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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

PROTEIN BIOCHEMISTRY, SYNTHESIS, STRUCTURE AND CELLULAR FUNCTIONS

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ACTIN: STRUCTURE, FUNCTIONS AND DISEASE

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Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

PROTEIN BIOCHEMISTRY, SYNTHESIS, STRUCTURE AND CELLULAR FUNCTIONS

ACTIN: STRUCTURE, FUNCTIONS AND DISEASE

VICTORIA A. CONSUELAS AND

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

DANIEL J. MINAS EDITORS

Nova Science Publishers, Inc. New York

Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

Copyright © 2012 by Nova Science Publishers, Inc.

All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers‟ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works.

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Library of Congress Cataloging-in-Publication Data Actin : structure, functions, and disease / editors, Victoria A. Consuelas and Daniel J. Minas. p. ; cm. Includes bibliographical references and index. ISBN:  (eBook) I. Consuelas, Victoria A. II. Minas, Daniel J. [DNLM: 1. Actins--physiology. 2. Actins--ultrastructure. 3. Neoplasms--physiopathology. 4. Respiratory Tract Diseases--physiopathology. QU 55.3] LC classification not assigned 362.196'2--dc23 2011031650

Published by Nova Science Publishers, Inc. † New York Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

Contents Preface

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Chapter I

vii Actin Dynamics and Remodeling of Cell-Cell Junctions in Epithelial Morphogenesis Marc D.H. Hansen and Michael R. Stark

Chapter II

Actin: Structure, Function and Disease YamilaTorres Cleuren and Johannes Boonstra

Chapter III

Osmotic Pressure: A Tool to Investigate the Polymeric Forms of Actin Enrico Grazi

Chapter IV

Two Communication Bridges to One Versatile Molecule Ricardo Mondragón and Doris Cerecedo

Chapter V

Stressed Out and Actin Up: Stress-Activated Protein Kinase Regulation of Actin Remodeling Directs Endothelial Cell Morphology and Migration Meron Mengistu, Joshua B. Slee and Linda J. Lowe-Krentz

1 61

97 137

177

Chapter VI

The Molecular Mechanisms of Actin Regulatory Proteins Barak Reicher and Mira Barda-Saad

207

Chapter VII

Actin Cytoskeleton Alterations: Are There Any Consequences? Silvia Versari, Livia Barenghi and Silvia Bradamante

229

Chapter VIII

Role of the Actin Cytoskeleton in Tumor Escape to Immune System and Acquisition of Tumor Resistance to Cytotoxic Treatment Rania Zaarour, Fui Goh, Meriem Hasmim, Bassam Janji and Salem Chouaib

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245

vi Chapter IX

Chapter X

Contents Insight into Force Transmission along Actin Filaments: Sliding Movement of Actin Filaments Containing Inactive Components on Myosin Molecules Syunsuke Matsushita and Kuniyuki Hatori Hypoxia Induces Endothelial Barrier Dysfunctions in Lungs: Nitric Oxide Perturbs Actin Dynamics in Endothelial Cells Swaraj Sinha, Hima Bindu Reddy and Suvro Chatterjee

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Index

Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

257

271 283

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Preface Actin is a globular protein found in all eukaryotic cells and plays a central role in cell morphology, cell adhesions, cell contractility and motility, signal transduction, transcription and its regulation, cytokinesis and synapse formation, as well as malfunctioning of actin which can lead to various diseases. This book presents topical research in the study of the structure, functions and diseases related to actin. Topics discussed include actin dynamics and remodeling of cell-cell junctions in epithelial morphogenesis; osmotic pressure and the polymeric forms of actin; the molecular mechanisms of actin regulatory proteins and actin cytoskeleton alterations Chapter 1- Cell-cell adhesion systems play a critical role in integrating individual cells into tissues. Crucial to cell-cell adhesion is the structure and dynamics of the actin cytoskeleton, particularly at cell-cell junctions. Such organization forms the basis of proper cell and tissue morphology. Control of actin dynamics at adhesion sites allows epithelial tissues to undergo dramatic remodeling events. These epithelial morphogenesis events occur throughout development. The staggering variety of cell and tissue architecture changes during epithelial morphogenesis has resulted in events often being categorized by specific types of cellular rearrangements, with little regard to understanding epithelial morphogenesis in a unified manner. All morphogenetic rearrangements rely on movements and shape changes made by individual cells within the tissue. When these movement and shape changes occur collectively, they result in repositioning of individual cells within the tissue or an overall deformation of the tissue. The unfiying feature of such events is that cell-cell adhesion are maintained, allowing individual cell morphology changes to alter the overall tissue architecture. Thus, actin dynamics in individual cells are coordinated thorough cell-cell contacts to drive changes in morphology at the tissue level. Epithelial morphogenesis also includes tissue remodeling events where cell shape changes and movement occur in isolation and do not impact the entire tissue, allowing cells to behave in a solitary fashion. An example is epithelial-mesenchymal transition (EMT), where cells completely detach from the epithelial tissue and migrate to new locations. Importantly, epithelial morphogenesis events that result in individual cells detaching from the tissue highlight how cells can uncouple actin dynamics from cell-cell adhesion systems to generate completely different outcomes on tissue architecture. Given the central role of actin-membrane connections at adhesion sites in determining whether actin dynamics of individual cells occur collectively or independently, it is important that the molecular basis of membrane connections at adhesion sites has been

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Victoria A. Consuelas and Daniel J. Minas

under recently renewed scrutiny. The prevailing dogma that cadherin-based adhesions are directly anchored to actin filaments, thus providing the majority of functional actin-membrane linkages at adhesion sites, has been challenged and the precise molecular organization of actin-membrane connections of cell-cell adhesions is under active revision. Actin-membrane anchor sites at epithelial cell-cell junctions are typically symmetrical, with actin membrane linkages occurring on both sides of the adhesion site. A benefit of such organization it that symmetrical connection points allow the actin cytoskeleton of individual cells to be integrated into a larger multi-cellular actin network, connected through points of cell-cell adhesion. This network can be regulated at the level of the whole tissue to drive collective epithelial morphogenesis events. From the perspective of actin dynamics and actin-membrane connections, the entire variety of epithelial morphogenesis events can be viewed as a spectrum of outcomes that result from changes in actin organization and contractility, as well as whether these changes are coordinated through actin-membrane connections with cell-cell adhesion systems. In this view, the extent, order, and location of changes in the actin cytoskeleton in each cell constrain the outcome of epithelial morphogenesis in terms of tissue architecture. A detailed understanding of how actin reorganization and contractility are coordinated through cell-cell contacts of epithelial cells will be critical in developing such a unified view of epithelial morphogenesis events. A major goal of cell biologists has been to characterize actin dynamics in live cells and identify how specific molecular machines generate those actin dynamic events. A major emphasis of developmental biologists has been to understand how signaling networks are regulated in time and space to drive specific developmental events. This chapter will focus on how spatiotemporal signaling and actin rearrangements of cell-cell adhesions interface during development to generate specific changes in tissue morphology. Chapter 2- Actin is a globular protein found in all eukaryotic cells. Depending on its location, it can form different structures and perform various functions. Actin monomers (Gactin) come together to form filaments (F-actin); it is found abundantly in the form of microfilaments and thin filaments in cells. With the help of different actin-binding proteins that regulate its structure and activity, it can assemble in several combinations giving rise to actin bundles and networks with differing functions. Playing a central role in cell morphology, cell adhesions, cell contractility and motility, signal transduction, transcription and its regulation, cytokinesis and synapse formation, malfunctioning of actin can lead to various diseases; among them congenital myopathies, compromised immunity, neurodegeneration, and cancer spread. The structure and function of actin and its role in different diseases are here discussed. Chapter 3- A very impressive property of actin is the versatility in the formation of supramolecular structures. G-actin is converted into filamentous or globular structures, it is ordered into paracrystalline arrays or in tubular structures. The formation of these almost regular structures is accompanied by distinct changes of the osmotic properties of the actin solution so that, in some cases, the geometric parameters of the supramolecular structure can be related, albeit indirectly, to protein osmotic pressure. With the increase of protein osmotic pressure protein-bound water is removed and the volume of the system decreases. These events are necessarily linked to changes of the geometric parameters of the supramolecular structure. The study of protein osmotic pressure may thus provide information on the geometry of the supramolecular assembly, even though the geometry depends in a complex manner on the concentration and charge of the protein. In addition, change of protein osmotic

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Preface

ix

pressure may signal the transition of the actin filaments into a network of filaments. The study of osmotic protein pressure allows to determine the activity and the change of the free energy of the solute in binary solutions and, in some cases, in ternary solutions. Two further points deserve consideration: A. The free energy of the free actin monomers is related to the protein osmotic pressure generated by polymeric actin solutions. B. With the model of Biron et al. [Europhys. Lett 2006, 73, 464] it is possible to relate the change of the free energy of the free actin monomers to the change of the length distribution of the actin filaments. It follows that the length distribution of the actin filaments is regulated by (1) the free energy of hydrolysis of ATP and (2) the protein osmotic pressure. Chapter 4- Actin helical filaments are the key tools of the cytoskeleton for adapting cells to the physical or chemical microenvironment signals organizing cell contents, coordinating movement, or changing shape. Actin polymerization is controlled by regulatory proteins including nucleation, depolymerizing and severing factors, capping proteins, polymerases, crosslinkers, and stabilizing proteins. The cell‟s exquisite sensitivity in responding to a wide range of physical or chemical stimuli, is translated into cytoskeleton reorganization and adhesion-site modulation. Living cells survive, proliferate, or differentiate while they are anchored to their extracellular matrix. It comprises a complex bulk of information integrated into a coherent environmental signal. This is achieved through integrins that are the major family of transmembrane adhesion receptors composed of α and β units. These heterodimers not only play an anchorage mechanical role they also transmit chemical signals into the cell concerning their microenvironment and adhesive state. Chapter 5- Actin remodeling in the vascular system is central for functions such as vascular remodeling and contractility. Endothelial cells (EC) that form a monolayer lining the vasculature undergo remodeling of their actin cytoskeleton in order to (1) change their polarity, which gives them contractility that allows them to reduce their height in order to decrease the magnitude of the strain they experience from the blood flow, (2) migrate during vascular remodeling, and (3) maintain cell-cell contacts for endothelial integrity and barrier function. Dysfunction in ECs is the first step of atherogenesis, where cell morphology, migration, and barrier integrity are affected. Atherosclerosis is a geometrically focal disorder where endothelial dysfunction and subsequent plaque formation occur in areas of blood recirculation, such as the outer wall of vessel bifurcations and areas near vascular branching points. Blood flow exerts different magnitudes of fluid shear stress (FSS) on endothelial cells, playing a crucial role in the localization of atherosclerotic lesions. ECs found in regions of arterial curvatures and bifurcations experience lower FSS conditions and are pre-disposed to atherosclerotic lesions, while ECs lining the straight parts of arteries are exposed to higher FSS conditions and are atheroprotected. FSS is an important regulator of EC morphology and migration. Under low FSS, ECs are polygonal in shape and experience high turnover, while they are elongated with their longer axes that align in the direction of flow when exposed to higher FSS conditions. Such morphological change is driven by FSS-stimulated fiber formation and alignment of these fibers in the direction of flow. Cell-cell integrity is also compromised during atherogenesis, where breaks in the monolayer of ECs provide openings for monocytes and low-density lipoprotein (LDL) to enter the wall of the artery and contribute to plaque formation. After atherosclerotic lesions have formed, one of the treatment options is angioplasty where the blocked or narrowed arteries are opened using stents. Endothelialization of stents, which requires EC migration, is important to control

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Victoria A. Consuelas and Daniel J. Minas

vascular tone and prevent restenosis, the re-narrowing of the artery. In order to remodel the actin cytoskeleton for morphology changes, migration and maintenance of the barrier function, FSS and other stimuli need to be translated into chemical signals. The signaling activities of Stress-Activated Protein Kinases (SAPKs) JNK and p38 are involved in actin remodeling events that regulate actin dynamics. Both JNK and p38 activities are transiently activated by the higher FSS treatments, and required to achieve actin alignment in the direction of flow. Other stimuli that alter actin remodeling and barrier changes also induce activation of stress kinases. JNK activity is detected in association with stress fibers and cortical actin, and its activity is required in EC morphology adaptation. p38 activity seems to play a role in actin remodeling near focal adhesions, where it is detected at the ends of stress fibers. In this chapter, we review the role of these SAPKs in actin remodeling that leads to EC alignment in the direction of flow, migration, and maintenance of cell-cell integrity. Chapter 6- Actin polymerization is the driving force behind multiple cellular processes including proliferation, motility, adhesion and endocytosis, providing the infrastructure for structural cellular remodeling and intracellular signal transduction. The variety and flexibility of cellular function is achieved by the formation of diverse actin structures. de novo formation of actin filaments is controlled by a combination of G-actin binding proteins, actin severing proteins, and capping factors. In order to overcome these regulatory checkpoints and support efficient actin polymerization, the orchestrated activity of various actin elongation and nucleation proteins, such as formins and the ARP2/3 complex is required. This delicate balance between actin polymerization and de-polymerization must be highly controlled to ensure the formation of the correct actin structures at the correct place and time. The highly controlled actin polymerization process is enabled by the activity of actin nucleation promoting factors (NPFs). These proteins are present in molecular complexes that associate with actin nucleation proteins and are involved in their stabilization and activation at actin-rich sites. These complexes work in concert to control the spatial and temporal formation of diverse actin structures. Recently, the development of cutting-edge imaging technologies have provided new insights into the regulatory mechanisms of the NPFs, and their modes of activation, dynamics and association with the actin nucleation proteins. In this review, we describe the activation mechanisms of actin regulatory proteins, their differential and overlapping functions in actin-dependent processes and structures, and their role in health and disease. Chapter 7- Actin is one of the most abundant and highly conserved proteins of all eukaryotes. It is the major component of the cytoskeleton, and is involved in many of the structural and dynamic aspects of cell growth, differentiation, division, membrane organisation, transport, and signal transduction. Alterations in such a critical component can lead to pathological conditions. We here describe the effects of actin cytoskeleton disorganisation and/or depolymerisation in the Saccharomyces cerevisiae yeast model system. The structure of the actin cytoskeleton was disorganised by subjecting yeast cells to simulated (Rotating Wall Vessel) or real microgravity (spaceflight), both of which activated the signal transduction cascade of the high osmolarity glycerol (HOG) MAP kinase pathway, which responds to cell swelling/shrinking, and the cell wall integrity (CWI) pathway, which is involved in cell wall biogenesis and actin cytoskeleton reorganisation. The same results were observed when the actin cytoskeleton structure was depolymerised by means of

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Preface

xi

treatment with dihydrocytochalasin B (DHCB) or (+)-(R)-trans-4-(1-aminoethyl)-N-(4pyridyl)cyclohexanecarboxamide dihydrochloride (Y-27632). Chapter 8- Cancer cell survival is a fundamental process essential for cancer related mortalities. This process could be related either to a defect in the immune system machinery or to an acquiring of survival properties by cancer cells allowing them to survive despite a functional immune system. How a cancer cell survives in an environment where competent immune cells are present remains an important question. Understanding basic mechanisms of tumor-host immune interactions will shed light on developing methods to eradicate tumor cell survival, to attenuate immunotherapy resistance and to develop targeted anti-cancer drugs. Actin and tubulin form highly versatile, dynamic polymers that can organize cytoplasmic organelles and intracellular compartments, define cell polarity, and generate both pushing and contractile forces. Therefore, it is not surprising that they are key players in many processes in cell biology. Accumulating evidence involve the actin cytoskeleton at the core of this question. Indeed, the cytoskeleton plays an essential role in regulating a plethora of molecular events that ultimately lead to death resistance by cancer cell. In this regard, cytoskeletal remodeling has been described to be responsible for the resistance of cancer cells to cytotoxic immune cell attack as well as in the activation of the attack by immune cells. In this chapter we focus on the role of the actin cytoskeleton in the processes of regulating T cell activity and evading T lymphocyte mediated killing events. Chapter 9- The sliding movement of actin filaments, consisting of heterogeneous components, on skeletal muscle myosin molecules was examined to specifically evaluate the effect of internal modulation of the actin filaments for force transmission on the sliding movement. Inactive actin molecules were prepared by conjugation with indocarbocyanine fluorescent dyes (IC3-OSu or Cy3-NHS) in molar ratios greater than a 3-fold excess. IC3OSu is an analogue of Cy3-NHS, and it can bind to primary amino groups. IC3-conjugated actin (IC3-actin) monomers were polymerized into the filaments which led to complete impairment of both motile activity and myosin-ATPase activation. Filaments of Cy3conjugated actin (Cy3-actin) exhibited a decrease in velocity to a third of the value (33%) observed for intact actin filaments. In the absence of ATP, dissociation rates of IC3- and Cy3actin filaments from myosin molecules were greater than those of intact actin filaments, indicating that IC3- and Cy3-actin act as smaller resistance components against sliding movement compared to intact filaments. Subsequently, two types of copolymer filaments were prepared. The first type of copolymer were filaments copolymerized homogeneously with intact actin monomers and IC3-actin monomers, while the second kind were block copolymer filaments composed of two short filaments of intact actin and IC3-actin. The sliding velocities of these copolymer filaments hyperbolically decreased as the fraction of IC3-actin monomer increased. In practice, 75% IC3-actin within homogeneous copolymer was required to reduce the velocity by half. In the case of block copolymer 65% IC3-actin led to the same decrease in velocity. For Cy3-actin copolymer filaments similar differences between homogeneous and block copolymer filaments were also observed. Drag ratio between IC3-actin (or Cy3-actin) and intact actin was estimated by consideration of the force balance between the power force and the drag force imposed on a filament during steady movement. Consequently, the drag ratios of IC3-actin to intact actin were 0.31 (homogeneous copolymer) and 0.47 (block copolymer), respectively. Thus, IC3-actin incorporated homogeneous copolymers exhibits a smaller resistance to sliding movement than IC3-actin modified block copolymers.

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Victoria A. Consuelas and Daniel J. Minas

Chapter 10- Endothelium forms a physical barrier that separates blood from tissues. Communication between blood and tissues occurs through the delivery of molecules and circulating substances across the endothelial barrier by directed transport either through or between cells. In response to different stress stimuli, the endothelial barrier becomes less restrictive that results in increased water and protein permeability. Two mechanisms together or independent of each other may account for such an increase in permeability: paracellular (i.e., between cell) and transcytotic (i.e., through cell) transport. The role of intercellular gaps in mediating paracellular permeability also has been determined by osmotically shrinking endothelial cells.Vascular remodeling in response to stress is a common complication of many pulmonary abnormalities such as pulmonary hypertension; vein graft remodeling and high altitude induced pulmonary edema (HAPE). The actin cytoskeleton is a dynamic structure that undergoes rearrangement under the control of various actin binding, capping, nucleating, and severing proteins, which are intimately involved in integration of endothelial monolayers. Therefore, actin filaments are of critical importance to endothelial cells permeability. Although significant progress has been made in understanding the cellular and molecular events that regulate permeability, the mechanistic insight of endothelial barrier functions in clinical conditions such as HAPE remains elusive. An alternative strategy to protect barrier integrity could be achieved by focusing on the downstream signaling of cytoskeletal rearrangements in the endothelial cells. To prevent the endothelial cells from cellular contraction via reductions in hyperphosphorylation known to be essential for several models of agonist-induced pulmonary edema and ventilator-induced lung injury might facilitate restoration following endothelial barrier dysfunctions. The permeability of epithelial surfaces has been studied more thoroughly than that of endothelial surfaces, and several studies have suggested a role of cytoskeleton in regulating epithelial permeability. We hypothesized that the cytoskeleton might also contribute to regulation of endothelial permeability. Alterations of the endothelial cell cytoskeleton under hypoxia may contribute to changes in pulmonary vascular permeability and thereby leads to HAPE. Many studies have been reported that hypoxia disrupts the endothelial barrier by creating gaps in cell- cell junctions that causes infiltration of blood proteins and cells into vessel wall. Few biophysical studies also proved that under hypoxia cytoskeletal changes occur that cause this remodeling of endothelium. Nitric oxide plays an important role in maintaining endothelial cell morphology. Our study has demonstrated that hypoxia attenuates nitric oxide levels that make the endothelium leaky [1]. In pathological conditions like HAPE, the transvascular leakage occurs due to low bioavailability of nitric oxide in endothelium. We deem that membrane permeability, cytoskeletal re-arrangements and vascular leakage under hypoxia are associated phenomena. Our work improvises the concept that nitric oxide aids to resist hypoxia induced endothelial leakage, which may lead to the development of therapeutic strategies for minimizing vascular leakage in various conditions under hypoxia.

Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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Chapter I

Actin Dynamics and Remodeling of Cell-Cell Junctions in Epithelial Morphogenesis Marc D.H. Hansen and Michael R. Stark Department of Physiology and Developmental Biology Brigham Young University 574 WIDB Provo, UT, 84602, U.S.A.

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Abstract Cell-cell adhesion systems play a critical role in integrating individual cells into tissues. Crucial to cell-cell adhesion is the structure and dynamics of the actin cytoskeleton, particularly at cell-cell junctions. Such organization forms the basis of proper cell and tissue morphology. Control of actin dynamics at adhesion sites allows epithelial tissues to undergo dramatic remodeling events. These epithelial morphogenesis events occur throughout development. The staggering variety of cell and tissue architecture changes during epithelial morphogenesis has resulted in events often being categorized by specific types of cellular rearrangements, with little regard to understanding epithelial morphogenesis in a unified manner. All morphogenetic rearrangements rely on movements and shape changes made by individual cells within the tissue. When these movement and shape changes occur collectively, they result in repositioning of individual cells within the tissue or an overall deformation of the tissue. The unfiying feature of such events is that cell-cell adhesion are maintained, allowing individual cell morphology changes to alter the overall tissue architecture. Thus, actin dynamics in individual cells are coordinated thorough cell-cell contacts to drive changes in morphology at the tissue level. Epithelial morphogenesis also includes tissue remodeling events where cell shape changes and movement occur in isolation and do not impact the entire tissue, allowing cells to behave in a solitary fashion. An example is epithelial-mesenchymal transition (EMT), where cells completely detach from the 

Corresponding author: 801.422.4998. [email protected]

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Marc D.H. Hansen and Michael R. Stark

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epithelial tissue and migrate to new locations. Importantly, epithelial morphogenesis events that result in individual cells detaching from the tissue highlight how cells can uncouple actin dynamics from cell-cell adhesion systems to generate completely different outcomes on tissue architecture. Given the central role of actin-membrane connections at adhesion sites in determining whether actin dynamics of individual cells occur collectively or independently, it is important that the molecular basis of membrane connections at adhesion sites has been under recently renewed scrutiny. The prevailing dogma that cadherin-based adhesions are directly anchored to actin filaments, thus providing the majority of functional actin-membrane linkages at adhesion sites, has been challenged and the precise molecular organization of actin-membrane connections of cell-cell adhesions is under active revision. Actin-membrane anchor sites at epithelial cell-cell junctions are typically symmetrical, with actin membrane linkages occurring on both sides of the adhesion site. A benefit of such organization it that symmetrical connection points allow the actin cytoskeleton of individual cells to be integrated into a larger multi-cellular actin network, connected through points of cell-cell adhesion. This network can be regulated at the level of the whole tissue to drive collective epithelial morphogenesis events. From the perspective of actin dynamics and actin-membrane connections, the entire variety of epithelial morphogenesis events can be viewed as a spectrum of outcomes that result from changes in actin organization and contractility, as well as whether these changes are coordinated through actin-membrane connections with cell-cell adhesion systems. In this view, the extent, order, and location of changes in the actin cytoskeleton in each cell constrain the outcome of epithelial morphogenesis in terms of tissue architecture. A detailed understanding of how actin reorganization and contractility are coordinated through cell-cell contacts of epithelial cells will be critical in developing such a unified view of epithelial morphogenesis events. A major goal of cell biologists has been to characterize actin dynamics in live cells and identify how specific molecular machines generate those actin dynamic events. A major emphasis of developmental biologists has been to understand how signaling networks are regulated in time and space to drive specific developmental events. This chapter will focus on how spatiotemporal signaling and actin rearrangements of cell-cell adhesions interface during development to generate specific changes in tissue morphology.

Introduction Compartmentalization and specialization of physiological functions within organs and tissues requires coordination of individual cells. While coordination of specific cellular activities plays a major role in tissue function, coordination of the position and morphology of cells within a tissue has important repercussions on establishment and maintenance of tissue architecture that are equally important to function. Proper tissue function cannot be achieved without proper form; it is the interplay between structure and function in tissues that generates normal tissue homeostasis. Interplay between actin organization and cell adhesion systems is a major component required to provide the framework for organizing individual cells into epithelial tissue structures. It is cell-cell adhesion systems in particular that integrate individual cells into tissues that exhibit a collective function that exceeds that of the individual cells composing the tissue [1]. Importantly, cell-cell junctions are highly dynamic [2]. Cell-cell adhesions undergo constant dissolution and renewal in order for the position and morphology of cells within tissues to be altered. In fact, the constant remodeling of cell-cell adhesions is thought to be

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Actin and Adhesion Dynamics in Morphogenesis

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required for normal tissue maintenance of many epithelial tissue types. Remodeling events become even more striking during epithelial morphogenesis events in development, where individual cellular movements are carefully choreographed to drive deliberate and dramatic changes in overall tissue architecture. This choreography depends greatly on cell-cell adhesion systems [3]. The status of cell-cell adhesions determines whether individual cellular movements and shape changes are coordinated within the tissue to generate a collective effect or whether individual cells will act independently to produce isolated, solitary cells that act apart from the tissue. Clearly the assembly and breakdown of cell-cell junctions must be carefully controlled. While dramatic, examples of wholesale cell-cell junction assembly or detachment are, in fact, more simple to explain than instances where collective movement of individual cells within a tissue are used to achieve a larger change in epithelial tissue architecture. Here, as before, it is actin reorganization and contractility that generate morphological changes in individual cells [4]. Connection of the actin cytoskeleton with cell-cell adhesion systems coordinates these individual movements to generate tissue-wide effects [5]. Spatiotemporal regulation of actin organization and contractility programs within each cell can have dramatic effects on the types of cell shape and movement changes that, when coordinated through points of cell-cell adhesion, alter epithelial tissue architecture in different ways [4-6]. The diversity of epithelial morphogenesis programs that are observed in development supports this. In this chapter we will explore how actin rearrangements and their connections to cellcell junctions drive the variety of epithelial tissue remodeling events. Our overall hypothesis is that actin reorganization and contractility events drive shape and motility changes in individual cells and that regulating connections between the actin cytoskeleton and cell-cell adhesion systems, thus linking the actin cytoskeleton of neighboring cells, allows these shape and motility changes to be either coordinated or independent (Figure 1). In collective epithelial morphogenesis events, cell-cell adhesions remain connected to the actin cytoskeleton. Thus, the effects of actin reorganization and contractility are transduced through the tissue. Actin reorganization and contractility are separately and locally regulated, allowing each to occur independently in time and space to generate the diversity of morphogenetic outcomes observed throughout development. In epithelial morphogenesis events that give rise to solitary cells detaching from the epithelium, connections between actin and cell-cell adhesion systems are disrupted. This prevents effects of actin reorganization and contractile forces from being applied throughout the tissue. The result is isolated morphological changes of shape and movement in solitary cells. This chapter will be devoted towards reviewing our understanding of actin organization and contractility during epithelial morphogenesis, with a particular focus on connections to cell-cell adhesion systems. Importantly, spatiotemporal regulation of epithelial morphogenesis requires an understanding of developmental signaling programs that provide this timing and location or direction information to cells and tissues. Thus, control of epithelial morphogenesis programs by developmental signaling networks, as well as their interface with actin dynamics and cell-cell adhesion, will also be discussed. The overall goal is to develop a unified model for how the diversity of tissue architecture changes that arise from epithelial morphogenesis programs can be explained by changes in actin dynamics.

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Figure 1. A model for how actin dynamics drive epithelial morphogenesis. Actin organization as cells transition from a stable epithelium to a dynamic epithelium undergoing collective morphogenesis and then to generation of solitary cells, with a view of the plane (left) or cross section (right) of the epithelium.

Actin Structure at Cell-Cell Junctions Actin associated with epithelial cell-cell contacts is arranged in one of two general organizations: in a cortical ring with actin filaments running roughly parallel to cell-cell junctions and circumscribing the cell or a medial network consisting of cables that span the cell and connect with cell-cell junctions at approximately perpendicular angles. The cortical actin ring is the most common organization in epithelial cells. Medial actin networks are more commonly observed in epithelial cell undergoing dynamic tissue remodeling. Medial actin networks are more visible by light microscopy, probably accounting for why actin networks were initially observed only in dynamic, and not stable, epithelia [7]. Importantly, these two organizations play a major role in epithelial morphogenesis, with transition of overall actin structure between these organization subtypes impacting how tissue architecture changes.

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Figure 2. The cortical actin ring network. Actin organization of epithelial cells with (left) and without (right) cell-cell junctions is depicted.

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Cortical Actin Ring Structures in Epithelial Cells A general actin organization consisting of a circumferential cortical ring of actin occurs in numerous cell types and is particularly visible when cells are cultured on two dimensional surfaces ( 2) [8, 9]. Cell-cell adhesion dramatically alters the organization of the cortical actin ring [10]. In solitary epithelial cells that are not participating in cell-cell adhesion, a cortical ring of very loosely bundled actin filaments circumscribes the cell. Associations of actin with the plasma membrane at the outermost cell periphery are not pronounced. Interestingly, assembly of actin structures that drives protrusion of the membrane during cell migration often result in a local disassembly of the cortical actin ring, demonstrating that sections of the ring can be disassembled without affecting the entire cortical ring structure. In epithelial cells with established cell-cell contacts, cells are also circumscribed by a ring of actin. In contrast to solitary cells, the cortical actin ring in cells with cell-cell contacts is tightly bundled so that individual filaments are often hard to resolve. Further, the cortical actin is immediately juxtaposed against cell-cell contacts, to a degree that rings of adjacent, contacting cells often cannot be resolves independently. Thus, the cortical actin ring closely follows the irregular outline of the cell periphery. Despite this tight abutment against cell-cell junctions, the actin of the circumferential cortical ring is highly dynamic, with a very fast rate of turnover that indicates active polymerization and treadmilling within this actin structure [11]. The nature of membrane anchoring of circumferential actin at cadherin-based adhesions of epithelial cells remains unclear, as will be discussed at the molecular level in the next section. Despite several proposed or described series of molecular interactions have would link the membrane of the cell-cell junctions to actin filaments, it remains unknown whether actin binding occurs at the end or sides of actin filaments. The difference in location of actin filament binding sets important constraints on how the overall cortical actin ring structure is organized in epithelial cells and tissues. Anchoring by adaptor proteins that bind the ends of actin filaments suggests that most actin-membrane connections occur at the vertices, where multiple cells make contact. In contrast, anchoring via proteins that bind the sides of actin filaments would suggest that actin filaments associate with the membrane the entire length of cell-cell junctions.

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Actin-membrane linkages at the ends of actin filaments would suggest that cells with established cell-cell junctions would only integrate their actin networks at discreet points. Since actin filaments of the cortical actin ring of contacting epithelial cells terminate at vertices [10, 12], it is primarily at the vertices where integration of actin networks from contacting cells would occur. In this model, the lengths of actin filaments connecting vertices would lie immediately against the intervening junctions between two adjacent cells, but with few direct physical interactions with the membrane. This higher order arrangement of individual actin filaments likely includes bundling of individual actin filaments into cables. Further, since anchoring is expected to occur via barbed ends and since these cables would be anchored at vertices at either end, cables must contain individual actin filaments with an antiparallel orientation or, like stress fibers, consist of shorter, repeating structural units with opposing cytoskeletal polarity. Either arrangement facilitates the action of myosin-based contractility that could generate tension along actin cables that connect neighboring vertices. This could ensure maintenance of strong cell-cell adhesions within a sheet of epithelial cells, even if cadherin-based adhesions at sites between vertices are not associated with the underlying actin cytoskeletal system. Importantly, this organization of actin easily accounts for cell-cell junction remodeling events observed in several specific morphogenetic processes, most particularly convergent extension. In our view, organization of the actin cytoskeleton at cell-cell junctions is most likely to follow this architecture, using actin-membrane anchoring proteins that interact with the ends of actin filaments. Anchoring of actin filaments to membranes via actin binding proteins that interact with the sides of actin filaments would easily explain how the circumferential cortical actin ring organization is achieved. Each actin filaments could be linked to cell-cell junctions by adaptor proteins along their entire lengths. Such binding would facilitate orienting of actin filament to run parallel to the membrane. Such interactions would also easily explain the tight proximity of actin filaments with cell-cell junction membranes observed in cells in cell-cell contact with their neighbors. Importantly, however, how reorganization of such an actin organization could lead to cell-cell junction remodeling events that are observed during tissue remodeling is far less clear than for a model where actin-membrane linkages occur at the ends of actin filaments. Further, actin-membrane linkages thorough the sides of actin filaments would be fundamentally different from those appearing in medial actin networks, whereas actinmembrane anchoring at the ends of actin filaments could account for cell-cell junction associations of both the cortical actin ring and medial actin network structures.

Medial Actin Networks Associated with Cell-Cell Junctions Medial actin networks consist of actin networks with the ends of actin cables is apparent association with cell-cell contacts. Despite controversy regarding the nature of such molecular connections, medial actin networks can demonstrably transmit contractile forces to cell-cell adhesions [13-15]. Thus, connections of actin to cell-cell adhesions must be robust enough to withstand force transmission. The most striking example of such robust connections of the ends of actin filaments to cell-cell junctions is observed in non-epithelial cells, namely at the intercalated discs of cardiomyocytes. Here a regularly and highly ordered sarcomeric actin structure, consisting of characteristic arrangement of parallel and aligned actin filaments, terminates at a perpendicular angle at cell-cell junctions. The intercalated disc is a specialized

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cell-cell junction mediated by N-cadherin adhesion receptors [16, 17], though also enriched in specific integrins and gap junction proteins [18]. Sarcomeric contractions in cardiomyocytes are directly focused on the intercalated disc; cardiomyocyte contraction is designed to shorten the cell and pull the intercalated discs at opposite ends of each cardiomyocyte towards each other. Within cardiac tissue, the concerted action of cardiomyocytes, contracting in near unison and with intercalated discs connecting many individual cardiomyocytes into a larger tissue, generates large scale tissue contractions that result in the beating function of the heart itself. Thus, the intercalated disc uniquely demonstrates the requirement for strong, robust actin-membrane associations at cell-cell junctions; it has been proposed that dilated cardiomyopathies are a result of perturbation in the integrity of the intercalated disc [19]. Individual cardiomyocytes work as a coordinated tissue when the sarcomeric actin organization of each individual cell is integrated into a larger, multi-cellular actin structure. It is the intercalated disc, containing cell-cell adhesion receptors, that integrates the actin structure of each individual cell across cell-cell adhesions [20]. In fact, since slight perturbations in the integrity of the intercalated disc generate such drastic cardiac phenotypes, the intercalated disc is a powerful model system that is uniquely suited to dissecting the molecular architecture of cell-cell junctions. Even only a small loss of force transduction across the membrane results in hypertrophy and, eventually, heart failure. This is illustrated in mice with tissue specific deletions in genes encoding cadherin complex component proteins that make up the intercalated disc. Function blocking N-cadherin antibodies or misexpression of an N-cadherin fragment result in loss of cardiomyocyte cellcell adhesions and in perturbations in alignment of sarcomeric actin [21, 22]. Cardiac tissuespecific deletion of the gene encoding N-cadherin results in dissolution of the intercalated disc, a result of the failure of individual cardiomyocytes to integrate into a function cardiac tissue by aligning sarcomeric actin structures of individual cells across cell-cell junctions [23]. At the organismal level, the result is moderate dilated cardiomyopathy and death. Death is thought to result from arrhythmia, a result of loss of coordinated contraction within cardiac tissue, which requires Gap junctions that are concentrated in normal intercalated discs [24]. Interestingly, an increase in expression of specific integrins is also observed, suggesting that cardiomyocytes may attempt to compensate for loss of cadherin expression with increased integrin levels. That this indicates integrins play a role in connecting cadherins to the actin cytoskeleton at adherens junctions remains untested in this system, but is a possible mechanism for linking cell-cell adhesions to actin filaments. Importantly, the cardiac-specific N-cadherin null phenotype can be reversed, at least partially, by exogenous expression of Nor E-cadherin [25]. Animals exogenously expressing N- or E-cadherin in a cardiac muscle specific N-cadherin null background do develop a dilated cardiomyopathy, though without arrhythmia [26]. Those expressing E-cadherin exhibit an earlier onset of the cardiomyopathy phenotype, possibly a result of cadherin-specific differences in recruitment of proteins to the adherens junction, most notably vinculin, which is recruited to the intercalated disc by Ncadherin, but not E-cadherin [26]. Genetic elimination of adaptor proteins that have been proposed to link cadherins to cytoskeletal elements also generates cardiomyopathy phenotypes. Cardiomyocytes express two α -catenin isoforms, αE- and αT-catenin [27]. However, targeted genetic perturbation of only αE-catenin in the heart results in the identical phenotype to the cardiac tissue-specific Ncadherin null [28]. Surprisingly, loss of β-catenin has no effect on heart function or on the macroscopic or microscopic organization of the intercalated disc [29, 30], but increased β-

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catenin levels by expression of a stabilized mutant did generate a heart defect [29]. Intercalated disc proteins are still localize properly, though this is associated with an increase in plakoglobin targeting to the intercalated disc, which may compensate for the loss of βcatenin [30]. Supporting a role for plakoglobin at the intercalated disc, a cardiomyopathy phenotype is observed in humans homozygous for a recessive mutant allele of the gene encoding plakoglobin [31]. It is possible that the role of plakoglobin demonstrates the particularly important function of desmosomes, points of cell-cell adhesion that are linked to intermediate filament systems [32], in mediating strong adhesion at the intercalated disc. Consistent with desmosomes acting as points of strong cell-cell adhesion, deletion of Ncadherin from cardiomyocytes does result in a loss of desmosomes from cell-cell adherens junctions of cardiomyocytes [21]. Despite a possible role for desmosomes at the intercalated disc, it is widely accepted that actin-membrane connections must connect the actin networks of adjacent cardiomyocytes at the intercalated disc. In support of this idea, several actin binding proteins that localize to both cell-cell junctions of epithelial cells and the intercalated disc of cardiomyocytes are required for normal intercalated disc function. These include the VASP relative Mena [33] and vinculin [34]. That loss, or even expression level changes, of adherens junction proteins results in cardiomyopathy phenotypes makes the intercalated disc a powerful, and dramatically underutilized, model system for dissecting the molecular architecture of actin-membrane connections at cell-cell junctions. While this model system has the major drawback of lacking for a tissue culture model system that successfully recapitulates the intercalated disc structure in vitro, application of genetic strategies to exacerbate or rescue subtle heart phenotypes arising from genetic perturbation of cadherin component proteins could be used to identify additional components of cell-cell adhesion system, including important actin regulatory systems. Epithelial cells display much smaller bundles of actin filaments that terminate obliquely at points of cadherin-mediated adhesion in the medial actin network structure. As with the intercalated disc, points of cell-cell adhesion are typically symmetrical in their organization, with cables of actin filaments from medial actin networks emanating from the point of adhesion on both sides of the junction [10, 15, 35, 36]. Though actin bundles of medial actin networks terminate at spots of cadherin-based adhesion are not organized into a sarcomeric structure, they are typically decorated with myosin and generate contractile forces, as the membrane of the cell-cell junction is typically deformed at areas where such structures are detected [15]. Transduction of myosin-based contractile forces to cell-cell junction is clear evidence that actin must make physical connections with membranes at adhesion sites. Interestingly, proteins localizing to these actin-membrane connections are also those that are observed at the intercalated disc: zyxin, VASP, vinculin, and cadherin-catenin complex components [10, 15, 35, 36]. Experimental perturbation of proteins required for normal intercalated disc function in epithelial cells generates cell-cell adhesion phenotypes, further demonstrating how the intercalated disc appears to be closely related to epithelial actin-membrane connections.

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Actin Dynamics and Contractility in Cell-Cell Junction Remodleing Cell-cell adhesions are often imagined to be highly static structures, simply acting to maintain rigid contacts between dormant cells and reinforcing tissues against mechanical disruption. Similarly, actin is often thought of as a structural element that simply underlies cell-cell adhesions in the manner of a static scaffold network system. Contrary to this view, epithelial tissues are highly plastic [2]. Cells rearrange their position within the tissue despite maintaining constant cell-cell interactions. Individual cells of the epidermis are replenished by cell division at the basement membrane, but these cells slowly move through the stratifications of the epidermis to the surface until being sloughed off. In the colon epithelium, individual cells are also replenished by cell division. This occurs at the base of the crypt and the resulting cells then migrate out of the crypt within the plane of the epithelium. In both cases, one can envision collective movement of cells within the tissue in a manner whereby cell-cell adhesions are constantly maintained: new cells are generated, old cells are lost from the tissue, and cell-cell contacts between cells require little remodeling as cells move through the tissue. More complicated is the example of the transitional epithelium, where cells rapidly convert between stratified and simple epithelium and where cell-cell junctions must be formed and broken during this conversion. An added level of complexity occurs in epithelial morphogenesis events, where developing tissues undergo highly dramatic rearrangements that illustrate how cell-cell junctions must be highly plastic. In this section the association of actin reorganization and contractility with cell-cell junction remodeling events will be examined, with a focus on studies using cell model systems.

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Actin Dynamics in Establishment of Cell-Cell Adhesions Rearrangements of the actin cytoskeleton during formation of epithelial cell-cell junctions have been characterized in several tissue culture model systems [10, 36-38]. At the morphological level, initiation of cadherin-based cell-cell adhesion is associated with a dramatic increase in membrane protrusions, particularly lamellipodia, that occur at or near the site of cell-cell contact [10, 36, 39]. This is followed by a compaction of the cell towards sites of cell-cell adhesions. When cells are placed in culture at low density, such that a single junction between two cells is established, the result is a net movement of one cell towards its newly contacting neighbor [10]. In cells placed in culture at higher density, where cell-cell adhesions are initiated at many points simultaneously, compaction of cells generates no net movement and arrays of retraction fibers (“adhesion zippers”) are observed between adjacent cells that pull a short distance apart during compaction [36]. Retraction fibers result from cells pulling apart except at distinct points of reinforced cell-cell adhesion, as illustrated by the appearance of cell-cell adhesion the remaining point of connection at retraction fiber tips [36]. Actin rearrangements that form the basis of these cell morphology changes have also been characterized at a high level of detail, particularly in instances where two cells collide and establish a single cell-cell junction (Figure 3) [10, 12, 40]. Here initial contact occurs at a small point between two cells.

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Figure 3. Actin organization during formation of cell-cell junctions between cells in culture.

Initially, these cells exhibit a loosely arranged cortical actin ring. At the point of first cellcell contact, cadherins accumulate into spot adhesions that become associated with radial actin filaments that abut the point of adhesion at a perpendicular angle. Additional points of cadherin-based adhesion continue to form as the nascent contact laterally expands. These newer points of adhesion are first formed within the center of the expanding contact, while older points of adhesion move laterally towards the ends of the contact and aggregate into a larger plaque. While points of cadherin-based adhesion are appearing, the cortical actin ring is disassembled along the length of the cell-cell contact. This may be associated with the formation of actin-based membrane protrusions that are associated with initial cell-cell contact. The result is that the cortical actin ring now becomes an arc. The ends of the cortical arc of actin terminate at either end of the expanding junction, appearing to make a tight physical connection with the larger plaques of cadherin-based adhesion that occur there. With this overall actin organization achieved, compaction of the cells towards each other begins in earnest, an event that tightly coincides with the apparent contraction of the cortical arc of actin in the abutted cells. The arc becomes smaller in circumference and the individual actin filaments become more densely packed. Eventually a network of actin that tightly associates with the length of the fully expanded cell-cell junction is assembled, though the detailed structure of this network has not been described at high resolution. The coordinated lateral mobility of points of cadherin-based adhesion strongly suggests that actin makes physical connections with cadherin-based adhesion systems in the plasma membrane in some way. It is presumably forces resulting from actomyosin-based contractility in the cortical actin system [41] that are laterally mobilizing spots of cadherin in the membrane and concentrating them into dense plaques at the ends of the nascent cell-cell adhesion (Figure 4). In fact, such actin-membrane connections could well explain how actomyosin-based contractility might drive cell-cell adhesion. With actin membrane connections at dense plaques of cadherin-based adhesion at either end of the cell-cell junction, contraction within the cortical arc of actin would result in a lateral pulling force that would expand the cadherinbased adhesion. Further, the plaque of cadherin would move laterally, remaining at the boundary between the contacting and non-contacting membrane as cell-cell junction expansion proceeded. Further, the lateral movement of new spots of cadherin-based adhesion, with their radial bundles of actin filaments, could be explained simply by the eventual incorporation of the radial actin filaments into the larger cortical actin network.

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Figure 4. A model for actin contractility in cell-cell junction formation. Arrows indicate the direction of contractile forces arising out of myosin activity in the cortical actin arc.

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Actin Dynamics in Disassembly of Cell-Cell Adhesions Detachment of epithelial cell-cell junctions in response to growth factor-induced scattering similarly reveals clues about the nature of actin organization at cell-cell contacts during tissue rearrangements [15, 42-45]. Such studies take advantage of tissue culture model systems, where cells can be stimulated with growth factor to undergo scattering in a synchronized manner throughout the culture [46]. Scattering is thought to mimic epithelialmesenchymal transition (EMT) events observed in development, and occurs when cadherinbased adhesion between epithelial cells are disrupted and cells become both solitary (lacking cell-cell interactions) and highly migratory. Prior to scattering, epithelial cells exhibit a cortical actin ring structure that is tightly associated with cell-cell junctions, as is typical of stable epithelial cells [43]. Following scattering, the cortical ring is more loosely arranged and stress fibers are more evident [43]. Cell morphology changes are dramatic during scattering [15, 44, 45]. Cells undergo a striking spreading phase, where the substrate surface area covered by individual cells roughly doubles. Cell migration is also strikingly increased, though early in scattering all motility is collective; cell migration occurs at the tissue level and with maintenance of cell-cell junctions. Gradually, an increase in cell-cell detachment events is observed until cells maintain cell-cell interactions solely through small, punctate spots of adhesion, usually at the end of retraction fibers. Finally even these recalcitrant points of robust adhesion are ruptured and cells can now migrate in true solitary fashion and without cell-cell contacts. Live fluorescent imaging also reveals how the actin cytoskeleton is reorganized as cells undergo scattering (Figure 5) [15]. Initially, the cortical actin that is abutted against the cellcell junction resolves into multiple smaller actin cables that move toward the cell interior. At cell vertices, where these small actin cables terminate, there is a burst of actin polymerization. The resulting spot of actin at the vertex fragments into smaller puncta, each at the end of a smaller actin cable and with lateral mobility that results in the lateral displacement of puncta away from the vertex. Laterally mobile spots of actin at cell-cell junctions and their connections to the ends of actin cables are symmetrical, meaning that actin cables are

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connected to these puncta on both sides of the junction as angular actin structures described in the previous section.

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Figure 5. Actin reorganization during disruption of cell-cell adhesions between epithelial cells in culture.

The overall result of this reorganization event is a rapid transition from a cortical ring arrangement to a medial actin network based on angular actin membrane connections. The symmetric organization of actin membrane connections at points of cell-cell adhesion results in the integration of actin networks of individual cells into a larger multi-cellular actin network shared by cells that still maintain cell-cell contacts. Further progression of scattering is associated with the disruption of actin-membrane connections at points of cell-cell adhesion at linkages between angular actin cables of adjacent cells.

Actin Regulatory Proteins Driving Epithelial Morphogenesis in Cell Model Systems The recruitment or redistribution of actin regulatory proteins at cell-cell contacts during cell-cell junction formation or detachment provides important clues as to how cell-cell adhesion and actin dynamics are coordinated at the molecular level. A number of regulators of actin dynamics, including proteins or protein complexes that bind actin filaments directly, have been localized to cell-cell junctions and implicated in cell-cell junction formation or maintenance. In our model for how actin is reorganized to contribute to tissue remodeling, it is proposed that actin organization, contractility, and membrane linkages are separately

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regulated to generate a variety of tissue remodeling events. Thus, it is predicted that actin regulatory systems that participate in actin organization, contractility, and membrane linkages will be separately regulated. Here we will discuss how actin regulatory proteins associated with these three major roles are reorganized during cell-cell junction assembly and disassembly in tissue culture model systems. Actin rearrangements during disruption of cell-cell adhesions highlight the relatedness of cortical actin of stable cell-cell junctions and radial actin networks of dynamic epithelial cells [15]. Stable epithelial tissues feature tight abutment of actin filaments to cell-cell contacts, where a number of actin bundling proteins localize. In our view, the cortical actin arrangement of stable epithelial tissues, where actin cables run parallel to the junction and are anchored at cell vertices, is maintained in part by actin bundling proteins. These serve to stabilize parallel and anti-parallel actin filaments into cables. Several actin bundling proteins localize specifically to cell-cell junctions and are dynamically recruited or dispersed in cellcell junction formation or disassembly, respectively. One well-characterized actin binding protein that has long been recognized to localize to cell-cell contacts is α-actinin [47]. In some experiments, α-actinin and α-catenin have been shown to interact directly [47], though this interaction has failed to account for the anchoring of cadherin complexes to actin filaments [11, 48]. α-Actinin also functions as part of the adaptor system that links integrinbased adhesions to actin filaments [49], and could serve a role at cell-cell junctions by mediating integrin-actin connections. More likely is the overall role of α-actinin in crosslinking actin filaments into larger cables, such as those that run parallel to cell-cell junctions. Another protein with actin bundling activity, VASP, also localizes to cell-cell junctions in a dynamic manner and does so in a manner that is similar to that of α-actinin [35, 36, 50]. Also like α-actinin, VASP has also been proposed to link actin to membranes. VASP maintains actin-membrane connections by binding molecules that bear proline rich FPPPPP repeats [51]. Such molecules include adaptor proteins, such as zyxin [52], and even adhesion receptors, like the non-classical FAT cadherin [53]. Both α-actinin and VASP are recruited to cell-cell junctions upon initiation of cell-cell contact [35, 36, 54] and are rapidly released from cell-cell contacts upon initiation of cell-cell junction remodeling [15]. Interestingly, αactinin and VASP are released from cell-cell contacts as cells initiate programs of cell-cell detachment, with a timing that immediately precedes dissolution of larger actin cables that are directly abutted against cell-cell contacts into smaller actin cables [15]. This release of individual actin filaments that are tightly associated with cell-cell junctions of stable epithelial cells specifically results in the formation of medial actin networks [15]. The timing of αactinin and VASP release from cell-cell contacts in tissue remodeling supports the idea that the bundling activity of these proteins may be required to maintain actin architecture at cellcell adhesions of stable epithelial tissues and that this activity must be released to allow changes in actin dynamics required for junction and tissue remodeling. Clearly other actin regulatory systems alter the overall architecture of actin as cells remodel cell-cell adhesions, though the identity of such factors and their precise role remains far less obvious than for actin bundling and crosslinking systems. Physical reorganization of individual cells within tissues requires force generation from the actin cytoskeleton. While formation of actin-based membrane protrusions rely directly on actin polymerization, most other cell shape changes are driven by actin-based contractility through myosin motor systems. Unconventional myosin motors, such as myosin VI [55], have been directly implicated in cell-cell adhesion, though the precise roles of these motors remain

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unclear. Better understood is the role of the conventional non-muscle myosin, myosin II, in generating actin-based tension forces in dynamic epithelial cells. Myosin II acts by generating tension forces in parallel arrays of actin filaments. Importantly, the role of certain isoforms of myosin II in cell-cell adhesion appears to occur independently of their motor activity [56], suggesting that some myosins might serve only as actin binding proteins in some instances. Myosin II displays a dynamic association with actin cables in epithelial cells, its appearance not surprisingly coinciding with the generation of tension forces. In cell-cell junction assembly, it has been proposed that myosin II generates the contractility that shortens the cortical actin ring in regions outside of the nascent cell-cell contact, thus driving cells towards their cell-cell contact as the cortical actin ring shrinks. During the early stages of cell-cell junction disassembly of MDCK cells treated with scatter factor in culture, myosin II is rapidly recruited to individual actin filaments that make up medial actin networks and tension forces from this network are focused onto their attachment sites at cell-cell adhesions [15]. Areas where myosin-decorated medial actin network connects with the cell-cell junction show deformations resulting from the transduction of actin-based tension forces. Findings in this model system are recapitulated in developmental processes that highlight how myosin function can generate different tissue remodeling outcomes depending on cellular actin organization and its connections to cell-cell junctions (Figure 6).

Figure 6. Myosin contractility on epithelial tissue architecture. Actin contractility generates different effects on epithelial tissue architecture if contractility follows reorganization of the cortical actin ring into a medial actin network (A) or occurs in cells that maintain the cortical actin ring structure of stable epithelial cells (B).

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Most comparable to observations in tissue culture system, medial actin networks are assembled near the apical surface of epithelial cells during Drosophila development and pulses of myosin II recruitment result in the generation of tension forces that are applied to the apical region cell-cell junction [13]. The overall effect here is on the three-dimensional shape of individual cells in the tissue, specifically converting cylindrically-shaped cells to conical shaped cells to drive bending of the overall tissue [13]. Tissue elongation in the same model organism occurs by a similar mechanism, but it is myosin-dependent contractility at the basal region of individual cells that generates tissue-wide effects [57]. During convergence-extension and intercalation events, where cells change relative position in order to alter the dimensions of the epithelium in two dimensions, myosin II is also recruited to actin and generates tension that is thought to be transduced upon and through connections with cell-cell junctions [2]. Though there is some controversy about how actin organization and contractility are coordinated to generate forces that shorten specific cell-cell junctions [14], the prevailing thinking is that actin retains the cortical actin ring organization of a stable epithelium, with actin tightly against each cell-cell junction and terminating in connections at vertices. The effect of contraction of the cortical actin in such cells is tension forces that bring adjacent vertices together. Maintenance of actin-membrane connections at sites of cell-cell adhesion plays a major role in determining how actin reorganization and contractility affect overall tissue architecture during epithelial morphogenesis. The molecular architecture of actin-membrane connections at cell-cell contacts are detailed in the next section, but a requisite feature of such connections in terms of tissue remodeling is their reversibility. When cells are integrated into the issue via cell-cell adhesions, actin-membrane connections are maintained. In contrast, the ability of cells to detach from their neighbors and use actin dynamics and contractility in a solitary fashion requires the disruption of actin-membrane connections at points of cell-cell adhesion.

The Molecular Basis of Collective Cellular Behaviors: Cell-Cell Adhesion Systems and Their Actin-Membrane Connections Given the fundamental role of adhesion systems in maintaining proper tissue architecture and function, defining the molecular organization of actin linkages with adhesion systems and their regulation has been a major endeavor of cell biologists. Adhesion systems of all types, both cell-cell and cell-substrate adhesions, appear to follow a general organization paradigm. In this paradigm, adhesion receptors become anchored to the underlying cytoskeleton, thus providing strength to the adhesion system (Figure 7). This paradigm has remained the dominant central hypothesis among cell biologists for roughly two decades and has been strongly applied to cell-cell adhesion systems. Interactions of adhesion systems with cytoskeletal elements is thought to provide constraints on the overall cellular cytoskeletal organization, generating coordinated cellular movements and connections that form the basis of the simplest tissues. When actin connections with adhesion systems are severed, collective cell behavior is abandoned for solitary cell behaviors.

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Figure 7. A paradigm for molecular organization of cell adhesion systems and their linkage to cytoskeletal elements.

In favor of a model for a direct linkage between adhesion systems and cytoskeletal systems, such as actin, is the need to integrate the regulation of individual cytoskeletal rearrangements as a multi-cellular network that can be collectively controlled. When actin networks of individual cells are connected through cell-cell adhesions, the forces of individual cell movements are instantly and fully transduced through the tissue. Both adult and developing tissues use cell-cell connections to transduce forces across cell-cell boundaries and properly distribute them within the tissue, the example of cardiac tissue having already been discussed. While the consequences of coordinated movements can vary, depending on the arrangement of both the adhesion points and the cytoskeleton, clearly actinadhesion connection must be robust enough to withstand significant forces focused on specific adhesion sites. Cell-cell adhesion receptors are transmembrane proteins of the cell surface whose extracellular domains bind proteins on the surface of contacting cells, thus resulting in ap lsama membrane that is sticky to the surfaces other cells. One major classification criteria for cell-cell adhesion receptor systems is whether they are homotypic or heterotypic in their trans interactions, meaning whether an adhesion receptor binds to cognate receptors of the same or different type on adjacent cells. Homotypic adhesion receptors mediate interactions of the same cell type within tissues and play a critical role in aggregating cells of the same type to form a primordial tissue during development. Heterotypic adhesion receptor systems allow cells of different types to interact, either within tissues or between tissues. Another major

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criterion for classification of cell-cell adhesion receptor systems is by the number of transmembrane domains. Most cell-cell adhesion receptors, and even adhesion receptors generally, contain a single transmembrane domain, but several adhesion receptor systems span the membrane several times. This classification is important because single transmembrane domain receptors, in isolation, cannot transduce extracellular binding events through the membrane to activate the receptor‟s cytoplasmic tail, whereas receptors with multiple transmembrane domains can convert extracellular binding to an intracellular conformational change that can alter protein function or activity. Importantly, current thinking is that adhesion systems work according to a general paradigm, by which adhesion receptors bind to either cognate receptors on adjacent cells or to non-cellular connective tissue (extracellular matrix) via their external domains and are linked to cytoskeletal element through cytosolic adaptor proteins that bind to the intracellular domains of the adhesion receptor system (Figure 7). In this section we will describe major cell-cell adhesion receptor systems that integrate cells into tissues and describe their connection to the actin cytoskeleton at the molecular level, discussing how these different cell-cell adhesion systems might fit into the general paradigm for adhesion systems generally.

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Cadherin Adhesion Receptors Cadherins are the primary molecule responsible for cell-cell interactions in most tissue types. The cadherin family consists of several smaller subgroups, including the so-called classical cadherins. The classical cadherins subfamily contains a large number of different cadherins which show tissue specific expression [58]. In addition to mediating cell-cell adhesion in a structural capacity [59], cadherins have been shown to participate in cellular signaling [60, 61]. This dual role is best illustrated in development, where cadherins function in aggregating cells of similar type and with the same cadherin expression profile (the structural capacity) [62], as well as in driving specific cell fates (the signaling capacity) [63, 64]. In fact, an experimental method used for predicting the tissue expression profile of a specific cadherin is to express it in teratoma cells, which drives differentiation of the teratoma cells into specific cell types [63, 64]. From the perspective of how cadherins participate in epithelial morphogenesis, an important consideration to the cell biologist is how cadherindependent signaling might direct changes in actin structure and dynamics without making any direct associations with actin filaments. Structural mechanisms of cadherin-mediated adhesion have been well studied. Classical cadherins are single transmembrane domain containing proteins that mediate cell-cell adhesion by homotypic trans interactions occurring between cadherins on juxtaposed cells [65]. Cadherin extracellular domains contain repeats of calcium binding domains [65]. Calcium binding is thought to be required for the cadherin extracellular domain to achieve its proper three dimensional structure, rather than as a mechanism to regulate protein function. In addition to trans interactions, it has been proposed that cadherins interact in cis, forming parallel dimers on the surface of the same cell that are required prior to trans interactions [66, 67]. Such cis interactions have the advantage of allowing cadherin engagement in trans to be transduced across the membrane and result in some kind of conformation event in the intracellular portion of the cadherin molecule. Importantly, however, cis interactions appear

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only to be observed following trans interactions and when cadherin concentrations become extremely high following lateral clustering [68]. Cadherins have been proposed to work according to the classical paradigm for prototypical adhesion receptors, by binding to the underlying actin cytoskeleton through adaptor proteins (Figure 8) [69, 70]. The intracellular domains of cadherins bind to a series of intracellular adaptors known as catenins [71-73]. More specifically, cytoplasmic domains of cadherin directly associate with β–catenin and p120-catenin. While the function of p120 catenin is primarily in controlling the surface residency of cadherins by determining whether they are targeted into endocytic pathways [74], it is β–catenin that is thought to provide the interactions for anchoring of cadherins to the actin cytoskeleton. β-Catenin in turn binds to another catenin, the vinculin family member α-catenin [75]. α-Catenin directly interacts with actin filaments, as well as several other actin binding proteins, including α–actinin [47] and vinculin [76]. Thus α-catenin has been generally accepted to tether cadherin adhesion complexes to actin filaments by simultaneously binding β-catenin and either actin or other actin binding proteins. Beyond the independent observation that, in independent experiments, α-catenin binds β–catenin and actin filaments, connections of cadherin-catenin complexes to the actin cytoskeleton is supported by the observation that cells expressing a fusion protein of cadherin and the actin binding portion of α-catenin can drive strong cell-cell adhesion [77]. However, reconstituted of complexes containing purified E-cadherin, β-catenin, and α-catenin fail to bind actin filaments [11, 48]. The reason is that though α-catenin can bind both β-catenin and actin filaments, the interactions are mutually exclusive [11, 48, 78]. α-Catenin occurs in dimeric and monomeric forms, but only monomers associate with β-catenin and only dimers associate with F-actin [78]. Additional factors, including α-actinin, vinculin, or even those found in a cytosolic extract, are not sufficient to result in cadherin-catenin-actin complex formation [11, 48]. Alternative mechanisms for anchoring cadherins to the actin cytoskeleton have been proposed.

Figure 8. Molecular organization of the cadherin adhesion system in the current paradigm for adhesion systems. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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The cytoplasmic domain of cadherin also binds AnkyrinG [79], whose interaction with spectrin could provide another connection to actin filaments [80]. However, the association of cadherins with ankyrin and spectrin is thought to function primarily in vesicle trafficking of cadherins, rather than in anchoring cadherins to actin networks at adhesion sites [79]. Further, perturbation of the actin cytoskeleton using small molecules have no effect on cadherin complex component turnovers at cell-cell junctions [11], a result that suggests that there is no direct, structural connection between the cadherin complexes and actin filaments, unless these connections are extremely dynamic. These results, reproduced in multiple model systems, cast significant doubt on the widely accepted idea that cadherin complexes directly associate with actin filaments to mediate strong adhesion.

Figure 9. Alternative linkages between actin filaments and membrane at sites of cadherin-based adhesion. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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It is possible that a currently undefined mechanism links cadherin-catenin complexes to actin filaments at cell-cell junctions, for which there are several general possibilities ( 9). Factors that serve as adaptors between α-catenin and F-catenin is one possibility, though if such factors exist it is unclear why cytosol, presumably containing such a protein, cannot facilitate linkages between cadherin complexes and actin in reconstitution experiments [11]. More likely is that cadherin complexes associate in lateral membrane protein complexes with other membrane proteins that are in turn tethered to actin filaments. Cadherin has notable interactions with other adhesion systems, including the nectin adhesion receptor system [81] and tetraspanin adhesion receptor systems [82]. The connection of these other adhesion systems to the cadherin system is following sections, as are the molecular details of their anchoring to actin filaments. Generally speaking, incorporation of the core cadherin-catenin complex into a larger complex consisting of other transmembrane adhesion receptor systems that bear actin linkages would allow cadherins to become indirectly tethered to actin filaments. A similar effect would also be achieved if cadherin-catenin complexes associated into large complexes that include transmembrane proteins that are not typically associated with a role in cell-cell adhesion, but interact with adaptor proteins systems that anchor to the actin cytoskeleton and with cadherins in larger complexes, such as the spectrin-ankyrin membrane skeleton [80]. In support of such a mechanism, α-catenin has been shown to interact with spectrin [83]. A transmembrane protein that interacts with the membrane skeleton system, the Na+/K+ ATPase, has been recently implicated to play a role in cell-cell junction maintenance in cardiomyocytes [84]. While such lateral associations might explain why reconstituted cadherin-catenin complexes cannot be made to bind actin filaments using in vitro experiments [11], even an indirect association of cadherin-catenin complexes with actin, if critical to function, would be expected to make cadherin turnover properties at the membrane susceptible to small molecules that target the actin cytoskeleton, an effect that is specifically not observed [11]. More recent and somewhat more controversial thinking suggests that cadherins do not directly associate with actin filaments to mediate actinmembrane linkages at all. Instead, cadherin adhesion receptors act to orchestrate changes in local actin organization or contractility at cell-cell junction that are required for strong adhesion [78, 85]. Cadherins do act in a dual role, as structural components of the cell-cell junctions and as signaling proteins. Interestingly, there is ample evidence that different cadherins initiate different downstream signaling events. First and already mentioned, different cadherin family members drive differentiation towards different outcomes when expression is forced in teratoma cells [63, 64]. Different cadherins could drive different signaling networks by lateral association with different integrin pairs via the teraspanin web. There is also lateral association of cadherins with transmembrane signaling receptors, which could initiate downstream signaling. Consistent with this idea, E-cadherin associates with EGF receptors [86, 87], while N-cadherin associates with FGF receptors [88-90]. Interestingly, ligand-dependent signaling from receptor tyrosine kinases is reduced by Ecadherin [91], but E-cadherin can also initiate EGF receptor signaling independently of EGF ligand [92]. Further, disruption of VE-cadherin prevents productive VEGF signaling in vascular endothelial cells [93]. Importantly, suggesting that cadherins‟ connection to actin filaments occurs through cellular signaling instead of by direct associations does not deny the importance of actin membrane linkages at cell-cell junctions, only that there may not be direct association of cadherin complexes with actin. Notably, a cell-cell junction containing cadherin-catenin complexes engaged in mediating cell-cell interactions could also be rich in

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membrane proteins that bind actin. Whether these other membrane proteins are adhesion receptor systems or not, they would still provide actin-membrane connections to the junction. Possibilities include a number of non-adhesion transmembrane proteins that bind adaptors with links to the actin cytoskeleton, or even peripheral membrane proteins with direct actin interactions. Annexins are cytoplasmic peripheral membrane proteins whose biochemical activity fits this description. Annexins bind membranes containing specific phospholipids in a calcium-dependent manner [94]. Annexins I and II associate directly with actin filaments [95] and localize to cell-cell contacts of epithelial cells [96, 97], providing a possible mechanism to connect actin to sites of cell-cell adhesion in a manner that does not require direct interactions with a cell-cell adhesion system.

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Nectin Adhesion Receptors Nectin adhesion receptors are single transmembrane domain proteins that mediate cellcell adhesion by binding in trans between contacting cells [98]. The nectin family is much smaller than the cadherin family with only four nectin genes. The function of nectins in mediating cell-cell adhesion is best characterized in epithelial cells. In adhesion between epithelial cells of the same type, nectin based adhesion is observed at the extreme apical edge of cell-cell junctions (the adherens junction proper), rather than being localized along the entire region of cell-cell contacts, as is the case for cadherins. Also unlike cadherins, nectins can mediate cell-cell adhesion by forming specific heteromeric pairs [99]. As such nectins can facilitate cell-cell adhesion between cells of different types within the same tissue. The cytoplasmic regions of nectin adhesion receptors closely follow the cytoskeletal anchoring paradigm for adhesion receptor systems (Figure 10). Nectin cytoplasmic tails bind the adaptor protein afadin [100, 101], a PDZ domain containing proteins that interacts directly with actin filaments [102]. Thus, nectin-afadin-actin complexes are thought to provide actin-membrane connections at the adherens junction proper.

Figure 10. Molecular organization of the nectin adhesion system. The dark grey cytoplasmic adaptor represents afadin and the light grey cytoplasmic adaptor represents ponsin. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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In addition to the afadin-actin linkage, afadin also interacts with ponsin [103], thus providing opportunities of connecting the nectin adhesion system to other membrane proteins. Such proteins also allow nectin to participate in signaling roles. Importantly, nectin adhesion receptor complexes also associate laterally with other adhesion systems. Nectin family members and afadin both bind ZO-1 [104, 105], an important cytosolic adaptor of the tight junction (discussed in the next subsection). In fact nectins play a critical role in the normal assembly of the tight junction [106, 107], strongly indicating that the connection between these adhesion systems is real. A weak interaction between afadin and α–catenin has also been described [81, 108], providing a possible interaction between the nectin system and cadherin adhesion receptors.

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Tetraspanin Adhesion Receptors Tetraspanins, as their name implies, are membrane proteins with four membranespanning transmembrane domains [109]. The orientation of these molecules is such that both ends and the center loop of the protein face the cytoplasm, while large and small outside loops of amino acids chains are exposed outside the cell [110]. The larger cellular function of the tetraspanin family occurs when tetraspanins associate into large lateral arrays that are termed the tetraspanin web [111]. These arrays act as membrane microdomains, clustering specific membrane proteins in a region of membrane, while excluding others. The tetraspanin web may contribute to cell-cell adhesion more indirectly, namely by controlling the internalization and recycling of adhesion receptor proteins [112]. In some cases, a direct cellcell adhesion function has been ascribed to tetraspanin proteins [113], and both homomeric and heteromeric adhesion pairings have been proposed. However, interpretation of experiments designed to examine trans interactions is greatly complicated by the numerous cis interactions that drive homotypic dimerization of individual tetraspanin proteins early in their trafficking to the cell surface and thus precedes tetraspanin web assembly [114]. Tetraspanins may be anchored to actin filaments using two mechanisms. The first, and more direct, mechanism is thought to occur when cytoplasmic domains of certain tetraspanins bind to adaptor proteins that also bind actin filaments. Though many tetraspanins contain Cterminal motifs that are consensus sequences for PDZ domain-containing proteins to bind [110], the identity of such PDZ domain-containing proteins remains unknown. Importantly, actin anchoring via such protein thus remains entirely theoretical. Much more clearly defined is an indirect anchoring system for tetraspanins. Generally speaking, this indirect anchoring system occurs when tetraspanin proteins laterally associate with other transmembrane proteins that in turn become tethered to actin filaments. More specifically, this occurs when tetraspanins associate via well-described binding to the integrin adhesion system [115-118]. In fact, integrin function in adhesion can be regulated by associated tetraspanins [119]. While the molecular basis of integrin regulation by tetraspanins remains unclear, it is known that this regulation does not pertain to integrin binding to extracellular matrices, but rather acts downstream to alter integrin signaling and integrin-actin linkages [120]. Tetraspanin-integrin networks have been most well-described in cell-substrate adhesion, but a role for these proteins in cell-cell adhesion has also been observed. Importantly, tetraspanin proteins are observed at cell-cell junctions and several tetraspanin proteins, associated with integrins, have been shown to associate laterally with cadherins [121, 122]. In

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fact, integrins have been shown to localize to cell-cell junctions, where they are presumably not participating in interaction with extracellular matrices. The role of integrins in cell-cell adhesion has been largely ignored, but association of integrins with the cadherin cell-cell adhesion system through tetraspanins could allow, in theory, for the cell-cell adhesion activity of the cadherin system to be coupled to the well-documented actin-membrane linkage function of integrin cell-substrate adhesion receptors. In such a scenario, integrins are activated by signal modulation controlled by cadherins, tetraspanins, or both, rather than by engagement with extracellular matrix. It is our opinion that dissection of tetraspanin and integrin function at cell-cell junctions, particularly with regards to a role for integrin-based actin-membrane connections, will provide important molecular insight into how cells control the strength of cell-cell adhesion.

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The Tight Junction Adhesion System The tight junction is a specialized region of cell-cell adhesion that delineates the lateral membrane domain from the apical membrane domain [123]. The prototypical adhesion receptor of the tight junction is occludin, the first transmembrane proteins of the tight junction to be identified. Subsequently, an unrelated family of proteins with a similar membrane topology, the claudins, was also found to be part of the tight junction [124]. Though discovered later, claudins are required for occludin to localize to the tight junction [125] and it is claudins that make up the physical strand-like structure of the tight junction [126]. The molecular architecture of occludin and of claudins is strikingly similar to that of the tetraspanins; occludin and claudins span the membrane four times, have large and small extracellular loops, and have short cytoplasmic tails that interact with PDZ domain proteins. The extracellular loops of occludin, and of claudins more generally, mediate a tight and lowproximity cell-cell interaction whose function is to act as a selective barrier to the diffusion of ions and solutes across epithelial barriers via paracellular transport [127]. In addition, the tight junction also limits the lateral diffusion of lipids and membrane proteins between the apical and lateral membranes, ensuring that the membrane identity of each of these membrane domains remains unique. Importantly, tight junction assembly is dependent on the prior engagement of other adhesion systems. Both cadherin-based [128] and nectin-based [107] adhesions are required for proper assembly of the tight junction. The intracellular domains of occludin interact with a series of adaptor proteins that provide an actin-membrane connection (Figure 11) [123]. The adaptors of tight junctions are the ZO MAGUK proteins, membrane-associated guanylate kinase homologues that actually bear no enzymatic activity [129]. Occludin binds ZO-1 [130, 131] and ZO-2 [132], while claudins associate with ZO-1, ZO-2, and ZO-3 [133]. Of these three ZO proteins, only ZO-1 and ZO-3 associate directly with actin filaments [130, 134]. ZO-2, in contrast, can only bind actin filaments indirectly, which is does through an interaction with protein 4.1R [135]. Though the map of interaction between actin, ZO proteins, and occludin is complex, there are multiple possible architectures that result in actin anchoring of the tight junction adhesion receptor system [123]. Importantly, there is ample evidence of crosslinking between the tight junction and other cell-cell adhesion systems; ZO-1 and ZO-2 both bind α-catenin [132, 136, 137] and ZO-1 binds afadin [105].

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Figure 11. Molecular organization of the tight junction. The darker grey cytoplasmic adaptors represent ZO-1 and ZO-3. The light grey adaptor represents ZO-2, which does not bind F-actin directly. The black adaptor represent protein 4.1R.

Regulation of Actin-Membrane Connections during Epithelial Morphogenesis Whatever the precise nature of actin-membrane connections at cell-cell junctions, it is clear that these linkages must be reversible. That is, controlled assembly and disassembly of these connections must be possible to allow tissue remodeling. Foremost candidates for regulation of actin-membrane linkages at cell-cell junctions are components of adhesion systems themselves. In the simplest method of driving detachment of cell-cell interactions within tissues, expression of critical components of adhesion receptor systems, usually the adhesion molecules themselves, is blocked. In many cases, however, detachment of cells precedes such changes in expression. In fact, in some cases fully detached epithelial cells retain the ability to establish cell-cell adhesions, though these are never strengthened to stable cell-cell junctions [43]. Thus, it is a more complex post-translational control of adhesion that accounts for rapid detachment of cells from tissues. The cadherin complex component βcatenin has been shown to be phosphorylated in a manner that changes its affinity for binding partners [138] and this phosphorylation has been proposed to direct changes in adhesion. Another notable candidate is the central cell-cell adhesion regulator α-catenin. Given recent finding regarding how this protein works at the molecular level, this very well known protein‟s precise role in cell-cell adhesion remains to be fully characterized. As pointed out previously, the prevailing dogma that α-catenin‟s actin-binding function serves to link

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cadherin molecules to actin filaments has been seriously challenged with direct experimentation [11, 48, 78]. In the primary alternative biochemical mechanism accounting for α-catenin function at cell-cell junctions, α-catenin acts on actin dynamics independently of cadherin binding, but uses cadherin binding as a mechanism for recruiting α-catenin to sites of cell-cell adhesion [78]. Thus cadherin engagement results in the concentrated sequestration of cadherin, and thus α-catenin, to sites of cell-cell adhesion. Free α-catenin in this area then dimerizes and acts as both an actin bundling protein and a blocker of branched actin structure formation [78]. Thus, α-catenin may serve to ensure that actin filaments at cell-cell contacts are unbranched and can be easily arranged into bundles, precisely the organization that is observed at stable epithelial cell-cell contacts. Alternatively, α-catenin could act as a tension sensor at cell-cell contacts [85]. Proteins implicated in maintaining actin structure at cell-cell adhesions, but not necessarily currently considered core components of cell-cell adhesion systems, are also candidates for maintaining actin-membrane connections that can be regulated by the cell. Such actin regulatory proteins include zyxin, VASP, and vinculin. These proteins are concentrated at points of cell-cell adhesion, where angular bundles of actin filaments terminate at cell-cell contacts on opposite sides of the junction [15, 35, 36]. VASP function appears required for normal cell-cell adhesion [36, 50]. Similarly, expression of zyxin mutants also alters cell-cell adhesion properties; expression of a dominant negative zyxin mutant elicits decreases in the rate of cell-cell junction formation and strengthening, while expression of a constitutively active zyxin mutant drives faster cell-cell junction formation and strengthening [35]. Expression of these same zyxin mutants also affects the ability of cell to detach during growth factor-induced scattering [15]. Zyxin and VASP, like β-catenin, are regulated by phosphorylation. In the case of VASP, this changes the affinity of VASP for actin filaments [139-141]. In the case of zyxin, it releases a head-tail interaction that allows interaction with binding partners, including VASP [142]. Vinculin is regulated by a head-tail interaction in a manner that is similar to that of zyxin, though regulation is controlled by a number of mechanisms [143-145]. All of these regulatory mechanisms allow the activity of these proteins to be switched on and off, allowing actin-membrane interactions to be maintained or disrupted during cellular signaling associated with tissue remodeling. Consistent with such an idea, expression of a zyxin mutant that mimics phosphorylation that prevents the head-tail interaction results in cell-cell junctions that are resistant to rupturing during growth factor-induced scattering [146].

Rho Gtpases as Central Players in Local Control of Actin Dynamics and Cell-Cell Adhesion Rho family GTPases play a central role in controlling cell morphology by acting as master regulators of both actin dynamics and adhesion systems. The Rho family of small GTPases is part of the Ras superfamily and contains many members, all of which appear to act as morphological regulators by affecting actin organization in cells [147]. Most characterized are Cdc42, Rac1, and RhoA. Expression of these proteins in mammalian fibroblasts drives filopodia, lamellipodia, or stress fiber formation, respectively [148-151]. These same GTPases play a major role in cell-cell adhesion [152]. In epithelial cells, expression of dominant negative forms of Rac1 and cdc42 decrease cell-cell adhesion, while

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expression of constitutively-active forms increases cell-cell adhesion [39, 153, 154]. This effect correlates with changes in actin staining at cell-cell junctions [155, 156], as well as overall changes in cell-cell junction morphology [156, 157]. Current thinking is that Rac1 drives actin rearrangements, while RhoA activity drives actin contractility, both of which combine to expand and reinforce initial cell-cell contacts [158]. In fact, localization of Rac1 and RhoA activity at expanding cell-cell junctions is restricted to the boundaries between cell-cell contact and non-contacting surfaces [158]. While Rho GTPases regulate cell-cell adhesion, cell-cell adhesion also impacts the levels of active forms of these GTPases, helping to establish spatial regulation of their activity during cell-cell junction formation. Cellular levels of Cdc42 and Rac1 activity is increased following initiation of cadherin-based cell-cell adhesion [159, 160], though this appears to be only a transient effect. Engagement of cadherin- and nectin-mediated adhesion in cells cultured in suspension results in a long term decreases in Rac1 activity following a transient period of activation [161]. This is consistent with observations that initiation of cadherinbased adhesion with a bead coated with cadherin extracellular domains results in the immediate, highly localized, and highly transient recruitment of Rac1 to the site of cadherin engagement with the bead [61]. Tetraspanin CD151, the same tetraspanin that links cadherins to integrins [122], activates signaling through Cdc42 in a manner that regulates cell-cell adhesion [162]. Further, and though its role at cell-cell junctions remains unclear, integrin signaling also results in activation of Rho GTPases, in particular Rac1 and RhoA [163, 164]. Tetraspanins serve to modulate integrin signaling [120], a role that alters epithelial morphogenesis programs [119, 165]. Rho GTPases function by switching to a GTP-bound state and then binding downstream effectors that, in turn, alter actin dynamics and adhesion. GTP-bound RhoA associates with and activates rho kinase [166]. The result is activation and subsequent phosphorylation of target proteins myosin light chain kinase and myosin phosphatase results in activation or inhibition, respectively, of these targets to induce myosin-based contractility [167-170]. GTP-bound Rac1 and cdc42 also associates with downstream effector kinases, the PAK family [171]. More importantly, however, the Rac1/Cdc42 binding motif in PAK is found in a number of actin regulatory proteins that control actin nucleation and polymerization. Rac1 and Cdc42 affect changes in actin dynamics by binding and activating several actin regulatory proteins, including NWASP, IRSp53, and IQGAP [172]. Besides directing changes in actin dynamics, Cdc42 works independently of actin regulatory systems to inhibit Rho-mediated actin contractility [173]. While the Rho family consists of many additional members outside of Rac1, RhoA, and Cdc42, the concerted regulation of activity of these three well-studied Rho GTPases could provide a framework for how cells exert spatiotemporal control of actin reorganization and contractility. In this framework Rac1 and Cdc42 activity drive actin reorganization events, while Rho drives myosin-dependent contractility. Superimposing areas of Cdc42 and RhoA activity would allow actin reorganization, but without myosin-based contractility. Superimposing areas of Rac1 and RhoA activity would allow actin reorganization and contractility to occur together. Thus, Cdc42 determines whether actin rearrangements, driven by Rac1 and Cdc42 itself, are accompanied by myosin-dependent contractility is response to RhoA activity.

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Epithelial Morphogenesis in Development: The Interface of Actin Dynamics, Cell-Cell Adhesion, and Cellular Signaling

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One of the most interesting places to observe cellular adhesion dynamics is in embryonic development, where changes in the shape, motility, and adhesion of individual cells impact tissue architecture in order to drive morphogenesis as required to form an organism. While the contribution of actin reorganization, contractility, and membrane connections to epithelial morphogenesis have been initially addressed by in vitro studies, as detailed in previous sections of this chapter, recent and emerging studies in diverse animal model systems undergoing development have provided important additional views about how actin and adhesion contribute to epithelial morphogenesis. Importantly, since cell signaling and the related signal transduction pathways driving morphogenesis have primarily been studied in the embryo, developmental biology can make major contribution in our understanding of how actin regulatory systems are controlled to generate the correct timing and positioning of specific actin dynamics events that underlie epithelial morphogenesis. Studies in Drosophila have been critical to identifying the several known molecules required in epithelial morphogenesis, from establishing cell polarity, bending and shaping the ectoderm, directional cell migration, and axonal outgrowth and pathfinding, to name a few. Most of the foundational morphogenetic processes identified in Drosophila have also been explored in vertebrate models, and many of the molecular components are now being studied concomitantly in multiple model systems. Here we highlight vertebrate examples of epithelial morphogenesis during embryonic development.

Detachment of Cell-Cell Adhesions and Dispersion of Cells from Dynamic Epithelial Tissues in Development Epithelial morphogenesis events include instances where cells fully detach from the epithelial tissue. Such events are often referred to as epithelial-mesenchymal transitions (EMT). In our model, release of individual cells from tissues results from uncoupling of actin connections with cell-cell adhesion systems. Importantly, analysis of scattering events suggests that, prior to cell-cell detachment, tissues recapitulate a series of actin rearrangements that are seen in other epithelial morphogenesis events, including reorganization of cortical actin into medial networks and contractility of actin to generate forces that impinge on cell-cell adhesions [15]. Nevertheless, it is the ultimate loss of cell-cell junctions that allows individual cells to isolate the effects of shape changes and contractility from other cells in the tissue. This loss of adhesion may result from reduced adhesion receptor system expression, cadherin “switching,” or loss of actin-membrane connections at adhesion sites. Detachment of cell-cell adhesions associated with complete scattering of an epithelial tissue into individual cells is known to follow reduced expression of adhesion receptors in many, but not all, instances. Negative regulators of cadherin transcription, such as Slug [174], Snail [175], and Twist [176], are upregulated in cells that will undergo scattering events in several model systems for epithelial morphogenesis events . Cadherin “switching” is a term

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that refers to changes in the specific cadherin expression profile in the cell [177]; cells change which cadherins are expressed and presented at the cell surface to participate in cell-cell adhesion. In many model systems, both in culture and in vivo, many cells undergoing dramatic epithelial morphogenesis events that end in cell-cell detachment undergo a transition from expression of E-cadherin to expression of N-cadherin [178, 179]. Cadherin switching at the molecular level is the regulation of proteins that bind to p120 catenin and alter its preference for associations with specific cadherin family members [180]. Though cadherin switching allows tissues to physically sort cells into different subpopulations based on their cadherin expression, it is unclear how changing cadherin expression can result in cell-cell detachment within each subpopulation when newly expressed cadherin family members can mediate cell-cell adhesions within the population. Thus cadherin switching is a possible mechanism that would allow a tissue to become partially disrupted into groups of cells that maintain cell-cell adhesion with one another, and not complete disruption of the tissue into individual, solitary cells, as would occur with downregulation of cadherin expression. Regulation of actin-membrane connections with cell-cell adhesion systems is the most poorly understood mechanism for allowing cells to uncouple their actin dynamics from neighboring cells. Control of adaptor proteins that link actin filaments to adhesion receptor systems has already been discussed in some detail in a previous section. An important point is that allowing cell-cell detachment by controlling actin-adhesion system linkages is not incompatible with the cadherin switching mechanism, since this would allow remodeling of cell-cell junctions within tissues expressing the same cadherins. Consideration of molecular mechanisms of uncoupling of cell-cell adhesions from actin filaments likely coincides with the precise architectural changes associated with different epithelial morphogenesis programs. In instances of complete dissolution of the epithelial tissue into individual, solitary cells, detachment of cell-cell junctions can occur globally, since target cells ultimately retain no cell-cell connections. Downregulation of cadherin expression or uncoupling actin connections with cell-cell adhesion are strong candidate mechanisms for allowing these cells to exhibit solitary behavior, whereas cadherin switching seems less likely to generate solitary cells. In instances of partial detachment of the epithelium, including extrusion of some cells from the monolayer, cell-cell junctions between populations of cells are broken, while cell-cell junctions within populations are retained. Such events are best explained by cadherin switching, or by local regulation of actin linkages with cell-cell adhesion systems. In the embryo we find several well-studied examples of EMT that allow evaluation of the molecular links between embryonic signal transduction and both actin regulatory system and cell-cell junction dynamics. The neural crest is probably the most thoroughly studied example of EMT in vertebrate development. Neural crest cells originate at the dorsal margins of the neural plate at an interface between neural ectoderm and surface ectoderm, as will be described in more detail in the following section. Early in neurulation, neural crest cells undergo EMT; neural crest cells break cell-cell adhesions, delaminate from the epithelial tissue, and migrate throughout the embryo to contribute a multitude of cell types. Neural crest-derived cell types include peripheral neurons and glia, melanocytes, bone and connective tissue, and many others. The differentiation of neural crest cells is partially dependent on cell-intrinsic determinants, but also relies on local microenvironmental cues present in the several migratory destinations. The migratory paths are rich in ECM molecules such as fibronectin and laminin, and neural crest cells express matrix-binding integrins, thereby promoting their general migratory abilities [181]. Neural crest cells are also directed

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by chemical cues as they migrate. A well described molecular example of this is found in neural crest cells that migrate through the medial somite (schlerotome). Here, the posterior halves of each somite express the ephrin ligands, while expression is absent in the anterior halves. Ephrin receptors are expressed by neural crest cells, and when activated by ephrin ligand, cause cells to retreat, thereby directing cell migration toward other more favorable environments, namely the anterior somite [182]. Similarly, axon growth cones expressing Eph-receptors collapse when encountering high levels ephrin ligand, thereby directing the axon in a different direction [183-185]. The cytoskeletal changes associated with ephrin activation are more relevant to a discussion of cell migration, and are therefore not discussed here. Instead we focus briefly on the regulation of neural crest EMT. Neural crest cells are specified within the dorsal-most region of the neural plate in a region referred to as the neural folds (Figure 12).

Figure 12. Epithelial morphogenesis in the formation of the neural tube and neural crest. Neural tube development occurs as depicted from the central region of the ectoderm. Flanked by surface ectoderm, the neural plate (A) soon begins to bend at the midline to form the neural groove (B). Dorsolateral hinge points soon bring the neural folds into close proximity (C), where dynamic changes in cell adhesion allow for midline fusion of the neural tube, and separation from the overlying surface ectoderm, which also fuses as the midline (D). Neural crest cells also form from the neural folds (#).

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One important aspect of neurulation is the early patterning of this tissue, which allows for specific cell types to be specified within specific regions along the dorsal-ventral axis. This is accomplished primarily by secreted molecules or morphogens produced at either the ventralmost or dorsal-most regions of the developing neural tube. Ventrally, sonic hedgehog (Shh) is expressed first by the underlying notochord, and then by the floorplate of the neural tube. Shh is able to diffuse dorsally, thereby creating a concentration gradient wherein high levels of Shh direct the differentiation of some cell types, while lower concentrations (dorsal) direct cells toward a different fate. Dorsal signals, such as BMP and Wnt act similarly to specify dorsal fates under high concentrations. Therefore high BMP/Wnt signaling and low Shh signaling results in the dorsal-most cell type, the neural crest [186]. Located adjacent (lateral and dorsal) to the neural crest zone is the surface ectoderm, which is molecularly distinct from the neural ectoderm in that it expresses E-cadherin in contrast to the N-cadherin-expressing neural crest and neural ectoderm. In the region of neural crest cell formation, N-cadherin has been shown to be downregulated just prior to the time neural crest cells undergo EMT [187]. Relevant to this topic, neural crest cells express the transcription factor Slug [188], which has been shown to downregulate expression of cadherins [174]. N-cadherin expression is maintained in the remaining neural plate, hinting that its downregulation is needed for detachment from the epithelium, and in agreement with the adhesion model where loss of cadherin expression can drive cells to detach completely from the epithelial tissue. More recent research has expanded what we know about embryo signaling and cadherins in neural crest development. BMP signaling has been shown to play a role in the downregulation of membrane-localized Ncadherin in neural crest cells, and cadherin-6B is thought to be expressed at higher levels in the neural crest just prior to EMT [189]. New data suggests cadherin 6B and BMP act to reciprocally induce one another, resulting in N-cadherin repression and neural crest deepithelialization [190]. Here it was shown that cadherin 6B is upregulated and maintained in pre-migratory and migratory neural crest cells and that blocking its function or expression resulted in reduced EMT, suggesting a cadherin switching event in generating the neural crest. Interestingly, neither cadherin 6B nor β-catenin were localized to the apical region of cell-cell contacts in pre-migratory neural crest cells, while N-cadherin and β-catenin were apically enriched in the adjacent neuroepithelium. Similar results were shown for other AJ proteins, hinting at a reduction or loss of cellular polarity in neural crest precursor cells. Ectopic cadherin 6B expression caused similar cell polarity disruption in non-neural crest cells, and even promoted EMT. This further supports the idea that cadherin switching is a key regulator of cellular adhesion changes associated with delamination of neural crest cells. A likely result in the loss of polarity and columnar morphology of neural cell precursors is that apical constriction of surrounding N-cadherin-expressing cells could force neural cell precursors to become extruded from the epithelium in a basal direction. Firm links between the developmental signals that specify neural crest cells and the mechanical proteins that regulate cell-cell adhesion are few and only recently have some of the cell adhesion and cytoskeletal effectors been characterized. In zebrafish it has been shown that breakdown of cellular junctions during formation of the neural crest relies on Rho-kinase and myosin II [191]. Additional signaling factors such as Notch, FGF and several Sox-family transcription factors have been identified as regulators of EMT in the neural crest [192], while more recently-described molecular contributors are αN-catenin, which must be repressed for

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normal EMT [193], and Rho family member RhoU, which is required for late EMT and cell migration in the cranial neural crest [194]. Another related example of EMT occurs in the nearby surface ectoderm, where in the head sensory neurons form from regions of the epithelia called neurogenic placodes. Here, cells delaminate from the ectoderm as individuals or as small clusters, then migrate to condense into neuronal aggregates called sensory ganglia (Figure 13). While consistent progress has been made toward understanding their differentiation, little is known about the mechanisms that drive EMT in these cells. It appears that most cells exit the epithelial through small gaps that form in the basal lamina. Detachment and migration are accompanied by obvious cell shape changes and classic migratory morphology. After EMT and cellular migration, placode cells condense into ganglia, sometimes only with other placode cells, but often with neural crest cells. Little is known about what drives re-adhesion of differentiating neurons. However, N-cadherin expression is required for proper re-aggregation of these cells, as is Slit/Robo signaling [195], which suggests a E-cadherin to N-cadherin switching events has occurred and this could contribute to detachment of placode cells from the ectoderm. As for the initial EMT, while little is known regarding the neuronforming placodes, it appears that Rho-dependent apical constrictions involving actin and myosin II are required, at least in the non-neurogenic lens placode [196]. This could be reminiscent of the role of apical constriction in surrounding cells to extrude neural crest cells, only now applied to extrude placode cells. In the trigeminal placodes, blocking FGF or Wnt signaling resulted in failed neurogenesis, and failure of cells to detach from the ectoderm [197, 198].

Figure 13. Epithelial morphogenesis in placodes. Neurogenic placodes, such as the trigeminal placode depicted here, form bilaterally in the surface ectoderm of the vertebrate head. Other placodes such as the non-neurogenic lens placode, and placodes with mixed derivatives, such as the otic placode also for bilaterally. Each has a somewhat unique molecular and morphological developmental program. Trigeminal placode cells undergo EMT over a protracted period of time similar to what is shown. Cells migrate and later condense as neurons to form sensory ganglia.

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Additionally, blocking Notch signaling resulted dramatically enhance neurogenesis, even within the surface ectoderm [199]. Though Notch inhibition did not drive individual cells to detach from the ectoderm, the ectoderm often appeared fragmented, indicating that cell-cell junctions were disrupted. It remains unclear whether differentiation and EMT are coupled. It is known that neurogenin, a Notch effector gene, directly regulates the expression of neuronal differentiation genes and cell adhesion genes, coupling neurogenesis with cell migration through the bHLH transcription factor Ngn2 [200, 201]. Little is known about the interface between cellular signaling and the actin regulatory machinery that drives epithelial morphogenesis in placodes. However, the simplicity of the system makes it an attractive developmental model to define this interface at the molecular level. Somite development is another instance where cell-cell adhesion changes drive an epithelial morphogenesis program (Figure 14). Somites are arranged as a segmented array of epithelial tissue blocks that form from mesoderm adjacent to the notochord and adjacent to the neural tube. Unassociated mesodermal mesenchymal cells aggregate into these epithelial blocks in a process that moves from head to tail, such that a new pair of somites is formed about every 90 minutes in the chick or mouse model systems. Extensive studies over the past decade have focused primarily on the dynamic wave-like patterns of gene expression that direct proper somitogenesis from within the unsegmented mesoderm [202, 203]. Once formed, the somites quickly reshape into somite compartments, namely the sclerotome, which is a ventral mesenchymal compartment that will primarily become vertebral bone, and the dermamyotome, which is a dorsolateral compartment that remains epithelialized and will primarily contribute muscle and dermal connective tissue to the organism. The sclerotome undergoes a dramatic EMT event to give rise to cells that assemble parts of the skeletal system.

Figure 14. Epithelial morphogenesis in somites. Somites form along the length of the vertebrate axis, first forming undifferentiated epithelial block with a central fluid-fill cavity. Over time, the somites are patterned by several signals, including Shh and Noggin from the notochord. As the somite differentiates, these patterns become apparent as the ventromedial sclerotome, and the dorsolateral dermamyotome. The sclerotome undergoes global EMT to become a mesenchyme tissue, while the dermamyotome remains epthelialized. Myoblast cells will eventually undergo EMT from the medial and lateral margins of the dermamyotome.

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During muscle cell differentiation, individual myoblasts located at the medial and lateral margins of the dermamyotome will undergo EMT, detaching from the tissue and migrating throughout the embryo to establish muscle tissue. As such, somite development provides two unique examples of EMT: in the sclerotome, where the immature somite epithelium is completely disrupted, breaking all cell-cell adhesions to fully return to a mesenchymal state, and in myoblasts, which delaminate one after the other from the dermamyotome over a protracted period and then migrate as individual cells. In sclerotome de-epithelialization, it has been shown that early Noggin and Shh expression from the notochord act to pattern the developing somite, including specifying the sclerotome compartment at higher concentrations [204]. Once specified little is known regarding the signals that induce EMT in the sclerotome. It may be similar to what is seen in neural crest cells, since matrix metalloprotease-2 (MMP2) has been shown to be required for EMT in both model systems [205]. In the dermamyotome, it has been shown that asymmetric cell division and preferential N-cadherin segregation into apical (dermamytomal) but not basal (post-mitotic myotomal) cells is needed for dermamyotomal cell adhesion and maintenance of the muscle precursor pool in this tissue [206]. Surprisingly, N-cadherin protein is maintained in differentiating myoblasts [207] as they undergo EMT at the medial and lateral borders of the epithelium, indicating that while N-cadherin is involved in somite and muscle development, its downregulation is not required for myoblast EMT. It is here that regulation of actin connections to cell-cell adhesion systems most likely accounts for cell-cell detachment. It has been shown that scatter factor/hepatocyte growth factor (SF/HGF) regulates myoblast EMT, as c-met mutants show deficiencies in myblast migration, and ectopic SF/HGF administration enhances myblast emigration [208, 209]. Importantly, cadherin surface expression is retained following HGF stimulation of tissue culture cells, despite which dramatic cell-cell detachment occur [43]. More recent studies have supported a role for SF/HGF and c-met in early muscle development, such as one identifying Met signaling as needed here and in neuromast release from the lateral line placode in zebrafish [210]. Interestingly, c-met is a direct target of the transcription factor Pax3 [211], which is highly expressed in pre-migratory and migratory myoblasts, and in pre-migratory and migratory trigeminal placode cells that were discussed earlier [212]. However, a role for HGF/met has not been investigated in placode cell EMT/migration. And other signaling mechanisms may contribute to EMT in this system: Wnt11 was shown to be required for neural crest and dorsal somite EMT in Xenopus. Here Wnt11 activates a non-canonical pathway distinct from the PCP pathway – the Wnt/Ca+ pathway. Here it was shown that calcium/calmodulin-dependent kinase II (caMKII) is a modulator of EMT, and is a target of Wnt11 signaling [213]. As we see from these vertebrate examples, EMT is critical for normal development, but there are many unanswered questions regarding the link between embryo patterning, instructive signals for EMT, and the cellular machinery that ultimately drives proper cell-cell adhesion and actin dynamics.

Actin Dynamics That Drive Tissue Deformations – Furrows, Tubes and Vesicles Another common type of epithelial remodeling during development occurs when a flat sheet of tissue bends, curves, or rolls into a tube or vesicle. Examples include the formation of

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the ventral furrow and posterior midgut invagination in Drosophila, the shaping of the neural tube, otic and optic vesicles in vertebrates, and gut tube morphogenesis in several species. Additional but more subtle examples include segment boundary formation in fruit flies, and rhombdomere boundary formation in the vertebrate brain, where simple furrowing does not progress to tube formation and tissue separation as is seen in the prior examples. For tissue bending morphogenesis, cells must precisely wedge or change shape in some other way to promote tissue morphogenesis in the select region. Currently, structural changes in the actomyosin network, as well as more subtle changes in cell-cell adhesion, are thought to be the primary contributors to tissue bending, with apicobasal polarity being key to directionality of the bending event. Here we focus on neural tube formation as an example of tissue bending, including the important signals required for proper specification and formation of the neural tube. Neural tube formation in vertebrates has been described foundationally in the chick embryo model system, where embryo accessibility was historically an advantage to studying neurulation. In chick, as in other vertebrates, the neural plate forms near the time of gastrulation in the central region of surface ectoderm overlaying the notochord and developing somites (Figure 12). Neural plate cells differ from adjacent ectoderm in that they quickly begin to take on a columnar shape, with the neural ectoderm becoming progressively more thickened as development proceeds. The thickening of the neural epithelia appears fairly uniform across the width of the plate, although early evidences of tissue shape changes are apparent at the midline, where the neural plate begins to form a visible groove, dipping down toward the notochord, with laterally adjacent regions of the plate becoming elevated in the dorsal direction. Some of the shape changes occurring in the neural plate are thought to be due in part to extrinsic factors, such as pushing forces from the surface ectoderm [214, 215], while additional internal forces due to cell movement and cell shape changes are also at play [216, 217]. The groove midline is sometimes referred to as the medial hinge-point (MHP). Soon the neural plate appears as a deep groove, with the left and right halves approximating one another and the adjacent surface ectoderm being drawn upward and adhering to the lateral margins of the neural plate (referred to as the neural folds), which are now located dorsally (Figure 12). As neural tube morphogenesis progresses, additional flexures become evident bilaterally. Referred to as the dorsolateral hinge-points (DLHPs), these flexures act to broaden the lumen in the lateral direction, and perhaps bring the neural folds into proper alignment for tube closure. The DLHPs are typically located dorsal to the dorsal-ventral (D-V) midline, a position likely dictated by D-V patterning signals such as the ventral sonic hedgehog (Shh) signal and dorsal Wnt or BMP signals. While the precise mechanisms have not been determined, surface ectoderm is required for, and Shh overexpression prevents, proper DLHP formation in the spine [218]. Interestingly, excess BMP signaling seen in Noggin mutants results in a loss of midbrain DLHP formation [219], BMP2 expression is found in areas lacking DLHPs, and BMP2 mutant mice display enhanced DLHP formation [220]. A mechanism wherein Shh suppresses dorsal Noggin expression is proposed to account for the interplay between D-V patterning cues (Figure 15). While neurulation is somewhat different in zebrafish, where a neural mass forms and subsequently undergoes cavitation to create the lumen, DLHPs still appear as in other vertebrates, and similarly require an apical medial actomyosin network. The zinc-finger transcription factors zic2a and zic5 are required for this process in zebrafish, and may act upstream of apical reorganization of F-actin and myosin II into the medial actin network, a process that is dependent on canonical Wnt signaling [221].

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With the neural folds abutting one another, dynamic changes in cell adhesion cause the two folds to fuse, and the overlying ectoderm to separate from the neural ectoderm and fuse bilaterally to form a continuous sheet of dorsal ectodermal tissue. Meanwhile, neural crest cells, which form at the surface ectoderm/neural ectoderm boundary, undergo EMT and begin to migrate as mesenchyme cells, as discussed in the previous section. From this brief description of cellular rearrangements, one can imagine the complex changes in cell-cell adhesion that are required for proper morphogenesis at the neural folds. It is known the Ncadherin and E-cadherin are differentially expressed in neural and surface ectoderm cells, respectively, and this differential expression is thought to be required for proper sorting of cells into ectoderm and neural plate. Midline fusion of the surface and neural ectoderm may be distinct processes, since disruption of NF-protcadherin (NFPC) only prevents neural tube closure, but not surface ectoderm fusion [222]. Neural crest cells form at the interface of the two epithelial types, and require the downregulation of N-cadherin prior to undergoing EMT [187]. Additionally, BMP signaling is thought to play a role in downregulation of membrane-localized N-cadherin in neural crest cells, and cadherin-6B is thought to be expressed at higher levels in the neural crest [189].

Figure 15. Proposed molecular regulators of dorsolateral hinge point (DLHP) formation in vertebrates. Shh from the notochord and floor plate restricts Noggin expression from expanding ventrally. Noggin, and known inhibitor of BMP signaling, restricts the inhibitory action of BMP, thereby allowing DLHP formation (after [40]). Additionally, Wnt signaling from the dorsal neural folds acts to regulate Zic2/5 expression, which may have a direct effect on actomyosin contractions and on adherens junctions (after [41]).

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New data suggests cadherin-6B and BMP act to reciprocally induce one another, resulting in N-cadherin repression and neural crest de-epithelialization [190]. Additionally, it was shown that cadherin-7 and -11 are upregulated by in neural crest cells, a phenomenon dependent on activation of canonical Wnt signaling [223]. As such, many signaling molecules are involved in neural crest formation, with cadherin-mediated adhesion/signaling also being a core regulator of the process. Back in the neural ectoderm, coupling of robust cell-cell adhesion at adherens junction with contraction of an apical medial actin network forces cylindrical cells to become conical in shapes, with the apical domain at the point of the cone. The end result at the tissue level is rolling of the neuroepethelium into a tube. One primary molecular regulator in the process is cadherin, which appears to be playing a broad role in shaping this tissue. As mentioned, zebrafish neurulation begins with a mass of cells which later sort out to become a defined neuroepithelium with a lumen. This process requires CE and intercalation, which are disrupted in N-cadherin mutants [224]. Hedgehog (Hh) signaling may negatively regulate integrins apically, resulting in a localized increase in cadherin activity that may help specific the apicobasal axis for constriction [225]. N-cadherin may act in concert with the tight junction protein ZO-1 to establish this apicobasal polarity, with their roles preceding those of Lin7a and Nok, which appear to be crucial for maintaining the already established polarity [226]. N-cadherin-mediated control of the plane of cell division may also be important [227], further supporting a critical role for N-cadherin in cellular polarity as required for neural tube morphogenesis. A very recent study evaluated the complex interplay between AJs, cadherins, Hh signaling and cell proliferation. Here, N-cadherin mutants displayed a cell proliferation increase in the neural tube resulting from ligand-independent Hh signaling activation. Hh mutants, on the other hand, displayed reduced AJ markers, leading to the hypothesis that Ncadherin-mediated adhesion regulates cell proliferation in the neural ectoderm due to reciprocal interactions between AJs and Hh signaling [228]. As such, in addition to adhesive and cell shape requirements for proper tube formation, cell proliferation must be tightly regulated throughout the process. Deformations of epithelial sheets can be oriented along an axis or towards a point, depending on what final tissue organization is required. Whichever the case, the underlying morphogenetic movements of individual cells are shared. Cylindrical shaped cells must be converted to conical shaped cells. In our model, this occurs when cells remodel their cortical actin ring into medial actin network with connections to cell-cell adhesion systems. Symmetrical myosin-based contraction of the medial network results in a narrowing of the cell. The conversion of cells to a conical shape occurs when contraction is restricted along the apicobasal axis of polarity. Contraction near the apical surface constricts the apical surface of the cell, while contraction of actin networks near the basal membrane constricts the basal area of the cell. Since the basal cell surface maintains connections through cell-substrate adhesions, contraction must be focused through cell-cell and cell-substrate adhesion systems. This model is supported by observations of actin organization and contractility in epithelial deformation programs during development [13]. The key to this process is the asymmetric contraction along the apicobasal axis. Conceptually, this is easy to explain, since the distinct compositions of the apical and basal regions could allow activation of local signaling networks that impact only local actin dynamics. This model highlights the need for local, polarized signaling during epithelial morphogenesis. Alternatively, the cell may take advantage of the distinct zones of adhesion that are apparent in cell-cell junctions along the

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apicobasal axis [229]. Most apical are the tight junction and adherens junction proper with its belt-like cortical actin ring, adhesion sites where cadherin-based adhesion is linked to nectin and claudin/occludin adhesion receptor systems. Below this region, cell-cell adhesion is more exclusively mediated by cadherin adhesion receptors. It is possible that cells can distinguish between actin networks associated with different regions of cell-cell adhesion. Remodeling of cortical actin associated with the adherens junction proper and tight junction into a contractile medial actin network would result in an apical constriction. Contraction of medial actin networks associated with integrin-based adhesions and cadherin-based adhesions below the adherens junction and tight junction would result in a basal constriction. In fact, maintenance of the actin architecture associated with cell-cell adhesion outside of the region of contraction would actually prevent contractile forces from constricting the opposite end of the cell, further ensuring the acquisition of a conical cell shape. Such a model would not necessarily require polarized signaling, as long as global changes in signaling could differentially impact specific actin networks.

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Cellular Rearrangements within Epithelia – Convergent Extension by Intercalation A frequent example of epithelial morphogenesis occurs when cell position changes within the epithelium combine to promote tissue elongation or other gross changes in the two dimensional shape of the epithelium. In most organisms, the shape of the embryo or the shape of a developing organ system undergoes elongation events, which require rapid and dramatic alterations in the two-dimensional shape of the shape of epithelia (Figure 16). At the cellular level, this occurs as groups of cells converge toward an axis within the plane of the epithelium, intercalate with one another, and thereby elongate and narrow the epithelial sheet. This process, referred to as convergent extension (CE), is relevant to several morphogenetic processes, such as early gastrulation in vertebrates, germ-band extension in Drosophila, and neural tube morphogenesis. Principles that drive CE include direction sensing by chemoattraction, changes in both cell-cell and cell-substrate adhesion systems, and cell shape changes, all of which combine to allow cells to alter their relative position within the plane of the epithelial tissue. One fairly well characterized example of epithelial morphogenesis based upon CE is the complex process of axis elongation and corresponding gastrulation in early vertebrate development. While the process has been studied in all vertebrate model systems, historical studies in Xenopus and more recent research in zebrafish have provided the primary framework for our current understanding of CE in vertebrate axis formation and gastrulation. During the initial phases, cells of the epiblast are undergoing rapid cellular rearrangements that will quickly generate the three primary germ layer cell types, while at the same time create a new, elongated shape to replace the sphere-like blastula embryo. Cells from the surface and sub-surface of the spherical mass converge toward the blastopore lip as they are instructed to take on new fates. Midline mesodermal cells are born as they migrate through the dorsal region of the lip, and are specified to intercalate and tightly adhere to one another as they form the notochord. Meanwhile, CE movements are occurring in the overlying epithelial sheet, where cells are specified to become neural ectoderm [230]. These CE events ultimately reshape an embryo with a recognizable but primitive shape, elongated along the head-to-tail axis [231].

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Figure 16. Remodeling of specific cell-cell junction to drive convergent extension.

How might actin reorganization, contractility and linkages with cell-cell adhesion systems by properly coordinated to drive CE? Live cell analysis of cell shape changes occurring in CE give some important clues. Hexagonal arrays of epithelial cells that make up the epithelium prior to CE undergo a series of shape changes that allow cells to alter position. More specifically, individual cell-cell junctions are shortened to a point, brining two cell vertices together into a single, larger vertex. In the second phase, single vertices then resolve into two vertices that move apart as a new cell-cell junction is established (Figure 16). Such repositioning requires few rearrangements of the actin cytoskeleton; the overall actin organization in epithelial cells does not change dramatically. In the prevailing view, contractility of actin cables connecting adjacent vertices is sufficient to reduce the length of a cell-cell junction between two cells in the epithelium, thus bringing the two vertices together until they meet. Less understood is the resolution of a vertex into two vertices separated by a cell-cell contact. It is possible that loss of myosin function in actin that has forced two

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vertices together is sufficient to allow a cell-cell contact to reappear. But there are clearly cellular controls, as the reappearance of a cell-cell adhesion is specifically oriented in a desired direction. This means that the overall positioning of vertices is altered and now different cells contribute a “corner” to the new vertices. This must require remodeling of actin connections with cell-cell adhesion systems at the vertices (Figure 17).

Figure 17. Actin contractility and membrane connections during convergent extension in the current model.

It is unclear how proper organization of actin-membranes at the vertex might be achieved as cells undergo CE. A high rate of binding and release in actin linkages with cell-cell adhesion systems would certainly facilitate rearrangements of actin-junction connections needed for the resolution phase of CE, where a new junction is formed on a new axis. Upon the formation of a four cell vertex, such a system would result in the association of actin filaments with random cell-cell junction attachment sites. Connections in the correct orientation could be reinforced in some manner until almost all connections are correctly aligned and the new junction could be established. Such a model is reminiscent of that proposed for organization of connections between spindle microtubules

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and centromeres of chromosomes during cell division, where connections form randomly and with a high rate of turnover, but correctly oriented connections are stabilized. Interestingly, it is tension forces that reinforce correctly oriented microtubule-chromosome connections. A tension sensing mechanism is an attractive mechanism for selecting correctly oriented actinadhesion system connections during the resolution phase of CE. There is evidence to support a connection between tension and actin dynamics at adhesion sites. Though this has been described for integrin-based focal adhesions, it is zyxin dependent [232-234] and zyxin also function in cell-cell adhesion. Further, both α-catenin and vinculin have both been proposed to be tension sensors that act to stabilize actin connections to cell-cell junctions [85, 235]. Recent work analyzing the localization of myosin-based contractility in CE challenges the accepted model where contraction of actin networks without prior alterations of standard epithelial actin organization serves to pull two or more vertices into a single vertex [14]. During convergent extension in Drosophila, actin was observed in medial actin networks that undergo pulsed contractions in a manner similar to those observed by Martin et al. during Drosophila gastrulation [13]. An alternative model for how actin-based contractility drives CE is that convergent extension movements do not dramatically differ from other collective epithelial morphogenetic events. CE begins with a reorganization of the cortical actin ring from the cell-cell junctions into a medial actin network. This is then followed by myosin-based contraction of the medial network. Cell-cell junctions are maintained, as are actin linkages with cell-cell adhesion systems, ensuring that contractile forces are transduced through the tissue. Essentially, a contractile network is assembled around the junctions that must be shortened and pulsed contraction force the two vertices together (Figure 18). Resolution of a larger vertex into two smaller vertices separating a junction oriented in the correct direction is less clear. While contractility without actin reorganization accounts for how epithelial sheets generate CE instead of a sheet deformation in the standard accepted model, here it is the control of the contractile vectors that are used to generate CE in this model. In fact, the alternative model for how actin dynamics drives CE actually serves to simplify our unifying model for how actin dynamics control epithelial morphogenesis generally, since all tissue remodeling events would share the same series of actin dynamic events, namely reorganization of cortical actin into medial actin networks followed by contractility. Junctions where contractile forces will be applied could be the only ones whose cortical actin is reorganized into the medial actin network, the result being that contraction of the medial actin network applies forces only upon specific cell-cell contacts and not others. Alternatively, the entire cortical actin ring might be remodeled into a medial actin network, but connections between this network and each cell-cell junction could be locally altered, with robust connections at target junctions and looser, unproductive connections elsewhere. How cells direct contractile forces to specific cell-cell junctions rely on polarized signaling that would alter local actin dynamics. That polarity of signaling occurs within the plane of the epithelial tissue suggests several known developmental signaling networks. The signals that and organize the embryonic axis and initiate CE have been the focus of many detailed studies. Early in vertebrate development, several pre-gastrulation events occur prior to the CE events that shape the embryo. These events are important for establishing the head-to-tail and dorsal-ventral axes, and for creating the organizing centers that drive gastrulation.

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Figure 18. An alternative model for how actin dynamics drive convergent extension.

In Xenopus, Dishevelled (Dsh) is localized to the dorsal side of the embryo during cortical rotation, resulting in the accumulation of β-catenin and the corresponding activation of β-catenin-dependent (canonical) Wnt signaling. In Zebrafish, β-catenin protein accumulates dorsally, again resulting in activation of the canonical Wnt signaling pathway. This activity, along with high Nodal signaling and upregulation of organizer-specific transcription factors and signaling molecules, results in this dorsal tissue acquiring the ability to organize the embryonic axis [236-238]. Many of the molecular signals that convey organizer activity are secreted factors that inhibit BMP or canonical Wnt signaling, and in their absence CE gastrulation movements do not occur properly. More recently the focus has shifted from understanding organizer formation and activity to translating that activity into the actual cellular movements of gastrulation. Interestingly, some of the same molecules involved in establishing the organizer seem to be critical for directing the cellular mechanics of CE. Dishevelled, for example, is a core node in both canonical and planar cell polarity (PCP) Wnt signaling. The early events of organizer formation are primarily dependent on canonical Wnt activity. CE It is difficult to imagine how canonical Wnt signaling would be effective in establishing the directionality of CE, since its focus on changing gene transcription through β-catenin is designed to generate global, rather than local, signaling outcomes. However, an interplay between canonical Wnt signaling and cell-cell adhesion has been proposed [239]. Since β-catenin functions as an adaptor protein in both cadherin-mediated adhesion and canonical Wnt signaling, it is possible that polarized Wnt signaling could impact properties of cell-cell adhesion on a local

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basis, an impact that would change interactions with global changes in actin dynamics [240]. As mentioned, Dsh is a critical node in Wnt signaling, as it serves as a core component of both canonical/ β-catenin-dependent signaling as well as non-canonical PCP signaling. Nodal has very recently been identified as another upstream regulator of both CE and gastrulation, modulating the migration and intercalation of mesoderm cells contributing to the notochord [241], though no direct link to PCP signaling could be shown. In the related developmental process of anterior visceral endoderm (AVE) morphogenesis in mouse, however, Nodal signaling was shown to regulate Dsh membrane localization [242], a process thought to be critical for PCP activity [243]. This new evidence points to a possible Nodal requirement for Dsh membrane localization and proper PCP signaling in CE/gastrulation. Finally, a recent paper studying core PCP components revealed novel molecular interplay between Wnt, Ror2 (an alternative receptor for PCP Wnt signaling), and Vangl2, and identified mechanisms wherein Vangl2 phosphorylation levels, as regulated by Wnt signaling, could serve as the key determinant of planar cellular polarity [244]. While this study focused on cell polarity of limb bud chondrocytes and limb bud outgrowth and patterning, the general principles of PCP pathway function described here could clarify how PCP signaling drives CE in epithelial tissues. At the heart of PCP signaling is the ability of Dsh to activate Rho GTPase family members. In the current model of PCP signaling in CE/gastrulation, Wnt/Frizzled signaling via Wnt5/11 and Fz3/7 activate Dsh in its non-canonical role, where the relative activity of Rac1, RhoA, and Cdc42 are altered to regulate changes in actin dynamics. Thus, PCP signaling could be directly and locally influencing actin reorganization, contractility, and linkages with adhesion systems to determine the direction of CE events. Another candidate for localized or polarized signaling that could be used to orient actin dynamics to specific cell-cell junction in CE is the Wnt/Ca+ pathway. Little is currently known about calcium influxes initiated by Wnt signaling. For calcium influxes to generate asymmetrical actin dynamics during CE, such influxes must be locally restricted within the plasma membrane. In support of this idea, calcium influxes have been observed in membranes of cells undergoing epithelial morphogenesis events in tissue culture model systems [245, 246]. In addition to Wnt signaling, cadherins have long been associated with cell adhesion/migration in development, including an adhesive role in CE and gastrulation. The role of cadherins in CE may require their signaling function, which includes the ability of cadherins to alter the local activity of Rho GTPases [247]. However, it remains unclear how cadherin signaling might be polarized to generate asymmetric changes in actin dynamics. New data also suggests that ephrin and ephrin receptors act to maintain germ layer separation and contribute to cellular polarity, migratrion, and convergence during gastrulation [248]. This study provides additional evidence that convergence of cells to the midline and axis elongation may be independently regulated processes, since blocking EphA4 primarily impacted cellular convergence. The cytoplasmic targets of ephrin signaling also include Rho GTPases, further demonstrating the central position of these proteins in converting cellular signaling events into changes in actin dynamics that drive epithelial morphogenesis. In the case of CE, it is likely a complex balance between signaling emanating from ephrin and ephrin receptors, cadherins adhesion receptors, Wnts, and additional signaling networks that correctly modulate Rho GTPases, which in turn regulate actin dynamics that are required for the physical events of CE.

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Conclusion Epithelial morphogenesis includes a wide array of tissue remodeling events. A unifying principle is that actin dynamics in individual cells drive the changes in cell shape and motility that underlie all epithelial morphogenesis programs. We propose that changes in actin organization, contractility, and linkages to adhesion systems form the basis of how individual cells interact to generate specific epithelial morphogenesis outcomes. Connections of actin filaments to cell-cell adhesion systems play the major role in determining whether actin dynamics in individual cells are collective or independent. Maintenance of actin connections to adhesion systems results in actin reorganization and contractility events being transduced through adjacent cells and generating larger effects on tissue architecture. An uncoupling of actin connections with cell-cell adhesion systems prevents transduction of actin dynamics between adjacent cells. Cells, therefore, alter shape and motility in solitude. While much remains to be discovered about the molecular architecture of actin-membrane connections at sites of cell-cell adhesion, our understanding of actin dynamics in tissue remodeling makes several key predictions possible. First, the ability of actin-membrane connections to undergo rearrangements during collective epithelial morphogenesis movements, such as those that must occur in convergent extension, suggests that connections are highly dynamic, with a fast turnover rate. The second prediction is that specific actin-membrane connections can be reinforced and stabilized. We propose that tension forces on connections between actin filaments and cell-cell adhesion systems facilitate molecular events that reinforce these connections. In this view, cell-cell adhesion could occur in weak and strong phases, depending on whether actin-membrane connections had been reinforced. In this light it is interesting that a strong and weak phase of cell-cell junction formation is observed and that perturbation of protein function can alter the ability of cells to transition from the weak to the strong state [35, 39]. Further supporting this idea is the localization of tension sensing proteins to cell-cell junctions or the identification of biochemical activities that are regulated by tension in certain cell-cell junction proteins, most notably zyxin, vinculin, and α-catenin [85, 233-235]. After modulation of actin-membrane connections at cell-cell junctions, actin organization and contractility account for the remaining variability of outcomes in epithelial morphogenesis. Contractility generates forces that deform cell and tissue shape. Actin organization determines the direction contractility forces will be applied. A simple comparison of how contractility and actin reorganization are different in convergence extension and tissue bending highlights this principle. In the current and widely-accepted model for convergence extension, contractile forces are thought to occur without dramatic actin reorganization and, given the actin organization of cells in a stable epithelium, are thus applied parallel to cell-cell junctions. In tissue bending events, actin reorganization precedes contractility. The result is that a new organization constrains that the forces of contractility are applied perpendicular to the cell-cell junctions. In convergence extension the result is shortening of cell-cell junctions. In tissue bending the result is the transition of cell shape from cylindrical to conical. Unfortunately, this popular conceptual model may oversimplify what really occurs in convergent extensions. An important report suggests that tissue remodeling in convergent extension occurs following reorganization of actin into a medial actin network and that contractile forces are somehow applied to specific cell-cell junctions

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and not adjacent ones [14]. This new view is more in keeping with our idea that all epithelial morphogenesis events, including convergent extension, occur with a conserved series of actin dynamics, namely and in order, reorganization of actin structure, contractility of actin networks, and regulation of actin linkages to cell adhesion systems. In our proposed working model, changes in the subcellular localization of actin dynamics that universally underlie epithelial morphogenesis account for variability of outcomes on tissue organization. Whether changes in actin dynamics are asymmetric in relation to the apicobasal or planar cell polarity axes generates the actin organization and contractility can account for the entire range of variability in epithelial morphogenesis outcomes. This highlights the role of signaling pathways in controlling epithelial morphogenesis, particularly pathways that exert local or polarized signaling. How these pathways interface with cellular machinery that controls action dynamics is only beginning to emerge. Central players appear to be small GTPases of the Rho family, and particularly the interplay between Cdc42 and RhoA that serves to restrict the subcellular localization of myosin-based contractions. In fact, it is entirely possible that actin reorganization occurs in a strikingly similar manner in all instances of epithelial morphogenesis and that it is only spatiotemporal control of contractility that determines how actin rearrangements impact tissue architecture. Much research attention is still needed to test ideas that will demonstrate the complex interactions between actin dynamics, cell-cell adhesion, cell polarity, and epithelial morphogenesis.

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[220] P. Ybot-Gonzalez, C. Gaston-Massuet, G. Girdler, J. Klingensmith, R. Arkell, N.D. Greene, A.J. Copp, Neural plate morphogenesis during mouse neurulation is regulated by antagonism of Bmp signalling, Development 134 (2007) 3203-3211. [221] M.K. Nyholm, S. Abdelilah-Seyfried, Y. Grinblat, A novel genetic mechanism regulates dorsolateral hinge-point formation during zebrafish cranial neurulation, J. Cell Sci. 122 (2009) 2137-2148. [222] D. Rashid, K. Newell, L. Shama, R. Bradley, A requirement for NF-protocadherin and TAF1/Set in cell adhesion and neural tube formation, Dev. Biol 291 (2006) 170-181. [223] A.J. Chalpe, M. Prasad, A.J. Henke, A.F. Paulson, Regulation of cadherin expression in the chicken neural crest by the Wnt/beta-catenin signaling pathway, Cell Adh Migr 4 (2010) 431-438. [224] E. Hong, R. Brewster, N-cadherin is required for the polarized cell behaviors that drive neurulation in the zebrafish, Development 133 (2006) 3895-3905. [225] C. Fournier-Thibault, C. Blavet, A. Jarov, F. Bajanca, S. Thorsteinsdottir, J.L. Duband, Sonic hedgehog regulates integrin activity, cadherin contacts, and cell polarity to orchestrate neural tube morphogenesis, J. Neurosci. 29 (2009) 12506-12520. [226] X. Yang, J. Zou, D.R. Hyde, L.A. Davidson, X. Wei, Stepwise maturation of apicobasal polarity of the neuroepithelium is essential for vertebrate neurulation, J. Neurosci. 29 (2009) 11426-11440. [227] M. Zigman, A. Trinh le, S.E. Fraser, C.B. Moens, Zebrafish neural tube morphogenesis requires Scribble-dependent oriented cell divisions, Curr. Biol. 21 (2011) 79-86. [228] K. Chalasani, R.M. Brewster, N-cadherin-mediated cell adhesion restricts cell proliferation in the dorsal neural tube, Mol. Biol. Cell 22 (2011) 1505-1515. [229] M.G. Farquhar, G.E. Palade, Junctional complexes in various epithelia, J. Cell Biol. 17 (1963) 375-412. [230] P. Ybot-Gonzalez, D. Savery, D. Gerrelli, M. Signore, C.E. Mitchell, C.H. Faux, N.D. Greene, A.J. Copp, Convergent extension, planar-cell-polarity signalling and initiation of mouse neural tube closure, Development 134 (2007) 789-799. [231] C. Yin, B. Ciruna, L. Solnica-Krezel, Convergence and extension movements during vertebrate gastrulation, Curr. Top. Dev. Biol. 89 (2009) 163-192. [232] H. Hirata, H. Tatsumi, M. Sokabe, Zyxin emerges as a key player in the mechanotransduction at cell adhesive structures, Commun. Integr. Biol. 1 (2008) 192195. [233] H. Hirata, H. Tatsumi, M. Sokabe, Mechanical forces facilitate actin polymerization at focal adhesions in a zyxin-dependent manner, J. Cell Sci. 121 (2008) 2795-2804. [234] M. Yoshigi, L.M. Hoffman, C.C. Jensen, H.J. Yost, M.C. Beckerle, Mechanical force mobilizes zyxin from focal adhesions to actin filaments and regulates cytoskeletal reinforcement, J. Cell Biol. 171 (2005) 209-215. [235] Q. le Duc, Q. Shi, I. Blonk, A. Sonnenberg, N. Wang, D. Leckband, J. de Rooij, Vinculin potentiates E-cadherin mechanosensing and is recruited to actin-anchored sites within adherens junctions in a myosin II-dependent manner, J. Cell Biol. 189 (2010) 1107-1115. [236] E. Agius, M. Oelgeschlager, O. Wessely, C. Kemp, E.M. De Robertis, Endodermal Nodal-related signals and mesoderm induction in Xenopus, Development 127 (2000) 1173-1183.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 61-96

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter II

Actin: Structure, Function and Disease YamilaTorres Cleurena and Johannes Boonstraa,b a

University College Utrecht, 3508 TC Utrecht b Cell Biology, Department of Biology, 3584 CH Utrecht, the Netherlands.

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Abstract Actin is a globular protein found in all eukaryotic cells. Depending on its location, it can form different structures and perform various functions. Actin monomers (G-actin) come together to form filaments (F-actin); it is found abundantly in the form of microfilaments and thin filaments in cells. With the help of different actin-binding proteins that regulate its structure and activity, it can assemble in several combinations giving rise to actin bundles and networks with differing functions. Playing a central role in cell morphology, cell adhesions, cell contractility and motility, signal transduction, transcription and its regulation, cytokinesis and synapse formation, malfunctioning of actin can lead to various diseases; among them congenital myopathies, compromised immunity, neurodegeneration, and cancer spread. The structure and function of actin and its role in different diseases are here discussed.

Introduction Actin is much more than just the protein known for its activity in muscle cells; it is present in all eukaryotic cells, in the cytoplasm and in the nucleus, holding up different structures or giving them the opportunity to move around. It is present in many forms due to the regulation of the different actin binding proteins. In this review, the structure of actin and its dynamics will firstly be discussed, followed by the different functions actin can have in the cells. Lastly, the most important actin-related diseases will be discussed and related to the 

Address correspondence: Prof. Dr. J. Boonstra, Cell Biology, Padualaan 8, 3584 CH Utrecht, The Netherlands, email: J. [email protected].

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corresponding function of actin. In these sections, the different sides to actin are revealed and how this little protein give cells their most prominent features, such as crawling or signaling. But these properties are also given to other “unwanted” cells, cancerous cells, giving them the option of detaching and migration to other tissues. How can actin give cells such power? It is all in its functions.

Actin Structure

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Actin Monomers and Filaments Monomeric or globular actin is known as G-actin. It is a rather flat molecule, comparable with a prism, with dimensions 55Å x 55Å x 35Å. There are three isoforms: α, β and γ, differing only in a few amino acids, especially towards the N terminus. α actin is mainly present in muscle cells (skeletal, smooth and cardiac), being the major constituent of the contractile apparatus (Herman, 1993). β and γ actin are present in most cells and form part of the cytoskeleton, mediating internal cell motility as well. Actin undergoes posttranslational modifications that will determine its different properties, but even though 80 different structures of actin have been reported, the actin monomer conformation is essentially the same (for review see Dominguez and Holmes, 2011). The polypeptide chain of actin is composed of 375 amino acids and folds into two major α/β-domains also known as the small and large domains (Hild et al., 2010). However, traditionally, it has been divided into four subdomains: subdomains 1 and 3 are structurally related, while 2 and 4 are inserted into subdomains 1 and 3, respectively. There is little contact between subdomains 1 and 3, having a sort of hinge between them that gives rise to two clefts between the domains (see Figure 1) (Dominguez and Holmes, 2011).In these clefts, a tightly bound adenosine-derived nucleotide is found forming a complex in the physiological state of actin with magnesium (Hild et al., 2010). The upper cleft provides an important linkage between the domains while the lower cleft is the major binding site for most actin binding proteins (ABPs) (see Figure 1). Although not very effective, G-actin has an intrinsic ATPase activity (Ha and McKay, 1994). G-actin can be bound to either ADP or ATP (see Figure 1). ATP is very important for holding the G-actin structure together; the ADP/G-actin form has a much lower affinity and stability, and is lost from the polymerized filament (Asakura and Oosawa, 1960). In fact, most structures found in vitro are in the ATP-bound form as the ADP state is less stable (Dominguez and Holmes, 2011). In other words, ATP is crucial for the regulation of polymerization and depolymerization of actin filaments. F-actin filaments are a double-stranded, right-handed helix formed from G-actin under physiological ionic conditions (Hild et al., 2010). The atomic structure of the actin filament is not known, but it is known that F-actin exists in a dynamic steady state depending on the concentration of monomeric actin (Furukawa and Fechheimer, 1997). G-actin bound to ATP start to assemble into dimers and trimers, forming stable oligomers before becoming filaments. Different processes take place along the growing filament for which the polarity of the filament is very important. First, the ATP-bound form G-actin comes to the fast-growing barbed (or +) end of the filament. Second, along the filament hydrolysis of the G-actin bound

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ATP occurs. And third, ADP-bound G-actin dissociates from the pointed (or -) end (see Figure 2). This is known as treadmilling, a process tightly regulated by different ABPs (Hild et al., 2010; Dominguez and Holmes, 2011). This process is very dynamic, being able to either form or dissociate the filament.

2

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3

Figure 1. G-actin monomer.The G-actin monomer is formed by 4 subdomains, as represented in the top of the figure. 1 and 3 are the main domains, contacting each other through a hinge, giving rise to two clefts for binding. In the upper cleft, ATP or ADP can bind, while in the lower cleft, actin-binding proteins (ABPs) can bind. The upper cleft is very important in holding the structure together, which is more stable when ATP is present.

Organization of Actin Filaments Actin filaments within the cell are known as microfilaments, the thinnest filaments of the cytoskeleton with a diameter of 80Å (Ikawa et al., 2007), and can come together to form higher-order structures: actin bundles or actin networks. In association with myosin, actin filaments are responsible for most types of cell movements (among others muscle contraction

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and cell division). Myofibrils are the main structure responsible for muscle contraction and are formed by actin filaments (thin filaments) together with myosin fibrils (thick filaments).

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Figure 2. Cycle of addition of ATP/G-actin subunits, hydrolisis of ATP and disassembly of ADP/Gactin along the F-actin filament. This is a highly dynamic process. ATP/G-actin is added to the barbed end of F-actin, while along the filament ATP is being hydrolyzed. At the other end of the filament, the pointed end, ADP/G-actin is released, so that G-actin is recycled. If the ADP bound to G-actin is exchanged for ATP, G-actin can reenter the cycle and be part of the filament.

Individual actin filaments can cross-link with each other and make two types of formations: actin bundles and actin networks. Whether one organization or the other is achieved depends greatly on the influence of actin-binding proteins. In the bundles, filaments cross-link tightly into parallel arrays while in the networks they form 3D structures. Bundling is tightly regulated by actin-bundling proteins (e.g. Arp2/3); their cross-linking will lead to the formation of actin bundles (Furukawa and Fechheimer, 1997). They can appear in different forms in the presence of F-actin: bound to F-actin or in a free state in solution (Furukawa and Fechheimer, 1997). Changes in free energy govern this process, as a more organized system is less stable than an organized system according to the second law of thermodynamics (Furukawa and Fechheimer, 1997). In the networks, actin filaments are cross-linked with the help of large actin-binding proteins (such as Filamin A) to form orthogonal filaments and create loose 3D meshworks, resembling semisolid gels. These networks together with the associated actinbinding proteins form the cell cortex; the underlying support structure of the plasma membrane, determining cell shape and allowing movement and dynamic changes in this area to occur. Further in this review the functions of actin on the cell membrane and the different existing structures will be discussed in more detail.

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Actin Metabolism and Regulation The actin cytoskeleton has very complex dynamic properties which are tightly regulated at different levels by several proteins that control actin polymerization, severing of actin filaments and crosslinking of actin filaments into networks (Uribe and Jay, 2009). Many of these actin-binding proteins (ABPs) have actin binding domains composed of two calponin homology (CH) regions. This domain is formed by about 110 amino acid residues found in cytoskeletal and signal-transduction proteins (Stradal et al., 1998). Many ABPs bind to the same loci on the surface of actin, competing with each other. However, some can bind cooperating with other proteins, forming ternary complexes (Dos Remedios et al., 2003). ABPs can be membrane-associated proteins (e.g. vinculin), membrane receptors or ion transporters (e.g., 43K protein), sarcomeric proteins (e.g. tropomyosin), toxins (e.g. iota toxin) or even drugs (e.g. Latrunculin A). Classification of ABPs can be done in different ways, here it is classified in 7 groups, depending on their function: monomer-binding proteins (e.g. thymosin β4 and DNase I), filament-depolymerizing proteins (e.g. cofilin), filament endbinding proteins (e.g. tropomodulin), filament severing proteins (e.g. gelsolin), cross-linking proteins (e.g. Arp2/3), stabilizing proteins (e.g. tropomyosin), and motor proteins (myosin family) (Dos Remedios et al., 2003).

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Monomer-Binding Proteins Monomer-binding proteins sequester G-actin and stop its polymerization. Thymosin β4 (Tβ4) binds to and controls the availability of actin monomers for their incorporation into filaments (Dos Remedios et al., 2003). DNase I is known for cleaving double-stranded DNA to yield 5‟-phosphorylated polynucleotides, leading to apoptosis of cells. However, another function related to the cytoskeleton is also found for DNase I; it binds G-actin very tightly, removing all free actin monomers from solution (Dos Remedios et al., 2003). Filament-Depolymerizing Proteins Depolymerizing proteins can cause the conversion of F-actin to G-actin. Cofilin is the primary player in actin filament disassembly. It does so from the pointed (or minus) end. Cofilin can sever actin filaments, accelerating actin monomer release, and thus by creating more filament ends, increasing the rate of actin assembly/disassembly. Many of these steps can be regulated by factors such as the Rho family of GTPases, Ca2+, and PtdI(4,5)P2 (Uribe and Jay, 2009). Dephosphorylation of cofilin at serine 3 leads to its activation, its translocation into the nucleus and is correlated with cellular transformation. Its role in the nucleus is not clear, but its accumulation in the nucleus in transformed cells favors depolymerization of F-actin, suggesting a nuclear function of actin. In addition, actin can form bundles with ADF/cofilin in a 1:1 ratio, so-called rods. These are formed in all types of cells, but especially in axons and dendrites of stressed neurons (because of ATP-depletion), which can lead to their dysfunction by blocking transport and synaptic activity (Minamide et al., 2009). Filament End-Binding Proteins The ends of actin filaments are capped by filament end-binding proteins in order to prevent the loss of monomers at the pointed and at the barbed end. Tropomodulin (Tmod) is

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one of the few proteins that bind to the pointed end of F-actin, stopping filament elongation and promoting the hydrolysis of ATP-actin to ADP-actin (Rickard and Sheterline, 1986). It binds to actin only in the presence of tropomyosin. On the other side, CapZ binds to the barbed end of F-actin. It has several biological roles, but its main function is its binding to monomers or oligomers of actin, forming nuclei for elongation and therefore facilitating polymerization (Dos Remedios et al., 2003).

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Filament Severing Proteins F-actin can also be severed, shortening the average length of filaments by binding of ABPs to its side and cutting it into two pieces (Dos Remedios et al., 2003). The gelsolin superfamily includes several ABPs (e.g. gelsolin, villin, adseverin, CapG, flil, and severin). They are all grouped as they all have three or six gelsolin repeats (Kwiatkowski et al., 1986). Intracellular gelsolin can circulate in blood plasma, severing and capping actin filaments that have been released into the circulation after cell death. Actin monomers are then sequestered by vitamin D-binding protein (Heling et al., 2000). Gelsolin gets its name for its effect in vitro, transforming gels with actin into a sol. As mentioned previously, cofilin also belongs to the group of severing proteins. Cross-Linking Proteins To form actin bundles or networks, cross-linking is necessary. Cross-linking ABPs contain at least two binding sites for F-actin that facilitate this process. The Arp2/3 complex creates branch points in actin filaments by nucleating filament assembly near ruffling membranes and can also cross-link these branches, creating different complex structures (Dos Remedios et al., 2003). Arp2/3 works together with cofilin to reorganize the actin filaments. Arp2/3 binds F-actin at the barbed end, causing the nucleation of a new F-actin branch, while cofilin causes depolymerization at the other side (Uribe and Jay, 2009). Filamin A (FLNA) is a homodimeric F-actin cross-linker that organizes actin filaments into parallel arrays or 3D webs, linking them to cellular membranes. It anchors several transmembrane proteins to the actin cytoskeleton (such as β-integrins), providing a scaffold for many cytoplasmic and signaling molecules (Uribe and Jay, 2009; Pudas, 2006). Thus, it regulates actin cytoskeleton organization by interacting with integrins, transmembrane receptor complexes, and second messengers. α-actinin has different isoforms, with α-actinin-4 cross-linking actin filaments and reorganizing the cytoskeleton for cell movement. It has been observed that α-actinin-4 is also present in the nucleus of a particular population of breast cancer cells (Uribe and Jay, 2009). In different cell types, it is found to have different roles. In nonmuscle cells, it is found at adherens junctions, binding actin to the membrane. In muscle cells, it can be found at the Z-disc, helping the actin filaments to anchor. Spectrin lines the intracellular part of the plasma membrane, forming a scaffold and playing an important role in maintenance of cytoplasm integrity and cytoskeletal structure. It can be arranged in an hexagonal way; tetramers of spectrin associate with short actin filaments at either end of the tetramer. In the nucleus, it crosslinks actin filaments as well and can be found at the nuclear envelope and in intranuclear granules (Uribe and Jay, 2009).

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Stabilizing Proteins Other proteins can bind to the sides of actin filaments to prevent depolymerization. In other words, they stabilize the actin filaments. Tropomyosin is one of such proteins. It competes with cofilin for binding on actin (Blanchoin et al., 2001). Thus, the depolymerizing activity of cofilin is stopped by tropomyosin. Drebrin is involved in the regulation of actin filament organization, especially during the formation of neurites and cell protrusions of motile cells. Drebrin A is localized at dendritic spines of mature cortical neurons, inhibiting the activity of tropomyosin, fascin and α-actinin, and the interaction between actin and myosin in vitro (Hayashi et al., 1996). Upregulation of drebrin leads to an elongation of dendritic spines of cortical neurons (seen in carcinomas), while its downregulation suppresses the accumulation of F-actin in the dendritic spines, changing their morphology (seen in Alzheimer‟s disease and Down Syndrome) (Dun and Chilton, 2010; Ivanov et al., 2009). Motor Proteins The myosin family of motor proteins can harness the polymerization of actin to produce movement. Myosin is an ATPase that moves along actin filaments, coupling the hydrolysis of ATP to conformational changes (DePina and Langford, 1999). In other words, actin provides the tracks on which myosin can move, with ATP being the required fuel. In later sections, the role of actin in movement will be discussed in more detail.

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Concluding Remarks The structure of actin monomers gives rise to two clefts in which ATP or ADP and actinbinding proteins (ABPs) can bind, modifying the fate of actin. With ATP bound, actin is much more stable, and polymerization of F-actin is facilitated. F-actin filaments assemble dynamically from G-actin monomers; at the barbed end monomers bound to ATP are added, while along the filament hydrolysis of ATP occurs, leaving monomers bound to ADP which will dissociate at the pointed end of the filament. But actin can also form much more complicated structures (networks and bundles) with the help of ABPs with very different functions which interact with each other to coordinate the process of actin assembly/disassembly and its directionality.

Actin Function Structural Function Cell Morphology The shape of animal cells is determined by the organization of internal structural elements, which includes the filamentous structures of the cytoskeleton, and external conditions. Actin filament dynamics influence the cell shape by pushing on the plasma membrane, creating tension. Actin is responsible for the particular shape of a cell as has been demonstrated that local effects of actin remodeling can manifest as global shape modifications of the cell (Lacayo et al., 2007). In fact, loss of actin can cause deformities.

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Plasma Membrane Underlying the plasma membrane are the peripheral proteins (bound to it by electrostatic interactions and hydrogen bonding with the hydrophilic phospholipid heads), where the cytoskeleton can be found. It provides the membrane with a scaffold for proteins to anchor to by interacting intimately with the cell membrane (Doherty and McMahon, 2008). Moreover, it is able to form appendage-like organelles that extend from the cell. In the case of the structures based on actin, filopodia are formed. These structures allow the cell to sense the external environment and/or make new contacts with the environment.

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Actin and Signal Transduction The first evidence of actin being related to signal transduction came from studies on the effects of growth factors on cell morphology, in which EGF produced the formation of membrane ruffles due to actin polymerization (Chinkers et al., 1979; Boonstra and Moes, 2005). It was later found that the EGF receptor was bound to actin (Wiegant et al., 1986; den Hartigh et al, 1992). Stimulation of cells with EGF causes rapid actin polymerization, formation of membrane ruffles and translocation of various downstream signaling proteins to the newly formed membrane ruffles, forming potential signaling complexes at the plasma membrane (Diakonova et al., 1995). Other signal transduction proteins associated with actin microfilaments are phosphoinositide kinase, diacylglycerol kinase, phospholipase C, and Akt/PKB (Janmey, 1998; Payrastre et al, 1991). Thus, actin might play an important role in growth factor-induced signal transduction. The Rho family of small G-proteins is tightly coupled to actin dynamics and its association with the membrane, regulating the addition of actin monomers at the barbed end (Uribe and Jay, 2009). They coordinate the cytoskeletal and adhesion modeling within cells, processes crucial for migratory responses. Rho GTPases have been shown to be the main actin regulators in neural and non-neural cells (Govek et al., 2005; Jaffe and Hall, 2005). They can regulate actin through effectors that can directly bind to or activate more downstream actin binding proteins (Taka et al., 2001). The most known Rho GTPases are RhoA (leads to stress fiber formation, focal adhesion formation and acto-myosin contraction), Rac (induces lamelipodial formations) and Cdc42 (induces filopodial formation) (Dubreuil and Van Vactor, 2011). Depending on what effectors are activated, signaling by RhoA can lead to different effects: actin polymerization, depolymerization and acto-myosin contraction. ROCK is the best-characterized Rho associated kinase that interacts with GTP-bound active RhoA, inducing acto-myosin contraction through phosphorylation of myosin light chain (MLC) and myosin light chain phosphatase (MLCP), thereby inducing MyosinII ATPase activity and promoting its association to actin filaments (Dubreuil and Van Vactor, 2011). P140mDia from the formin family stimulates actin polymerization. After activation by GTP-bound RhoA, it can bind profilin (nucleating actin) and IRSp53 (assembling actin). Rac induces actin polymerization through p21-activated kinase (PAK) and WAVE (WASP-verprolin homologous). PAK leads to actin reorganization through LIMK phosphorylation and inhibition of cofilin, enhancing actin filament elongation (Dubreuil and Van Vactor, 2011). PAK can also affect contraction in two ways. The first, by phosphorylating and inactivating MLCK to restrict myosin contraction, and the second by

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phosphorylating MLCII, inducing actomyosin contractility (Zhao and Manser, 2005; Dubreuil and Van Vactor, 2011). WAVE does not directly interact with Rac but forms a complex instead that will lead to actin polymerization. Moreover, Rac binds to the previously described Arp2/3, also leading to actin polymerization. Cdc42 can lead to actin polymerization via the effector proteins PAK, WASP (WiskottAldrich syndrome protein) and IRSp53. Activation of PAK occurs in a similar way as with Rac, affecting cofilin phosphorylation which induces actin polymerization (Zhao and Manser, 2005; Dubreuil and Van Vactor, 2011). Cdc42 can also induce polymerization by the Arp2/3 complex through activation of WASP via direct binding (Tomasevic et al., 2007). Polymerization through IRSp53 occurs with the interaction with Mena (mammalian Ena). These proteins antagonize the capping proteins at the barbed ends of actin filaments, ensuring actin polymerization into long unbranched filaments (Dubreuil and Van Vactor, 2011).

Cell Tension

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The cytosol applies a pressure on the plasma membrane, contributing to its cell tension. This tension is modulated by the strength of membrane-cytoskeleton interactions (Dai and Sheetz, 1999). During actin filament polymerization, the filaments push the cell membrane from within, generating tension that is rapidly distributed across the lipid bilayer (Raucher and Sheetz, 2000). Membrane tension provides a resistance that has to be overcome if any deformation takes place, such as endo- or exocytosis. While exocytosis decreases membrane tension by increasing local surface area and reducing the cytosolic pressure, endocytosis increases the tension by the contrary movement (Dai et al., 1997). The endocytic machinery must overcome the great resistance to deform the plasma membrane. When successful, it results in an increased cytosolic pressure and reduced surface area, inhibiting further endocytosis (Itoh et al., 2005).

Endocytosis Endocytosis is the process in which the plasma membrane invaginates into the cell, producing a vesicle that will be able to fuse with endosomes and enter the endo-lysosomal membrane system (Smythe and Ayschough, 2006). In endocytosis, these vesicles formed are coated with clathrin, which forms domains on the plasma membrane called clathrin-coated pits, which can concentrate different receptors for the endocytosis of ligands. Actin structures help in the remodeling of the cell surface, allowing the inward movement of vesicles. There are several routes that can provide the directional changes in morphology observed at endocytic sites. Actin covering the cytoplasmic side of the plasma membrane‟s surface provides it with mechanical flexibility. Two roles have been argued for actin that allows for endocytic vesicle formation: actin polymerization in the provision of force for membrane deformation, and actin depolymerization to remove it from an otherwise very tensive barrier (Smythe and Ayschough, 2006). Thus, actin plays a very helpful role in endocytosis; actin polymerization facilitates endocytosis, increasing their speed of formation, providing directionality and a mechanical resistance to relaxation of the newly deformed membranes (Doherty and McMahon, 2008). Moreover, connection of the border of a pit that is being

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formed to actin can allow actomyosin help pull in the deforming membrane (Buss et al., 2001) and provide the tract to allow the trafficking of these vesicles. Actin polymerization at the neck of the coated pit helps the membranes of the site to come together, aiding membrane fission (Doherty and McMahon, 2008).

Protrusions of Cell Membrane

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Movement of cells begins with the formation of protrusions of the cell membrane and then adhesions at the cell front, linking the actin cytoskeleton to the substratum, traction forces moving the cell forward, and disassembly of adhesions at the cell rear (Parsons et al. 2010). These protrusions comprise large filopodia that are driven by the polymerization of actin filaments (Pollard and Borisy, 2003). The fast growing rate of actin filaments is very important for the success of cell movement. Cells contain a pool of actin monomers bound to profilin. When signaling pathways activate nucleation-promoting factors (such as members of the WASP family) and these stimulate Arp2/3 complex to initiate a new branch on the filament, these filaments grow and push the membrane forward, and thus allowing for movement. Protrusions are stabilized by adhesions that link the actin cytoskeleton to the ECM proteins, also allowing the actomyosin contraction to occur, generating traction forces on the substratum. With contractility, adhesions must be disassembled at the cell rear so that the cell can move forward. There are bidirectional interactions coordinating these adhesion changes: signals from new and stable adhesions influence cytoskeletal organization and actin polymerization, while cytoskeletal structures affect the formation and disassembly of the adhesions (Ridley et al., 2003). Adhesion formation, maturation and assembly is a continuous process driven by the balance of actin polymerization and actomyosin contraction (Parsons et al., 2010).

Cellular Adhesions Correct cellular adhesions are essential in maintaining multicellular structure, given that they bind the cell to another cell surface or extracellular matrix. They are especially important anchoring points between epithelial cells, where various types can be found: desmosomes, hemidesmosomes, adherens junctions, tight junctions and focal adhesions (Buckley et al., 1998). The adhesive structures are connected either to intermediate filaments (desmosomes and hemidesmosomes) or to actin filaments (adherens junctions, tight junctions and focal adhesions). This connection is needed for a stable adhesion formation. Even though adherens and tight juntions comprise different proteins, there is resemblance in the roles of specialized transmembrane proteins in forming extracellular adhesive contacts between cells and intracellular links to actin (Hartsock and Nelson, 2008). Adherens Junctions Adherens junctions are not just anchors to the environment; they have functions in the initiation and stabilization of cell-cell adhesion, regulation of the actin cytoskeleton, intracellular signaling and transcriptional regulation (Hartsock and Nelson, 2008). Classical

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cadherins are the major transmembrane proteins of the adherens junction. They initiate the intercellular contacts via trans-pairing between cadherins on opposing cells (Gumbiner, 2005). They can also bind to many cytoplasmic proteins, mainly the catenin family of proteins that locally regulate the organization of the actin cytoskeleton. The core of the adherens junction is indeed formed by the classical cadherin superfamily (such as E-cadherin) and the catenin family members (p120-catenin, β-catenin and α-catenin) (Hartsock and Nelson, 2008). Catenin provides the link between cadherin and the actin cytoskeleton. βcatenin connects the cytosolic parts of cadherins with α-catenin. While binding to β-catenin, α-catenin can also bind and bundle actin filaments. It exists in either a monomeric or homodimeric state. In this dimeric state, α-catenin is not able to bind β-catenin; and in the monomeric state, it is the other way around, it can bind β-catenin but not actin filaments. Homo-dimeric α-catenin binds actin filaments competing with the binding of the Arp2/3 complex to actin, thereby suppressing actin polymerization. Branching of the actin filaments is inhibited, and actin is reorganized from branched to bundled arrays (for review see Hartsock and Nelson, 2008).

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Tight Junctions Tight junctions, on their part, have two opposing functions: preventing the mixing of membrane proteins between the apical and basolateral membranes (in cells with a polarity) and controlling the paracellular passage of ions and solutes in-between cells (Hartsock and Nelson, 2008). In these junctions, occludins and claudins (transmembrane proteins) are associated with cytoplasmic proteins that can link them to the actin-cytoskeleton and the adherens junction. ZO-1, ZO-2 and ZO-3 (members of the membrane-associated guanylate kinase homolog family) can bind adherens and tight junction proteins and the actin cytoskeleton, and could potentially affect actin polymerization (Rajasekaran et al., 1996; Hartsock and Nelson, 2008).

0 Figure 3. Formation of cell-cell junctions.During cell-cell junction formation, actin is the main player. First, the epithelial cells extend protrusions. Second, cadherin comes to the tips of these protrusions forming “cadherinpuncta” and connecting to the circumferential actin via radial actin bundles. In some cells, keratinocytes, an intermediate configuration is adopted: the adhesion zipper, as a consequence of myosin-mediated tension pulling inward on cadherinpuncta. As the cell-cell contacts mature, actin arcs focus on the edges of the contact. This actin remodeling results in the formation of the adhesion belt.

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Cell-Cell Contact Formation Actin is the main player in the formation of cell-cell contacts. Before the process of contact formation starts, actin forms a circumferential actin cable or ring around the cells and a dense meshwork between the ring and the plasma membrane. As these contacts get formed, cadherin comes into play, forming cadherin puncta connected to the actin ring via radial actin bundles. Under these regions of contact, actin bundles then get replaced by thinner bundles forming the adhesion belt while the thick bundles focus on the contact edges (Cavey and Lecuit, 2009).

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Junction Stability The stability of the junctions depends on the integrity of the actin cytoskeleton (Pilot et al., 2006). Once a junction has been established, branched networks associated with lamellipodial protrusions are replaced by parallel contractile bundles. Factors promoting branched actin polymerization (Arp2/3 among others) are relatively depleted from “old” regions of cell contacts (Helwani et al., 2004). As the junctions mature, α-catenin is more present, controlling this transition by repression of Arp2/3 activity (Cavey and Lecuit, 2009). Myosin-II creates tension that also aids in the stabilization of the cell-cell junctions, inhibiting the formation of protrusions by its ability to align actin filaments parallel to the plasma membrane (Gloushankova et al., 1997). Moreoever, the tension along a cell can bring the cell membranes of two proximal cells into close contact, favoring new homophilic cadherin interactions to form and trapping these molecules in the region under tension (Cavey and Lecuit, 2009; Delanoe-Ayari et al., 2004). A disruption in the actin structure could lead to misregulated adhesions, affecting specially the permeability of endothelial cells, and thus leading to edema and increased interstitial pressure (Dejana et al., 2009) Remodeling of Junctions An important event that occurs is the remodeling of intercellular junctions; they are dynamic structures that can change not only upon exposure to agents increasing permeability but also in resting cells (Dejana et al., 2009). Cadherins (in particular VE-cadherin) can show a flow-like movement in a basal to apical direction which is accompanied by actin reorganization (Kametani and Takeichi, 2007). Although the junctional actin networks are very complex, a fraction of thin actin fibers run along the basal-to-apical axis of cell junctions associated with VE-cadherin clusters. It has been shown that VE-cadherin clusters move on actin bundles, being able to detach from one actin fiber and go on to another actin fiber (Kametani and Takeichi, 2007). Actin molecules dynamically reorganize themselves at the cell junctions. This actin flow depends on myosin II activities; actin filament turnover and organization are tightly controlled together with myosin-II activity to produce the mechanical forces that drive the assembly, maintenance, and remodeling of junctions (Cavey and Lecuit, 2009).

Actin in Vascular Smooth Muscle In vascular smooth muscle, actin filaments of smooth muscle cells are in close contact with dense plaques on the plasma membrane, linking to dense bodies in the myoplasm. These

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actin filaments concentrate around thick filaments of myosin in a hexagonal array, forming what is known as the contractile apparatus. The actin filaments in this apparatus are called the contractile actin, while the actin filaments that are not associated with myosin are called cytoskeletal actin, which maintains the structural integrity of smooth muscle cells (Tang and Anfinogenova, 2008). In the contraction-relaxation cycle of smooth muscle, actin filaments polymerize and depolymerize accordingly; when contraction occurs, the F-actin to G-actin ratio is increased from 1:8 to 5:8, going back down again when the muscle is relaxed (Tang and Anfinogenova, 2008). Cortical actin assembly can strengthen the linkage of actin filaments to integrins and enhance the transmission of contractile force. In addition, with actin assembly, the number of contractile units increases and also the length of the filaments, providing more and efficient contractile elements for force development (Tang and Anfinogenova, 2008). Regulation of Actin in Vascular Smooth Muscle ROCKs are important regulators of actin cytoskeleton and can mediate vascular function. Increased ROCK activity is observed in hypertension or vascular inflammation (Zhou et al., 2011). ROCKs are downstream targets of RhoA and mediate Rho-induced actin cytoskeleton changes through effects on myosin light-chain phosphorylation, facilitating the interaction of myosin with F-actin (Zhou et al., 2011). ROCKs also regulate the phagocytic activity of macrophages via actin cytoskeleton membrane protrusion and mediate cell permeability through effects on tight and adherens junctions (Zhou et al., 2011).

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Actin at Synapses Dendritic Synapses Neurons transmit information via specialized cell junctions called synapses. A tight control of the development and connectivity of synapses is of prime importance for good neural activity and brain functioning. These synapses are formed at tiny dendritic protrusions: the dendritic spines (Bourne and Harris, 2008). It is now well known that any changes in spine morphology account for functional differences at the synaptic level (Kasai et al., 2010). Dendritic spine formation and dynamics are determined by the actin cytoskeleton, as it is central to membrane dynamics. At synapses, actin does not only make up the structure of synapses but also organizes the postsynaptic density and anchors postsynaptic receptors to facilitate trafficking of synaptic cargos and localizing the translation machinery (Hotulainen and Hoogenraad, 2010). Therefore, various memory disorders involve defects in the regulation of the actin cytoskeleton, such as Alzheimer‟s and Parkinson‟s disease. Spines consist of three basic compartments: a δ-shaped base at the junction with the dendritic shaft, a constricted neck in the middle, and a bulbous head contacting the axon (Hotulainen and Hoogenraad, 2010). They contain the postsynaptic machinery; glutamate receptors, postsynaptic density (PSD), actin cytoskeleton, membrane-bound organelles. The PSD is an organizing structure that clusters receptors, adhesion molecules, and channels, assembling a great variety of signaling molecules (Hotulainen and Hoogenraad, 2010). Actin is the major cytoskeletal component of these spines, forming a network of long and short branching filaments in the spine neck and short-branched actin filaments in the spine head

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right under the PSD. In mature spines, actin stabilizes postsynaptic proteins (Renner et al., 2009) and modulates the spine head structure in response to postsynaptic signaling. Both Gactin and F-actin are present at the spines and as with vascular smooth muscle, the degree of actin polymerization (G-actin to F-actin ratio) affects dendritic spine morphology (Cingolani and Goda, 2008). Synaptic stimulation changes the equilibrium between F-actin and G-actin; long-term potentiation induces a shift towards F-actin, increasing spine volume, while longterm depression induces a shift towards G-actin, resulting in spine shrinkage (Okamoto et al., 2004). During dendritic spine enlargement, it has been found that three actin pools are dynamically regulating the structure and plasticity of the spines (Honkura et al., 2008). The actin pool at the tip forms filaments to generate an expansive force in the spine head, being the major determinant of spine volume. At the spine neck, actin filaments are arranged longitudinally. At this location, a mixture of branched and straight actin filaments is found. During neuronal development, thin dendritic filopodia can become more stable mushroom spines upon synaptic contact with the axon (Yoshihara et al., 2009). Dendritic filopodia initiate from preexisting patches of branched actin directly from the dendritic shaft. The actin-rich sites where this process is initiated becomes the base of the filopodia (Hotulainen and Hoogenraad, 2010). Dendritic filopodia formation can either be started randomly or by signal-induced initiation by glutamate or Arp2/3 complex. Once the dendritic filopodia are established and an axonal contact is made, their motility decreases to give place to the spine structure. At first, these spines are thin elongates with a small spine head, but thanks to the action of the Arp2/3 complex, actin polymerization takes place, allowing the spine head to grow. However, myosin II activity can modify the size and the shape of spines, and other actin cross-linking proteins can modify and stabilize the formation (Hotulainen and Hoogenraad, 2010). Even though numerous research has focused on actin, still the exact molecular mechanisms underlying spine development and plasticity need to be elucidated. Immunological Synapses When an immune response is mounted, cell-to-cell contacts need to be established between lymphocytes (naïve T cells, natural killer cells or B cells) and antigen-presenting cells (APCs: dendritic cells, macrophages, B cells and some target cells) (Gordón-Alonso et al., 2010). The immunological synapse is the structure formed by the interaction between lymphocytes and antigen-presenting cells. T cell receptors are stimulated during the formation of immunological synapses, producing rearrangements of actin at the T cell/APC contact zone to increase the contacting area. Accumulation of actin at the synapse must occur for lymphocyte activation (Gomez and Billadeau, 2008). The structure of the immunological synapse has been studied in vivo. It is formed by concentric rings of molecular aggregates. T cell receptors and major histocompatibility complexes (MHC) are concentrated in a central cluster, the central supramolecular activation cluster (cSMAC). This is surrounded by a ring outside that contains integrins and their ligands; the peripheral activation cluster (pSMAC). The pSMAC structure is supported by an F-actin ring (Dustin, 2009). Actin polymerization occurs early in the formation of the synapse, creating a lamellipodium at the intercellular contact zone, increasing the interacting surface area (Gordón-Alonso et al., 2010). Microfilament reorganization follows, creating a structure known as the “actin cloud”. Then, actin microfilaments can associate with several transmembrane proteins, moving to produce the observable segregation of proteins in concentric rings. After this segregation, actin cytoskeleton helps stabilize the synapse.

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Defective actin would lead to a deficient T cell activation, Ca2+ influx, cytokine secretion and T cell proliferation (Gomez and Billadeau, 2008).

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Nuclear Structure Actin helps assemble and maintain the nuclear envelope. Therefore, it interacts with the nuclear lamina and with a structural protein of the nucleus, the enhanced adult sensory threshold (EAST) endoskeleton (Bettinger et al., 2004; Wasser and Chia, 2000). EAST forms a network throughout the nucleus, excluding the chromosomes and the nucleolus (Wasser and Chia, 2000). Actin has been shown to interact with the C-terminal domain of lamin A and with emerin (a lamina-associated protein) (see Figure 4) (Fairley et al., 1999). These interactions result in the formation of a complex that stabilizes the nuclear envelope against mechanical stress (Holaska and Wilson, 2007). It has been demonstrated in Xenopugus laevis egg extracts; as chromatin assembled, actin accumulated and formed a network in the nuclei (Bettinger et al., 2004; Zhang et al., 1996). Actin is not only present in the nuclear envelope; it is also present in the matrix of the nucleus. It has been found to be present along the fibrogranular structures in nuclear matrix preparations (see Figure 4) (Nakayasu and Ueda, 1985). It plays a role in maintaining the structure of the nuclear matrix, the chromatin-free portion of the nucleus, and is thought to play a crucial role in maintaining the spatial order within the nucleus. Even though the existence of such a matrix is still controversial (Radulescu and Cleveland, 2010), there is a growing amount of data confirming an intranuclear filament network in the Xenopus model, and actin is an integral component (Kiseleva et al., 2004).

Figure 4. Actin in the nuclear structure. Actin interacts with the nuclear lamina and with a structural protein of the nucleus, the enhanced adult sensory threshold (EAST) endoskeleton. EAST forms a network throughout the nucleus, excluding the chromosomes and the nucleolus. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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Function in Motility Cellular Motility Directional cellular motility is an essential cellular process for many processes in which cells need to migrate in an organized fashion, e.g., embryonic development, immune responses, and development of tissues. Migration is also a very important feature of many diseases, including cancer and chronic inflammation. Knowing how all the different structures and regulators come into play can give insight into how these diseases can be tackled. Regulation of actin seems to be a key player and targeting its function could give some promising advances. Crawling motility of cells involves a four-step-cycle: protrusion of the leading edge, adhesion to the substratum, retraction of the rear, and de-adhesion (Pollard and Borisy, 2003). Lamellipodia on the cell cortex expand forward as the cell moves in a self-organized way, a kind of “molecular autopilot” (Pollard and Borisy, 2003). Actin is the most abundant structural component of lamellipodia.

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Membrane Deformation For movement to occur, the plasma membrane has to be deformed. They are predominantly flat structures, in their lowest energy state, thus energy must be supplied to allow membrane deformation. Polymerization of actin towards the membrane will provide force necessary to deform it. In fast moving cells, it has been demonstrated that actin remains stationary relative to the substrate while the cell advances, indicating that protrusion of the leading edge is coupled with polymerization of actin (Theriot and Mitchison, 1991). If a cell is static, actin filaments assemble at the margin of the cell and move away from the edge, reflecting the same relationship to the cell surface as in locomotion. However, the most common event is the transformation of actin polymerization into protrusions and partially into retrograde flow (Lin and Forscher, 1995). In the central and rear regions of migrating cells, actin filaments are organized into thick bundles; the stress fibers (Amano et al., 1997). Dorsal stress fibers connect with the substrate via focal adhesions at one end. Transverse arcs are formed (bundles of actin filaments formed parallel to the leading edge, undergoing retrograde movements towards the cell center) which are not anchored at first. When Rho is activated, ventral stress fibers arise from the already formed dorsal stress fibers and the transverse arcs, which get anchored to focal adhesions at both ends (Parsons et al., 2010).

Fast Polymerization of Actin Cells can produce barbed ends by three possible mechanisms: severing existing filaments, uncapping existing filaments, or de novo nucleation (Zigmond, 1996; Pollard and Borisy, 2003). The latter is considered to be the dominant mechanism of production in the leading edge. They contain a pool of unpolymerized actin monomers. New filaments arise when signaling pathways activate nucleation-promoting factors, such as the members of the

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WASp/Scar family of proteins (Pollard and Borisy, 2003). Active nucleation-promoting factors then stimulate Arp2/3 complex to initiate a new filament as a branch on the side of an existing filament. New branches grow rapidly and push the membrane forward. Actin monomers are being recycled constantly, following the cycle described above. Fast growing is a very important feature of actin polymerization. The rate of this process is proportional to the concentration of actin monomers. To achieve the highest rates, the concentration of actin monomers needs to be one hundred times the concentration of actin filaments (Pollard and Borisy, 2003). Cells have gained several mechanisms to maintain this pool of “polymerization-ready actin” at a high concentration; actin-binding and filamentcapping proteins cooperate in this process. Profilin binds to ATP-actin monomers at the barbed-end, allowing actin-profilin complexes to elongate the barbed ends of filaments at a normal rate but blocking the binding to filament pointed ends and the spontaneous nucleation of actin filaments. Thymosin-β4 associates to ATP-actin monomers as well. In platelets and leukocytes, the amount of unpolymerized actin exceeds the concentration of profilin, and so the thymosin-β4 makes up for the difference (De La Cruz et al., 2000). However, binding to thymosin-β4 is problematic, as it blocks all actin assembly reactions, stopping the growing of actin filaments. Profilin is more effective than thymosin-β4 in binding actin monomers. As both proteins exchange on and off actin very quickly, thymosin-β4 allows profilin to maintain a pool of actin ready for elongation while it holds the rest of the monomers in reserve (Pollard and Borisy, 2003). If this growth at barbed ends continues at a fast rate, it would deplete the unpolymerized actin pool very quickly, slowing down the process. Two mechanisms compensate for this depletion. On the one hand, there is capping of many barbed ends, reducing the rate of drawdown on the pool. It might seem counterproductive, but it is actually very useful to limit the lengths of the growing branches, as short filaments are stiffer than long filaments, and thus more effective at pushing on the membrane. Also, capping controls where the actin filaments “push”. Since only the barbed ends that contact the lamellipodial surface are effective generators of propulsive force, capping of barbed ends avoid the nonproductive consumption of actin subunits in other places of the cell. On the other hand, there are ADF/cofilin proteins that accelerate actin depolymerization, replenishing the monomer pool. Both these processes allow cells to maintain a high concentration of unpolymerized actin far from equilibrium (Pollard and Borisy, 2003). Moreover, directionality is provided for the growth of the actin filaments by cues from the plasma membrane. Their components transduce stimulatory cues from the exterior into PtdIns(4,5)P2/PtdIns(3,4,5)P3 accumulation, polarizing the assembly of actin (Doherty and McMahon, 2008).

Adhesion Assembly and Disassembly Apart from the formation of cell protrusions, adhesions are formed that will allow the cell to attach to the new surface. The earliest detectable adhesions form in the lamellipodium right behind the leading edge. Their assembly depends on the rate at which protrusions are being made at the leading edge with Arp2/3 complex-mediated actin polymerization. As the leading edge moves forward, new adhesions either elongate and grow or disassemble, depending on the cell type (Alexandrova et al., 2008; Choi et al., 2008; Parsons et al., 2010). Disassembly occurs when the new adhesions encounter the zone of depolymerizing actin at the juncture of

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the lamellipodium and lamellum. New adhesions can mature into focal complexes when a pause is taken from forward movement. How nascent adhesions are nucleated, elongate and disassemble is still not clear. Still, two possible not mutually exclusive models have been proposed. In the first, nucleation of adhesions is initiated by binding of integrins to proteins of the extracellular matrix. Ligand-mediated formation of clusters follows with a subsequent assembly of new adhesion complexes, ultimately linking to actin filaments (Parsons et al., 2010). In the second model, assembly is initiated by actin polymerization and uses dendritic actin as a template for the nucleation of adhesion complexes. To rearrange the cytoskeleton, myosin II comes into play; it moves antiparallel actin filaments past each other, it can bundle up the filaments and it can contribute to the maturation of newly formed adhesions to focal complexes and focal adhesions, by generating tension and cross-linking to other proteins (Alexandrova et al., 2008; Vicente-Manzanares et al., 2008; Parsons et al., 2010). In order for the moving cell to continue going forward, the adhesions formed with the extracellular matrix need to be disassembled. This occurs at the cell front and the rear. At the front, disassembly occurs most prominently at the lamellum-lamellipodium interface, owing to actin depolymerization and reorganization. Disassembly also occurs at regions where retraction is taking place, which is usually accompanied by a sliding of adhesions and then the dispersal of adhesion structures. This adhesion sliding seems to be a Rho GTPase- and myosin II-dependent form of treadmilling; the peripheral edge of the adhesion disassembles while the central edge assembles (Parsons et al., 2010). So, although the adhesion is seemingly moving, the components are actually being exchanged in and out.

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Interaction Actin and Motor Proteins Muscle Cell Contraction Actin is mainly known for giving muscle cells the ability to contract and give strength to different organisms. The current model accounting for skeletal muscle contraction is the sliding filament model (Krans, 2010). Under the microscope, skeletal muscle has a striped pattern. Each stripe is called a sarcomere and is formed by many parallel actin filaments and myosin filaments. The interaction of both filaments will give rise to sarcomere shortening, and thus muscle contraction. Myosin is able to pull upon actin, shortening the sarcomere. The globular end of each myosin (S1) has hinged segments that can bend and help in contraction. The slimmer tail region of myosin (S2) is also flexible, rotating in concert with the S1 contraction. When contraction takes place, myosin reaches forward, binding to actin, contracts, releases actin, and reaches forward to bind actin again. To bind to actin, crossbridges are formed between the filaments. This contraction movement of the S1 region of myosin is known as the power stroke, an energy-requiring process. Without ATP, myosin cannot release actin, and is thus stuck in a position. Two proteins regulate muscle contraction by blocking the binding of myosin to Factin: troponin and tropomyosin (Krans, 2010). But in the presence of Ca2+, these proteins can move aside and allow myosin to contact actin again. Thus, for muscle contraction, ATP and Ca2+ are the required cofactors.

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Vesicle Transport Actin filaments and myosin motors are needed for the transport of vesicles and the retention of organelles at specific sites. Secretory vesicles, endosomes and mitochondria are transported on actin filaments for short-range movement. The direction of the movement is established by the orientation of the plus and minus ends of the filaments (DePina and Langford, 1999). Actin filaments are required for the transport and retention of ER in the submicrovillar region and the localization of Golgi stacks and transport of vesicles. In secretory cells, actin filaments are involved in the release of exocytic vesicles (DePina and Langford, 1999). Thus, actin filaments do not just provide the tracks for myosin motors, they also serve as scaffold for organelles and organelle localization. However, how actin filaments link to the organelles is poorly understood. Cytokinesis After mitosis takes place, cells need to be divided to give rise to two daughter cells, i.e. cytokinesis. During cytokinesis, a contractile ring of F-actin filaments and myosin-II assemble at the equatorial cortex. Movement of myosin along actin filaments generates the force required for membrane constriction and the division into two daughter cells. Moreover, myosin regulates actin dynamics and recruits actin to the ring; it pulls on dynamic actin filaments, constricting the cleavage furrow cortex (Reichl et al., 2008). These actin networks are in general further organized into concentric antiparallel arrays, so that the myosin motors can pull the filaments and contract the membrane in a purse-string fashion (Reichl et al., 2008). Actin crosslinking proteins hold the filaments together so that when myosin-II pulls against the filament, tension on the filament can propagate into the whole crosslinked network. In the assembling of the contractile ring, both myosin heavy chain and phosphorylated myosin regulatory light chain accumulate at the equatorial cortex. Myosin II can localize to the ring prior to actin in diverse types of cells (Yumura, 2001) and needs actin to maintain it at the contractile ring site (Dean et al., 2005). After cytokinesis takes place, cells attach to the substratum via binding of integrins to the ECM substratum (e.g., fibronectin and laminin), getting into early G1 phase and spreading right after completion. This latter process is dependent upon actin metabolism (Boonstra and Moes, 2005). After attachment, spreading is achieved with membrane protrusions (lamellipodia and filopodia) and formation of actin stress fibers. Activation of integrins results in the activation of small GTPases of the Rho family of proteins. Rho family proteins activated by extracellular signal molecules (growth factors and extracellular matrix components) and downstream effects of the Rho proteins lead to changes in actin morphology (Boonstra and Moes, 2005).

Regulation of Transcription and Gene Expression Chromatin-Remodeling Different polymerization states of actin co-exist in the nucleus (Schoenenberger et al., 2005). Actin binds to three types of structures in the nucleus: ATP-dependent chromatin remodeling complexes, ribonucleoprotein particles, and the three eukaryotic RNA polymerases (Miralles and Visa, 2006). In response to extracellular signals, chromatin-

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remodeling complexes modify nucleosome structures (Miralles and Visa, 2006). Actin forms part of chromatin-remodeling and histone acetyl-transferase complexes. It has been identified as a component of the chromatin remodeling complex BAF (Brg or hBrm associated factors), a complex involved in T-lymphocyte activation (Olave et al., 2002). Actin binds directly to the BRG1 ATPase subunit of BAF, giving it its maximal ATPase activity and allowing BAF to bind to chromatin. The BAF complex might be binding to actin branch points to alter or maintain the chromatin structure (Bettinger et al., 2004). More actin-related proteins (ARPs) have been identified in the nucleus, with chromatin remodeling complexes of the SWI/SNF family. However, the molecular mechanisms by which actin and its related proteins help in chromatin remodeling complexes still have to be established (Dion et al., 2010).

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Regulation of Gene Expression Changes in actin dynamics can also regulate nuclear gene expression by modulating the subcellular localization of transcriptional regulators (Miralles and Visa, 2006). MAL, a transcriptional activator, binds directly to monomeric actin in the cytoplasm. When the Gactin-MAL complex dissociates, MAL accumulates in the nucleus. This shows that actin can act as a regulator of MAL accumulation in the nucleus (Miralles et al., 2003). Actin also affects the subcellular localization of other transcription factors, without actin polymerization PREP2 homeoprotein localizes to the nucleus while the transcriptional repressor YY1 is localized in the cytoplasm when actin polymerization is increased (Haller et al., 2004; Favot et al., 2005). Actin and Transcription Actin filaments are directly involved in transcription (Bettinger et al., 2004). Actin interacts physically with the three RNA polymerases. It is needed for a step in RNA-Pol I transcription that occurs after initiation (Miralles and Visa, 2006). With RNA-Pol II, it forms the preinitiation complexes of transcription. Disruption of the interaction between actin and the pre-mRNA-binding protein hrp65 blocks RNA-Pol II transcription at the elongation level. With RNA-Pol III it forms a complex and it is also needed for transcription (Miralles and Visa, 2006). Actin can function in conjunction with pre-mRNA-binding proteins as a platform for the localization of a HAT activity along active transcription units. Thus, actin plays an indispensable role in basal transcription, independently of chromatin. However, it is not known how the RNA polymerases use actin, but it has been proposed that all three share a common binding patch composed of RPABC2 and RPABC3 (Miralles and Visa, 2006). Actin in Pre-Mrnp Actin has been identified as one of the components of pre-mRNP (messenger ribonucleoprotein) complexes. Even though actin can work as an adaptor for mRNA export, it is not the major pathway. The cotranscriptional incorporation of actin into newly assembled pre-mRNPs affect chromatin structure at transcribed genes. Actin has been localized at nucleoplasmic filaments of the nuclear pore complexes and in a small nuclear-RNPassociated protein complex, through an interaction with nuclear DNA helicase-II (Bettinger et al., 2004).

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Concluding Remarks Actin can be found both in the cytoplasm and in the nucleus, playing different roles. Underlying the plasma membrane, it supports cell morphology by creating cell tension and is involved in the deformation of the plasma membrane, allowing the formation of protrusions, adhesions, exo- and endocytosis. In specialized cells such as neurons and cells of the immune system, actin is present at the synapses, playing a critical role in their establishment and during signaling. But actin is not a static structural protein, it is highly dynamic and is involved in cell movement (via formation of protrusions and assembly/disassembly of adhesions), intracellular vesicular transport, cytokinesis, and muscle cell contraction (with myosin motor proteins). All these processes are tightly regulated by actin-binding proteins, which coordinate the different processes and provide it with directionality.

Actin-Related Diseases Most diseases related to actin are caused by a malfunction in the formation of different actin structures. This can be either caused by a mutation in the actin structure itself or by the process being misregulated (errors in the actin-binding proteins). Actin has many different functions, and any little mistake in any of these roles could cause a disease. But the one function that is most involved in diseases is its structural role in cell membranes, leading to very severe and common diseases such as Alzheimer‟s and metastasis of cancer cells. The most prominent diseases are here reviewed and briefly linked to actin function.

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Muscular Diseases Actin is one of the most prominent proteins of muscle and any changes in actin structure may have a large impact on muscle function. As outlined above, muscle contraction depends on the interaction between actin and myosin on one hand and its interaction with the membrane on the other. Muscular dystrophy refers to hereditary muscle diseases characterized by progressive muscle weakness, muscle protein defects and ultimately dead of muscle cells and encompasses 40-50 disorders, amongst them Duchenne, Becker and EmoryDreyfuss dystrophy (for review see: Ozawa, 2010). Duchenne muscular dystrophy has been demonstrated to be due to mutations of dystrophin. Dystrophinlocalizes to costameres, large subsarcolemmal protein assemblies. At these sites dystrophin interacts with the cortical actin cytoskeleton, composed of γ-actin and the transmembrane dystroglycan, which on its turn binds to the extracellular protein laminin (Rybakova et al. 2000). Through interactions between γ-actin, dystroglycan and other proteins, dystrophin forms the dystrophin glycoprotein complex which forms a mechanical strong link between the cortical actin cytoskeleton and the extracellular matrix (Ervasti, 2007; Prochniewicz et al., 2009).Modulation of the composition of these complexes, for example through loss of dystrophin expression, causes a destabilization of the complexes and consequently in sarcolemmal damage and muscle cell necrosis (Ervasti, 2007). In addition to a loss of dystrophin expression that lead to muscular dystrophy, also many mutations have been

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described in the actin binding domain of dystrophin that lead to muscular dystrophy (Roberts et al, 1994). Of particular interest are the observations that, in addition to a structural role as described above, the dystrophin-glycoprotein complex may be involved in Ca2+ homeostatis and in signal transduction through the MAP kinase pathways and G-proteins (for review see: Batchelor and Winder, 2006). Furthermore, proteomic profiling of dystrophin-deficient heart demonstrated drastic changes in the levels of many metabolic and contractile proteins, indicating that the loss of dystrophin causes in addition to the contractile machinery also abnormalities in cell metabolism, cell stress response and the cytoskeleton (Lewis et al. 2010). The genes of actin isoforms in muscle cells are ACTA1 and ACTC, coding for the skeletal and cardiac muscle isoforms respectively (D‟Amico et al., 2006). During development, these two genes are co-expressed in the different muscle tissues, and continue to be so during adulthood with ACTA1 being most predominant in skeletal and ACTC in cardiac muscle. Over 100 ACTA1 mutations have been identified in patients with congenital skeletal muscle myopathies (D‟Amico et al., 2006). These myopathies are disorders characterized by skeletal muscle weakness ranging in their severity from neonatal lifethreatening disorders to mild muscle weakness in adulthood (Clarkson et al., 2004). Mutations in ACTA1 primarily cause three types of congenital myopathies; actin myopathy, intranuclear rod myopathy and nemaline myopathy (Schöder et al., 2004). Nemaline myopathy stands out for having the highest incidence of the three and having cases with filamentous nemaline bodies or rods which are composed of actin; the mutations in the actin gene affect the stability or conformation of actin, interfering with contacts between actin and other molecules (Ilkovsky, 2008). Actin myopathy is typified by homogenous filamentous insertions containing actin that occupy sarcomeres whereas intranuclear rod myopathy is characterized by the presence of giant rod bodies in the muscle cell nuclei. In addition, ACTA1 mutations can also affect the size and distribution of muscle fiber types. Depending of the mutations, different congenital myopathies occur. Neither the form of inheritance nor the severity of the disease can be predicted from the mutated gene, as each gene can give rise to dominant or recessive mutations and severe, typical or mild forms (Clarkson et al., 2004). Mutations in ACTC are more rare, being present in only some families, resulting in dilated or hypertrophic cardiomyopathy, which can also be caused by ACTA1 gene mutations (D‟Amico et al., 2006). Interestingly, the balance of actin assembly and disassembly has been demonstrated to control the transcriptional programme regulated by serum-response factor during muscle development and maintenance (Kuwahara et al. 2005). Thus actin-based myopathies may also result from altered transcriptional activities due to the role of serumresponse factor (Visegrady and Machesky, 2009).

Neurological Diseases Many neurological degenerative diseases, including Alzheimer‟s disease, Parkinson‟s disease, Huntington‟s disease, fronto-temporal dementia, are characterized by the formation of protein deposits in specific regions of the brain (Lee et al., 2011). Alzheimer‟s disease, one of the most prominent diseases, is famous for the formation of extracellular amyloid plaques in the neural tissue, leading to neurologic degeneration, and thus cognitive impairment. In addition to the extracellular plaques also intracellular neurofibrillary tangles (NFTs) are

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formed (Bamburg and Bloom, 2009). For a long time it has been thought that the amyloid plaques were the main cause of Alzheimer‟s and it has been the target of new therapies. However, new discoveries have demonstrated that cognitive impairment is evident before the development of plaque formation (Penzes and VanLeeuwen, 2011). It seems that synapse degeneration occurs prior to the deposition of the plaques. Thus, it has been argued whether synaptic degeneration is the driving force rather than the byproduct of Alzheimer‟s disease, leading to memory impairment (Penzes and VanLeeuwen, 2011). As described above, synaptic function is largely dependent upon synapse morphology, and on its turn synapse morphology is determined by the components of the cytoskeleton, including the microtubule system and actin microfilaments, both of these systems were demonstrated to be modulated in Alzheimer patients. The formation of intracellular NFTs has been described already by the discoverer of the disease (Alzheimer, 1907). These NTFs appeared densely packed fibers (20 nm in diameter) of variable length, one of the main components appeared to be the tau protein in addition to other neurofilamental proteins (for review see Bamburg and Bloom, 2009). Tau is a microtubule binding and stabilizing protein and accumulates preferentially in axons in brain tissue. In brain tissue of Alzheimer patients, tau appears to be heavily modulated, including hyperphosphorylation and C-terminal proteolysis, leading to a lower affinity for microtubules and consequently a higher tau polymerization (for review see Bamburg and Bloom, 2009). Interestingly, Alzheimer‟s is not the only disease that exhibited a modulated tau metabolism, but it appeared also in for example frontotemporal dementia with Parkinsonism, corticobasal degeneration and Pick‟s disease (for review see Bamburg and Bloom, 2009). In addition to the microtubules, also actin plays an important role in establishment of proper synapse morphology and function as described above.Among the most important regulators of actin dynamics in brain tissue are members of the ADF/cofilin family. As described above, cofilin is an actin severing protein and its activity results in the formation of many filament ends. The newly formed barbed ends may enhance nucleation and filament formation or the pointed ends may result in depolymerization, depending upon the external conditions. The ability of cofilin to enhance actin polymerization depends upon its phosphorylation by LIM kinase and other kinases and its dephosphorylation or inactivation is induced by highly regulated phosphatases (for review see Bamburg and Bloom, 2009). ADF/cofilin and actin in rod-like structures have been identified in frontal brain areas and the hippocampus in Alzheimer‟s patients (Bamburg and Wiggan, 2002). All dense-core amyloid plaques have ADF/cofilin-actin aggregates or rods in clusters around them, whereas about 45% of the aggregates and rods present are without amyloid plaques. These rods appeared to be required for pMAP/tau recruitment, indicating the connection between the actin microfilaments and the NTFs (Whiteman et al, 2009). Similar cofilin-actin rods are formed in many cell types in response to heat shock, osmotic stress and ATP depletion, in most cases these rods are reversible and do not cause irreversible damage to the cells (for review see Bamburg and Bloom 2009). In neurites, the rods have however been demonstrated to block transport (Maloney et al, 2005). In addition to its effects on rod formation, cofilin has also direct effects on the structure and dynamics of the postsynaptic terminals, i.e. the dendritic spines. Spine dynamics are driven by actin treadmilling (Honkura et al., 2008) mediated by cofilin and drebrin, an F-actin stabilizing protein (Kojima and Shirao, 2007). In Alzheimer‟s brains, upstream modulators of cofilin phosphorylation are down-regulated, resulting in enhanced cofilin activity, while an

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excessive release occurs of drebrin, stabilizing F-actin (Zhao et al, 2006). These effects may result in enhanced rod formation which may block transport to spines on one hand and sequestering of active cofilin on the other (for review see Bamburg and Bloom 2009). So, on the one hand cofilin induces actin destabilization, and there is increasing evidence that supports a role for cofilin in neurodegeneration and Alzheimer‟s disease (Maloney and Bamburg, 2007; Shankar et al., 2007; Penzes and VanLeeuwen, 2011). On the other hand, drebrin binds and stabilizes actin in dendritic spines and has been reported to be present in reduced levels in the hippocampal formations of patients with Alzheimer‟s and in other cortical areas (Harigaya et al., 1996). The pathways that underlie rod formation have been partly solved. ATP-depletion causes the release of the phosphatase chronophin from Hsp90 (Huang et al, 2008), resulting in chronophin activation and cofilin dephosphorylation. Silencing of chronophin in neurons with siRNA causes a decrease but not elimination of rod formation in response to ATP-depletion, suggesting that other phosphatases may be involved in rod formation as well (Huang et al. 2008). Interestingly, knock-out of cdc42 or inhibition of cdc42 activity caused a strong reduction of neuron forming rods, indicating also the involvement of cdc42 in actin rod formation (for review see Bamburg and Bloom, 2009). Another disruption in Alzheimer‟s patients is found in PAK, a critical regulator of actin assembly and dendritic spine modulation. In the hippocampus of these patients, total PAK is reduced, especially active PAK (Zhao et al., 2006). The pathology of Alzheimer‟s also mislocalizes PAK in neurons, which is followed by a loss in F-actin in dendrites and dendritic spines. Moreover, RhoGTPase has also been found to have a reduced expression and an altered localization in the hippocampus of Alzheimer‟s patients (Huesa et al., 2010). Furthermore, addition of a PAK inhibitor to cultured neurons induced the formation of cofilin rods (Zhao et al., 2006). All of these misregulations can end up in the formation of actin aggregates and thus rods. Of particular interest are the observations that gelsolin, an actin capping and severing protein, has been demonstrated to regulate the fibrillization of amyloidβ protein (Ji et al., 2010). The second most common neurodegenerative disease is Parkinson‟s disease, characterized by loss of dopaminergic neurons in the substantia nigra coupled with the formation of intracellular Lewy bodies (Olanow and Tatton, 1999). Recently genome-wide analysis have indicated a number of genes that are linked to Parkinson‟s disease, a major role being implicated for α-synuclein, LRRK2, parkin, PINK1 and DJ-1 (for review see Martin et al. 2011). α-Synuclein is a small protein enriched at the pre-synaptic terminals of almost all neurons. Recent evidence indicated that α-synuclein has a role in synaptic vesicle transport and the regulation of exocytosis (Bellani et al., 2010). In the brain of Parkinson‟s disease patients, α-synuclein is usually detected in Lewy bodies of neurons in the brain stem and cortex, aggregates of α-synuclein being the main component of the Lewy bodies (Lee and Trojanowski, 2006). It has been shown that α-synuclein interacts also with actin (Zhou et al.,2004) and is able to alter actin dynamics (Sousa et al., 2009). α-Synuclein is able to sequester actin monomers and consequently causes a reduction in the pool of polymerized actin and by that α-synuclein might influence exocytosis (Bellani et al., 2010). Another important protein related to Parkinson‟s disease is PINK1. Mutations in PINK1 have been demonstrated to cause early onset familial Parkinsons‟s disease and PINK1 gene knockdown was shown to increase the binding of parkin to actin filaments and an increased phosphorylation of cofilin (Kim and Son, 2010), again indicating the potential role of the

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actin microfilaments in Parkinsons‟s disease. Furthermore, Parkin, a ubiquitin E3 ligase implicated in Parkinson‟s disease, has been shown to bind to tubulin and to stabilize microtubules (Yang et al.,2005) but also was implicated in actin remodeling through cofilin. LRRK2, another protein involved in Parkinson‟s disease, is also connected to actin through the phosphorylation of ERM proteins (Parisiadou et al., 2009). Phosphorylated ERM proteins link F-actin to the plasma membrane by binding the N-terminal to the plasma membrane while the C-terminal binds directly to F-actin. Altogether, these observations strongly suggest an important role of actin in the development of Parkinson‟s disease.

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Immunological Diseases As described above, the actin-related processes are essential for virtually all cell types, but a strong relation has been oberserved between impaired regulation of the actin cortical filaments and immunodeficiency and autoimmune diseases (Zhang et al. 2009; Wickramarachchi et al. 2010), because essential actions of the immune system rely amongst others on migration, phagocytosis, secretion and cell-cell interaction. Proper actin dynamics are mandatory for correct activation of the lymphocytes. In the immunological synapse, actin dynamics are normally tightly regulated. This type of synapses is defined as the structure formed by the interaction between lymphocytes and antigen-presenting cells (APCs) (Gordón-Alonso et al., 2010). If actin dynamics are abnormal, immune diseases result. Neutrophil actin dysfunction is an autosomal recessive disease in which there is a severe neutrophil disorder as a result of a defect in neutrophil actin assembly, due to a defect in an actin-associated protein, not in the actin molecule itself (Nunoi et al., 2001). In leufactin disease, neutrophils show abnormalities in motility as a consequence of partial defects in actin polymerization and scattering (Nunoi, 2008). In these patients, there is a decrease in an 89 kd protein and an increase in a 47 kd protein. This 47 kd protein (now called leufactin) is known to interact with F-actin via the C-terminal domain, being important for the adjustment of actin polymerization, and thus is the cause of this disease. And in a more common X-linked recessive disorder, Wiskott-Aldrich Syndrome (WAS), actin nucleation is regulated abnormally, leading to severe immunodeficiency primarily in chemotaxis of macrophages (Nunoi, 2008). Mutations in the WAS gene lead to distinct syndrome variations depending on the mutation; premature termination and deletion abrogate Wiskott-Aldrich syndrome protein (WASp) expression, leading to severe WAS, whereas missense mutations lead to reduced protein expression and usually to attenuated WAS. WASp is key in transducing signals from the surface of the cell to actin, acting as an adaptor to bring together downstream mediators that facilitate Arp2/3 mediated actin polymerization and also being crucial for the subcellular organization of signaling complexes to actin (Blundell et al., 2010). As such, a lack of WASp can result in actin defects, compromising several aspects of normal cellular activity (e.g., phagocytosis, proliferation and immune synapse formation).

Vascular Diseases Vascular disease is a pathological state of large and medium sized muscular arteries and is triggered by endothelial cell dysfunction. The microvascular endothelium consists of a

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layer of closely apposed endothelial cells. Under normal conditions the equilibrium between endothelial cell-cell adhesions and actin-myosin centripetal tension controls the permeability properties of the microvascular barriers (Shen et al. 2010; Kliche et al.2011). Endothelial cells form a monolayer lining the luminal surface of the entire vascular system. One of their main functions is to control vascular permeability by providing a semi-permeable barrier between the blood and the underlying tissues. This barrier function is regulated to a great extent by endothelial cell-cell junctions, namely adherens and tight junctions. The maintenance of endothelial cell-cell junction integrity is a crucial process for the regulation of vascular homeostasis. As has been described above, actin plays a central role in the proper composition and functioning of cell-cell junctions and changes in actin properties or metabolism may have severe effects on endothelial permeability. In addition, actin plays a central role in contraction of smooth muscle cells underlying the endothelium, again indicating the essential role of actin in microvascular endothelial functioning. A wide variety of pathologies are caused by misregulated endothelial permeability; such as edema, increased interstitial pressure, inflammation, vascular damage, formation of microthrombi, and even hemorrhages (van Nieuw Amerongen and van Hinsbergh, 2002; Dejana et al., 2009). It has become clear during recent years that many of these pathologies have a common theme, that is the rapid and dynamic reorganization of actin microfilaments controlled by Rho-kinase signaling (Nunes et al. 2010; Zhou et al., 2011). Rho-kinase is the downstream effector of RhoA. Activation of RhoA increases actomyosin contractility and causes junction disruption whereas activation of Rac1 and of Cdc42 promotes endothelial barrier stabilization (van Nieuw Amerongen et al., 2007; Nunes et al. 2010; Shen et al. 2010). During ischaemic episodes, blood supply is reduced, leading to a shortage of ATP in the affected cells. In endothelial cells especially, there is a reversible and temporary disruption of the endothelial cell microfilament architecture that can lead to endothelial cell dysfunction (Hinshaw et al., 1988). As previously described, ATP is very important in maintaining the Gactin structure and in the F-actin filament dynamics. The effects of ischaemia on actin cytoskeleton have been extensively studied in renal vascular cells and have revealed how the actin cytoskeleton is involved in many cellular processes (Kwon et al., 2002). After an ischaemic process, disorganization (disruption and/or clumping of the filamentous structures) and an abnormal distribution of F-actin is observed (Kwon et al., 2002).

Cancer Cancer cells are characterized by the fact thatthey exhibit uncontrolled proliferation. In addition cancer cells are sometimes able to metastasize, that is to move to other parts in the organism. This latter property causes usually the biggest problems for the patients. Increased metastasis is due to the property that cancer cells become less dependent upon attachment (to other cells or the extracellular matrix) dependent proliferation. As described above, cell-cell attachment and cell-extracellular matrix attachment is heavily determined by actin through the establishment of cell junctions. In cancerous cells, by changing their actin structure or regulation, adhesions can be modified, detaching these cells from their environment, allowing them to move to other locations. Not only that, but actin structures can help these cells in their migration, by polymerizing under the plasma membrane, forming membrane protrusions and ultimately lamellipodia (Yamaguchi and Condeelis, 2006). Protrusions are formed in response

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to migratory and chemotactic stimuli. Molecules linking these migratory signals to the actin cytoskeleton (eg, cofilin) have been found to be upregulated in invasive and metastatic cancer cells (Yamaguchi and Condeelis, 2006). Protein products of classic oncogenes perturb the actin cytoskeleton. For instance, oncogenic Src causes loss of organized actin and adhesion structures in cells (cell to cell and cell to matrix adhesions). The presence of a kinase-defective v-Src protein (which inactivates RhoA) in fibroblasts causes stabilization of actin stress fibers and their associated focal adhesions, resulting in impaired migration (Fincham et al., 1995; Frame and Brunton, 2002). Research has focused on how the Rho family of small GTPases can be linked to oncogenesis and cancer development (Frame and Brunton, 2002). Members of this family control the actin cytoskeleton and cell shape via several characteristic effector proteins. Rac1 and Cdc42 work together at leading edges of migrating cells to coordinate the formation of lamellipodial and filopodial extensions, linking to the actin polymerization machinery (Frame and Brunton, 2002). There is quite important evidence that lamellipodia formation is an important process early in chemotactic attraction of tumor cells, and that Rho proteins, and downstream Arp2/3-dependent actin polymerization at the leading edge, are involved (Condeelis et al., 2001; Frame and Brunton, 2002).

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Conclusion All diseases described above depend also on the function of actin which in turn depends on its structure and regulation. Actin can be found in two forms: G-actin or F-actin. G-actin are the monomers which can be bound to ATP or ADP and to actin-binding proteins. F-actin are the filaments that are formed when G-actin is bound to ATP and polymerizes. In addition, actin filaments can combine with each other and form different structures such as networks and bundles. All these processes are regulated by actin-binding proteins, which coordinate it and provide it with directionality. Depending on the structure formed and the actin-binding proteins, actin can perform different functions. In the plasma membrane, it gives the cell its shape and creates cell tension, allowing different processes to occur (such as formation of protrusions). In neurons and cells of the immune system, actin is very important at the synapses. In cell movement, the dynamic character of actin is very useful, allowing cell movement to occur, intracellular vesicular transport or even cytokinesis. When any of these processes is misregulated or actin itself is changed, diseases can occur affecting any of its functions. From this, muscular, vascular and immune diseases, neurodegenerative disorders, and cancer can occur because of actin malfunctioning. Knowing more about the dynamics of actin and its role especially in diseases, could help in the discovering of new treatments.

Acknowledgements We would like to thank Naïma Cleuren for helping in the preparation of the figures. Due to space constraints and the huge body of literature, we were not able to cite all original and review papers dealing with actin and its metabolism. Therefore we would like to apologize to authors whose work in this area may have been omitted.

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Tang, D.D. and Anfinogenova, Y. (2008). Physiologic properties and regulation of the actin cytoskeleton in vascular smooth muscle.J. Cardiovasc. Pharmacol.therapeut, 13, 130140. Theriot, J.A. and Mitchison, T.J. (1991). Actin microfilament dynamics in locomoting cells.Nature, 352, 126–131. Tomasevic, N., Jia, Z., Russell, A., Fujii, T., Hartman, J.J., Clancy, S., Wang, M., Beraud, C., Wood, K.W. and Sakowicz, R. (2007). Differential regulation of WASP and N-WASP by Cdc42, Rac1, Nck, and PI(4,5)P2. Biochem.,46, 3494-3502. Uribe, R. and Jay, D. (2009). A review of actin binding proteins: new perspectives. Mol. Biol. Rep., 36, 121-125. van Nieuw Amerongen, G.P., and van Hinsbergh, V.W. (2002). Targets for pharmacological intervention of endothelial hyperpermeability and barrier function.Vascul. Pharmacol.39, 257-272. van Nieuw Amerongen, G.P., Beckers, C.M., Achekar, I.D., Zeeman, S., Musters, R.J., and van Hinsbergh, V.W. (2007). Involvement of Rho kinase in endothelial barrier maintenance.Arterioscler Thromb Vasc. Biol.27, 2332-2339. Vicente-Manzanares, M., Koach, M. A., Whitmore, L., Lamers, M. L. and Horwitz, A. F. (2008). Segregation and activation of myosin IIB creates a rear in migrating cells. J. Cell Biol., 183, 543–554. Visegrady, B. and Machesky, L.M. (2009). Myopathy-causing actin mutations promote defects in serum response factor signaling. Biochem. J., 427, 41-48. Wasser, M. and Chia, W. (2000). The EAST protein of Drosophila controls an expandable nuclear endoskeleton. Nature Cell Biol.,2, 268–275. Whiteman, I.T., Gervasio, O.L., Cullen, K.M., Guillemin, G.J., Jeong, E.V., Witting, P.K., Antao, S.T., Minamide, L.S., Bamburg, J.R. and Goldsbury, C. (2009). Activated ADF/cofilin sequesters phosphorylated microtubule-associated-protein during the assembly of Alzheimer-like neuritic cytoskeletal striations. J.Neurosci., 29, doi: 10.1523/JNEUROSCI.3531-09.2009 Wickramarachchi, D.C., Theofilopoulos, A.N. and Kono, D.H. (2010). Immune pathology associated with altered actin cytoskeleton regulation. Autoimmun, 43, 64-75. Wiegant, F.A.C., Blok, F. J., Defize, L.H.K., Linnemans, W.A.M., Verkleij, A.J. and Boonstra, J. (1986). Epidermal growth factor receptors associated to cytoskeletal elements of epidermoid carcinoma (A431) cells. J. Cell Biol., 103, 87-94. Winslow, A. and Rubinsztein, D.C. (2011). The Parkinson disease protein α-synuclein inhibits autophagy. Autophagy, 7, 429-431. Yamaguchi, H. and Condeelis, J. (2006). Regulation of the actin cytoskeleton in cancer cell migration and invasion.Biochim. Biophys.Acta, 1773, 642-652. Yang, F., Jiang, Q., Zhao, J., Ren, Y., Sutton, M.D. and Feng, J. (2005). Parkin stabilizes microtubules through strong binding mediated by three independent domains. J. Biol.Chem., 280, 17154-17162 Yin, H.L. and Janmey, P.A. (2003).Phosphoinositide regulation of the actin cytoskeleton.Ann. Rev.Physiol., 65, 761–789. Yoshihara, Y., De Roo, M. and Muller, D.(2009). Dendritic spine formation and stabilization.Curr. Opin.Neurobiol., 19, 146–153. Yumura, S. (2001). Myosin II dynamics and cortical flow Turing contractile ring formation in Dictyostelium cells. J. Cell Biol., 154, 137-146.

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Zhang, C., Jenkins, H., Goldberg, M. W., Allen, T. D. and Hutchison, C. J. (1996). Nuclear lamina and nuclear matrix organization in sperm pronuclei assembled in Xenopus egg extract. J. Cell Sci., 109, 2275–2286. Zhang, J., Dong, B. and Siminovitch, K.A. (2009). Contributions of Wiskott-Aldrich syndrome family cytoskeletal regulatory adapters to immune regulation.Immunolog.Rev., 232, 175-194. Zhao, Z.S. and Manser, E. (2005). PAK and other Rho-associated kinases--effectors with surprisingly diverse mechanisms of regulation.Biochem. J., 386, 201-214. Zhou, Y., Gu, G., Goodlett, D.R., Zhang, T., Pan, C., Montine, T.J., Montine, K.S., Aebersold, R.H. and Zhang, J. (2004). Analysis of α-synuclein-associated proteins by quantitative proteomics.J. Biol.Chem., 279, 39155-39164. Zhou, Q., Gensch, C. and Liao, J.K. (2011). Rho-associated coiled-coil-forming kinases (ROCKs): potential targets for the treatment of atherosclerosis and vascular disease. Trends Pharmacol.Sci., 32, 1-7. Zigmond, S.H. (1996). Signal transduction and actin filament organization.Curr. Opin. Cell Biol, 8, 66–73.

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ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter III

Osmotic Pressure: A Tool to Investigate the Polymeric Forms of Actin Enrico Grazi Dept. of Biochemistry and Molecular Biology, 44100 Ferrara, Italy.

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A very impressive property of actin is the versatility in the formation of supramolecular structures. G-actin is converted into filamentous or globular structures, it is ordered into paracrystalline arrays or in tubular structures. The formation of these almost regular structures is accompanied by distinct changes of the osmotic properties of the actin solution so that, in some cases, the geometric parameters of the supramolecular structure can be related, albeit indirectly, to protein osmotic pressure. With the increase of protein osmotic pressure protein-bound water is removed and the volume of the system decreases. These events are necessarily linked to changes of the geometric parameters of the supramolecular structure. The study of protein osmotic pressure may thus provide information on the geometry of the supramolecular assembly, even though the geometry depends in a complex manner on the concentration and charge of the protein. In addition, change of protein osmotic pressure may signal the transition of the actin filaments into a network of filaments. The study of osmotic protein pressure allows to determine the activity and the change of the free energy of the solute in binary solutions and, in some cases, in ternary solutions. Two further points deserve consideration: A. The free energy of the free actin monomers is related to the protein osmotic pressure generated by polymeric actin solutions. B. With the model of Biron et al. [Europhys. Lett 2006, 73, 464] it is possible to relate the change of the free energy of the free actin monomers to the change of the length distribution of the actin filaments. It follows that the length



Phone: 0039 0532 455421; Fax: 0039 0532 454442; e-mail: [email protected]

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Enrico Grazi distribution of the actin filaments is regulated by (1) the free energy of hydrolysis of ATP and (2) the protein osmotic pressure.

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1. Introduction The actin filament provides the physical basis for the structure and dynamic properties of the cell cytoskeleton and the muscle sarcomere. Metal ions, the state of hydrolysis of the bound nucleotide, actin-binding proteins strongly influence the properties of monomeric (G) as well as filamentous (F) actin in vitro. In the presence of either monovalent cations at high concentration or mM Ca2+ or mM Mg2+ G-actin polymerizes into double stranded F-actin (Kasai et al., 1962). The elongation of the actin filament takes place, simultaneously, through two processes: 1. additions of monomers to a presumably trimeric nucleus (Frieden and Goddette 1983) 2. condensation of small oligomers (Grazi, 1989). With Ni2+ as the polymerizing ion, the formation of short over long actin filaments prevails while, with Zn2+, globular aggregates are formed (Strzelzcka-Golassewka et al., 1978). Paracrystalline bundles of filaments are formed at Mg2+ concentrations above 5 mM or in the presence of polyamines (Oriol-Audit, 1978). Protamine induces the formation of various supramolecular structures from G-actin, depending on the molar ratio of the two proteins and on the ionic strength of the medium (Grazi and al., 1982). Lanthanides induce the formation of microcrystalline aggregates from G-actin that, by further addition of 0.1 M KCl, are converted into tubular structures (Dos Remedios and Dickens, 1978). These tubular structures, at difference with F-actin, contain ATP and not ADP (Barden and Dos Remedios, 1980). Furthermore, the substitution of Ca2+ with Tb3+ at the high affinity site is incompatible with the typical F-actin structure (Ferri and Grazi, 1981). The formation of actin bundles in vivo is regulated by many actin binding proteins such as tropomyosin, caldesmon and filamin. To reproduce in vitro the effects displayed by these proteins in vivo it is necessary to mimic the condition of the cell sap by the addition of macromolecules such as polyethylene glycol 6000. It is then realized that each bundling protein requires a different concentration of the macromolecule. With caldesmon-decorated actin bundles are formed in 3% poly(ethyleneglycol), with filamin-decorated actin at 4-5% poly(ethyleneglycol), with caldesmon-tropomyosin-decorated actin at 5-7% poly(ethyleneglycol), with F-actin alone at 6-7% poly(ethyleneglycol) and with tropomyosindecorated actin at 9-10% poly(ethyleneglycol) (Grazi and al., 1989; Cuneo and al., 1992). Furthermore the interplay between the concentration of the macromolecule, state of aggregation of actin and distribution of the ancillary cytoskeleton proteins is quite complex. Caldesmon binds tighter to tropomyosin-decorated F-actin than to F-actin (Moody and al., 1985), as a consequence a system composed by tropomyosin-decorated actin and by caldesmon-decorated actin evolves toward a system composed by F-actin and by tropomyosin-caldesmon-decorated actin. This has a profound effect on the proportion of actin filaments over actin bundles (Cuneo and al., 1992). Based on in vitro experiments, many actin bundling proteins were proposed to regulate the rapid formation and dissociation of actin bundles (Sobue and al., 1981; Sobue and al., 1982; Bahler and Greengard, 1987; Husai and al., 1988; Ikebuchi and Waisman, 1990). To our knowledge the dissociation of actin bundles was not checked under macromolecular

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crowding conditions similar to those of the cytosol. When the proposed dissociating agent is a protein the main drawback is the lack of penetration into the actin bundle, due to steric hindrance. Small molecules such a Mg2+ may overcome the drawback more easely (Grazi and al., 1992; Grazi, 1994). In the cell the gelsolin to actin molar ratio is reported to be about 1:100 (Pollard and Cooper 1986). In vitro this ratio is adequate to support nucleation, capping and cutting actin filaments. In the cell, however, due to high macromolecular crowding, that favours the formation of bundles of actin filaments (Suzuki and al. 1989), gelsolin is probably inadequate to the cutting function. This is because the strong latero-lateral association of the actin filaments in the bundles probably neutralizes the cutting by gelsolin (Grazi and al. 1991). Arps 2 and Arps 3, after activation through several actin binding proteins and ATP, form a tight complex with actin and, apparently, induce the branching of the actin filaments (Kiselar and al., 2007). However, in vivo, this branching role is not generally accepted (Koestler and al., 2008). A further branching mechanism may occur through the incorporation of the “Lower Dimer” of actin into the growing F-actin filaments (Steinmetz and al. 1997). The integration of the concept of macromolecular crowding with that of macromolecular osmotic pressure, i.e. the pressure contributed by the macromolecules in solution, offers a more rigorous approach to the study of polymeric actin in vivo. Pioneering work in this respect was performed by Ogston and Phelps (1961) and by Laurent and Ogston (1963) who first observed that 1. hyaluronic acid influences the partition of serum albumin between solutions of the polysaccharide and buffer 2. solutions containing both hyaluronic acid and serum albumin have osmotic pressure larger of the sum of osmotic pressure of solutions containing hyaluronic acid and serum albumin separately at the same concentration. The phenomenon was attributed to the “exclusion” of albumin from part of the solution occupied by hyaluronic acid. The thermodynamic properties of these ternary systems were discussed by Ogston (1962), Kuntz and Kautzmann (1974), Arakawa and Timasheff (1985a, 1985b). It was concluded that phenomena similar to those described by Ogston and Phelps (1961) were of general occurrence in the cell sap where the association and dissociation of macromolecules is influenced by the presence of other macromolecules. In this paper the study of macromolecular osmotic pressure is used as a tool to define the properties of actin in water solutions. Under “THEORY” ideal and non-ideal solutions are defined and their energetic is treated. A special ternary solution is studied where two macromolecular solutes undergo association, the reaction being at the chemical equilibrium. The relation between the chemical potential of water and the chemical potential of the free actin monomer in F-actin solutions is investigated. The distribution and energy of the actin filaments in the F-actin solution are described. The geometric constraints imposed by the association of the actin filaments into hexagonally packed bundles of filaments are exploited to estimate the inter-filament distance. The stiffness and the yield strength of the actin filament are related to protein osmotic pressure. These theoretical considerations are exploited in the “APPLICATIONS TO THEORY” where the „fluttering wing‟ model of the actin filament is proposed and the osmotic properties of the Ca2+-regulated actin filament are defined. The energetic of equimolar solutions (as monomers) of actin and myosin is assessed and the chemical potential of the 1:1 actomyosin complex is determined. Te free energy of the free actin monomer and the length distribution of the actin filaments, as a function of protein osmotic pressure, are investigated. Methods to estimate the yield strength and the stiffness of thin filament in vivo are proposed.

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2. Methods 2.1. Measurement of the Macromolecular Osmotic Pressure Macromolecular osmotic pressure is the fraction of total osmotic pressure generated by the macromolecular components of the system. Macromolecular osmotic pressure is measured by secondary osmometry, i.e. by equilibrating through a dialysis membrane a small volume (1 mL) of the protein solution of unknown macromolecular osmotic pressure against a large volume of the reference solution of known macromolecular osmotic pressure. Since small solutes diffuse freely through a dialysis membrane, osmotic stress (the macromolecular osmotic pressure) is due only to the macromolecular components. At equilibrium the macromolecular osmotic pressure of the unknown and of the test solution are the same. Furthermore, since the volume of the unknown solution is much smaller than that of the test solution, the macromolecular osmotic pressure of this latter is essentially equal to its own pressure at the beginning of the experiment. Thus the macromolecular osmotic pressure of the equilibrated protein solution is known. Equilibration of protein solutions is carried out for 4896 h, at 22°C, in stopped bottles, immersed in a shaker water-bath thermostatically controlled to within ±0.1°C. At the end of the equilibration, a bag at the time is processed. It is withdrawn from the bottle, wiped off gently, and opened with a pair of scissors. The content is rapidly transferred, by means of a spatula, onto preweighed 2 x 2 cm cover glasses previously stored in a desiccator over P205 (Schwienbacher and al. 1995).

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2.2. Determination of the Wet Weight of the Sample From the weight of the sample, determined at time intervals, in a 15 min period, the weight at 3 h is substracted (at this time evaporation is essentially complete). The plot, against time of the logarithm of these differences is linear and is extrapolated to zero time (the time of opening of the bag) to yield the wet weight of the sample.

2.3. Determination of the Dry Weight of the Sample The sample is stored for 17 h at 80 °C and weighed and the weight of the cover glass is substracted.

2.4. Determination of the Water Weight of the Sample The dry weight is substracted from the wet weight of the sample.

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2.5. Determination of the Protein Weight a) The weight of salt is substracted from the dry weight. The weight of salt is calculated from the salt content of the equilibrating solution and from the water weight. b) The cover glass is immersed for 4 h in a 1 M NaOH solution (10 mL). The absorbance at 290 nm of the resulting protein solution is then determined against a blank of 1 M NaOH. The original protein solution is utilized as a standard. Measurements with the two methods are usually within 5% and are averaged.

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2.6. Special Care for Long Lasting Dialysis at Low Macromolecular Osmotic Pressures 2.6.1. The Dialysis of F-Actin F-actin solutions, 120–168 µM as the monomer, are dialyzed for 24 h at 2°C (first dialysis) against buffer A (1000 g of water; KCl, 0.1 mol; triethanolamine, 0.01 mol; MgC12, NaN3, 2-mercaptoethanol, 2 mmol each; ATP and CaCl2, 0.1 mmol each. pH is taken to 7.45 with 6N HC1). The dialyzed actin solution is diluted with the same buffer to 12–48 µM. The resulting actin solution (10–12 mL) is dialyzed overnight (second dialysis) at 21°C against 2 L of buffer A. The dialysis tube is open at the upper extremity and care is taken to maintain the inner and the outer solutions at the same level. At the end of the dialysis, the concentration and the specific viscosity of F-actin are measured. The experiment is performed on 1 mL aliquots of the actin solution, transferred by means of 1 mL Pedersen micropipette into dialysis bags. These are then dialyzed (third dialysis), with stirring, against 100mL aliquots of buffer A taken from the same solution used for the second dialysis. Dialysis is performed at 21°C in stopped bottles (beside a small hole in the stop cock), immersed in a water-bath thermostatically controlled to within ±0.1 °C. Also in this case the dialysis bag is open at the upper extremity and care is taken to maintain the inner and the outer solutions at the same level. Dialysis is prolonged up to 12 days. F-actin is stable even for 12 days, as judged from its specific viscosity, provided that the solutions are supplemented with 2 mM 2-mercaptoethanol. At the end of the experiment the volume of the protein solutions is measured by means of Pedersen micropipettes and the viscosity is determined. Usually the 1mL protein solution is fully recovered. 2.6.2. The Dialysis of F-Actin Against Poly(Ethylene Glycol) 40,000 Solutions F-actin solutions are prepared as described in 2.6.1. with the difference that at the time of the third dialysis to the 100mL aliquots of buffer A, 0.03 to 1.6 g of poly(ethylene glycol) 40,000 are added. Equilibration is then performed up to 9 days at 21°C. At the end of the experiment the volume of the protein solutions is measured by means of Pedersen micropipettes and the viscosity is determined.

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2.6.3. The Macromolecular Osmotic Pressure Induced by Poly(Ethylene Glycol) Osmotic pressure associated with poly(ethylene glycol) 40,000 solutions (up to 5 g per 100 g of water) is measured by means of osmometers equipped with UH 100125 Schleicher and Schuell membranes, Mr, cutoff 25,000. The macromolecular osmotic pressure generated by poly(ethylene glycol) is related to the concentration of the macromolecule by, Macromolecular osmotic pressure = 98 (9c + c2.71) Pa, where, c, is the concentration of poly(ethylene glycol) (weight per 100 ml of solution.

3. Theory In this section are treated: 1. the ideality and the non-ideality of the solutions, i.e. the relation between the concentration of the macromolecules and their chemical potential, 2. the conversion of the actin filaments into actin bundles. This conversion introduces a geometric constraint that relates the concentration of actin, the diameter of the actin filament and the inclination of the long axis of the actin monomer with respect to the axis of the filament. 3. Some mechanic properties of the actin filament such as the stiffness and the yield strength. These parameters, even though related to the chemical potential of actin, are more conveniently treated in a separate sub-section.

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3.1. Ideal and Non-Ideal Solutions The study of the osmotic pressure is a profitable way to investigate the properties of a solution since the change of the chemical potential of the solvent and of the solute are mutually linked through the Gibbs-Duhem relation (Eq. 1) and the change of the chemical potential of the solvent is linked to the osmotic pressure of the solution (Eq. 2): (nWdµW + nSdµS)T, P,nS = 0

(1)

ΔµW (T, Π, ai) = RT ln [aW] = -ΠΔV

(2)

where nW and nS are the numbers of moles of water and of the solute, µW and µS are the chemical potentials of water and of the solute, aW, is the activity of water, V is the partial molal volume of water (18 x 10-6 m3) and Π is the osmotic pressure. In an ideal solution the following equation holds: mW V Π / (RT x mS) = 1

(3),

where, mW and mS, are the molalities of water and of the solute, respectively. In concentrated solutions the ratio deviates from unity and equals the function, φ, the molal osmotic coefficient: Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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mW V Π / (RT x mS) = φ

(4)

These solutions are defined as non-ideal solutions. The non-ideal behaviour arises when the molecules of the solute compete for water. The competition, especially in protein solutions, in some cases favours the association between the molecules of the solute, always perturbs their hydration shell, influences their conformation and, sometime, their functions. 3.1.1. The Energetic of a Non-Ideal Solution The chemical potential of the solute, ΔµS, is calculated from Eq. 1, nWdµW + nSdµS = 0 (1) and Eq. 5. mW V Π / RT = - mW ln [aW] = φ mS

(5),

this latter is obtained by rearranging Eq. 2 and Eq. 4. RT d ln[aW] + nS / nW RT d ln[aS] = 0, RT d ln[aW] + mS / mW RT d ln[aS] = 0,

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mS RT d ln[aS] = - mW RT d ln[aW] = RT d(φ mS), mS RT d ln[aS] = RT (mS dφ + φ dmS),

(6)

dµS = RT (dφ + φ/mS dmS)

(7)

Integration of Eq. 7 yields, ΔµS. The activity coefficient of the solute, γS, is calculated from Eq. 6, (Edsall and Wyman 1958): mS d ln[aS] = mS d ln[mS] + mS d ln[γS] = (mS dφ + φ dmS), d ln[γS] = - d ln[mS] + dφ + φ/mS dmS = dφ + (φ – 1)/mS dmS d ln[γS] = - 1/mS + dφ + φ/mS dmS = dφ + (φ – 1)/mS dmS

(8) In our experiments the solutions are not strictly binary, in fact they are composed by water, the macromolecule and low molecular weight solutes. However, if the medium exchangeable through the dialysis membrane is considered as a single multi-component

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system, one can speak of chemical potentials of all species, the multi-component system, mu, the stressing polymer, p, and the macromolecule of interest, mc, by the Gibbs-Duhem relation for each phase:

nmudµmu = - nmc dµmc nmu dµmu = - nPdµP The chemical potential of each is related to the osmotic pressure of water in the two solutions (Parsegian and al., 1986). The fact that the solute is charged does not impair the validity of the measurements since the overall free energy change is referred to the system composed by the solute molecules plus the surrounding cloud of the neutralysing ions. This means that „the component is electrically neutral even when the species is charged‟ (Scatchard 1946). 3.1.2. Solution of Two Macromolecular Solutes Undergoing Association, the Reaction Being at Chemical Equilibrium So far we have dealt with binary solutions, in some cases, however, the treatment may be extended to ternary solutions. An example is the ternary solution composed by water and by the 1:1 myosin-actin complex formed from equimolar solutions of myosin and of F-actin as monomer (Grazi et al., 2001). The treatment below indicates how to determine the chemical potential of the 1:1 acto-myosin complex.

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A solution of F-actin plus myosin (Eq. 9) and a solution of myosin alone (Eq. 10) are equilibrated at the same osmotic pressure: nWdµW + nmdµm + nadµa + nmadµa + nma2dµma2 = 0

(9)

dµW = - (n*m / n*W) dµ*m

(10).

In the actomyosin solution, nW, nm, na, nma, nma2 indicate the number of moles of water, of free myosin, of free F-actin monomer, of the 1:1 actin-myosin complex and of the 2:1 actinmyosin compex, respectively. In the pure myosin solution, n*W and n*m indicate the number of moles of water and of myosin. In the acto-myosin solution the total number of moles of myosin equals the total number of moles of F-actin as monomer, furthermore, the two solutions contain the same amount of total myosin (n*m). It follows that: n*m = nm + nma + nma2 = na + nma + 2 nma2 and Eq. 9 becomes: nWdµW + (n*m – nma – nma2) dµm + (n*m – nma –2nma2) dµa + nma dµma + nma2 dµma2 = 0

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nWdµW + n*m(dµa + dµm) – nma(dµa + dµm) - nma2(dµa +dµm) – nma2dµa + namadµma + nma2dµma2 = 0

(11)

Since the system is at the equilibrium the following relationships hold: dµm + dµa = dµma; dµma + dµa = dµma2 and Eq. 11 becomes: dµW = -(n*m / nW) dµma

(12).

Since osmotic pressure is the same in both systems, Eq. 10 and Eq. 12 are comparable. If now, as it is experimentally feasible, the two systems undergo the same infinitesimal change of µW, the right terms of Eq. 10 and of Eq. 12 can be equated: (n*m / nW) dµma = (n*m / n*W) dµ*m and dµma = (nW / n*W) dµ*m = (m*m / mm) dµ*m

(13).

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where, m*m, is the molality of myosin in the solution containing myosin alone and, mm, is the concentration of total myosin in the solution containing both myosin and F-actin.

3.2. The Relationship between the Chemical Potential of Water and the Chemical Potential of the Free Actin Monomer in F-Actin Solutions In our studies we have usually related the free energy change of actin polymers, under osmotic stress, to the total concentration of actin as the monomer. The choice is justified by the fact that the system is highly non-ideal and that, in any case, its behaviour ought to be described phenomenologically by making use of an activity coefficient. It would thus appear an unnecessary complication to try to relate the free energy change to the number density of the actin polymers. However, and more rigorously, the change of the water chemical potential of the system can also be related to the change of the chemical potential of the free monomer of actin (Grazi and Pozzati 2008). This is possible when the system is at the equilibrium: n ADP-G-actin F-ADP-actin. Usually in the experiments we start from a system that is not at the equilibrium because monomeric actin is in the ATP-G-actin form. However the initial, low (0.1 mM) concentration of ATP and the long times of incubation allow the system to reach the

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equilibrium due to the treadmilling of the actin polymers (Wegner 1976; Wegner and Neuhaus 1981). At the equilibrium, according to the Gibbs–Duhem relationship, the following relation holds: -wdµw = a1 dµ1 + a2 dµ2 + a3 dµ3 + ……. + an dµn, (14) where w is the number of mol of water, a1 is the number of mol of the actin monomer, a2 to an are the number of mol of the actin polymers and the dµ are the chemical potential differentials of the single species. Since the system is at the equilibrium, we also write 2µ1 = µ2 µ1 + µ2 = µ3 µ1 + µ(n-1) = µn. The expression holds if we write 2µ1 + 2 dµ1 = µ2 + dµ2, µ1 + dµ1 + µ2 + dµ2 = µ3 + dµ3, µ1 + dµ1 + µ(n-1) + dµ(n-1) = µn + dµn We also write

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2 dµ1 = dµ2, dµ1 + dµ2 = dµ3, dµ1 + dµ(n-1) = dµn. Hence, Eq. 14 becomes -w dµw = dµ1 (a1 + 2 a2 + 3 a3 + ….. n an).

(15)

We observe now that m = a1 + 2 a2 + 3 a3 + …… + n an

(16)

is the total concentration of the monomers, free plus embedded in the polymers and by substituting the expression in Eq. 15 we obtain -w dµw = m dµ1

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(17)

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Thus, from the change of the water chemical potential we calculate the change of the chemical potential of the free monomer in the system. Since µw = µ°w + RT ln[aW] and Π V = RT ln[aW], where, aw, is the water activity, V is the water partial molal volume and Π is the macromolecular osmotic pressure, we write dµw = - V d Π

(18)

by introducing this expression in Eq. 17, we obtain dµ1 = (w / m) V d Π

(19)

Integration of Eq. 19 under the form: dµ1 = (w / m) V (d Π / dm) dm

(20)

yields Δµ1.

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3.3. Distribution and Energy of the Actin Filaments in the F-Actin Solution The distribution and the energy of the actin filaments can be calculated by making use of the model of Biron and al. (2006). The model shows that, at the equilibrium, short-range attractions enhance the tendency of filaments to align parallel to each other, and may 1. increase the average filament length 2. decrease the relative width of the distribution. With the same model the free energy (per unit volume) of the solution containing the actin filaments is calculated. The essential features of the model are summarized below. When the interactions are absent the distribution of the filaments is exponential, ρl(0) = ρ0 exp[-a l]

(21)

with an average filament length, < l >0 = 1/a = (ρm / ρ0)1/2 and Cσ = 1. In the presence of short-range attractions the distribution is described by the two expressions ρl = ρ0 exp[-al + (gl* ρ*) l2] when l ≤ l* ρl = ρ0 exp[-a + (gl*2 ρ*) l] when l ≥ l*

(22)

With three unknown a, l*, ρ*, the distribution function, ρl, can be calculated by solving the following three equations:  

the monomer conservation condition, ∫ dll ρ1 = ρm the consistency condition,

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ρ* = ρ0 exp[-al + gl*3 ρ*] 

(23)

the condition that < l > = l*.

The free energy of the filaments in an F-actin solution is, f = ∑l ρl (ln[ρl v0] – 1 + αl + b) + ∑l0 = average length of the filaments; m, molality µ, chemical potential (J/mol) µ°, standard chemical potential (J/mol) µ, free energy of the actin monomer; Π, macromolecular osmotic pressure (Pa) R, gas constant; ρm, total monomer concentration; ρ0 = exp[-b] / v0; ρ1, numerical concentration of the free actin monomer; ρl(0), numerical concentration of the filament in the absence of interactions; ρ*, numerical concentration of the filament of average length; ρl, numerical concentration of the filament, l, in a distribution characterized by short range interactions; σ1, width of the distribution; T, absolute temperature; u0 < 0, short range attraction per monomer; V, partial molal volume of water (18 x 10-6 m3); v0, volume of the monomer; z ≡ (l1 / d) u0;

4. Applications to Theory

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4.1. The Actin Filament, the ‘Fluttering Wing’ Model Actin monomers assemble spontaneously into a filamentous helical structure, F-actin, that may undergo association into hexagonally packed bundles of filaments (De Rosier and Tilney, 1982). This packing is not impaired by large osmotic stress increases even though more and more protein-bound water is withdrawn and the volume of the protein solution decreases. It is thus possible to relate osmotic pressure to the geometric parameters of the supramolecular structure. By making use of this technique (Theory, 3.4.) it was shown that osmotic pressure dictates the diameter of the actin filament (Table 1) (Grazi and al. 1993). At that time the available models devised the actin filament as a rigid structure. This is true: 1. For the models generated from optical diffraction of actin paracrystals where the diameter of the actin filament ranges between 8 nm (Moore and al. 1970) and 7 nm (Spudich and al. 1972). 2. For the Heidelberg model generated by comparing the X-ray fiber diffraction pattern of an oriented gel of F-actin with the X-ray diffraction pattern of the actin monomer in the actin-DNAse I complex (Lorenz and al. 1993; Tirion and al. 1995). In the Heidelberg model the location and nature of a number of possible intermolecular bonds between adjacent actin monomers were defined and a diameter of 9.5 nm was assigned to F-actin (Lorenz and al. 1993; Tirion and al. 1995). 3. For the ribbon to helix transition model, proposed by Schutt and al. (1995) on the basis of the 2.55 Å resolution of the structure of crystalline profilin-β actin complex. In the model the hypothesis was formulated that actin may form a ribbon (diameter 9.5 nm) capable of undergoing a ribbon to helix transition into the actin filament. 4. Hegelman and Padron (1984) generated a model by using the integrated intensity of the 5.9

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nm layer line from the X-ray diffraction pattern of live relaxed frog sartorius muscle and the phases determined by electron microscopy of negatively stained, isolated actin filaments. They concluded that in vivo the actin filament displays a diameter of 10 nm. Table 1. The average diameter of the hydrated actin filament as a function of the osmotic stress Osmotic pressure, kPa 10.0 18.1 46.4 85.0 90.0

Actin filament diameter, nm 9.00 7.95 7.56 7.28 6.80

The value of the diameter of the actin filament is calculated as described under Theory, 3.2 (Grazi and al. 1993).

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At variance with the above models the „fluttering wing model‟ predicts that the diameter of actin is variable, it depends on the macromolecular osmotic pressure and, eventually, on the presence of actin bundling proteins. The fact that the diameter of the actin filament changes as a function of the macromolecular osmotic pressure implies also that the location and nature of the intermolecular bonds between adjacent actin monomers is likely to change. In conclusion the actin filament in vivo is not a rigid structure but changes in shape. This change is modeled by the decrease of the angle formed between the long axis of the monomer and the pointed end of the filament axis (Theory, 3.4.4.). As a result the actin monomer acts as a „fluttering wing‟ with the change of osmotic pressure (Grazi 1997).

4.2. Comparison of the Osmotic Properties of F-Actin and of Tropomyosin-F-Actin Figure 2 shows that, at a set of different osmotic pressures, the interfilament distance of tropomyosin-F-actin is larger than that of F-actin. This suggests that, in agreement with the data of Milligan and Flicker (1987) and of Lorenz and al. (1995), the two tropomyosin helices project out of the contour of the actin filament. Moreover, the atomic model of Lorenz and al. (1995), shows also that the closest distance between Cα atoms of tropomyosin and actin is about 10.5 Å which is too great to allow any but ionic interactions. Therefore the interaction between actin and tropomyosin must be essentially electrostatic. At the physiological protein osmotic pressure of frog muscle 24 kPa, (vertical line in the figure), (Maughan and Godt 2001), the average diameter of hydrated F-actin is ~8.2 nm and that of hydrated tropomyosinF-actin is ~10.4 nm.

4.3. Osmotic Properties of the Calcium-Regulated Actin Filament In 2 mM EGTA and at osmotic pressures ranging between 10 and 54 kPa, the interfilament distance of troponin-tropomyosin-decorated F-actin is practically constant, the

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average value is ~13.3 nm. The interfilament distance is significantly reduced in 0.2 mM CaC12, being 12 nm at 10 kPa and 11.4 nm at 54 kPa. The values of the interfilament distance converge to those found in the presence of EGTA at pressures larger than 800 kPa (Figure 3). At the physiologic protein osmotic pressures, 24 kPa ((Maughan and Godt 2001), the interfilament distance is ~13.3 nm in 2 mM EGTA and ~12.3 nm in 0.2 mM Ca2+. According to the model of Pirani and al. (2006) the bulk of the core domain of troponin does not move significantly on actin regardless of Ca2+ level, furthermore, at high Ca2+, tropomyosin shifts from the actin outer domain toward the actin inner domain, without any significant movement toward the center of the actin filament. This model therefore does not seem to explain the decrease of the interfilament distance of troponin-tropomyosin-decorated-F-actin in the presence of Ca2+. The possibility remains that, in the presence of Ca2+, the orientation of the monomer in the actin filament changes because of a decrease of the angle α (Theory 3.4.4.) thus pulling at a lower radius both tropomyosin and troponin.

Figure 2. The interfilament distance of F-actin (filled circles) and of tropomyosin-F-actin (open circles) as a function of the protein osmotic pressure. The vertical line indicates the physiological protein osmotic pressure of frog muscle, 24 kPa (Maughan and Godt 2001). The interfilament distance is calculated according to Theory 3.2.3. Data are taken from Schwienbacher and al. (1995).

Figure 3. Interfilament distance of Troponin-Tropomyosin-decorated F-actin, in the presence (open circles) and in the absence (filled circles) of Ca2+, as a function of the protein osmotic pressure. The vertical line indicates the physiological protein osmotic pressure of frog muscle, 24 kPa (Maughan and

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Osmotic Pressure Godt 2001). The interfilament distance is calculated according to Theory 3.2.3. Data are taken from Schwienbacher and al. (1995).

An additional, likely alternative is that the addition of Ca2+ contributes to the neutralization of the negative charge of the actin filaments thus decreasing the interfilament distance.

4.4. The Energetic of Solutions with 1. A Single Macromolecular Solute 2. Two Macromolecular Solutes Undergoing Association, the Reaction Being at Chemical Equilibrium The investigation of the osmotic properties of a binary solutions allows to define the energetic of the solute. Furthermore the same technique can be applied, in some cases, to a ternary solution composed by water and by two macromolecules, associated and at the chemical equilibrium (Grazi and al. 2001). 4.4.1. The Energetic of F-Actin Solutions By equilibration against solutions of increasing macromolecular osmotic pressure the concentration of an F-actin solution increases from 5.67 mmolal at 1.7 kPa to 22.2 mmolal at 223.8 kPa. The relation between the concentration of actin and the osmotic pressure is described by the equation:

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Π = 2.45 x 106 (mA – mA1.04 + 1.28 x 107 mA5.09) Pa

(32)

Eq. 32 allows to calculate: 1. the osmotic molal coefficient, φ, 2. the change of the chemical potential of actin, ΔµA, and 3. the activity coefficient of actin, γA. The osmotic molal coefficient is obtained from Eq. 4 (Theory 3.1.) 1000 Π / (RT x mA) = φ (4). The, ΔµA, is calculated from the equation,

(33) by taking as the reference 5 mmolal F-actin (as monomer) that corresponds to the protein osmotic pressure of 2.4 kPa. The logarithm of the activity coefficient of actin is calculated by Eq. 8 (Theory 3.1.1.)

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Table 2. Calculated protein osmotic pressure, molal osmotic coefficient, ln γ and Δµ for F-actin as a function of F-actin, as monomer Osmotic Pressure Molal osmotic Ln[γ] Δµ (J.mol-1) kPa coefficient 5 2.4 0.196 - 21.028 0 6 2.9 0.195 - 21.175 72.7 7 3.4 0.199 - 21.295 135 8 4.1 0.209 - 21.391 191 9 5.0 0.226 - 21.466 244 10 6.2 0.253 - 21.520 297 11 7.8 0.290 - 21.553 352 12 10.0 0.340 - 21.562 412 13 12.9 0.407 - 21.546 478 14 16.9 0.492 - 21.502 553 15 22.0 0.599 - 21.427 639 16 28.6 0.731 - 21.317 739 17 37.1 0.891 - 21.168 854 18 47.8 1.084 - 20.975 986 19 61.2 1.314 - 20.735 1140 20 77.7 1.585 - 20.441 1320 21 97.8 1.901 - 20.089 1520 22 122.2 2.268 - 19.672 1750 Osmotic pressure is calculated from the equation: Π = 2.45 x 106 (ma - ma1.04 + 1.28 x 107 ma5.09) Pa. Molal osmotic coefficient, ln[γ] and Δµa are calculated as described in the text, with reference to 5 mmolal F-actin, as monomer, corresponding to the calculated protein osmotic pressure of 2.4 kPa. Data are taken from Grazi et al. (2001). Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

Actin mmolal

Due to the high polymerization degree of Factin, the molal osmotic coefficients are lower than one and are expected to tend to one at infinite dilution of actin and accompanying solutes. Ln[γA] is negative even at F-actin concentrations larger than 18 mmolal, where the molal osmotic coefficient is larger than one. This is because Ln[γA] is obtained by integration starting from zero F-actin concentration, Eq. 8. The high negative value of Ln[γA] indicates the much higher stability (lower energy) of the F-monomer as compared to the G-monomer in the ideal solution. 4.4.2. The Energetic of the Myosin Solutions By equilibration against solutions of increasing macromolecular osmotic pressure the concentration of a myosin solution increases from 0.72 to 1.75 mmolal with the increase of the pressure from 13.5 to 479 kPa. The phenomenon is described by the equation, Π = 2.45 x 106(m*m + 100 m*m2 + 1.7 x 1010 m*m4), Pa,

(34).

Eq. 34 allows to calculate: 1. the osmotic molal coefficient, φ, 2. the change of the chemical potential of myosin, ΔµM, and 3. the activity coefficient of myosin, γM. The osmotic molal coefficien is obtained from Eq. (4) (Theory 3.1.)

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Osmotic Pressure 1000 Π / (RT x mm) = φ (4). The, Δµm, is calculated from the equation,

(35) by taking as the reference 0.368 mmolal myosin which corresponds to the protein osmotic pressure of 1.7 kPa. The logarithm of the activity coefficient of myosin is calculated by Eq. 8 (Theory 3.1.1.)

The results are summarized in Table 3. 4.4.3. The Energetic of a Solution of Myosin and F-Actin, Equimolar as Monomers and at the Chemical Equilibrium Equimolar solution, as the monomer, of myosin and F-actin are equilibrated against macromolecular solutions at increasing osmotic pressures. As a result the concentration of total myosin, mm, increases from 0.45 to 1.8 mmolal with the increase of the osmotic pressure from 1.7 to 229 kPa. Eq. 36 describes the phenomenon:

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Π = 2.45 x 106(mm + 2600 mm2 + 7 x 109 mm3.99), Pa

(36).

To calculate the free energy of formation of actomyosin from F-actin and myosin, the behaviour of the actomyosin solutions is compared with that of the solutions containing myosin alone. At the time of mixing the actomyosin solutions contain n*m moles of myosin and an identical number of moles of F-actin (as monomer). At the equilibrium the solutions contain, moles of free myosin = (n*m – nma – nma2) moles of F-actin (as monomer) not bound to myosin = (n*m – nma – 2 nma) moles of the 1:1 actin-myosin complex: nma moles of the 2:1 actin-myosin complex: nma2. The pure myosin solutions (Table 3) contain n*m moles of myosin and are equilibrated at the same osmotic pressures as the actomyosin solutions. To compare the two sets of solutions, it is convenient to express the molality of myosin as a function of the protein osmotic pressure. Therefore, Eq. 34 and Eq. 36 are converted into their equivalents: 1. Eq. 37 that holds for the solutions of pure myosin and is equivalent to Eq. 34,

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Enrico Grazi m*m = 0.00034977 + 1.16659 x 10-5 Π0-4 – 2.00647 x 10-9 Π

(37)

and Eq. 38 that holds for the actomyosin solutions and is equivalent to Eq. 36, mm = 0.0000171694 (1+ 1.51356 Π0.18 + 2.04174 Π0.31)

(38).

We now take into account that, dµma, is expressed by Eq. 13 (Theory 3.1.2.) dµma = (nW / n*W) dµ*m = (m*m / mm) dµ*m (13), and that, dµ*m, refers to the pure myosin solution, a binary solution, defined by Eq. (7) (Theory 3.1.2.) dµ*m = RT (dφ* + φ*/m*m dm*m) (7) where the asterisk indicates the solutions with myosin alone. Substitution of Eq. (7) into Eq. (13) yields, dµma = (m*m / mm) x RT x (dφ* + φ*/m*m x dm*m)

(39)

where, φ* and m*m are the molal osmotic coefficient and the molality of myosin in the solutions with myosin alone while mm is the molality of total myosin in the F-actin plus myosin solutions.

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Table 3. Protein osmotic pressure, molal osmotic coefficient, lnγM and Δµ of myosin as a function of myosin molality Osmotic Pressure Molal osmotic ln[γM] Δµ J.mol-1 (kPa) coefficient 0.7 11.8 6.90 7.91 1.80 x 104 0.8 19.1 9.78 11.76 2.77 x 104 0.9 29.7 13.48 16.70 4.01 x 104 1.0 44.3 18.10 22.86 5.55 x 104 1.1 63.9 23.73 30.38 7.42 x 104 1.2 99.6 30.49 39.41 9.65 x 104 1.3 122.5 38.48 50.06 12.28 x 104 1.4 163.9 47.72 62.47 15.34 x 104 1.5 215.0 48.52 76.80 18.86 x 104 1.6 277.5 70.79 93.16 22.89 x 104 1.7 352.7 84.69 111.70 27.44 x 104 1.8 442.4 100.32 132.55 32.57 x 104 Osmotic pressure is calculated from the equation: Π = 2.45 x 106(mT + 100 mT2 + 1.7 x 1010 mT4), Pa. Molal osmotic coefficient, ln[γ] and ΔµM are calculated as described in the text, with reference to 0.368 mmolal F-actin, as monomer, corresponding to the calculated protein osmotic pressure of 1.7 kPa. Data are taken from Grazi et al. (2001). Myosin mmolal

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Osmotic Pressure

The expressions for the molality as a function of protein osmotic pressure, Eq. (37) and (38), are then introduced into Eq. (39) that by integration, becomes:

(40) The free energy of formation of actomyosin is calculated by numerical integration of Eq. 40, after analytical differentiation of the terms, d*mm/dΠ and dφ*/ dΠ. In the equation the terms marked by the asterix belong to the system containing myosin alone. The ratios of the activities are calculated from the equation: Δµma / RT = ln[a‟ma / ama]

(41)

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4.5. The Free Energy of the Free Actin Monomer and the Length Distribution of the Actin Filaments At the steady state the length distribution of actin filaments is exponential (Kawamura and Maruyama 1970; Burlacu and al., 1992; Edelstein-Keshet and Ermentrout, 1998; Ermentrout and Edelstein-Keshet, 1998; Sept and al., 1999; Fujiwara and al., 2002; Biron and Moses, 2004) and is reported to display a mean length of 7 µm (2600 actin monomers). This mean is independent of the initial concentration of the actin monomer from 5 to 100 µM (Sept and al., 1999). Various factors may alter the exponential distribution. These are: 1. the charge neutralization with multivalent cations (Strzelecka-Golaszewka and al., 1978; Grazi and al., 1982) 2. the increase of protein osmotic pressure that promotes the conversion of the actin filaments into bundles of filaments (Suzuki and al., 1989; Schwienbacher and al., 1995) 3. the actin crosslinking protein, such as actinin and the capping proteins (Biron and Moses 2004). Biron and al. (2006) further confirm these findings and predict that, at the equilibrium, shortrange attractions enhance the tendency of filaments to align parallel to each other, eventually leading to an increase in the average filament length and a decrease in the relative width of the distribution of filament lengths. The model of Biron and al. (2006) offers a tool to calculate the length distribution of the actin filaments and their free energy. 4.5.1. Modeling the Length Distribution of the Actin Filaments as a Function of the Free Energy of the Free Actin Monomers The osmotic tool, applied to the model of Biron and al. (2006), relates the free energy of the free actin monomers to the length distribution of the actin filaments, provided that the Factin system is in equilibrium: n (ADP-G-actin) = F-ADP-actin. Under these circumstances, as it is shown later, the actin monomer and all the actin filaments display the same concentration. Furthermore, the perturbation of the osmotic equilibrium due to the addition of an external macromolecule, induces a new actin filament length distribution, due to the onset of short-range attractions. This new filament length

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distribution is reproduced provided that the corresponding value of the short-range attractions is known. This latter is the value of the short-range attractions that makes the change of the free energy of the free actin monomer, calculated according to Biron et al. (2006), equal to that obtained from the osmotic experiments.

4.5.1.1. Protein Osmotic Pressure Associated with F-Actin Solutions of Moderate Concentrations

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F-actin in Buffer A solution, being a non-diffusible species, is expected to generate an inward flux of water when dialyzed against Buffer A (Methods). In the experiment 1 mL aliquots of F-actin, 10–60 µM as monomer, are dialysed at 21°C, even for 288 h without a significant change of volume. The volume recovered is 1 ± 0.005mL. The change of the water chemical potential induced by F-actin is thus undetectable (Figure 4 open circles).

Figure 4. Macromolecular osmotic pressure (Pa) as a function of total actin concentration. Open circles: F-actin, either 19.18 or 60 µM, dialysed against Buffer A. Filled circles: F-actin dialysed against Buffer A plus increasing concentrations of poly(ethylene glycol), initial actin concentration 19.18 µM. Open triangles: F-actin dialysed against Buffer A plus increasing concentrations of poly(ethylene glycol), initial actin concentration 60 µM. Data are taken from Grazi and Pozzati (2008).

4.5.1.2. Equilibration of F-Actin Solutions against Poly(Ethylene Glycol) 40,000 Solutions Equilibration of F-actin solutions against poly (ethylene glycol) solutions of increasing concentration promotes a significant decrease of the volume of the protein solution. The final actin concentration is calculated by the formula: Final concentration = (initial volume x initial concentration) / (final volume) In Figure 4 the macromolecular osmotic pressure (Π) at the equilibrium is plotted against the actin concentration, as monomer. In the first series of experiments (filled circles) the

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starting concentration of actin is 19.18 µM, as monomer. The samples are equilibrated for 168 h in the presence of increasing concentrations of poly(ethylene glycol). At the end of the experiments the volume of the samples is measured and the actin concentration is calculated. The relation between total actin concentration, c (molarity), and the macromolecular osmotic pressure, Π, is given by, Π = -1750.666+1.876 x 108 c – 2.7338 x 1010 c1.5 + 1.2539 x 1012 c2 – 8.7418 x 1014 c3, Pa (42). The equation is only aimed at describing the data and is valid only in their range. In the second series of experiments (open triangles) the starting concentration of F-actin is 60 µM, as monomer. The samples are equilibrated for 173 h in the presence of increasing concentrations of poly(ethylene glycol). At the end of the experiments the volume of the samples is measured and the actin concentration is calculated. The relationship between total actin concentration, c, molarity, and the macromolecular osmotic pressure is given by, Π = - 1376.1 + 2.839 x 107 c - 9.3939 x 1010 c2 + 9.544 x 1013 c3 + 7.837 x 1010 c4, Pa (43) The equation is only aimed at describing the data and is valid only in their range. These results show the divergent properties of resting and perturbed F-actin solutions. Resting solutions of either 19.18 or 60 µM actin do not develop a detectable osmotic pressure and the water chemical potential of the solution is essentially that of Buffer A. On the contrary the actin solution concentrated from 19.18 to 54.86 µM develops a macromolecular osmotic pressure of 1085.2 Pa, i.e. the change of the water chemical potential is

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ΔµW = - ΠV = - 1085.2 x 18 x 10-6 = 0.0195336 J/mol of water, or - 7.877 x 10-6 RT/mol of water. where, V, is the partial molar volume of water, 18 x 10-6 m3. This means that the water chemical potential of the actin solution is lower by 7.877 x 10-6 RT/mol of water than the water chemical potential of buffer A. The different behaviour of the two actin solutions, of almost identical concentration is probably explained by a different actin filament length distribution.

4.5.1.3. The Macromolecular Osmotic Pressure and the Free Energy of the Actin Monomer It can be shown that the free energy of the free actin monomer changes with the macromolecular osmotic pressure associated to the actin solution (Theory 3.2.). Integration of Eq. 20 yields the change of the chemical potential of the free monomer, dµ1 = (w / m) V (d Π / dm) dm (20)

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The integration is performed after substitution of the term Π either with Eq. 42 or with Eq. 43 that relates the macromolecular osmotic pressure to the total concentration of actin. In the first experiment the increase of total actin concentration from 19.18 to 48 µM and to 226 µM (filled circles) is accompanied by the increase of the free energy of the free actin monomer by 13.36 and by 16.77 RT/mol of free monomer, respectively. In the second experiment the increase of total actin concentration from 60 to 98.18 µM and to 135 µM is accompanied by the increases the free energy of the free actin monomer by 3.13 and by 4.5 RT/mol of free monomer, respectively (Figure 5).

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Figure 5. The change of the free energy of the free actin monomer as a function of the total actin concentration. Filled circles: initial actin concentration 19.18 µM; open triangles: initial actin concentration 60 µM. Data are taken from Grazi and Pozzati (2008).

4.5.1.4. Calculating the Actin Filament Length Distribution at the Steady State The mean of the length distribution of the actin filaments is ~7 µm and is independent of the actin concentration up to 100 µM total actin, as monomer (Sept and al. 1999). This length distribution is reproduced by making use of Eq. 22 (Theory 3.3.). This is done by setting the average numerical length, l*, to 2560 = 7000 nm / 2.73 nm, where 2.73 nm is the length increment per actin subunit (Hanson and Lowy, 1963), by setting the maximum numerical length of the filaments to 25,600 (69,888 nm) and by setting the short-range attraction per monomer, u0, to the very low value of -10-15 kBT, in order to minimize the short-range attractions and to obtain an exponential curve (Theory 3.3.). These operations are performed at two F-actin concentrations, 19.18 and 110 µM, as the monomer (Figure 6). In both distributions the average length of the filaments is 7 µm but the concentration of the free monomer is 2.923 and 16.744 pM at 19.18 and 110 µM total actin, respectively.

4.5.1.5. Filament Distribution Associated with the Free Energy Minimum of the Actin Filaments By Eq. 23 and Eq. 24 (Theory 3.3.) the total free energy of the actin filaments, displaying the length distribution shown in Figure 6, is calculated. These free energies are 7.495 x 10-9 RT and 4.29 x 10-8 RT (per liter of solution) for the solutions containing 19.18 µM and 110 µM total actin, respectively.

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Figure 6. Distribution of the actin filaments in F-actin solutions at the steady state. Upper line: total monomer concentration, 110 µM; free monomer concentration, 1.6744 x 10 -11 M; lower line: total monomer concentration, 19.18 µM; free monomer concentration, 2.923 x 10 -12 M. The vertical line indicates the average filament length, 7 µm. Data are taken from Grazi and Pozzati (2008).

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However the free energy minima of the actin filaments are attained at the average filament length of 34,944 nm (12,800 monomers) and are, 1.5 x 10-9 RT and 8.594 x 10-9 RT (per liter of solution) at 19.18 µM and 110 µM total actin. At the free energy minimum all the actin filaments, monomer included, display the same concentration, 0.0586 and 0.3357pM at 19.18 µM and at 110 µM total actin, respectively (Figure 7). The distribution of the actin filaments described by Sept et al. (1999) is not at the free energy minimum because the system is not at the equilibrium.

Figure 7. Distribution of the actin filaments in F-actin solutions at the equilibrium. Upper line: total monomer concentration, 110 µM; free monomer concentration, 3.357 x 10 -13 M; lower line: total monomer concentration, 19.18 µM; free monomer concentration, 5.8638 x 10 -14 M. The vertical line indicates the average filament length, 34.94 µm. The value of Cσ = 0.557328 and of u0 = - 10-15 kBT is the same for the two samples; the value of a equals 1.4195 x 10-30 kBT for 19.18 µM actin and 8.1269 x 10-30 kBT for 110 µM actin. Data are taken from Grazi and Pozzati (2008).

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In fact in the experiments of Sept and al. (1999) polymerization is started from ATP-Gactin, thus is coupled to the hydrolysis of ATP. Only after the complete hydrolysis of the ATP in the solution, the system is simply described by the reaction, n (ADP-G-actin = F-ADP-actin, the actin filaments reach their free energy minimum and their distribution is characterized by all the filament concentrations being equal.

4.5.1.6. The Effect of the Dialysis against Poly(Ethylene Glycol) on the Length Distribution of the Actin Filaments The actin solutions dialysed either against Buffer A or against Buffer A plus poly(ethylene glycol) behave differently (Figure 4). We made the hypothesis that the difference is due to a rearrangement of the distribution of the actin filaments, so that the distribution of the actin filament of a 60 µM resting actin solution is different from that obtained by concentrating to 60 µM actin an originally 19.18 µM actin solution. This is because: 1. In resting solutions the long time of dialysis allows the complete hydrolysis of ATP and the attainment of the equilibrium. 2. In the solution perturbed by dialysis against poly(ethylene glycol), short range attractions are induced (Biron and al. 2006) and the system is shifted toward a new filament length distribution. Table 4. Observed and calculated concentrations of total myosin in the F-actin-myosin solutions. Activities ratios and Δµma of the 1:1 actin-myosin complex as a function of protein osmotic pressure Total myosin Activity ratios of Δµma J.mol-1 calculated, the 1:1 actomyosin mmolal complex 1.7 0.450 0.458 1.0 0 3.6 0.600 0.574 4.8 0.38 x 104 6.9 0.635 0.687 39.4 0.90 x 104 8.5 0.830 0.729 97.0 1.12 x 104 10.0 0.830 0.763 219.0 1.32 x 104 13.2 0.770 0.824 1166 1.73 x 104 14.0 0.800 0.838 1683 1.82 x 104 16.0 0.860 0.870 4483 2.06 x 104 18.0 0.890 0.900 11005 2.28 x 104 20.9 0.860 0.938 40629 2.60 x 104 32.3 1.000 1.060 4.26 x 106 3.74 x 104 10 56.2 1.325 1.240 1.90 x 10 5.80 x 104 14 85.1 1.560 1.400 1.51 x 10 8.00 x 104 29 229.0 1.800 1.860 4.00 x 10 16.70 x 104 Total myosin concentration is calculated by Eq. 38. The ratios of the activities of the 1:1 actomyosin complex are calculated by Eq. 41. The Δµ for the 1:1 actin-myosin complex is calculated by Eq. 40. The pressure of reference is 1.7 kPa, the lowest experimental pressure attained in our experiments.

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Π, kPa

Total myosin observed, mmolal

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Table 5. Comparison of the properties of resting F-actin solutions with the properties of perturbed F-actin solutions concentrated by dialysis against poly(ethylene Glycol)

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Total actin monomers Free actin Cσ u0 kBT / mon (µM) monomers (pM) Resting F-actin solutions 38.40 0.113774 0.577328 - 10-15 2.80 x 10-8 -15 54.86 0.162581 0.577328 - 10 2.70 x 10-8 -15 93.66 0.277516 0.577328 - 10 3.56 x 10-8 Perturbed F-actin solutions 38.40 0.11371 0.577302 -5.10 x 10-7 11.376 54.86 0.16258 0.577287 -5.33 x 10-7 14.215 93.66 0.277452 0.577278 -4.51 x 10-7 15.778 213.44 0.669419 0.577271 -3.10 x 10-7 16.770 The original concentration of the perturbed F-actin solutions, concentrated by dialysis against poly(ethylene glycol) is 19.18 mM. The concentrations indicated in the Table are reached after equilibration with poly(ethyleneglycol) solutions generating the macromolecular osmotic pressure of 753.5, 1085.2, 1328.1, 1730.5 Pa, respectively.

To mimic this situation it is sufficient to decrease the value of u0 (the short-range attraction per monomer) until the change of the free energy of the free actin monomer, calculated according to Biron et al. (2006), matches that obtained in the osmotic experiments. At this point the distribution of the actin filaments is calculated. The results of the calculations are presented in Table 5 where the properties of the resting F-actin solutions are compared with the properties of the perturbed F-actin solutions of the same concentration, obtained starting from actin solutions of lower concentration. As shown in Table 5, the concentration of the free actin monomer of the perturbed Factin solutions is very slightly decreased, and Cσ is slightly decreased compared to that of the resting F-actin solutions. The ratio, free energy/free actin monomer, increases from ~10-8 kBT (resting solutions) to 14.215 kBT (perturbed solutions). Concomitantly u0 (the short-range attraction per monomer) decreases from –10-15 kBT (resting solutions) to -5.33 x 10-7 kBT (perturbed solutions). Surprisingly u0 reaches its lowest value, -5.33 x 10-7 kBT, at 54.86 µM total perturbed actin while it increases at larger total actin concentrations, being, -3 x 10-7 kBT, at 213.44 µM total perturbed actin. It is unreasonable that short-range attractions decrease with the increase of the concentration; it is thus likely that one or more influent parameters escaped recognition in the model. The effects of the change of u0 on the distribution of the actin filament is presented in Figure 8 where it is shown the behavior of a 19.18 µM actin solution concentrated, by dialysis against different concentrations of poly(ethylene glycol), to 38.4 µM (upper figure), 54.84 µM (middle figure) and 93.66 µM (lower figure). As shown in the figure, the concentration of the actin filaments increases with their length while, at equilibrium (Figure 7), the concentration is independent of the length. As predicted by the theory, all the curves display a break at the mean length.

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Figure 8. Proposed actin filament length distribution in F-actin solutions equilibrated against poly(ethylene glycol) - Abscissa, filament length, µm; ordinate, filament concentration, pM. Initial Factin concentration, 19.18 µM, as the monomer. Final F-actin concentration: upper figure, 38.4 µM; middle figure, 54.86 µM; lower figure, 93.66 µM. The break coincides with the mean length of the actin filament, 34.94 µm. The maximum filament length is 69.89 µm. Data are taken from Grazi and Pozzati (2008).

4.6. The Linear Relationship between Stiffness and Yield Strength Allows to Estimate the Yield Strength of Thin Filament In Vivo When stretched, the actin filament elongates. The force required to elongate by 1 nm a filament 1 µm in length is defined as the stiffness (pN.nm-1). If elongation exceed a given limit the actin filament breaks: the force required to break the filament is defined as the yield strength (pN). As indicated in Table 6, at comparable ionic strength, the stiffness and the yield strength of different models of the actin filament tested changes by more than two orders of magnitude while the specific elongation is essentially constant (0.0071–0.0097). A plot of the yield strength against the stiffness shows a linear behaviour, described by the equation: Yield strength = -2.368 + 9.889 stiffness (pN)

(44).

The linearity between stiffness and yield strength depends on the almost constant value (0.0071 – 0.0097) of the specific elongation at the yield point, that is only slightly influenced by the chemical modifications (coupling with tetramethylrhodamine iodoacetamide) and by Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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the decoration with either phalloidin or tropomyosin or myosin subfragment 1. The yield of the filament occurs when the average distance between the contact surfaces of the monomers increases by 0.19 to 0.26 Å. Also the work required to break the various models of the actin filament differs by more than two orders of magnitude, as calculated from the equation: Work = stiffness . (elongation at the yield point)2 /2 (pN.nm)

(45).

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These behaviours are reasonable since: 1. The specific elongations at the yield point (i.e. the bonding length) depends on the nature of the inter-monomer contacts, mostly hydrophobic interactions (Holmes and al., 1990), a feature which does not change with the model of the actin filament. 2. The work required to break the actin filament depends on the extent of the inter-monomer contact area that, on the contrary, is likely to change significantly in the different models (Grazi, 1997). By means of Eq. 44 the yield strength of the actin filament is estimated. As an example from the stiffness, 11.26 pN.nm, of the actin filaments decorated with alexa-fluor tropomyosin the yield strength of 111 pN is calculated (Figure 9). From the stiffness in vivo of molluscan smooth muscle in the active, 33 pN.nm, and in the catch, 26 pN.nm, states (Tojima et al. 1994), the corresponding yield strengths are calculated to be 324 pN in the active state and 255 pN in the catch state (Figure 9).

Figure 9. Relationship between yield strength and stiffness. Data are taken form Table 6. The straight line is described by Eq. 44. Filled circles: observed yield strengths (Tsuda and al. 1996; Adami and al. 2002). Calculated yield strengths: Open circle, alexa-fluor tropomyosin F-actin; Empty diamond, smooth muscle thin filament active state; Filled diamond, smooth muscle thin filament catch state.

4.7. Protein Osmotic Pressure and Cross-Bridge Attachment Determine the Stiffness of Thin Filaments in Muscle Ex Vivo I describe here the effects of ex vivo conditions, particularly of the macromolecular osmotic pressure and of the attachment – detachment of the cross-bridges, on the stiffness of the actin filament. These effects may explain the greater stiffness of thin filaments ex vivo

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with respect to that of F-actin in vitro. I tentatively compute the stiffness of thin filaments in relaxed muscle and assess the upper limit of thin filament stretching ex vivo (Grazi and Di Bona 2006).

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4.7.1. The Effect of Protein Osmotic Pressure on the Stiffness of Both Cross-Bridge and the Actin Filament In vitro protein osmotic pressure influences the stiffness of the cross-bridges (Grazi and al. 1996), furthermore cross-bridge attachment increases the stiffness of the actin filament (Adami et al. 2002, Trombetta and al. 2005). Ex vivo from the overall compliance of the frog muscle half sarcomere, 1.05 mm in length, Linari and al. (1998) computed the compliance of the remaining free part of the thin filament. At 4°C, the estimated thin filament stiffness was 58.46 pN nm-1 µm-1 in isometric contraction and 87.69 pN nm-1 µm-1 in rigor. These values were obtained from an isometric tension of 226 kN·m-2 in the intact fiber and on the assumption that there are 1015 thin filaments·m-2. It is illuminating to compare these values with the stiffness of two models of the actin filament: tropomyosin-decorated tetramethyl-rhodamine-phalloidin F-actin, stiffness 65.3 ± 6.3 pN nm-1 µm-1 (Kojima and al. 1994) and F-actin decorated with alexa-fluor tropomyosin, stiffness 11.26 ± 2.4 pN nm-1 µm-1 (Adami and al. 2003). The stiffness of the model of Kojima and al. (1994) approaches that of thin filaments in muscle while that of the model of Adami and al. (2003) is 5–7.8 times lower even though both the species are composed of actin decorated with tropomyosin. 4.7.2. Possible Reasons for the Different Behavior of the Two Models We have repeatedly criticized the use of phalloidin F-actin as a model for thin filaments (Cintio and al. 2001, Cuneo and al. 1995, Trombetta and al. 2002). Here I only observe that phalloidin F-actin exhibits an extremely low critical concentration as compared to F-actin (Wielan and al. 1975, Kishino and Yanagida 1988) and that the critical concentration is inversely related to the stiffness of the actin filament (Adami et al. 1999). Thus it is not surprising that the model of Kojima et al. (1994) exhibits stiffness as great as 65.3 ±6.3 pN nm-1 µm-1. The model of Adami et al. (2003), on the contrary, exhibits a lower stiffness than that of thin filaments. We propose that this is due to the effect of protein osmotic pressure. The protein osmotic pressure in muscle is ~24 kPa (Maughan and Godt 2001) while in a system composed of F-actin decorated with alexa-fluor tropomyosin (20 nM F-actin as the monomer) the protein osmotic pressure is almost not detectable. The effect of protein osmotic pressure on the critical concentration of actin is shown in Figure 10, which is obtained from Figures 2 and 3 of Tellam and al. (1983) by converting the concentration of poly(ethylene glycol) 6000 into the corresponding macromolecular osmotic pressure. The conversion is done by means of Eq. 46 (Money 1989), Π = - 2.7 x 10-4 c + 1.5 x 10-5 c2 (ΜPa)

(46)

where, c = g.L-1. The increase of macromolecular osmotic pressure lowers the critical concentration of actin (Figure 10), thus it increases the stiffness of the actin filament. With 0.4 mM MgCl2 as the polymerizing agent the critical concentration is 5.7 µM in the absence

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of poly(ethylene glycol) and ~2.4 µM at 24 kPa, the physiological protein osmotic pressure of frog skeletal muscle (Maughan and Godt 2001). At 1.6 mM MgCl2 the critical concentration decreases from 0.738 µM in the absence of poly(ethylene glycol) to 0.113 µM at 8.92 kPa. Figure 11 presents the estimated relative stiffness in the range of physiological macromolecular osmotic pressure.

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Figure 10. Effect of macromolecular osmotic pressure on the critical concentration of actin. Total Gactin, 6.4 µM; temperature, 25°C; Filled circles, 0.4 mM MgCl 2; Open diamonds, 1.11 mM MgCl2; Open triangles, 1.68 mM MgCl2. The vertical line indicates the muscle physiological macromolecular osmotic pressure, 24 kPa. Data are taken from Figures 2 and 3 of Tellam and al. (1983).

Figure 11. Relative stiffness of the actin filament as a function of macromolecular osmotic pressure. In the calculation stiffness is assumed to parallel – ΔGCc (Eq. 31). The critical concentration is expressed as mol.m-3. Relative stiffness is obtained by normalization as to the calculated value of the stiffness at ~0 osmotic pressure. The vertical line indicates the physiological macromolecular osmotic pressure, 24 kPa (Grazi and Di Bona 2006).

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4.7.3. Putative Stiffness for the Fully Non-Overlapped Thin Filament Linari and al. (1998) assigned to the non-overlapped part of thin filament (sarcomere length 2.00 – 2.15 µm) the stiffness of 58.46 pN nm-1 µm-1 in isometric contraction and 87.69 pN nm-1 µm-1 in rigor. There is, however, evidence, that the structure of the entire myosin filament (shaft included) changes on activation (Linari and al. 2000). It is thus likely that the thin filament behaves similarly and when the stiffness changes it changes concordantly for both the overlapped and the non-overlapped portions. The question thus remains about the stiffness of the fully non-overlapped thin filament (i.e. when the cross-bridges are all detached). If we assume that stiffness increases linearly with the fraction of attached crossbridges and that cross-bridge stiffness is equal both in rigor and in the isometric condition, the stiffness, x, for the fully non-overlapped thin filament is obtained by solving the equation, (58.46 – x) / 0.43 = (87.69 –x) / 1.00 The solution is, x = 36.39 pN nm-1 µm-1. What happens, however, in the more likely case that cross-bridge stiffness is not equal but is greater in rigor than in the isometric condition? To answer this question a more general expression must be utilized,

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(58.46 – x) / (0.43 / n) = (87.69 –x) / 1.00 where n = (stiffness of the isometric cross-bridge/stiffness of the rigor cross-bridge). Analysis of this expression shows that for n = 0.7, the stiffness, x, of the relaxed thin filament in vivo equals 11.91 pN nm-1 µm-1 and approaches the stiffness of the model of Adami et al. (2003), i.e. of alexa fluor tropomyosin F-actin in vitro (11.26 ± 2.4 pN nm-1 µm-1). This prediction contradicts the experimental evidence, namely that the stiffness of the actin filament increases with the protein osmotic pressure. We conclude, therefore, that ex vivo the ratio (stiffness of the isometric cross-bridge/stiffness of the rigor cross-bridge) must be lower than 1 and larger than 0.7. 4.7.4. The Upper Limit of Thin Filament Stretching The models of thin filament so far described exhibit very different stiffness, 0.38 pN nm-1 µm-1 (Adami and al. 1999) to 65 pN nm-1 µm-1 (Kojima and al. 1994), but their specific elongation at the yield point is very similar (7.1 – 9.7 nm µm-1) and is not at all related to the stiffness (Kojima and al. 1994, Cuneo and al. 1995, Adami and al. 1999, Adami and al. 2002, Adami and al. 2003, Grazi and al. 2004). It is thus likely that also ex vivo the thin filament cannot afford stretching by greater than 10 nm µm-1. Since the compliance of thin filaments in active muscle represents about 29% of the half sarcomere compliance (Linari and al. 1998) it seems safe to assume that stretching greater than 24–29 nm per half sarcomere is accompanied either by cross-bridge detachment or thin filament breaking. 4.7.5. The Energy Required to Stretch the Thin Filament The elongation of the actin filament is proportional to the force applied (Adami and al. 2003) thus the work performed on the filament while stretching is, W = stiffness.Δl2 / 2.

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Accordingly, to stretch by 10 nm a 1 µm filament of either alexa-fluor-tropomyosin Factin, stiffness 11.26 pN nm-1 (Adami and al. 2003), or a thin filament in isometric contraction, stiffness 58.46 pN nm-1 (Linari and al. 1998), requires 563 and 2923 pN.nm, respectively. Incidentally, the energy released on the hydrolysis of one ATP molecule under muscle conditions, 74.4 pN.nm (Kushmerick 1969), is only sufficient to stretch by 1.59 nm the thin filament in isometric contraction (Grazi and Di Bona 2006).

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Conclusions The cellular medium is a highly non-ideal solution where proteins compete for water. In such a concentrated solution structure and non-ideality are interconnected. The change of the structure implies a change of the non-ideality of the solution and vice versa. Reasoning on non-ideal systems requires the knowledge of activities. Determination of activities requires an osmotic approach to the system. Osmotic measurements are slow, hold at the equilibrium, apply to binary solutions and, in some cases, to ternary solutions. Nevertheless the skilled application of this technique may help to obtain a reasonable sketch of the free energy changes in vivo. The weakness of the osmotic technique is that it holds at the equilibrium so that the investigation of either a sudden reaction change or a steady state is precluded. An example of the potentiality of the method is offered by the analysis of the chemical potential changes of solutions of actin and myosin, as a function of their concentration. Even though F-actin and myosin filaments are complex structures, their free energy change under osmotic stress is easily related to the concentrations of the total monomeric species of the two proteins. The free energy change of the actomyosin complex, 1:1 as monomer, is also studied as a function of the osmotic stress. These data, obtained directly and without any limiting assumption from osmotic pressure measurements, provide the first evaluation of the chemical potential of the 1:1 actin-myosin complex under conditions similar to those of skeletal muscle. The studies show also that the formation of actomyosin influences protein osmotic pressure thus the water chemical potential. Alteration of the water chemical potential necessarily perturbs the energetic of all the protein structures eventually present in the medium (Grazi and al. 1998, Grazi and al. 2001). Osmotic measurements allow to relate the free energy of the free actin monomer to the water chemical potential. This feature, merged with the theory of Biron and al. (2006), allows to define three conditions for the F-actin solutions: a) the steady state where the concentration of the actin filaments is exponentially related to their length, b) the equilibrium where the concentration of the actin filaments is independent of their length, c) the equilibrium perturbed by an external macromolecule where the concentration of the actin filaments increases very slightly as a function of their length, with a break at the mean length. The prediction is made that, while the free energy of hydrolysis of ATP approaches zero, the total free energy of the actin filaments decreases and their average length increases, so Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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that, at the equilibrium, the total free energy of the actin filaments reaches a minimum and the concentration of the actin filaments becomes independent of their length. The prediction is also made that the free energy of the free actin monomer as well as the length distribution of the actin filaments are dictated by the macromolecular osmotic pressure, not only for a system in equilibrium but also at any steady state (Grazi and Pozzati 2008).

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References Adami, R., Choquet, D. and Grazi, E. (1999). Rhodamine phalloidin F-actin: critical concentration versus tensile strength. Eur. J. Biochem. 263, 270–275. Adami, R., Cintio, O., Trombetta, G., Choquet, D. and Grazi, E. (2002). Effects of chemical modification, tropomyosin and myosin subfragment 1 on the yield strength and critical concentration of F-actin. Biochemistry, 41, 5907–5912. Adami, R., Cintio, O., Trombetta, G., Choquet, D. and Grazi, E. (2003). On the stiffness of the natural actin filament decorated with alexa-fluor-tropomyosin. Biophys. Chem., 104, 469–476. Arakawa, T. and Timasheff, S.N. (1985). Mechanism of poly (ethylene glycol) interaction with proteins. Biochemistry, 24, 6756-6762. Arakawa, T. and Timasheff, S.N. (1985). Theory of protein solubility. Methods Enzymol., 114, 49-77. Bahler, M. and Greengard, P. (1987). Synapsin I bundles F-actin in a phosphorylationdependent manner. Nature, 326, 704-707. Barden, J.A. and Dos Remedios, C.G. (1980). Cristalline actin tubes. I Is the conformation of the lanthanides-induced actin tube monomer like F-actin than G-actin? Biochim. Biophys. Acta, 624, 163-173. Biron, D. and Moses, E. (2004). The effect of a-actinin on the length distribution of F-actin. Biophys. J., 86, 3284–3290. Biron, D., Moses, E., Borukhov, I. and Safran, S.A. (2006). Inter-filament attractions narrow the length distribution of actin filaments. Europhys. Lett. 73, 464–470. Burlacu, S., Janmey, P.A. and Borejdo, J. (1992). Distribution of actin filament lengths measured by fluorescence microscopy. Am. J. Physiol., 262, C569–C577. Cintio, O., Adami, R., Choquet, D. and Grazi, E. (2001). On the elastic properties of tetramethylrhodamine F-actin. Biophys. Chem., 92, 201–207. Cuneo, P., Magri, E., Verzola, A. and Grazi, E. (1992). Macromolecular crowding is a primary factor in the organization of the cytoskeleton. Biochem. J., 281, 507-512. Cuneo, P., Trombetta, G., Magri, E., and Grazi, E. (1995). The "in vitro motility assay" and phalloidin-F-actin. Biochem. Biophys. Res. Commun., 211, 614–618. De Rosier, D. J., and Tilney, L. G. (1982). How actin filaments pack into bundles. Cold Spring Harbor Symp. Quant. Biol., 46, 525-40. Dos Remedios, C.G. and Dickens, M. (1978). Actin micocrystals and tubes formed in the presence of gadolinium ions. Nature, 276, 731-733. Edelstein-Keshet, L. and Ermentrout, B.G. (1998). Models for the length distributions of actin filaments: I. Simple polymerization and fragmentation. Bull. Math. Biol., 60, 449–475.

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Edsall, J.T. and Wyman, J. (1958). Biophysical Chemistry, vol. I, Academic Press, New York, p. 195. Egelman, E.H. and Padron, R. (1984). X-Ray Diffraction Evidence that Actin is a 100Å Filament. Nature, 307, 56-58. Ermentrout, B.G. and Edelstein-Keshet, L. (1998). Models for the length distributions of actin filaments: II. Polymerization and fragmentation by gelsolin acting together. Bull. Math. Biol., 60, 477–503. Ferri, A. and Grazi, E. (1982). Mechano-chemical energy transduction in biological systems. The effect of mechanical stimulation on the polymerization of actin: a kinetic study. Biochem J., 205, 281-284. Frieden, C. and Goddette, D.W. (1983). Polymerization of actin and actin-like systems: evaluation of the time course of polymerization in relation to the mechanism. Biochemistry, 22, 5836 – 5843. Fujiwara, S., Takahashi, H., Tadakuma, T., Funatsu, S. and Ishiwata, S. (2002). Microscopic analysis of polymerization dynamics with individual actin filaments. Nat. New Biol., 4, 666–673. Grazi, E., Magri, E. and Pasquali-Ronchetti, I. (1982). Multiple supramolecular structures formed by interaction of actin with protamine. Biochem J., 205, 31-37. Grazi, E. (1989). An alternative pathway of actin filament elongation. The condensation of small oligomers. J. Muscle Res. Cell Motil., 19, 275-279. Grazi, E., Trombetta, G. and Guidoboni, M. (1990). Divergent effects of filamin and tropomyosin on actin filament bundling. Biochem. Biophys. Res. Communs., 167, 11091114. Grazi, E., Magri, E., Cuneo, P. and Cataldi, A. (1991). The control of cellular motility and the role of gelsolin. FEBS Letters, 295, 163-166. Grazi, E., Cuneo, P. and Cataldi, A. (1992). The control of cellular shape and motility. Mg2 and tropomyosin regulate the formation and dissociation of microfilament bundles. Biochem. J., 288, 727-732. Grazi, E., Schwienbacher, C., and Magri, E. (1993). Osmotic stress is the main determinant of the diameter of the actin filament. Biochem. Biophys. Res. Commun., 197, 1377-81. Grazi, E. (1994). Cytoskeleton, motile structures and macromolecular crowding. Adv Exp Med Biol. 358, 123-30. Grazi, E., Magri, E., Schwienbacher, C. and Trombetta, G. (1996). A model relating protein osmotic pressure to the stiffness of the cross-bridge components and the contractile force of skeletal muscle. Eur. J. Biochem., 241, 25–31. Grazi, E. (1997). Hypothesis. What is the diameter of the actin filament? FEBS Letters, 405, 249-252. Grazi, E., Cuneo, P., Magri, E., Adami, R. and Trombetta, G. (1998). Protein cross talking through osmotic work: the free energy of formation of the MgADP-myosin complexes at the muscle protein osmotic pressure. Biochim. Biophys. Acta, 1388, 419-427. Grazi, E., Adami, R., Cintio, O., Cuneo, P., Magri, E. and Trombetta, G. (2001). Dissecting the free energy of formation of the 1:1 actomyosin complex. Biophys. Chem., 89, 181191. Grazi E., Cintio, O. and Trombetta G. (2004). On the mechanics of the actin filament: the linear relationship between stiffness and yield strength allows estimation of the yield strength of thin filament in vivo. J. Muscle Res. Cell Motil., 25, 103–105.

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Grazi, E. and Di Bona, C. (2006). Protein Osmotic Pressure and Cross-Bridge Attachment Determine the Stiffness of Thin Filaments in Muscle Ex Vivo. J. Biochem., 140, 39–42. Grazi, E. and Pozzati, S. (2008). Protein osmotic pressure modulates actin filament length distribution. J. Theoret. Biol., 251, 411–420. Hanson, J. and Lowy, J. (1963). The structure of F-actin and of actin filaments isolated from muscle. J. Mol. Biol., 6, 46-58. Holmes, K., Popp, D., Gebhard, W. and Kabsh, W. (1990). Atomic model of the actin filament. Nature, 347, 44–49. Husai-Chisti, A., Levin, A. and Branton, D. (1988). Abolition of actin bundling by phosphorylation of human erythrocyte protein 4.9. Nature, 334, 718-721. Ikebuchi, N.W. and Waisman, D.M. (1990). Calcium-dependent regulation of actin filament bundling by lipocortin-85. J. Biol. Chem., 265, 3392-3400. Kawamura, M. and Maruyama, K., (1970). Electron microscopic particle length of F-actin polymerized in vitro. J. Biochem., 67, 437–457. Kiselar, J.G., Mahaffy, R., Pollard, T.D., Almo, S.C. and Chance, M.R. (2007). Visualizing Arp2/3 complex activation mediated by binding of ATP and WASp using structural mass spectrometry. Proc Natl. Acad. Sci. USA, 104, 1552–1557. Kishino, A. and Yanagida, T. (1988). Force measurements by micromanipulation of a single actin filament by glass needles. Nature, 334, 74–78. Koestler, S.A., Auinger, S., Vinzenz, M., Rottner, K. and Small, J.V. (2008). Differentially oriented populations of actin filaments generated in lamellipodia collaborate in pushing and pausing at the cell front. Nat. Cell. Biol., 10, 306–313. Kojima, H., Ishijima, A., and Yanagida, T. (1994). Direct measurement of stiffness of single actin filaments with and without tropomyosin by in vitro nanomanipulation. Proc. Natl. Acad. Sci. USA, 91, 12962–12966. Kuntz, I.D. and Kautzmann, W. (1974). Hydration of proteins. Adv. Protein Chem., 28, 239345. Kushmerick, M.J. (1969). Appendix. Free energy of ATP hydrolysis in the sarcoplasm. Proc. Roy. Soc. Lond. B, 174, 348–353. Laurent, T.C. and Ogston, A.G. (1963). The interaction between polysaccharides and other macromolecules. 4. The osmotic pressure of mixtures of serum albumin and hyaluronic acid. Biochem. J., 89, 249-253. Linari, M., Dobbie, L., Reconditi, M., Koubassova, N., Irving, M., Piazzesi, G., and Lombardi, V. (1998). The stiffness of skeletal muscle in isometric contraction and rigor: the fraction of myosin heads bound to actin. Biophys. J., 74, 2459–2473. Linari, M. Piazzesi, G., Dobbie, L., Koubassova, N., Reconditi, M., Marayanan, T., Diat, O., Irving, M., and Lombardi, V. (2000). Interference fine structure and sarcomere length dependence of the axial x-ray pattern from active single muscle fibers. Proc. Nat. Acad. Sci. USA, 97, 7226–7231. Lorenz, M., Popp, D. and Holmes, K.C. (1993). Refinement of the F-actin model against Xray fiber diffraction data by the use of a directed mutation algorithm. J. Mol. Biol., 234, 826-836. Lorenz, M., Poole, K.J.V., Popp, D., Rosenbaum, G., Holmes, K.C. (1995). An Atomic Model of the Unregulated Thin Filament Obtained by X-ray Fiber Diffraction on Oriented Actin-Tropomyosin Gels. J. Mol. Biol., 246, 108–119.

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Magri, E. Cuneo, P., Trombetta, G. and Grazi, E. (1996). The osmotic properties and free energy of formation of the actomyosin rior complexes from rabbit muscle. Eur. J. Biochem., 239, 165-171. Maughan, D.W. and Godt, R.E. (2001). Protein osmotic pressure and the state of water in frog myoplasm. Biophys. J., 80, 435–442. Milligan, R. A. and Flicker, P. F. (1987). Structural relationships of actin, myosin, and tropomyosin revealed by cryo-electron microscopy. J. Cell Biol., 105, 29-39. Money, N.P. (1989). Osmotic pressure of aqueous polyethylene glycols. Relationship between molecular weight and vapor pressure deficit. Plant Physiol., 91, 766–769. Moody, C.J., Marston, S.B. and Smith, C.W.J. (1985). Bundling of aorta calsedmon by thin filaments is not related to its regulatory function. FEBS Letters, 191, 107-112. Moore, P.B., Huxley, H.E. and De Rosier, D.J. (1970). Three-dimensional reconstruction of F-actin, thin filaments and decorated thin filaments. J. Mol. Biol., 50, 279-295. O‟Brien, E. J., Gillis, J. M., and Couch, J. (1975). Symmetry and molecular arrangement in paracrystals of reconstituted muscle thin filaments. J. Mol. Biol., 99, 461-475. Ogston, A.G. and Phelps, C.F. (1961). Exclusion of inulin from solutions of hyaluronic acid. Biochem. J., 78, 827-831. Ogston, A.G. (1962). Some thermodynamic relationships in ternary systems, with special reference to the properties of systems containing hyaluronic acid and proteins. Arch. Biochem. Biophys. Suppl., I, 39-51. Oriol-Audit, C. (1978). Polyamine induced actin polymerization. Eur. J. Biochem., 87, 371376. Parsegian, V.A., Rand, R.P., Fuller, N.L. and Rau, D.C. (1986). Osmotic stress for the direct measurement of intermolecular forces. Methods Enzymol., 127, 400 - 416. Pirani, A., Vinogradova, M.V., Curmi, P.M.G., King, W.A., Fletterick, R.J., Craig, R., Tobacman, L.S., Xu, C., Hatch, V. and Lehman, W. (2006). An Atomic Model of the Thin Filament in the Relaxed and Ca2C-Activated States. J. Mol. Biol., 357, 707–717. Pollard, T.D. and Cooper, J.A. (1986). Actin and actin-binding proteins. A critical evaluation of mechanisms and functions. Annu. Rev. Biochem., 55, 987-1035. Scatchard, G.J. (1946). Physical chemistry in protein solution. I. Derivation of the equations for the osmotic pressures. J. Amer. Chem. Soc., 68, 2315 - 2319. Schutt, C.E., Rozycki, M.D., Chik, J.K. and Lindberg, U. (1995). Structural studies on the ribbon-to-helix transition in profiling-actin crystals. Biophys. J., 68, 12s-18s. Schwienbacher, C., Magri, E., Trombetta, G. and Grazi, E. (1995). Osmotic Properties of the Calcium-Regulated Actin Filament. Biochemistry, 34, 1090-1095. Sept, D., Xu, J., Pollard, T.D. and McCammon, J.A., (1999). Annealing accounts for the length of actin filaments formed by spontaneous polymerization. Biophys. J., 77, 2911– 2919. Sobue, K., Muramoto, Y., Fujita, M. and Kakiuchi, S. (1981). Purification of a calmodulinbinding protein from chicken gizzard that interacts with F-actin. Proc. Nat. Acad. Sci. USA, 78, 5652-5655. Sobue, K., Morimoto, K., Kanda, K., Maruyama, K. and Kakiuchi, S. (1982). Econstitution of Ca2+-sensitive gelation of actin filaments with filamin, caldesmon and calmodulin. FEBS Letters, 138, 289-292.

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Spudich, J.A., Huxley, H.E. and Finch, J.T. (1972). Regulation of skeletal muscle contraction. II Structural studies of the interactions of the tropomyosin-troponin complex with actin. J. Mol. Biol., 72, 619-632. Steinmetz, M.O., Goldie, K.N. and Aebi, U. (1997). A Correlative Analysis of Actin Filament Assembly, Structure, and Dynamics. J. Cell Biol., 138, 559–574. Strzelzcka-Golassewka, H., Prochniewicz, E. and Drabikowsli, W. (1978). Interaction of actin with divalent cations. 1. The effect of various cations on the physical state of actin. Eur. J. Biochem., 88, 219-227. Suzuki, A., Yamazaki, M. and Ito, T. (1989). Osmoelastic coupling in biological structures: formation of parallel bundles of actin filaments in a crystalline-like structure caused by osmotic stress. Biochemistry, 28, 6513-6518. Tellam, R.L., Sculley, M.J., Nichol, L.W., Wills, P.R. and Michael, P.R. (1983). The influence of poly(ethylene glycol) 6000 on the properties of skeletal-muscle actin. Biochem. J., 213, 651–659. Tirion, M.M., ben-Avraham, D., Lorenz, M. and Holmes, K.C. (1995). Normal modes as refinement parameters for the F-actin model. Biophys. J., 68, 5-12. Trombetta, G., Adami, R., Cintio, O. and Grazi, E. (2002). Differential response of fast and slow myosin ATPase from skeletal muscle to F-actin and to phalloidin F-actin. Biochim. Biophys. Acta., 1569, 135–138. Trombetta, G., Di Bona, C. and Grazi, E. (2005). The transition of polymers into a network of polymers alters per se the water activity. Int. J. Biol. Macromol., 35, 15–18. Tsuda, Y., Yasutake, H., Kishino, A. and Yanagida, T. (1996). Torsional rigidity of single actin filaments and actin-actin bond breaking force under torsion measured directly by in vitro manipulation. Proc .Nat. Acad. Sci. USA, 93, 12937–12942. Wegner, A. (1976). Head to tail polymerization of actin. J. Mol. Biol., 108, 139-150. Wegner, A. and Neuhaus, J.M. (1981). Requirement of divalent cations for fast exchange of actin monomers and actin filament subunits. J. Mol. Biol., 153, 681-693. Wieland, T., de Vries, J.X., Schafer, A.J., and Faulstich, H. (1975). Spectroscopic evidence for the interaction of phalloidin with actin. FEBS Lett., 54, 73–75.

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In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 137-176

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter IV

Two Communication Bridges to One Versatile Molecule Ricardo Mondragón1 and Doris Cerecedo2 1

Departamento de Bioquímica, 4Unidad de Microscopía Electrónica, Centro de Investigación y de Estudios Avanzados (CINVESTAV) del IPN, México DF, México. 2 Laboratorio de Hematobiología, Escuela Nacional de Medicina y Homeopatía, Instituto Politécnico Nacional (IPN), México DF.

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Abstract Actin helical filaments are the key tools of the cytoskeleton for adapting cells to the physical or chemical microenvironment signals organizing cell contents, coordinating movement, or changing shape. Actin polymerization is controlled by regulatory proteins including nucleation, depolymerizing and severing factors, capping proteins, polymerases, crosslinkers, and stabilizing proteins. The cell‟s exquisite sensitivity in responding to a wide range of physical or chemical stimuli, is translated into cytoskeleton reorganization and adhesion-site modulation. Living cells survive, proliferate, or differentiate while they are anchored to their extracellular matrix. It comprises a complex bulk of information integrated into a coherent environmental signal. This is achieved through integrins that are the major family of transmembrane adhesion receptors composed of α and β units. These heterodimers not only play an anchorage mechanical role they also transmit chemical signals into the cell concerning their microenvironment and adhesive state. Integrin-based interaction networks follow an ordered series of events that range from their activation to focal adhesion assembly involving the participation of actin and actin binding proteins. Activated integrins link directly to the signalling and cytoskeletal



Correspondence to: Dr Doris A. Cerecedo Mercado, Laboratorio de Hematobiologıía, Escuela Nacional de Medicina y Homeopatía, IPN, Guillermo Massieu Helguera no. 239, Col. La Escalera Ticomán, 07320 México, DF, México. E-mail: [email protected]

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Ricardo Mondragón and Doris Cerecedo systems regulating multiple cellular features, such as cell anchoring, locomotion, substrate deformation and matrix remodeling, in which actin possesses an essential role. Integration of incoming signals and whether to respond to these depends not only on integrin properties; an additional glycoprotein system named dystroglycan has demonstrated specific binding patterns to certain extracellular matrix components, supporting cell adhesion and translating signals. Dystroglycan was identified as Dystrophin glycoprotein complex (DGC) component; it comprises α- and β subunits. Alfa-dystroglycan is located at the extracellular peripheral membrane interacting with extracellular matrix proteins, while β-dystroglycan binds to α-dystroglycan on the extracellular face, and on its intracellular face to F-actin. Dystroglycan potential adhesion is due to its privileged position between cytoskeleton and extracellular matrix combined with their differential glycosylation patterns, tissue-specific expression, and multiple potential interactions. In this regard, dystroglycan has been found as a component of podosome adhesion structures, as well as in focal adhesions, interacting with vinexin, a vinculin binding partner. In this chapter, we focus on the relationship between the two specific transmembrane proteins that link the extracellular matrix and connect with F-actin to develop microenvironment-triggered responses, particularly regarding the focal adhesions, stress fibers, podosomes, and filopodia in which integrins and dystroglycans are involved.

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Integrins Integrins are a superfamily of heterodimeric cell-surface receptors that are found in a broad range of animal species [1]. As their name implies, integrins integrate the cell cytoskeleton with adhesion points of extracellular matrix ligands and cell-surface ligands in order to mediate essential cellular processes such as cell-cell and cell-extracellular matrix interactions, polarization in response to extracellular cues, cell migration, differentiation, and survival, and even in cell-pathogen interactions [2]. Integrins also participate in the pathological properties of many diseases such as neoplasia, tumor metastasis, immune dysfunction, ischemia injury, viral infections, osteoporosis, and coagulopathies [3,4]. Integrin-dependent physiological processes include tissue morphogenesis, inflammation, wound healing, and regulation of cell growth and differentiation [5]. In general, integrins are dimers that are approximately 280 Å in length and consist of one  (150-180-kD) and one β (90-kD) subunits held together by non-covalent bounds [6,7]. Both integrin subunits are type I one transmembrane pass proteins with a large N-terminal extracellular domain of 700-1,100 residues inserted into the lipid membrane through a stoptransfer anchor sequence followed by a short 30-50-residue cytoplasmic domain. Like other receptors, integrins transmit signals to the cell‟s interior (so-called “outsidein” signaling in an event associated with conformational changes in the integrin extracellular segment), which regulate cytoskeleton organization, activate kinase signaling cascades, and modulate the cell cycle and gene expression; therefore, integrins attest the activation state of the cell (reviewed in [8]). These changes vary with cell type and the state and nature of the ligand and are modulated by the divalent cations that are also required for integrin ligand interaction [9-11]. In vertebrates, 19 different integrin  subunits and 8 different β subunits have been reported, forming by mutual combination about 25 β heterodimers [5] (Figure 1). Integrin α and β subunits are totally distinct, with no detectable homology between them; sequence

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identity among α subunits is about 30% and among β subunits, 45%, indicating that both the α and the β gene families evolved by gene duplication [7].

Figure 1. Integrin  and β subunits from 24 heterodimers that differ in ligand recognition and inside-out and outside-in signaling. Integrins with I domain in a subunit are indicated with asterisks. (Modified from [7]).

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Classification of Integrins The majority of combinations of /β subunits can be organized into three fundamental groups based on the subunit type (β1, β2, and β3, or v chains, on the extracellular matrix protein type recognized, or on the specific adhesion motifs [12] (Table 1). β1 integrins form the first and largest group of integrins and are ubiquitously distributed in nucleated cells as well as in platelets. With the exception of the 4β1 complex, β1 integrins are generally unrestricted in their expression to any given cell lineage. β1 integrin receptors generally mediate the adhesion of mesenchymal and epithelial cells to extracellular matrix proteins. The 4β1 integrin possesses the unique characteristic of mediating cell-cell and cellmatrix binding. It is expressed in bone marrow-derived cells (except for neutrophils), in certain tumor cells, and in muscle development. The 4β1 integrin mediates binding to Vascular cell adhesion molecule 1 (VCAM-1) and fibronectin (CS-1 region) by an Arginine-GlycineAspartic acid (RGD)-independent mechanism. A second major group of integrins shares either the β3 or the v subunit (Table 1). Receptors such as IIBβ3, also known as platelet glycoprotein IIb/IIIa, and vβ3 recognize different ligands from a broad of cell and tissue sources. Integrins with the v subunit may form dimers with at least five different β chains, including the β1 chain. Subunits v and β3 recognize RGD domains present in extracellular matrix proteins.

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Table 1. Classification of integrins according to ligand motifs and distribution

Abbreviations: BC = B cells; Col = Collagen; EC = endothelial cells; Eos = Eosinophils; EPC = Epithelial cells; Fb = Fibroblasts; Fib = Fibrinogen; Fn = Fibronectin; iC3b = inactivated component of complement; Lm = Laminin, LGL = Large granular lymphocytes; Macros = Macrophages; Mega = Megakaryocytes; Monos = Monocytes; OPN = Osteopontin; Plt = Platelets; PMN = Neutrophils; SMC = Smooth muscle cells; TC = T cells; TSP = Thrombospondin; Vn = Vitronectin; vWf = von Willebrand disease. (Modified from [12]).

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The third group of integrins shares the β2 integrin chain, whose expression is restricted to leukocytes [13] (Table 1). Receptors such as 4β2, also known as the LFA-1 integrin, determine the capability of leukocytes in the transmigration of the endothelial epithelium and recognize members of the Intercellular adhesion molecule (ICAM) family of adhesion proteins. In contrast, the expression of Mβ2 is restricted to monocytes, macrophages, and granulocytes; it recognizes fibrinogen and inactivated C3b, playing an important role in the phagocytosis of opsonized particles and bacteria [14]. The fourth group of integrins includes three integrins (6β4, 4β7, and Eβ7) with different subunits in comparison with the three previous groups (Table 1). These integrins recognize components of the extracellular matrix as well as adhesion molecules of the Immunoglobulin superfamily (IgSF), Mucosal addressin cell adhesion molecule (MAdCAM) and Vascular cell adhesion molecule 1 (VCAM-1), and intercellular-junction adhesion molecules of epithelial cells such as E-cadherin. These have a diverse distribution including endothelial and epithelial cells, T cells, and Schwann cells. The binding of T-cells expressing the Eβ7 integrin to E-cadherin may represent a homing mechanism to maintain lymphocytes within their respective epithelial tissues.

Integrin Structure The general structure of integrins has been analyzed and determined by electron microscopy and analysis of protein crystals. Integrins consist of multiple structural domains with different properties (Figure 2).

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Properties of Integrin Domains Integrins contain extracellular domains of >940 and >640 residues, respectively, that span the plasma membrane and that in general have very short cytoplasmic domains of about 40– 70 amino acids, with the exception of the β4 subunit, which possesses a cytoplasmic domain of 1,088 amino acids that is specialized to connect with the keratin cytoskeleton [16]. Apparent molecular weights (kDa) of  and β subunits vary according to the respective isoform. For the  subunit (in kDa), these are 1 (210), 2 (165), 3 (130), 4 (150), 5 (135), and 6 (120), and for the β subunit (kDa), these are β1 (130), β2 (95), β3 (105), β4 (220), and β5 (110) (Apparent molecular weights (kDa) by SDS-Page. Millipore Technical Library; www.millipore.com/tech publications). The extracellular portion of integrin heterodimers consists of the following multiple extracellular domains: (see Figure 1): Alpha subunit has the following domains: a) the βpropeller domain constituted of seven amino acid repeats forming β-sheets arranged in a circle that contains the place to bind the ligand and a putative Mg2+ ion [1,5,17]; b) the I domain is a 200 amino acid region that is located between the 2-3 repeats of β-sheets of the propeller domain (Figure 1). It represents the major ligand-binding site and contains a divalent cation coordination site denominated as the Metal ion-dependent adhesion site (MIDAS), which binds negatively charged residues in ligands [17-19]; c) the tight domain and Calf-1 and 2 domains are β sandwich domains that contribute to interdomain rotation

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movement around the contact area and additionally contribute to providing rigidity to the structural entity [6].

Figure 2. Integrin domains. (A) Ribbon diagram of the straightened extracellular segment of the Vβ3 integrin within the 3-dimensional crystal structure of the integrin. Amino acid domain boundaries are indicated in parenthesis. (B) Organization of domains within the primary structure of Lβ2. (Modified from [5,15]).

The β subunit contains the following: a) the PSI domain (Plexin-semaphorin-integrin) at the N-terminal end is characterized as having seven cysteines in two  helices that form longrange intrachain disulfide bonds with the Epidermal growth factor (EGF) cysteine-rich domain [5,20]; b) the I-like domain is a highly conserved domain of about 240 residues with a putative metal binding (DXSXS) sequence motif similar to MIDAS. The I-like domain binds directly to ligand in integrins that lack the I domain and to indirectly regulate ligand binding by integrins that contain I domains [6]; c) two hybrid domains flanking I-like domain; d) four EGF-like repeats rich in cysteine residues that is important for signal transduction [5,20], and e) β tail domain. The headpiece, which contains the ligand-binding site, is formed by the interactions of the β-propeller domain of the α subunit with the PSI domain, the β I-like or βA domain, and the hybrid domain of the β subunit. Integrins with αV, α8, α5, and αIIb subunits

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make up a family of non-I domain α subunits that recognizes the RGD motif; in addition, integrins with α3, α6, and α7 subunits are classified as the laminin-binding family [15,21]. Integrin Vβ3 (CD51/CD61), whose structure has been resolved, is one of the most promiscuous integrins, is an important receptor in tumor angiogenesis and metastasis, inflammation, and bone resorption, and was previously described because it binds to multiple ligands including vitronectin, angiostatin, and osteopontin and also serves as a receptor for several viruses such as foot-and-mouth disease virus, adenovirus, and Human immunodeficiency virus (HIV) [6,22]. The Vβ3 integrin can take two types of conformations: a bent conformation that is inactive, and an extended conformation that is commonly observed in electron micrographs that represents the active conformation [6]. During activation, the integrin dimer in the bent conformation or inactive state is subjected to upward movement of the headpiece that contains the β propeller-I domain and tight domains both in β subunit and I-like and hybrid domains from the β domain, exposing the ligandbinding domains [5].

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Recognition of RGD Sequences by Integrins The RGD sequence (Arginine-Glycine-Aspartic acid) is the cell attachment site of a large number of adhesive extracellular matrix, blood, and cell-surface proteins, and nearly one half of the >25 known integrins recognize this sequence in their adhesion protein ligands. The RGD was initially identified as a cell-adhesion sequence in fibronectin and was then recognized as a broad sequence distributed in a variety of proteins involved in cell attachment and suggesting that the RGD sequence plays a central role in cell adhesion biology as the prototype adhesion signal [23]. Each of the  and β subunits contributes to ligand specificity and contains potential binding sites for it through the RGD motif. It is believed that residues outside the RGD motif contribute in specificity and affinity during ligand recognition. These „secondary‟ sites are generally assumed to interact directly with the α subunit of the integrin, whereas the RGD motif binds primarily to the β subunit [24]. At least eight different integrins recognize the RGD sequence in their ligands through a domain located at the β subunit (see Table 2). All  integrin subunits (5, 8, v, and IIb) show greater sequence similarity to each other than to the other  subunits [25]. A partial list of adhesion proteins with RGD sites includes fibronectin, vitronectin, fibrinogen, von Willebrand factor, thrombospondin, laminin, entactin, tenascin, osteopontin, bone sialoprotein, and, under some conditions, collagen. There are a variety of proteins from pathogenic bacteria (Escherichia coli, Bordetella pertussis, Yersinia pseudotuberculosis) and viruses (Coxsackie-type virus, Human immunodeficiency virus [HIV]) that contain the RGD sequence and that can be recognized by host cell integrins as a initial step in adhesion or invasion [23]. By assays of cross-linking between RGD peptides and the integrins and site-directed mutagenesis, a ligand-binding site has been localized in the first 200 amino acid residues of the β subunit N-terminus of the IIbβ3 integrin [26,27] and in the β3 domain of vβ3 integrins. In both cases, the ligand-binding site is located near a binding site for divalent cations. The  subunit also contains one or more ligand-binding sites for the RGD sequence within divalent cation-binding sequences [28].

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Table 2. Integrins classified according to recognition of the RGD sequence (arginine-glycine-aspartic acid). (Modified from [23])

The recognition site for collagen binding by the 2β1 integrin and for fibrinogen by the Mβ2 integrin is located in the I domain, which is an insertion into the divalent cationbinding region of the  subunit present in some integrins [19]. Slight variations in the RGD sequence can preserve the binding properties of several integrins [23]. Peptides containing the KGD (Lys-Gly-Asp) sequence can bind to the IIbβ3 integrin [29]. The glycine position can be occupied by a number of different individual amino acid residues, or even by two residues, in proteins, with retention of integrin-binding activity [30]. Recognition properties of peptides containing the RGD sequence by integrins and their inhibitory effect in adhesion, cell migration, and cell proliferation has been considered a promising alternative for prevention of cell attachment as a treatment strategy for different diseases [31]. Inhibition of octeoclast adhesion to vβ3 during osteoporosis would prevent bone degradation during osteoporosis [32]. Other possibilities include inhibition of angiogenesis by inhibiting the endothelial cell adhesion that interferes in the growth and spread of tumor cells [33].

Integrin-Associated Proteins In different cell processes, integrins become associated with supramolecular complexes constituted of different integral or peripheral membrane proteins that allow access to signaling cascades with which integrins do not interact outside the complexes, increasing in

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this manner the possibility for cell activation. There are examples of integrin-associated membrane proteases, growth factor receptors, immunologic receptors, transporters, and channels that have been found employing co-immunoprecipitation [34] (Table 2). Some supramolecular complexes include Matrix metalloproteinases (MMPs) which are initially secreted and then associated with cell-surface integrins [35]. The integrin-MMP association induces a degradation of Extracellular matrix (ECM) components and promotes adhesion and cell migration. MMP1 associated with divalent cation binds to the I domain of the 2β1 integrin [36]. MMP2 interaction with Vβ3 requires the carboxy-terminal hemoPexin-like (PEX) domain of the protease [37]. Blockade of association between MMP2 and the integrin inhibits angiogenesis in vivo, indicating that the physical association of protease and integrins is required for endothelial proliferation or migration, although the mechanism is unknown [38]. Another protein that can bind integrins is the P2Y2 nucleotide receptor, which contains an RGD sequence in the extracellular portion of the protein that permits its association with Vβ3/β5 integrins, enhancing the signaling role of G-protein-coupled receptor (GPCR) and affecting the recognition of EMC ligands by the integrin [39]. A subset of integrins 3β1 and 6β1 associates with proteins of the Transmembrane 4 superfamily (TM4SE, or tetraspans) through the  chain [61] Several TM4SF cell-surface proteins (including CD9, CD63, CD81, CD82, and CD151) may contribute significantly to adhesion, cell motility, and tumor cell metastasis probably through the recruitment of signaling enzymes such as PI 4-kinase and Protein kinase C (PKC) into complexes with integrins [40,41]. Binding of uPAR and MMPs with integrins does not prevent the binding of the ECM ligands of integrin, but under particular conditions, this appears to enhance integrin ligand binding. Other GPI-anchored proteins include CD14, and FcRIIIB complexes with integrins [40-44]. It has been hypothesized that 3β1-CD151-PI4-kinase complexes should provide localized production of Ptdlns-4-P, and subsequently Ptdlns-4, 5-P. Both lipid derivatives can then be substrates for PI 3-kinase, as well as regulators of the actin cytoskeleton. Thus, via these TM4SF links, PKC and PKCβ can be recruited into complexes with integrins 3β1, 4β1, and 6β1 [61], producing the phosphorylation of 3 and 6 cytoplasmic tails by PKC. Certain integrins bind immunoglobulin superfamily proteins. The Vβ3 and llbβ3 integrins associate with extracellular domains of CD47, an Integrin associated protein (IAP) belonging to the Ig superfamily of proteins [40]. Several functions of Vβ3-dependent neutrophils can be affected by exposure to anti-CD47 antibodies and by deletion of the CD47 gene [45]. The extracellular Ig domain of CD47 is also necessary for cell binding to vitronectin and for acting as a receptor for thrombospondin activating signaling through a heterotrimeric G protein pathway for presentation of a chemotactic response [46,47]. CD147 or Extracellular metalloproteinase inducer (EMMPRIN) or basigin is another Ig superfamily protein that associates with the  subunits of 3β1 and 6β1 integrins [40]. CD147 participates in the regulation of matrix metalloproteinase production and in cell migration through 3β1-dependent cell migration. Some Glycophosphatidylinositol (GPI) anchor-linked proteins can associate with specific β1, β2, and β3 integrins (Table 3), as occurs in the interaction between the β-propeller domain and the associated protein Urokinase-type plasminogen activator receptor (UPAR). Such an interaction is an essential step in cell migration, tumor invasion, and host defense [58, 59].

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Table 3. Membrane proteins that interact directly with integrins. (Modified from [52, 61])

The 3βl integrin also strongly associates with other transmembrane proteins such as CD46, which is widely expressed in normal human tissues and that acts as a co-factor for Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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factor I, a serum protease that inactivates cell-bound C3b/C4b complement proteins, thus protecting cells from complement-mediated damage [40]. The membrane protein caveolin can be co-immunoprecipitated with both β1 and β2 integrins [40], but this interaction also has not been shown to be direct. Nevertheless, this may be an important interaction, considering that caveolin may recruit signaling molecules and non-receptor tyrosine kinases (e.g., src and fyn) into complexes with integrins. Upon cell stimulation with insulin or Platelet-derived growth factor (PDGF), a highly phosphorylated and activated subfraction of the receptors for these molecules can be co-immunoprecipitated with the Vβ3 integrin. These complexes contain multiple components and it is not yet clear which of these may be directly bound to the integrin.

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Membrane Rafts in Supramolecular Complex Formation and Function Membrane rafts are thought to be regions of the plasma membrane with more highly ordered lipids that are rich in cholesterol and sphingolipids and that are sites of concentration of a variety of cytoplasmic membrane-associated signaling molecules [48]. Some integrincontaining supramolecular complexes may be concentrated in membrane rafts, including those with several GPI-linked protein partners (uPAR, FcγRIIIB, and CD14), tetraspanins, CD98, and CD47 [40]. TM4SF complexes with α3β1 have been shown to form and to associate with at least one signaling molecule (phosphatidylinositol 4-kinase) independent of membrane rafts. Insulin-like growth factor-1 treatment of smooth muscle cells decreases CD47 association with rafts. This change in distribution is associated both with an increase in its co-precipitation with αVβ3, with increased avidity of αVβ3 for vitronectin, and with the ability of an anti-CD47 monoclonal antibody to block cell migration [49]. Thus, association of integrins with rafts can influence supramolecular complex formation. These examples raise the possibility that there may be significant cellular regulation of integrin-containing membrane complexes.

Integrin Function and Signaling Numerous signaling pathways have been described as activated by interactions of cells with matrix proteins via integrin receptors, which ultimately promote not only cell binding but also lead to cell spreading [50,51]. Both phenomena are critical for cell survival because cells that adhere, but cannot spread, may undergo programmed cell death [52,53]. Despite this high degree of redundancy, the majority of integrins appear to possess specific biological functions, raising the possibility of signaling differences among integrins. Integrins not only bind ligands present in the extracellular matrix; certain integrins can also bind to soluble ligands such as fibrinogen, or to counter-receptors such as Intracellular adhesion molecules (ICAMs) on adjacent cells. The capacity of many integrins to bind ligand is regulated by cellular signaling mechanisms through a process frequently called integrin “activation” or “inside out signal transduction”, resulting in re-organization of the cytoskeleton, motility, migration

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differentiation, and gene expression, among others. Activation of integrin receptors triggers signaling pathways involved in (1) cytoskeletal organization, (2) cell proliferation, and (3) cell survival [50]. After cell adhesion to ECM substrates by recognition of integrin receptors, integrinligand membrane clustering occurs with the formation of actin stress fiber-rich focal adhesions and regulation of cell adhesion, changes in the cell shape during spreading, and locomotion [54]. A diverse number of structural and signaling proteins (integrins, cytoskeletal proteins, actin binding proteins such as -actinin, talin, tensin, paxillin, vinculin, and tensin, and protein kinases such as Focal adhesion kinase [FAK], c-src, protein kinase C, and Integrin-linked kinase [ILK]) become concentrated at focal adhesion sites [40,53,55-57] (Table 4) (Figure 3). The binding of β1 and v integrins to their ligand in the formation of focal adhesions activates tyrosine phosphorylation of FAK, a non-receptor tyrosine kinase [58,59], followed by a cascade of phosphorylation events on target proteins [50].

Figure 3. Cytoskeleton proteins associated with integrins in focal adhesion sites. (Modified from [73]).

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Table 4. Cytoskeleton proteins and intracellular proteins associated with integrins. (Modified from [61])

FAK interacts with cytoskeleton proteins talin, vinculin, and paxilin [60,61]. Talin is required for integrin αIIbβ3 activation downstream of a number of physiologically relevant signaling pathways; its loss results in a failure of integrin activation [62]. Talin has an Nterminal 47-kDa globular head domain and a C-terminal 190-kDa rod domain that binds to the cytoplasmic tails of integrins β1A, β1D, β2, and β3 through the Ezrin, Radixin, Moesin homology (FERM) domain located in the head domain. Phosphatidylinositol 4,5bisphosphate (PIP2) promotes talin binding to the integrin β1 tail associated with a conformational change in talin that is thought to expose the integrin binding site [63]. Phosphorylation of serine and threonine residues in talin comprises one of the mechanisms of regulation of binding to integrin β tails. The clustering of integrin-ligand complexes results in oligomerization of FAK and its activation by trans-autophosphorylation [50]. FAK is linked with a number of intracellular signaling pathways. Initial recruitment of FAK to activated integrins is indirect, mediated by talin, and followed by a conformational change of FAK and its interaction through its amino terminal domain with the integrin‟s β subunit tail. Autophosphorylation of FAK at residue 397 results in the binding and activation of Src and Fyn, members of the Src family of kinases, which then phosphorylate a number of FAK-associated proteins including paxilin, tensin, and p130 CAS [64,65]. Src can also phosphorylate FAK at tyrosine residue 925, creating a binding site for the Grb2-mSOS complex [66]. In addition, autophosphorylated FAK can bind to and activate the Phosphatidyl-inositide-3 kinase (PI-3K) [67]. Integrin-linked kinase (ILK) also appears to regulate cell migration properties; therefore, during integrin-mediated cell adhesion, the activity of ILK, which interacts with the β1 subunit of the integrin, is inhibited [68].

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Integrin-ligand clustering activates an important group of proteins known as Rho proteins (RAS-related GTP binding proteins), with consecutive assembly of cortical actin as stress fibers in focal adhesions (Figure 4). The Rho family is made up of Rho A, B, C, D, and E, in addition to Racs 1 and 2 and Rac E, and Cdc42, Rho G, and TC10 proteins [69,70]. Rho family members are subjected to sequential activation: Cdc42 activates Rac, and Rac activates Rho. Rac is directly responsible for membrane ruffling and lamellipodia extension, and Cdc42 uniquely controls filopodia formation [69,71,72]. Integrins activate Rho through the production of Phosphatidyl inositol biphosphate (PIP2) possibly through its interaction with a type I isoform of Phosphatidyl inositol 4-phosphate 5kinase (PIP4-5K) [73,74].

Figure 4. Role of Rho and Ras family GTPases in focal adhesion sites mediated by integrin recognition. (Modified from [50]).

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Due to that profilin and gelsolin are modulated by PIP2, it is possible that the increase of PIP2 by Rho and its enrichment in focal adhesion sites contribute to the formation of the adhesion plaques [73]. The association of PIP2 with vinculin induces a conformational change in vinculin, allowing it to interact with talin [75]. The involvement of phosphoinositide kinase PIP4-5K, a serine/threonine kinase that is associated with protein kinase (ROCK) and protein kinase N have been involved in the activation of the Rho A protein as well as p190, a Rho GTPase-activating protein (GAP) that regulate integrin-clustering [57,76]. PI-3 kinase, which phosphorylates PI(4) phosphate (PIP) or PI(4,5) biphosphate (PIP2) to generate PI (3,4)P2 or PI (3,4,5)P3, respectively, has been shown to associate with integrin-associated focal adhesion complexes [67]. Rho can also activate a serine/threonine protein kinase known as Rho kinase, which plays an important role in regulation of the assembly of focal adhesions by phosphorylation of the myosin light-chain phosphatase, thereby suppressing the activity of the enzyme [77]. Phosphorylation of the myosin light chain induces a conformational change in myosin, increasing its binding to actin filaments and promoting actomyosin contractility and the formation of focal complexes [78]. In addition to Rho family members, R-Ras, a GTP-binding protein, has been shown to activate integrins during the formation of focal adhesion plaques. In some way, R-Ras converts integrins from a low-affinity to a high-affinity condition [79]. Certain integrins such as laminin receptor 6β4, lamin/collagen receptor 1β1, fibronectin receptor 5β1, and vitronectin receptor vβ3 are linked with the RAS-ERK signaling pathway by the adaptor protein Shc, an SH2 and Phosphotyrosine binding (PTB)-domain adaptor protein that links tyrosine-phosphorylated signal transducers with Ras. A detailed review of the signal transductions involved in integrin receptors was reported by Kumar [50]. Activation of the RAS-ERK pathway by integrin ligation has been shown to require an intact cytoskeleton, suggesting that integrin-dependent cytoskeletal complexes play a key role in activation of the MAP kinase pathway. Integrins and growth factors appear to synergize or directly stimulate the RAS-MAP kinase pathway, which in turn activates a number of transcription factors such as SRF and c-Myc, which are involved in regulating growth and differentiation [50]. During cell migration, integrins activate ERKs, and these phosphorylate the Myosin light-chain kinase (MLCK), resulting in its activation and consecutive phosphorylation of Myosin light chains (MLC) as an initial step in the association with actin filaments to induce motility [80]. Adhesion to substrates influences survival for many specialized cell types [50]. Endothelial and epithelial cells, when detached from substrate and are displaced from their natural environment, undergo apoptosis through a mechanism named as “anoikis”, from the Greek for homelessness [81-84]. Cell adhesion to extracellular matrix by integrins promotes cell survival mediated by Shc [50] (Figure 5). Phosphoinositide 3-Kinase and PKB/Akt also participate in anoikis [85]. PI-3 kinase lipid products provide protective signal acting through PKB/Akt, which blocks entry into apoptosis. When epithelial cells are detached from the matrix, PI-3 kinase and PKB/Akt become inactive and an apoptotic pathway is initiated. RAS activation leads to the constitutive activation of PI-3 kinase and then to the PKB/Akt pathway through FAK in order to favor cell survival. Alternatively, because PI3-K is a major target effector of RAS [86], activation of PI3-K by integrins could be mediated by the ability of Shc to activate RAS.

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Figure 5. Signaling pathways activated by recognition of integrins and involved in cell survival and proliferation. (Modified from [50]).

Figure 5 summarizes the integrin-mediated signaling pathways leading to cell proliferation and cell survival. A possible physiological significance of anoikis is that it may prevent cells from attaching themselves to inappropriate places in the body. Interestingly, evidences have shown that cell spreading following cell attachment by means of integrin receptors activates an anti-apoptotic mechanism, such as a raise in expression of the anti-apoptotic protein Bcl-2 and the blocking of anoikis [50]. In addition,

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integrin type could determine cell survival; thus, vascular endothelial cells appear to depend on vβ3 receptor for their survival [33]. Knowledge of the basis of the role of integrins in adhesion and cell activation has been recently used in the understanding of the highly dynamic properties of migrating malignant cells during metastasis. Characterization of the molecular basis in ligand recognition by integrins will provide the necessary tools for the design of pharmaceutical strategies focused on inhibiting the dynamic and vital properties of cancer cells [87]. The previously mentioned information refers to a direct association of integrins with several transmembrane proteins. According to confocal Fluorescence resonance energy transfer (FRET) and immunoprecipitation assays performed in adhered platelets, a feasible association has been suggested between β-1-integrin- and -DG-modulating focal adhesions [88]. Dystroglycan (DG) is a ubiquitously expressed cell adhesion protein present in focal adhesions in non-muscle cells including platelets [89,90]. In myoblasts, DG is recruited to adhesion structures interacting directly on vinculin and mediating cell adhesion and spreading. In addition, cellular adhesion to fibrinectin, agrin, or laminin 2 triggers the phosphorylation of a tyrosine at the DG cytodomain [91] as well as binding of the SH3 domain of Grb2 involved in actin-cytoskeleton reorganization [92] The molecular relationship between β1 integrins and DG can involve relevant cell roles during substrate adhesion; thus, additional studies will be required in order to clarify the precise contribution of such an association in cell adhesion and the remodelling of actin cytoskeleton.

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Identification and Cloning of DG Dystroglycan (DG) was first named cranin after its discovery as a novel laminin-binding protein at the plasma membrane of diverse cell types [93]; in 1992, it was first cloned and characterized. In mammals, the dag1 gene encodes for a 895 amino acid polypeptide precursor, which undergoes post-translational proteolytic cleavage resulting in a protein complex composed of two subunits, namely α and β [94]. α-DG is extensively decorated with multiple and heterogeneous N- and O-linked sugar chains [8], and the central mucin-like central region is particularly important for establishing a network of interactions with extracellular proteins such as laminins, agrin, perlecan, neurexin, and biglycan [53,95-98]. Its C-terminal domain interacts non-covalently with the Nterminal extracellular domain of the β-subunit. The β-DG subunit is a type I transmembrane protein subjected to some N-linked glycosylation binding to the C-terminal domain of -DG on the extracellular side, and on its intracellular side, it anchors either directly to actin or through an actin-binding protein with its cytosolic domain. This same domain also binds other proteins, such as rapsyn, caveolin-3, and Grb2 [99,100]. DG belongs to the Dystrophin-glycoprotein complex (DGC), where it plays a central role together with sarcoglycans, dystrobrevins, syntrophins, and sarcospan (reviewed in [101]).This glycocomplex includes other peripheral members or associated proteins, such as nitric oxide synthase and caveolin-3 [102]; via its cytodomain, β-DG is associated with rapsyn and the Ras/MAPK signaling pathway through the adapter protein Grb2 [103-105] and entangling the innumerable interactions that comprise the basement

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membrane, cross the plasma membrane, and that are eventually translated through the cell cytoskeleton [106] (Figure 6).

Figure 6. A representation of the central role of dystroglycan (subunits alpha and beta) and their association with sarcoglycans, dystrobrevins, syntrophins, and sarcospan, as well as other peripheral members such as nitric oxide synthase and caveolin-3.

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DG Subdomains Alpha and β-DG subunits are organized into subdomains, representing autonomously folding units. α-DG is constituted of two domains (N- and C-terminal) separated by an elongated, mucin-like region rich in prolines, serines, and threonines and that is highly Oglycosylated [107], while β-DG is composed of an N-terminal extracellular domain, a transmembrane region, and a cytoplasmatic, proline-rich C-terminal domain (Figure 2). The biochemical dissection of DG domains based on its expression as a recombinant protein have allowed to identify subdomains [108]. For α-DG, the solved crystallographic structure of its N-terminal domain shows the presence of two subdomains, the first an Ig-like domain and the second, similar to ribosomal protein S6, both connected by a flexible loop [109]. In this regard, the β-DG extracellular domain is capable of binding the C-terminal domain of α-DG, and although it belongs to the group of the “natively unfolded proteins” [108], a computational model suggests the presence of at least some secondary structure, which is in part the β-DG ectodomain [110]. No structural data are available on the cytosolic domain of β-DG, but its high content of proline residues points to a rather disordered conformation. It has been established that DG domains are particularly important for maintaining the multiple interactions that are translated into several tissue functions. The link between the extracellular matrix and the cytoskeleton [106] is important for initiating signaling, and scaffolding roles are suggested to occur through the cytoplasmic tail of dystroglycan [104,105,111-113]. The binding of extracellular ligands by α-DG is dependent on post-

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translational processing [114-116]. Although primary sequence of α-DG predicts a molecular mass of 67 kDa, the molecular weight observed in skeletal muscle is 156 kDa due to the posttranslational addition of glycans [93,94]. Both N- and O-linked sugar moieties are present in α-DG, but chemical removal of Nlinked sugars reduces the molecular mass of the protein by only 4 kDa [99,117]. Thus, the greater-than-predicted molecular weight is largely due to the addition of O-linked sugars. Disruption of N-linked glycosylation exerts no effect on the ability of dystroglycan to bind laminin, while disruption of O-linked glycosylation reduces its binding to ligand [106]. Therefore, the ligand-binding affinity of dystroglycan is dependent on the post-translational addition of glycans to α-DG [104], suggesting that the diversity of the DG function may be attributable to cellular differences in glycosylation and to the diversity of dystroglycan ligands (Figure 7). The N-terminal actin-binding regions of utrophin and dystrophin act as simple anchors to the sub-membranous F-actin cytoskeleton in the cell, providing the intracellular tether for the dystroglycan complex. The C-terminus of dystrophin or utrophin interacts with the cytoplasmic domain of β-DG via necessary, but not sufficient, WW domain-mediated interaction with the WW motif PPxY in β-DG [118-120].

Figure 7. Post-translational process of DG propeptide: (a) the product of the DAG1 gene of the 895 amino acid polypeptide has Signal sequence (SS), a mucine-like region, a Transmembrane region (TM), and positions of WW-domain interaction motifs PPxY. (b) the propetide is post-translationally cleaved by an unknown protease at residue 653 (P) to yield an -DG subunit at the extracellular surface, and a -DG subunit at the plasma membrane. (Modified from [129]).

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An elegant structural work [121] provided insight into the specificity of dystrophindystroglycan WW-domain interaction, which is a ~30 amino acid motif containing two conserved tryptophan residues that binds to proline-rich sequences in a manner analogous to that of SH3 domains [122]. The presence of the dystrophin or utrophin EF hand-like region is required to stabilize the WW domain, and both stabilize and orient the unstructured Cterminus of β-DG [121]. The potential connections made by α-DG on the extracellular surface with agrin, perlecan, and several laminin isoforms take place via an LG module-dependent mechanism that is calcium-dependent, and dystroglycan–laminin-α1 interaction (but not that of laminin α2) can be inhibited by heparin, because the positively charged surface of the LG module would effectively inhibit interaction with α-DG [106] and requires novel O-mannosyl-type [8] or sialic-acid [123] oligosaccharide moieties, whereas biglycan does not appear to require carbohydrate moieties on α-DG for binding [124]. The key linkages mediated by the dystroglycan complex between the actin cytoskeleton and the extracellular matrix (Figures 6) via utrophin or dystrophin are crucial for dystroglycan complex integrity. Mutations disrupt dystrophin-actin interaction and underlie a reduction in dystrophin-dystroglycan interaction, leading to a severe Becker muscular dystrophy phenotype [125]. In addition, disruption of dystroglycan-dystrophin interaction leads to a Duchenne muscular dystrophy phenotype, and mutations disrupting laminin-dystroglycan interaction lead to congenital muscular dystrophy [126]. Total deletion of DG in mouse leads to embryonic lethality [127], while a deletion in mouse muscle lead to muscular dystrophy [128].

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β-DG as an Outside-in Signal Transducer The major clue supporting signaling role of DG derives from the original observation that cellular adhesion to fibronectin, agrin, or laminin-α2 triggers the phosphorylation of a tyrosine within the β-DG cytodomain (Tyr 892), which is one of the residues involved in the interaction with the WW domain of dystrophin or utrophin [91,118] and in the binding of the SH3 domain of Grb-2, an adaptor protein involved in signal transduction and cytoskeleton organization both in skeletal muscle and brain [92]. Moreover, in bovine brain synaptosomes, β-DG binds phosphorylated focal adhesion (FAK) through the SH2 domain of Grb2 [130]. As a general mechanism, phosphorylation of β-DG might modulate its interaction with dystrophin or utrophin, functioning as a molecular switch between WW, SH3 and SH2 domains. Phosphorylation might alter the triggering of β-DG targeting to specific cellular compartments; in skeletal muscle tissues, phosphorylated β-DG has been localized to recycling endosomes together with c-src [91].

Functional DG Molecular Plasticity The DG gene has been widely described in vertebrates, while some surrogate forms, defined as DG-like, have been found in invertebrates. In Caenorhabditus elegans, deletion of the major DG-like gene (dgn-1) does not cause a muscle phenotype [131], but in Drosophila,

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similar connections with laminins and the dystrophin-linked cytoskeleton have been observed [132]. In lower vertebrate species, the primary structure of DG and associated proteins is highly conserved in several mammals and maintains a high degree of homology, suggesting that domain organization and the function of the complex have been conserved during evolution. However, knock-out DG experiments performed in lower species such as zebrafish does not affect the embryo, stopping their development and confirming the crucial role played by DG in the establishment of the first extraembryonic basement membrane, or Reichert‟s membrane [59,127]; however, DG plays a major role in morphogenesis in mice [114]. DG possesses a wide tissue distribution and is expressed in muscle, in the central and peripheral nervous systems, and in epithelia and endothelia [133]. The generation of mice with conditional knockout of DG in skeletal muscle and brain contributed to the understanding of its role in the muscular and central nervous systems; it is also well known that DG gene mutations do not link with muscular diseases, but are primarily linked with alterations of their maturation process and/or of their membrane localization, as have been observed in many neuromuscular disorders [134]. While chimeric mice develop severe muscular dystrophy [128], the skeletal muscle-specific ablation of DG results in a mild form of muscular dystrophy, revealing a role of DG in muscle regeneration promoted by satellite cells [135]. In brain, selected deletion of DG leads to important structural defects [136] and conditional DG knockout in peripheral nerves demonstrates its crucial role in myelination and in the architecture of Ranvier‟s node [137]. Although prior studies indicated the dispensable role of an α-DG cytoplasmic domain as a stabilizer of the glial-limit basement membrane during the development of the cerebral cortex and adult brain [138-140], recent studies have shown that deletion of DG in neurons blunted hippocampal long-term potentiation, indicating a novel role in synaptic plasticity [141]. At the neuromuscular junction, where its localization depends on its interaction with ankyrin [142], DG binds agrin with high affinity [143] and is involved in the stabilization of post-synaptic acetylcholine receptor clusters [144]. Recently, a specific role for DG was additionally demonstrated at retina, where it binds pikachurin, an extracellular matrix protein [98]. Dystroglycan can bind - and β-neurexins [96], promoting neuron-glia adhesions, in addition to its role as receptor for extracellular matrix proteins (laminin, perlecan, and agrin), which are important for the organization and development of both the peripheral nervous and central nervous systems [53,93,95,145-148].

The Intriguing Location of β-DG at the Nuclei Recently, the presence of some members of the DGC in nuclear matrix protein fractions and in in situ nuclear matrix has been reported [149-151]. The presence of a DGC complex was also confirmed in the nucleus of C2C12 muscle cells [152]. Its localization in the nucleus could reveal a role for β-DG as a transcription factor, but the lack of DNA binding motifs within its primary structure renders this highly unlikely, although it may function indirectly in binding yet unidentified transcription factors. In this respect, at the present stage it would be difficult to formulate any reliable hypothesis on its definite functional role in the nucleus for β-DG.

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In HeLa cells, the association of members of nuclear DAPC with nuclear matrix proteins lamin B1 and nuclear actin suggested that they may work as scaffolding proteins involved in nuclear architecture. As primary structure analysis showed, β-DG is the only component of the DGC that contains a potential Nuclear localization sequence (NLS), localized within the juxtamembrane region of its cytoplasmic domain. It has been suggested that this NLS triggers its nuclear translocation, together with all the of the previously associated DGC components, through nuclear pore complexes [153]. Upon tyrosine phosphorylation, β-DG undergoes a change in its localization from the plasma membrane to an internal-vesicle membrane compartment, which represents a subset of recycling endosomes in which it co-localized with c-Src [91]. Ultimately β-DG would release its anchorage to dystrophin or utrophin and undergo internalization in endosomes, increasing the possibility of relocating the β-subunit to the nucleus [154]. There is, perhaps, a temptation to speculate that β-DG, together with other elements of the DGC, could offer stability to the nucleus membrane or could even form part of the cell division system, stabilizing nuclear membranes, supporting or facilitating their adhesion and/or disassembly/assembly to other subcellular structures. Whatever the biological function of β-DG in the nucleus, it does not require the presence of α-DG, because it has not been detected in the cellular models studied to date, which provides further support for the idea that the DG precursor is necessarily cleaved into two subunits in order for these to carry out distinct functions. .

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Involvement of β-DG in Dystrophies Dystroglycan as a central element for the DGC is a key glycoprotein for its integrity in reducing or disrupting interactions with dystrophin, provoking severe Becker or Duchenne muscular dystrophy, respectively [125,126]. However, abnormal post-translational processing of DG is associated with congenital muscular dystrophies with Central nervous system (CNS) involvement ranging from mental retardation to structural defects, including cobblestone (type II) lissencephaly and hydrocephalus [99,155]. Analysis of conditional knock-out DG mice revealed that although neuronal DG does participate in forebrain development, DG expression in neurons is required for normal hippocampal long-term potentiation; in addition to the presence of DG, the integrity of its glycosylation is important for maintaining and potentiating the synapsis of CA3 and CA1 pyramidal neurons [99,114,116,138]. The common feature of different forms of secondary dystroglycanopathies, which range from severe congenital muscular dystrophy with CNS involvement to Muscle-eye–brain disease (MEB), Fukuyama congenital muscular dystrophy (FCMD), Walker-Warburg syndrome (WWS), and congenital Muscular dystrophies types 1C and 1D (MDC1C and MDC1D), to milder forms of Limb-girdle muscular dystrophy (LGMD2I and LGMD2N) [156], which is the defect in α-DG O-mannosyl glycosylation. In many cases, it has been reported that the severity of the dystrophic phenotype does not correlate with sarcolemma integrity or with mechanical defects, but rather with changes in the expression level of some proteins involved in signal transduction [157], suggesting an important contribution of one or more signal transduction pathways to myofiber degeneration [158]. Considering the central role for DG in the pathogenic mechanism of the human disease, including cases for which no causative mutation has been found, addressing the role

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of DG in signal transduction pathways might provide precious insights into the pathogenesis of many muscular dystrophies (Figure 8).

Figure 8. Glycosylation of -DG in normal muscle where glycans attached to the -DG link the dystrophin-glycoprotein complex to the extracellular matrix (agrin, neurexin, and laminin-2). (B) If the glycosylation pattern is lost in -DG, this impedes the link between -DG and the extracellular matrix.

DG in Signaling Pathways MEK/ERK pathway transmission of signals is usually initiated by the activation of a small G protein (e.g., Ras) followed by sequential activation of several sets of cytoplasmic protein kinases that are modulated by DG. It has been postulated that α-DG competes with the α6β1 integrin for binding with laminin, reducing the ERK activity triggered by α6β1 integrin engagement in laminin [159]; other mechanisms postulate that DG possesses a scaffold role in recruiting specific components of the signaling cascade, either by preventing their exposure to other components of the signaling cascade and/or by limiting their activity to defined cellular compartments, as has been described for Cos-7 cells, in which β-DG interacts with MEK within the membrane ruffles, uptaking it and preventing the phosphorylation of MEKcatalyzed ERK [160]. On the other hand, α-DG binding of full-length laminin α1, as well as its LG4-5 domain that contains the α-DG binding sites, causes the first of a series of phosphorylation reactions of factors in a signal cascade that culminates in the activation of the transcription factor c-jun,

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which might sustain myoblast proliferation, stimulating muscle hypertrophy. This same signal cascade appears to be activated by muscle tension [112,113]. Notwithstanding this, many studies have been focused on the role played by DG in signal transduction, but the precise molecular events in which DG is effectively involved and their real biological relevance require clarification.

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β-DG and the Many Faces of Actin The conversion of physical signals such as contractile forces or external mechanical perturbations into chemical signaling events enabling cells to adapt to their physical surroundings has been denominated mechanotransduction. This fundamental cellular process occurs at cell-extracellular matrix contacts known as focal adhesions, where transmembrane receptors are associated via their cytoplasmic domains with the actin cytoskeleton. The development, homeostasis, and regeneration of complex organ systems require extensive cellcell and cell-extracellular matrix communication to ensure that different cells proliferate, migrate, differentiate, assemble, and function in a coordinated and timely fashion. Cell adhesion molecules, once believed to function primarily only in cells tethering to extracellular ligands, are now widely recognized as having key functions in cellular signaling cascades. DG and its ligands are critical regulators of cell contact-dependent signaling and patterning and, depending on cell type and context, a limited and conserved set of receptor-ligand interactions is translated into a large variety of downstream signaling processes. β-DG has been localized in microvilli structures in a number of cell types, where it associates with the cytoskeleton adaptor ezrin through a cluster of basic residues in the juxtamembrane region of β-DG, modulating the actin cytoskeleton. Mutations preventing ezrin binding affect the induction of peripheral filopodia and microvilli [161]. Interestingly, dominant-negative Cdc42, which is capable of activating PAK kinase, inhibits DG-induced cell-surface protrusions, suggesting that the formation of membrane protrusions request a signal transduction pathway involving Cdc42 [162]. Recently, a role for DG involvement in the formation of podosomes has been proposed in myoblasts, where a ternary complex among β-DG, src kinase, and Tks-5 has apparently been detected [163]. Despite that DG is a widely expressed, well-characterized, and unique general adhesion protein, and although it has a well-established adhesion and mechanotransductive role in skeletal muscle [129,164], its role in other tissues is not as well understood. The presence of both - and β-DG in focal adhesions and a specific role for both of these in these structures has long been debated. It has been reported previously that -DG is present in focal-adhesionlike structures [161,165]. Other DG-associated proteins such as utrophin are present in focal adhesions in nonmuscle cells including platelets [89,90]. It is therefore assumed that in conditionally immortalized myoblast cell lines, β-DG participates in the formation of cell adhesions during cell spreading and in whether it can interact with proteins such as vinculin [166]. In platelets, β-DG was found to act as an interplay protein between actin and microtubules, and additional communication between these two cytoskeleton networks was maintained through proteins of the focal adhesion complex [88].

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DG and Cancer In addition to linking the ECM with the actin cytoskeleton of skeletal muscle, stabilizing the sarcolemma during contraction, and mediating a range of signaling events [94,164], DG is expressed in all other tissues, in which it performs a range of functions including basement membrane assembly, cell polarity, actin cytoskeleton organization, and transduction of extracellular signals [129]. Despite the critical function of -DG glycosylation in the muscular system, there are several reports that have shown its association with breast, colon, oral, and prostate carcinomas [167-170]. Reduced or altered expression of -DG has been demonstrated in pediatric tumors [171], prostatic cancer [172], cervical and vulval cancer [173], breast cancer, colon cancer [174], gliomas [175], and oral cancer [168]. There are also published studies on breast cancer cell lines [167,176,177]. It is well-known that normal epithelial cells are tightly associated with each other and with the underlying basement membrane in order to maintain tissue architecture and function. During cancer progression, primitive cancer cells escape from this control by modifying the binding affinities of their cell membrane receptors, concerning which several receptors have been described as important for this process, integrins being the most widely studied [178]. The -DG receptor has been reported to be required for the development and maintenance of epithelial tissues [133,179] and the profound effect that loss of DG expression has on cell polarity and laminin binding has been recorded in cultured mammary epithelial cells. This findings strongly suggest that -DG is not only important in the establishment and maintenance of epithelial structure, but also, that it serves as an effective means by which cancerous cells modify their adhesion to the ECM [167,180]. In addition, reduced expression of N-acetyl-lactosaminidebeta-1,3-N-acetylglucosaminyltransferase (β3GnT1) leads to diminished synthesis of laminin-binding glycans, greater migration of cancer cells, and increased tumor formation, playing a critical role in the synthesis of laminin-binding glycans in -DG. Restoration of laminin- binding glycans by forced expression of β3GnT1 restored laminin-binding glycans, reducing cell migration and tumor formation. Previous studies of epithelium-derived cancers [167,168,173,174,180,181] demonstrated that loss of immunoreactivity of -DG antibodies correlates with tumor grade and poor prognosis. This reduced detection of -DG, however, is based on a loss of -DG reactivity to antibodies (known as IIH6 and VIA4-1) that recognize the laminin-binding glycoepitope of -DG, therefore their functionality (functional glycosylation). However, in the majority of cancer samples studied to date, β-DG is expressed at normal levels in the cell membrane. Thus, the aforementioned cancer-associated loss of -DG expression may reflect a failure in the post-translational processing of DG, rather than in the synthesis of -DG itself. A similar defect in DG has been reported in a group of congenital muscular dystrophies [116]. This spectrum of human developmental syndromes involves brain, eye, and skeletal muscle and shows a dramatic gradient of phenotypic severity ranging from the most devastating in Walker-Warburg Syndrome to the least severe in limb-girdle muscular dystrophy. Six distinct known and putative glycosyltransferases have been shown to underlie these syndromes: Protein O-mannosyl-transferase 1 (POMT1); Protein O-mannosyltransferase 2 (POMT2); Protein O-linked-mannose beta-1,2-N-acetylglucosaminyltransferase

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1 (POMGnT1); like-glycosyltranferase (LARGE), and Fukutin and Fukutin-related protein (FKRP) [182-187]. The ability of DG to bind to laminin G domains containing ECM proteins and to mediate their polymerization is highly dependent on DG glycosylation status; however, it is necessary to consider a feasible epigenetic silencing of LARGE as the cause of the cancer-associated loss of functional dystroglycan-glycosylation. It has been confirmed by independent methods that -DG is hypoglycosylated and non-functional in terms of laminin binding in metastatic cell lines, although -DG is present at the cell membrane in all cases. These -DG hypoglycosylated cells are not able to anchor to the basement membrane or to organize ECM polymerization as effectively as normal epithelial cells, and their integration into the epithelium is compromised. This cancer-associated form of -DG may have a different function than the fully glycosylated one, shifting its affinity from laminin to a different ECM ligand. There has been very little work conducted on the β-DG molecule in human cancer, which may well be a more functionally significant protein in that it contains the cytoplasmic structural and signaling region; however, its expression has been found reduced or absent in prostatic and breast cancer, but only 15 and six cases, respectively, were studied [181] Given the fundamental cellular role of β-DG in cell adhesion, cell polarity, and cell signaling, it is likely that disruption of its expression and function could be implicated in carcinogenesis or tumor progression. An immunohistochemical survey of the expression of β-DG in a large number of samples, normal and neoplasic, from a wide variety of tissues have shown that in the majority of human cancers, there is loss or weak expression of -DG at both intercellular and basal cellular junctions. The only cancers to exhibit complete retention of -DG expression were cutaneous basal cell carcinomas, and it may be relevant that these tumors nearly never metastasize. [188]. In one previous small-scale study, DG loss in epithelial tumors suggested that DG may act as a tumor suppressor [167]; although evidence for this role is lacking, re-expression of DG in breast tumor cells implanted into nude mice significantly reduces tumor size [177]. However, although reduction of DG levels in normal cells does not induce tumor formation, the initial steps of DG loss are likely to be mediated by matrix metalloproteinase cleavage of the extracellular domain, generating a 31 kDa fragment [189], although the pathways leading to this fragment or to its complete degradation remain to be elucidated [190].

DG and Integrins Share Some Functions and Activation Pathways Considering DG as a cell adhesion molecule and its properties as a signaling molecule involving Src family kinases [91,118,191-193], as well as the information and evidence previously mentioned, a comparison to describe similarities and differences between integrins and DG comes to the fore. Although there is greater evidence related with the role of DG that impacts on its mechanical function, survival signaling and pathways including PI 3-kinase and Akt involving small and heterotrimeric G-proteins have been described. As was previously noted, during ligand recognition, integrin structure is subjected to structural modifications from the inactive bent form to the upright active conformation, with induction of signaling activation pathways. The different intracellular integrin responses are

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rendered by the diversity of combination options between  and β subunits and their ECM ligands and are represented in multiple genes. But unlike the different heterodimeric combinations, cellular distributions, and expressions, there is only one ubiquitously expressed -/β-dystroglycan heterodimer, which is not an obligate heterodimer. As has been previously mentioned, β-DG may possess functions in the absence of -DG. DG is represented by one gene, but the -DG cytoplasmic domain entertains the possibility of several predicted proteinprotein interaction sites that could mediate multiple functions. The best studied molecules involved in adhesion are integrins, while DG is beginning to achieve a place in this entangled process. With the aid of data derived from published experimental studies, Zaidel-Bar et al. in 2007 [194] addressed the molecular basis for integrin-mediated adhesion and signaling at multiple hierarchical levels. A 156-compartment network forming 690 potential interactions was created in which adhesion sites, such as focal complexes, focal adhesions, fibrillar adhesions, and podosomes, were included. This complex cellular adhesion system depends on spatiotemporal and organism-, tissue-, or cell-specific factors that determine the interactions that can and do take place. Despite DG ubiquitous distribution and diverse roles in multiple tissues, it has been studied most extensively in muscle, due to that DG was first considered a central element of the DGC. The podosomes and focal adhesion structures found in migratory myoblasts and mature myofibers highlight the versatility of this adhesion receptor, indicating multiple modes of interaction and regulation by tyrosine phosphorylation. Both integrins and DG have also been implicated in the assembly of laminin-based basement membranes. DG functions as a receptor essential for initial binding of laminin to the cell surface, whereas the β1 integrin and perlecan are required for laminin matrixassembly processes after it binds to the cell [3,195-197]. On the other hand, and although the receptor repertoire of all known laminins is not fully elucidated, it is well-known that laminins bind integrins and DG [198-200]. In this regard, integrins and DG appear to play complementary roles; the 6β1 integrin or the 7β1 integrin [201,202] possess the property of binding laminins with different affinities and specific mechanical functions and signaling. -DG-mediated cellular interactions with laminins are important for branching epithelial morphogenesis, such as in kidney, lung, or salivary gland [179,203]. Laminin fragments that activate the α1β1 integrin, the α6β1 integrin, and DG contribute to the attachment of monolayers of astrocytes in culture; therefore, integrins are essential for generating polarity, while DG is indispensable for extending adhesion processes [204]. Correct cell anchoring to the ECM is crucial to orchestrating and maintaining cellular structure and function. In the epithelium, these effects result from cooperation between the β1 integrin and DG, which connect the ECM to intracellular actin and intermediate filaments and that also elicit intracellular signal transduction and differentiation [160,167,178]. In fact, the changes in integrins β-1 and β-4, and in -2, -3, and -6 that have been reported in mammary tumors and in cell lines lead to a loss of polarity and invasiveness, and it has been documented that integrin misplacement can lead to the loss of polarity in epithelial cells [205]. Nevertheless, examples of opposing roles between integrins and DG have been reported. There is evidence that indicates that binding of DG to laminin, and most likely to perlecan and agrin, counteracts the signals initiated by integrin binding to extracellular molecules [206]. -DG competes with the 6β1 integrin for binding with laminin, reducing the ERK activity triggered by the 6β1 integrin engaged with laminin [159]. In a second

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mechanism proposed according to experiments performed in Cos-7 cells, β-DG recruits specific components of the signaling cascade (MEK), preventing exposure to other components of the signaling cascade (ERK) [161] (Figure 9). It is noteworthy that in addition to these direct linkers, there are dozens of potential connections consisting of two or more links, and the list of newly discovered components continues to grow. Intercellular adhesion and adhesion to the substrate define cell shape and tissue organization; knowing how cells adhere is important in understanding disease processes such as cancer and some muscular dystrophies, as both diseases closely related with a failure in cell adhesion.

Figure 9. Signaling pathways activated by recognition of integrins and DG involved in cell locomotion, survival and proliferation.

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[177] Sgambato A, Camerini A, Faraglia B, Pavoni E, et al. Increased expression of dystroglycan inhibits the growth and tumorigenicity of human mammary epithelial cells. Cancer Biol. Ther. 2004; 3: 967-975. [178] Muschler J, Lochter A, Roskelley CD, Yurchenco P, et al. Division of labor among the alpha6beta4 integrin, beta1 integrins, and an E3 laminin receptor to signal morphogenesis and beta-casein expression in mammary epithelial cells. Mol. Biol. Cell 1999; 10: 2817-2828. [179] Durbeej M, Larsson E, Ibraghimov-Beskrovnaya O, Roberds SL, et al. Non-muscle alpha-dystroglycan is involved in epithelial development. J. Cell Biol. 1995; 130: 7991. [180] Weir ML, Oppizzi ML, Henry MD, Onishi A, et al. Dystroglycan loss disrupts polarity and beta-casein induction in mammary epithelial cells by perturbing laminin anchoring. J. Cell Sci. 2006; 119: 4047-4058. [181] Henry MD, Cohen MB,Campbell KP. Reduced expression of dystroglycan in breast and prostate cancer. Hum. Pathol. 2001; 32: 791-795. [182] Beltran-Valero de Bernabe D, Currier S, Steinbrecher A, Celli J, et al. Mutations in the O-mannosyltransferase gene POMT1 give rise to the severe neuronal migration disorder Walker-Warburg syndrome. Am. J. Hum. Genet 2002; 71: 1033-1043. [183] van Reeuwijk J, Janssen M, van den Elzen C, Beltran-Valero de Bernabe D, et al. POMT2 mutations cause alpha-dystroglycan hypoglycosylation and Walker-Warburg syndrome. J. Med. Genet. 2005; 42: 907-912. [184] Yoshida A, Kobayashi K, Manya H, Taniguchi K, et al. Muscular dystrophy and neuronal migration disorder caused by mutations in a glycosyltransferase, POMGnT1. Dev. Cell 2001; 1: 717-724. [185] Longman C, Brockington M, Torelli S, Jimenez-Mallebrera C, et al. Mutations in the human LARGE gene cause MDC1D, a novel form of congenital muscular dystrophy with severe mental retardation and abnormal glycosylation of alpha-dystroglycan. Hum. Mol. Genet 2003; 12: 2853-2861. [186] Kobayashi K, Nakahori Y, Miyake M, Matsumura K, et al. An ancient retrotransposal insertion causes Fukuyama-type congenital muscular dystrophy. Nature 1998; 394: 388-392. [187] Brockington M, Blake DJ, Prandini P, Brown SC, et al. Mutations in the fukutin-related protein gene (FKRP) cause a form of congenital muscular dystrophy with secondary laminin alpha2 deficiency and abnormal glycosylation of alpha-dystroglycan. Am. J. Hum. Genet 2001; 69: 1198-1209. [188] Yamada H, Saito F, Fukuta-Ohi H, Zhong D, et al. Processing of beta-dystroglycan by matrix metalloproteinase disrupts the link between the extracellular matrix and cell membrane via the dystroglycan complex. Hum. Mol. Genet. 2001; 10: 1563-1569. [189] Singh J, Itahana Y, Knight-Krajewski S, Kanagawa M, et al. Proteolytic enzymes and altered glycosylation modulate dystroglycan function in carcinoma cells. Cancer Res. 2004; 64: 6152-6159. [190] Cross SS, Lippitt J, Mitchell A, Hollingsbury F, et al. Expression of beta-dystroglycan is reduced or absent in many human carcinomas. Histopathology 2008; 53: 561-566. [191] Ilsley JL, Sudol M,Winder SJ. The WW domain: linking cell signalling to the membrane cytoskeleton. Cell Signal 2002; 14: 183-189.

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[192] Ilsley JL, Sudol M,Winder SJ. The interaction of dystrophin with beta-dystroglycan is regulated by tyrosine phosphorylation. Cell Signal 2001; 13: 625-632. [193] Sotgia F, Lee H, Bedford MT, Petrucci T, et al. Tyrosine phosphorylation of betadystroglycan at its WW domain binding motif, PPxY, recruits SH2 domain containing proteins. Biochemistry 2001; 40: 14585-14592. [194] Zaidel-Bar R, Itzkovitz S, Ma'ayan A, Iyengar R, et al. Functional atlas of the integrin adhesome. Nat. Cell Biol. 2007; 9: 858-867. [195] Gardner H, Kreidberg J, Koteliansky V,Jaenisch R. Deletion of integrin alpha 1 by homologous recombination permits normal murine development but gives rise to a specific deficit in cell adhesion. Dev. Biol. 1996; 175: 301-313. [196] Fleischmajer R, Utani A, MacDonald ED, Perlish JS, et al. Initiation of skin basement membrane formation at the epidermo-dermal interface involves assembly of laminins through binding to cell membrane receptors. J. Cell Sci. 1998; 111 ( Pt 14): 1929-1940. [197] Sasaki T, Forsberg E, Bloch W, Addicks K, et al. Deficiency of beta 1 integrins in teratoma interferes with basement membrane assembly and laminin-1 expression. Exp Cell Res. 1998; 238: 70-81. [198] Mercurio AM. Laminin receptors: achieving specificity through cooperation. Trends Cell Biol. 1995; 5: 419-423. [199] Henry MD,Campbell KP. Dystroglycan inside and out. Curr. Opin. Cell Biol. 1999; 11: 602-607. [200] Aumailley M,Smyth N. The role of laminins in basement membrane function. J. Anat. 1998; 193 ( Pt 1): 1-21. [201] Sonnenberg A. Laminin receptors in the integrin family. Pathol. Biol. (Paris) 1992; 40: 773-778. [202] von der Mark H, Durr J, Sonnenberg A, von der Mark K, et al. Skeletal myoblasts utilize a novel beta 1-series integrin and not alpha 6 beta 1 for binding to the E8 and T8 fragments of laminin. J. Biol. Chem. 1991; 266: 23593-23601. [203] Durbeej M,Ekblom P. Dystroglycan and laminins: glycoconjugates involved in branching epithelial morphogenesis. Exp. Lung Res. 1997; 23: 109-118. [204] Peng H, Shah W, Holland P,Carbonetto S. Integrins and dystroglycan regulate astrocyte wound healing: the integrin beta1 subunit is necessary for process extension and orienting the microtubular network. Dev. Neurobiol. 2008; 68: 559-574. [205] Giancotti FG. Signal transduction by the alpha 6 beta 4 integrin: charting the path between laminin binding and nuclear events. J. Cell Sci. 1996; 109 ( Pt 6): 1165-1172. [206] Bao X, Kobayashi M, Hatakeyama S, Angata K, et al. Tumor suppressor function of laminin-binding alpha-dystroglycan requires a distinct beta3-Nacetylglucosaminyltransferase. Proc. Natl. Acad. Sci. USA 2009; 106: 12109-12114.

Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 177-205

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter V

Stressed Out and Actin Up: Stress-Activated Protein Kinase Regulation of Actin Remodeling Directs Endothelial Cell Morphology and Migration Meron Mengistu, Joshua B. Slee and Linda J. Lowe-Krentz

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Department of Biological Sciences, Lehigh University, Bethlehem, PA 18015.

Abstract Actin remodeling in the vascular system is central for functions such as vascular remodeling and contractility. Endothelial cells (EC) that form a monolayer lining the vasculature undergo remodeling of their actin cytoskeleton in order to (1) change their polarity, which gives them contractility that allows them to reduce their height in order to decrease the magnitude of the strain they experience from the blood flow, (2) migrate during vascular remodeling, and (3) maintain cell-cell contacts for endothelial integrity and barrier function. Dysfunction in ECs is the first step of atherogenesis, where cell morphology, migration, and barrier integrity are affected. Atherosclerosis is a geometrically focal disorder where endothelial dysfunction and subsequent plaque formation occur in areas of blood re-circulation, such as the outer wall of vessel bifurcations and areas near vascular branching points. Blood flow exerts different magnitudes of fluid shear stress (FSS) on endothelial cells, playing a crucial role in the localization of atherosclerotic lesions. ECs found in regions of arterial curvatures and bifurcations experience lower FSS conditions and are pre-disposed to atherosclerotic lesions, while ECs lining the straight parts of arteries are exposed to higher FSS conditions and are atheroprotected. FSS is an important regulator of EC morphology and migration. Under low FSS, ECs are polygonal in shape and experience high turnover, while they are elongated with their longer axes that align in the direction of flow when

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Meron Mengistu, Joshua B. Slee and Linda J. Lowe-Krentz exposed to higher FSS conditions. Such morphological change is driven by FSSstimulated fiber formation and alignment of these fibers in the direction of flow. Cell-cell integrity is also compromised during atherogenesis, where breaks in the monolayer of ECs provide openings for monocytes and low-density lipoprotein (LDL) to enter the wall of the artery and contribute to plaque formation. After atherosclerotic lesions have formed, one of the treatment options is angioplasty where the blocked or narrowed arteries are opened using stents. Endothelialization of stents, which requires EC migration, is important to control vascular tone and prevent restenosis, the re-narrowing of the artery. In order to remodel the actin cytoskeleton for morphology changes, migration and maintenance of the barrier function, FSS and other stimuli need to be translated into chemical signals. The signaling activities of Stress-Activated Protein Kinases (SAPKs) JNK and p38 are involved in actin remodeling events that regulate actin dynamics. Both JNK and p38 activities are transiently activated by the higher FSS treatments, and required to achieve actin alignment in the direction of flow. Other stimuli that alter actin remodeling and barrier changes also induce activation of stress kinases. JNK activity is detected in association with stress fibers and cortical actin, and its activity is required in EC morphology adaptation. p38 activity seems to play a role in actin remodeling near focal adhesions, where it is detected at the ends of stress fibers. In this chapter, we review the role of these SAPKs in actin remodeling that leads to EC alignment in the direction of flow, migration, and maintenance of cell-cell integrity.

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I. Introduction Mechanical forces in the body have been shown to play important roles in regulating morphologies and functions of cells. Perhaps the most visible cellular response to mechanical forces occurs in the cardiovascular system, where vascular cells are constantly exposed to forces from blood flow. Hemodynamic forces primarily acting on endothelial cells (ECs), which form a monolayer that lines the vasculature, dictate the structure and function of these cells. ECs lining the vasculature in regions of arterial curvature experience low fluid shear stress (FSS) (an average of less than 4 dynes/cm2) from the re-circulating, non-laminar, blood flow, and adopt a different morphology and function compared to those found in the straight parts of the artery where blood flow is rapid, laminar and axial, and exerts a relatively higher FSS (greater than 15 dynes/cm2) on these cells (Caro et al., 1969; Girard and Nerem, 1995; Malek et al., 1999). These different magnitudes of FSS, exerted by the blood flow on ECs, play a crucial role in the localization of atherosclerotic lesions. Atherosclerosis is a cardiovascular disorder, and dysfunction of ECs is believed to be the first step of atherogenesis. Factors circulating throughout the vasculature such as high levels of modified-low-density lipoprotein (modLDL) and glucose, low levels of HDL, and radicals from cigarette smoke, as well as conditions such as obesity, diabetes, hypertension, stress, and viral infections have all been shown to contribute to endothelial dysfunction and plaque formation (Ross, 1999), but the localization of these atherosclerotic lesions appears to be dictated by FSS. ECs experiencing low FSS as a result of blood re-circulation in areas such as the outer wall of vessel bifurcations and areas near vascular branching points are prone to atherosclerotic plaque formations, while cells in straight parts of arteries where blood flow is smooth experience higher magnitudes of FSS, and are protected from plaque formation (reviewed in Chiu and Chien, 2011; Cunningham and Gothlieb, 2005 and summarized in figure 1).

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Figure 1. FSS plays a role in EC dysfunction that leads to atherogenesis. In regions where FSS ≤ 4 dyn/cm2, corresponding to regions of disrupted blood flow, ECs become dysfunctional, becoming “sticky,” and provide adhesion sites for monocytes and platelet aggregates, which can then penetrate the endothelium (“leaky” endothelium). In regions of a bifurcating artery where blood flow is smooth, and the FSS ≥ 15 dyn/cm2 FSS, ECs do not provide adhesion sites for monocytes and platelets, and the endothelium remains intact. Cell morphology is different under the different FSS conditions, where ECs elongate and align in the direction of flow under high FSS, but remain cuboidal and have random orientations under low FSS. These morphological changes are driven by remodeling of their actin cytoskeleton which rearranges to align in the direction of flow in the former, but is randomly oriented under the later FSS condition.

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The macroscopic changes that lead to atherogenesis result from changes at the microscopic level within endothelial cells, partly induced by different flow conditions in the arterial circulation system. In this chapter, endothelial cell responses to different FSS conditions that mimic the athero-prone and athero-protected regions of the artery will be examined, and the role of stress activated protein kinases (SAPKs) JNK and p38 in mechanotransduction of these FSS stimuli into actin cytoskeleton remodeling to induce EC morphology changes will be the focus. Endothelial barrier function and EC migration also dysfunction during atherogenesis. Their dependence on SAPK-mediated actin cytoskeleton remodeling and will be discussed for comparison.

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II. Fluid Shear Stress-Induced Changes in Endothelial Cell Morphology Result from Actin Reorganization Endothelial cells exposed to low FSS conditions, like in the athero-prone regions of arterial curvature, adopt a different morphology and function than ECs found in regions experiencing higher FSS. These FSS-induced EC responses are summarized in Figure 1. Fluid shear stress-induced morphological changes in ECs are results of reorganization of cytoskeletal structures. The cytoskeleton is a network of filamentous proteins that is distributed throughout the cytoplasm of a cell, and it plays a role in maintaining mechanical integrity of the cell. It is composed of three major types of filaments: actin filaments (or microfilaments), microtubules, and intermediate filaments. Even though the cytoskeleton provides the cell‟s framework, it is dynamic; adapting to changes in the environment by constantly polymerizing and de-polymerizing its components, depending on the necessary functions of the cell. All three cytoskeletal filaments undergo rapid displacement and deformation to reorient in the direction of flow (Flitney et al., 1996; Franke et al., 1984; Galbraith et al., 1998; Helmke et al., 2001; Malek and Izumo, 1996; Sato et al., 1987; Wechezak et al., 1985). The cytoskeleton has been shown to be responsible for the transmission of mechanical stresses from the cell surface to various intracellular locations such as cell-cell adhesion sites, focal adhesion sites and the nucleus (Barakat and Davies, 1998). Cytoskeletal remodeling as a response to FSS may serve various functions, such as movement of neutrophils, and pro- and anti-inflammatory stimuli (Wang and Doerschuk, 2001). The actin cytoskeleton undergoes the most significant rearrangement in response to FSS stimuli. Actin makes up 10% of the total endothelial cell proteins (Lehoux and Tedgui, 2003), and actin filaments form a network that spans the intracellular space, making connections between most cellular structures. This makes actin crucial to the transmission of forces throughout the cell, possibly by playing a role in scaffolding and/or translocation of signaling molecules and organelles.

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a) Endothelial Cells Exposed to Low Fluid Shear Stress Do Not Remodel Their Actin Cytoskeleton to Align in the Direction of Flow Regions of the artery that are prone to atherosclerotic lesion formations are marked by relatively low FSS conditions (less than or equal to 4 dyn/cm2), which result from disrupted blood flow that undergoes abrupt cessation and re-circulation. ECs conditioned with disrupted flow and characteristically lower FSS adopt a polygonal cell shape, and have thin cortical actin and filaments that are randomly oriented within the cell. Their cell-to-cell connections are compromised, possibly due to the absence of a thicker cortical actin that allows cells to maintain an intact monolayer that can withstand forces from the blood. This outcome results in a permeable (“leaky”) endothelium that is penetrated by monocytes and platelet aggregates which contribute to plaque buildup. ECs in these regions also undergo increased cell turnover (Davies et al., 1986; Langille et al., 1986) and have impaired repair responses during endothelial denudation after injury (Vyalov et al., 1996), which is discussed below in section VI. Low FSS conditions in these regions also induce increased cell proliferation through the generation of reactive oxygen species (ROS) (Milovanova et al., 2008).

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b) High Fluid Shear Stress Induces Three-Phase Actin Remodeling Under the athero-protective higher FSS conditions, ECs elongate and align their long axes in the direction of flow (reviewed in Chiu and Chien, 2011; McCue et al., 2004). FSSinduced actin remodeling in ECs occurs in three phases (Mengistu et al., 2011) which are described in figure 2. Phase1: Within 5 minutes of high FSS treatment, stress fibers were formed. Stress fibers are actomyosin bundles composed of 10-30 actin filaments held together by cross-linking proteins such as -actinin and filamin, and non-muscle myosin and tropomyosin (Cramer et al., 1997; Lazarides and Burridge, 1975; Wang et al., 1975; Weber and Groeschel-Stewart, 1974). Stress fiber formation is a conserved adaptation of eukaryotic cells, but its assembly is not very well understood. Formation of these structures as a response to FSS is believed to play a role in giving ECs contractility. Stress fibers have also been shown to be mechanosensitive, and play a role in mechanical homeostasis by allowing cells to undergo contraction forces and maintaining their shape by resisting stretching forces (reviewed in Deguchi and Sato, 2009). After 15 minutes of high FSS treatment, a cluster of actin, which could represent a pool of newly made actin, was often found near nuclei. These clusters could not be detected in the presence of cycloheximide, which inhibits protein synthesis (Mengistu et al., 2011). Similar actin behavior has been reported in ECs exposed to FSS (Noria et al., 2004) and biaxial stretching (Wang et al., 2001). Phase 2: After 30 minutes of exposure to the high FSS, a dense peripheral band of actin could be seen, inducing ECs to become more regular and round in appearance. This actin band is composed of actin, myosin, tropomyosin, and α-actinin, and reinforces a robust cellto-cell contact important for an intact endothelium. The contractile cortical endothelial ring not only helps maintain EC-to-EC adhesions, but also gives these cells the capability to contract and regulate permeability of the endothelium (Schnittler, 1998; Schnittler et al., 2001). After 30 minutes of exposure to 15 dyn/cm2 FSS, remodeling resulted in a cortical

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actin ring after 30 (see figure 2). The appearance of this dense cortical actin was preceded by the appearance of a pool of actin near the nucleus in phase 1, believed to be newly synthesized actin, which could have been transported to the cell periphery accounting for the increase in actin signal (Mengistu et al., 2011). Phase 3: Between 60 and 120 minutes of 15 dyn/cm2 FSS treatment, endothelial cells began to form stress fibers that ran parallel to the longer axes of the cells, and the fibers also started aligning in the direction of flow. This actin formation induced ECs to elongate and align their longer axes in the direction of flow as well. Such morphology also allows ECs to decrease their height in order to decrease the magnitude of the strain they experience from hemodynamic forces (Barbee et al., 1994; Hu et al., 2003; Karcher, 2003; Pellegrin and Mellor, 2007).

Figure 2. Summary of the high fluid shear stress-induced changes in EC morphology, actin cytoskeleton (red), as well as the spatial and temporal distribution of the active phosphorylated forms of SAPKs JNK (green), and p38 (blue) relative to actin. These high FSS-induced changes are divided into 3 phases based on responses in actin remodeling.

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Nuclear Actin: Actin staining was detected in the nucleus of ECs in all 3 phases of FSSinduced actin remodeling. Some increased nuclear actin was observed during the first phases of actin remodeling, but nuclear actin returned to resting levels by phase 3 (Mengistu, et al., 2011). These nuclear actin structures are discussed in depth below in section II.C.

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c) Fluid Shear Stress Induces Nuclear Actin Formation Published data indicate that not only do entire cells and the actin cytoskeleton elongate in the direction of FSS, but the nuclei of those cells elongate in the direction of FSS as well (Flaherty et al., 1972). These results suggest that the nucleus is in some way “sensing” the FSS and evoking an appropriate response. This is not entirely surprising given that there are well-defined links between cytoskeletal structures such as actin and the nucleus, which could transmit signals between the nucleus and the cytoplasm (reviewed in Starr and Fridolfsson, 2010). Even more interesting is the fact that actin, which when localized to the cytoplasm responds to FSS, is also found in the nucleus (Bittinger et al., 2004; Castano et al., 2010; Mengistu et al., 2011). This presents an exciting possibility that nuclear actin may be mediating certain responses to various types of cellular stress, a possibility which is gaining popularity in the field of nuclear actin research (Visa and Percipalle, 2010). Although to the knowledge of these authors, there have been no published studies specifically examining nuclear actin responses to FSS to date, there have been studies investigating the role of actin and actin-binding proteins in the nucleus. The existence of nuclear actin has been of much historical debate, but the consensus now is that nuclear actin exists, albeit with slightly different behavior than its cytoplasmic counterpart. These subtle differences, mainly structural, made characterizing nuclear actin difficult. These structural differences include the lack of long filamentous actin typical of that found in the cytoplasm which can be bound by phalloidin (Castano et al., 2010). Further evidence for actin in the nucleus was gained from the discovery of actin-binding proteins also present in the nucleus implicating actin in a wide range of essential nuclear processes, such as chromatin remodeling, gene expression, DNA replication and repair, nucleocytoplasmic transport, and sensing outside the nucleus (for detailed reviews see: Castano et al., 2010; Visa and Percipalle, 2010). Thereby suggesting that actin once thought of as solely a cytoplasmic protein is critical for proper functioning of many nuclear processes. Specifically related to this chapter, it has been shown that upon different forms of cellular stress, actin translocates to the nucleus and that this process is mediated by cofilin, a member of the actin depolymerizing factor family of proteins (Pendleton et al., 2003). The authors noted increased actin staining in the nuclei of mast cells treated with latruculin B or ATP depleted cells and that treatment with an anti-cofilin antibody prevents this accumulation in the nucleus (Pendleton et al., 2003). These results suggest that as actin lacks a nuclear localization sequence, it must associate with a protein such as cofilin in order to gain entry into the nucleus through traditional nuclear pore complex mechanisms. A pioneering study of cofilin indicated that in rat fibroblastic 3Y1 cells exposed to heat shock or dimethyl sulfoxide stresses, cofilin is dephosphorylated and accumulates in the nucleus (Ohta et al., 1989). Taken together these two studies indirectly suggest that under cellular stress cofilin is dephosphorylated and escorts actin to the nucleus, where it likely plays a role in mediating cellular responses to various stressors, possibly including shear stress.

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A recent article published by Goyal and colleagues (Goyal et al., 2011) suggests that a novel serine/threonine kinase, STK35L1, which they show associates with nuclear actin, is involved in the regulation of the cell cycle and migration in ECs (HUVECs – human umbilical vein ECs). The authors noted that depletion of STK35L1 by siRNA accelerated the G1 to S phase transition of the cell cycle, indicating a normal role for STK35L1 in regulating this transition (Goyal et al., 2011). The protein playing a role in mediating this affect was determined by cell cycle-specific gene array analysis to be p16INK4a (Goyal et al., 2011). Silencing of STK35L1 in cells grown on matrigel results in decreased EC sprouting, an essential step for angiogenesis, suggesting a role for STK35L1 in promoting migration and new blood vessel formation (Goyal et al., 2011). STK35L1 was also found to co-localize with actin in the nucleus of HUVECs, and bioinformatic searches led to the discovery of a predicted class III PDZ domain binding motif which could mediate this interaction (Goyal et al., 2011). This study suggests a potential link between the cell cycle and migration in ECs mediated through STK35L1 and its association with nuclear actin. Although the authors did not determine the role of nuclear actin association, it is likely important for localizing or supporting the effects of STK35L1. While much remains to be discovered regarding nuclear actin, emerging evidence suggests that it is playing a major role in many nuclear processes. It is easy to foresee a mechanism where cells exposed to FSS transduce this form of mechanical stress (Wang et al., 2009), resulting in cofilin escorting actin to the nucleus, where it and actin binding proteins, such as STK35L1, are capable of altering gene transcription and chromatin structure to alter protein expression in order to respond to and manage the stressor. As is the case for STK35L1, these changes may lead to inhibition of the cell cycle, migration away from the stressor, or modulation to a morphology better suited to handle certain types of cellular stressors.

III. Stress Activated Protein Kinases JNK and p38 Are Involved in Mechanotransduction of Fluid Shear Stress into Actin Adaptation Although FSS-induced endothelial remodeling is a phenomenon that has been reproduced in numerous laboratories (reviewed in Chiu and Chien, 2011; Cunningham and Gothlieb, 2005) and is being extensively studied, the underlying mechanisms by which these cells achieve these changes are largely unknown. ECs have the capability of sensing differences between low and high FSS conditions (mechanosensing), and translating these mechanical stimuli into biochemical signals (mechanotransduction). While only the apical surface of ECs is exposed to this mechanical force, sites removed from the stimulus, like the nucleus, focal adhesion sites, and cell-cell junction sites; need to also undergo changes in order to attain such cellular deformations.

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a) Fluid Shear Stress Is Detected by Mechanosensors on Endothelial Cell Surfaces The plasma membrane (PM), which is constantly exposed to blood flow, has been shown to undergo changes as a response to FSS, increasing in fluidity (Butler et al., 2001; Haidkker et al., 2000), and inducing the activation of MAPK signaling cascades (Butler et al., 2002). Other than being a mechanosensing surface, the PM also contains components that have been previously identified as mechanosensors which locally respond to changes in blood flow by inducing intracellular signaling. Integrins are transmembrane glycoproteins that form at focal adhesion (FA) sites, where their extracellular domain forms attachments with extracellular matrix (ECM) proteins and their cytoplasmic domain connects with cytoskeletal and signaling proteins (Hynes, 1999; Li et al., 1997), making them desirable as mechanosensors. FSS modulates integrin affinity and strength of binding to the ECM (Bazzoni and Hemler, 1998; Tzima et al., 2001), and induces rapid remodeling and alignment of FAs in the direction of FSS, which cells achieve through flow-induced phosphorylation of focal adhesion kinase (FAK) and Src (Davies et al., 1993; Davies et al., 1994; Ishida et al., 1996; Tseng et al., 1995). FA remodeling is an event that is important for ECs to achieve actin alignment in the direction of flow. In vivo, integrin expression levels were found to be higher in athero-prone regions of arteries compared to atheroprotective regions (Tzima et al., 2001), indicating that their levels are modulated by FSS. Integrins have also been shown to activate the MAPK signaling proteins JNK via Shc (Chen et al., 1999; Jalali et al., 2001), and ERK through Rho (Jalali et al., 2001; Li et al., 1999; Tzima et al., 2001). Together they modulate FA and actin alignment in the direction of flow. These mechanotransduction events through integrin mechanosensors are summarized in figure 3 (red arrows). Another transmembrane glycoprotein playing a role in mechanosensing is Platelet Endothelial Cell Adhesion Molecule (PECAM), also known as CD31 and endoCAM. PECAM is mostly found at cellcell junctions, and FSS has been shown to result in phosphorylation of its cytoplasmic domain (Harada et al., 1995; Osawa et al., 1997), which induces the recruitment of Shp-2, a tyrosine phosphatase, which then activates the Raf-MEK-ERK pathway through Ras (Fujiwara et al., 2001; Masuda et al., 1997) (figure 3, green arrows). This PECAM pathway has also been shown to be involved in alignment of actin filaments in the direction of FSS, where PECAM1-/- mice exhibited defects in response to flow (Tzima et al., 2005). In vivo, PECAM-1 tyrosine phosphorylation is also seen in regions of arteries exposed to higher FSS conditions (Kano et al., 2000), which are atheroprotected regions, where it might play a role in maintaining the integrity of the endothelium at cell-cell junctions. The transmission of forces from PECAM to other intracellular sites could be achieved through associations with the actin cytoskeleton or intermediate filaments. PECAM-1 has been shown to interact with - and catenin, which are molecular linkers (Green and Jones, 1996). Caveolae, which are spherical invaginations of the plasma membrane, have also been implicated in playing a role in EC mechanosensing. Caveolin-1, a crucial constituent of caveolae, allows these structures to sequester signaling molecules and regulate several endothelial functions (Minshall et al., 2003). Caveolin-1 is associated with G protein -subunits (Gudi et al., 1996), Src family kinases (Minshall et al., 2000), Ras, Rac (Fanger et al., 1997), and calmodulin (Wang and Abdel-Rahman, 2002), concentrating them at caveolar sites of the cell and regulating their activities, usually by rendering them quiescent.

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Figure 3. A model for shear-stress mediated mechanotransduction events through various mechanosensors (integrin, PECAM, caveola, K+ & Ca2+ channels, G protein & Tyrosine Kinasecoupled receptors) that activate the MAPK signaling cascade through upstream signaling small GTPases such as Src, Ras, Rac and Rho. Signaling events through integrins are outlined in red, caveolae in blue, and PECAM in green. Double-line arrows represent signaling pathways originating from the nucleus, and dashed arrows represent pathways with unknown intermediates. Nuclear actin translocation to the nucleus in a cofilin-dependent fashion has been shown in other models, but not as a response to FSS and therefore represented with “?”.

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FSS has been shown to induce increased numbers of caveolae formatted on the luminal surfaces of ECs (Boyd et al., 2003; Rizzo et al., 2003; Sun et al., 2002). The selective localization of many signaling molecules to caveolar domains, suggests that these structures function by compartmentalizing various signaling molecules and serving as organizing centers, which are crucial for the cell‟s responses to external stimuli. This caveolar organization of the plasma membrane into different functional units might be important for rapid activation of signaling molecules, which can occur by posttranslational modification events, such as phosphorylation of caveolin-1. Caveolin-1 also associates with 1-integrins (Radel et al., 2007) and caveolae have possible linkages to the actin cytoskeleton (Mundy et al., 2002), which will allow them to relay mechanical stimuli to other sites within the cell (figure 3, blue arrows). In vivo, caveolae are also responsive to FSS exposure where caveolin1-deficient mice were shown not to undergo vascular changes in response to changes in FSS (Yu et al., 2006). Other mechanosensors such as G protein- (Gudi et al., 1996; Gudi et al., 2003; Jo et al., 1997) and tyrosine kinase-coupled receptors (Chen et al., 1999; Jin et al., 2003; Wang et al., 2002), ion channels [K+/Na+ (Alevriadou et al., 1993; Hutcheson and Griffith, 1994; Olesen et al., 1988), and Ca2+ influx (Ando et al., 1988; Helmlinger et al., 1996; Kanai et al., 1995; Yamamoto et al., 2000)]; have also been identified. A 300-400 nm thick coating consisting of glycoproteins, proteoglycans, and glycosaminoglycans, known as the glycocalyx, has also been implicated as a mechanosensor in ECs (Florian et al., 2003; Orr et al., 2006; Smith et al., 2003; Weinbaum et al., 2003). Florian et al. (Florian et al., 2003) showed that when this layer was enzymatically removed, fluid shear stress-induced nitric oxide production was inhibited. Mechanosensing through several structures and mechanotransduction through MAPK signaling cascades is schematically represented in figure 3.

b) Stress Activated Protein Kinases JNK and p38 Mediate Fluid Shear Stress Mechanotransduction Events In order to alter cytoskeletal dynamics, FSS stimuli detected by mechnosensors are translated into chemical signals, a phenomenon termed mechanotransduction. Fluid shear stress has been shown to activate mitogen-activated protein kinase (MAPK) signaling cascades, each of which is a series of phosphorylation and activation events that start with MAPKKKs (or MEKKs) that activate MAPKKs (or MEKs), which in turn finally activate the 3 MAPKs, extracellular signal-regulated kinase (ERK), c-jun NH2-terminal kinase (JNK), and p38 (figure 3). ERK is activated within 5 minutes of high FSS exposure, and has been shown to mediate endothelial nitric oxide synthase (eNOS) production of nitric oxide (NO), a vasodilator and an inhibitor of adhesion molecules. ERK activity is dependent on Gi, Ras, Src, FAK, and PKC protein kinases (Boyd et al., 2003; Cunningham and Gothlieb, 2005; Park et al., 2000). ERK activity, which promotes cell growth and survival, is possibly required for the cell‟s anti-inflammatory response. Although ERK has been shown to regulate actin organization by modulating focal adhesions (Han et al., 2007) only JNK and p38 have thus far been shown to be directly involved in endothelial actin alignment in the direction of flow (Azuma et al., 2001; Chen et al., 1999; Chien, 2007; Jalali et al., 1998; Kadohama et al., 2006; Mengistu et al., 2011; Yoshizumi et al., 2003).

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The SAPKs JNK and p38 are transiently activated for up to an hour of FSS exposure, and their activity is also very important for many cellular functions including pro- and antiinflammatory events that play a role in atherosclerotic plaque localization (Wang et al., 2001). JNK and p38 on one side and ERK1/2 on the other have competitive effects, and balance is achieved through “crosstalk” between these MAPK signaling cascades (Azuma et al., 2001). Such interactions are made evident through JNK-ERK and p38-ERK interactions. ERK has been shown to deactivate JNK by regulating MAPK phosphatase-2 expression, and p38 interacts with and negatively regulates ERK1/2 by preventing its phosphorylation by MEK1/2 (Azuma et al., 2001). Having to share the same signaling molecules for their activation, the cell can attain ERK and JNK specificity by compartmentalizing these signaling molecules in microdomains such as caveolae. These cholesterol-sensitive microdomains have been shown to be involved in the mechanosensitive activation of ERK, which is regulated through caveolin-1, but not JNK (Rizzo et al., 1998).

c) Actin Cytoskeleton Remodeling That Leads to Endothelial Cell Morphology Changes Is Mediated by Stress Kinase Activities

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The SAPKs JNK and p38 are differentially regulated by FSS. The temporal activation and spatial distribution of JNK and p38 required for the 3-phase actin remodeling as a response to FSS are summarized in figure 2. JNK JNK is an effector kinase which is activated through phosphorylation by MAPK kinases MKK4 and MKK7, which are brought together in close proximity by the actin binding protein filamin A (FLNa) (Nakagawa et al., 2010). JNK is also found downstream from the small GTPases Rac and Cdc42, which are involved in actin reorganization (Garrington and Johnson, 1999; Hall, 1998; Ip and Davis, 1998). G/, phosphatidylinositol-3-kinase- (PI3K-), Ras, Src and FAK protein kinases have been found to induce JNK activation and actin depolymerization leading to its remodeling (Azuma et al., 2001; Bagrodia et al., 1995; Hoefen and Berk, 2002; Li et al., 1997; Otto et al., 2000), and a study by Otto et al. (Otto et al., 2000) identified a downstream target of JNK, p150-Spir, which has been shown to be involved in actin reorganization (Bi and Zigmond, 1999; Ramesh et al., 1999). In vitro, the lower FSS treatments that mimic athero-prone regions of the artery did not induce large changes in cytoplasmic JNK activity levels, while athero-protective FSS substantially increased its activity. The spatio-temporal tracking of active phosphorylated JNK under higher FSS conditions revealed most of the changes in JNK activity levels did not occur in the nucleus where FSS could be inducing activation of transcription factors, but rather occurred in the cytoplasm. JNK co-localized with three different forms of actin; (i) stress fibers, (ii) a pool outside the nucleus, and (iii) cortical actin (Mengistu et al., 2011). In phase 1 of high FSS-induced actin remodeling, active phosphorylated JNK was found to localize with stress fibers, but its activity was not required for stress fiber formation. Active JNK was also found to co-localize with a pool of actin outside the nucleus, which could be a cluster of newly synthesized G-actin. In vascular smooth muscle cells, JNK has been shown to play a role in the regulation of smooth muscle alpha-actin expression (Garat et al., 2000),

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and this kinase activity has also been shown to be involved in the regulation of the expression of four different β–actin forms (Ha et al., 2008). At least 2 different JNK-dependent pathways are possibly involved in phase 2 of FSSinduced actin remodeling. (1) JNK activity is involved in the maintenance of cell-cell associations through regulation of the dense cortical actin, in an ATP-dependent process. (2) JNK also appears to play a role in the delivery of newly made actin to cell peripheries, and regulation of the amount of peripheral actin (Mengistu et al., 2011). Since JNK does not directly phosphorylate actin, it could be acting on components of the contractile cortical endothelial ring, which is composed of actin, myosin, tropomyosin, and -actinin. Yang et al. (Yang et al., 2007) reported JNK association with stress fibers depends on α–actinin, which contributes to the activation of this kinase (Yang et al., 2007). In phase 3 of actin remodeling, JNK activity appears to be required for the alignment of stress fibers in the direction of flow, which is dependent on disbanding the dense cortical ring formed in phase 2 (Mengistu et al., 2011). JNK substrate phosphorylation appears to be required for this step, but the substrate(s) has not yet been identified. ECs exposed to lower FSS conditions do not undergo this phasic actin remodeling. The multiple stages of actin rearrangement could reflect the need to first enhance barrier function, an important role for cortical actin in ECs (Prasain and Stevens, 2009). The low levels of cytoplasmic active phosphorylated JNK could be responsible for the lack of dense cortical actin that supports EC-to-EC contacts, leading to a permeable, “leaky,” endothelium seen in athero-prone regions of the artery. p38 p38 is involved in FSS-induced actin alignment and EC elongation, where inhibition of its activity prevented this actin alignment and cell orientation change as a response to flow (Kadohama et al., 2006). One mechanism through which p38 can achieve this is by phosphorylating MAPK-activated kinase-2 (MAPKAP-2), an activator of heat shock protein27 (HSP27) whose activity uncaps actin filaments and destabilizes them (Azuma et al., 2001; Hoefen and Berk, 2002). p38 is an important regulator of actin and focal adhesion remodeling as a response to FSS (Azuma et al., 2001; Hoefen and Berk, 2002; Kadohama et al., 2006; Rousseau et al., 1997), but the mechanisms through which it achieves these functions are not well understood. FSS exposure induces p38 activation and differential spatial and temporal distributions (figure 4). In the absence of flow, active, phosphorylated p38 (phospho-p38) was evenly distributed in the cytoplasm of ECs (figure 4, top green). When exposed to high FSS (15 dyn/cm2) for 30 min, fluorescence labeling of active phosphorylated p38 in ECs exhibited an up-regulation of punctate fluorescence signals, (figure 4, middle green). ECs also form a dense cortical actin band (figure 4, middle red), and phospho-p38 signals co-localize at ends of actin filaments at focal adhesion sites of actin, where cells make contact with the ECM. Focal adhesion remodeling plays a significant role in FSS-induced BAEC remodeling, and p38 could be playing a role in signaling to and/or from these ECM contact sites. After 60 minutes of high FSS treatment, phospho-p38 levels were very low, where active p38 was primarily detected in the nuclei of ECs (figure 4, bottom, green). At this time point, phosphop38 did not co-localize with the actin cytoskeleton that is beginning to align in the direction of flow (figure 4, bottom, red and Merge).

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Figure 4. Spatial and Temporal Distribution of Phospho-p38 in ECs exposed to 15 dyn/cm2 FSS and Association of Phospho-p38 with Actin. Phospho-p38 was fluorescently labeled (green) in ECs exposed to no flow, or 15 dyn/cm2 FSS for 30 and 60 minutes. Acquisition settings for ECs that were not treated with flow were saved and used for subsequent imaging of these FSS exposed cells to make possible comparison of the relative intensity levels of phospho-p38 labels. Phospho-p38 (green) and actin (red) were simultaneously labeled with primary antibodies against the proteins, and their spatial distribution was visualized in ECs exposed to FSS for 30, and 60 minutes. Co-localization between phospho-p38 and actin can be seen as yellow signals (Merge). Range bars = 10 m.

ECs treated with SB203580, a very specific p38 inhibitor, have enhanced cortical actin formation in the Phase 2 of FSS-induced actin remodeling (figure 5B) compared to cells not treated with this inhibitor (figure 5A). This inhibitor also affected the transition from phase 2 to phase 3, where the cortical actin could not remodel into stress fibers (figure 5B). These results suggest that p38 activity is necessary for signaling to and/or from focal adhesions and is needed for cortical actin remodeling into flow-aligned stress fibers. On the other hand, JNK inhibition with SP600125 dramatically limited the formation of a dense cortical actin in phase 2, which compromised EC-EC contacts resulting in cells being washed off by increased exposure to high FSS (Mengistu et al., 2011). When both p38 and JNK SAPKs were inhibited simultaneously in ECs with SB203580 and SP600125, respectively, an intermediate response to the individual treatments was observed for phase 2 actin remodeling, where cortical actin, less dense than in untreated cells (figure 5A) but slightly more dense than with SP600125 treatments (Mengistu et al., 2011) was formed (figure 5C).

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Figure 5. Effects of p38 inhibitor SB203580 alone and in combination with JNK inhibitor SP600125 on FSS-induced actin remodeling. ECs were treated with 10-M of SB203580 for 1 hour (B), or with 5M of SB203580 + 5-M SP600125 (10-M total) (C) then exposed to 15 dyn/cm2 FSS for 0, 30, 60, and 120 minutes. The actin cytoskeleton was then visualized using TRITC-conjugated phalloidin. The effect of the inhibitor treatment on actin adaptation is made evident through comparisons with the actin distribution of untreated ECs treated with high FSS for 0, 30, 60, and 120 minutes (A). ECs treated with 10-M SP600125 then exposed to 15 dyn/cm2 FSS for 0, 30, 60, and 120 minutes were previously published in Mengistu et al., 2010. Image acquisition settings were the same for all treatments, and increase in intensity levels indicates increase in protein levels. Scale bars = 10 m.

The cortical actin also seems to remodel into stress fibers after 60 minutes of high FSS exposure, although this phase 3 remodeling led to weakening of EC-EC contacts where cells are washed off with increased exposure to flow (figure 5C, 120 min). Blocking both JNK and p38 activities appears to partially overcome inhibition of phase 2-to-phase 3 transitions seen in figure 5B, suggesting that these SAPKs might have synergistic roles in FSS-induced actin remodeling. However, the presence of a dense cortical actin in these cells did not prevent cell detachment; therefore, these SAPKs do not simply balance each other in maintaining the integrity of the endothelium monolayer through effects on actin control. Actin realignment in ECs has been studied extensively in other physiologically important situations, such as endothelial barrier function, which is compromised during atherogenesis, and EC migration required for wound healing (i.e. Endothelialization of stents). These events require SAPK-mediated actin remodeling and are discussed below in sections V and VI, respectively.

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IV. Actin Cytoskeleton-Induced Endothelial Barrier Function Is Mediated by Stress Kinase Activity Endothelial cell layers provide barriers between the blood and surrounding tissues (Bogatcheva and Verin, 2008; Komarova and Malik, 2010). Specific situations of inflammatory stress result in actin realignment as a critical step in modulation of endothelial barrier function (Bogatcheva and Verin, 2008; Capaldo and Nusrat, 2009; Fernandez-Borja et al., 2010). Relatively few studies identify roles for JNK in barrier remodeling despite significant information about the process, and much evidence for JNK modulation of gene expression. JNK activity does appear to be important for HUVEC responses to bFGF (through FGFR1) in the post-translational down-regulation of cell surface cadherin expression. Blocking JNK activity allows cadherins to return to the cell surface (Wu et al., 2008). The response of human coronary artery endothelial cells to TNF-α involves activation of JNK and NF-κB. TNF-α induces increased permeability by blocking mRNA and protein expression of tight junction proteins zonula occluden-1 and claudin-1 through activation of JNK and NF-κB. Human stanniocalcin blocks TNF-α-induced endothelial permeability (Chen et al., 2008). It blocks both of these TNF-α-induced signaling pathways and similar direct blocking of JNK also inhibited TNF-α-induced increased permeability (Chen et al., 2008). A specific test of whether TNF-α-induced JNK activity directly controlled actin realignment in pulmonary microvascular endothelial barrier function suggested that Rho kinase activation was responsible for actin realignment and JNK activation in response to TNF-α treatment (Mong et al., 2008). However, JNK was not an intermediate between Rho kinase and actin realignment (Mong et al., 2008). In addition, hypothermia blocked TNF-α-induced HUVEC barrier dysfunction while decreasing JNK and p38 activity. MKP-1 levels are increased by the hypothermia, and blocking MKP-1 induction inhibited the effects of hypothermia on TNF-αinduced barrier dysfunction. However, directly blocking JNK activity did not have an effect on the barrier dysfunction, while directly blocking p38 did prevent TNF-α-induced barrier dysfunction (Yang et al., 2010b). 4-hydroxy-2-noneal, produced in peroxide-treated cells, also caused changes in bovine lung endothelial barrier function and actin remodeling along with activation of ERK, JNK and p38. Specific inhibitors for each MAPK were able to significantly block both the loss of barrier function and actin remodeling (Usatyuk and Natarajan, 2004), suggesting that each MAPK might have some role. Together these results support JNK involvement in endothelial barrier changes, but indicate that JNK involvement in barrier dysfunction actin changes is observed in only some situations. While the evidence implicating JNK in some aspects of endothelial barrier dysfunction is limited, many studies link p38 activity to direct modulation of actin in barrier dysfunction. Induction of p38 activity in porcine aortic endothelial cells by TGF-β activation of a Racmediated pathway (Varon et al., 2008) and in HUVEC by VCAM-1 mediated activation through Rac (van Wetering et al., 2003) resulted in stress fiber formation, contraction, and intercellular gaps. In both studies, p38 activity was required for the stress fiber formation. Similar studies in microvascular ECs where ICAM was engaged resulted in activation of p38 and signaling through HSP27 to actin filament formation (Wang and Doerschuk, 2001; Wang et al., 2005). Thrombin-induced permeability increases (Borbiev et al., 2004), angiotensin II-

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induced permeability increases (Yang et al., 2010a), modulated barrier function induced by serum from burn patients (Chu et al., 2010), melanoma cell interaction-induced barrier changes (Khanna et al., 2010), mechanical ventilation-activated barrier dysfunction (Damarla et al., 2009), and hypoxia-induced changes (Liu et al., 2009) all involve p38 activity as a necessary component of the signaling pathway to actin rearrangements. Barrier structures differ between different EC phenotypes in different vessels and locations (e.g. Prasain and Stevens, 2009). Differences in modulation of endothelial barriers between different endothelial cell types also exist (Cai et al., 2008). However, a shared dependence on the p38 stress kinase for actin changes and endothelial barrier decreases is solid evidence that stress kinase activity is intimately tied to modulation of actin filament dynamics and resulting control of cell physiology.

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V. Actin Cytoskeleton-Induced Endothelial Cell Migration Is Mediated by Stress Kinase Activity Endothelial migration is important in angiogenesis (during development and in response to specific signals generated in cancer, wound healing, and other specific situations) and in repair of endothelial denudation after injury. Angiogenesis and endothelial migration involve modulation of actin structures. Evidence for the involvement of stress kinases in these processes has been published, and the data indicate stress kinase activities that appear to vary by EC type, matrix type and assay system. Wounded endothelial layers typically begin migration and proliferation without special treatments. Scratch wounded cultures of bovine large artery ECs activate cells at the edge of the wound to begin migration. These cells exhibit active JNK associated with their actin fibers as do proliferating sub-confluent cultures of the cells, while active p38 is associated with focal adhesions (Hamel et al., 2006). The JNK association was stable to actin filament isolation procedures suggesting involvement of signaling through JNK to or from the stress fibers. Migrating human dermal microvascular ECs forced to express constitutively active ALK1 (activin receptor-like kinase, a TGF β receptor family member in ECs) are less migratory (David et al., 2007). The silencing of endogenous ALK1 increases migration which can be blocked by blocking JNK activity. Conversely, the active ALK1 construct decreases wound-induced JNK activity. Expression of dominant negative JNK also completely blocks the cell migration (David et al., 2007). Migration of wounded human pulmonary artery ECs, and the use of various caldesmon constructs suggested that caldesmon was important for cell migration. p38 phosphorylation of caldesmon was critical for actin remodeling and migration of these cells, as evidenced by effects of specific p38 inhibitors (Mirzapoiazova et al., 2005). Similar inhibition of ERK, also known to phosphorylate caldesmon, was ineffective. Thus, EC migration in wound repair appears to involve JNK and/or p38 activity associated with actin fibers. VEGF is often cited as required for induction of both new vessel formation and as inducing migration of ECs. In response to VEGF, the MAPKs ERK, JNK, and p38 are typically activated (Kinney et al., 2008; Liao et al., 2010), though the time frames for each, durations of activation, and outcomes vary over a wide range of conditions and between endothelial tissues and species. For example, migration of placental artery ECs induced by

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VEGF could be blocked by inhibition of Src, PI3K or JNK activity. Each of these inhibitors suppressed VEGF induced cofilin phosphorylation and actin stress fiber formation (Liao et al., 2010). In these studies, inhibition of p38 activity was not effective in blocking actin stress fiber formation. In human microvascular ECs, VEGF induced JNK activity, and the JNK induced tube formation and migration could be decreased by natriuretic peptides (Pedram et al., 2001). In murine aortic ECs, VEGF induces MKP-1 synthesis which is required for cell migration, and in turn requires JNK activity for its induction indicating a critical balance for JNK activation and inactivation in migration (Kinney et al., 2008). HUVECs also respond to VEGF by expression of DUSP1 (MKP-1) and DUSP5. However in the HUVECs, the DUSP1 modulates p38 activity which is critical for migration (Bellou et al., 2009). Again in HUVECs, VEGFA induces p38-mediated phosphorylation of LIMK which phosphorylates cofilin (Kobayashi et al., 2006) and annexin 1 (Côté et al., 2010) thereby decreasing actin dynamics and facilitating cell migration and tube formation. In addition, in HUVECs, VEGF induced p38 activation resulted in the activation of the actin regulator HSP27. In this particular situation, the involvement of JNK was ruled out (McMullen et al., 2004; Rousseau et al., 1997). Interestingly, VEGF induction of HUVEC migration often involved p38-induced actin changes, while in other types of ECs noted above, VEGF-induced migration and actin rearrangements were reported to involve JNK activity. Quite a few molecules besides VEGF enhance migration of various types of ECs and require p38 stress kinase activity for migration to occur. Several of these molecules also function through p38-induced actin rearrangements. For example, sesamin is a natural antioxidant that prevents endothelial dysfunction. It enhances neovascularization in animal models though multiple angiogenic signal modulators. Inhibition of p38 activity inhibited sesamin-induced HUVEC migration (Chung et al., 2010). Hydrogen sulfide can modulate growth of HUVECs seeded on matrigel thereby stimulating angiogenesis-like activity. Blocking p38 or the downstream HSP27 both blocked cell migration (Papapetropoulos et al., 2009). Sphingosine 1 phosphate induces migration of human aortic ECs along with ERK and p38 activation. Blocking p38 activity blocked migration of the cells in a Boyden chamber assay (Kimura et al., 2000). As in the VEGF studies, there is also significant evidence for JNK-induced actin changes and modulation of endothelial migration in response to other agents. Examples include the cytokine fractaline, plentiful in rheumatoid arthritis, which induces activation of JNK and ERK and migration of HUVEC and human microvascular ECs. Migration can be retarded by blocking either JNK or ERK activity (Volin et al., 2010). Hepatocyte growth factor induces migration of brain microvascular ECs, a property that requires JNK and ERK activity (Rush et al., 2007). Endothelin also induces migration of brain microvascular ECs. Blocking JNK activity blocks endothelin-induced migration, and blocking p38 activity partly decreases migration suggesting the two stress kinases might play different roles in endothelin-induced migration (Milan et al., 2006). Several types of ECs migrate in response to the cytokine SDF1, and microvascular endothelial cells form tubes as well. This requires specific activation of JNK3 which is facilitated by nitrosylation and inhibition of MKP7 (Pi et al., 2009). ADP also stimulates HUVEC migration while activating JNK, p38, and ERK though the P2Y1 receptor. Specifically blocking JNK activity inhibited migration, while blocking p38 only partly blocked cell migration (Shen and DiCorleto, 2008). Angiogenesis involves tube formation as well as migration, and this occurs effectively in vivo under conditions where the matrix is permissive. In matrigel, lung ECs align into tubes, a

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process that can be blocked by blocking JNK, or partly decreased by blocking p38 activity (Medhora et al., 2008). Induction of MKP-1 (which inactivates both p38 and JNK) blocks tube formation in HUVECs, but induction of MKP-3 which only inactivates ERK does not (Medhora et al., 2008). Most likely, differences in EC types, matrix employed, and culture conditions can all modulate the importance of JNK and p38 for migration. The studies implicating dual specificity phosphatase functions in endothelial migration further indicate that fine-tuning of the stress kinase activity is critical for the final outcome. Together the data indicate that both p38 and JNK activities play roles in modulating actin fiber reorganization that is involved in endothelial migration.

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Conclusion Blood flow exerts different magnitudes of fluid shear stress on endothelial cells, playing a crucial role in the localization of atherosclerotic lesions. Endothelial cells can distinguish differences in FSS, and adopt different morphologies driven by actin cytoskelton remodeling as a response to these stimuli. In order for these actin adaptations to occur, cells need to translate their responses to FSS (mechanotransduction), into biochemical signals. The stressactivated protein kinases, JNK and p38, mediate FSS mechanotransduction into differential actin adaptation, which occurs in 3 phases under high FSS conditions. Stress-activated protein kinases, JNK and p38, also mediate signaling to and/or from the actin cytoskeleton. These signals play roles in modulating endothelial barrier function and EC migration. ECs in different parts of the artery have different phenotypes and vary in migration which may be controlled through their differential actin adaptations. Athero-prone regions have decreased endothelial barrier function, which is dependent, in part, on p38-mediated actin phenotype. The time-dependent phasic actin adaptations to high FSS treatments seen in vitro do not typically occur in vivo. Endothelial cells found in regions of arterial bends and bifurcations are constantly exposed to lower FSS, and cells lining straight arteries are constantly exposed to higher FSS conditions. Understanding the control mechanisms involved in actin remodeling may be useful for several applications. Endothelialization issues, which involve the laying down of new ECs on stent implants, the low FSS regions, which are atheroprone, where EC turnover is high, or vascular development where new endothelial cells are forming, are a few applications where endothelial cells experience a temporal gradient of flow, and therefore undergo time-dependent actin remodeling. About 20% of stents that are used to keep blocked or narrowed arteries open result in restenosis, which is the re-narrowing of the artery (Hoffman et al., 1996; Spanos et al., 2003). When stents are introduced into arteries, the endothelium is often denuded, which leads to smooth muscle cell proliferation that thickens the intima, and these regions also become prone to thrombosis (Luscher and Tshuci, 1993). Endothelial cells control vascular tone, thrombosis, inflammation and cell proliferation, so the endothelialization of stents is very important. One way that problem is being addressed is by coating stents with growth factors and/or other growth-facilitating molecules to promote EC growth (Asahara et al., 1995; Van Belle et al., 1997). The stent repopulating ECs will still be exposed to the low FSS conditions present in treated regions, making it critical that FSS issues are also addressed. EC barrier function and migration

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patterns will also be dictated by their location. Understanding the role of SAPK signaling in FSS-induced actin remodeling, endothelial barrier function, and EC migration could lead to drug design that targets these pathways in the treatment of atherosclerosis.

Acknowledgements Research from the LJL-K laboratory presented in the chapter was initiated with support from the Biosystems Dynamics Summer Institute, funded through an HHMI undergraduate education grant to Lehigh University. Ongoing research in the lab is funded by NIH award HL54269 to LJL-K. The authors also acknowledge Hannah Brotzman for her involvement in FSS studies and thank Elizabeth Miller and Timothy Krentz for critical reading of the manuscript.

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Wang Q, Doerschuk C. 2001. The p38 mitogen-activated protein kinase mediates cytoskeletal remodeling in pulmonary microvascular endothelial cells upon intracellular adhesion molecule-1 ligation. Journal of Immunology 166:6877-6884. Wang Q, Yerukhimovich M, Gaarde W, Popoff I, Doerschuk C. 2005. MKK3 and -6dependent activation of p38 MAP kinase is required for cytoskeletal changes in pulmonary microvascular endothelial cells induced by ICAM-1 ligation. American Journal of Physiology - Lung Cell Molecular Physiology 288:L359-L369. Wang X, Abdel-Rahman A. 2002. Estrogen modulation of eNOS activity and its association with caveolin-3 and calmodulin in rat hearts. American Journal of Physiology - Heart and Circulatory Physiology 282(6):H2309-2315. Wang Y, Miao H, Li S, Chen K, Li Y, Yuan S, Shyy JY, Chien S. 2002. Interplay between integrins and FLK-1 in shear stress-induced signaling. American Journal of Physiology Cell Physiology 283:C1540-C1547. Weber K, Groeschel-Stewart U. 1974. Antibody to myosin: the specific visualization of myosin-containing filaments in nonmuscle cells. Proceedings National Academy of Sciences, USA 71(11):4561-4564. Wechezak A, Viggers R, Sauvage L. 1985. Fibronectin and F-actin redistribution in cultured endothelial cells exposed to shear stress. Laboratory Investigation 53(6):639-647. Weinbaum S, Zhang X, Han Y, Vink H, Cowin S. 2003. Mechanotransduction and flow across the endothelial glycocalyx. Proceedings National Academy of Sciences, USA 100:7988-7995. Wu J, Yan H, Chen W, Chen W, Wang C, Chi Y, Kao W. 2008. JNK signaling pathway is required for bFGF-mediated surface cadherin downregulation on HUVEC. Experimental Cell Research 314:421-429. Yamamoto K, Korenaga R, Kamiya A, Ando J. 2000. Fluid shear stress activates Ca(2+) influx into human endothelial cells via P2X4 purinoceptors. Circulation research 87:385391. Yang C, Patel K, Harding P, Sorokin A, Glass WF, II. 2007. Regulation of TGF-1/MAPKmediated PAI-1 gene expression by the actin cytoskeleton in human mesangial cells. Experimental Cell Research 313:1240-1250. Yang D, Liu J, Tian C, Zeng Y, Zheng Y, Fang Q, Li H. 2010a. Epigallocatechin gallate inhibits angiotensin II-induced endothelial barrier dysfunction via inhibition of the p38 MAPK/HSP27 pathway. Acta Pharmacologica Sinica 31:1401-1406. Yang D, Xie P, Guo S, Li H. 2010b. Induction of MAPK phosphatase-1 by hypothermia inhibits TNF-α-induced endothelial barrier dysfunction and apoptosis. Cardiovascular Research 85:520-529. Yoshizumi M, Abe J, Tsuchiya K, Berk BC, Tamaki T. 2003. Stress and vascular responses: atheroprotective effect of laminar fluid shear stress in endothelial cells: possible role of mitogen-activated protein kinases. Journal of Pharmacological Sciences 91:172-176. Yu J, Bergaya S, Murata T, Alp I, Bauer M, Lin M, Drab M, Kurzchalia T, Stan R, Sessa W. 2006. Direct evidence for the role of caveolin-1 and caveolae in mechanotransduction and remodeling of blood vessels. Journal of Clinical Investigation 116:1284-1291.

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In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 207-227

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter VI

The Molecular Mechanisms of Actin Regulatory Proteins Barak Reicher and Mira Barda-Saad The Mina and Everard Goodman Faculty of Life Sciences, Bar-Ilan University, Ramat-Gan 52900, Israel.

Abstract

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Actin polymerization is the driving force behind multiple cellular processes including proliferation, motility, adhesion and endocytosis, providing the infrastructure for structural cellular remodeling and intracellular signal transduction. The variety and flexibility of cellular function is achieved by the formation of diverse actin structures. de novo formation of actin filaments is controlled by a combination of G-actin binding proteins, actin severing proteins, and capping factors. In order to overcome these regulatory checkpoints and support efficient actin polymerization, the orchestrated activity of various actin elongation and nucleation proteins, such as formins and the ARP2/3 complex is required. This delicate balance between actin polymerization and de-polymerization must be highly controlled to ensure the formation of the correct actin structures at the correct place and time. The highly controlled actin polymerization process is enabled by the activity of actin nucleation promoting factors (NPFs). These proteins are present in molecular complexes that associate with actin nucleation proteins and are involved in their stabilization and activation at actin-rich sites. These complexes work in concert to control the spatial and temporal formation of diverse actin structures. Recently, the development of cutting-edge imaging technologies have provided new insights into the regulatory mechanisms of the NPFs, and their modes of activation, dynamics and association with the actin nucleation proteins.



Correspondence to Dr. Mira Barda-Saad, Faculty of Life Sciences, Bar-Ilan University, Ramat-Gan 52900, Israel. Tel.: +972-3-5317311. Fax: +972-3-7384058. Email: [email protected]

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Barak Reicher and Mira Barda-Saad In this review, we describe the activation mechanisms of actin regulatory proteins, their differential and overlapping functions in actin-dependent processes and structures, and their role in health and disease.

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Glossary Abi: Abelson tyrosine kinase (ABL)-interacting adaptor proteins. Part of the WAVE regulatory complex, and binds the Abelson tyrosine kinase. ARP2/3 complex: A complex composed of seven proteins, including ARP2, ARP3 and ARP complex protein 1 (ARPC1)–ARPC5. On its own, the complex has little activity but, when bound to an NPF, it is activated to promote branched actin elongation, nucleated from pre-existing filaments. Calpain: A group of Ca2+-activated cytoplasmic cysteine proteases that are found in many tissues and that hydrolyze various endogenous proteins, including cytoskeletal proteins. Capping proteins: Ubiquitously expressed proteins in eukaryotic cells, that are able to bind the barbed ends of actin filaments, thereby preventing both association of actin monomers and dissociation of filaments. Coiled coil: A structural domain that consists of two or more helices that twist around each other to form a stable, supercoiled structure; such domains can mediate protein oligomerization. FAM21: A family of four homologues. The FAM21 family members are highly phosphorylated early endosomal membrane proteins that function as components of the WASH complex. F-BAR: A dimerization domain with affinity for negatively charged lipids. The F-BAR dimer allows proteins containing them to sense a particular membrane curvature or to deform membranes to a defined curvature imposed by its structure. Filopodia: Finger-like cellular extensions composed of unbranched F-actin that elongate to drive membrane protrusions. Formins: A group of multidomain actin- nucleating proteins, that function as dimers to assemble unbranched actin filaments. Formins generate actin polymerization nuclei by stabilizing actin dimers through their homodimeric formin homology2 (FH2) domains. Guanine nucleotide exchange factor (GEF): A protein that activates GTPases by catalyzing exchange of GDP for GTP. HSPC300: Hematopoietic stem progenitor cell-300. Part of the WAVE regulatory complex. Insulin receptor substrate protein of 53 kDa (IRSp53): A membrane bound protein, which belongs to the RCB (Rac binding) domain-containing protein family, a class of proteins with similar properties to that of the EFC/F-BAR proteins. IRSp53 contains an SH3 domain that binds the proline-rich domain of mammalian WAVEs and simultaneously binds to activated Rac, which contributes to the Rac-dependent localization of WAVE. Invadopodia: Large membrane protrusions produced by invasive cells that extend into the extracellular matrix and are rich in F-actin and matrix-degrading proteolytic proteins. Lamellipodia: Sheet-like cellular extensions composed of a branched F-actin meshwork, which elongates to drive membrane protrusion. Lipid raft: A microdomain of the plasma membrane, enriched with cholesterol and sphingolipids that allows compartmentalization of various signaling molecules.

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Matrix metalloproteinases (MMPs): Zinc-dependent endopeptidases capable of degrading many types of extracellular matrix proteins. MMPs play a major role in cellular invasion, are found in invadopodia, and contribute to metastasis. Nap1: Nck-associated protein-1. Part of the WAVE regulatory complex, and binds the adaptor protein, Nck. Nck: Noncatalytic region of tyrosine kinase. Ubiquitously expressed, SH2 and SH3 domaincontaining adapter proteins. Nck proteins (Nck and Nck) connect receptor and nonreceptor tyrosine kinases to the machinery of actin polymerization through binding to actin regulatory proteins, e.g., WASp or WAVE. Podosome: An adhesion structure of small (0.5 μm) diameter that is found in various malignant cells and in some normal cells, including macrophages. Podosomes are comprised of an actin core surrounded by a ring containing typical focal-adhesion proteins, such as vinculin and paxillin. Profilin: A ubiquitously expressed actin-binding protein that is present in all eukaryotic organisms. Most profilin in the cell is bound to actin and is important for actin cytoskeleton dynamics by binding proline-rich domains of cytoskeletal proteins and putting them in proximity to pools of G-actin. Pseudopodia: Large cellular extensions that contain a network of F-actin, which mediate the protrusion of the leading edge of an amoeboid cell or a phagocyte during crawling migration. Rho GTPase: A conserved family of small enzymes that convert GTP to GDP and act as a „molecular switch‟ that is active in the GTP-bound form and inactive in the GDP-bound form. Active Rho proteins bind to effectors, such as NPFs, to trigger cytoskeletal remodeling. Sra: Specifically-Rac1 associated protein-1. Part of the WAVE regulatory complex; forms a heterodimer with Nap1. Src homology 2 domain (SH2): A protein domain, present on intracellular signal-transducing proteins that commonly binds phosphotyrosine residues. Src homology 3 domain (SH3): A protein domain, present on intracellular signal-transducing proteins that commonly binds polyproline motifs. Tandem-monomer-binding nucleators: A group of actin nucleators characterized by tandem G-actin- binding motifs, which bring together monomers to form a polymerization seed. Transducer of Cdc42 activity-1 (TOCA-1): A membrane-associated protein that belongs to the F-BAR domain-containing family of proteins. TOCA-1 binds both to N-WASp and to the Rho GTPase, Cdc42, which contributes to the Cdc42-dependent localization and activation of N-WASp. WASp interacting protein (WIP): An actin-binding adaptor protein that is found in a stable complex with WASp, regulating its function and protecting it from degradation. WIP also binds the G-actin-binding protein profilin and provides a pool of G-actin for WASpmediated polymerization. Wiskott-Aldrich syndrome (WAS): An X- linked recessive disease characterized by immunodeficiency, thrombocytopenia and eczema. Untreated, most patients die from recurrent infections, bleeding or autoimmune disease. Genetic deficiency of WASp was identified as the causative defect in WAS.

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Introduction The actin cytoskeleton is deeply involved in a variety of cellular processes, including migration, adhesion, intracellular trafficking, cell polarity, cell division, endocytosis, and the formation of distinct cellular morphologies. These processes depend on the rapid assembly of filamentous actin (F-actin), which provides the basis for structural cellular remodeling and intracellular signal transduction [1, 2, 3]. The diversity and flexibility of cellular function are achieved by the formation of different actin structures, i.e. filopodia and lamellipodia for motility, podosomes for adhesion, and invadopodia for tissue invasion. Actin is one of the most ubiquitous and conserved proteins in eukaryotic cells, which contain a large pool of globular monomeric actin (G-actin). However these actin monomers are mostly bound by G-actin-binding proteins such as profilin, which, along with actin severing and capping proteins, suppress the spontaneous nucleation and elongation of new actin filaments [4]. Thus, in order to support efficient actin polymerization, eukaryotic cells harness the activity of various actin elongation and nucleation proteins, such as Formins and the actinrelated protein 2/3 (ARP2/3) complex [5, 6]. These proteins enable actin polymerization by initially stabilizing actin dimers or trimers, creating a "seed" of actin. Then, elongation continues from the existing seed. The activity of these actin nucleation and elongation factors must be highly controlled, both spatially and temporally, to ensure the formation of proper actin structures. Furthermore, actin nucleation proteins such as ARP2/3 are by themselves inefficient nucleators and require other proteins to be efficiently activated. Thus, another set of proteins, known as actin nucleation promoting factors (NPFs), has evolved in eukaryotic cells. These proteins associate with actin nucleation proteins and are involved in their stabilization and activation at actin rich sites. In their native form, NPFs never function alone, but are associated with other proteins to form dynamic molecular complexes. These molecular complexes regulate the activity of NPFs, recruit them towards specific sub-cellular locations, and control their association with the actin machinery. The proteins in the NPF complex also stabilize the NPFs and protect them from degradation [7-12]. Not surprisingly, NPFs were found to be involved in a variety of pathological conditions, ranging from immunodeficiency (e.g. Wiskott - Aldrich syndrome; WAS) to cancer [13-16]. In this review, we will focus on the activation mechanisms of NPFs, their differential and overlapping functions in actin-dependent processes and structures, and their role in health and disease.

Actin Nucleators: Sowing the Seeds of Actin Spontaneous F-actin polymerization involves the formation of highly unstable polymerization intermediates that are short-lived and rapidly dissociated, making spontaneous polymerization kinetically unfavorable. However, once a stable trimeric nucleus of actin is formed, filaments elongate at their fast-growing ('+' or 'barbed') ends at a rate linearly proportional to the concentration of available actin monomers. Filament elongation at the

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slower-growing ('-' or 'pointed') ends is blocked by proteins that are bound to these ends, making such a potential process physiologically irrelevant. Additionally, the vast majority of G-actin in eukaryotic cells is bound to profilin, which raises another obstacle to spontaneous nucleation. This makes actin nucleation the rate-limiting step in the process of actin polymerization. Therefore, actin nucleation proteins catalyze this step, enabling efficient actin polymerization [5]. So far, three classes of nucleation proteins have been identified: the ARP2/3 complex, the formins, and the tandem-monomer-binding nucleators (Cordon-bleu (Cobl), Leiomodin (Lmod), and Spire). The ARP2/3 complex facilitates the polymerization of branched F-actin, while the other two classes drive the polymerization and elongation of linear, unbranched Factin forms. Formins are dimeric, multidomain proteins that are characterized by their formin homology (FH) domains, FH1 and FH2. The interaction between two FH2 domains creates a "doughnut shape" dimer that catalyzes F-actin nucleation, while the FH1 domain binds profilin-bound G-actin, increasing its effective concentration. In addition to their actin nucleation activity, formins function as actin elongation factors that associate with growing barbed ends and thereby prevent polymerization termination by capping proteins [6]. As the most recently identified actin nucleators, much less is known about the molecular mechanisms of actin nucleation by the tandem-monomer-binding nucleators. These nucleators share little structural homology; however, they share a common feature of multiple actin binding motifs that allow them to promote actin nucleation [17]. Each class of nucleators uses a different mechanism to catalyze nucleation: ARP2/3 structurally mimics the polymerization intermediates, formins associate strongly with spontaneously formed actin dimers, and the tandem-monomer-binding nucleators bring together actin monomers through their clustered actin-binding motifs to form an actin nucleus. Formins and the tandem-monomer-binding nucleators have been reviewed in detail elsewhere [6, 17], and we focus here mainly on the ARP2/3 complex.

NPFs: Kick Starting the ARP2/3 Machine The first actin nucleator to be identified is ARP2/3, a 220 kDa complex of seven stably associated polypeptides. The complex includes a core of ARP2 and 3, plus five additional subunits, ARPC1–ARPC5. The ARP2/3 complex produces branched F-actin networks by initiating new actin filaments on the sides of preexisting filaments. These dendritic actin networks, created by ARP2/3, commonly form the structure of the lamellipodium [5]. However, a recent study contradicts this convention by showing, using electron tomography, that the organization of actin in the lamellipodia of vitreously frozen cells consists mainly of unbranched filaments [18]. Although disputed by this recent study, branched actin polymerization by ARP2/3 still remains the canonical mode of action of this complex. In order to support stable actin nucleation, the ARP2 and ARP3 subunits mimic an actin dimer and act as a template for the formation of the branch-point of F-actin. Alone, the ARP2/3 complex is an inefficient nucleator, requiring the binding of actin filaments to increase its activity and couple it with nucleation and branching. This coupling of actin binding with nucleation and branching is mediated by NPFs.

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Figure 1. Suggested molecular mechanisms of NPFs activation. In resting cells (Left), NPFs are either autoinhibited (WASp) or trans-inhibited (WAVE, WASH, and presumably WHAMM). Following cellular activation, multiple signals contribute to the release of the inhibition (Right): (A) WASp GBD domain binds the GTP bound Rho GTPase Cdc42, which releases the autoinhibitory interaction between the VCA domains and the GBD, allowing VCA to activate the ARP2/3 complex to facilitate branched actin polymerization. Additional phosphorylation by Src kinases on tyrosine 291 stabilizes the active conformation of WASp. WASp associates to TOCA-1 which, along with the SH3 domaincontaining adaptors Grb-2, Nck and the acidic phospholipid PIP2, recruits WASp to the plasma membrane. (B) WAVE is trans-inhibited by the members of the WAVE regulatory complex (WRC) Nap1, Sra1, Abi and HSPC300. Its VCA domain is sequestered within a niche in Sra1 and also connected to a part of its N-terminus. Three main signals release WAVE from its inhibition: (1) The

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binding of the GTP bound Rho GTPase Rac1 to a conserved patch in Sra1; (2) phosphorylation of WAVE by the c-Abl kinase and other kinases; and (3) binding of PIP3, which -along with IRSp53mediates the recruitment of WAVE to the membrane and its activation. (C) WASH is also transinhibited by the members of its own regulatory complex (SHRC), comprised of Strumpellin, SWIP, FAM21, and CCDC53. Although the SHRC is genetically and structurally related to the WRC, its activation mechanisms are still unknown, and might include activation by the Rho GTPase, Rho1. In addition to actin polymerization by its VCA domains, the tubulin binding region of WASH (TBR) binds the microtubule cytoskeleton. (D) WHAMM does not seem to be autoinhibited and is probably trans-inhibited by factors that interact with its N-terminal WHAMM membrane interaction domain (WMD) and/or its coiled-coil (CC) regions, and that also interact with the microtubule cytoskeleton. (E) The mechanisms of JMY inhibition and activation are unknown. In addition to its ARP2/3 dependent branched actin polymerization activity, by its VCA domains, JMY functions as a tandem-monomer actin nucleator through its three V domains, which facilitate the nucleation of unbranched, linear Factin. Additionally, JMY function as a transcription co-factor through the binding of the transcription factor CBP/p300 at its N-terminus. Question marks indicate unknown factors or speculative mechanisms of activation. WASp, Wiskott-Aldrich syndrome protein; GBD, GTPase binding domain; V, Verprolin homology; C, cofilin homology; A, acidic domain; PRD/P, proline rich domain; WH1, WASp homology1; B, basic region; WIP, WASp interacting protein; TOCA-1, transducer of Cdc42dependent actin assembly; PIP2, PtdIns-4,5-bisphosphate; WAVE, WASp family-verprolin homologous protein; WRC, WAVE regulatory complex; WHD, WAVE homology domain; Abi, Abelson tyrosine kinase (ABL)-interacting adaptor proteins; Nap1, Nck-associated protein-1; Sra, specifically-Rac1 associated protein-1; HSPC300, hematopoietic stem progenitor cell-300; PIP3, phosphatidylinositol-3,4,5-trisphosphate; IRSp53, insulin receptor substrate protein of 53 kDa; WASH, Wiskott–Aldrich syndrome protein and SCAR homologue; SHRC, WASH regulatory complex; WAHD1, WASH homology domain 1; TBR, tubulin-binding region; CP, capping protein ; WHAMM, WASp homologue associated with actin, membranes and microtubules; WMD, WHAMM membrane-interacting domain; JMY, junction-mediating regulatory protein; L, linker; F-actin, filamentous actin; G-actin, globular actin.

When activated by NPFs, the ARP2/3 complex initiates the formation of a new actin filament that emerges from an existing filament, resulting in a typical branched Y-shape, with a branch angle of ~70o. The mechanism of ARP2/3 activation by the NPFs lies in the common modular structure of NPFs; the largest group of NPFs activate the ARP2/3 complex using their C-terminal VCA domains (Figure 1). All of these proteins contain a VCA (also termed WCA) domain, which consists of a verprolin-homology (V) domain, a cofilin-homology (C; also termed central or connector) domain, and an acidic (A) domain. These domains act together to bind the ARP2/3 complex, activate it and supply it with actin monomers for nucleation. In the current model of ARP2/3 activation by NPFs, the V domain delivers an actin monomer to the complex, and the C and the A domains bind the complex, bringing together the ARP2 and ARP3 subunits to an active conformation, facilitating the nucleation of new actin filament on the branch point of the mother filament [19].

Controlling the Controllers: Regulatory Mechanisms of the NPFs There are currently eight known eukaryotic NPFs (Figure 1). The Wiskott-Aldrich Syndrome protein (WASp) was the first to be discovered and is exclusively expressed in hematopoietic cells. The WASp homologue, neuronal-WASp (N-WASp) was initially

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discovered in the central nervous system (CNS), but is expressed in most tissues. WASp and N-WASp localize primarily to the plasma membrane, where they enhance formation of lamellipodia and filopodia, and are involved in endocytosis. The WASp family-Verprolin homologous protein (WAVE) is also widely expressed. This sub-family of NPFs is comprised of three members (WAVE 1-3) [20]. The WAVEs localize mainly to the plasma membrane, to support cellular motility by creating lamellipodia. Recently, three novel NPFs were identified: The Wiskott–Aldrich syndrome protein and SCAR homolog (WASH) [12], WASp homolog associated with actin, membranes and microtubules (WHAMM) [21], and junction mediating regulatory protein (JMY) [22]. Despite their similar modular structure based on the C-terminal VCA domain, different NPFs possess distinct actin-related cellular activities. These proteins have different modes of activation and regulation; their expression is sometimes tissue specific; they are recruited to a diversity of sub-cellular locations and interact with different proteins. The question is: what makes them so different? To answer this question, we must first look at the structure of NPFs. As mentioned above, all of these proteins contain a C-terminal VCA domain. However, they mainly differ in their N-termini and other domains. These differences give them their distinct characteristics.

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Distinct NPF Domains Are Important for Their Stability, Function and Cellular Localization WASp and N-WASp are characterized by their WASp-homology 1 (WH1) domains that are known to interact with the WASp-interacting protein (WIP), or alternatively, with its homologous WICH/WIRE and CR16. This interaction, mediated by the WH1 domain, was shown to control WASp activity, dynamics and stability [7, 23, 24]. An example of the importance of the WASp-WIP interaction is the Wiskott - Aldrich syndrome (WAS). Genetic deficiency of WASp was identified as the causative defect in WAS. This X-linked recessive disease is characterized by immunodeficiency, thrombocytopenia and eczema. Untreated, most patients die by recurrent infections, bleeding or autoimmune disease. Cells from WAS patients are characterized by their relatively smooth surface, due to profound defects in their ability to create actin structures such as filopodia and lamellipodia [13, 14]. The vast majority of WAS mutations map to the WIP binding region and lead to a decrease or a complete absence of WASp protein expression [25-27]. Moreover, in resting T-cells, over 95% of WASp is found in complex with WIP [24]. These findings hinted at a protective role for WIP on WASp, and indeed, several studies showed that WIP protects WASp from degradation [7, 23, 28]. Recently, Massaad et al. showed that overexpression of WIP, or even of the minimal WASp binding region of WIP (nWIP), restored WASp levels to normal in B-cells from XLT (a mild version of WAS) patients, and corrected the defective actin polymerization of their T cells in response to anti-CD3 stimulation [29]. These data demonstrate the pivotal role of the WASp-WIP interaction for WASp stability, mediated by its unique WH1 domain. In addition to its protective role, WIP was also suggested to inhibit WASp activation [24, 30], though a matter of debate [31], and to recruit WASp to areas of actin assembly in T-cells [24]. In addition to the WH1 domain, WASp contains a proline rich domain (PRD]. This domain binds Src-homology 3 (SH3)

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domain- containing proteins, which recruit WASp to the membrane, where actin polymerization takes place (see below). The other NPFs also have distinct N-terminal domains. The WAVE proteins and the WASH proteins are characterized by their WAVE homology domains (WHDs) and WASH homology domain 1 (WAHD1), respectively. The WHD of WAVE was shown to be the main site of interaction with the other proteins in the WAVE regulatory complex (WRC) [32, 33]. This pentameric hetero-complex, also comprising Nap1 (HEM1), Sra1 (PIR121), Abi and HSPC300, regulates WAVE stability, activation and localization (as will be discussed later). Like WASp, WAVE also contains a PRD. This domain binds to the SH3 domains of the membrane binding protein insulin receptor substrate IRSp53. This interaction contributes to the recruitment of WAVE to the plasma membrane, linking WAVE with its activator, the Rho GTPase, Rac1 [34-36]. Interestingly, WASH was also found to stably interact with four other proteins, through its WAHD1, to form the WASH regulatory complex (SHRC) [34]. Biochemical and bioinformatic analysis, along with electron microscopy revealed that this complex is structurally related to the WRC, and it has been suggested to function analogously to the WRC [37]. Therefore, it is assumed that WASH undergoes a similar mode of regulation to that of WAVE, though this assumption is a subject to extensive investigation. In addition to its WAHD1, WASH contains a unique tubulin binding region (TBR). Additionally, Drosophila WASH displays microtubule bundling activity [38]. Therefore, WASH may link the actin cytoskeleton with the microtubule cytoskeleton. However, further research is required in order to reveal the molecular mechanisms by which WASH links the two cytoskeletal arms. The two most recently identified NPFs, WHAMM and JMY have a sequence identity of nearly 35%, but differ more strongly from each other in their N-termini, which share only 25% sequence identity [22, 39]. Unlike WAVE and WASp (or even WASH), our current knowledge of WHAMM and JMY regulation is limited. The defining study on WHAMM characterized its N-terminal WHAMM membrane-interacting domain (WMD) as required for its recruitment and interaction with various intra-cellular membranes, mainly the cis-Golgi and the tubulovesicular ERGolgi intermediate compartment (ERGIC) [22]. This recruitment to the cis-Golgi and the ERGIC is unique to WHAMM, and potentially broadens the spectrum of ARP2/3 mediated actin related functions to include participation in ER-Golgi transport. Indeed, RNAi mediated silencing or overexpression of WHAMM resulted in deformation of the Golgi apparatus, and the depletion of WHAMM also impaired transport of the VSV virus G protein from the ER to the Golgi [22]. However, the mechanism of participation by WHAMM and/or the ARP2/3 complex in such transport under physiological conditions is yet to be determined. Similar to WASH, WHAMM has microtubule binding abilities. WHAMM binds microtubuli directly through its coiled-coil (CC) domain [22]. However, the functional relevance of this interaction is yet to be determined. JMY, the most recently discovered NPF, was initially identified as a cofactor of the transcriptional regulator p300/CBP, which upregulates the p53 response. DNA damage triggers its release from the Mdm2 ubiquitin ligase, resulting in nuclear accumulation [40-42]. The N-terminus of JMY then binds the p300/CBP transcriptional regulator to augment the p53 response. In addition to its unique function as a co-transcriptional regulator, JMY is the only NPF that has three actin-binding V domains (the other NPFs have 1-2 V domains) adjacent to the ARP2/3-binding CA domains [39, 42]. This unusual VCA configuration enables JMY to function both as an NPF for the ARP2/3 complex and as an ARP2/3-

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independent actin nucleator, by bringing together actin monomers through its clustered V domains to form an actin nucleus (as described for the tandem-monomer class of actin nucleators). This transcriptional co-activation, combined with JMY mediated ARP2/3 dependent and independent actin nucleation activity, suggests that JMY may integrate the p53 response and actin driven cellular motility.

Autoinhibition and Trans-Inhibition: An on/off Switch for the NPFs Another important layer of NPF regulation is their mode of activation. NPFs can be switched "on" and "off" in order to support proper spatial and temporal actin nucleation. The subject of this layer of regulation is usually the VCA domain. The accessibility of this domain to the ARP2/3 complex determines the NPF activation state. NPFs can be either autoinhibited through intramolecular interactions between their VCA and N-terminal domains as in WASp and N-WASp, or trans-inhibited by interacting proteins, as shown for the WAVEs, and recently for WASH.

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WASp and N-WASp Are Autoinhibited Apart from their distinctive WH1 domain, WASp and N-WASp differ structurally from the other NPFs in their basic region, and in their GTPase-binding domain (GBD), C-terminal to the WH1 domain. In resting cells, WASp is mainly present in an autoinhibited form in which its VCA domain interacts with a hydrophobic patch within the GBD (Figure 1), thereby masking the binding sites for the ARP2/3 complex [20]. Several coincident signals contribute to the activation of WASp: The Rho family GTPase, Cdc42, activated by the Rho GTPase exchange factor (GEF), VAV, and binds to the WASp GBD site. This binding, together with phosphorylation of WASp on tyrosine residue 291 by protein tyrosine kinase (PTK), induces a dramatic conformational change. The hydrophobic patch is then disrupted, releasing the VCA domain, enabling its interaction with the ARP2/3 complex, and thereby promoting actin polymerization. Another important factor that contributes to WASp activation is the membrane lipid phosphatidylinositol (4,5)-bisphosphate (PIP2). This factor was shown to bind a basic region adjacent to the GBD domain, and together with Cdc42 facilitate the release of the VCA domain [43-45]. As a membrane lipid, PIP2 not only activates WASp but can also restrict WASp-driven actin polymerization to or from the proximity of defined membrane domains such as lipid rafts and neuromuscular junctions, thereby spatially controlling actin polymerization [44, 46, 47]. Additional factors that contribute to WASp activation are SH3 domain-containing proteins, such as Nck [48, 49, 50] that bind to WASp PRD. However, the binding of these proteins to the PRD does not directly release the autoinhibition; instead, it enhances WASp interactions with its activators i.e. Cdc42 and PIP2, via recruitment of WASp to signaling complexes at the plasma membrane [50-53].

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The WASp interacting protein, WIP, provides another mode of regulation for WASp. Studies have shown that WIP is bound to WASp and inhibits its activity in vitro [51], however this is a metter of debate, since it was also demonstrated that WASp and WIP constitutively interact and WIP does not affect WASp function [52]. Unlike recombinant, purified WASp whose autoinhibition is relatively weak, the native WIP-WASp complex is much more strongly inhibited, and the addition of Cdc42 alone is insufficient for native WASp activation [30, 54]. It is not clear however, how WIP inhibits WASp. A possible explanation is that WIP stabilizes the autoinhibited conformation of WASp, though this assumption requires further investigation. In order to relieve WASp from WIP inhibition, additional activities are required, including WASp binding to the transducer of Cdc42 activity-1 (TOCA-1) protein [30]. This protein binds both Cdc42 and WASp, thereby enhancing WASp activation by Cdc42. Moreover, TOCA-1 has been shown to recruit the WIP-N-WASp complex to artificial tubular membrane structures of defined curvatures through its F-BAR domain [35]. The diameter of these tubular structures appeared to correspond to the diameter of clathrin-coated pits. Therefore, TOCA-1 may stimulate actin polymerization in a membrane curvature-dependent manner. Indeed, a recent study demonstrated that TOCA-1 interacts with N-WASp in vivo and induces the formation of dynamic membrane tubules and vesicles, in a Cdc42 dependent manner [55].

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Trans-Inhibition: The WAVE and WASH Regulatory Complexes The WAVE proteins represent a different mode of NPF regulation. In their native form, these proteins are part of a large ~400 kDa pentameric complex comprising WAVE, Nap1 (HEM), Sra1 (PIR121), Abi and HSPC300 (Figure 1). The components of this complex (also termed WAVE regulatory complex, WRC) are interdependent, stabilizing one another against degradation [9, 10, 56, 57]. Unlike WASp and N-WASp, purified WAVE is not autoinhibited. Instead, WAVE is trans-inhibited; its VCA domain is sequestered within the WRC, rendering it inaccessible to the ARP2/3 complex. The characteristics of WRC, the basal state of the WAVE complex, and its activation upon signaling have been a subject of debate. WAVE complexes were found to be basally inactive, however, their binding to Rac1 and Nck caused dissociation of the complex, which released active WAVE–HSPC300 and led to actin nucleation [58]. However, other studies showed that WAVE complex is basally phosphorylated and activated [59, 60]. These reports led to the suggestion that WAVE activation is mainly regulated by its localization. Using advanced purification and reconstitution techniques, recent studies have resolved most of the controversy about WAVE regulation, determining that the WAVE complex is intrinsically inactive and inhibited [11, 32, 61]; however, it is activated via integrated signals, as described below: Multiple signals contribute to the activation of WAVE. The best known activator of WAVE is the small GTPase, Rac1, which like Cdc42, can be activated by the GEF, VAV [62]. Through WAVE, Rac1 mediates the formation of membrane structures [10, 35, 63, 64]. Following activation, Rac1 binds the WRC through Sra1, thereby activating WAVE. An additional factor that contributes to WAVE activation is the binding of acidic phospholipids, mainly the Phosphatidylinositol 3-kinase product, Phosphatidylinositol (3,4,5)-triphosphate

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(PIP3). PIP3 was shown to directly bind the basic region of WAVE, to recruit the WAVE complex to the plasma membrane, and to cooperate with Rac1 and IRSp53 in its activation [32, 34, 65]. Phosphorylation provides another way to activate WAVE. Abi-1 of the WRC was shown to mediate Abelson (Abl) kinase-evoked phosphorylation of WAVE2 on tyrosine 150 [66]. This phosphorylation is required to couple WAVE2 and activated Rac1 with actin polymerization [67]. The Abl kinase was also found to phosphorylate and activate WAVE3, independently of Abi [68]. Phosphorylation of the WAVE2 complex on serine and/or threonine residues were shown to be required for activation of the complex by Rac1 and acidic phospholipids [32]. Although the key factors that activate the WAVE complex were identified, the molecular mechanism of this activation has been unclear. An early study suggested dissociation of the complex triggered by Rac1 and Nck binding [58]. Later studies ruled out this model by showing that the complex remains intact throughout the activation process [10, 32, 61, 66]. Recent analysis of the WRC crystal structure provided a more precise model for the molecular activation mechanism of WAVE [69]. Based on these new data, it appears that the WAVE1 VCA domain is bound by residues 82-184, to a conserved niche of Sra1, keeping it inaccessible and inactive. Rac1 binding to Sra1, in a position adjacent to the interface with the VCA, competes with the binding of the VCA to WAVE itself and to Sra1, thereby causing a conformational change that facilitates the release of the VCA. Interestingly, these data reflect a different mode of regulation for WAVE, in which WAVE is not only trans-inhibited, but also might be slightly auto-inhibited, resembling the mechanism of WASp regulation. WAVE complex activation by phosphorylation was also examined in this study. Two putative tyrosine residues, located at the interface between WAVE1 and Sra1, Tyr151 (equivalent to Tyr150 of WAVE2) and Tyr125, were mutated and the resulting protein was checked for lamellipodia formation. The expression of both constitutively active mutants led to upregulation of lamellipodia structure formation suggesting activation of the WRC by these two tyrosines [69]. As mentioned above, acidic phospholipids, mainly PIP3, recruit the WAVE complex to the cell membrane and synergize with Rac1 (among others ) to activate WAVE. In this recent study, mapping the surface distribution of charge of the entire complex, defined two opposite faces: a negatively charged face that includes the WAVE C-terminal, Abi and HSPC300, and a positively charged face that includes the WAVE basic region and the basic surface of the Nap1/Sra1 dimer. This polarized arrangement is proposed to orient the complex to bind the membrane through the interactions between the acidic phospholipids and positively charged face of the complex, exposing the negatively charged face (including the VCA domain) to the cytoplasm, where actin polymerization takes place [69]. Thus, based on these data, the current model suggests that in resting cells, the VCA domain is coiled, bound and sequestered within Sra1, making it inactive. Upon activation, Sra1 binds to activated Rac1, and the entire complex is recruited to the plasma membrane, where the VCA domain is released and associates with ARP2/3. The interactions between the positively charged face of the WRC and negatively charged phospholipids e.g. PIP3, reorients the WRC so that the VCA domain is exposed to the cytoplasm, where it can interact with the actin polymerization machinery. Like WAVE, WASH is also regulated as part of a stable complex (Figure 1). This ~700 kDa WASH regulatory complex is termed SHRC and composed of five proteins, including WASH, Strumpellin, KIAA1033 (also termed Strumpellin and WASH-interacting proteinSWIP), FAM21, and CCDC53 [33, 37]. Like the WRC, the SHRC components stabilize one

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another against degradation [12, 70]. Some studies suggested that the , capping protein dimer (CP) also be considered as part of the complex [70]. Interestingly, CP inhibits actin dynamics by binding the barbed end of the filament. Thus, CP binding to the SHRC, probably via the early endosomal membrane protein, FAM21 [37], may promote local F-actin polymerization by both recruitment to barbed ends, and attenuation of capping protein activity. Purified WASH is active, meaning that unlike WASp, it is not autoinhibited [12, 37, 70]. Nevertheless, the basal activation state of WASH is a matter of debate. Previously, it was found that the SHRC is constitutively active [70], while a recent study showed that highly purified recombinant SHRC has no detectable activity toward ARP2/3 complex [37]. Unlike WASp and WAVE, which are the effectors of Cdc42 and Rac1, respectively, WASH activators remain unknown. Nevertheless, studies in Drosophila demonstrated the ability of WASH to crosslink/bundle linear F-actin as an effector of Rho1 [38]. Interestingly, this effect was mutually exclusive with the binding of the ARP2/3 complex, which inhibited WASHdependent linear actin bundling, and promoted branched actin nucleation, suggesting ARP2/3 as a molecular switch between WASH-dependent linear and branched actin polymerization [38]. The mechanism by which WASH is regulated by the SHRC is still unknown. However, the structural and sequence resemblance between the WRC and SHRC, suggest a similar WAVE-like mechanism of regulation for WASH [37]. This proposed similarity, provides a foundation for further research that will address several questions: Is WASH regulated by phosphorylation, or binding to phospholipids? What controls WASH localization? What is the molecular mechanism of WASH inhibition? Much less is known of the regulatory mechanisms of WHAMM (Figure 1). Purified fulllength WHAMM and the VCA domain alone, have the same actin nucleation activity, suggesting that WHAMM is regulated by a mechanism other than autoinhibition. Accordingly, overexpression of WHAMM in mammalian cells induces ARP2/3 dependent actin polymerization. However, further research must be conducted to determine whether WHAMM is part of a regulatory complex, like WAVE and WASH, or whether it is regulated through a different mechanism(s).

NPF Degradation Mechanisms- Control of NPF Stability Unlike the vast knowledge we have regarding the positive control mechanisms of NPFs, i.e. factors that activate NPFs to support actin polymerization, the negative regulation mechanisms of NPFs are far from being clear. The unregulated activity of NPFs might lead to improper and/or exaggerated actin polymerization activity, resulting in increased cellular motility and the progression of malignancies or autoimmune diseases. In order to avoid cellular hyper-responsiveness, cells have developed several mechanisms for signaling attenuation, by controlling protein expression. Unsurprisingly, overexpression of cytoskeletal proteins was found in several malignancies (described in detail below) [71], though the mechanisms of their function and correlation between protein overexpression and pathogenesis of the disease are yet to be determined. Conversely, insufficient NPF expression, or NPF deficiency, might lead to diminished actin polymerization, resulting in cellular hypo-responsiveness, as in the case of the WAS immunodeficiency [14].

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For NPFs, protein degradation was found to control protein expression, stability and turnover. Several studies showed WASp to be a target of the calpain, cysteine protease and the proteasome [7, 23, 72, 73]. Furthermore, WIP, the binding partner of WASp, protects WASp from degradation, as WIP deficient cells show diminished levels of WASp [7, 8, 23, 28]. WASp degradation seems to occur following cellular activation; in platelets, WASp is degraded by calpain, following activation by either Ca2+ ionophore, thrombin, or collagen [72, 73]. A phospho-mimicking, constitutively active WASp mutant (Y293E), when expressed in dendritic cells of WASp-deficient mice, was unstable, undergoing spontaneous degradation [28]. These data suggest that the active conformation of WASp is a target for degradation. The observation that WASp degradation is coupled with its activation, further emphasize NPF degradation as a possible mechanism for signal attenuation. Previously, by using live imaging techniques, we showed that following T cell antigen receptor (TCR) activation, WASp is recruited to the membrane, where actin polymerization takes place. Late in the activation process, vesicles containing clusters of WASp and the adaptor protein SLP76 leave the membrane periphery and are endocytosed to a central structure, where they are degraded [52]. Therefore, in T lymphocytes, WASp degradation can serve as a mechanism for TCR signaling control (Reicher et al, in preparation). WAVE was also shown to be a target for degradation. Similar to WASp, all three isoforms of WAVE undergo calpain dependent degradation in activated platelets [74]. In Drosophila, Abi protects WAVE from proteasomal-dependent degradation. Nevertheless, the molecular mechanisms of these degradation reactions are not clear. Interestingly, in a study that monitored WAVE expression levels in macrophages over time under various culture conditions, WAVE1 expression levels decreased in adherent cells. This reduction was due to cleavage mediated by calpain, which removed the VCA domain of WAVE, abrogating its activity [75]. These results may suggest a role for WAVE1 downregulation in the process of macrophage maturation. Thus, NPF downregulation by protein degradation is likely to be a general mechanism that balances actin polymerization during cellular activation and differentiation.

NPFs and Cancer: The Actin-Metastasis-Invasion Connection Metastasis, the cause of most cancer deaths, is the re-location of a cancer cell to a distant organ, where it develops into a metastatic lesion [76]. In order to metastasize, cancer cells must detach from the primary tumor, invade and move through the surrounding tissue, travel through blood or lymph vessels, and finally, invade and colonize at a distant organ. The dynamic remodeling of the actin cytoskeleton provides the structural basis for the formation of different membrane protrusions. These specialized, actin based protrusions, termed invadopodia, provide the metastatic cells with enhanced ability to degrade the cellular matrix (ECM) and to emigrate from the original tumor site. Recently, the ARP2/3 complex and its associated NPFs were found to be involved in cancer progression. ARP2/3 gene expression is upregulated in a rat model of mammary invasive tumor [77], and following the 7q21-q22 gene amplification that causes invasive pancreatic cancer [78]. Accordingly, RNAi silencing of the ARPC1A gene of the ARP2/3

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complex in pancreatic cancer cells, led to a dramatic reduction in cell migration and invasiveness [78]. Additionally, upregulation of ARP2, together with co-localization with WAVE2, was associated with highly invasive breast carcinoma and with liver metastasis of a colorectal tumor [79, 80]. As an NPF, N-WASp is required for the formation of invadopodia [15, 81-83]. The WAVE proteins are also involved in tumor invasion. WAVE2 is essential for lamellipodia formation in carcinoma cells [82], while WAVE1 and WAVE3 are involved in the secretion of matrix degrading metaloproteases (MMPs) that enable ECM modulation [84, 85]. Based on these data, it is not surprising that NPFs are mostly overexpressed in several types of invasive tumor, including breast, colorectal and lung cancer [71]. The overexpression of NPFs in malignancies and their involvement in tumor invasiveness, highlight the importance of actin cytoskeleton homeostasis. Although the exact linkage between NPF overexpression and malignancies is yet to be determined, current results suggest that dysregulation of the actin polymerization machinery is one of the causes that promote cancer progression.

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Conclusion As master regulators of the actin cytoskeleton, the ARP2/3 complex and its associated NPFs are the subject of intensive ongoing research. A great deal of progress has been made since the first identification of ARP2/3 as an actin nucleator, until the identification of the recently discovered NPFs, WASH, WHAMM, and JMY. We now know that NPFs are essential to almost every aspect of cellular activity; ranging from cellular motility, through endocytic trafficking, to transcriptional activity. Using advanced high resolution microscopy, along with gene silencing and improved biochemical techniques, we are beginning to decipher the molecular mechanisms underlying NPF regulation. The more we progress in our research, the more complexity of NPF regulation is revealed. Furthermore, it appears that NPFs not only have additional roles in cellular processes, but generally function as part of multi-protein complexes, that form both stable and transient interactions. In light of this, researchers must consider multiple actin remodeling pathways to grasp the entire process. Many open questions remain. The regulatory mechanisms of the recently discovered NPFs are still unknown. Moreover, even after the high resolution structure determination of the WRC, the activation mechanism of this complex is yet to be established. Regulation of NPF protein expression has implications for pathological conditions including immunodeficiency and cancer. Therefore, understanding the molecular mechanisms regulating NPF expression is important for the development of new specific therapies for these diseases.

Acknowledgment MBS thanks the following agencies for their research support: The Israel Science Foundation for grants no.1659/08, 971/08, 1503/08 and 491/10, the Ministries of Health & Science for grant no. 3-4114 and 3-6540, the Israel Cancer Association through the Estate of

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the late Alexander Smidoda, and the Taubenblatt Family Foundation for the Bio-medicine excellence grant.

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Aldrich syndrome protein (N-WASp) with actin-related protein (ARP2/3) complex. J. Biol. Chem. 2000 Jul 21;275(29):21946-52. Zeng R, Cannon JL, Abraham RT, Way M, Billadeau DD, Bubeck-Wardenberg J, et al. SLP-76 coordinates Nck-dependent Wiskott-Aldrich syndrome protein recruitment with Vav-1/Cdc42-dependent Wiskott-Aldrich syndrome protein activation at the T cellAPC contact site. J. Immunol. 2003 Aug 1;171(3):1360-8. Rivera GM, Vasilescu D, Papayannopoulos V, Lim WA, Mayer BJ. A reciprocal interdependence between Nck and PI(4,5)P(2) promotes localized N-WASp-mediated actin polymerization in living cells. Mol. Cell. 2009 Nov 13;36(3):525-35. Barda-Saad M, Braiman A, Titerence R, Bunnell SC, Barr VA, Samelson LE. Dynamic molecular interactions linking the T cell antigen receptor to the actin cytoskeleton. Nat. Immunol. 2005 Jan;6(1):80-9. Barda-Saad M, Shirasu N, Pauker MH, Hassan N, Perl O, Balbo A, et al. Cooperative interactions at the SLP-76 complex are critical for actin polymerization. Embo J. 2010 Jun 18. Martinez-Quiles N, Rohatgi R, Anton IM, Medina M, Saville SP, Miki H, et al. WIP regulates N-WASP-mediated actin polymerization and filopodium formation. Nat. Cell Biol. 2001 May;3(5):484-91. Takano K, Toyooka K, Suetsugu S. EFC/F-BAR proteins and the N-WASP-WIP complex induce membrane curvature-dependent actin polymerization. Embo J. 2008 Nov 5;27(21):2817-28. Bu W, Lim KB, Yu YH, Chou AM, Sudhaharan T, Ahmed S. Cdc42 interaction with N-WASP and Toca-1 regulates membrane tubulation, vesicle formation and vesicle motility: implications for endocytosis. PLoS One. 2010;5(8):e12153. Nolz JC, Gomez TS, Zhu P, Li S, Medeiros RB, Shimizu Y, et al. The WAVE2 complex regulates actin cytoskeletal reorganization and CRAC-mediated calcium entry during T cell activation. Curr Biol. 2006 Jan 10;16(1):24-34. Zipfel PA, Bunnell SC, Witherow DS, Gu JJ, Chislock EM, Ring C, et al. Role for the Abi/wave protein complex in T cell receptor-mediated proliferation and cytoskeletal remodeling. Curr Biol. 2006 Jan 10;16(1):35-46. Eden S, Rohatgi R, Podtelejnikov AV, Mann M, Kirschner MW. Mechanism of regulation of WAVE1-induced actin nucleation by Rac1 and Nck. Nature. 2002 Aug 15;418(6899):790-3. Kim Y, Sung JY, Ceglia I, Lee KW, Ahn JH, Halford JM, et al. Phosphorylation of WAVE1 regulates actin polymerization and dendritic spine morphology. Nature. 2006 Aug 17;442(7104):814-7. Innocenti M, Zucconi A, Disanza A, Frittoli E, Areces LB, Steffen A, et al. Abi1 is essential for the formation and activation of a WAVE2 signalling complex. Nat. Cell Biol. 2004 Apr;6(4):319-27. Ismail AM, Padrick SB, Chen B, Umetani J, Rosen MK. The WAVE regulatory complex is inhibited. Nat. Struct. Mol. Biol. 2009 May;16(5):561-3. Scheele JS, Marks RE, Boss GR. Signaling by small GTPases in the immune system. Immunol. Rev. 2007 Aug;218:92-101. Miki H, Suetsugu S, Takenawa T. WAVE, a novel WASP-family protein involved in actin reorganization induced by Rac. Embo J. 1998 Dec 1;17(23):6932-41.

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[64] Yan C, Martinez-Quiles N, Eden S, Shibata T, Takeshima F, Shinkura R, et al. WAVE2 deficiency reveals distinct roles in embryogenesis and Rac-mediated actin-based motility. Embo J. 2003 Jul 15;22(14):3602-12. [65] Oikawa T, Yamaguchi H, Itoh T, Kato M, Ijuin T, Yamazaki D, et al. PtdIns(3,4,5)P3 binding is necessary for WAVE2-induced formation of lamellipodia. Nat. Cell Biol. 2004 May;6(5):420-6. [66] Stuart JR, Gonzalez FH, Kawai H, Yuan ZM. c-Abl interacts with the WAVE2 signaling complex to induce membrane ruffling and cell spreading. J. Biol. Chem. 2006 Oct 20;281(42):31290-7. [67] Leng Y, Zhang J, Badour K, Arpaia E, Freeman S, Cheung P, et al. Abelson-interactor1 promotes WAVE2 membrane translocation and Abelson-mediated tyrosine phosphorylation required for WAVE2 activation. Proc. Natl. Acad. Sci. USA. 2005 Jan 25;102(4):1098-103. [68] Sossey-Alaoui K, Li X, Cowell JK. c-Abl-mediated phosphorylation of WAVE3 is required for lamellipodia formation and cell migration. J. Biol. Chem. 2007 Sep 7;282(36):26257-65. [69] Chen Z, Borek D, Padrick SB, Gomez TS, Metlagel Z, Ismail AM, et al. Structure and control of the actin regulatory WAVE complex. Nature. 2010 Nov 25;468(7323):533-8. [70] Derivery E, Sousa C, Gautier JJ, Lombard B, Loew D, Gautreau A. The Arp2/3 activator WASH controls the fission of endosomes through a large multiprotein complex. Dev. Cell. 2009 Nov;17(5):712-23. [71] Kurisu S, Takenawa T. WASP and WAVE family proteins: friends or foes in cancer invasion? Cancer Sci. 2010 Oct;101(10):2093-104. [72] Shcherbina A, Miki H, Kenney DM, Rosen FS, Takenawa T, Remold-O'Donnell E. WASP and N-WASP in human platelets differ in sensitivity to protease calpain. Blood. 2001 Nov 15;98(10):2988-91. [73] Lutskiy MI, Shcherbina A, Bachli ET, Cooley J, Remold-O'Donnell E. WASP localizes to the membrane skeleton of platelets. Br. J. Haematol. 2007 Oct;139(1):98-105. [74] Oda A, Miki H, Wada I, Yamaguchi H, Yamazaki D, Suetsugu S, et al. WAVE/Scars in platelets. Blood. 2005 Apr 15;105(8):3141-8. [75] Dinh H, Scholz GM, Hamilton JA. Regulation of WAVE1 expression in macrophages at multiple levels. J. Leukoc. Biol. 2008 Dec;84(6):1483-91. [76] Chaffer CL, Weinberg RA. A perspective on cancer cell metastasis. Science. 2011 Mar 25;331(6024):1559-64. [77] Wang W, Wyckoff JB, Goswami S, Wang Y, Sidani M, Segall JE, et al. Coordinated regulation of pathways for enhanced cell motility and chemotaxis is conserved in rat and mouse mammary tumors. Cancer Res. 2007 Apr 15;67(8):3505-11. [78] Laurila E, Savinainen K, Kuuselo R, Karhu R, Kallioniemi A. Characterization of the 7q21-q22 amplicon identifies ARPC1A, a subunit of the Arp2/3 complex, as a regulator of cell migration and invasion in pancreatic cancer. Genes Chromosomes Cancer. 2009 Apr;48(4):330-9. [79] Iwaya K, Norio K, Mukai K. Coexpression of Arp2 and WAVE2 predicts poor outcome in invasive breast carcinoma. Mod. Pathol. 2007 Mar;20(3):339-43. [80] Iwaya K, Oikawa K, Semba S, Tsuchiya B, Mukai Y, Otsubo T, et al. Correlation between liver metastasis of the colocalization of actin-related protein 2 and 3 complex and WAVE2 in colorectal carcinoma. Cancer Sci. 2007 Jul;98(7):992-9.

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[81] Yamaguchi H, Lorenz M, Kempiak S, Sarmiento C, Coniglio S, Symons M, et al. Molecular mechanisms of invadopodium formation: the role of the N-WASP-Arp2/3 complex pathway and cofilin. J. Cell Biol. 2005 Jan 31;168(3):441-52. [82] Oser M, Yamaguchi H, Mader CC, Bravo-Cordero JJ, Arias M, Chen X, et al. Cortactin regulates cofilin and N-WASp activities to control the stages of invadopodium assembly and maturation. J. Cell Biol. 2009 Aug 24;186(4):571-87. [83] Lorenz M, Yamaguchi H, Wang Y, Singer RH, Condeelis J. Imaging sites of N-wasp activity in lamellipodia and invadopodia of carcinoma cells. Curr. Biol. 2004 Apr 20;14(8):697-703. [84] Suetsugu S, Yamazaki D, Kurisu S, Takenawa T. Differential roles of WAVE1 and WAVE2 in dorsal and peripheral ruffle formation for fibroblast cell migration. Dev. Cell. 2003 Oct;5(4):595-609. [85] Sossey-Alaoui K, Ranalli TA, Li X, Bakin AV, Cowell JK. WAVE3 promotes cell motility and invasion through the regulation of MMP-1, MMP-3, and MMP-9 expression. Exp. Cell Res. 2005 Aug 1;308(1):135-45.

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In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 229-244

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter VII

Actin Cytoskeleton Alterations: Are There Any Consequences? Silvia Versari, Livia Barenghi and Silvia Bradamante CNR-ISTM Institute of Molecular Science and Technologies Via Golgi 19 20125 Milan, Italy.

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Abstract Actin is one of the most abundant and highly conserved proteins of all eukaryotes. It is the major component of the cytoskeleton, and is involved in many of the structural and dynamic aspects of cell growth, differentiation, division, membrane organisation, transport, and signal transduction. Alterations in such a critical component can lead to pathological conditions. We here describe the effects of actin cytoskeleton disorganisation and/or depolymerisation in the Saccharomyces cerevisiae yeast model system. The structure of the actin cytoskeleton was disorganised by subjecting yeast cells to simulated (Rotating Wall Vessel) or real microgravity (spaceflight), both of which activated the signal transduction cascade of the high osmolarity glycerol (HOG) MAP kinase pathway, which responds to cell swelling/shrinking, and the cell wall integrity (CWI) pathway, which is involved in cell wall biogenesis and actin cytoskeleton reorganisation. The same results were observed when the actin cytoskeleton structure was depolymerised by means of treatment with dihydrocytochalasin B (DHCB) or (+)-(R)trans-4-(1-aminoethyl)-N-(4-pyridyl)cyclohexanecarboxamide dihydrochloride (Y27632). The HOG and CWI activation indicate a response to a variation in cell volume. Under such conditions, yeast activates volume-sensitive ion channels that alter the ion flux to restore normal volume. These alterations are not pathological per se but, in the case of significant environmental stress (such as oxidative stress), they can lead to clear signs of damage. We observed massive protein carbonylation and a marked loss of the antioxidant glutathione through chloride channels. These findings suggest interrelationships between the actin cytoskeleton, cell volume regulation and the loss of antioxidant defences, and provide new insights into the

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Silvia Versari, Livia Barenghi and Silvia Bradamante underlying cause of the GSH depletion associated with many human diseases, such as cancer, neurodegenerative disorders, and HIV- and aging-related diseases.

Introduction The findings of a large number of studies using localisation and drug-based disruption methods suggest a close relationship between the structural organisation of the actin cytoskeleton and alterations in cell volume [1]. Various lines of evidence indicate that alterations in actin structure may act directly on the membrane transporters responsible for volume homeostasis. This coupling is complex as it involves a number of upstream and downstream regulatory partners, and cell responses depend on the cell type. For this study, yeast was chosen as the model system because it has short generation times, is less demanding than mammalian cells in terms of culture conditions, and yeast osmoadaptation mechanisms have been extensively studied. Many agents can induce osmotic perturbations and/or actin cytoskeleton alterations. This chapter considers mechanical forces (microgravity) and the chemical compounds dihydrocytochalasin B (DHCB) and (+)-(R)-trans-4-(1-aminoethyl)-N-(4-pyridyl) cyclohexanecarboxamide dihydrochloride (Y-27632). The effects of applying an additional stress to the already altered cell system are also considered. Among the various environmental stresses, oxidative stress has been chosen since it is a widely used approach and the cell responses are well characterized.

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1. Mechanical Forces and Chemical Compounds Acting on the Actin Cytoskeleton 1.1. Real Microgravity Many attempts have been made to explain the effects of gravity at cell level, and it is now known that cells adapt to a new gravitational environment by modifying their metabolism and/or molecular organisation. Under conditions of microgravity (µg), nutrients and oxygen are only supplied by diffusion because of the lack of sedimentation and free convection, and in the cell-solution border the membrane potential can change causing alterations in solute concentrations. These conditions lead to the formation of stationary boundary layers around the cells and, as the velocity of oxygen and nutrient consumption exceeds the rate of diffusion, cell metabolism can be disordered [2]. These changes in the distribution of solute concentrations possibly lead to osmotic volume perturbations. Many mathematical models have been proposed to explain these effects [2-4]. Fernàndez-Sempere et al. used real-time holographic interferometry (previously used to measure concentration profiles in the polarised layer during membrane processes) to visualise buoyancy effects on the dead-end reverse osmosis of salts by rotating the cell 90° and 180° from its original position (0°) [4]. However, less has been done at cell level. On the basis of what is known so far, cell osmoadaptation in µg implies critical cytoskeletal rearrangements that have always been observed in the different cell types cultured in space. Hughes-Fulford et al. described a uniquely abnormal

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morphology, fewer stress fibres, and significant alterations in the actin cytoskeleton of MC3T3-E1 osteoblasts launched on the STS-56 shuttle flight [5]; Lewis et al. found diffuse shortened microtubules extending from poorly defined microtubule organising centres (MTOCs) in human T lymphoblastoid cells (Jurkat) flown on the space shuttle [6]; and Walther et al. found significant differences in the distribution of bud scars and increased random budding of Saccharomyces cerevisiae yeast cultured for eight days in µg during the Spacelab IML-2 mission [7].

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1.2. Simulated Microgravity Gravitational force on the space station or on a space craft is approximately 10−4 to 10−6 g. There are bioreactors for cell cultures that are capable of modelling aspects of µg on ground, and these are valuable for developing hypotheses concerning gravitational cell biology, as well as for directing the design and scope of orbital space studies, and substantiating the findings [8-11]. They reduce the average gravitational force acting on the cells to between 10−2 and 10−3. For the sake of simplicity, these conditions of hypogravity are referred to as simulated µg (sim-µg). Two commonly used commercial simulators are the Rotating Wall Vessel (RWV, Synthecon Inc., Houston, TX, USA) [12], and the Random Positioning Machine (RPM, Dutch Space, NL) [13, 14]. The RWV is a suspension culture vessel optimised to produce laminar flow and minimise mechanical stress on cultured cell aggregates. It provides fluid dynamic operating principles characterised by: 1) solid body rotation about a horizontal axis that is characterised by the co-localisation of cells and aggregates with different sedimentation rates, optimally reduced fluid shear and turbulence, and three-dimensional spatial freedom; and 2) oxygenation by diffusion. Oxygenation takes place through a gas-permeable silicone rubber membrane. Regardless of whether they are suspended or seeded on microcarriers, cells grown in the RWV are cultured in “suspended animation” as a result of which they are continuously free-falling and promoting the assembly of 3D cell aggregates which allow more efficient cell-to-cell interactions and the exchange of growth factors. The RPM is essentially a 3-axis clinostat that creates a condition in which the gravity vector is continually and randomly reoriented. When the changes in direction are faster than the object‟s response to gravity, the effects are comparable to the effects of real µg [15]. The device consists of two cardanic frames and an experimental platform. The frames are driven by means of belts and two electro-motors, which are controlled on the basis of feedback signals generated by encoders mounted on the motor axes and „null position‟ sensors on the frames [16]. The RPM allows cells to be cultured in standard plates.

1.3. Dihydrocytochalasin B (DHCB) Cytochalasins are a group of small, naturally occurring cell-permeable organic molecules that bind to actin filaments and block its polymerisation and elongation. They have been widely used to study the role of actin in many biological processes, and as models for actinbinding proteins. Functionally, cytochalasins resemble capping proteins, blocking one end of actin filaments, nucleating polymerisation, and shortening filaments. They bind to the barbed

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end of actin filaments, which inhibits the association and dissociation of subunits at that end [17]. This chapter concentrates on the saturated derivative of cytochalasin B (DHCB), a wellknown inhibitor of actin polymerisation that induces changes in morphology and motility, and shortens actin filaments by blocking monomer additions at the fast-growing end of polymers, but has little effect on sugar transport.

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1.4. (+)-(R)-trans-4-(1-aminoethyl)-N-(4pyridyl)cyclohexanecarboxamide dihydrochloride (Y-27632) Y-27632 is a potent and selective inhibitor of the Rho-associated coiled-coil forming protein serine/threonine kinase (ROCK) family of protein kinases Y-27632 [18]. Rho GTPases are members of the Ras superfamily of monomeric GTP-binding proteins. Many different Rho GTPases have so far been identified (some with multiple isoforms); the most extensively characterised members are Rho, Rac and Cdc42. The main function of the Rho GTPase family is to regulate the assembly and organisation of the actin cytoskeleton by means of direct and/or indirect action on actomyosin assembly and contraction, actin filament stabilisation, and increased actin polymerisation. Although this family is best characterised for their effects on the actin cytoskeleton, they are also involved in other cell functions including cell adhesion, cell motility, vascular and smooth muscle contraction, cytokinesis, cell proliferation, and gene transcription [19, 20]. The Rho GTPase proteins are highly sensitive to cell volume changes and play important roles in a number of volume-dependent changes in the actin cytoskeleton, including shrinkage- and swellinginduced F-actin rearrangements [21]. They are also good candidates for the regulation of ion channel activity, but the mechanism is complex as it is modulated by many factors, including cross-talk with other intracellular signalling pathways, interactions with cytoskeletal elements, and probably other factors yet uncharacterized [22]. Given the involvement of Rho GTPases in such a wide variety of important cell processes, much effort has been made to identify their cell targets or effector proteins. The Rho, Rac and Cdc42 effectors involved in actin cytoskeleton reorganisation are respectively ROCK (Rho-associated coiled-coil forming protein serine/threonine kinase), WAVE (WASPlike verprolin-homologous protein)/PI-4-P5K (Phosphatidylinositol-4-phosphate 5kinase)/PAK (p21-activated kinase), and PAK/N-WASP (Wiskott Aldrich syndrome protein). This study focuses on the Rho-associated kinase ROCK and its potent and selective inhibitor Y-27632.

2. Yeast as a Model System Yeast has many characteristics shared by higher organisms (including humans) [23-25], particularly lots of homolog genes [26]. Saccharomyces cerevisiae is the most commonly used system because of industrial interest, the many laboratories studying it, and the large number of optimised and already tested protocols. This chapter highlight only some of the specific features involved in actin alterations.

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2.1. Osmoadaptation

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It has been shown that changes in medium osmolarity affect various signalling pathways in yeast. By far the best-characterised systems are the high osmolarity glycerol (HOG) and cell wall integrity (CWI) pathways [27, 28]. HOG is activated by osmotic changes in less than one minute. An osmotic up-shift causes a striking transcriptional response that affects the expression of about 10% of yeast genes. The inability of mutants with an inactive HOG pathway to adapt properly to high osmolarity medium, and the known function of genes whose expression is stimulated via the HOG pathway, confirm that the pathway‟s cellular role is to coordinate a significant part of the transcriptional response of yeast cells to high osmolarity. The HOG pathway also mediates post-transcriptional effects. The CWI pathway orchestrates changes in cell morphology by controlling the expression of genes encoding the enzymes involved in cell wall metabolism, and by taking part in the reorganisation of the actin cytoskeleton. It is a network of interacting signalling routes that diverge from or converge to protein kinase C (Pkc1p) and G-protein Rho1p, as well as the proteins controlling them. The many pathways interacting physically and/or genetically with these central components of the CWI pathway include the Slt2/Mpk1 MAP kinase cascade and the HOG pathway. In brief, the CWI pathway controls cell wall metabolism during growth and development, and the response to cell expansion or cell wall damage. The HOG and the CWI MAP kinase pathways collaborate in monitoring cell swelling or shrinking: the HOG pathway at plasma membrane level, and the CWI pathway at cell wall level. Together, the two pathways set the osmotic conditions and appropriate turgor pressure for cell morphogenesis [27, 28].

2.2. Actin Cytoskeleton Yeast can adapt and grow under a variety of environmental conditions [25], and may sense and respond to physical forces. The actin cytoskeleton is the cell component that is mainly involved in the adaptation of yeast to environmental changes. Mammalian cells have many actin isoforms, but Saccharomyces cerevisiae has only one essential gene for actin (ACT1), and it has been found that diploid cells containing a single copy of ACT1 are osmosensitive [29]. The cytoskeleton has long been assigned important roles in mechanosensing and transduction, and has been the subject of much interest as a player in the events initiated by cell volume perturbations. Extensive cytoskeleton reorganisation takes place rapidly after osmotic volume perturbations in the great majority of cell types. Yeast has three different types of cytoskeletal filaments: long flexible polymers of actin called microfilaments (Factin), polymers of tubulin called microtubules, and intermediate filaments [30]. Of the three, F-actin plays a major role because its net increase/decrease is associated with osmotic shrinkage/swelling.

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2.3. Cell Volume Regulation and Chloride Channels

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It is known that an osmotic shock disrupts the actin cytoskeleton, which is reorganised during recovery and adaptation [1]. The findings of a number of studies using the actin filament disrupting agents, cytochalasins D and B, indicate a close relationship between actin and the regulatory volume decrease (RVD) response. In most cells, the RVD is caused by an efflux of K+, Cl– and organic osmolytes such as polyols and amino acids (e.g. glycine, glutamate and aspartate). The voltage-dependent chloride channels (ClCs) activated by cell swelling play a key role in this process [31]. Among the various proteins that may be responsible for this chloride conductance in mammalian cells, it has been suggested that ClC-3 (a member of the ClC family) may be the chloride channel involved in cell volume regulation. In a number of mammalian cell types (renal cortical collecting duct cells, astrocytes, intestinal epithelial cells, etc.), it has been found that the RVD-activated chloride channel is activated by the cytochalasin-mediated disruption of actin, and inhibited by the phalloidin-mediated stabilisation of actin [32-35]. ClCs are the most evolutionarily conserved family of chloride channels, having homologues in both prokaryotic and eukaryotic organisms. The properties of the protein channel in yeast have not been extensively studied. Saccharomyces cerevisiae possesses only one gene (GEF1) that encodes for the Gef1p protein that is regarded as a member of the voltage-gated chloride channel. Gef1p, which possesses homology with the mammalian ClC-3, transports chloride anions through the plasma membrane and plays a crucial role in ion homeostasis [36, 37].

3. The Effects on Yeast of Reduced-Gravity Environments and the Two Actin-Disrupting Drugs This chapter describes a series of experiments aimed at evaluating the responses of Saccharomyces cerevisiae to conditions that are known to induce osmotic perturbations and/or actin cytoskeleton alterations (reduced-gravity environments and actin disrupting drugs). The effects of µg as a mechanical force capable of inducing osmotic perturbations were evaluated in space by means of the Saccharomyces cerevisiae Oxidative stress Response Evaluation (SCORE) experiment [38] and in sim-µg on ground using the RWV bioreactor. The results have been compared with those obtained on ground (1g) by treating the cells with DHCB and Y-27632, two chemical compounds that directly or indirectly induce cytoskeletal modifications. The data are summarised in Table 1.

3.1. Reduced Gravity Environments The SCORE experimental hardware and protocol are described in Figure 1. During the spaceflight experiment, the yeast cells were metabolically active as they consumed glucose, produced ethanol, and maintained a high intracellular trehalose content.

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Figure 1. SCORE spaceflight experiment. The SCORE experimental hardware is an automated apparatus for cell culture and fixation designed for spaceflight that consists of a battery pack, an electronic board, a peristaltic pump, and two separate bioreactors, each of which contains two variable volume modules (VVMs). Each VVM (20 mL) consists of two chambers separated by a sliding piston. One VVM of each bioreactor is dedicated to cell culture and culture medium, the other to fixatives. Two parallel 3-step experiments were carried out: one used oxygen saturated medium (hyperoxia) and the other aerated medium (normoxia). The 24-hour experiments were prepared seven days before the FOTON-M3 launch, and activated seven days after the launch. At the end of the experiment, the yeast cells and culture medium were separated and preserved using suitable fixative.

In comparison with the related ground controls, the spaceflight samples showed a twofold increase in random budding and the activation of two major MAP kinase pathways in response to the imposed stress: HOG and CWI. Attention was concentrated on the Hog1 and Slt2 proteins, which were found to be markedly activated. As a further result of HOG pathway activation, a significant concentration of glycerol was detected in the culture medium. As spaceflight constraints limited the amount of material and the number of analyses, SCORE was repeated and validated on the ground in sim-µg using the RWV bioreactor and following the same experimental and analytical protocol. Our previous experiences with various µg-simulators confirm the reliability of this approach [39-41] although the values are lower than those obtained during a spaceflight.

3.2 Actin- Disrupting Drugs Using the same experimental and analytical protocol as above, the cells were treated with DHCB under conditions of normal gravity. There was a 1.5-fold increase in random budding, and significant Hog1 and Slt2 activation. Furthermore, methylene blue staining and the determination of colony-forming units revealed no alteration in cell viability, and high osmolarity, sodium dodecyl sulfate, and Calcofluor White sensitivity assays indicated no alteration in cell wall integrity or permeability.

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Y-27632 treatment validated the DHCB data: a 1.6-fold increase in random budding and Hog1 and Slt2 activation. Table 1. Effects of reduced-gravity environments and actin disrupting drugs on Saccharomyces cerevisiae cultured in normoxia

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% Random buddinga, c P-Hog1/Hog1b,c P-Slt2/Slt2b,c 1g control 35,50 ± 0,72 0,14 ± 0,01 0,21 ± 0,01 µg (SCORE) 78,0 1,33 7,36 sim-µg 70,70 ± 1,30 2,20 ± 0,09 3,40 ± 0,09 DHCB 62,50 ± 1,68 2,07 ± 0,04 2,71 ± 0,21 Y-27632 63,00 ± 0,60 1,85 ± 0,02 2,44 ± 0,02 1g control: earth gravity control; µg: microgravity; SCORE: spaceflight experiment; sim-µg: microgravity simulated using the Rotating Wall Vessel (RWV); DHCB: dihydrocytochalasin B; Y27632: (R)-(+)-trans-4-(1-aminoethyl)-N-(4-pyridyl)cyclohexanecarboxamide dihydrochloride monohydrate. a % random budding. Bud scar distribution was examined by means of UV fluorescence microscopy after chitin staining with Calcofluor White. In comparison with the related ground control, there was a significant increase in random budding in all of the experiments. b MAP kinase analysis. Hog1 and Slt2 activation were detected by immunoblotting using an anti-phospho-p38 antibody that recognises phospho-Hog1 (P-Hog1) or an anti-phospho-p44/42 MAP kinase antibody that recognizes phospho-Slt2 (P-Slt2). The values shown in the table represent the densitometric quantification of at least three blots of phospho-protein normalised to the amounts of the specific core proteins. Hog1and Slt2 were activated under all of the experimental conditions. c The results are expressed as the mean values ± SEM of an appropriate number of experiments (except for the single SCORE experiment).

In all of these experiments, the HOG and CWI pathway activation indicated the yeast‟s response to a variation in cell volume. Under such conditions, yeast activates volumesensitive ion channels that alter the ion flux in order to restore normal volume. The alterations are not pathological per se but, in the case of significant environmental stress, can lead to clear signs of damage.

4. Oxidative Stress It is well known that cells have developed numerous antioxidant factors, in order to counteract the effects of reactive oxygen species (ROS), such as superoxide, hydrogen peroxide and highly reactive hydroxyl radicals. These include detoxifying enzymes such as superoxide dismutases, catalases, and peroxidases that detoxify ROS directly, and many defence systems such as glutathione (GSH), thioredoxin and methionine sulfoxide reductases, which maintain a cellular redox state and repair oxidatively damaged proteins, DNA and lipids [42, 43]. This study concentrated on GSH [44] and, in particular, any alterations in its homeostasis. GSH is widely found in micro-organisms, plants and animals, and has various defensive functions against oxidative stress and xenobiotic toxicity by working as a scavenger in a complex with GSH peroxidase (GPx). It plays a critical role in many biological processes, acting directly as a co-factor in enzymatic reactions and indirectly as the major thiol-disulfide redox buffer in mammalian cells. It also provides a critical defence system that

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protects cells against many different forms of stress. In this study, in order to avoid using of free radical-generating compounds [45], mild oxidation was applied to prevent the extensive chemical deterioration of the system. Molecular oxygen was chosen as the oxidant because of the restrictions imposed by the experimental set-up, particularly the SCORE spaceflight experiment and the reported evidence of hyperoxic damage [42]. The responses of Saccharomyces cerevisiae to hyperoxia were evaluated when it was cultured in reduced-gravity environments or treated at 1g with the two actin-disrupting drugs.

4.1 Extracellular GSH Release and Protein Carbonylation

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The results indicated a high rate of GSH release in the extracellular medium (GSH ER), which reached up to 40% w/dw in the SCORE spaceflight experiment, and 8-12% w/dw under the other experimental conditions (Figure 2).

Figure 2. GSH extracellular release (GSH ER) in hyperoxia. The extracellular GSH concentration was determined by means of reverse-phase HPLC. The data were validated by analysing the 1H NMR spectra of the supernatant or by using the Glutathione Assay Kit (Sigma) in accordance with the manufacturer's instructions. In comparison with the related ground control, hyperoxia induced enormous GSH ER (% w/dw) under all of the experimental conditions. The results are expressed as the mean values ± SEM of an appropriate number of experiments. In the case of the single SCORE spaceflight experiment, the results are the mean value ± SEM of eight independent measures.

Furthermore, the extracellular release of GSH was accompanied by a marked increase in protein carbonylation in all of the experiments. Protein carbonylation is the major and most common oxidative alteration. It has been shown that protein carbonyls affect the function and stability of the modified proteins, and it is likely that they also play an important role in the

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pathophysiology of disorders associated with considerable oxidative stress. There was also a significant increase in random budding and Hog1 and Slt2 activation (Table 2). Table 2. Effects of reduced-gravity environments and of actin disrupting drugs on Saccharomyces cerevisiae cultured in hyperoxia % Random P-Hog1/Hog1b,c buddinga, c 1g control 54,00 ± 0,51 0,27 ± 0,01 µg (SCORE) 81,00 14,04 sim-µg 71,00 ± 2,23 4,67 ± 0,17 DHCB 59,50 ± 1,20 3,01 ± 0,06 Y-27632 64,00 ± 0,85 3,09 ± 0,04 Hyperoxia was obtained by saturating the culture medium with oxygen. See details, results and abbreviations.

P-Slt2/Slt2b,c 0,55 ± 0,01 17,64 5,18 ± 0,08 4,44 ± 0,06 4,27 ± 0,03 Table 1 for experimental

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4.2. Dynamics of GSH ER The dynamics of the GSH ER (Figure 3) indicated that it occurs within one hour and is accompanied by a parallel increase in the glycerol concentration of the extracellular medium. The release is concomitant with the transcriptional activation of GSH1, GSH2, HSP26 (heat shock protein 26) and CTT1 (cytosolic catalase T), as detected by means of reversetranscription (RT) PCR. GSH1 and GSH2 encode two gamma glutamylcysteine synthetases that catalyse the two main steps of GSH biosynthesis, whereas HSP26 and CTT1 are two genes involved in the stress response and their transcription is mediated by the MAP kinase pathway [46], which is in line with the observed activation of Hog1. On the contrary, GTT1 (which encodes GSH transferase 1), BPT1 and YCF1 (which encode two vacuolar GSH Sconjugate transporters of the ATP-binding cassette family) were not modulated, thus indicating that vacuolar transport was not involved in the observed GSH ER.

4.3. Involvement of Chloride Channels in GSH ER Although their final effects are very similar, the reduced-gravity environment and the two actin-disrupting drugs act differently. Real or simulated microgravity alters the organisation of the cytoskeletal machinery, whereas DHCB and Y-27632 depolymerise actin fibres. In all of the considered cases, the cells responded to the imposed stresses by producing and releasing an enormous amount of GSH into extracellular space. Working on the hypotheses that yeast activates the volume-sensitive ion channels in order to restore normal volume, and that anionic GSH is permeant in chloride channels [47] (as observed in cystic fibrosis), the sim-µg, DHCB, and Y-27632 experiments were repeated in hyperoxia with the potent chloride channel blocker 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) being added to the culture medium. In all cases, NPPB completely blocked GSH ER, thus confirming the key involvement of chloride channels (Figure 4). A similar

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relationship between cell volume alteration, actin structure and RVD-activated chloride channel has also been found in a number of other cell types [32-34, 48].

Figure 3. Twenty-four hour profiles of intra- and extracellular GSH, ethanol and glycerol concentrations, and pH in earth gravity control (1g ctr), sim-µg, and DHCB in hyperoxia. The cells were harvested and their intra- and extracellular GSH (% w/dw), ethanol and glycerol concentrations and pH were determined at selected time points. (a, a‟) In the 1g ctr experiment, GSH ER was very low, whereas the release of ethanol and glycerol were as expected. (b) In sim-µg. GSH ER occurred 10-11 hours after t=0 and, at the same time, intracellular GSH peaked before returning to its initial level. (c) Similarly, but four hours earlier, the presence of DHCB under hyperoxic conditions caused GSH ER in one hour. (b‟-c‟) Twenty-four hour profiles of ethanol and glycerol release in sim-µg and DHCB. See Table 1 for abbreviations.

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Figure 4. Twenty-four hour profiles of intra- and extracellular GSH, ethanol and glycerol concentrations, and pH in (a) sim-µg, and (b) DHCB in hyperoxia with the addition of 5-nitro-2-(3phenylpropyl-amino) benzoic acid (NPPB). (a – b) The addition of NPPB completely blocked GSH ER, whereas pH remained in the range 4.8-5.4. (a‟-b‟) Twenty-four hour profiles of ethanol and glycerol release in sim-µg and DHCB. See Table 1 for abbreviations.

Conclusion The actin cytoskeleton plays a central role in many cellular processes, which may be altered by changes in its organisation and polymerisation dynamics. Taking advantage of the space extreme environment and of two well-known actin depolymerising compounds, some of the yeast responses to alterations in actin structure and/or cell volume have been investigated. In these conditions, the Saccharomyces cerevisiae yeast, chosen as a model system, adapts and survives by activating the MAPK signalling cascade and volume-sensitive ion channels. In addition, in the presence of a mild oxidative stress huge amounts of intracellular GSH is released. Attention has been concentrated on GSH, since it protects against oxidative damage caused by ROS, being one of the major defences under oxidative stress [49]. Briefly, all of the reported experiments demonstrate that: 1) microgravity induces alterations of cell volume, and this causes critical actin cytoskeleton rearrangements that determine the activation of the volume-sensitive ion channels though which GSH is released; 2) the two chemical compounds DHCB and Y-27632 induce alterations in the structure of actin inducing similar GSH extracellular release. Being aware that some of the experiments have a number of limitations, in terms of number of samples, choice of fixatives and

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environmental stress, nevertheless the straightforward conclusion is that in all the considered conditions, the actin-related activation of chloride channels is the main process responsible for GSH loss. The evident inter-relationship between the actin cytoskeleton structure, volume-sensitive ion channels, and the loss of antioxidant defences will provide new insights into the underlying causes of many human pathologies.

Acknowledgements The study was supported by the Italian Space Agency (ASI: Contract No. 1-006/06/1 “DCMC” 1B1118, coordinator: S. Bradamante). We would like to thank Thales Alenia Space for the design, development and support of the SCORE experimental hardware, and dr. Ivan Orlandi and prof. Marina Vai for molecular biology support.

References

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Henson, JH. Relationships between the actin cytoskeleton and cell volume regulation. Microsc. Res. Tech, 1999, 47, 155-162. [2] Slezak, A; W sik J; Dworecki K. Gravitational effects in a passive transmembrane transport: the flux graviosmotic and gravidiffusive effects in non-electrolytes. Journal of Biological Physics, 2000, 26, 149-170. [3] Slezak, A; Jasik-Slezak J; Sieron A. Gravitational effects in the passive osmotic flows across polymeric membrane of electrolytic solutions. Polim. Med., 2000, 30, 21-44. [4] Fernández-Sempere, J; Ruiz-Beviá F; Salcedo-Díaz R; García-Algado P. Measurement of concentration profiles by holographic interferometry and modelling in unstirred batch reverse osmosis. Industrial and Engineering Chemistry Research, 2006, 45, 7219-7231. [5] Hughes-Fulford, M; Lewis ML. Effects of microgravity on osteoblast growth activation. Exp. Cell Res., 1996, 224, 103-109. [6] Lewis, ML; Reynolds JL; Cubano LA; Hatton JP; Lawless BD; Piepmeier EH. Spaceflight alters microtubules and increases apoptosis in human lymphocytes (Jurkat). FASEB J., 1998, 12, 1007-1018. [7] Walther, I; Bechler B; Muller O; Hunzinger E; Cogoli A. Cultivation of Saccharomyces cerevisiae in a bioreactor in microgravity. J. Biotechnol, 1996, 47, 113-127. [8] Meaney, DF; Johnston ED; Litt M; Pollack SR. Experimental and numerical investigations of microcarrier motions in simulated microgravity. Adv. Heat Mass Transf Biotechnol. HTD, 1998, 362, 103-107. [9] Pampaloni, F; Reynaud EG; Stelzer EH. The third dimension bridges the gap between cell culture and live tissue. Nat. Rev. Mol. Cell Biol., 2007, 8, 839-845. [10] Rucci, N; Rufo A; Alamanou M; Teti A. Modeled microgravity stimulates osteoclastogenesis and bone resorption by increasing osteoblast RANKL/OPG ratio. J. Cell Biochem., 2007, 100, 464-473.

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[11] Unsworth, BR; Lelkes PI. Growing tissues in microgravity. Nat. Med., 1998, 4, 901907. [12] Hammond, TG; Hammond JM. Optimized suspension culture: the rotating-wall vessel. Am. J. Physiol. Renal. Physiol., 2001, 281, F12-F25. [13] Klaus, DM. Clinostats and bioreactors. Gravity Space Biol. Bull, 2001, 14, 55-64. [14] van Loon, JJWA. Some history and use of the random positioning machine, RPM, in gravity related research. Advances in Space Research, 2007, 39, 1161-1165. [15] Borst, AG; van Loon JJWA. Technology and developments for the Random Positioning Machine, RPM. Microgravity Science and Technology, 2009, 21, 287-292. [16] Huijser, RH. Desktop RPM: new small size microgravity simulator for the bioscience laboratory. Fokker Space, 2000, 1-5. [17] Cooper, JA. Effects of cytochalasin and phalloidin on actin. J. Cell Biol., 1987, 105, 1473-1478. [18] Ishizaki, T; Uehata M; Tamechika I; Keel J; Nonomura K; Maekawa M, et al. Pharmacological properties of Y-27632, a specific inhibitor of rho-associated kinases. Mol. Pharmacol., 2000, 57, 976-983. [19] Bishop, AL; Hall A. Rho GTPases and their effector proteins. Biochem. J., 2000, 348, 241-255. [20] Ridley, AJ. Rho family proteins: coordinating cell responses. Trends Cell Biol., 2001, 11, 471-477. [21] Tatebayashi, K; Yamamoto K; Tanaka K; Tomida T; Maruoka T; Kasukawa E, et al. Adaptor functions of Cdc42, Ste50, and Sho1 in the yeast osmoregulatory HOG MAPK pathway. The EMBO Journal, 2006, 25, 3033-3044. [22] Pochynyuk, O; Stockand JD; Staruschenko A. Ion channel regulation by Ras, Rho, and Rab small GTPases. Exp. Biol. Med., 2007, 232, 1258-1265. [23] Qi, M; Elion EA. MAP kinase pathways. J. Cell Sci., 2005, 118, 3569-3572. [24] Tong, AH. et al., Global mapping of the yeast genetic interaction network. Science, 2004, 303, 808-813. [25] Gasch, AP; Spellman PT; Kao CM; Carmel-Harel O; Eisen MB; Storz G, et al. Genomic expression programs in the response of yeast cells to environmental changes. Mol. Biol. Cell, 2000, 11, 4241-4257. [26] Botstein, D; Chervitz SA; Cherry M. Yeast as a model organism. Science, 1997, 277, 1259-1260. [27] Hohmann, S. Osmotic stress signaling and osmoadaptation in yeasts. Microbiol. Mol. Biol. Rev., 2002, 66, 300-372. [28] Rodriguez-Pena, JM; Garcia R; Nombela C; Arroyo J. The high-osmolarity glycerol (HOG) and cell wall integrity (CWI) signalling pathways interplay: a yeast dialogue between MAPK routes. Yeast, 2010, 27, 495-502. [29] Chowdhury, S; Smith KW; Gustin MC. Osmotic stress and the yeast cytoskeleton: phenotype-specific suppression of an actin mutation. The Journal of cell biology, 1992, 118, 561-571. [30] Botstein, D; Amberg D; Mulholland J; Huffaker T; Adams A; Drubin D, et al. The yeast cytoskeleton. Cold Spring Harbor Monograph Series, 1997, 21, 1-90. [31] Nilius, B; Droogmans G. Amazing chloride channels: an overview. Acta Physiol Scand, 2003, 177, 119-147.

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[32] Lascola, CD; Nelson DJ; Kraig RP. Cytoskeletal actin gates a Cl- channel in neocortical astrocytes. J. Neurosci., 1998, 18, 1679-1692. [33] Schwiebert, EM; Mills JW; Stanton BA. Actin-based cytoskeleton regulates a chloride channel and cell volume in a renal cortical collecting duct cell line. J. Biol. Chem., 1994, 269, 7081-7089. [34] Tilly, BC; Edixhoven MJ; Tertoolen LG; Morii N; Saitoh Y; Narumiya S, et al. Activation of the osmo-sensitive chloride conductance involves P21rho and is accompanied by a transient reorganization of the F-actin cytoskeleton. Mol. Biol. Cell, 1996, 7, 1419-1427. [35] Mazzochi, C; Benos DJ; Smith PR. Interaction of epithelial ion channels with the actinbased cytoskeleton. Am. J. Physiol. Renal Physiol., 2006, 291, F1113-1122. [36] Lopez-Rodriguez, A; Trejo AC; Coyne L; Halliwell RF; Miledi R; Martinez-Torres A. The product of the gene GEF1 of Saccharomyces cerevisiae transports Cl- across the plasma membrane. FEMS Yeast Res, 2007, 7, 1218-1229. [37] Wolfe, DM; Pearce DA. Channeling studies in yeast: yeast as a model for channelopathies? Neuromol. Med., 2006, 8, 279-306. [38] Bradamante, S; Villa A; Versari S; Barenghi L; Orlandi I; Vai M. Oxidative stress and alterations in actin cytoskeleton trigger glutathione efflux in Saccharomyces cerevisiae. Biochim. Biophys. Acta, 2010, 1803, 1376-85. [39] Versari, S; Villa A; Bradamante S; Maier JAM. Alterations of the actin cytoskeleton and increased nitric oxide synthesis are common features in human primary endothelial cell response to changes in gravity. Biochim. Biophys. Acta, 2007, 1773, 1645-1652.. [40] Villa, A; Versari S; Barenghi L; Maier JAM; Bradamante S. Effects of spaceflight simulation on human cells. European Space Agency, [Special Publication] SP, 2005, SP-585, villa/1-villa/2. [41] Villa, A; Versari S; Maier JA; Bradamante S. Cell behavior in simulated microgravity: a comparison of results obtained with RWV and RPM. Gravit Space Biol Bull, 2005, 18, 89-90. [42] Outten, CE; Falk RL; Culotta VC. Cellular factors required for protection from hyperoxia toxicity in Saccharomyces cerevisiae. Biochem. J., 2005, 388, 93-101. [43] Kohen, R; Nyska A. Oxidation of biological systems: oxidative stress phenomena, antioxidants, redox reactions, and methods for their quantification. Toxicol. Pathol., 2002, 30, 620-650. [44] Ballatori, N; Krance SM; Notenboom S; Shi S; Tieu K; Hammond CL. Glutathione dysregulation and the etiology and progression of human diseases. Biol Chem, 2009, 390, 191-214. [45] Thorpe, GW; Fong CS; Alic N; Higgins VJ; Dawes IW. Cells have distinct mechanisms to maintain protection against different reactive oxygen species: oxidative-stressresponse genes. Proc. Natl. Acad. Sci. USA, 2004, 101, 6564-6569. [46] Schuller, C; Brewster JL; Alexander MR; Gustin MC; Ruis H. The HOG pathway controls osmotic regulation of transcription via the stress response element (STRE) of the Saccharomyces cerevisiae CTT1 gene. EMBO J., 1994, 13, 4382-4389. [47] Linsdell, P; Hanrahan JW. Glutathione permeability of CFTR. Am. J. Physiol., 1998, 275, C323-326. [48] Cantiello, HF. Role of the actin cytoskeleton in the regulation of the cystic fibrosis transmembrane conductance regulator. Exp. Physiol., 1996, 81, 505-514.

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[49] Jones, DP. Radical-free biology of oxidative stress. Am. J. Physiol. Cell Physiol., 2008, 295, C849-868.

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Chapter VIII

Role of the Actin Cytoskeleton in Tumor Escape to Immune System and Acquisition of Tumor Resistance to Cytotoxic Treatment Rania Zaarour1, Fui Goh1, Meriem Hasmim2, Bassam Janji3 and Salem Chouaib2

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1

University of Sharjah, College of Medicine, Department of Basic Medical Sciences, Sharjah, United Arab Emirates. 2 INSERM U753, Institut Gustave Roussy, 94805 Villejuif – France. 3 Public Research Center for Health, Department of Oncology, Laboratory of Experimental Hemato-Oncology, L-1526 Luxembourg City, Luxembourg.

Introduction Cancer cell survival is a fundamental process essential for cancer related mortalities. This process could be related either to a defect in the immune system machinery or to an acquiring of survival properties by cancer cells allowing them to survive despite a functional immune system. How a cancer cell survives in an environment where competent immune cells are present remains an important question. Understanding basic mechanisms of tumor-host immune interactions will shed light on developing methods to eradicate tumor cell survival, to attenuate immunotherapy resistance and to develop targeted anti-cancer drugs. Actin and tubulin form highly versatile, dynamic polymers that can organize cytoplasmic organelles and intracellular compartments, define cell polarity, and generate both pushing and contractile forces. Therefore, it is not surprising that they are key players in many processes in cell biology. Accumulating evidence involve the actin cytoskeleton at the core of this question. Indeed, the cytoskeleton plays an essential role in regulating a plethora of molecular events

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that ultimately lead to death resistance by cancer cell. In this regard, cytoskeletal remodeling has been described to be responsible for the resistance of cancer cells to cytotoxic immune cell attack as well as in the activation of the attack by immune cells. In this chapter we focus on the role of the actin cytoskeleton in the processes of regulating T cell activity and evading T lymphocyte mediated killing events.

Actin as a Key Component of Cell Regulation

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Actin is essential for the survival of most cells: it provides the internal skeletal support and provides the tracts for movement of intracellular cargo. Eukaryotic cells use actin, along with more than one hundred accessory actin regulating proteins, to modulate cellular activities dependent on actin polymerization and turnover. Under physiological conditions, actin monomers polymerize to form filaments, with a helical arrangement of subunits [1]. Polymerization into filaments occurs sequentially: following the formation of short oligomers (nucleation) that are unstable and disassemble readily, rapid elongation occurs in which subunits add quickly onto the ends of the nucleated filament. Actin filaments are polarized because the subunits point in the same direction. Actin subunits each have a binding site for ATP. ATP is rapidly hydrolysed after assembly into filaments resulting in phosphate dissociation (reviewed in [2]). Several accessory proteins regulate the assembly of actin filaments. Their role is to respond to intracellular and extracellular cues that are translated into filament restructuring. These regulatory proteins can function to maintain a free pool of actin monomers, initiate nucleation and polymerization, restrict length, cross-link, and cap or sever actin filaments [3].

Actin and Effector Cytotoxic Cells We have previously demonstrated that shift in cytoskeletal organization can be used by tumor cells to resist to CD8+ cytotoxic T lymphocytes (CTL)-mediated lysis [4]. In this regard, it has been also reported that the activity of the actin binding protein WASp (WiskottAldrich syndrome protein) is required for a number of immune cell functions including migration, phagocytosis and immunological synapse (IS) formation between T cells and tumor cells [5]. The formation of this synapse is believed to be important for effective tumor clearance. Indeed, when in contact with tumor cells, CTL infiltrating human tumors underwent granzyme B (a serine protease produced by CTL and Natural Killer cells) polarization and cytoskeleton rearrangement to form a specific structure in the contact area. It has been suggested that tumor cells targeted by Granzyme B show a fragmentation of the microtubular system and an increase in the expression level of cleaved caspase 3, which suggests that CTL likely provoke changes in tumor cells and subsequently induce cell death [6]. Taken together, these studies plead for an important role of the actin cytoskeleton in the formation of the CTL immunological synapse. Understanding the molecular mechanisms of actin cytoskeleton remodeling in CTL may be relevant for an effective immune-mediated clearance of tumorigenic cells, therefore opening up new avenues for tumor immunotherapy.

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Actin Cytoskeletal Control of T-cell Activation Tumor cells express immunogenic proteins which, when presented at the cell surface by the Major Histocompatibility Complex (MHC) class I molecules, could serve as a trigger for the adaptive immune response. This phenomenon of immune surveillance was initially proposed more than 50 years ago and describes the role of the immune system in eradicating newly emerged tumor cells through identification of tumor-associated antigens [7] [8]. T lymphocytes play a central role in adaptive immunity, and the CTL subset represents the major effector cells recognizing and engaging tumor antigen peptide / MHC complexes (pMHC) via T-Cell Receptor (TCR). This subsequently activates a series of intracellular signaling events that ultimately leads to actin cytoskeleton reorganization and T cell activation. Here we summarize the underlying signal transduction mechanisms of T cell activation and discuss the pivotal role of actin cytoskeleton in poising T cells for their effector functions and tumor rejection.

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1. Proximal Signaling Linking TCR to Actin Cytoskeleton Network Upon engagement of pMHC by TCR, the Src protein tyrosine kinases Lck and Fyn are activated leading to phosphorylation of Immunoreceptor Tyrosine-based Activation Motifs (ITAMs) that lie within the signal transduction subunit of TCR complex, CD3 (reviewed in [9]). Phosphorylated ITAMs recruit and activate Syk-kinase zeta chain-associated protein 70 (ZAP-70), which subsequently initiates the phosphorylation of several downstream signaling molecules including the linker for the activation of T cells (LAT) [10]. Following this, Grb-2 like adapter (Gads) binds LAT and recruits Src homology 2 (SH2) domain-containing Leukocyte Phosphoprotein of 76 kDa (SLP-76) using its SH2 and SH3 domains respectively [11]. Importantly loss-of-function studies support the role of LAT and SLP-76 as key intermediary proteins in linking TCR ligation to distal signal transduction pathways. Indeed, cells deficient in LAT and SLP-76 proteins showed abrogation of TCR-mediated signaling (reviewed in [12]). Moreover, LAT and SLP-76 are required to stabilize Phospholipase-C (PLC)- which is a bifurcating point in TCR downstream signaling leading to activation of calcium- and diacylglycerol (DAG)- dependent pathways resulting in transcriptional activation (reviewed in[9]). SLP-76 functions as a scaffold protein to bind Nck and the guanine nucleotide exchange factor (GEF) Vav1. Vav1 stimulates the activation of the Rho family GTPases members Cdc 42 and Rac1 that play pivotal role in regulating downstream nucleation promoting factors (NPFs). Particularly, Cdc42 activates WASp; Rac1 mediates the activation of WASp family Verprolin homologous protein-2 (Wave2). The association of WASp and Wave2 proteins with the actin-nucleating polypeptides complex, actin-regulated protein 2/3 (Arp 2/3), represents the fundamental machinery governing the initiation of actin filament formation. Arp 2/3 is a multimeric complex consisting of seven polypeptides including Arp2 and Arp3, both of which possess a structure analogous to actin (reviewed in [13]). The formation of actin trimer serves as the trigger point, referred to as nucleation, for actin assembly and subsequent filamentous actin (F-actin) formation [14]. It has been postulated that Arp 2 and

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Arp3 could replace two actin monomers during nucleation thereby initiating cytoskeleton remodeling [15].

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2. Dynamics of Actin Cytoskeleton in Immunological Synapse and Microcluster (MC) Formation It appears that upon TCR ligation there is temporal and spatial hierarchical organization of several actin-regulators leading to actin nucleation and filament formation, two fundamental mechanisms underlying the configuration of IS and the preceding MC formation. IS refers to the specialized subcellular region between a lymphocyte and an antigenpresenting cell (APC), necessary for intercellular communication with and in the immune system [16] [17]. IS formation was initially observed in CD4+ T helper cells, but this phenomenon was rapidly extended to other immune cells including B cells, natural killer cells and CTLs (reviewed in [16]). Significantly, through various biochemical studies and live cell imaging, in vitro and in vivo formation of IS at the interface of CTL/APC has been demonstrated [18, 19] [20]. Currently there are several postulations for the role of IS in T cell response, including T cell priming, endocytosis and exocytosis, exerting constraints on cytotoxic granules to target APC and terminating T cell signaling (reviewed in [17]). Intensive efforts have gone into characterizing the molecular actors of IS formation. It is now known that reorganization of TCR and signaling molecules occurs at IS, exemplified by segregation of these components into specific domains called supramolecular activation clusters (SMACs) [17]. Particularly, TCR-pMHC complexes and specific signaling mediators such as PKC and Lck are located medially in central SMAC (cSMAC), whilst adhesionrelated molecules are found laterally in peripheral SMAC (pSMAC) forming an outer ring around cSMAC. This concentric arrangement is known as the “bull‟s eye” structure, a distinctive feature of T cell IS [21] [22]. It was initially hypothesized that sustained TCR signaling is required for T cell activation; and this could be attributed to the formation of the IS. The segregation of cSMAC and pSMAC during IS formation was generally observed more than 10 minutes after TCR ligation. However, several studies have reported the occurrence of signaling events preceding IS formation. For example, calcium mobilization and tyrosine kinase activity were found to peak within minutes of conjugate formation and appeared to be IS-independent [23] [24]. These observations raised the significant question of what could be the earlier underlying molecular events prior to IS formation. It was then revealed that assembly of TCR with signaling complexes into MCs could be detected at the periphery of T cell-APC interface immediately after ligand conjugation [25] [26]. Intermediary signaling proteins such as ZAP70, LAT, GRB2, SLP76 and Vav1 were found to translocate into TCR-MCs within seconds after TCR ligation [24] [27]. The formation of TCR-MCs was found to be dependent on Factin polymerization as in the presence of latrunculin, an actin filament modulator, de novo formation of TCR-MC was prevented [26] [28] [29]. Interestingly, the presence of formed TCR-MCs was only transient in the periphery as it has been observed that these MCs move inwards to form cSMAC by utilizing frictional coupling to the actin cytoskeletal network, similar to the mechanism underlying cell motility [29] [28] [30]. Additionally, treatment of T cells with actin depolymerizing agent also inhibited TCR-induced calcium mobilization [29].

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Taken together, these results support the pivotal regulatory role of actin skeleton network in T cell activation by governing the formation and centripetal translocation of TCR-MCs, TCR signaling and IS configuration. However, relatively little is known regarding the linker proteins involved in these processes and awaits further exploration. The exact roles of MCs and IS in T cell signaling is still an area of constant debate. IS was originally thought to be central for sustaining signaling due to high abundance of TCR/pMHC and Lck in cSMAC [22] [31]. But some other studies showed that continuous formation of TCR-MCs is required to sustain signaling and that retention of TCR-MCs in periphery results in signal augmentation [25] [26] [32]. Thus, at present, the general consensus is that TCR-MCs serve as the core of signaling whereas cSMAC might contribute to endocytosis and signal termination [25, 26, 29].

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Role of Actin Cytoskeleton in Tumor Resistance to CTL-Induced Death Most studies on the role of actin cytoskeleton during the interaction between CTL and their target cells were focused on CTL. However, target cell cytoskeleton seems to play a role as well. From sensitive tumor cells (IGR-Heu) and following in vitro pressure by specific CTL, we have isolated tumor variants resistant to CTL-mediated killing (IGR-HeuR8) [4]. These variants were characterized by a morphological change visualised by phalloidin staining and resulting in cell rounding (Figure 1). Gene profiling analysis performed by microarray on CTL-resistant and -sensitive tumor cells revealed that two actin related genes, ephrin-A1 and scinderin, were overexpressed in resistant cells [4]. Analysis of the resistant clones revealed unaltered antigen presentation and CTL activation (perforine and granzym B synthesis as well as lytic granule polarisation), ruling out a default of CTL activation by resistant variants. On the other hand, immune synapse formation was loose (Figure 2) and actin polymerization at the cell synaptic contact was reduced (Figure 3) in the resistant variants compared to the sensitive cells that formed a tight synapse.

Figure 1. Morphology and actin cytoskeleton organization of CTL sensitive tumor cells (IGR-Heu) and CTL resistant (IGR-HeuR8) variant. Actin cytoskeleton was stained with 568 Alexa Fluor-Phalloidin and cells were analysed with Zeiss Axiovert 200 inverted fluorescence microscope. From [4].

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Figure 2. Electron microscopic analysis of ultrathin sections of CTL-sensitive and -resistant tumor cell conjugates. (Upper image) After 15 min of contact, very close contacts are observed between the sensitive target and the CTL (see arrows for examples). Note the electron-dense filaments accumulated at the contact points. (Lower image) Resistant targets are observed in close proximity to the CTLs, but contact between points is not very close. Arrowheads: lytic granule N (nucleus). (Scale bars, 200 nm). From (4).

Silencing both ephrin-A11 and scinderin in the resistant variants resulted in decreased resistance to CTL-mediated lysis and restoration of parental morphology and synapse formation [4]. These results highlight the active contribution of actin cytoskeleton in the interaction between target tumor cells and CTL, and suggest that an original immune escape mechanism for tumor cells could be actin disorganisation. These resistant variants have also a decreased phosphorylation of FAK correlating with a decreased focal adhesion formation and a decreased adhesion to extra-cellular matrices. Activation of Rho-GTPases in the resistant cells using a GTPase-activating bacterial toxin (CNF1) restored these defects and also increased susceptibility of the resistant variants to CTL-induced death. Moreover, suppressing FAK expression or inhibiting its phosphorylation in the sensitive cells was able to decrease their susceptibility to CTL [33]. Therefore, FAK activation, usually described to favour tumor progression and survival, has in the context of interaction with CTL an opposite effect resulting in tumor target death. Cytoskeleton remodelling being a mechanism of tumor escapes despite immunogenicity; it should be taken into consideration in anti-tumor therapies due to the possible emergence of resistant variants.

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Figure 3. Microscopic analysis of synaptic polymerized actin. Tumor cells were incubated with autologous CTL for 30 min at 2:1 ratio. After washing to eliminate nonadherent lymphocytes, conjugates were stained with 568 Alexa Fluor–phalloidin and analyzed with a Zeiss Axiovert 200 inverted fluorescence microscope. upper images: true color; Lower images: staining intensity (blue to red, low to high). Data are from one representative experiment of at least thre for each image.From [4].

Actin Cytoskeleton and Tumor Cell Resistance to Therapy Several studies reported that alterations of actin cytoskeleton organisation are associated with several pathologies including cancer. Indeed, regulated expression of some actin-binding proteins has been associated with an increased proliferation and motility of tumor cells [34]. Expression and profiling experiments have led to the identification of transformationassociated actin-binding proteins, such as profilin [35], thymosins [35, 36], gelsolin [37], tropomyosin [38, 39], L-plastin [40, 41] and ABP 280 [42]. It is noteworthy that scaffold proteins such as WASP, zyxin or ezrin have emerged as valuable therapeutic targets. Thus, altered expression of gelsolin, moesin, ezrin, tropomyosin, CAP-G, HSP27, HSP70, TCP-1, and stathmin were associated with in vivo resistance of childhood acute lymphoblastic leukaemia to vincristine [43]. We have also reported that the acquisition of breast adenocarcinoma cell resistance to the cytotoxic effect of TNF-α (Figure 4) was associated with an increase in the expression of Lplastin [44]. In addition, it has been described that patients carrying mutation in WASp exhibit immunological disorders and are prone to cancer. Taken together, these studies support a central role of the actin cytoskeleton in tumor resistance to anticancer therapies (Figure 4).

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Figure 4. Actin cytoskeleton remodeling in breast adenocarcinoma cells following the acquisition of resistance to TNF-α. TNF-sensitive MCF-7 cells and TNF-resistant clone designated 1001 were stained with 488 Alexa-Fluor Phalloidin to visualize F-actin and analyzed using a Zeiss laser scaning confocal microscopy (LSM-510 Meta). MCF-7 cells contained sub-membranous cortical actin, whereas 1001 cells exhibited well-organized actin stress fibres. From [44].

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Conclusion The actin cytoskeleton plays a very important role in the regulation of many cellular functions. Understanding the mechanisms influencing the organization of the actin network within cells is therefore paramount to understand how extracellular and intracellular cues influence the multitude of processes dependent upon actin dynamics. Cytoskeletal remodeling is responsible for cell plasticity and facilitates differentiation, motility and adhesion related functions. In this regard, actin filament structures are established as key determinants in the control of cell shape and migration. In addition it has become clear that the distinctive shape of a cell is dependent not only on the organization of actin filaments but also on proteins that connect the filaments to the membrane. Accumulating evidence indicate that actin cytoskeleton plays a critical role in the activation of T cells including regulation of T cell shape, development and movement and more importantly in the creation of the molecular basis for immunological synapse formation and T cell signalling. Evidence was also provided suggesting that actin cytoskeleton is required for early apoptosis signaling. Various cytoskeleton modifications were found to be associated with malignant cell transformation and have been used as prognostic factors or markers of malignancy in certain epithelial cancers. In this regard, intermediate filaments could be a marker associated with tumor resistance to cytotoxic treatments. In this respect, actin cytoskeleton abnormalities in tumor cells were found to mediate tumor cell resistance to CTL lysis and to TNF. Clearly, microtubule targeting agents may help and could be useful agents to combat cancer.

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[19] Stinchcombe, J. C., G. Bossi, S. Booth, and G. M. Griffiths. 2001. The immunological synapse of CTL contains a secretory domain and membrane bridges. Immunity 15:751761. [20] Barcia, C., C. E. Thomas, J. F. Curtin, G. D. King, K. Wawrowsky, M. Candolfi, W. D. Xiong, C. Liu, K. Kroeger, O. Boyer, J. Kupiec-Weglinski, D. Klatzmann, M. G. Castro, and P. R. Lowenstein. 2006. In vivo mature immunological synapses forming SMACs mediate clearance of virally infected astrocytes from the brain. J. Exp. Med. 203:2095-2107. [21] Grakoui, A., S. K. Bromley, C. Sumen, M. M. Davis, A. S. Shaw, P. M. Allen, and M. L. Dustin. 1999. The immunological synapse: a molecular machine controlling T cell activation. Science 285:221-227. [22] Monks, C. R., B. A. Freiberg, H. Kupfer, N. Sciaky, and A. Kupfer. 1998. Threedimensional segregation of supramolecular activation clusters in T cells. Nature 395:82-86. [23] Lee, K. H., A. D. Holdorf, M. L. Dustin, A. C. Chan, P. M. Allen, and A. S. Shaw. 2002. T cell receptor signaling precedes immunological synapse formation. Science 295:1539-1542. [24] Bunnell, S. C., D. I. Hong, J. R. Kardon, T. Yamazaki, C. J. McGlade, V. A. Barr, and L. E. Samelson. 2002. T cell receptor ligation induces the formation of dynamically regulated signaling assemblies. J. Cell Biol. 158:1263-1275. [25] Yokosuka, T., K. Sakata-Sogawa, W. Kobayashi, M. Hiroshima, A. Hashimoto-Tane, M. Tokunaga, M. L. Dustin, and T. Saito. 2005. Newly generated T cell receptor microclusters initiate and sustain T cell activation by recruitment of Zap70 and SLP-76. Nat. Immunol. 6:1253-1262. [26] Campi, G., R. Varma, and M. L. Dustin. 2005. Actin and agonist MHC-peptide complex-dependent T cell receptor microclusters as scaffolds for signaling. J. Exp. Med. 202:1031-1036. [27] Miletic, A. V., K. Sakata-Sogawa, M. Hiroshima, M. J. Hamann, T. S. Gomez, N. Ota, T. Kloeppel, O. Kanagawa, M. Tokunaga, D. D. Billadeau, and W. Swat. 2006. Vav1 acidic region tyrosine 174 is required for the formation of T cell receptor-induced microclusters and is essential in T cell development and activation. J. Biol. Chem. 281:38257-38265. [28] Kaizuka, Y., A. D. Douglass, R. Varma, M. L. Dustin, and R. D. Vale. 2007. Mechanisms for segregating T cell receptor and adhesion molecules during immunological synapse formation in Jurkat T cells. Proc. Natl. Acad. Sci. USA 104:20296-20301. [29] Varma, R., G. Campi, T. Yokosuka, T. Saito, and M. L. Dustin. 2006. T cell receptorproximal signals are sustained in peripheral microclusters and terminated in the central supramolecular activation cluster. Immunity 25:117-127. [30] DeMond, A. L., K. D. Mossman, T. Starr, M. L. Dustin, and J. T. Groves. 2008. T cell receptor microcluster transport through molecular mazes reveals mechanism of translocation. Biophys. J. 94:3286-3292. [31] Irvine, D. J., M. A. Purbhoo, M. Krogsgaard, and M. M. Davis. 2002. Direct observation of ligand recognition by T cells. Nature 419:845-849.

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[32] Mossman, K. D., G. Campi, J. T. Groves, and M. L. Dustin. 2005. Altered TCR signaling from geometrically repatterned immunological synapses. Science 310:11911193. [33] Abouzahr-Rifai, S., M. Hasmim, H. Boukerche, J. Hamelin, B. Janji, A. Jalil, C. Kieda, F. Mami-Chouaib, J. Bertoglio, and S. Chouaib. 2008. Resistance of tumor cells to cytolytic T lymphocytes involves Rho-GTPases and focal adhesion kinase activation. J. Biol .Chem. 283:31665-31672. [34] Weed, S. A., and J. T. Parsons. 2001. Cortactin: coupling membrane dynamics to cortical actin assembly. Oncogene 20:6418-6434. [35] Clark, E. A., T. R. Golub, E. S. Lander, and R. O. Hynes. 2000. Genomic analysis of metastasis reveals an essential role for RhoC. Nature 406:532-535. [36] Pedersen, M. W., T. Thykjaer, T. F. Orntoft, L. Damstrup, and H. S. Poulsen. 2001. Profile of differentially expressed genes mediated by the type III epidermal growth factor receptor mutation expressed in a small-cell lung cancer cell line. Br. J. Cancer 85:1211-1218. [37] Brandt, B. H., A. Roetger, T. Dittmar, G. Nikolai, M. Seeling, A. Merschjann, J. R. Nofer, G. Dehmer-Moller, R. Junker, G. Assmann, and K. S. Zaenker. 1999. c-erbB2/EGFR as dominant heterodimerization partners determine a motogenic phenotype in human breast cancer cells. FASEB J. 13:1939-1949. [38] Gabbiani, G., J. Csank-Brassert, J. C. Schneeberger, Y. Kapanci, P. Trenchev, and E. J. Holborow. 1976. Contractile proteins in human cancer cells. Immunofluorescent and electron microscopic study. Am. J. Pathol. 83:457-474. [39] Bharadwaj, S., and G. L. Prasad. 2002. Tropomyosin-1, a novel suppressor of cellular transformation is downregulated by promoter methylation in cancer cells. Cancer Lett. 183:205-213. [40] Leavitt, J. 1994. Discovery and characterization of two novel human cancer-related proteins using two-dimensional gel electrophoresis. Electrophoresis 15:345-357. [41] Lapillonne, A., O. Coue, E. Friederich, A. Nicolas, L. Del Maestro, D. Louvard, S. Robine, and X. Sastre-Garau. 2000. Expression patterns of L-plastin isoform in normal and carcinomatous breast tissues. Anticancer Res. 20:3177-3182. [42] Cunningham, C. C., J. B. Gorlin, D. J. Kwiatkowski, J. H. Hartwig, P. A. Janmey, H. R. Byers, and T. P. Stossel. 1992. Actin-binding protein requirement for cortical stability and efficient locomotion. Science 255:325-327. [43] Verrills, N. M., N. L. Liem, T. Y. Liaw, B. D. Hood, R. B. Lock, and M. Kavallaris. 2006. Proteomic analysis reveals a novel role for the actin cytoskeleton in vincristine resistant childhood leukemia--an in vivo study. Proteomics 6:1681-1694. [44] Janji, B., L. Vallar, Z. Al Tanoury, F. Bernardin, G. Vetter, E. Schaffner-Reckinger, G. Berchem, E. Friederich, and S. Chouaib. 2010. The actin filament cross-linker L-plastin confers resistance to TNF-alpha in MCF-7 breast cancer cells in a phosphorylationdependent manner. J. Cell Mol. Med. 14:1264-1275.

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In: Actin: Structure, Functions and Disease Editors: V. A.Consuelas et al. pp. 257-269

ISBN: 978-1-62100-191-1 © 2012 Nova Science Publishers, Inc.

Chapter IX

Insight into Force Transmission along Actin Filaments: Sliding Movement of Actin Filaments Containing Inactive Components on Myosin Molecules Syunsuke Matsushita and Kuniyuki Hatori* Department of Biosystem Engineering, Graduate School of Science and Engineering, Yamagata University, Yonezawa, 992-8510 Japan.

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Abstract The sliding movement of actin filaments, consisting of heterogeneous components, on skeletal muscle myosin molecules was examined to specifically evaluate the effect of internal modulation of the actin filaments for force transmission on the sliding movement. Inactive actin molecules were prepared by conjugation with indocarbocyanine fluorescent dyes (IC3-OSu or Cy3-NHS) in molar ratios greater than a 3-fold excess. IC3-OSu is an analogue of Cy3-NHS, and it can bind to primary amino groups. IC3conjugated actin (IC3-actin) monomers were polymerized into the filaments which led to complete impairment of both motile activity and myosin-ATPase activation. Filaments of Cy3-conjugated actin (Cy3-actin) exhibited a decrease in velocity to a third of the value (33%) observed for intact actin filaments. In the absence of ATP, dissociation rates of IC3- and Cy3-actin filaments from myosin molecules were greater than those of intact actin filaments, indicating that IC3- and Cy3-actin act as smaller resistance components against sliding movement compared to intact filaments. Subsequently, two types of copolymer filaments were prepared. The first type of copolymer were filaments copolymerized homogeneously with intact actin monomers and IC3-actin monomers, while the second kind were block copolymer filaments composed of two short filaments *

To whom correspondence should be addressed: Kuniyuki Hatori, Phone: +81 238 26 3727. Fax: +81 238 26 3727. E-mail address: [email protected]

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Syunsuke Matsushita and Kuniyuki Hatori of intact actin and IC3-actin. The sliding velocities of these copolymer filaments hyperbolically decreased as the fraction of IC3-actin monomer increased. In practice, 75% IC3-actin within homogeneous copolymer was required to reduce the velocity by half. In the case of block copolymer 65% IC3-actin led to the same decrease in velocity. For Cy3-actin copolymer filaments similar differences between homogeneous and block copolymer filaments were also observed. Drag ratio between IC3-actin (or Cy3-actin) and intact actin was estimated by consideration of the force balance between the power force and the drag force imposed on a filament during steady movement. Consequently, the drag ratios of IC3-actin to intact actin were 0.31 (homogeneous copolymer) and 0.47 (block copolymer), respectively. Thus, IC3-actin incorporated homogeneous copolymers exhibits a smaller resistance to sliding movement than IC3-actin modified block copolymers.

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Introduction Actin filaments exhibit flexible motion such as bending during sliding movement along myosin molecules fixed on a glass surface with ATP hydrolysis [1, 2]. In addition, the internal mobility of actin constituents within the filament, which are fundamental to filamental flexibility, has been revealed by a number of spectroscopic methods [3–5]. Kim et al. have reported that inter-monomer cross-linking among actin monomers by chemical crosslinkers inhibits the sliding motility of actin filaments without affecting ATPase activation [6]. Also, internal cross-linking of an actin monomer has been found to reduce the sliding motility of the filaments [7]. Therefore the internal mobility within the filament may be essential to coordinating the sliding movement along myosin molecules. Furthermore a filament structure composed of actin monomers may mediate the force transmission from myosin molecules. We have previously demonstrated that transversal fluctuations occurring in actin filaments during the sliding movement propagate along the longitudinal axis of the filament at a faster rate compared with the sliding velocity [8]. Propagation of filamental fluctuations may be generated by progressive mitigation of filamental distortions resulting from receiving of active forces of myosin power strokes. In this situation, association and dissociation of myosin molecules with/from actin filaments are likely to occur with cooperative effects in temporal and spatial dimensions. In fact, cooperativity of actin filaments is observed in the structural changes themselves and in the binding of myosin and tropomyosin molecules to the filaments [9–11]. In eukaryotic cells, actin filaments are a platform for binding of various proteins in addition to myosin molecules. The functional and structural nature of actin filaments is dependent on the operation of these binding proteins [12, 13]. If heterogeneous binding of different type of protein takes place on an actin filament, the question is raised of how the force transmitted along the actin filament with spatial modulation so that sliding movement is performed. Two key topics are discussed in this chapter: the first aspect is describes how both motile activity and ATPase activation of actin can be decrease by excessive conjugation of fluorescence dyes to actin monomers. Secondly, we report the effects of internal defects along the filaments upon the sliding velocity of entire filaments. An increase of defects leads to a reduction in sliding velocity. The influence of different arrangements of actin components upon the motile properties of the filament is also examined. It appears that nature of

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Insight into Force Transmission along Actin Filaments

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neighboring molecules has a significant impact on the force transmission with cooperative effects.

Reagents and Fluorescent Dyes

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N-Ethyl-N‟-[5-(N”-succinimidyloxycarbonyl)pentyl]indocarbocyanine chloride (IC3OSu) and Cy3-NHS were purchased from Dojindo Laboratories and from GE Healthcare, respectively (Figure 1). These dyes (0.1 mg) were dissolved in 40 µL of DMSO, and were stored at –30oC. Tetramethylrhodamine-phalloidin (TMR-phallodin) was obtained from Fluka. Actin and myosin were extracted from rabbit skeletal muscle. Actin was purified by the method of Spudich and Watt [14]. Myosin was purified by the method of Perry, and then was digested with alpha-chymotrypsin for the preparation of heavy meromyosin (HMM) [15]. Other reagents were special reagent grade.

Figure 1. Structure of IC3-OSu (A) and Cy3-NHS (B). IC3-OSu was obtained from Dojindo Laboratories and Cy3-NHS from GE Healthcare. The molecular weights of IC3-OSu and Cy3-NHS are 604.18 and 765.95, respectively. IC3 is more hydrophobic than Cy3 because it is lacking in sulfonate groups.

Preparation of Actin Monomers Conjugated with Fluorescent Dyes Filamentous actin solutions (1 mg/mL) were prepared in 0.1 M KCl, 2 mM MgCl2, 50 mM NaHCO3 solutions. In excess of 6-fold molar equivalents of IC3-OSu per actin monomer was added to the above F-actin solution, and the resulting mixture was incubated for 90 min.

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at 25oC. The reaction was terminated by addition of 0.1 vol. of 2 M Tris-HCl (pH 7.5). After dialysis of the mixture against G-buffer (2mM Tris-HCl (pH 8.0), 0.1 mM CaCl2, 0.2 mM ATP, 0.05% NaN3) for 2 days, unbound dyes were excluded by gel-filtration. Finally actin and bound dye concentrations were estimated from absorbances at 290, 310, and 552 nm by UV-spectrometry (Shimazu, UV-1200). The values of molar extinction coefficient for IC3 at 290, 310, and 552 nm were 5760, 2720 and 51300, respectively. Cy3-conjugated actin monomers were also prepared by similar procedure described for IC3-conjugated actin. We employed both IC3-conjugated actin at a 6-fold molar ratio (IC3-actin) and Cy3-conjugated actin at a 3-fold molar ratio (Cy3-actin).

Preparation of Heterogeneous Actin Filaments

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Two kinds of actin filaments were prepared: (a) Block copolymer filaments consisting of short filaments of intact actin fragments and either IC3- or Cy3-actin fragments. IC3- or Cy3actin filaments and intact actin filaments were stabilized by phalloidin and TMR-phalloidin, respectively. After mixing in a 1:1 ratio, the filaments were fragmented ultra-sonically. Heterogeneous block copolymer filaments were developed for annealing of short fragments to each other. It was possible to determine the IC3-actin content of an entire filament because the IC3-actin portions were bright due to excess of fluorescent dyes. This allowed them to be clearly distinguished from TMR-phalloidin-labeled actins under a fluorescence microscope. (b) The second filament prepared was a homogeneous copolymer. It could be obtained by mixing intact actin monomers and IC3- or Cy3-actin monomers at various molar ratios prior to filamentous formation.

Measurement of Dissociation Rate of an Actin Filament from a HMM Molecule In the absence of ATP, the dissociation rate of an actin filament from a HMM molecule fixed on a glass surface was recorded under a fluorescent microscope (Nikon, Diaphoto-TMD equipped with TMD-EF2, objective DIC 100x oil and Omega, optical interference filters XF101-2). HMM solution (1 µg/mL) was dispersed on the collodion-coated slide glass. After 60 s, the solution was replaced with 10 mg/mL of BSA solution. Short actin filaments prepared by ultrasonication were then added. The final conditions were 25 mM KCl, 25 mM imidazole-HCl (pH 7.4), 4 mM MgCl2, 0.5% 2-mercaptoethanol, 0.02 mg/mL catalase, 0.1 mg/mL glucose oxidase, 6 mg/mL glucose and 0.05 mg/mL hexokinase. The fluorescent images from a highly sensitive camera (Hamamatsu, C2400-08) were recorded on a PC (Apple Co., Power Mac G3) using a video grabber board (Scion Co., LG-3). The dwell time from the binding of an actin filament to an HMM molecule to its subsequent detachment was measured. Two hundred filaments were recorded under each condition. The dissociation rate constants (koff) for three kinds of actin filament were determined by an exponential plot of the frequency distribution (N) versus dwell time (t) where, N(t) = N0 exp ( – koff t).

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Insight into Force Transmission along Actin Filaments

Measurement of ATPase Activity ATP hydrolysis was monitored by measuring the concentration of inorganic phosphate (Pi) using a malachite green method [16]. Phalloidin-stabilized actin filaments (0.05 mg/mL) of IC3-actin and intact filaments were incubated at 25oC under the following conditions: 25 mM KCl, 25 mM imidazole-HCl (pH 7.4), 4 mM MgCl2, and 2 mM ATP. HMM molecules (0.05 mg/mL) were added to the actin solution to initiate the reaction. At intervals of 10 min, an aliquot (0.1 mL) of the reaction solution was mixed with 0.1 mL of 0.6 M PCA to terminate the ATPase. After a centrifugation process, 0.1 mL of the supernatant was added to 2.0 mL of the malachite green reagent (0.15 g/l malachite green, 6.15 g/l disodium molybdate (VI) dihydrate, 0.25 g/l triton-X, 0.5 M HCl, 0.15 M PCA) and 0.2 mL of citric acid (34% w/v) was then added to the mixture. After incubation for 15 min at 25oC, the absorbance of the sample at 650 nm was determined by spectrometry. The ATPase activity was recorded as the molar ratio of Pi released to HMM molecule per second. Actin-activated ATPase activity was the differences between ATPase activity in the presence and absence of actin.

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Motility Assay Sliding velocities of various types of actin filaments were determined using an in vitro motility assay [17]. HMM solution of 0.05 mg/mL concentration was perfused on a collodion-coated slide glass. After removal of unbound HMM, actin filaments were added to HMM-fixed surface. Sliding movement of actin filaments was initiated by addition of ATP solution (25 mM KCl, 25 mM imidazole-HCl (pH 7.4), 4 mM MgCl2, 0.5% 2mercaptoethanol, 3 mg/mL glucose, 0.02 mg/mL catalase, 0.1 mg/mL glucose oxidase, and 2 mM ATP) and observed under a fluorescence microscope. The sliding velocity of the actin filaments was determined by measuring the moving distance at 0.5 s intervals using image analysis software (NIH, ImageJ). In each case, the sliding velocity of actin filaments was determined by averaging 100 independent samples. Spacing between the nearest-neighbor pixels in the image was 0.083 µm.

Simple Model for Sliding Movement of Actin Filaments Containing Heterogeneous Components We assumed that the sliding velocity is dependent on the driving force exerted from myosin molecules and on the drag force derived from binding between actin and myosin. At steady state this velocity becomes constant, and total amount of driving forces is equal to total amount of drag forces as follows: fn (1 – p) + fm p = (gn (1 – p) + gm p) V.

(1)

In eq. (1), f and g denote the driving force and drag coefficient, respectively. Subscripts n and m denote components for intact actin and for modified actin (IC3- or Cy3-actin), Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

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respectively. The fraction of modified actin in the entire actin constituents of the filament is represented by p. solving for the sliding velocity V gives, V = (fn/gn)((1 + p( – 1))/(1 + p( – 1)))

(2).

Parameter  is defined as the ratio of driving forces (fm/fn) and  is the ratio of drag coefficients (gm/gn).

Binding Ratio of Fluorescence Dye to Actin

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Actin filaments were mixed with IC3-OSu amino reactive dyes in various molar ratios. IC3-OSu compounds bound to actin constituents in the filament up to a molar ratio of 6.0. In the case of Cy3-NHS, a binding molar ratio of up to 3.0 was observed when the actin and dye were mixed in ratio of 6.0. Figure 2 illustrates the intense brightness of actin filaments conjugated with either IC3 in 6-fold excess or with Cy3 in 3-fold excess compared to TMRphalloidin-labeled filaments under a fluorescent microscope. At present we have not identified the binding sites on the actin for these dyes. Lu et al. have already determined the reactivity of lysine residues, which possess a primary amino group, for filamentous actin [18]. They have reported that the order of reactivity of lysine residues is K336, K328, K238, K326, K191, and K84, which are candidates for the binding sites of compounds containing reactive amino moieties.

Figure 2. Fluorescent photographs of filaments of TMR-phalloidin-bound actin at 1:1 (A), IC3conjugated actin at 6:1 (B) and Cy3-conjugated actin at 3:1 (C). These photographs were taken under the same conditions (gain, exposure time, and sensitivity). Actin filaments, in the absence of ATP, were bound to HMM molecules fixed on a glass surface. Scale bar indicates 10 µm. Actin: Structure, Functions and Disease : Structure, Functions, and Disease, edited by Victoria A. Consuelas, and Daniel J. Minas, Nova Science

Insight into Force Transmission along Actin Filaments

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Influence of Binding of Fluorescence Dye on the Motility of Actin Filaments

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

The sliding velocity of the filaments decreased with an increase of binding ratio in both dyes. Figure 3 shows that above molar ratio of 3 the movement of IC3-actin filaments was completely suppressed while these filaments were bound to HMM molecules fixed on glass surface. On the other hand, Cy3-actin filaments at a 3-fold molar ratio the sliding velocity decreased to a value of 33% of the original level. The average length of the filaments was almost comparable to the proportion of intact filaments. Thus, the formation of filaments was not inhibited by the conjugation of IC3- or Cy3-dyes.

Figure 3. Sliding velocities of filaments of IC3-actin (filled circles) and Cy3-actin (open circles) at various binding ratios.

Binding Affinity of IC3- or Cy3-Actin for HMM Molecule When HMM molecules are bound to a glass surface at low density, a short actin filament