoglou S, Lutzmann M, Santos-Rosa H, Leonard K, Mueller S, Aebi U, Hurt E. 2000. Structure and assembly of the Nup84p complex. J Cell Biol 149: 41–54. Stavru F, Hulsmann BB, Spang A, Hartmann E, Cordes VC, Görlich D. 2006. NDC1: a crucial membrane-integral nucleoporin of metazoan nuclear pore complexes. J Cell Biol 173:509–19. Stewart M. 2000. Insights into the molecular mechanism of nuclear trafficking using nuclear transport factor 2 (NTF2). Cell Struct Funct 25:217–25. Strambio-De-Castillia C, Niepel M, Rout MP. 2010. The nuclear pore complex: bridging nuclear transport and gene regulation. Nat Rev Mol Cell Biol 11:490–501. Strawn LA, Shen T, Shulga N, Goldfarb DS, Wente SR. 2004. Minimal nuclear pore complexes define FG repeat domains essential for transport. Nat Cell Biol 6:197–206. Suntharalingam M, Wente SR. 2003. Peering through the pore: nuclear pore complex structure, assembly, and function. Dev Cell 4:775–89. Terry LJ, Wente SR. 2009. Flexible gates: dynamic topologies and functions for FG nucleoporins in nucleocytoplasmic transport. Eukaryot Cell 8:1814–27. Timney BL, Tetenbaum-Novatt J, Agate DS, Williams R, Zhang W, Chait BT, Rout MP. 2006. Simple kinetic relationships and nonspecific competition govern nuclear import rates in vivo. J Cell Biol 175:579–93. Unwin PNT, Milligan RA. 1982. A large particle associated with the perimeter of the nuclear pore complex. J Cell Biol 93:63–75. Van de Vosse DW, Wan Y, Wozniak RW, Aitchison JD. 2011. Role of the nuclear envelope in genome organization and gene expression. WIREs Syst Biol Med 3: 147–66. Vetter IR, Arndt A, Kutay U, Görlich D, Wittinghofer A. 1999. Structural view of the Ran-importin β interaction at 2.3 Å resolution. Cell 97:635–46.

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Watkins JL, Murphy R, Emtage JL, Wente SR. 1998. The human homologue of Saccharomyces cerevisiae Gle1p is required for poly(A)+ RNA export. Proc Natl Acad Sci U S A 95:6779–84. Watson MLJ. 1955. The nuclear envelope: its structure and relation to cytoplasmic membranes. J Biophys Biochem Cytol 1:257–70. Weis K. 2002. Nucleocytoplasmic transport: cargo trafficking across the border. Curr Opin Cell Biol 14: 328–35. Wente SR, Rout MP. 2010. The nuclear pore complex and nuclear transport. Cold Spring Harb Perspect Biol 2:a000562. doi: 10.1101/cshperspect.a000562. Whittle JR, Schwartz TU. 2009. Architectural nucleoporins Nup157/170 and Nup133 are structurally related and descend from a second ancestral element. J Biol Chem 284:28442–52. Winey M, Hoyt MA, Chan C, Goetsch L, Botstein D, Byers B. 1993. NDC1: a nuclear periphery component required for yeast spindle pole body duplication. J Cell Biol 122:743–51. Wischnitzer S. 1958. An electron microscope study of the nuclear envelope of amphibian oocytes. J Ultrastruct Res 1:201–22. Wozniak RW, Blobel G. 1992. The single transmembrane segment of gp210 is sufficient for sorting to the pore membrane domain of the nuclear envelope. J Cell Biol 119:1441–9. Wozniak R, Burke B, Doye V. 2010. Nuclear transport and the mitotic apparatus: an evolving relationship. Cell Mol Life Sci 67:2215–30. Wozniak RW, Blobel G, Rout MP. 1994. POM152 is an integral protein of the pore membrane domain of the yeast nuclear envelope. J Cell Biol 125:31–42. Wozniak RW, Rout MP, Aitchison JD. 1998. Karyopherins and kissing cousins. Trends Cell Biol 8:184–8. Yang Q, Rout MP, Akey CW. 1998. Three-dimensional architecture of the isolated yeast nuclear pore complex: functional and evolutionary considerations. Mol Cell 1:223–34. Zabel U, Doye V, Tekotte H, Wepf R, Grandi P, Hurt EC. 1996. Nic96p is required for nuclear pore formation and functionally interacts with a novel nucleoporin, Nup188p. J Cell Biol 133:1141–52.

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25

CYTOPLASMIC RNA DOMAINS Henry Parker Tom C. Hobman

DEFINITION RNA domains are discrete areas/structures within the cellular cytoplasm where messenger RNAs (mRNAs) and associated molecules are concentrated for the purpose of mRNA processing. Contained in these domains are RNA-binding proteins and those molecules involved in RNA transport, storage, degradation, translation inhibition, and activation, all for the purpose of controlling localization, stability, and translation. The necessity of transducing an environmental signal to changes in either or both transcript and proteome populations requires a more rapid approach than transcriptional control alone. RNA domains provide a quick response through posttranscriptional regulation. Localization of mRNAs is a key component of this speedy reaction to signals.

HISTORICAL BACKGROUND The discovery of spontaneous splicing of RNAs (Kruger et al. 1982) as well as the capacity of such RNAs to catalyze RNA–RNA reactions has bolstered the view of RNA being the primordial molecule that establishes replicative genetic specificity. Moreover, the fact that RNA is the agent of peptide-bond synthesis in the modern ribosome and that modified ribozymes act as aminoacyl esterases, thereby mimicking the activity of aminoacyl tRNA synthetase, establish RNAs as both messenger and ribosome (Piccirilli et al. 1992). Collectively, these findings indicate that RNA has the capacity to catalyze its own replication plus the ordered polymerization of proteins. As cellular functions evolved, so too did the roles of RNA in cellular physiology. DNA, which is more stable than RNA, eventually replaced RNA as the primary genetic material and proteins, which are more efficient promoters of chemical reactions than RNA, now function in most metabolic activities. At the same time, RNA became specialized for carrying genetic messages for protein production and for regulating gene expression at multiple levels. This evolutionary process included co-opting of RNA-binding proteins that served to concentrate the activities of RNA molecules, thereby creating RNA domains with a variety of functions. Localization of mRNAs in eukaryotic cells was first described 45 years ago (Bodian 1965; Koenig 1965), and today, it is generally accepted that virtually all RNAs function in the context of ribonucleoprotein particles or complexes (RNPs) that are distributed throughout the cell nucleus and cytoplasm (see also Chapter 23 on nuclear domains).

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For many years, it has been recognized that RNPs regulate critical functions including mRNA splicing and protein translation in eukaryotic cells. These processes have been studied extensively, and comprehensive reviews can be found in most biochemistry and molecular biology textbooks. More recently, it has become clear that a new class of RNPs that are targeted to specific mRNAs and nuclear loci regulate gene expression on a global level. This phenomenon, which is known as RNA interference (RNAi) (Fire et al. 1998), is thought to affect the expression of more than 50% of all human genes. Argonaute proteins form the cores of these RNPs (Hannon 2002) whose specificities are determined by small interfering RNAs (siRNAs) or microRNAs (miRNAs). These small RNAs direct argonautecontaining RNPs to specific mRNA or loci via complementarity-based targeting.

RNPS AND POSTTRANSCRIPTIONAL GENE REGULATION RNP dynamics play critical roles in determining if, how, and when a particular mRNA is translated or degraded. Nascent mRNA transcripts are first incorporated into nuclear RNPs that contain cap- and RNA-binding proteins. These RNPs facilitate nuclear export and the processing of mRNAs in the cytoplasm. Studies from yeast have shown that the composition of the messenger RNA-protein particles (mRNPs) often reflects the immediate fate of the mRNA. For example, before decapping, an early step in mRNA degradation, occurs, translation initiation factors are replaced by decapping factors, resulting in an mRNP that can no longer initiate translation (Tharun 2008). Moreover, when decapping activators are expressed in decapping null yeast mutants, they can act as translational repressors (Coller and Parker 2005; Pillai et al. 2005). Conversely, remodeling of mRNA-containing RNPs by replacement of decapping complexes with translation factors can allow for rapid translation. After translation, polysomes are disassembled and mRNAs can then be deadenylated and subsequently degraded or stored in RNPs for future rounds of translation. Other newly transcribed mRNAs may be programmed for delayed translation and are either stored locally or transported to a distal location. mRNAs that are translationally idle may become concentrated in one of a number of related cytoplasmic RNA granules for processing (Table 25.1). These cytoplasmic RNPs include processing bodies (P-bodies, also known as GW bodies due to the characteristic Gly-Trp repeat found in the scaffold 182kDa protein), stress granules (SGs), and related structures such as polar bodies (found in germ line cells) and transport granules. Once concentrated in these cytoplasmic domains, an mRNA may be steered into the translation, storage, or decay pathways (Fig. 25.1). There are other RNA granules/domains within the cells, but with the exception of ribosomes, the four discussed in this chapter are the best characterized with respect to RNA metabolism in the cytoplasm. P-bodies and SGs are key RNP domains that share a large cohort of common components (>50%) and are associated with differential processing of mRNAs in response to signals. Polar and neuronal (transport) granules may in fact be specialized versions of P-bodies and SGs in germ line cells and neurons, respectively. A list of known protein components in each RNP is shown in Table 25.2. From this table, it can be seen that the RNPs share common components as well as unique subunits that may provide different functions to each RNA granule.

Transport Granules Many mRNAs are transported to specific intracellular destinations before they are translated (reviewed in Chartrand et al. 2001; Piper and Holt 2004). This provides an important

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TABLE 25.1.

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General Characteristics of Cytoplasmic RNA-Protein Complexes

RNP Processing body

Stress granule* Polar body Transport granule

Organism

Size (nm)

Number (Per Cell)

M D S M S M M D

100–300

3–30 3–8 3–4 >20 1–5 2–10 >20 10–15

100–2000 200–900 150–1000

*Only present in stressed cells. M, mammals; D, D. melanogaster; S, S. cerevisiae.

level of control by restricting the activities of certain proteins to specific regions of the cell. Within mammalian systems, this process is best characterized in neurons, cells that have particularly complex architectures. Most notably, they possess extremely long cytoplasmic projections that require unique transport logistics for proper functioning. Local control of translation in neurons is important for regulating synaptic plasticity as well as for development and growth of axons and dendrites (Hillebrand et al. 2007) (see also Chapter 22). Neuronal transport granules are defined by the presence of two conserved proteins: fragile X mental retardation 1 protein (FMRP) and the double-stranded RNA (dsRNA)-binding protein Staufen (Mazroui et al. 2002; Thomas et al. 2005; Barbee et al. 2006; Kiebler and Bassell 2006). Transport of mRNA involves active targeting through the interaction of cis elements in the RNA and RNA-binding proteins. For example, mammalian brain mRNAs contain 11-nucleotide A2RE11 or 21-nucleotide A2RE sequences that bind to the heterologous nuclear ribonucleoproteins A2 and A3. In turn, these proteins direct the bound mRNAs into cytoplasmic transport granules, which are subsequently delivered to specific sites in the cytoplasm. Movement of the transport granules to distal sites requires microtubules and associated motor proteins such as the kinesin protein KIF5 (Hirokawa 2006; Kiebler and Bassell 2006) or cytoplasmic dynein (Smith 2004). In other cell types, actin and intermediate filaments may be employed instead of microtubules (see Chapters 12, 14, and 16). While en route, mRNAs are translationally inactive, but upon reaching their destination, the mRNAs are either translated immediately or stored and later translated upon release from the inactive state. Translationally inactive mRNAs can be stored in more than one type of RNP/RNA domain (see Table 25.2), not all of which contain ribosomal components (e.g., P-bodies). It has been observed that approximately half of all P-bodies and transport granules transiently interact with each other (Zeitelhofer et al. 2008). This may indicate that the RNAi pathway regulates translation of dendritic mRNAs in response to signaling, for example, by calcium flux.

Polar Granules (P-Granules) P-granules, also known as germinal granules, were first described more than 100 years ago (Hegner 1914) and were later shown to define sites of primordial germ cell differentiation. The germ line is the cell lineage that transmits genetic information between generations of a species. P-granules are found in the cytoplasm of these cells and are enriched in maternal mRNAs required for germ cell specification and establishment/maintenance of polarity (Schisa et al. 2001; Leatherman and Jongens 2003), processes that involve

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AAA

AAA

Figure 25.1. Model depicting known and putative functions of the four major cytoplasmic RNA granules. Processing of miRNA precursors is shown in the nucleus. As well as incorporating nascent mRNAs and miRNAs, some of the RNA granules exchange protein and RNA components.

K

RNPS AND POSTTRANSCRIPTIONAL GENE REGULATION

TABLE 25.2.

433

Asserted Protein Components of RNPs

RNP Type

Function

Protein

P-Bodies

Stress Granules

Polar Bodies

Transport Bodies

Organism

DCAP1,2 DCS2 DDH/RCKp54/DEAD-box 6 DHH1 EDC1,2,3 GE-1/HEDLS PAT1 RAP55 SBP XRN1 CPEB Cup Eap 1 eIF2 eIF2B eIF3 eIF4A eIF4E eIF4E-T eIF4G eRF1,3 hMex3A hMex3B hnRNP A1 hnRNP A3 hnRNP K hnRNP Q Lin28 Nrp1 Pat1 PCBP2 TIA-1/Pub1 TIAR/Ngr1 Calreticulin CIRP CUG-BP1 FAK FXR1P,2P FMRP Grp7 Musashi NAT1/P97 PRTB Pumilio 1

+ + + + + + + + + + + ND + ND ND + ND + + + + + + ND + ND + + + + + + + ND ND ND ND ND + ND ND ND ND ND

– ND + + ND – ND + ND + + ND + + + + + + ND + ND + ND + ND + + + + ND + + + + + + + + + + + + + +

ND ND + + ND ND ND ND ND + + + ND ND ND ND ND + ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND + ND ND ND ND ND

ND ND + + ND ND ND ND ND + + + ND + ND ND ND + ND ND ND ND + ND ND ND + ND ND ND ND ND ND ND ND ND ND + + ND ND ND ND +

C, D, M, S S C, D, M, S C, D, M, S D, M, S D, M D, S M S M, S M D S M M M, S M D, M, S M M, S M, S M M M M M M M S M, S M M, S M, S M M M M M D, M M M M M M

(continued)

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TABLE 25.2.

CYTOPLASMIC RNA DOMAINS

(continued) RNP Type

Function

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Protein

P-Bodies

Stress Granules

Polar Bodies

Transport Bodies

Organism

Pumilio 2 40S ribosome 60S ribosome Sam68 Src3 SGNP Smaug 1 Me31B AIN-1 Argonautes Dicer GW182 mRNA miRNA siRNA TNRC6B Ataxin-2 DDX/Ded1 DDX1 Lsm1-7 Ppb1 NXF7 Rpb1 Rpb4 Staufen Ygr250c p48&p52 BTZ Caprin-1 Cgh-1 GRTH MVH RanBPM RBM42 RHAU YPS APOBEC3F APOBEC3G CCR4-CAF1-NOT complex Hrp1 Importin-8 PABP1/Pab1 Pan2/3 Roquin Vts

ND – – ND ND ND ND + + + ND + + + + + + + ND + + + + + + + ND ND ND + ND ND ND ND ND ND + + + + + + + + +

+ + ND + + + + + ND + + – + + ND ND + + + ND + + ND + + + ND ND ND + ND ND ND + + ND ND + ND + + + ND + ND

ND ND ND ND ND ND ND + + + + + + + ND ND ND ND ND ND ND ND ND ND + ND + + + + + + + ND ND + ND ND ND ND ND ND ND ND ND

ND + + ND ND ND ND + + + + + + + ND ND ND ND ND + ND ND ND ND + ND ND + ND ND ND ND ND ND ND + ND ND ND ND ND + ND ND ND

M M, S M M M M M D C, D, M C, D, M M M M M M M M, S M, S M, S M, S M, S M S S D, M S M, Z D, M M C,M M M M M M D M M M, S S M D, M, S M M S

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TABLE 25.2.

435

(continued) RNP Type

Function

mRNA decay

Assembly structure movement

Apoptosis

Miscellaneous

Protein HuR/D MBNL1 MEX67 TDP-43 SERBP1 ZBP1 MLN51 YB-1 BRF1 FBP&KSRP PMR1 TTP Ebs1 Nam7 SMG5,7 UPF1 UPF2 UPF3 DIS1 Rap55 DIC1/DHC1 KHC/KLC Actin Vimentin Cathepsins CCAR1 FAST Prohibitin 2 RACK1 RSK2 Gemin5 MTR-1 SMN snRNPs AcH2B,3 H2B H4 Rpm2 Tra1 LINE 1 ORF1p COX1 CytoC Enolase F1a,b

P-Bodies

Stress Granules

Polar Bodies

Transport Bodies

Organism

+ ND ND ND ND ND ND + + ND + + + + + + + + ND + ND ND ND ND ND ND + ND ND ND + ND ND ND ND ND ND + ND ND ND ND ND ND

+ + + + + + + + + + + + ND ND ND ND ND ND + + + + ND ND ND + + + + + ND ND + ND ND ND ND ND ND + ND ND ND ND

+ ND ND ND ND ND ND ND ND ND ND ND ND ND ND + ND ND ND ND ND ND + + + ND ND ND ND ND ND + ND + + + + ND + ND + + + +

+ ND ND ND ND + ND ND ND ND ND ND ND ND ND + ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND + ND ND ND ND ND

M M M M M M M M M X M M S S M D, M, S M, S M, S M M, S M M M M M M M M M M M M M M M M M S D M M M M M

(continued)

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TABLE 25.2.

CYTOPLASMIC RNA DOMAINS

(continued) RNP Type

Function

Protein IP5K LDH PHGPx HC-antigen Plakophilin1,3 Htt Hsp27 Hsp70 TDRD3 G3BP TRAF2 AKAP350 E2 Nanos PA700 LAMP1,2 Acid phosphatase LAP DNase NADpase CMPase Ubiquitin

P-Bodies

Stress Granules

Polar Bodies

Transport Bodies

Organism

ND ND ND ND ND + ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND

+ ND ND ND + ND + ND + + + + ND ND ND ND ND ND ND ND ND ND

ND + + + ND ND ND + ND ND ND ND + ND + + + + + + + +

ND ND ND ND ND ND ND ND ND + ND ND ND + ND ND ND ND ND ND ND ND

M M M M M M M M M M M M M D M M M M M M M M

The data for this table were obtained from multiple articles (Anderson and Kedersha 2006, 2008; Barbee et al. 2006; Eulalio et al. 2007; Katahira et al. 2008; Yokota 2008; Buchan and Parker 2009; Moser and Fritzler 2010). In some cases, reports of localization of proteins to specific granules differ among the publications. C, C. elegans; D, D. melanogaster; M, mammals; S, S. cerevisiae; X, X. laevis; Z, zebrafish; ND, not detected.

epigenetic reprogramming and genetic recombination. In addition to maternal mRNAs, P-granules contain proteins that are involved in mRNA translation and decay (Table 25.2). These include CAR-1, an Sm protein related to Lsm proteins that regulate splicing, decapping, and decay (Audhya et al. 2005; Boag et al. 2005); CGH-1, an RNA helicase that is related to Dhh1 and p54/Rck (enzymes associated with translational silencing and decapping) (Navarro and Blackwell 2005); and the mRNA decapping enzyme DCP1 that removes the 5′cap structures of mRNAs. The latter is an important step in the regulated turnover of mRNAs (Lall et al. 2005). In addition to mRNA decay enzymes, P-granules may also contain translation initiation factors such as eIF4E (Amiri et al. 2001). In Drosophila melanogaster, Caenorhabditis elegans, and Xenopus laevis, P-granules are asymmetrically distributed among prospective germ cells. During early embryogenesis, they direct the timing of maternal mRNA translation in order to establish the progeny germ line (Schisa et al. 2001; Leatherman and Jongens 2003). Conversely, in mammalian cells, P-granules are only observed in later stages of germ cell differentiation (spermatogenesis and oogenesis) (Chuma et al. 2009). Accordingly, even though the molecular compositions of these RNPs are highly conserved, their functions may differ according to species-specific differential developmental programs.

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Chromatoid bodies (CBs) are a subset of P-granules found in mammalian male germ line cells. They were first described in early rat spermatids more than 100 years ago by von Brunn (von Brunn 1876). Whereas P-granules direct spatial and temporal expression of maternal mRNAs in germ cells, CBs direct expression of paternal mRNAs in spermatogenic cells. Electron microscopic studies revealed that these RNPs are composed of irregular aggregations of dense fibrillar material. They exhibit nonrandom movements and occasionally make transient contact with the Golgi membranes (Parvinen and Jokelainen 1974), but do not have membranes of their own. The number of CBs per germ cell varies according to the developmental stage. CBs are thought to transport mRNAs to specific locations in the cytoplasm (Parvinen 2005). Their role in regulating gene expression is likely dependent on the RNAi pathway as evidenced by the observation that the argonaute family member Miwi is a component of these RNPs. Movement of CBs likely depends on the Miwi-binding protein KIF17b, a testis-specific kinesin that migrates between nuclear and cytoplasmic compartments (Kotaja et al. 2006). Other RNAi pathway components such as the RNase Dicer and the DEAD-box helicase mouse VASA homolog (MVH) are also found in CBs. The picture that emerges from these combined studies is that nuclear haploid gene products are assembled into RNPs, transported through nuclear pores via the motor protein KIF17b and then targeted to CBs where they may be translated, stored for specific temporal and spatial translation, or degraded (see also Chapter 24 on the nuclear pore).

SGs In 1989, Nover et al. observed that tomato cell cultures stressed by heat concentrated a specific subset of mRNAs in cytoplasmic RNPs, which they called heat shock granules (Nover et al. 1989). Later, these RNPs would become known as SGs. We now know that SGs form as a protective mechanism against diverse stresses such as exposure to high temperatures, toxins, unfolded proteins, ischemia, or viral infection (Buchan and Parker 2009). Formation of these RNPs may serve, among other things, to silence the translation of housekeeping mRNAs, thereby allowing the cell to conserve energy for the repair/ recovery from molecular damage caused by the stress. Unlike housekeeping mRNAs, the mRNAs encoding stress-induced factors such as heat shock proteins are excluded from SGs (Anderson and Kedersha 2009). SG formation appears to be highly coordinated as it involves recruitment of proteins and nucleic acids from multiple cellular compartments. Data from a large-scale RNAi screen revealed that SG biogenesis is dependent on more than 100 human genes (Ohn et al. 2008). Cellular stress can result in the phosphorylation of eIF2α, a component of the eIF2 complex, which is responsible for loading the initiator tRNA (Met-tRNAMet) onto the 40S ribosomal subunit. This phosphorylation event can lead to global inhibition of protein synthesis (Berlanga et al. 1998; Gray and Wickens 1998; Srivastava et al. 1998). Following eIF2α phosphorylation, a number of key RNA-binding proteins, T cell intracellular antigen (TIA)-1, TIA-related protein (TIAR), and poly(A)+-binding protein I (PABP-I) together with poly(A)+ RNA, are concentrated in nascent SGs (Kedersha et al. 1999). Normally, PABP-I resides in the cytoplasm where it promotes the stability and translation of mRNAs (Adam et al. 1986; Matunis et al. 1993). In contrast, TIA-1 and TIAR are concentrated in the nucleus at steady state but are known to shuttle between the nucleus and the cytoplasm. TIA-1 and TIAR act downstream of the stress-induced phosphorylation of eIF2α to promote the recruitment of untranslated mRNAs to SGs. Current opinion holds that SGs function in translational repression of mRNAs, and this view is supported by the observation that formation of SGs coincides with a decrease

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in overall translation (Anderson and Kedersha 2009). However, it is important to point out that global reduction in translation does not require SG formation per se (Buchan et al. 2008; Fujimura et al. 2009; Loschi et al. 2009), and as such, SGs may form as a consequence of blocking protein translation. In this regard, SGs could act as mRNA triage sites during/following cellular stress. Indeed, certain mRNAs are stabilized by SGs. Moreover, cellular stress can inhibit deadenylation, an early step in mRNA degradation pathways (Laroia et al. 1999; Hilgers et al. 2006). Nevertheless, deadenylation does not occur when mRNAs are trapped in polysomes of stressed cells; in which case, SGs are not formed. While SGs most certainly play critical roles in downregulating protein synthesis on a global scale, evidence suggests that when translation resources are limiting, SGs can actually promote assembly of translation initiation complexes. For example, mRNAs that contain internal ribosome entry sites are preferentially translated during stress conditions (Spriggs et al. 2008). This phenomenon may involve SG components such as hnRNP A1 and PCBP2, which bind to internal ribosome entry sites and promote translation (Bonnal et al. 2005). In addition to proteins that regulate RNA metabolism, SG components include several apoptotic regulators such as CCAR1, Prohibitin2, RSK2, and RACK1 (Buchan and Parker 2009). This suggests that SGs have additional functions including regulating signaling pathways that control apoptosis and inflammation. The fact that cell survival decreases when SG formation is inhibited during stress is consistent with this theory (Eisinger-Mathason et al. 2008). There are multiple mechanisms by which SGs can modulate apoptotic signaling. One involves blocking the pro-apoptotic kinase MTK1. The activity of this kinase is enhanced by the SG component RACK1, and in response to some types of stress, sequestration of RACK1 in SGs may limit activation of MTK1, therefore inhibiting apoptosis (Arimoto et al. 2008). Another mechanism involves interaction with the pro-apoptotic SG resident TIA-1. This protein blocks the anti-apoptotic action of Fas-activated serine/threonine phosphoprotein (FAST) (an inhibitor of caspase-3 activation) but can be prevented from doing so by complex formation with RSK2 and SGs (Eisinger-Mathason et al. 2008). Finally, it is worth discussing the mechanistic link between SGs and the antiinflammatory action of cellular stress (Kim et al. 2005). During stress, mammalian cells have been observed to sequester the signaling protein TRAF2 into SGs. Under normal conditions TRAF2 interacts weakly with eIF4GI, but during/following heat stress, this translation initiation factor is recruited to SGs where it interacts strongly with TRAF2. This in turn, reduces nuclear factor κ-light-chain-enhancer of activated B cells (NF-κB) activation by affecting tumor necrosis factor signaling, a process that is dependent on TRAF2.

P-Bodies Localization of mRNA decay enzymes to discrete cytoplasmic foci, which later became known as P-bodies, was first reported more than 10 years ago. The first reported P-bodyassociated RNA-degrading enzyme is XRN1, a 5′ → 3′ exoribonuclease (Bashkirov et al. 1997). Later, it was reported that the decapping enzyme DCP2 and associated cofactors colocalized with XRN1 in the same cytoplasmic foci (Ingelfinger et al. 2002; LykkeAndersen 2002; van Dijk et al. 2002; Sheth and Parker 2003; Cougot et al. 2004). In parallel studies, Eystathioy et al. described cytoplasmic foci that reacted with autoimmune serum from a patient presenting with motor and sensory neuropathy (Eystathioy et al. 2002). The reactive antigen was a large RNA-binding protein, GW182, named for the numerous glycine, tryptophan (GW) repeats. GW182 is required for P-body formation and

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is now known to play an important role in blocking translation of mRNAs (Yang et al. 2004). These foci, which at the time were called GW bodies, were later found to contain the P-body residents and mRNA-degrading enzymes XRN1 and DCP2. Subsequent studies revealed that P/GW-bodies contain other mRNA-degrading enzymes including the decapping enzyme GE-1 and the coactivator LSm14 (Albrecht and Lengauer 2004; Anantharaman and Aravind 2004; Fenger-Gron et al. 2005; Yu et al. 2005; Bloch et al. 2006; Yang et al. 2006), thus firmly establishing these RNPs as sites of mRNA degradation. A link between P-bodies and RNAi surfaced when it was discovered that pools of argonaute proteins localize to these structures and to SGs as well. Argonautes are RNA-binding proteins that use small guides (miRNAs and siRNAs) for targeting to P-bodies and SGs in mammalian cells (Liu et al. 2005; Sen and Blau 2005; Leung et al. 2006; Pauley et al. 2006). P-bodies are found in the cytoplasm of most eukaryotic cells. These RNPs are highly dynamic in that their size, number, and even composition vary with cell type, rate of proliferation, and phase of the cell cycle (Yang et al. 2004; Moser et al. 2007). Furthermore, there are significant differences between P-bodies in lower and higher eukaryotes. For example, argonaute proteins, LSm14 and GE-1 homologs are not associated with P-bodies of the budding yeast, Saccharomyces cerevisiae. Similarly, the genome of this organism does not encode argonaute proteins, which are an important component of mammalian P-bodies. Mammalian P-bodies are also larger (100–300 nm in diameter) than their yeast counterparts (Table 25.1). Proteomic studies by several laboratories have revealed much about the composition of P-bodies (Table 25.2 lists the components of P-bodies from different organisms). The data suggest that in addition to mRNA degradation, P-bodies function in mRNA triage, storage, and RNAi and translational repression. Compared with SGs, which are largely immobile, P-bodies are actively transported throughout the cytoplasm and the nuclear periphery using microtubule-based motors (Carmichael et al. 2006; Aizer et al. 2008) (see also Chapter 14 on microtubules). Bidirectional movement along microtubules may facilitate interactions with mRNA transcripts that are destined for degradation by RNAi or may reflect coupling of the mRNA transport and silencing machineries. As a mechanism to coordinate temporal and spatial regulation of gene expression, a subset of mRNAs is actively transported in the context of RNPs via microtubules to specific intracellular locations. These mRNAs are often translationally repressed until they arrive at their final destination. In this regard, the fact that RNAi effector complexes and some mRNAs use microtubules for transport is unlikely to be a coincidence. Rather, it may reflect the underlying mechanism by which a subset of nascent mRNAs that emerge from nuclear pores is silenced until they reach their sites of action. In addition, mobility of the RNAi apparatus may be important for efficient surveillance of the cytoplasm for mRNAs that are destined for degradation or repression. In mammalian cells, P-body formation is linked to the miRNA pathway (Pauley et al. 2006) and activity of argonaute proteins (Pare et al. 2009). With respect to recruitment of proteins to P-bodies, there appears to be a specific order by which this occurs. In general, Q/N-rich elements within proteins seem to foster P-body aggregation (Decker et al. 2007; Mazzoni et al. 2007; Reijns et al. 2008). Studies in budding yeast revealed that accumulation of Lsm1-7 or Dcp1 in P-bodies requires Pat1 and Dcp2, respectively (Teixeira and Parker 2007). While there has been considerable progress, the mechanisms governing P-body formation are not completely understood. This is perhaps not surprising as no less than 39 genes are required for biogenesis of P-bodies in human cells (Ohn et al. 2008). P-body assembly is also linked to the presence of nontranslating mRNA (Pillai et al. 2005; Teixeira et al. 2005). The rate and extent of P-body formation is directly proportional to the amount of available inactive mRNA, but not all nontranslating mRNA is found in P-bodies. Moreover, simply dissociating mRNA from polysomes is not sufficient

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to form P-bodies. Rather, the mRNA must contain elements that attract P-body components (Franks and Lykke-Andersen 2007). For example, the P-body residents tristetraprolin (TTP) and butyrate response factor 1 (BRF) can localize adenine-uracil (AU)-rich mRNA into P-bodies. A question that still remains is whether P-bodies form around specific mRNAs and associated proteins or are they targeted to preexisting mRNP structures that contain similar mRNAs and proteins that shared between various cytoplasmic RNPs. Although P-bodies are enriched in enzymes that mediate mRNA destruction, they are not necessarily a dead end for mRNAs because at least some P-body-associated mRNAs can return to the translational pool (Brengues et al. 2005; Bhattacharyya et al. 2006). P-bodies and SGs have been shown to physically interact, suggesting that components can be exchanged between these RNPs (Wilczynska et al. 2005; Buchan et al. 2008). In yeast, assembly of SGs is promoted by the presence of preexisting P-bodies. One model contends that cytoplasmic mRNAs cycle among polysomes, P-bodies, and SGs (Balagopal and Parker 2009). This model is supported by the observation that inhibition of translation initiation causes the loss of mRNAs from polysomes and a corresponding increase in the amount of mRNA that associates with P-bodies and SGs. However, if translation elongation is blocked, the mRNAs remain preferentially associated with polysomes as opposed to P-bodies and SGs. The precise role of P-bodies in mRNA decay and posttranscriptional gene silencing is a matter of some controversy. For example, although decapping enzymes are concentrated in these RNPs, decapping of mRNA in yeast does not require formation of P-bodies. Similarly, translational repression and stabilization of mRNA during stress and RNAi occurs in the absence of microscopically visible P-bodies (Chu and Rana 2006; Decker et al. 2007; Kwon et al. 2007; Ohn et al. 2008). However, because they have been conserved throughout evolution, P-bodies must serve an important role in RNA metabolism. One theory is that P-bodies serve to concentrate factors involved in mRNA decay and RNA silencing, thereby making the processes more efficient. Alternatively, concentration of decay factors in P-bodies may serve to lower their activities in the bulk cytoplasm. Finally, it is worth noting the potential role of P-body components in human disease. Recently, it was reported that a pool of the Huntingtin protein (Htt, Huntington’s disease protein) localizes to P-bodies where it associates with the RNAi effector protein Ago2 (Savas et al. 2008). Interestingly, the Huntington-disease-associated expansion of the polyQ expansion in Htt, results in reduced binding to Ago2 and a reduction in P-body numbers and RNAi. Subsequent data from this laboratory demonstrated that Htt plays a role in silencing of transport-granule-associated mRNAs (Savas et al. 2010). Together, these data suggest that altered translational regulation may contribute to pathology in one or more serious human diseases.

P-Bodies and SGs: Close Cousins? While all of the cytoplasmic RNPs discussed in this chapter share some common components, P-bodies and SGs appear to be much more closely related in terms of composition (slightly more than half of SG components are also found in P-bodies) and function. This is particularly evident for translational regulation. Despite the obvious similarities, recent studies indicate that these RNPs are quite different from each other with respect to morphology and biogenesis. Electron microscopic analyses indicate that SGs are loosely organized fibrillo-granular aggregates of moderate electron density, whereas P-bodies are more dense and fibrillar (Souquere et al. 2009). As mentioned above, SGs dynamically interact with P-bodies, which led to the suggestion that mRNAs are transferred from SGs to PBs for degradation (Kedersha et al. 2005; Wilczynska et al. 2005). Despite the frequent interaction between these two types of RNPs, they remain structurally and compositionally

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distinct (Souquere et al. 2009). Initially, it was believed that protein components are exchanged between SGs and P-bodies; however, it was later shown that the SG components came from the cytosol and not from adjacent P-bodies (Mollet et al. 2008). In some organism though, SG assembly is dependent on P-body formation (Buchan et al. 2008). The studies described above are consistent with the notion that biogenesis of P-bodies and SGs are governed by different mechanisms. At least 31 common genes coordinate assembly of both types of RNPs, while over 80 additional unique gene products are specific to P-body or SG assembly (Ohn et al. 2008). More recent work has shed light on the factors involved in these processes. For example, Hsp90 activity is essential for assembly and/or maintenance of P-bodies, whereas nascent SG formation does not require this molecular chaperone (Pare et al. 2009). Studies from multiple laboratories suggest that biogenesis and disassembly of both SGs and P-bodies is regulated by microtubules (Ivanov et al. 2003; Carmichael et al. 2006; Sweet et al. 2007; Nadezhdina et al. 2009). A number of microtubule-based motors have been linked to homeostasis of these RNPs in yeast and mammalian systems (Loschi et al. 2009; Tsai et al. 2009; Stoica et al. 2010). Importantly, evidence from these studies suggests that these microtubule-based motor proteins function in posttranscriptional gene silencing.

Future Perspectives The studies discussed above review how a family of interrelated RNPs functions to regulate mRNA transport, storage, expression, and decay in the cytoplasm of eukaryotic cells. Based on their shared constituents, it is tempting to speculate that these complexes perform a number of common functions. One of the major challenges that lie ahead is to understand how the functions of these RNPs are coordinated to regulate gene expression on a global level. This is particularly important given the current race to develop RNA-based therapies (i.e., RNAi) for treatment of genetic and infectious diseases.

ABBREVIATIONS BRF-1 CBs DEAD dsRNA FAST GW miRNA mRNA mRNP MVH

butyrate response factor 1 chromatoid bodies asp-glu-ala-asp double-stranded RNA Fas-activated serine/threonine phosphoprotein earlier name for P-bodies; based on Gly-Trp-repeat scaffold protein microRNA messenger RNA messenger RNA-protein particle Mouse VASA homolog

NF-κB

Nuclear factor kappa-light-chain-enhancer of activated B cells P-bodies processing bodies PBs GW bodies P-granules polar granules RNAi RNA interference RNP ribonucleoprotein particle SGs stress granules siRNA small interfering RNA TIA-1 T cell intracellular antigen-1 TIAR T cell intracellular antigen-related protein TTP tristetraprolin

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Index

abscission, 159 acantholysis, 328 ACBD3, 141 acetylation, 241 actin, 198, 267. See also filamentous actin at adherens junction, 314 improper regulation of, 197 microfilaments, 225 at microvillus tip, 216 in phagocytosis, 171 in polarized cells, 205–206 polymerization of, 16, 214, 216–217 in regulation of CCP formation, 16 actin cytoskeleton, 195, 209 biochemical regulation of, 202–203 components of, 198, 199–200 definition of, 197–198 historical perspectives on, 198–199 kinase and phosphoprotein components regulating, 203–204, 204 mechanosensing function of, 203 membrane, 198, 198 and neuritogenesis, 206–207 plasticity of, 202–204 and polymerization and branching, 200–202, 201, 202 proteome of, 208, 209 signaling studies of, 207–209 actin fences, and plasma membranes, 18 actin skeleton mesh, size distribution of, 8 activating transcription factor 6 (ATF6), 123 acyl-anchored proteins, in lipid rafts, 65 adaptor complexes AP-2, 27, 28, 29 monomeric adaptors, 29–30

adaptor protein, phosphotyrosine interaction, PH domain, and leucine zipper containing 1 (APPL1), 157, 158 adenosine diphosphate (ADP), hydrolyzing of ATP to, 200 adenosine-5’-diphosphate (ADP), 97 adenosine-5’-triphosphate (ATP), 87 adherens junction (AJ) cytoplasmic machinery of, 306–309 cytoskeleton regulation of, 314–315 definition of, 303 dynamic organization of, 315 and epithelial cell polarity, 311–312 historical perspective on, 303–305 maintenance and remodeling, 313 metazoan development and organization through, 311 peripheral membrane components of, 305, 306–309, 310–311 protein-protein interactions of, 305 protein trafficking and polarity at, 312–313 adhesion. See also extracellular matrix; focal adhesions, 286 imaging dynamics of, 297 adhesion plaque, 285, 287 adipocytes, caveolae in, 41 afadin, 310 Afipia felis, phagocytosis of, 173 agriculture, role of microtubules in, 240 α-catenin, 310 α-subunit, in basolateral localization of Na+ pump, 359 altered-in-glycosylation (ALG) genes, 136 alternative reading frame (ARF), 398

Alzheimer’s disease (AD), 91, 117, 117, 386 AMOG, 360 AMPA, 380 amphiphysin, 31 amyotrophic lateral sclerosis (ALS), 387 anaerobic mitochondria, 89 anaphase, microtubules during, 239 anchored-protein picket model, 5, 6, 7 anchored-protein pickets, 18 anemia, 12 ankyrins, 115 ankyrin-spectrin cytoskeleton, retention at PM by, 359–360 antiinflammatory action, of cellular stress, 438 AP180, 30 apoptosis conventional, 275 Fas-induced, 276 argonaute proteins, 430, 439 armadillo family members, of desmosome, 330–331 Arp2/3 complex, 12, 200–201, 202 arrhythmogenic right ventricular cardiomyopathy (ARVC), and desmosomal alteration, 332 artificial membranes, diffusion coefficients for, 9–10 assembly polypeptides (APs), 24 associated proteins (APs), 24 ataxia, and human mitochondrial genome, 91 atherosclerosis, 116, 117 atomic force microscopy (AFM), of membrane organization, 72 autism, 386 autocrine motility factor receptor (AMFR)/GP78, 188 autosomal recessive hypercholesterolemia (ARH), 30 avalanche photodiodes (APDs), 80 axo-dendritic polarization, 384

Cellular Domains, First Edition. Edited by Ivan R. Nabi. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc.

445

bindex.indd 445

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446

INDEX

axonal domains, 373, 376 axon initial segment, 376, 376–377, 378 internode, 378 juxtaparanode, 378 nodes of Ranvier, 378 paranode, 378 presynapse, 376–377, 378 synaptic cleft, 379 axonal trafficking disorders, 387 axonal transport, 237 axoneme, of cilia, 245 molecular structure of, 255 proteins related to, 250–251 axon guidance, 385 axon initial segment (AIS), 376, 376–377, 378 axons, 371 cytoskeletal specializations of, 383 molecular composition of, 373, 376 Bardet-Beidl syndrome (BBS), 258 Barth syndrome, 95 basal body (BB), ciliary, 245, 246, 247 centrioles of, 253–254 proteins related to, 249 released from TZ, 261 structure of, 253 basket neuron, 374 B-cell receptor (BCR) in membrane domain studies, 76 signal transduction, 12–13 B-cells receptor microclusters in, 72 role of membrane rafts in, 75 β-arrestins, 28, 29–30 β-catenin, 305, 310 β1-integrin, 275 β-subunit, in sorting of Na,KATPase, 360–361 bin/amphiphysin/rvs (BAR), 31–32, 149 bin/amphiphysin/rvs (BAR) domaincontaining proteins, 155 bone morphogenic proteins (BMPs), 385 breast cancer, 117 EGFR in, 139 signaling pathway related to, 17 breast tumor kinase/Src-related intestinal kinase (BRK/SIK), 405 Brownian diffusion, in plasma membrane of PtK2 cell, 11 Brucella abortus, phagosomelysosome fusion inhibited by, 171

bindex.indd 446

brush border, of microvilli, 213, 214 discovery and identification of, 214–215 dynamics of, 221–224, 222 ultrastructural organization of, 215 cadherin, 303, 315 desmosomal, 329 in different cell types, 304 naming of, 304 tissue expression of, 331 cadherin adhesion protein, 304 Caenorhabditis elegans, P-granules in, 436 cajal bodies, in situ structure of, 396 calcium in cell-cell adhesion, 304 at ERJs, 179 calcium stress, and microvillar morphology adaptation, 222 calnexin, 119, 126 calpain, 296 calreticulin, 119, 121–122 cancer, 117, 117, 139 desmosomes and, 332 and ErbB2, 206 impaired function of ER in, 126 and microtubule function, 240 and nucleolar size, 397 role of caveolae in, 51 SNBs implicated in, 406 tight junction altered in, 327 cannabinoid type 1 receptor (CBIR), 167 carboxyl terminal domain (CTD), 403 cardiolipin, 94 catastrophe, in microtubule dynamics, 233 Cav1, 43–44, 45 Cav2, 44, 45 Cav3, 44, 45 caveolae biogenesis of, 45, 47, 49 cellular functions of, 49–51 definition of, 39 distribution of, 40–41, 41, 42, 43 endocytosis of, 49–50 and formation of invaginations, 44–45 historical perspective on, 39–40 lipids in, 46 molecular components of, 43–46 morphology of, 40–41, 41, 42, 43 naming, 40 neck region of, 47 physiological roles of, 51

role in disease of, 40, 51 structure of, 51 and transducing mechanical stimuli, 50–51 caveolar coat, 42, 43 cavins, 45–46 cell-cell junctions, electron microscopy studies of, 304 cell polarity, definition of, 349 cell signaling. See also signaling intermediate filaments in, 268 role of membrane rafts in, 75 central nervous system (CNS), perineuronal nets in, 382 centrosome protein 55 (CEP55), 159 centrosomes, 238 ceramide transfer protein (CERT), 185 cerebellar purkinje neuron, 374 cerebrospinal fluid (CSF), and ciliary function, 260 channels, in ERJs, 178 chaperone-mediated autophagy (CMA), 170 Charcot-Marie-Tooth neuropathy, 98, 117, 117 Chlamydomonas ciliary functions in, 259 ciliary motility in, 261 for studying cilia, 247 Chlamydomonas flagella, TEM of, 257 cholesterol of Golgi membranes, 135 ordering effect of, 10 in TCR membrane domains, 66 cholesterol/sphingolipid-rich microdomains, raft hypothesis of, 137 choroid plexus (CP) cell polarity in, 351 epithelial cell polarity of, 354, 355 chromatin, organization of, 393–395 chromatoid bodies (CBs), 437 chromosome territory (CT), 395 ch-TOG gene, 241 cilia definition for, 245 functional implications of cell proliferation and differentiation, 261 motility, 258–261 sensory antennae, 256, 258, 259 historical perspective on, 245, 247 molecular composition of, 247, 248, 248, 249–253 basal body, 253–254

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INDEX

ciliary tip, 255 cilium, 254–255 determination of, 248 machinery of ciliary assembly, 256, 257 proteins, 249–253 transition zone, 254 signal transduction in, 259 structure of, 245, 246–247 ciliary membrane, 245 ciliary tip basic structure of, 255 and IFT, 257 cisternal maturation model, 137 clathrin, 24, 27, 156, 358, 364, 420 clathrin-associated sorting proteins (CLASPs), 29, 33 clathrin-coated pits (CCPs), 8, 8 assembly, 34 cargo-dedicated, 34, 35 cargo incorporated by, 33–34 constructing, 32–33 deep-etch cryo-electron microscopy of, 32 definition of, 23 functions of, 23 generating membrane curvature for, 25, 33 historical perspective on, 23–24, 25, 26 microscopic observation of, 25 molecular components of, 25, 26 adaptor complexes, 27, 28, 29–30 clathrin, 27 regulatory proteins and lipids, 30–31 structural proteins, 31–32 clathrin-coated plaques, 34, 35 clathrin-coated vesicles (CCVs), 148 clathrin-independent carrier (CLIC), 148 clathrin triskelion, 25, 27, 28, 33 claudin family of proteins, 325 cloning of cDNAs for clathrin and adaptors, 26 DNA, 358 of PTRF, 45 coat protein complex (COP), 114 coilin protein, 401 coincidence detection, in construction of CCPs, 32 colchicine antimitotic effects of, 231 in microtubule biology, 230 compartmentalization, membrane skeleton-based, 17

bindex.indd 447

congenital disorders of glycosylation type 1 (CDG), 136 connexins, 339 functions of, 345 in gap junction domains, 341, 342 life cycle of, 340 molecular combinations of, 342 turnover time for, 342 COPII, 420 cortactin, 12 cortical actin rim, 198, 199 cortical ER (cER), in yeast, 182–183 Coxsackie adenovirus receptor (CAR), 365 CREB-binding protein (CBP), 399 Creutzfeldt-Jakob disease, 117 cristae apoptotic remodeling of, 98 formation of, 97–98 organization of, 95 crista junction (CJ) during apoptosis, 98 formation of, 97–98 organization of, 95 crista membrane (CM), 87 Crumbs complex, 312 cryo-electon microscopy, structural studies by, 26 cuchromatin, 394 cyan fluorescent protein (CFP), in membrane domain studies, 76 cystic fibrosis, 117, 117, 125–126 cystic kidney disease, and sensory cilia dysfunction, 258 cytokinesis, 159 cytoplasmic linker proteins (CLIPs), 115 cytoplasmic/scaffolding proteins, of tight junction, 326 cytoskeletal specializations, 383 cytoskeleton AJ regulation of, 314–315 components of, 195 function of, 195 influence on plasma membrane of, 3 and neuronal domains, 381–382, 382 cytoskeleton-linking membrane proteins (CLIMPs), 115 cytoskeleton-linking protein (CLIMP63), 238 deafness, and human mitochondrial genome, 91 death-inducing signaling complex (DISC), 275 dendrite guidance, 385 dendrites, 371, 379

447

cytoskeletal specializations of, 383 disorders of, 386 dendritic domains dendritic spine, 380–381 postsynapse, 379–380 and synaptic integration, 381 dense fibrillar component (DFC), 396–397 depolymerization, in regulation of CCP formation, 16 desmoglein terminal domain (DTD), of desmogleins, 330 desmosomal plaque, major regions of, 328 desmosomes. See also hemidesmosomes alterations in function of, 331–332 constituent protein iosforms of, 331 definition of, 327 discovery of, 322, 327–328 dysfunction of, 329 as intercellular adhesive cement, 328–329 keratin IFs and, 273, 274 structure of, 329–331 detergent-resistant membranes (DRMs), 167, 220 and alteration of phagosomal functions, 172 enrichment of, 62, 62 quantitative proteomic analysis of, 62 diabetes mellitus, and human mitochondrial genome, 91 digestive process, intestinal microvilli in, 221 1,2-dioleoyl-sn-glycero-3phosphoethanolamine (DOPE), SPT experiments with, 78–79 diphosphatidylglycerol, 94 disabled-2 (Dab2), 30 diseases, human, 406 associated with connexins, 345 and desmosome function, 331–332 endoplasmic reticulum-related, 125 IF-pathies, 269 and keratin gene mutations, 276–277 mitochondrial, 100–101 and nuclear splicing speckles, 403 pemphigus, 328–329 role of caveolae in, 51 and sensory dysfunction of cilia, 258 and tight junction proteins, 326

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448

INDEX

DM protein kinase (DMPK), 403–404 DNA mitochondrial (mtDNA), 88, 90 and role of RNA, 429 DNA cloning, 358 domain formation, 18 double-stranded RNA (dsRNA)binding protein Staufen, 431 Down’s syndrome, 386 Drosophila melanogaster, P-granules in, 436 Drosophilia studies of adherens junctions, 312, 313 of desmosomes, 330–331 dsRNA-activated protein kinase-like endoplasmic reticulum kinase (PERK), 123 dynamin, 31 dyneins, in ciliary motility, 260–261 early endosomal antigen 1 (EEA-1), 153 early endosomes (EEs), 148 EB1, 255 E-cadherin, 313 Edman degradation, 167 EGF receptor (EGFR), surface halflife of, 139 electron microscopy (EM), 81 ER identified with, 114 introduction of, 39 of membrane organization, 72 for microtubule research, 231 of polarized epithelial tissues, 322 electron spectroscopic imaging, 406 electron tomography, of plasma membrane, 7–8, 8 embryonic development desmosomal proteins in, 332 dynamic processes during, 225 enabled/vasodilator-stimulated phosphoprotein (ENA/VASP), 207 endocytic proteins, and CCP construction, 35 endocytic system, 148 endocytosis, 166 of cadherin-nectin complex, 315 of caveolae, 49–50 CCP-mediated, 15 clathrin-mediated, 23 lipid rafts in clathrin-mediated, 67 and protein trafficking, 313 endophilin, 31 endoplasmic-reticulum-associated degradation (ERAD), 113, 116, 118, 119, 120

bindex.indd 448

endoplasmic reticulum degradationenhancing 1,2-mannosidase-like protein (EDEM), 120 endoplasmic reticulum (ER), 85, 195 and Ca2+ homeostasis, 121, 126 Ca2+ buffers, 120–122 Ca2+ transport, 121, 122 and cis-Golgi, 133 composition of, 113 definition of, 113 diseases associated with breakdown in, 116–117, 117 dynamics of, 114 and ERAD, 120 folding within, 117–118, 119 functions of, 113 historical perspectives on, 113–114 impaired function of, 125 multifunctions of, 115–117, 126 PM proteins synthesized in, 357, 357 and protein synthesis, 118 proteomic analysis of, 115, 116 quality control, 118, 119 resident oxidoreductases in, 122 selected proteins identified from, 116 stress and UPR, 123–125, 124 structure of, 114–115 endoplasmic reticulum (ER) junctions, 85 endoplasmic reticulum (ER) membrane, 415 endoplasmic reticulum (ER) protein deficiency, mouse models of, 125–126 endosomal membranes, 155–157, 160 endosomal network endocytic system, 148–149, 149 and formation of MVBs, 151 historical considerations, 147–148 membrane retrieval routes, 149, 149–151, 150 endosomal sorting complex required for transport (ESCRT), 151, 152, 157, 159, 161 endosomal system dynamics of, 150, 151, 152, 153 organization of, 160–161 Rab domains as functional units of, 153–155 spatiotemporal dynamics of, 150 endosomes, 85, 195 biochemical characterization of, 147 and cellular homeostasis, 158–159

composition of, 147 definition of, 147 functional domains on, 152 geometry of, 148 and membrane remodeling, 159 multiple populations of, 161 and shaping tissues, 160 signaling machinery of, 157–158 endosymbiotic theory, of mitochondria origin, 88 endothelia cells, caveolae in, 41 enteropathogenic Escherichia coli (EPEC), 223–224 enteropathogenic Escherichia coli (EPEC) infection, 222 enzymes in ERJs, 178 on intestinal microvilli, 220 epidermal growth factor (EGF), 199–200 epidermal growth factor (EGF) receptor (EGFR), coordination of functions of, 13–14 epidermolysis bullosa simplex (EBS), 276 epithelia cells, caveolae in, 41 epithelial cell asymmetry, and keratin IFs, 276 epithelial cells, 351 AJ of, 303 and apical-basolateral localization, 356 and apical-basolateral polarization, 351–355, 353, 354 PM proteins in, 352 polarized organization of, 325 targeting mechanisms of, 353, 355 tight junction strands, 322 variations in sorting machinery of, 364–365 epithelial-mesenchymal transition (EMT), 160, 278 epithelial polarity, 349, 356, 357 epithelial protein lost in neoplasm (EPLIN), 314 epithelial tissues permeability of, 332 “spot welds” in, 327 epsin/Eps15 N-terminal homology (ENTH) domains, 30 ER-associated degradation (ERAD) pathway, 340 ER-Golgi intermediate compartment (ERGIC), 137 ER junctions (ERJs) components of, 178 definition of, 177 fine organization of, 189

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INDEX

formation of, 181 historical perspective on, 177 location of, 177 molecular composition of Golgi-ERJ, 184–186 mito-ERJ, 186–188 NVJ, 179–181, 180–181 PM-ERJ, 182–184 structural components of, 180–181 unresolved issues for, 188–189 ER-mitochondria encounter structure (ERMES), 181, 187–188 erythrocytes, membrane skeleton organization in, 12 espin, 217, 218, 218 exocytosis of cadherin-nectin complex, 315 and protein trafficking, 313 extracellular matrix (ECM), in developing nervous system, 382 extracellular matrix (ECM) adhesion, and Rho activity, 288 extracellular matrix (ECM) molecules, 285 ezrin, 218, 220–221 ezrin-radixin-moesin (ERM) family, 213 F-actin cytoskeleton, 197, 198, 201 F-actin cytoskeleton interactome, 201 F-actin modulators, 209 F-actin phosphorylation interactome, 209 familial hypercholesterolemia, 117 F1AsH/ReAsH pulse-chase study, 343 FcεRI, 14 fence-picket model, 5, 6, 9, 78 fenestrated diaphragms (FDs), 47, 48 FERM domain, 294 FG-Nups, 421–422, 423 fibrillar centers (FCs), 396–397 filamentous actin (F-actin), 32, 215. See also F-actin at adherens junction, 303 of membrane skeleton, 12 regulation of, 204 filensin, 269 fission, mitochondrial, 99 Fj, Golgi expression of, 142 flagella use of term, 245 waveform of, 259, 260 flagellar assembly (FLA), 247 flotillin-1, 168, 170

bindex.indd 449

fluctuation correlation spectroscopy (FCS), 297 fluorescence correlation spectroscopy (FCS), 11, 71, 81 “calibration-free,” 80 purpose of, 80 stimulated emission depletion, 11 fluorescence microscopy, 81, 387, 406 fluorescence recovery after photobleaching (FRAP), 71, 297 and diffusion coefficients, 9 and diffusivity of membrane proteins, 10 early observations, 4 and lipid diffusion, 17 in membrane domains, 73–74 fluorescence recovery after photobleaching (FRAP) experiments, in caveolae, 49 fluorescence resonance energy transfer (FRET), 71, 74–76 fluorescent proteins, genetically encoded, 387 focal adhesion (FA) assembly integrin-mediated, 288, 289 role of Syn4 in, 292 focal adhesion kinase (FAK), 200, 275, 290, 294–295, 298 focal adhesion kinase (FAK) cells, 294–295 focal adhesions (FAs), 271 assembly of, 287–288 complexity of cytoskeletal proteins, 293–294 integrins, 290, 291 Syndecan-4, 292–293 cytosolic composition of, 291 definition for, 285, 286 disassembly of, 296–297 dynamics of, 297, 298 historical perspective on, 285, 287 and keratin IFs, 273, 274, 275 signaling mediators in, 294–296 with stress fibers, 285, 286 focal adhesion targeting (FAT) domain, 294 focal complexes (FCXs), 285 assembly of, 288 disassembly of, 296–297 maturation of, 288, 289, 290 with small actin filaments, 286 focal contacts, 285 formin family of proteins, 12 förster resonance energy transfer (FRET)-based biosensors, 205 fragile X-associated tremor/ataxia syndrome (FXTAS), 406

449

fragile X mental retardation 1 protein (FMRP), 431 freeze-etching technique, 7 FRET-based technologies, 207 fungicides, microtubules targeted by, 240 fusion in endosomal system, 151 of mitochondria, 98–99 galactosyltransferase activity, localization of, 134 galectin-4, 220 galectins, 139 gap junction domain molecular compositions of, 341–343 significance of, 344–345 gap junction plaques, 344 gap junctions definition of, 339 growth of, 343–344 historical perspective on, 339–341 internalization of, 344 and lipid composition of plaques, 343 during mitosis, 344 molecular composition of, 339, 340, 341–343, 342 proteome of, 345 turnover time for, 342 G-Cyclin-associated kinase (GAK), 31 genome, mitochondrial, 89–90, 91 glial cell-derived neurotrophic factor (GDNF), 65 glial fibrillary acidic protein (GFAP), 270 glucose transporter 2 (GLUT2), 221 glycoforms, 138 glycolipid transfer protein (GLTP), 180–181, 185 glycosphingolipids, FRET of, 75 glycosphingolipid synthesis, 185–186 glycosylation Golgi, 135–136 Golgi models for, 138 receptor, 139 glycosylphosphatidylinositol (GPI), 61 glycosylphosphatidylinositol (GPI)anchored proteins, 72, 75, 116 GM1, 170 gold-tagged phospholipid (DOPE), trajectory of, 8 Golgi apparatus, 85, 195 and cell polarity, 139–141, 140, 141

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450

INDEX

Golgi apparatus (cont’d ) and cellular signaling, 139 definition of, 133 and disassembled microtubules, 237–238 domain partitioning in, 134, 135 EM analyses of, 134 glycosylation and, 135–136 historical perspective on, 133– 135, 135 mitosis and, 141 and N-glycan remodeling, 139 and planar cell polarity, 141–142 trafficking to, from, and within, 136–139 ultrastructure of, 134 Golgi-ERJs, functions for, 184 Golgi-ER-lysosome (GERL), 179 Golgi matrix composition of, 134 EM analysis of, 134 Golgi matrix proteins, 140, 140 Golgi reassembly stacking proteins (GRASPs), 134–437 GPI-anchored proteins, in lipid rafts, 64 G-protein-coupled receptors (GPCRs), 29–30, 150, 167 granular component (GC), 396–397 green fluorescent protein (GFP), 207, 231, 403 green fluorescent proteinglycosylphosphatidylinositol (GFP-GPI), 49 growth cone dynamics of, 206–207 membrane, 385 GTPase activation factor (GAF), and Rho family proteins, 205 GTPase exchange factor (GEF), 205, 295 GTPases, in microtubules, 229 GTPase-activating proteins (GAPs), 295 guanine nucleotide exchange factor (GEF), 155, 156 guanosine-5’-triphosphate (GTP), 99 GW bodies, 430 heart disease, and desmosomal alteration, 332 Hedgehog protein, 65 hemidesmosomes, keratin IFs and, 273, 274, 275 heparin-binding domain (HBD), 295 hepatitis C virus, 326 hepatocellular carcinoma (HCC), 399

bindex.indd 450

hepatocyte growth factor (HGF), 158 herbicides, microtubules targeted by, 240 hereditary spastic paraplegias (HSPs), 387 heterochromatin, 394, 395 high-affinity immunoglobulin E (IgE) receptor pathway, 65 Hippo kinase pathway, 141 hnRNP A1-associated protein (HAP), 405–406 hop diffusion, 4 inability to detect, 11 of phospholipids, 5, 11 Hox genes, 404 Hsp90 activity, 441 Huntingtin protein, 440 Huntington’s disease, 387 hybridization, in situ, 395 hydrogenosomes, 89 IgE-Fc receptor, FcεRI, 14 image correlation spectroscopy (ICS), 297 immunity, role of phagosomes in, 165 immunochemical analyses, of IFs, 268 immunoelectron microscopy analysis, 418 immunofluorescence microscopy, 231, 287 immunolocalization studies, of connexins, 341 infectious disease, and desmosome function, 331. See also diseases, human infertility, and microtubule function, 240 inflammatory conditions and desmosome function, 331 and tight junction function, 326 influenza virus, 357 ingenuity software program (IPA), 199 inner boundary membrane (IBM), 87, 95 inner membrane (IM), 95 inner nuclear membrane (INM), 416 inositol-requiring 1 kinase (IRE1), 123, 124 insulin receptors, at neck of adipocyte caveolae, 47 integrin-linked kinase (ILK), 275 integrins, 298 activation of, 292 discovery of, 287

GFP-tagged, 297 keratin IF linkage to, 278 in mature FA structure, 290, 292 interchromatin granule clusters (IGCs), 402 interchromatin particles, 402 interchromosome domain compartment (ICD) model, 395 intermediate filaments (IFs), 195 in cell signaling, 268, 272–273 definition of, 267 functions of, 268 historical perspective on, 267–269 interaction with desmosomes of, 331 as multiple gene family of proteins, 269, 269–270 structure of, 267–268 and surface microdomains, 278 intermediate filaments (IFs), keratin and desmosomes, 273, 274 and epithelial cell asymmetry, 276 and FAs receptor, 275–276 and hemidesmosomes, 273, 274, 275 multifunctional features of, 277 intermediate filaments (IFs)-pathies, 269, 277 intermembrane space (IMS), 87 internalization processes, 165–166 internodes, 378 intraflagellar transport (IFT) in ciliogenesis, 256 components of, 257 intraflagellar transport (IFT) particles, 255 intralumenal vesicles (ILVs), 149 ion channels, light-activated, 387 Jerker mice, 218 JNK interacting proteins (JIPs), 158 Joubert syndrome (JBTS), 258 junctional adhesion molecule C (JAMC), 365 junctional adhesion molecule (JAM) family, 321 junctophilins, 182 juxtaparanodes, 378 Kaps, 422–423 Kartagener’s syndrome, 260 katanin, 234 KDEL retrieval motif, 137 keratins, 270 and cancer metastasis, 278 and cell signaling, 272–273 in colon cancer cells, 278 structural features of, 270, 271

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INDEX

and surface membrane domains, 272, 273–276, 274 kidney stone disease, role of caveolae in, 51 KIF2A, 236 kinases, in actin cytoskeleton, 203– 204, 204 Kinesin-1 family proteins, 236 kinesins, and microtubule catastrophes, 236 knockout mouse models, 218, 284 of alteration in desmosome function, 331–332 and associated diseases, 117 K8-null mouse model, 273 Kv2.1 potassium channels, and membrane skeleton, 14–15 lactate transporters polarity of, 364 proton-coupled, 362 LAMP-1, phagosome membrane identified with, 170 LAMP-2, 138 LAMP-2A, formation of multimeric, 170 leading edge, 384 Leishmania donovani, phagosome fusion inhibited by, 171–173, 172 Lek protein, 79 leucine-based sorting signal, 26 LGN relay neuron, 374 LIM and SH3 domain protein one (LASP-1), 208 LIM kinase (LIMK), 204 lipid analysis, of immunoisolated adipocyte caveolae, 46 lipid rafts, 170, 220, 358. See also detergent-resistant membranes acceptance of, 67–68 biophysics of, 66–67 classical markers of, 63 definition of, 61 historical perspective on, 61–63, 62 in host-pathogen interactions, 67 lipid composition of, 66 and membrane skeleton-induced compartmentalization, 16 protein composition of, 63–66 role for, 61 SPT study of, 79 lipids in caveolae, 46 diffusion of, 9 nonraft, 62 lipid transfer protein (LTP), 181, 186, 188–189 lipoarabinomannan (LAM), 173

bindex.indd 451

lipophosphoglycan (LPG), 171, 172 liposome studies, gap junction channels in, 343 live-cell imaging, 406 liver disease, and keratin gene mutations, 277 low-density lipoprotein (LDL), 148 lysosome-associated membrane proteins (LAMPs), 151 lysosomes, 85 definition of, 165 historical perspective on, 166 and membrane microdomains, 173 microdomains of, 170 major histocompatibility complex (MHC), 165 malignant melanoma, 117 MAMs, 186–187 Markov model, “hidden,” 77 mass spectrometry, 167 mast cells FCS measurements for, 80 role of membrane rafts in, 75 matrix-assisted laser desorption/ ionization (MALDI-TOF-MS), 167 MCAK, 236 MDCK cell line, 271, 355 MDCK model, 356, 357 mean square displacement (MSD) for Brownian diffusion, 72–73 in classical SPT analysis, 76–77 Meckel syndrome (MKS), 258 medicine, role of microtubules in, 240–241 membrane-associated guanylate kinase (MAGUK), 321 membrane bilayers, 1 membrane contact sites (MCSs), 177 membrane domains. See also plasma membrane characteristics of, 71 FCS measurements of, 80 fluorescence microscopy for studying, 81 FRAP in study of, 73–74 FRET microscopy in, 74–76 historical perspective on, 71–72 interconnectedness of, 189 SPT of, 76–80 theoretical basics for, 72–73 membrane rafts, definition of, 72 membrane skeleton, 3 in CCP formation during endocytosis, 15 and cell functions, 17–18

451

and diffusivity of membrane molecules, 10–11 effects on signaling of, 12–15 and Kv2.1 potassium channels, 14–15 and micron-to-cell-sized diffusion, 15 molecular composition of, 12 and regulation of lipid rafts, 16 membrane skeleton-based compartments, effects on reaction kinetics of, 16 membrane skeleton fence model, 5, 6, 78 mental retardation (MR), abnormal dendritic development associated with, 386 mesenchymal-epithelial transition (MET), 208 mesostructures, fabricated, artificial regulation of membrane molecular dynamics with, 17 messenger RNA-protein particles (mRNPs), 430 messenger RNAs (mRNAs), 429 translationally inactive, 431, 433–436 transport of, 431 Met30, 187 metastasis, keratins and, 277 metazoans, development of AJ in, 311 methyl β-cyclodextrin (mβCD) lipid rafts disrupted by, 63, 67 sensitivity of mitochondrial proteins in DRMs to, 65–66 MIA complex, 93 micron-to-cell-sized diffusion barriers, and membrane skeleton, 15 microRNAs (miRNAs), 430 microtubule-associated proteins (MAPs), 229, 240–241, 382 microtubule organizing centers (MTOCs), 133, 139, 140, 232, 253, 276 microtubules, 195 at adherens junctions, 314 basic biology of, 241 cellular polarity determined by, 237 composition of, 230 definition of, 229 under fluorescence microscope, 236 historical perspective on, 230–231 lability of, 231, 232 MAP stabilization of, 234–235 in mature neurons, 382

5/6/2011 6:09:57 PM

452

INDEX

microtubules (cont’d ) molecular composition of, 231 in plant cells, 237 polymerization rates of, 233 protein regulation of, 233–235, 234, 236 spatial organization of as cellular machines, 237–239 nuclear and cell division, 238–239 as tracks for intracellular transport, 235–237 microvilli definition of, 213 F-actin parallel bundles, 217–219 functions of, 224–225 historical perspective on, 214–215 intestinal, 219 and glucose transporter 2, 221 lipid composition of, 220 plasticity of, 221–223, 222 and retention of absorptive factors at membrane, 220–221 and vesicles in intestinal lumen, 221 and pathogens, 223–224 pathologies associated with defects of, 224 proteins associated with, 224 shape of, 215–219, 218 microvillus inclusion disease, 224 mitochondria, 195 biogenesis of, 92, 93 crosstalk with ER of, 100 definition of, 87 discovery of, 87–88 dynamics of fission in, 99–100 fusion, 98–99 quality control, 99–100 functions of, 85 membrane lipids of, 94–95, 95 mitochondrial genome, 89–90, 91 morphology of, 96 origin of, 88–89 pathologic conditions associated with, 100–101 phospholipid composition of, 95 proteome of, 92, 94, 94 research in, 89, 90 mitochondrial DNA (mtDNA), 88, 90 mitochondrial phospholipase D (MitoPLD), 99 mitochondrial proteins, in lipid rafts, 65–66 mitochondrial research in crosstalk with ER, 100

bindex.indd 452

in disease, 100–101 in mitochrondrial dynamics, 95, 96, 97–98 in mitochondrial ultrastructure, 95, 96, 97–98 mitochondrion, scheme of, 88 Mito-ERJs MAMs, 186–187 structural bridge for, 187–188 mitogen-activated protein kinase (MAPK), 203 mitosis, in microtubules, 239 mitosomes, 89 Miwi-binding protein, 437 molecular markers, for identification of nuclear domains, 393, 394 monocarboxylate transporters (MCTs) polarized distribution of, 362–363 proton-coupled, 362–364, 364 variable polarity of, 363 monomeric adaptors, 29–30, 34 Monte Carlo simulations, 7 mouse genetics, 224 mouse knockout studies, 294 mRNA decay, role of P-bodies in, 440 MTK1, 438 mucus, on microvilli, 223 multivesicular body (MVB), formation of, 151 MURC4, 46 murine double minute 2 (MDM2), 398 muscle-cell-specific cavin-4/ MURC4, 39 muscular dystrophy, role of caveolae in, 51 mycobacterium tuberculosis, phagosome membrane microdomains inhibited by, 173 myocardial infarction, 116 myoclonus, and human mitochondrial genome, 91 myosin, 198 myosin-1a, 218, 220 myosin light chain (MLC), and ROCK, 288 myosin light-chain kinase (MLCK), 203 myosin XVa, 217 myotonic dystrophy (DM), 403 Na,K-ATPase α- and β1-subunits, 364 mechanism involved in sorting of, 359–361, 360–361 in transporting epithelia, 351–352, 353, 354

nano-meso structure, controllable, 17 Naxos disease, 332 Ndc1p, protein, 418 nectin, 304–305, 310, 312, 315 Neisseria gonorrhoeae, in HEK 295 cells, 173 nephronophthisis (NPHP), 249, 258 N-ethylmaleimide-sensitive factor (NSF), 153 neural cell adhesion molecule (NCAM), 365 neural networks, formation of, 371 neurological disorders, and microtubule function, 240 neuromuscular synapse (NMJ), 378 neuronal domains, 372–373, 374, 375, 376 axonal domains, 373, 376, 376– 377, 378–379 cytoskeleton and, 381–382, 382 dendritic domains, 379–381 during development, 384 axon and dendrite guidance, 385–386 during migration, 384 polarity established, 384–385 and extracellular matrix, 382 and mental disorders, 386–387 research in, 387 soma, 373 neurons definition for, 371 diversity of, 373 historical perspective on study of, 371–372 molecular composition of specialized membrane domains in, 376, 376–377, 378 schematic of, 375 subdomains for, 373 vertebrate, 374, 374 visualizing individual, 372 nicotinamide adenine dinucleotide phosphate (NADPH), 171 Niemann-Pick C1 (NPC1) protein, 167 NMDA, 380 Nobel Prize recipients, 90 nodes of Ranvier, 378 normal human diploid fibroblasts (NHDFs), 396 normal rat kidney (NRK) cells, 3 Notch pathways, 160 nuclear bodies, identification of, 406 nuclear domains cajal bodies, 401–402

5/6/2011 6:09:57 PM

INDEX

chromatin, 393–395 definition for, 393, 394 molecular markers for identification of, 393, 394 nuclear gems, 401–402 nuclear splicing speckles, 402–404 nucleolus, 395–399, 396 PML nuclear bodies, 399–401 PNC, 404–405 polycomb bodies, 404 SLM nuclear bodies, 405–406 nuclear envelope (NE), 415, 416 nuclear gems, 401–402 nuclear magnetic resonance, structural studies by, 26 nuclear pore definition of, 415 functions of, 425 historical perspective on, 416–417 nuclear pore complex (NPC), 415, 416, 425 establishing directionality for, 424–425 model of, 417 molecular organization of, 417– 418, 419, 420–422 structural complexity of, 416 structural model of, 419 transport through, 422–424 nuclear speckles, in situ structure of, 396 nuclear transport factor (NTF), 422, 425 nucleoids, 90 nucleolar organizing regions (NORs), 396 nucleoli, 393, 395–399, 396 in cancer cell biology, 399 during cell cycle, 398 first observations of, 397 formation of, 396 in p53 regulation, 398 in situ structure of, 396 nucleolus-derived foci (NDF), 398 nucleus and movement of macromolecules, 425 role of, 393 various nuclear domains of, 394 nucleus-vacuole junctions (NVJs), molecular composition of, 179–181, 180–181 Nup84-containing subcomplexes, 420–421 Nups identification of, 417–418 subgroups of, 418

bindex.indd 453

“oligomerization-induced trapping” effect, 9 oligosaccharyltransferase (OST), 135–136 oncoproteins, viral, 327 online mendelian inheritance in man (OMIM), 199 OPA1, 98 oryzalin, 246 OSBP-related domain (ORD), 180–181 OSBP-related proteins (ORPs), 183, 184 Osh proteins, 183, 189 osteogenesis imperfecta (OI), 403 outer nuclear membrane (ONM), 416 Oxa1 complex, 93 oxysterol-binding protein (OSBP), 181, 183, 185 PACS-2, 188 paranodes, 378 parathyroid hormone receptor (PTHR), 365 PAR complex, 312 Parkinson’s disease, 91, 125, 125–126, 387 pathfinding, 206 P-bodies, 438–440 formation of, 439 SGs compared with, 440–441 pemphigus, 328–329, 331 peptide correlation profiling-stable isotope labeling by amino acids in cell culture (PCP-SILAC), 66 perichromatin fibrils (PFs), 402 perinucleolar compartment (PNC), 393, 404, 405 p53, 400 phagocytes, key roles of, 167 phagocytosis, 85, 166, 168 phagolysosomes, maturation of phagosomes into, 169 phagosome membrane composition of, 169 microdomains on, 168–170, 169 phagosomes definition of, 165 early, 166–167 formation of, 165, 166, 168 host-pathogen interactions of, 171–173, 172 and membrane microdomains, 173 proteome of, 167–170 phakinin, 269 phenylalanines in acidic tract (FFAT) motif, 184

453

phosphatases, in actin cytoskeleton, 203–204, 204 phosphatidylinositol 4,5-biphosphate (PIP2), 294 phosphatidylinositol-4-phosphate adapter proteins (FAPPs), 185–186 phosphatidylinositol 4-phosphate (PtdIns(4)P), 32 phosphatidylinositol transfer protein (PITP), 180–181, 186 phosphatidylinositol transfer protein (PITP) family proteins, 189 phosphoinositides (PIPs), 182, 184 phospholipids, hop diffusion of, 5, 11 phosphoproteins, in actin cytoskeleton, 203–204, 204 phosphoproteome, mitochondrial, 92 phosphoproteomic analyses, of actin cytoskeleton, 208 photo-activated localization microscopy (PALM), 81 physiological disorders, and ciliary dysfunction, 261 picket-induced slowdown effect, 6, 7 pickets, protein, and lipids, 5, 6, 7 PIPK1γ90, 293 plakin family members, of desmosome, 330 plakoglobin, 305, 310, 330 planar cell polarity (PCP), 261 plasma membrane (PM) compartmentalized view of, 5 diffusivity in, 4 domain organization of, 1 electrical potential difference across, 373, 376 electron tomography of cytoplasmic surface of, 7–8, 8 and F-actin parallel bundle, 219 influence of cytoskeleton on, 3 mesocale domains in, 3 molecular transport across, 1 properties of, 4 turnover of E-cadherin at, 313 plasma membrane (PM) apical-basolateral polarization, 356–359 in different epithelia, 352–354, 354 plastin-1, 217–218, 218 pleckstrin homology (PH) domain, 157

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454

INDEX

pleckstrin homology (PH) domain containing, family A member 7 (PLEKHA7)-Nezha complex, 314–315 plectin, 275 plus-end tracking proteins (+TIPs), 235 polar bodies, 430 polar granules (P-granules), 431, 432, 436–437 polarity cell, 349 and sorting machinery, 365 Pol I and transcript release factor (PTRF), 39, 45 poly(A)+-binding protein I (PABP-I), 437 polycomb (Pc-G) bodies, 404 polycystic kidney disease (PKD) flagellar assembly mutant associated with, 247 and microtubule function, 240 and sensory dysfunction of cilia, 258 Poms, 418 p130cas, 298 pore complex, 416 posterior sex combs (Psc), 404 postsynaptic density (PSD), 380 postsynaptic specializations, 379–380 posttranslational modifications (PTMs), 241 prenucleolar bodies (PNBs), 398 presynapses, 376–377, 378 primary ciliary dyskinesia (PCD), 260 processing bodies (P-bodies), 430 prohibitin, 168 prometaphase, microtubules during, 239 promyelocytic leukemia (PML) gene, 400 promyelocytic leukemia nuclear bodies (PML NBs), 393, 399–401 and antiviral response, 401 as catalytic surfaces, 400 formation of, 399 in situ structure of, 396 as tumor suppressors, 400 promyelocytic leukemia (PML) protein, 399 protein disulfide isomerase (PDI), 119–120 protein disulfide isomerase (PDI)like proteins, 115 protein folding, energy landscape model of, 35 protein kinase A/C (PKA/C), 204

bindex.indd 454

protein kinase C (PKC), 275 protein kinase C delta-binding protein (PRKCDBP), 45 protein-lipid dynamics, on mesoscale, 7 proteins. See also specific proteins acyl-anchored, 65 at adherens junction, 306–309 Cav family of, 43 cilia-related, 249–253 diffusion of, 9 ER Ca2+ buffering, 121–122 in ERJs, 178 from F-actin cytoskeleton network, 201 GPI-anchored, 64 identification of phagosomal, 167–168 intraflagellar transport (IFT), 251 mitochondrial, 65–66, 93 in muscle tissue, 198 nucleolar, 396, 397 signaling, 64–65 spatiotemporal dynamics of, 80 proteins, intermediate filaments (IFs), 267 assembly, 270–271, 271 classes of, 269 and cytoskeletal interplay, 271– 272, 272 proteomes, for identifying components of cilia, 248, 249–253 proteomics definition of, 115 era, 167 proximal tubule (PT), kidney epithelial cell polarity of, 352– 354, 354 lactate levels in, 363 Pseudomonas aeruginosa, 173 pseudopodium-enriched atypical kinase one (PEAK1), 208 PTB/hnRNP1, 404–405 PtdIns(4,5)P2 lipid, 30–31 PTP-1B, 184 p21-activated kinase (PAK), 203 pyramidal neuron, 374 quality control, of mitochondria, 99–100 Rab conversion, 155 Rab domains concept of, 153–154, 154 and directionality of membrane traffic, 155, 156 endocytic routes and, 149 Rab GTPase code, 153 self-assembly, 154

rab8, 224, 253 raft domains, exclusion effect of, 10. See also lipid rafts Ran GTPase-activating protein (RanGAP), 424–425 rapid-freeze deep-etch techniques, caveolae demonstrated by, 47 Ras, 140 raster-scan image correlation spectroscopy (RICS), 297 Rdgb, 184 reaction kinetics, and membrane skeleton-based compartments, 16 reactive oxygen species (ROS), 296 receptor for activated C kinase 1 (RACK1), 272–273, 275, 438 recycling endosome (RE), 151 red fluorescent protein (RFP), 188 repeating unit domains (RUDs), of desmogleins, 330 replication, and chromatin, 395 rescue, in microtubule dynamics, 233 residency times, determination of, 11 retinal bipolar neuron, 374 retinal pigment epithelium (RPE) cell polarity in, 351 epithelial cell polarity of, 354, 355 retinitis pigmentosa, 249, 258 retinoic acid receptor-alpha (RARA), 400 Rho-associated kinase (ROCK), 204, 206, 288 Rho family proteins, 205, 206 Rho GTPases, in maturation of FCXs into FAs, 295, 295–296 ribonucleoprotein particles (RNPs), 429–430 functions of, 441 known protein components of, 433–436 P-bodies, 438–440 P-granules, 431, 432, 436–437 and posttranscriptional gene regulation, 430, 431, 432 stress granules, 432, 437–438 transport granules, 430–431, 431 ribosomes, biogenesis of, 397 rippling muscle disease, role of caveolae in, 51 RNA-based therapies, 441 RNA-binding proteins, 429 RNA domains definition for, 429 historical background for, 429–430 RNA interference (RNAi), 430

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INDEX

RNAs, in nucleolus, 397 ROCK (Rho kinase), 204, 206, 288 salivary gland, lactate levels in, 363 SAM complex, 93 Sam68/SLM nuclear bodies (SNBs), 405–406 sarcoplasmic reticulum (SR), 114 scaffolding domain, of Cav molecule, 44 scaffold proteins, 290 SCAR/WASP family verprolinhomologous protein (WAVE), 12 sdr-related gene product that binds to c-kinase (SRBC), 39, 45 serine-argenine-rich (SR) protein family, 402–403 serum deprivation protein response (SDPR), 39, 45 Shigella flexneri, 223–224 sialylation, 134 signaling effects of membrane skeleton on, 12–15 in ERJs, 178 signaling protein oligomeric transduction structures (SPOTS), 276 signaling proteins, in lipid rafts, 64–65 signal transduction pathways, receptor redistribution and clustering in, 14 Singer-Nicolson fluid mosaic model, 71 single fluorescent-molecule tracking (SFMT), 4 single-particle tracking (SPT), 4, 71 analysis of particle tracks with, 76–77, 78 models of plasma membrane domains studied with, 77–79 phospholipid hop diffusion revealed by, 8 in T-cell signaling, 79–80 single-particle trajectories, compartment sizes deduced from, 5 site of flagellar autotomy (SOFA), 254 SLC16 gene family, 362 sliding window statistical analysis, 77 SMAD anchor for receptor activation (SARA), 157, 158 small interfering RNAs (siRNAs), 430

bindex.indd 455

small nuclear ribonucleoprotein (snRNP), 401, 402 small ubiquitin-like modifier (SUMO)-interacting motif (SIM), 399 soluble NSF attachment protein (SNAP) receptors (SNAREs), 153, 154, 154 soma, 371, 373 sorting nexin 9, 31 sorting nexin (SNX), 155–156 sorting signals basolateral vs. apical, 365 in epithelial cell polarity, 358 recognized by AP-2, 29 Spalt, 160 spastin, 234 speckles, nuclear splicing, 402–404 spectrin tetramer dissociation gate model (SPEQ gate model), 12 sphingolipids, caveolae in, 46 sphingomyelin, in TCR membrane domains, 66 sphingomyelin synthesis, 185 spinal motor neuron, 374 spinal muscular atrophy, 387, 401 spindle assembly, 238–239 spines, dendritic, 380–381 SPT techniques, quantum-dot-based, 13 Src family kinases (SFKs), 290, 294 Src homology-3 (SH3) domains, 32 stathmin, 233 stathmokinetic effect, 231 stereocilia actin polymerization at tips of, 217 of microvilli, 213, 214 proteins associated with, 225 steroidogenic acute regulatory (STAR)-related domain (START), 185 sterol traffic, role for PM-ERJs in, 183 stimulated emission depletion (STED), 81 stimulated emission depletion FCS (STED-FCS), 11 stomatal diaphragm (SD), of caveolae neck, 47, 48 stomatin, 168 store-operated calcium entry (SOCE), 182 stress granules (SGs), 430, 432, 437–438, 440–441 stromal interacting module 1 (STIM1), 238 stromal-interacting molecule 1 (STIM1), 114, 122

455

subretinal space (SRS), lactate levels in, 362–363 sucrase-isomaltase deficiency, 220 suppressor two of zeste, 404 synapses and dendritic geometry, 381 isolation of, 372 synaptic biology, 15 synaptic cleft, 379 synaptic disorders, 386 synaptojanin, 31 Syndecan-4 (Syn4), 292–293 systems biology, 160 talin, 288, 290, 298 in FA, 293 and FA disassembly, 296 TANGO1 protein, 137 taxol, and microtubule dynamics, 240 T-cell receptor (TCR) clustering, 141 GEF associated, 140–141 pathway, 65 recovery of fluorescently labeled, 74 T-cells FCS measurements for, 80 receptor microclusters in, 72 T-cell signaling, SPT in, 79–80 telophase, microtubules during, 239 terminal web, 218 tetraspan proteins, of tight junction, 325 TGN-to-plasma membrane carriers (TPCs), 137 tight junction definition of, 321 early references to, 322 electron microscopy, 322 molecular composition of, 323, 324, 325–326 as paracellular gate, 324 pathophysiology of, 326–327 tight junction proteins, 323 TIM22 complex, 93 TIM23 complex, 93 TOM complex, 93 total internal reflection fluorescence (TIRF), 76, 208 toxoplasma, oryzalin resistance in, 240 TRAF2, 438 transcription, and chromatin, 395 transendothelial channels (TECs), SDs on, 47 transfection techniques, 358

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456

INDEX

trans-Golgi network (TGN), 133, 135, 137, 148, 186 formation of APs at, 24 PM proteins synthesized in, 357, 357 transition zone (TZ), of cilia composition of, 254 proteins related to, 249–253 releasing BB from, 261 transmembrane proteins crowding effect of, 10 diffusion coefficient for singlepass, 10 membrane-skeleton fence model for, 4–5, 6 of tight junction, 326 transmission electron microscopy (TEM), caveolar coat on, 42, 43 transport granules, 430–431, 431 triad junction, 177, 182 Tsc13, 180, 181 tubulin gene-in, 232 tubulins, 267 binding of stathmin to, 233 in microtubules, 229 multigene family of proteins of, 232–233 posttranslational modification of, 241 tumor cells, keratins and, 277 tumorigenesis, role of junction proteins in, 332–333 twinfilin-2 transport, 217 tyrosine-based sorting signals, 26

bindex.indd 456

ubiquitination, 314 ubiquitin-interacting motifs (UIMs), 30 UDP-glucose: glycoprotein transferase (UGGT), 117, 117 unfolded protein response (UPR), 116, 118, 119 activation of, 124, 124–125 and ER stress, 123 protein synthesis inhibited by, 136 urokinase-type plasminogen activator (uPA), 277 vacuolar membrane-associated protein (VAMP)-associated protein (VAP), 183 valosin-containing protein (VCP), 120 vascular diseases, 116 vesicular stomatitis virus (VSV), 49, 357 Golgi-associated, 138 G protein, 134 vesiculo-vacuolar organelle (VVO), 47, 48 villin, 217, 218, 222–223 vimentin, 270 and cancer metastasis, 278 endothelial cells and, 278 vinblastine, and microtubule dynamics, 240 vinculin, 288, 290, 298 discovery of, 287 in FAs, 294 and Syn4, 292

vinculin null cells, 294 viruses, and tight junction proteins, 326 whirlin, 217 Wiskott-Aldrich syndrome protein (WASP), 12 Wiskott-Aldrich syndrome protein (WASP) and SCAR homolog (WASH), 156–157 Wnt/Wingless, 160 X-box-binding protein 1 (XBP1), 124 Xenopus laevis, P-granules in, 436 XMAP215 family, 235 X-ray crystallography, 26 yeast cortical ER in, 182–183 homeostasis of RNPs in, 441 NPC analysis in, 417–418 Osh proteins in, 183, 189 P-bodies of, 439 PI transfer proteins, 186 yellow fluorescent protein (YFP), 76, 343 zonulae adherens, 328 zonula occludens (ZO), 322, 328. See also tight junction function for, 325 original description of, 321–322 zonula occludens (ZO) proteins, 321

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