Cell Movement : New Research Trends [1 ed.] 9781608766642, 9781606925706

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Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved. Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved. Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

CELL MOVEMENT: NEW RESEARCH TRENDS

Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved.

No part of this digital document may be reproduced, stored in a retrieval system or transmitted in any form or by any means. The publisher has taken reasonable care in the preparation of this digital document, 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 herein. This digital document is sold with the clear understanding that the publisher is not engaged in rendering legal, medical or any other professional services.

Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved. Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

CELL MOVEMENT: NEW RESEARCH TRENDS

T. ABREU AND

G. SILVA Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved.

EDITORS

Nova Biomedical Books New York

Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

Copyright © 2009 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.

Copyright © 2008. Nova Science Publishers, Incorporated. All rights reserved.

Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Library of Congress Cataloging-in-Publication Data Cell movement : new research trends / [edited by] T. Abreu & G. Silva. p. ; cm. Includes bibliographical references and index. ISBN 978-1-60876-664-2 (eBook) 1. Cells--Motility. I. Abreu, T. (Thomas), 1967- II. Silva, G. (Gregorio), 1966[DNLM: 1. Cell Movement. 2. Cell Differentiation. 3. Neoplasm Metastasis. 4. Neoplasms-metabolism. QU 375 C3934 2009] QH647.C463 2009 571.6'7--dc22 2008047198

Published by Nova Science Publishers, Inc.

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Contents Preface

vii

Research and Review Chapters

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Chapter I

Scaffolding Proteins that Regulate the Actin Cytoskeleton in Cell Movement S.J. Annesley and P.R. Fisher

1

Chapter II

Systems Dynamics Behind Cell Movement Mikiya Otsuji and Shinya Kuroda

51

Chapter III

Mast Cells in Injury Response Stefano Bacci, Aurelio Bonelli and Paolo Romagnoli

81

Chapter IV

Role of Rho GTPases in Tumor Cell Migration and Metastasis Yong Tang and Daotai Nie

123

Role of Serine Proteases and Their Receptors in Cellular Motility V.M. Shpacovitch, M.D. Hollenberg and M.Steinhoff

159

Tuberin and Hamartin in Moving Breast Cancer Cells: Expression, Localization, and Function Marina A. Guvakova and William S.Y. Lee

187

Chapter V

Chapter VI

Chapter VII

Role of Chemokines in Colorectal Cancer and Metastasis Kathrin Rupertus, Otto Kollmar, Michael D. Menger and Martin K. Schilling

Chapter VIII

The Proteoglycan Versican: An Important Regulator of Cell Locomotion in Development and Disease Carmela Ricciardelli, Andrew Sakko, Miranda Ween, Darryl Russell and David Horsfall

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209

227

Contents

vi Chapter IX

Chapter X

The Role of Endometrial Stromal Cell-Mediated Contractility in Endometrial Tissue Remodeling Kaei Nasu, Akitoshi Yuge, Harunobu Matsumoto, Akitoshi Tsuno and Hisashi Narahara The Role of Transmission Electron Microscopy Analysis in Different Cases of Reduced Motility in Human Spermatozoa Giulia Collodel, Nicola Antonio Pascarelli, Elena Moretti

Chapter XI

Reproducing In Vivo Cell Migration Lilian Soon and Alastair Stewart

Chapter XII

Chlamydomonas as the Unicellular Model for Chemotaxis and Cellular Differentiation E.V. Ermilova

247

265

295

313

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Short Communications A

Fluorescence Recovery after Photobleaching Macarena Peran, Fernando Rodriguez-Serrano, Houria Boulaiz, Juan Antonio Marchal, Elena Lopez, Manuel Picón, Pablo Alvarez, Ana R. Rama, Raúl Ortiz, Alberto Ramirez and Antonia Aranega

331

B

Improvement in Sperm Cell Kinetics Fábio F. Pasqualotto, Luana Venturin Lara and Eleonora Bedin Pasqualotto

345

Index

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355

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Preface Cell movement is a complex phenomenon primarily driven by the actin network beneath the cell membrane, and can be divided into three general components: protrusion of the leading edge of the cell, adhesion of the leading edge and deadhesion at the cell body and rear, and cytoskeletal contraction to pull the cell forward. Each of these steps is driven by physical forces generated by unique segments of the cytoskeleton. This review examines the specific physics underlying these phases of cell movement and the origins of the forces that drive locomotion. Cell movement or motility is a highly dynamic phenomenon that is essential to a variety of biological processes such as the development of an organism (morphogenesis), wound healing, cancer metastasis and immune response. For example, during morphogenesis there is a targeted movement of dividing cells to specific sites to form tissues and organs. For wound healing to occur, cells such as neutrophils (white blood cells) and macrophages (cells that ingest bacteria) move to the wound site to kill the microorganisms that cause infection, and fibroblasts (connective tissue cells) move there to remodel damaged structures. This new book presents important research in the field from around the globe. Chapter I - Actin is the main component of the microfilament system in all eukaryotic cells and is essential for most intra- and inter-cellular movement including muscle contraction, cell movement, cytokinesis, cytoplasmic organisation and intracellular transport. The polymerisation and depolymerisation of actin filaments in nonmuscle cells is highly regulated and the reorganisation of the actin cytoskeleton can occur within seconds after chemotactic stimulation. There are many proteins which are involved in the regulation of the actin cytoskeleton. These include receptors which receive chemotactic stimuli, G proteins, second messengers, signalling molecules, kinases, phosphatases and transcription factors. These proteins are varied and numerous and are involved in multiple pathways. Despite the large number of proteins, there are not enough to coordinate the various responses of the cytoskeleton. An additional level of regulation is conferred by scaffolding proteins. Due to the presence of numerous protein interaction domains, scaffolding proteins can tether various proteins to a certain location within the cell to facilitate the rapid transfer of signals from one protein to the next. This colocalisation of the components of a particular pathway also helps to prevent unwanted crosstalk with components of other pathways. Tethering receptors, kinases, phosphatases and cytoskeletal components to a particular

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T. Abreu and G. Silva

location within a cell helps ensure efficient relaying and feedback inhibition of signals to enable rapid activation and inactivation of responses. Scaffolding proteins are also thought to stabilise the otherwise weak interactions between particular proteins in a cascade and to catalyse the activation of the pathway components. There are numerous scaffolding proteins involved in the regulation of the cytoskeleton and this chapter has focussed on examples from several groups of scaffolding proteins including the MAPK scaffolds, the AKAPs, scaffolds of the post synaptic density and actin binding scaffolding proteins. Chapter II - Recent studies have shown that the interactions of various molecules are responsible for cell movement. Despite a consensus that systems of molecular interactions mediate the various dynamic properties of cells, it is still intuitively difficult to understand the ‘systems dynamics’ underlying cell movement. One way to visualize these dynamics is to use a systems-biological approach. The cause-result relationship between molecular interactions and cell movement can be understood as differential equations and their solution. Conceptual models, rather than detailed models, are often of greater use for extracting the essence of systems dynamics. In the first section, we review several mathematical conceptual models proposed for the analysis of cell polarity in migrating cells. The models are classified into four groups: the ‘LEGI + MSNL system’, the ‘CTSP system’, the ‘MCRD system’ and the ‘Meinhardt system.’ Chemotactic cells, even under normal conditions, do not always migrate with a stable front-back axis, but rather exhibit dynamic behaviors, depending on the conditions. In the second section, we show that MCRD systems composed of multiple species generate dynamic properties, which, in fact, can be observed in chemotaxing cells. The MCRD models with multiple species of molecules shed light on systems dynamics behind cell movement. Chapter III - Mast cells are bone marrow derived cells capable of secreting many active molecules, ranging from the mediators stored in specific granules, some of which are known since several decades (histamine, heparin), to small molecules produced immediately upon stimulation (membrane lipid derivatives, nitric oxide), to a host of constitutively secreted, multifunctional cytokines. With the aid of a wide array of mediators the activated mast cells control the key events of inflammation and healing and participate to the regulation of local immune response. On the basis of the structure, origin, principal subtypes, localization and function of these cells, their involvement in injury repair is therefore to be considered in acute and chronic conditions respectively. The importance of mast cells in regulating healing processes is underscored by the proposed roles of a surplus or a deficit of their mediators in the formation of exuberant granulation tissue (such as keloids and hypertrophic scars), the delayed closure or dehiscence of wounds and the transition of acute to chronic inflammation. Chapter IV - Small GTPase Rho signaling pathways regulate the growth, motility, invasion and metastasis of a variety of cancer cells. Aberrant Rho signaling, as results from alterations in the levels of Rho GTPase proteins, the status of activation, or the abundance of effector proteins, endows cancer cells with elevated invasive and metastasis capability. Alterations of Rho signaling particularly impact the cytoskeleton, whose organization and reorganization underpin the motility of cancer cells during the invasive growth and metastasis process. Progress is being made to elucidate the underlying mechanisms by which Rho GTPases activate the downstream signaling effectors. Further investigations are required for

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development of novel tumor therapeutic strategies targeting the Rho GTPase signaling pathways to inhibit invasion and metastasis of cancer cells. Chapter V - Serine proteinases constitute a family of proteolytic enzymes with diverse biological functions ranging from the digestion of extracellular matrix and dietary proteins to complement formation. However, recent groundbreaking studies revealed a role of certain serine proteinases as signalling molecules which are able to regulate cell function via cell surface receptors. Urokinase plasminogen activator receptor (uPAR) and proteinase-activated receptors (PARs) are two receptor types via which serine proteinases can trigger intracellular signalling cascades. Currently, the role of these receptors in serine proteinase-mediated signalling is under intensive investigation. Both types of receptors have been reported to influence the migratory behaviour of different cells under physiological and pathophysiological conditions. Among these cells are human leukocytes, smooth muscle cells, sperm, and invading cancer cells. Moreover, activation of uPAR as well as PARs is known to affect the expression of cell adhesion molecules, actin polymerisation and to induce cell shape changes, subsequently acting on cell movement. Thus, serine proteinases are capable of regulating cell motility via two major pathways. By one pathway, the enzymes induce changes in cell microenvironment (for example, via digestion of extracellular matrix proteins). By the other pathway, serine proteases may directly trigger the signalling cascades resulting in changes of cell migratory behaviour. Understanding of mechanisms involved in both pathways could lead to new therapeutic approaches in inflammatory, allergic, and neoplastic diseases. Chapter VI - While the role of hamartin and tuberin complex as a negative regulator of protein translation and cell growth is well established, little information is available about the role of these proteins in regulating cell migration [1]. Hamartin, the protein encoded by the tuberous sclerosis complex (TSC) 1 gene, has been implicated in cell-matrix adhesion through its interaction with the ezrin-radixin-moesin (ERM) family of actin-binding proteins and regulation of the GTPase Rho [2]. Tuberin, encoded by the TSC2 gene, shows homology with Rap1 GTPase activating protein (GAP) and acts as a Rap1GAP in vitro [3]. Rap1, a member of the Ras oncogene superfamily, plays a role in the control of cell adhesion and movement [4]. Here, by laser-scanning confocal microscopy, we revealed that hamartin is present, whereas tuberin is excluded from lamellipodia of moving breast cancer cells stimulated with insulin-like growth factor I (IGF-I). The dissimilar intracellular patterns suggest that hamartin and tuberin may have independent roles in the moving cells. Hence we tested the hypothesis that tuberin regulates intracellular activity of Rap1 and thus might contribute to cell motile behavior. We showed that a partial knockdown of tuberin expression by TSC2 siRNA increased GTP-loading of Rap1 in response to IGF-I. We then utilized a set of the tuberin mutants, containing or lacking a putative GAP domain, to investigate if tuberin decreases levels of active GTP-bound Rap1. As expected, tuberins with a truncated GAP domain had no effect on Rap1 activation by IGF-I. Surprisingly, over-expression of the fulllength tuberin had no effect on Rap1 either, whereas expression of a deletion mutant of tuberin, lacking the hamartin-binding domain (HBD) spanning a leucine-zipper (amino acids 81-98) and a coiled-coil (amino acids 346-371), displayed Rap1GAP function. Further deletion of HBD and the central region of tuberin, including the coiled-coil (amino acids 1008-1021) failed to promote hydrolysis of Rap1-bound GTP. These data reveal that the

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region spanning amino acid residues 1-460 inhibits, whereas both the central regions of tuberin (amino acids 461- 1114) and the GAP domain (amino acids 1531-1758) promote Rap1 GTPase function of tuberin in vivo. We propose that tuberin may act as a Rap1GAP in the moving breast cancer cells; however, the interaction of hamartin with tuberin inhibits tuberin’s GAP activity towards Rap1. Chapter VII - Background: Chemotactic cytokines (chemokines) are inflammatory cytokines that stimulate the migration of distinct subsets of cells, including leukocytes, lymphocytes and tumor cells. The chemokines MIP (macrophage inflammatory protein)-2 and SDF (stromal cell-derived factor)-1, which are both members of the CXC-chemokine superfamily, are supposed to have an important impact on tumor progression and metastasis. In our recent studies we investigated the chemotactic response of CT26.WT colorectal cancer cells on MIP-2 and SDF-1 in vitro and the influence of these chemokines on tumor growth in vivo. Material and methods: Using flow cytometry and chemotaxis chambers, we investigated the expression of the chemokine receptors CXCR2 and CXCR4 in vitro and the chemotactic response of CT26.WT colon carcinoma cells. To determine the influence of chemokines on tumor growth in vivo, we analyzed the growth of CT26.WT-tumors after exposure to MIP-2 and SDF-1. Additionally, we investigated the growth characteristics of CT26.WT tumors which were implanted into the left liver lobe of BALB/c mice after exposure to MIP-2 or after treatment with a neutralizing anti-MIP-2 antibody. MIP-2 and tumor growth: Our studies revealed that in vitro 98.8% of the CT26.WT cells were CXCR-2 receptor positive and showed a dose-dependent migration along a MIP-2 gradient. MIP-2 also promoted angiogenesis, hepatic engraftment and tumor growth of liver metastases, as well as tumor growth of established extrahepatic tumors. Blockade of MIP-2 inhibited the augmentation of angiogenesis and metastatic tumor growth of intrahepatic tumors after liver resection and suppressed engraftment of CT26.WT tumor cells at extrahepatic sites. These results indicate a significant role of the chemokine MIP-2 during tumor engraftment, progression and metastasis and point out to the promising approach of targeting the MIP-2/CXCR2 pathway for anti-tumor therapy. SDF-1 and tumor growth: Despite a lower surface expression of the SDF-1 receptor CXCR4 (31.5%), CT26.WT cells showed a significant and dose-dependent migration in response to a SDF-1 gradient. Exogenously applied SDF-1 was capable to stimulate the growth of already established extrahepatic CT26.WT tumors. It also induced an angiogenic response with persistent changes in tumor vasculature. These results also encourage for further investigation on the influence of chemotactic cytokines in tumor growth and may also be useful to elucidate the pathophysiology of other diseases which are related to angiogenesis and cell migration. Conclusion: The results of our studies demonstrate that chemotactic signaling does not only contribute to the metastatic spread of tumor cells but is also essential for tumor engraftment and progression. Therefore, chemotactic cytokines which are a part of the tissuespecific environment could be promising targets for anti-tumor therapy. Chapter VIII - One of the major keys to understanding tissue reorganization during embryological development and pathophysiology will be to decipher the mechanisms that regulate cell locomotion. This review focuses on what is known about versican, a large

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extracellular proteoglycan intimately involved in cellular relocation and tissue reorganization during normal processes and pathological states, including cancer metastasis. A member of the lectican family, versican plays diverse roles in cell adhesion, proliferation, migration and angiogenesis. These wide ranging functions have been attributed to the N-(G1) and C-(G3) terminal globular domains and the central glycosaminoglycan-binding region of versican, which collectively interact with a large number of extracellular matrix and cell surface structural components. Consequently, the diverse roles of versican may depend on the level of specific isoforms and/or levels of G1 and G3 domains which can be generated as a result of processing by specific proteases. Little is known about versican catabolism and whether its proteolytic fragments have altered biological activity compared to the intact molecule in physiological and disease processes. Studies over the last ten years have confirmed a significant role for versican in regulating cell locomotion in normal development and disease. Chapter IX - The human uterine endometrium is a dynamic organ that undergoes cyclic phases of remarkable periodic growth, remodeling, and breakdown throughout the reproductive years. The endometrial tissue remodeling, which depends on the menstrual cycle, is thought to be regulated by ovarian steroids, various cytokines, neuropeptides, and growth factors that are produced and secreted in an endocrine, a paracrine, and/or an autocrine manner. An important event during endometrial tissue remodeling is the contraction of connective tissue, which is carried out by the endometrial stromal cells. Through the use of threedimensional collagen matrices, investigations on endometrial stromal cell biology have provided new opportunities to understand the reciprocal and adaptive interactions between the cells and the surrounding matrix in a tissue-like environment. Such interactions are integrated with the regulation of endometrial tissue morphogenesis and characterized as endometrial tissue homeostasis during the cyclic tissue remodeling. This paper is a review of the recent information concerning the mechanism of endometrial tissue remodeling during menstruation, with a focus on the contractility of endometrial stromal cells. Chapter X - The aim of this review is to highlight the pivotal role of transmission electron microscopy (TEM) analysis in the diagnosis of human sperm pathologies related to flagellum and affecting motility. The analysis of sperm motility plays a central role in the evaluation of male fertility, as it is known that a high percentage of poorly motile or immotile sperm will not be able to fertilize. The study of sperm flagellum and its abnormal forms is generally carried out by light microscope with obvious technical limitations. Only the use of TEM allows for the performance of an ultrastructural evaluation of sperm flagellar assembly with a precise characterization of anomalies, also extrapolating the functional aspect. Severely reduced or completely absent sperm motility in subfertile or infertile men is associated with submicroscopic alterations in the cytoskeletal structure of sperm flagellum, in the mitochondria structure and assembly, in the axonemal pattern and periaxonemal accessory structures, such as the outer dense fibres and fibrous sheath. Sperm anomalies are classified as non-specific, or non-systematic sperm defects, or as systematic sperm defects. The first and most frequent type is related to a heterogeneous combination of randomly distributed alterations affecting the head and the tail organelles in a varied percentage in sperm ejaculate.

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These alterations can be related to andrological pathologies or to other endogenous or exogenous factors. Necrozoospermia, an extreme manifestation of asthenozoospermia, is, a rather rare phenomenon characterized by an increase in non-viable sperm up to the persistent presence of 100% dead spermatozoa in different ejaculates from the same patient. TEM has proved to be the only tool able to specifically characterize the morphological features of the systematic sperm defects in which an identical and specific alteration affects the vast majority of the sperm population in sterile patients. These defects tend to show family clustering and are significantly more frequent in individuals with a history of consanguinity. The most documented sperm defects of supposed genetic origin affecting the tail structures include: “Dysplasia of Fibrous Sheath”, “Primary Ciliary Dyskinesia”, “Detached tail”, “Absence of fibrous sheath”, “Absent axoneme”. Chapter XI - The imaging field has provided unprecedented insight into the behaviours of cancer cells in tumours particularly, cell migration. From in vivo imaging data, we now understand that the heterogeneity of cancer cells, the microenvironment of the tumour, and host cell participation have an impact on cell migration with significant implications on how we study these cells in vitro. Cell migration performed on two-dimensional surfaces has provided many breakthroughs in the elucidation of the molecular regulation of the distinct steps of migration from sensing of the extracellular milieu to protrusion, adhesion and translocation. There is evidence that these studies are relevant to some in vivo situations for example, the in vivo display of the steps of the chemotaxis cycle. There is also a trend towards creating or revisiting methods that emulate specific in vivo conditions for cell migration taking into account for example, linear trekking along fibres and vessels, the multidimensionality of tumours, tumour cell characteristics such as chemokinetic and chemotactic movements and amoeboid-like and mesenchymal-like migration. In order to meet the environmental criteria, specific approaches are required and include the use of intravital imaging, 3D spheroids as models, micropatterning of substrate, gradient-generating devices and fibrillar collagen. Chapter XII - Chlamydomonas has long been one of the most successful unicellular organism for genetic and biochemical studies of the photosynthesis, organelle genomes and flagellar assembly. The availability of the new molecular genetic techniques is increasing interest in Chlamydomonas as a model system for research in areas like swimming behavior where it previously has not been widely exploited. The swimming behavior of Chlamydomonas reinhardtii is influenced by several different external stimuli including chemical attractants. Chemotaxis of the green alga is altered during gametic differentiation. Gametogenesis results in the conversion of chemotactically active vegetative cells into chemotactically inactive gametes. This experimental system offers the opportunity to study cellular behavior and differentiation at the molecular level with use of a wide range of molecular genetic approaches, including gene tagging by insertional mutagenesis, quantitative PCR and RNA interference. In this chapter I discuss recent progress in the field of chemotaxis in Chlamydomonas. Emphasis is placed on the signal pathways by which the two environmental cues – ammonium and light control chemotaxis and gametic differentiation. Short Communication A - Dynamics of cells proteins and membrane component can be measured by the use of Fluorescence Recovery After Photobleaching (FRAP) method. FRAP

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is a powerful tool that permits the lateral motions of molecules in cell membranes of single, living cells to be examined under physiological conditions. In FRAP experiments fluorescently labelled molecules are irreversibly photo bleached by a high-intensity laser beam that briefly illuminates a small area of the cell. Diffusion of surrounding non-bleached molecules leads to recovery of the fluorescence. A higher mobility of the molecules results in a shorter time of recovery. From the recovery curve, it is possible to obtain estimates of the diffusion coefficient and immobile fraction. Here we reviewed the use of Fluorescence Recovery After Photobleaching in the study of lateral mobility of cell membrane elements. Short Communication B - Although evolution has put strong selective pressure on ensuring that individuals have healthy, functioning gametes, at least 15% of human couples have fertility problems. The infertility can be due to either partner, but fecundation failure in the male partner occurs in one third of the cases. A large number of different causes can lead to this male infertility, but the most striking ones are those that affect the mobility of the sperm because the main function of this cell is to deliver the male nuclear package to the ovocyte. In cases of motility problems, the normal beating of this flagellum may be decreased, abnormal, or totally absent, depending on the flagellar ultrastructural defects. Sperm motility requires the interaction of intracellular factors, including an adequate level of adenosine triphosphate (ATP), functional axonemal dynein ATPases, and an intact axoneme bathing in a proper ionic environment. These factors also represent the minimal conditions required to initiate and maintain the motility of modeled spermatozoa (i.e.; spermatozoa demembranated by a detergent treatment in which motility is initiated by the addition of ATP and ions. Extracellular factors that affect these minimal requirements will cause an arrest in sperm motility. Factors such as sperm agglutinating antibodies that act at the surface of cells by forming a physical network of spermatozoa bound to each other obviously act via different mechanisms. Through their adhesion to spermatozoa, bacteria such as E. coli can trigger membrane damage and a subsequent loss of motility. High concentrations of cytokines such as interferon-a and g, tumor necrosis factors, IL-1 and IL-6 and TNF-a decreased sperm motility and/or penetration rates in zona-free hamster oocytes assays. Interleukin-6, the most specific marker of male accessory gland infection, was associated with lipid peroxidation of human sperm membrane. It also increased capacitation and acrosome reaction. ROS generated in semen originated mainly from PMN and abnormal spermatozoa and the levels of these ROS were inversely correlated to the percentage of motile spermatozoa and fertility in vivo. ROS, of which hydrogen peroxide is responsible for the inhibition of sperm motility, acted by depletion of intracellular ATP and subsequently on the cAMP-dependent phosphorylation of axonemal proteins. The impact of ROS on sperm motility depends of several factors, including the type of ROS involved, the site and the duration of exposure, and the types of ROS scavengers present. The time at which the effects produced by ROS are evaluated is also of paramount importance because ROS often initiate a cascade of events that will trigger a biological response. Antioxidants, as well as antibiotics may improve motility.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 1-50

ISBN: 978-1-60692-570-6 © 2009 Nova Science Publishers, Inc.

Chapter I

Scaffolding Proteins that Regulate the Actin Cytoskeleton in Cell Movement S.J. Annesley and P.R. Fisher Microbiology Department, La Trobe University, Melbourne, Australia

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Abstract Actin is the main component of the microfilament system in all eukaryotic cells and is essential for most intra- and inter-cellular movement including muscle contraction, cell movement, cytokinesis, cytoplasmic organisation and intracellular transport. The polymerisation and depolymerisation of actin filaments in nonmuscle cells is highly regulated and the reorganisation of the actin cytoskeleton can occur within seconds after chemotactic stimulation. There are many proteins which are involved in the regulation of the actin cytoskeleton. These include receptors which receive chemotactic stimuli, G proteins, second messengers, signalling molecules, kinases, phosphatases and transcription factors. These proteins are varied and numerous and are involved in multiple pathways. Despite the large number of proteins, there are not enough to coordinate the various responses of the cytoskeleton. An additional level of regulation is conferred by scaffolding proteins. Due to the presence of numerous protein interaction domains, scaffolding proteins can tether various proteins to a certain location within the cell to facilitate the rapid transfer of signals from one protein to the next. This colocalisation of the components of a particular pathway also helps to prevent unwanted crosstalk with components of other pathways. Tethering receptors, kinases, phosphatases and cytoskeletal components to a particular location within a cell helps ensure efficient relaying and feedback inhibition of signals to enable rapid activation and inactivation of responses. Scaffolding proteins are also thought to stabilise the otherwise weak interactions between particular proteins in a cascade and to catalyse the activation of the pathway components. There are numerous scaffolding proteins involved in the regulation of the cytoskeleton and this chapter has focussed on examples from several groups of scaffolding proteins including the MAPK scaffolds, the AKAPs, scaffolds of the post synaptic density and actin binding scaffolding proteins.

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1. Introduction Actin is the main component of the microfilament system in all eukaryotic cells and is essential for a wide variety of functions including muscle contraction, cell movement, cytokinesis, cytoplasmic organisation to intracellular transport [Van Troys et al., 1999]. Actin is ubiquitous in eukaryotes and is one of the most highly conserved proteins in evolution. For example, actin of humans and the protist Euplotes crassa still share 61 % identity. Actin is even more conserved in evolution than histone H4 [Van Troys et al., 1999]. Actin is present in two forms - the 42 kDa globular monomeric actin [G-actin] which can reversibly assemble into filamentous actin [F-actin] [McGough, 1998]. In resting cells about half of the actin is present in the monomeric state and the other half is present in the filamentous state [Eichinger et al., 1998]. The polymerisation and depolymerisation of actin filaments in nonmuscle cells is highly regulated. The reorganisation of the actin cytoskeleton can occur within seconds after chemotactic stimulation and these changes correspond to morphological changes. The signalling pathways leading to reorganisation of the cytoskeleton generally involves activation of a receptor through binding of a ligand and transmission of this signal through G proteins, signalling molecules such as GTPases, second messengers such as Ca2+ and its associated molecules, kinases, phosphatases and cytoskeletal components. The movement of the cell is concomitant with the formation and breakdown of cell adhesion sites at the leading edge and trailing edge respectively. These cell adhesion sites contain integrins which mediate attachment to extracellular matrix components. Clearly the rearrangement of the cytoskeleton is complex and involves numerous proteins. The signalling pathways leading to movement of the actin cytoskeleton involve many proteins, yet the number of specific cellular behaviours that are elicited by and are reliant on these pathways is even greater. Additionally a single stimulus can elicit more than one response. Hence there are not enough proteins for each to have an individual biological role, which implies that the signalling proteins must work in conjunction with each other [Pawson & Nash, 2000]. So how do the pathways maintain specificity and prevent cross talk between alternate pathways? One way that cells do this is via scaffolding proteins. Scaffolding proteins have many postulated roles in signalling pathways. They are thought to tether various proteins to a certain location within the cell to facilitate the rapid transfer of signals from one protein to the next [Smith & Scott, 2001]. The colocalisation of the components of a particular pathway also helps to prevent unwanted crosstalk with components of other pathways [Whitmarsh & Davis, 1998]. Tethering receptors, kinases, phosphatases and cytoskeletal components to a particular location within a cell helps ensure efficient relaying and feedback inhibition of signals to enable rapid activation and inactivation of responses. Scaffolding proteins are also thought to stabilise the otherwise weak interactions between particular proteins in a cascade and to catalyse the activation of the pathway components [Burack & Shaw, 2000].

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Table 1. Protein Interaction Domains Domain

Size

Structure

Recognition Site

Examples

References

Short peptide motifs containing PTyr residues. Group I prefer PTyrhydrophilic-hydrophilic-hydrophobic and Group II prefer PTyr-hydrophobichydrophobic recognition sequence. Residues C-terminal to PTyr determine substrate specificity. Binding to peptide with optimal sequence is relatively high. Bind to proline-rich target sequences containing the PXXP motif. The 1st X tends to be a proline and the 2nd X tends to be an aliphatic residue. The target peptides generally adopt a polyproline type II helix which can bind the domain in either orientation. Specificity is determined by residues N- or C-terminal to the PXXP motif. Most bind phosphoinositides or inositol phosphates and they do this with varying affinities. PH domains with low affinities may localise to membranes through the presence of multiple PH domains on a single protein.

Found in proteins involved in protein tyrosine kinase pathways such as phospholipid metabolism, tyrosine phosphorylation and dephosphorylation, gene expression, protein trafficking and cytoskeletal architecture. Found in proteins involved in signal transduction, cytoskeletal organisation and membrane traffic.

Kuriyan & Cowburn, 1997 Pawson, 1995 Cohen et al., 1995 Machida & Mayer, 2005

(amino acids)

SH2 (Src homology 2)

~100

A spine consisting of an antiparallel β sheet dividing the domain into two distinct sides. One side binds PTyr [phosphotyrosine] the other contains residues which interact with residues Cterminal to the PTyr on the target. A conserved pocket binds the PTyr.

SH3 (Src homology 3)

50-70

Consists of 2 antiparallel β sheets each comprised of 3 β strands which fold into a β sandwich. One side contains a hydrophobic pocket which binds the recognition motif.

PH (Pleckstrin homology)

100120

PH fold contains 2 antiparallel β sheets consisting of 3 and 4 strands respectively. The structure is capped by an amphiphatic C-terminal α helix. On one side of the PH domain is a narrow cleft containing 3 variable loops and clusters of positive charges, this is the site for binding ligand.

Found in a wide range of signalling and cytoskeletal proteins. Proteins are generally cytoplasmic but associate with the plasma membrane & are often involved in the intracellular localisation of proteins.

Morais et al., 1996 Bar-Sagi et al., 1993 Pawson, 1995 Kuriyan & Cowburn, 1997 Cesareni et al., 2002

Ludbrook et al., 1997 Chung et al., 2001 Funamoto et al., 2001 Maffucci & Falasca, 2001 Lemmon et al., 2002

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Table 1. (Continued) Domain

Size

Structure

Recognition Site

Examples

References

Proteins containing PDZ domains are ubiquitously expressed and have diverse roles eg. Assembly & regulation of neuronal synapses and establishment & maintenance of epithelial cell polarity & are found in a range of proteins including protein kinases, tyrosine phosphatases and a variety of proteins involved in ion-channel and receptor clustering. Cell adhesion molecules, membrane bound receptors such as tyrosine receptors, transcription factors and cytoskeletal proteins.

Morais et al., 1996 Pawson & Scott, 1997 Kuriyan & Cowburn, 1997 Huber,2001 Hung & Sheng, 2002 Zimmermann, 2006

(amino acids)

PDZ (PSD95,discs large, zona occludens 1)

80-90

Comprise 6 β strands (βAβF) and 2 α helices (αA & αB) which fold into a 6stranded β sandwich. Target proteins bind as an antiparallel β strand in a cavity formed by the βB strand, the αB helix and a loop connecting βA & βB strands. PDZ interactions are reversible and versatile and the binding affinities are moderate.

Recognise motifs of 3-7 residues or PDZ binding motifs (e.g. the E[S/T]DV motif of certain ion channel subunits) generally at the C-terminus of target proteins. PDZ domains can form homotypic or heterotypic interactions with other PDZ domains. Proteins with PDZ domains often contain other protein interaction domains. Specificity appears to be determined by the interaction of the first residue of the αB helix and the side chain of the -2 residue of the C-terminal ligand.

Immunoglobulin (Ig)-like

~100

Each domain contains 2 β sheets which are comprised of 3-5 β strands. The 2 β sheets are packed tightly together to form a β sandwich which is further stabilised by a conserved disulfide bond, however this bond is absent in some examples.

Most Ig-domains are able to interact with other Ig-like domains. The fold also facilitates interactions with other proteins and DNA molecules. The domains interact with other proteins mainly through the faces of the β sheets & therefore can interact with other proteins through any accessible part of their surface.

Williams & Barclay, 1988 Wisemann et al., 2000 Nagata et al., 1999 Noegel et al., 1989

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Table 1. (Continued) Domain

Size

Structure

Recognition Site

Examples

References

Contains a core fold which consists of 2 perpendicular antiparallel β sheets (β1-4 & β5-7) packed to form a β sandwich capped by a C-terminal helix (α3). Ligand binds in the groove between β5 and α3, phospholipid binds in a highly basic area composed of residues from loop regions near the N-terminus.

Recognise peptide sequences containing variations of the NPXY motif which is typically located in the juxtamembrane regions of integral membrane proteins. Also bind to the head group of phospholipids.

Occurs in proteins involved in signal transduction, membrane trafficking and cytoskeletal dynamics, such as neuronal development, immune response, tissue homeostasis and cell growth.

DiNitto & Lambright, 2006 Uhlik et al., 2005

(amino acids)

PTB (Phosphotyrosine binding)

EH (Eps15 homology)

~100

Each domain contains 2 calciumbinding helix-loop-helix motifs (EF hands, however not all bind calcium), which are linked by an antiparallel β sheet. Most EH domains are at the Nterminus of the protein.

Recognises peptides containing a NPF motif. The NPF motif enters a conserved hydrophobic pocket which binds the peptide, bringing the asparagine residue of the peptide in close proximity to the highly conserved tryptophan residue in the EH domain.

Found in proteins which regulate endocytic membrane transport events, the regulation of actin dynamics, and various roles in signal transduction.

Naslavsky & Caplan, 2005

WW (refers to 2 tryptophan residues [W] present in these domains)

40

Fold as a stable antiparallel 3 stranded β sheet structure which forms a shallow binding pocket for ligands.

Recognise peptides containing a proline-rich core motif (generally PPXY or PPLP) which is usually flanked by additional prolines.

Found in proteins involved in numerous cellular processes including RNA processing & transcription, receptor signalling and protein trafficking.

Kay et al., 2000 Sudol et al., 2005

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Table 1. (Continued) Domain

Size

Structure

Recognition Site

Examples

References

(amino acids)

LIM (lin-11, isl-1 & mec-3)

50-60

Double zinc finger motifs with the consensus sequence (CX2CX1623HX2C(X2)CX2CX16-21CX2H/D/C). The conserved cysteine, histidine & aspartic acid residues form 2 tetrahedral pockets which bind zinc & stabilise the secondary & tertiary structure.

No common LIM binding motif has been identified however, LIM domains can associate with each other, with motifs found on transcription factors and other protein interaction domains.

Proteins containing LIM domains are involved in various cellular processes including cytoskeletal organisation, cell lineage specification and organ development.

Bach, 2000 Brown et al., 2001 Khurana et al., 2002

EVH1 (enabled/vasodilator -stimulated phosphoprotein homology 1)

115

The overall fold is highly conserved and consists of a compact β sandwich closed along one edge by a long α helix. There is a cluster of three aromatic sidechains which form the recognition site for the proline rich targets.

Bind to proteins containing proline rich sequences. Two classes bind different motifs: Class I recognise FPxфP motif [where ф is a hydrophobic residue] and Class II recognise PPxxF motif. The binding is of low affinity but high specificity. Specificity is achieved through residues flanking the core motif.

Exist primarily in proteins which are involved in the regulation of the actin cytoskeleton and in postsynaptic proteins.

Ball et al., 2002

IQ

~20

General sequence is [I,L,V]QXXXRXXXX[R,K] which forms a basic amphipathic helix. Critical hydrophobic residues reside at positions 1, 8 and 14. Basic amino acids often flank the core motif.

Bind calmodulin.

Present in wide range of proteins including myosins, phosphatases, Ras exchange proteins, neuronal growth proteins and voltage operated channels.

Bahler & Rhoads, 2002

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Table 1. (Continued) Domain

Size

Structure

Recognition Site

Examples

References

The V region binds to G actin and the CA region binds to the Arp2/3 complex promoting actin nucleation.

Present in WASP family members which are involved in the regulation of actin polymerisation. Exist in a wide number of proteins such as signalling proteins and transcription factors and also to RNA. Proteins are present in all subcellular locations, and are involved in diverse biological functions from chromatin remodelling to apoptosis.

Cory et al., 2003 Kato et al., 2002 Yamaguchi et al., 2000

(amino acids)

VCA (Verprolin, cofilin homologyacidic) or VPH (Verprolin homology) SAM (Sterile Alpha Motif)

~70

V region contains the motif KLKK which is essential for actin binding. Also contains two conserved serines which are phosphorylated, enhancing interaction with Arp2/3 complex. The core consists of a five helix bundle comprised of hydrophobic residues. Helices 3,4 and 5 are essential for the high affinity interactions.

Can self associate to form homoor hetero-SAM domain interactions and also form heterotypic interactions with nonSAM containing proteins.

Kim & Bowie, 2003 Kwan et al., 2006

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Most scaffolding proteins are large in size presumably due to their ability to simultaneously bind various proteins. However some smaller scaffolding proteins such as MP1 (MEK partner 1) which is only 14.5 kDa in size have also been identified and MP1 has been shown to bind at least two members (MEK1 and ERK1) of a signalling cascade [Vomastek et al., 2004]. They also often contain many protein-protein interaction domains such as SH2, SH3, PH, and PDZ domains amongst many others, which aid in tethering different proteins involved in a particular cascade. The protein interaction domains have been conserved throughout evolution due to their biological significance and are represented in nearly all eukaryotes [Liu et al., 2006]. In some cases the prevalence of a domain increases with multicellularity. For example the PDZ domain is found in only two proteins in yeast whereas 545 PDZ domains have been identified in humans [Zimmermann, 2006]. Additionally some domains such as the SH2 domain are not represented in fungi, mainly due to the lack of tyrosine phosphorylation pathways [Liu et al., 2006]. Many protein interaction domains have been identified and some of these are listed in Table 1. Many of the scaffolding proteins identified to date, especially the MAPK family members, dimerise to form homo- and hetero-oligomers and this dimerisation appears to be a conserved feature [Elion, 2001]. The dimerisation may allow phosphorylation in trans between protein kinases attached to different subunits of the dimer [Hunter, 2000]. As exemplified by overexpression experiments, the stoichiometry of the scaffolding protein to other components in the pathway is tightly regulated to ensure sufficient concentrations of all components of the pathway are present. Too much scaffold can titrate out and separate the binding partners, and consequently inhibit signalling [Symons et al., 2006]. An example of this is the scaffolding protein Ksr which was identified as a positive and negative regulator of signalling. When Ksr was overexpressed it was shown to suppress signalling, however at normal levels it acts as a positive regulator, increasing signalling [Levchenko et al., 2000]. There are several major families of scaffolding proteins involved in the regulation of the cytoskeleton. These include amongst others the MAPK (Mitogen Activated Protein Kinase) scaffolding proteins, the AKAPs (A-Kinase Anchoring Proteins), the post synaptic density (PSD) scaffolding proteins and actin binding scaffolding proteins including IQGAP and filamin. This article will focus on these groups.

1.1. MAPK Scaffolding Family The mitogen activated protein kinases (MAPKs) are present in all eukaryotic organisms and are expressed in virtually all mammalian tissues [Sabbagh et al., 2001]. MAPKs respond to a variety of extracellular stimuli such as growth factors, mitogens, cytokines and environmental stress [Whitmarsh & Davis, 1998] and they relay these messages mainly to transcription factors to alter gene expression. The signalling pathways these proteins are involved in are diverse, including those controlling proliferation, differentiation, apoptosis and cell movement [Pullikuth et al., 2005]. MAPK cascade scaffolds are widely divergent, indicating that they have evolved independently for the individual requirements of subsets of kinases and upstream activating events [Elion, 2001].

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A MAPK cascade generally consists of three protein kinases: a MAPK, a MAPK activator (MEK or MKK) and a MKK activator (MEKK or MKKK) and the particular cascade is named after the last kinase of the series [Schaeffer & Weber, 1999]. Some MAPK cascades also contain an additional kinase, a MAPKKKK which phosphorylates the MKK activator. The first kinase in the series, MKKK, is activated in response to various stimuli binding to a receptor. Ligand binding to the receptor results in the recruitment of adaptor proteins specific to the receptor type and/or activation of small GTP-binding proteins, both of which can activate MKKK proteins. For example tyrosine kinase receptors often result in recruitment of Rho GTPases which can then activate a MKKK [Davis, 2000]. Activation of MKKK leads to MKK serine/threonine phosphorylation which in turn activates the MAPK by dual phosphorylation on threonine and tyrosine residues in the activation loop [Ito et al., 1999]. Phosphorylation of both residues on the MAPK is essential for it to perform its various functions as shown by mutational analysis proving that phosphorylation of one of the residues does not activate the kinase whereas dual phosphorylation at these sites results in a >1000 fold increase in the activity of the MAPK [Widmann et al., 1999]. The MAPK’s are proline-directed serine/threonine kinases. The phosphorylation of the MAPK by the upstream kinase results in a conformational change that creates a surface pocket which is specific for a proline residue. The MAPK generally recognizes a proline at the +1 position of a potential substrate [Schaeffer & Weber, 1999]. Potential substrates of MAPKs are more often than not transcription factors, however, other targets include cytoskeletal proteins, protein kinases and phosphatases [Symons et al., 2006]. In mammals MAPK cascades have been shown to be involved in diverse pathways regulating cell fate including differentiation and proliferation, response to extracellular stress and apoptosis. Many of the MAPK substrates are transcription factors, however other substrates such as cytoskeletal proteins, protein kinases and phosphatases have also been identified [Ho et al., 2006; Widmann et al., 1999]. Three major mammalian MAPK cascades have been identified, the ERK (extracellular signal-related kinase), the JNK/SAPK (c-Jun Nterminal kinase/stress-activated protein kinase])and p38 [Fusello et al., 2006] cascades. 1.1.1. The ERK Cascade Eight ERK proteins have been identified in mammals, numbered 1 through 8, and the cascades involving these proteins are required for regulating processes as diverse as cell growth and differentiation as well as cytoskeleton-dependent focal adhesion and cell spreading. ERK1 and 2 were the first ERKs to be identified. They share 90 % sequence identity and participate with the same members of a MAPK cascade [Bogoyevitch & Court, 2004]. Both proteins have been extensively studied and hence will be the focus of this section. The members of this pathway include Raf-1 (a MKKK), MEK1/2 (a MKK) and ERK1/2 (a MAPK), and many other associated regulatory proteins. Raf-1 acts as a scaffolding molecule by binding several components of the pathway and localizing them to the plasma membrane upon stimulation. The ERK1/2 cascade is initiated through the binding of growth factors such as EGF (epidermal growth factor) and PDGF (platelet-derived growth factor) to numerous receptors. These receptors include tyrosine kinase receptors, cytokine receptors, G-protein coupled receptors and integrins [Widmann et al., 1999].

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As seen in Figure 1, ligand binding to the various receptors ultimately results in tyrosine phosphorylation of the receptor and the recruitment of various adaptor proteins to the receptor where they facilitate the activation of Ras through the exchange of GDP for GTP at its active site [Pouyssegur et al., 2002]. Raf-1 binds a group of proteins called the 14-3-3 proteins. These proteins bind to the kinase domain of Raf-1 and stabilise the inactive configuration of Raf-1 in which the regulatory domain masks the catalytic domain. Ras has a high affinity for Raf-1 and once activated it is able to bind to the NH2-terminal regulatory domain of Raf-1 and displace the 14-3-3 protein. The displacement of the 14-3-3 protein alters the structure of Raf-1 so the catalytic domain is accessible to deposphosphorylation by PP2A. Only Raf-1 which is dephosphorylated at these residues is able to initiate the MAPK cascade [Dhillon & Kolch, 2002]. The binding affinity of Ras for Raf-1 is very high, higher even than the affinity of Ras for Ras-GAP. Termination of Raf-1 activation occurs through phosphorylation of Raf-1 at another serine residue by the cAMP-dependent kinase, PKA thereby diminishing the affinity of Ras for Raf-1 [Dhillon & Kolch, 2002]. Ras activated Raf-1 binds its downstream target kinase, MEK1 through a proline-rich sequence and phosphorylates MEK1 at a serine/threonine residue. MEK1 in turn binds ERK1/2 and activates it through dual phosphorylation on a threonine and tyrosine residue in the Thr-Glu-Tyr motif. This phosphorylation causes a conformational change in the ERK2 activation lip creating a proline-specific pocket [Schaeffer & Weber, 1999]. Activated ERK1/2 can then dissociate from the scaffold and translocate to specific locations within the cell such as the nucleus where it can bind to transcription factors or to the cytoplasm where it binds cytoskeletal proteins. ERK1/2 phosphorylates a serine and threonine residue on its substrates in a proline directed manner. The most stringent recognition sequence for the ERK1/2 proteins is Pro-Leu-Ser/Thr-Pro [Widmann et al., 1999]. The ERK1/2 cascades are activated by numerous growth factors binding to many different receptors and have various substrates which they bind. They are also present in nearly all types of mammalian cells. To accommodate all these varied responses, additional levels of regulation exist to ensure flexibility yet specificity to each cascade and also insulation from regulation by other stimuli [Pullikuth et al., 2005]. In addition to the scaffolding protein Raf-1, other scaffolding proteins seem to be employed in the ERK1/2 cascades. An example of an additional scaffolding protein is Ksr (Kinase suppressor of Ras). Ksr appears to be evolutionarily conserved with homologs identified in mammals, Drosophila, C. elegans and Xenopus [Denouel-Galy et al., 1997]. Ksr shows high structural homology to Raf-1, but despite a putative kinase domain in the Cterminus it is devoid of any kinase activity [Roy & Therrien, 2002]. Figure 1 provides an illustration of this MAPK cascade and how many of the components interact due to the two scaffolding components Raf-1 and KSR. Like Raf-1, Ksr is bound by a 14-3-3 protein which may serve to sequester Ksr to the cytoplasm. In response to stimuli such as ceramide, activated Ras induces the translocation of Ksr to the plasma membrane where it binds to the γ subunits of heterotrimeric G-proteins. Ksr has also been shown to constitutively associate with 14-3-3, MEK and a Rho family member regardless of Ras activation, whereas its interaction with Raf-1 and MAPK is Ras dependent. It appears then that Ras-dependent localisation of Ksr may bring MEK in close

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proximity to Raf-1 and a MAPK and aids in assembling of the cascade [Roy & Therrien, 2002].

[Morrison, 2001]

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Figure 1. Scaffolding proteins Raf-1 and Ksr in an ERK (MAPK) cascade. The two scaffolding proteins Raf-1 and Ksr both bind a protein, 14-3-3 which holds these proteins in the cytoplasm. Upon activation of Ras, Raf-1 is recruited to the membrane where it binds Ras with a very high affinity. The binding of Ras displaces the 14-3-3 protein, changing the structure of Raf-1 to an active configuration. The activation of Ras also recruits Ksr to the membrane where it binds to the β subunit of a heterotrimeric G protein (Gβ). Ksr brings MEK and ERK (MAPK) in close contact with the other kinase members and initiates the phosphorylation cascade from Raf-1 to MEK and finally to ERK [Morrison, 2001]. (Reproduced with permission of the Company of Biologists).

Another additional scaffolding protein of an ERK1/2 cascade is MP1 (MEK partner 1). MP1 has been shown to interact in ERK cascades in response to adhesion factors but not in response to other factors such as PDGF. MP1 tightly associates with p14, a protein that acts as an adaptor linking MP1 to late endosomes [Teis et al., 2002]. This association is very stable and the MP1/p14 complex is able to directly bind two members of the ERK1/2 cascade, these being ERK1 and MEK1 but not their closely related isoforms ERK2 and MEK2 respectively. This selectivity suggests that there may also be different but as yet unidentified components which would selectively regulate ERK2 and MEK2 [Pullikuth et al., 2005]. MP1/p14 associates with MEK1 through a proline-rich sequence which contains a site for PAK (p21-activated kinase) phosphorylation. MP1/p14 is able to bind activated PAK (p21-activate kinase) which can then phosphorylate and activate MEK1. PAK is activated in response to cell adhesion factors and by phosphorylating MEK1 it can couple the MEK1ERK1 activation to upstream adhesion signals [Pullikuth et al., 2005]. MP1/p14 is also able to interact with Rho (a GTPase) and ROCK (a Rho kinase). ROCK is able to phosphorylate various kinases and a cytoskeletal protein, cofilin which can then counteract the membrane protrusion stimulated by the Rho proteins. This inhibition is known to be important for focal adhesion turnover and cell spreading [Pullikuth et al., 2005]. Once activated ERK1 is released from the complex with MEK1 and MP1 allowing the movement of ERK1 molecules to their sites of action [Sharma et al., 2005]. Taken together this particular ERK cascade provides an example of how different stimuli (adhesion factors), can recruit members of the ERK cascade through adaptor proteins (p14) and scaffolding proteins (MP1) to specific

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compartments of the cell (late endosomes) in order to elicit a particular behaviour (cell adhesion and spreading). Rho

ROCK

Cell Adhesion Factors PAK

P MEK1

MP1 p14

Late Endosomes

ERK1

P ERK1

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Figure 2. The MP1 Scaffold. MP1 is able to interact directly with two members of the MAPK cascade, MEK1 and ERK1. MP1 associates very strongly with the adaptor protein p14. This association links MP1 and the members of the complex to the late endosomes. This MAPK cascade is initiated by cell adhesion factors which lead to the activation and phosphorylation of PAK1. Once activated, PAK1 can phosphorylate MEK1 and MEK1 in turn phosphorylates ERK1 (as denoted by the broken arrows). Activated ERK1 is released from the complex and translocates to its sites of action. MP1 is also able to interact directly with Rho and ROCK. Rho is a GTPase and ROCK is a Rho kinase which, in addition to phosphorylating and activating Rho, can phosphorylate various kinases and the cytoskeletal protein cofilin.

The strength and duration of signals transmitted through ERK1/2 can also affect the type of response. For example in PC-12 cells stimulation of the ERK1/2 cascade through EGF stimulates proliferation yet stimulation by NGF stimulates differentiation of cells. The different responses are thought to be linked to the duration of the ERK1/2 activation. EGF stimulates ERK1/2 transiently with activity levels returning to basal standards within 1-2 hours. Conversely NGF stimulates ERK1/2 in a more sustained manner [Widmann et al., 1999]. It seems that there are many factors which aid in the regulation of MAPK cascades, including adaptor and scaffolding proteins and duration of expression. 1.1.2. JNK/SAPK Cascades This family of MAPKs are activated by a variety of genotoxic and cytotoxic cellular stresses including heat shock, UV irradiation, hyperosmolarity, ischemia and cytokines such as interleukin-1 [Nihalani et al., 2001; Ho et al., 2006]. To date 14 associated MKKKs (including MEKK1-4, ASK1, TAK1, MST, SPRK, MLK1-3, DLK, MUK and TPL2), 2 MKKs (MKK4 and 7) and three JNK kinases (JNK1-3) and their 10 isoforms resulting from alternative splicing have been identified in mammals [Ho et al., 2006; Nihalani et al., 2001; Widmann et al., 1999]. JNK3 is expressed predominantly in the brain and testis whilst JNK1 and 2 are ubiquitously expressed [Ito et al., 1999]. The JNKs, like other MAPKs are activated

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by dual phosphorylation on threonine and tyrosine residues of the Thr-Pro-Tyr motif and they phosphorylate substrates at a Ser/Thr-X-Pro motif [Widmann et al., 1999]. The numerous members of these cascades reflect the number of distinct signalling pathways that these cascades are involved in [Ito et al., 1999]. Distinct members of JNK cascades have been implicated in various responses including apoptosis, proliferation, and embryonic morphogenesis [Nihalani et al., 2001]. There are a number of signalling proteins which are able to activate the JNK cascade and this activation can operate through a variety of cell surface receptors including the TNF receptor family, GPCR, tyrosine kinase receptors and cytokine receptors [Widmann et al., 1999]. The type of receptor that is stimulated by a particular ligand determines which adaptor proteins are recruited. For example, GPCRs are linked to the adaptor proteins β-arrestins which control the desensitization and internalization of the receptors. They have also been shown to act as MAPK scaffolds themselves binding many members of the cascade and recruiting the signalling complexes to activated GPCRs. An example involves the AT1aR (angiotensin type 1a receptor) GPCR. β-arrestin 2 binds all three members of the MAPK cascade, JNK3, MKK4 (indirect interaction) and ASK. Upon activation of the receptor, βarrestin 2 is recruited to the receptor along with its associated members of the MAPK cascade and aids in the activation of JNK3 [Willoughby & Collins, 2005]. Tyrosine kinase receptors activate members of the Rho family of GTPases which in turn phosphorylate and activate MLK and MEKK family members. Cytokine receptors and the IL1 receptor appear to mediate JNK activation through a group of adaptor proteins of the TRAF group which have been shown to phosphorylate MEKK1 and ASK1. Alternatively the adaptor protein Nck may mediate JNK activation by Eph receptors [Davis, 2000]. Most JNK substrates identified are transcription factors including c-Jun, Elk-1, p53 and NFAT4. Phosphorylation of these substrates can lead to increased or decreased rates of transcriptional activity, it can also lead to structural changes which may protect the transcription factor from subsequent modifications, thereby stabilizing them [Widmann et al., 1999]. However, there are some non-nuclear substrates such as the Bcl-2 family member BAD and the 14-3-3 protein [Willoughby & Collins, 2005]. The diversity and localisation of these cascades is often aided by scaffolding proteins. Several JNK scaffolding proteins have been identified, these include JIP (JNKinteracting protein, JSAP1 (JNK stress activated protein 1) and JLP (JNK-associated leucine zipper protein) [Ito et al., 1999; Nihalani et al., 2001; Lee et al., 2002a]. These scaffolding proteins appear to bind particular members of a JNK cascade in response to specific stress stimuli.

1.1.2.1. The JIP Scaffolds Four JIP genes and several splice isoforms have been identified in mammals. Two of the encoded proteins JIP1 and JIP2 are closely related, whereas JIP3 is structurally unrelated [Davis, 2000]. Unlike the other JIP proteins, JIP4 is involved in activating the p38 MAPK and not the JNK MAPK [Kelkar et al., 2005]. The JIP1 and 2 proteins contain an SH3 domain and a PTB domain within their carboxyl-terminal domain. The PTB domain has been shown to interact with several receptors and members of the Rho family. The function of the SH3 domain is unclear, however, this domain has been shown in other proteins to function in

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protein-protein interactions [Davis, 2000]. All JIP proteins interact specifically with members of the MLK (mixed lineage kinase) family of MKKKs but not with other MKKKs such as the MEKK proteins. They also interact with MKK7 but not MKK4 [Nihalani et al., 2001] and JNK1 and 2. As seen in Figure 3 JIP binds to a MLK protein in its monomeric inactive state through its leucine zipper domain as well as binding a receptor or adaptor protein. Upon appropriate stimulation mediated through the adaptor protein the MLK protein dissociates allowing the two leucine zipper domains to interact and form a MLK dimer. Dimerisation initiates autophosphorylation and subsequent catalytic activation. The interaction of the two leucine zipper domains in the MLK protein to form stable dimers is of high affinity and is the most favourable configuration. The JIP protein therefore provides a mechanism of regulation for this JNK cascade by tethering MLK in an inactive state prior to stimulation. The dissociation and activation of the MLK protein results in recruitment of JNK to the JIP scaffold by an unknown mechanism and also results in phosphorylation of JIP1-bound MKK7. MKK7 can then phosphorylate the bound JNK protein [Nihalani et al., 2001]. By binding and aggregating all three components of the cascade JIP may therefore act as a scaffolding protein.

MLK

MLK

M K K 7

M K K 7

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MLK

JIP

JIP

JNK Nucleus

Figure 3. The JIP scaffold. JIP associates with inactivated MLK and MKK7. An adaptor protein (not shown on this diagram) facilitates the activation of MLK which stimulates the protein to dimerise, autophosphorylate and dissociate from the complex. This dissociation results in the phosphorylation of MKK7 and the recruitment of JNK to the complex which is phosphorylated by MKK7. Phosphorylated JNK can then dissociate from the complex and translocate to its substrate targets such as transcription factors in the nucleus.

Like other scaffolding proteins JIP proteins can form homo- and hetero-oligomers and belong to the group of phosphoproteins [Nihalani et al., 2001]. JIP1, 2 and 3 have been shown via a yeast two hybrid screen to interact with the tetratricopeptide repeat (TPR) domain of the light chain of the microtubule motor protein kinesin [Kelkar et al., 2005]. JIP1 and JIP2 interact with kinesin through their COOH terminus whereas JIP3 interacts via a leucine zipper domain. In immunoprecipitation experiments, kinesin, JIP1, a MKKK protein and a receptor, Reelin, coprecipitated suggesting that kinesin, a microtubule motor protein, acts to localise signalling complexes through a scaffolding protein [Verhey et al., 2001].

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1.1.2.2. The JSAP Scaffold JSAP is another scaffolding protein of the JNK cascade and is expressed as a result of alternative splicing of the JIP3 gene [Nihalani et al., 2001]. JSAP preferentially binds all three JNKs (with a higher preference for JNK3 [Ito et al., 1999]), the MKKs MKK/SEK1, 4 and 7, and the MKKK MEKK1 [Takino et al., 2005]. All three interactions occur at unique sites on JSAP1. Figure 4 is a cartoon depicting the JNK cascade which involves JSAP as a scaffolding protein. In this figure it can be seen that MEKK1 interacts with inactive SEK1 and JSAP1, however inactive SEK1 does not interact directly with JSAP1 until it is activated. It appears that when stimulated MEKK1 phosphorylates SEK1 and facilitates its binding to the scaffold protein JSAP1. SEK1 can then activate JNK through dual phosphorylation. Once activated JNK phosphorylates JSAP1 and is subsequently dissociated from the scaffold. Phosphorylated JSAP1 has little binding affinity for JNK. Once dissociated, JNK is free to translocate to the nucleus and to phosphorylate its transcription factor substrates. JSAP1 contains a leucine zipper motif which may mediate its homo- or hetero-oligomerisation [Ito et al., 1999]. It is unknown whether this dimerisation is required for its function, however it is known to occur in many other scaffolding proteins [Elion, 2001].

Figure 4. JSAP Scaffold. Prior to initiation of the cascade JSAP interacts directly with two members of the MAPK cascade, MEKK1 and JNK. Once MEKK1 is stimulated it phosphorylates SEK1 which, in this activated state can bind to the scaffold, JSAP. SEK1 is then able to phosphorylate JNK which subsequently phosphorylates JSAP and results in the dissociation of JNK from the scaffold. Dissociated JNK is then free to translocate to the nucleus and phosphorylate its various transcription factor substrates.

1.1.2.3. The JLP Scaffold The third JNK scaffolding protein is JLP. JLP shows high sequence homology (69 % identity) to the other two scaffolding proteins discussed (JSAP1 and JIP). JLP is a 180 kDa

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protein that is ubiquitously expressed and is located primarily in the cytoplasm. However, JLP is translocated to the nucleus in response to stress signals [Lee et al., 2002a]. JLP associates with the MKKK MEKK3, the MKK MKK4, as well as JNK1 and a member of a different MAPK cascade p38α. MKK4 has previously been shown to activate MAPK members of both the JNK and p38 cascades. Unlike other scaffolding proteins JLP is able to directly bind target transcription factors such as Max and c-Myc which are substrates for both JNK and p38 members [Lee et al., 2002a]. JLP has also been shown to directly interact with the Gα12 or Gα13 subunit of the heterotrimeric G protein G12 or G13 respectively. G proteins are known to activate JNK cascades however JLP is the first mammalian scaffold identified that links Gα12 or Gα13 to the JNK signalling module [Kashef et al., 2005].

G12 β α γ

MEKK3

MEKK3

JLP MKK4

MKK4

JLP

JNK

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p38

Max

c-Myc

c-Myc

Max

Figure 5. The JLP Scaffold. JLP is able to directly bind all three members of the MAPK cascade - the MKKK, MEKK3; the MKK, MKK4; and two different MAPKs, JNK and p38. JLP is a dimer and as such is able to bind both MAPKs simultaneously within the cell. This MAPK cascade is initiated by stress signals which activate G proteins. JLP directly binds the alpha subunit of the heterotrimeric G proteins G12 or G13 (G12 depicted in this figure). Activation of G proteins stimulates the phosphorylation cascade leading to phosphorylation-mediated activation of JNK and p38, and translocation of JLP to the nucleus where it can directly bind to transcription factors such as Max and c-Myc. JNK or p38 then phosphorylate its transcription factor targets.

JLP contains two leucine zipper domains and three putative SH2 and SH3 binding sites. The leucine zipper domains have been shown to be the regions which bind the transcription factors and recently the second leucine zipper domain has been shown to bind kinesin light chain 1 (KLC1) [Nguyen et al., 2005]. The other JNK scaffold mentioned previously, the JIP family members and JSAP also interact with kinesin, albeit through a different domain. Kinesin’s are a family of motor proteins that move cargo along microtubules in an ATP dependent manner. JLP therefore serves as a link between the kinesin motor proteins and the JNK signalling complex proteins [Nguyen et al., 2005].

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The SH2 and SH3 binding sites in JLP could potentially facilitate interactions with SH2 and SH3 containing kinases which could link to other signalling pathways. JLP binds JNK1 and p38α at different sites and can therefore tether both molecules simultaneously. JLP can form oligomers and this may further facilitate the tethering of JNK and p38. Thus JLP acts as a scaffolding protein by bringing all three members of a MAPK cascade together as well as downstream targets of both the JNK and p38 MAPKs. Binding of the transcription factors to its upstream activators may accelerate transmission of signals. Interactions of JLP with other signalling molecules, possibly through the SH2 and SH3 binding sites could also facilitate subcellular trafficking of a signalling complex to different subcellular sites [Lee et al., 2002a]. 1.1.3. p38 MAPK Cascade The mammalian p38 cascade is homologous to the yeast osmosensing Hog1 pathway. These MAPK cascades are activated like JNK cascades by cellular stress such as UV irradiation, heat shock, protein synthesis inhibitors and certain cytokines. The members of these pathways include the MAPK members, p38α, β, γ and δ, the MKK members MKK4, 3 and 6 and the MKKK members which include TAK1, ASK1, MLK3 and SPRK [Widmann et al., 1999; Ito et al., 1999]. p38α and β share more than 70 % sequence identity and are ubiquitously expressed, whereas p38γ and δ share 60 % sequence identity to p38α and are only expressed in specific tissues such as the brain [Katsoulidis et al., 2005; Dean et al., 2004]. The p38 kinases are activated by dual phosphorylation at a Thr-Gly-Tyr motif. Several potential scaffolding proteins have been identified which interact with specific members of the p38 cascade and are expressed in select tissues. These include the JIP family members JIP2, JLP and JIP4, and OSM (osmosensing scaffold for MEKK3) [Robidoux et al., 2005; Uhlik et al., 2003]. As the p38 kinases like the JNK kinase cascades are activated by cellular stresses it is not surprising that many of the proteins which act as scaffolds for JNK members also act as scaffolds for p38 members. This is the case for many of the JIP family members such as JIP2 and the JLP protein whose role as a scaffold for p38 was described in the preceding section. Another JIP family member, JIP4 has been identified by Kelkar et al. [2005]. It is functionally distinct from other JIP members as it does not activate JNK but has been shown to activate p38 MAP kinase. The jip4 gene encodes at least three proteins JIP4, the previously described JLP and SPAG9 (sperm-associated antigen 9 protein) of unknown function. These proteins all result from alternative splicing [Kelkar et al., 2005]. JIP4 is structurally most similar to JIP3. Like JIP3, JIP4 contains a putative transmembrane domain which indicated that this protein may span the cytoplasmic membrane. However, JIP4 was found to be widely distributed throughout the cytoplasm with no localisation to any organelles. Like other JIP members JIP4 also binds the motor protein kinesin and does this through a leucine zipper domain. In studies conducted by Kelkar et al. [2005] JIP4 was shown to interact with the MAP3K, ASK1, and the MAPK family members, JNK and p38 as illustrated in Figure 6. Even though JIP4 can bind JNK and p38 it is only able to activate p38. Why JIP4 binds yet does not activate JNK is unclear and the authors suggested that JNK may act as a regulator of JIP4 by competing with p38 for binding. No MKK member was shown to interact directly

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with JIP4, so this interaction may occur indirectly via the ASK1 or p38 proteins as has been shown in other MAPK cascades such as the cascade involving β2-arrestin as a scaffold. In this cascade β2-arrestin interacts with MKK4 indirectly via interactions with the MKKK, ASK1, and the MAPK JNK3 [McDonald et al., 2000]. Kelkar et al. [2005] did however demonstrate that in MKK3 and MKK6 double knockout mice, JIP4 is unable to activate p38. Hence these two MKK members may be involved in the JIP4 cascade.

ASK1

JIP4

MKK3/6 p38

Figure 6. The JIP4 scaffold. JIP4 is able to interact with all three members of a p38 signalling module. JIP4 interacts directly with the MKKK ASK1 and the MAPK p38 and indirectly with the MKKs, MKK3 and 6.

Rac1 Actin

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MEKK3

OSM

MKK3 p38

Figure 7. The OSM scaffold. OSM is located in the cytoplasm and interacts with all three members of the p38 module, MEKK3, MKK3 and p38. Upon stimulation with sorbitol OSM and its bound proteins are relocalised to ruffle like structures which are rich in actin. OSM then binds to actin and Rac1 which results in the initiation of the cascade and the activation of p38.

Another potential p38 scaffolding protein is OSM. This protein was identified by Uhlik et al. [2003] in a two hybrid screen using MEKK3, a known p38 MKKK protein as bait. As seen in Figure 7 OSM interacts directly with all three members of a MAPK cascade: p38, MKK3 and MEKK3. OSM also interacts directly with actin and a GTPase Rac1. OSM is located throughout the cytoplasm however upon exposure of cells to sorbitol, OSM and the bound members of the cascade were relocalised to ruffle-like structures rich in actin and Rac1. Once stimulated OSM binds actin and Rac1 and activates p38. Other stimulants were used such as anisomycin and no relocalisation or activation of p38 was observed [Uhlik et

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al., 2003]. This indicates that OSM acts as a scaffolding molecule for a Rac-MEKK3-MKK3 cascade in response to hyperosmotic shock caused by treatment with sorbitol.

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1.2. AKAPs (A-Kinase Anchoring Proteins) The second messenger cyclic AMP (cAMP) is involved in a signal transduction cascade that has been extensively studied. Most of the effects that are mediated by cAMP are through its binding to PKA (cAMP-dependent protein kinase A). Mammalian PKA is a tetramer which is composed of two catalytic subunits and two regulatory subunits. The catalytic subunits are found in three isoforms encoded by three different genes (Cα, Cβ and Cγ) and all isoforms have essentially the same kinetic and physiochemical properties [Colledge & Scott, 1999]. The R subunits are expressed from four different genes RIα, RIβ, RIIα and RIIβ. The PKA tetramer assembles to form a heterotetrameric enzyme of two different forms depending on the R subunits involved. PKA type I contains RIα and RIβ dimers and is primarily cytoplasmic, whereas PKA Type II contains RIIα and RIIβ dimers and associates with specific cellular structures and organelles [McConnachie et al., 2006]. The discrete localisation of Type II PKA within the cell is due to its association with scaffolding proteins called AKAPs. AKAPs are a family of functionally related proteins which are defined based on their ability to bind the R subunits of PKA. AKAPs have two conserved domains. The first is the PKA-binding motif which forms an amphipathic helix of 14-18 residues that interacts with a hydrophobic pocket formed on the N-terminus of the R subunit dimer [Schillace & Scott, 1999; Diviani & Scott, 2001]. Most AKAPs identified have been shown to bind to RII dimers, however some interactions have been shown between AKAPs and RI dimers such as AKAPCE in C. elegans [Angelo & Rubin, 1998]. The affinity of AKAPs for RII is about 100 fold higher than that for RI subunits [Carlisle-Michel & Scott, 2002]. The other conserved feature of AKAPs is a targeting motif which directs the PKA-AKAP complex to discrete locations within the cell such as the plasma membrane, actin cytoskeleton, mitochondria, endoplasmic reticulum, nuclear membrane, centrosomes and vesicles [Schillace & Scott, 1999]. AKAPs are defined as scaffolding proteins as they can colocalise many components of a signal transduction pathway in close proximity to specific substrates [Dell’Acqua et al., 2006]. As these complexes contain signal termination proteins such as phosphatases, and phosphodiesterases and signal transduction enzymes such as kinases, they are capable of upregulating and downregulating specific signalling pathways [McConnachie et al., 2006]. About 50 AKAPs have been identified [McConnachie et al., 2006], however this section will discuss four in detail due to their involvement in regulation of the cytoskeleton. They are gravin, ezrin, CG-NAP/AKAP450 and WAVE-1 a member of the WASP family. Like all AKAPs these proteins not only bind to PKA but also other kinases and phosphatases and they can regulate the bidirectional phosphorylation events at specific compartments in the cell. 1.2.1. Gravin Gravin is a 250 kDa protein which acts as a scaffold by binding PKA, PKC, PP2B (protein phosphatase 2B), β2-AR (β2-adrenergic receptor), GRK2 (G-protein-linked receptor

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kinase 2) and by transiently binding to β-arrestin and clathrin [Fan et al., 2001; Diviani & Scott, 2001]. Gravin contains many domains or sites which facilitate its binding with the associated proteins. Membrane association is achieved through three domains, the first of which is a putative N-myristoylation site. N-myristoylation is a post-translational modification which provides weak and reversible interactions between gravin and the membrane. A second membrane domain was identified which shows homology to the membrane effector domain of MARCKS (myristoylated alanine-rich C-kinase substrate) proteins. This domain has been shown in MARCKS proteins to provide binding to membranes in a manner reversible by Ca2+/calmodulin binding [Malbon et al., 2004a]. Recently this has also been demonstrated for Gravin [Tao et al., 2006]. The third membrane association domain is provided by a set of sites located near the midpoint of the protein. These sequences are conserved amongst Gravin and AKAP79 and their homologs and are involved in the binding of the β2-adrenergic receptor (β2–AR) [Malbon et al, 2004b]. Gravin binds the protein kinases and protein phosphatase PP2B at unique conserved sites. A putative F-actin binding domain resides within the PKC binding domain and may be involved in the translocation of the complex. Also a putative SH3 binding domain was identified with which the non-tyrosine kinase Src can associate. These kinases are known regulators of GPCR-signalling complexes [Malbon et al., 2004b]. A fundamental feature of cell signalling is the attenuation of a signal following stimulation, this process is referred to as desensitization. Protein phosphorylation is often a key feature of this process. In the case of the β2-AR, the desensitization involves phosphorylation of the receptor via the protein kinases PKA, PKC and GRK2 which is illustrated in Figure 8. Gravin has been shown to associate with the β2-AR in unstimulated cells, however upon agonist stimulation this association is enhanced. This brings the other gravin-associated proteins such as PKA, PKC, GRK and PP2B into close proximity to the receptor, permitting an initial attenuation of the signal through phosphorylation of the receptor via the associated kinases (PKA, PKC and GRK). Phosphorylation of the C-terminal region of the membrane bound receptor by GRK allows the recruitment of β-arrestin from the cytosol which can associate with clathrin and translocate the complex to an intracellular endosomal pool via clathrin-dependent endocytosis. This process is necessary for subsequent resensitisation of the receptor. Once the complex has been sequestered to the endosome, PP2B dephosphorylates the receptor thereby disrupting its association with β-arrestin and clathrin and the complex is then translocated back to the plasma membrane [Lin et al., 2000]. Gravin is an example of a mobile scaffold which maintains the ensemble of protein kinases and phosphatases through translocation from the plasma membrane to vesicles involved in trafficking the complex during desensitization and resensitisation [Fan et al., 2001].

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Plasma Membrane

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β2-AR

GRK2

GRK2

β-arrestin

Gravin

Gravin

PKC

PKC PKA

PKA PP2B

PP2B

β2-AR Clathrin Endosomal Pool

GRK2

β-arrestin

Gravin PKC PKA PP2B

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Figure 8. The Gravin Scaffold. Gravin binds several protein kinases and a phosphatase PP2B. The protein is also able to interact with the β2-AR. Upon stimulation of the receptor the gravin-β2-AR interaction is enhanced bringing the gravin-associated proteins in close proximity to the receptor which would allow for attenuation of the signal. GRK2 is phosphorylated at the membrane and recruits β-arrestin from the cytosol. β-arrestin associates with clathrin which translocates the complex to an endosomal pool where PP2B dephosphorylates the receptor. This disrupts the association of β-arrestin and clathrin and the complex is translocated back to the plasma membrane.

1.2.2. Ezrin Ezrin belongs to a family of closely related cytoskeletal proteins termed ERM proteins, named after its three members ezrin, radixin and moesin. These AKAPs link integral membrane proteins to the actin cytoskeleton. ERM proteins are found in all multicellular organisms and share a high degree of sequence identity and structural conservation [Fievet et al., 2007]. The conserved structure consists of three major domains, an N-terminal globular FERM (four-point one ERM) domain of approximately 300 residues, followed by a 160 residue region which is an α-helical domain predicted to form coiled coils and an actin binding C-terminal domain of approximately 90 residues [Finnerty et al., 2003]. The FERM domain is composed of three subdomains, F1, F2 and F3 which are arranged in the shape of a cloverleaf. The central α-helical domain connects the FERM domain to the F-actin binding site present in the C-terminus [Fievet et al., 2007]. The three ERM members are expressed early in development in different tissues or cells within the organism. In mammals Ezrin is expressed mainly in epithelial cells and it is enriched at the apical surface [Berryman et al., 1993], while radixin is present in the liver [Tsukita et al., 1989] and also in the cochlear stereocilia [Pataky et al., 2004; Kitajiri et al., 2004] and moesin is found primarily in

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endothelial cells [Lankes & Furthmayr, 1991] and in some epithelial cells [Berryman et al., 1993]. ERM proteins are conformationally regulated by a head to tail interaction that masks sites for membrane associated proteins and F-actin [Gary & Bretscher, 1995; Reczek & Bretscher, 1998]. As seen in Figure 9 in the inactive configuration the ERM proteins are inactive and localise mainly to the cytoplasm. Activation of ERM proteins is achieved through disruption of the head to tail interaction by binding to PIP2 and phosphorylation of a conserved threonine in the actin binding site of the C-terminus. PIP2 binds in a cleft between subdomains F1 and F3 in the FERM domain, this binding is thought to alter the conformational change leading to disruption of the interaction between C- and N-termini [Hamada et al., 2000]. This interaction has been shown to be critical for translocation of ERM proteins to the plasma membrane. Once localised to the plasma membrane ERM proteins can integrate with additional membrane associated proteins. Plasma membrane localised ERM proteins are then phosphorylated at the conserved threonine residue which stabilises the active configuration [Yonemura et al., 2002; Fievet et al., 2004]. Several protein kinases can phosphorylate ezrin at this residue including Protein kinase C which has been shown to carry out this phosphorylation in vitro [Ng et al., 2001] Additionally, Akt can phosphorylate ezrin at the conserved threonine in response to Na+/glucose transport, leading to association of ezrin with the actin cytoskeleton and the translocation of the Na+/H+ exchanger NHE3 [Fievet et al., 2007]. Ezrin may be phosphorylated and its active state stabilised by different kinases depending on the type of complex ezrin is involved in and the stimuli present. α-helical domain

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F2 F!

F2 F3

C-terminal domain

P

α-helical domain F!

C-terminal domain

F3

PIP2

Cytoplasm [Inactive Configuration]

Plasma Membrane [Active Configuration]

Figure 9. Activation of ERM proteins. ERM proteins contain a conserved structure consisting of an Nterminal FERM domain constructed from three subdomains F1, F2 and F3, followed by a central α-helical domain and a C-terminal domain. ERM proteins are held in the cytoplasm in an inactive configuration through the C-terminal domain binding to the FERM domain. Binding of PIP2 in a cleft between the F1 and F3 subdomains and phosphorylation of a conserved threonine residue in the C-terminus disrupts the interaction of the C-terminus with the FERM domain. This active configuration leads to translocation of ERM proteins to the plasma membrane where it can interact with additional plasma membrane associated proteins.

Of the three family members only ezrin has been shown to be essential for cell survival with ezrin-deficient mice dying shortly after birth [Saotome et al., 2004]. Further discussion Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

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will focus on the A-kinase anchoring, ERM protein ezrin. Ezrin is able to interact with many proteins and these interactions occur either through direct binding or through association of Ezrin with an adaptor protein with the best characterised adaptor being EBP50 (ERM-binding protein of 50kDa). Ezrin interacts directly with membrane proteins in addition to PIP2 and these interactions help localise ezrin to the plasma membrane. Ezrin can directly bind the Na+/H+ exchanger NHE1 and this interaction is required for control of cell shape [Denker et al., 2000]. Ezrin also directly binds the hyaluronan receptor CD44 and this interaction has been shown to be important for cell motility [Legg et al., 2002]. Additional interactions with membrane proteins occur through the interaction of ezrin with EBP50. EBP50 is another scaffolding protein that contains two PDZ domains which can interact with numerous proteins, and a Cterminal region which binds to the FERM domain of Ezrin [Reczek & Bretscher, 1998]. EBP50 interacts with the cystic fibrosis transmembrane conductance regulator (CFTR) [Short et al., 1998], the β2-adrenergic receptor [Cao et al., 1999] and the Na+/H+ exchanger NHE3 [Yun et al., 1998] via its PDZ domains. Ezrin is an AKAP and binds the RII subunit of PKA. Through Ezrin’s interaction with numerous membrane proteins Ezrin is able to localise PKA to the membrane where it can phosphorylate its substrates including CFTR and NHE3 which it can activate or inhibit, respectively [Sun et al., 2000a; Sun et al., 2000b; Dransfield et al., 1997; Lamprecht et al., 1998; Weinman et al., 2000]. The complexes formed are important for trafficking of membrane proteins such as the recycling of β2-andrenergic receptors from the endosome to the plasma membrane after agonist stimulated endocytosis [Cao et al., 1999] and the localisation of the CFTR [Moyer et al., 1999, 2000] Ezrin forms multiprotein complexes which are essential for the morphogenesis of the apical domain of epithelial cells, including orginisation of actin filaments and delivery of membrane proteins. Ezrin has also been shown to play an important role in Fas-mediated apoptosis. Fas is a member of the death family proteins and when activated recruits numerous proteins to form the death inducing signalling complex that initiates the apoptotic cascade. Ezrin directly binds to Fas via its FERM domain and may play a role in localising Fas and its associated molecules to the plasma membrane to receive signals from apoptotic stimuli [Fais et al., 2005]. The Fas-Ezrin interaction is essential for Fas-mediated apoptosis as mutants in which Ezrin was down regulated showed defects in apoptosis [Parlato et al., 2000]. Through its association with EBP50 ezrin is also important for linking lipid rafts in T cells to the actin cytoskeleton, and plays a negative role in immune synapse formation [Itoh et al., 2002]. These examples show how ezrin can act as a scaffold binding numerous receptors, cell adhesion molecules and scaffolding proteins to link various proteins from the plasma membrane to the underlying actin cytoskeleton. 1.2.3. CG-NAP (Centrosome and Golgi Localised Protein Kinase N (PKN)-Associated Protein) Also Called AKAP450/AKAP350 CG-NAP is a giant protein predicted to form a coiled coil over most of its length except for small regions (~200 amino acids) at both the C- and N-terminal regions [Gillingham & Munro, 2000]. The protein is localised to the centrosome throughout the cell cycle, the midbody at telophase and the Golgi apparatus at interphase in cultured cell lines [Takahashi

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et al., 1999]. The protein interacts with many components of signal transduction cascades including three protein kinases (PKA, PKCε and PKN) and two phosphatases (PP1 and PP2A) [Takahashi et al., 2000], phosphodiesterase 4D [Tasken et al., 2001], casein kinase 1δ/ε [Sillibourne et al., 2002], cdc42 interacting protein 4 [Larocca et al., 2004] and Ran [Keryer et al., 2003]. CyclinE-cdk2 has been shown to be involved in the process of centrosome duplication [Hinchcliffe et al., 1999] and Nishimura et al. [2005] have shown that CG-NAP recruit Cyclin E-cdk2 to the centrosome and therefore may be involved in centrosome duplication. CG-NAP contains two RII binding sites at its C-terminus and may therefore bind two PKA holoenzymes in one signalling complex. The centrosomal targeting domain resides in the C-terminus and shows high homology to another centrosomal AKAP, pericentrin. These two AKAPs may compete with each other for centrosome binding or they may work in a coordinated fashion [Carlisle-Michel & Scott, 2002]. CG-NAP also binds to the N-terminal region of PKN, a Rho-activated kinase which phosphorylates intermediate filament proteins indicating that the complex may be important for cytoskeletal reorganisation events. PKN is a substrate of PP2A which can dephosphorylate PKN and decrease its kinase activity. PP2A binds to CG-NAP through its regulatory subunit. An immature, non- or hypophosphorylated PKCε can also associate with CG-NAP until it is sufficiently phosphorylated whereby it dissociates from the scaffold and can respond to incoming second messenger signals [Takahashi et al., 2000]. CG-NAP therefore acts as a scaffold by localising many kinases and phosphatases and it coordinates the activity of these enzymes at centrosomes and in the Golgi apparatus. 1.2.4. WAVE1 WAVE1 is a member of the WASP family of proteins. This family is named after its founding member Wiskott-Aldrich syndrome protein (WASP) which, when mutated, causes a rare X-linked immunodeficiency disease [Westphal et al., 2000]. WAVE1 functionally couples small Rho GTPases, Rac and Cdc42 (activated by ligand binding a receptor) to the Arp2/3 complex, which is a group of seven related proteins that nucleate actin polymerization, and can thereby elicit changes in the actin cytoskeleton in response to the signal [Yamazaki et al., 2005]. The C-terminus of WAVE1 contains an acidic domain which is responsible for binding and activating the Arp2/3 complex [Calle et al., 2004]. WASP proteins also contain a central proline rich sequence which facilitates binding to several SH3 containing proteins and to an actin binding protein, profilin. WASP members also contain a verprolin homology (VPH) domain that is responsible for binding monomeric actin [Westphal et al., 2000]. WAVE1 has been shown to bind to the RII subunit of PKA and it does this through the VPH domain. It appears that actin competes for the RII binding site, the significance of this is unclear but sufficient concentrations of actin may displace the anchored PKA. Thus, PKA may be dynamically regulated at sites of actin reorginisation [Westphal et al., 2000]. WAVE1 can also form homo- or hetero-dimers with other WAVE isoforms (WAVE2 & 3). WAVE1 may therefore tether PKA to the cytoskeleton through the actin based interactions of WAVE2 or 3. WAVE1 can also bind a tyrosine kinase Abl (Abelson) at its SH3 domain. Rac-activated stimulation by Platelet-derived growth factor (PDGF) results in the rapid redistribution of

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WAVE1, PKA and Abl from focal adhesions to sites of actin polymerization. The assembly of the WAVE1 complex is dependent on this extracellular signal and it appears that WAVE1 not only facilitates the coupling of Rac to the Arp2/3 complex but also coordinates the location and action of PKA at sites of actin reorganisation [Diviani & Scott, 2001].

[Diviani & Scott, 2001]. Figure 10. WAVE1, an AKAP at the cytoskeleton. WAVE1 forms homo- or hetero-dimers with other WAVE isoforms, WAVE2/3. Activated Rac1 or Cdc42 binds to WAVE1 and causes the relocalisation of WAVE1 and its associated proteins, such as the kinases Abl and PKA to sites rich in actin. WAVE1 can directly bind actin, actin binding proteins and the Arp2/3 complex to promote actin polymerisation. Actin can compete with PKA for binding to WAVE1 and therefore sufficient concentrations of actin may displace PKA. (Reproduced with permission of the Company of Biologists).

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1.3. PSD (Post Synaptic Densities) Scaffolding Proteins PSDs are disc-like structures approximately 30-40 nm thick and 400 nm wide which are observed under electron microscopy as thickenings of the postsynaptic membranes [Ziff, 1997]. These areas are specialised complexes which contain and organise many components of the postsynaptic response including receptors for the neurotransmitter glutamate, receptor associated proteins, signal transduction molecules, cytoskeletal and regulatory proteins [Irie et al., 2002]. There is an abundance of proteins in the PSD, some estimating the total number to be as many as a few hundred [Sheng, 2001]. The PSD is highly enriched in three types of glutamate receptors - the NMDA (Nreceptors, the AMPA (α-amino-3-hydroxy-5-methyl-4methyl-D-aspartate) isoxazolepropionic acid) receptors and the mGluRs (group 1 metabotropic glutamate receptors) [Sheng, 2001]. These receptors are specifically targeted to the postsynaptic membrane where they are responsible for fast signalling at these synapses. The NMDA receptors are a consistent, stable feature of excitatory synapses, they are permeable to monovalent and calcium ions both of which induce synaptic plasticity, such as long term potentiation. AMPA receptors are glutamate-gated monovalent cation channels whose activation provides most of the postsynaptic depolarisation that drives neuronal firing [Fukata et al., 2005]. Synaptic transmission at low frequency is dependent primarily on AMPA receptors

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[Leonard et al., 1998]. The presence of AMPA receptors in excitatory synapses is highly variable with a significant fraction of the synapses not containing these receptors [Sheng, 2001]. The distribution of AMPA receptors in excitatory synapses is dependent on synaptic activity. AMPA receptors are dynamically regulated with receptors being delivered into and out of the synapse rapidly in response to stimulus [Sheng, 2001]. The mGluRs are linked to the activation of phospholipase C and mediate excitatory affects by eliciting a release of calcium from intracellular stores [Tappe & Kuner, 2006]. The N terminus of these receptors is extracellular followed by a transmembrane region and the C terminus is located on the intracellular side of the membrane [Leonard et al., 1998]. There is a complex network of proteins which transmit the glutamate signal received by the various receptors through signalling cascades to effector substrates such as the cytoskeleton. These proteins interact with each other via numerous scaffolding proteins. There are a large number of scaffolding proteins present in the PSD and this section will focus on those involved in the regulation of the cytoskeleton including the MAGUK (membrane-associated guanylate kinases) family, cortactin, homer, and shank scaffolding proteins. 1.3.1. MAGUK (Membrane-Associated Guanylate Kinases) Family The MAGUK family of scaffolding proteins are membrane-associated cortical cytoskeletal proteins which are involved in the trafficking of glutamate receptors. Members of this family include PSD-95/SAP90, PSD93, SAP97 and SAP102 [Schluter et al., 2006]. The most extensively studied of these are PSD-95 and SAP97 and they are the focus of this section. The MAGUKs consist of three PDZ domains at the N-terminus followed by an SH3 domain, and a C-terminal guanylate kinase (GK) domain. Both PSD-95 and SAP97 are able to bind to AMPA receptors, however they do so via different mechanisms. SAP97 is able to directly bind to the GluR1 subunit of AMPA receptors via its PDZ domain [Schluter et al., 2006], whereas PSD-95 binds to the receptor indirectly through a protein called stargazin. Stargazin is a transmembrane protein which can bind to all four subunits of AMPA receptors and traffic them to the plasma membrane. The C-terminus of stargazin binds PSD-95 and links the scaffolding protein to the receptor [Fukata et al., 2005]. PSD-95 also directly binds to the C-terminus of NR2 subunits of the NMDA receptors via the first and second PDZ domains whereas SAP97 does not coprecipitate with NMDA receptors and hence, may be involved in the regulation of AMPA but not NMDA receptors [Leonard et al., 1998]. PSD-95 and SAP97 are able to interact with protein kinases, phosphatases, cytoskeletal proteins and signal transduction molecules via their five proteinprotein interaction domains. Some examples of PSD-95 and SAP-97 binding partners are given below. PSD-95 and SAP97 associate with each other as a heteromeric complex via the Nterminal segment of SAP97 and the SH3 domain of PSD-95. This interaction may influence the interactions of each of these MAGUKs with other proteins. For example SAP97 which is normally not located in dendritic spines was strongly recruited to these spines when PSD-95 was overexpressed. Conversely, PSD-95 normally triggers the accumulation of AMPA receptors to synaptic spines however overexpression of SAP97 strongly inhibited this process [Cai et al., 2006]. It seems that each of these MAGUKs may work in synergy with each other promoting and inhibiting interactions with other proteins. The MAGUKs are important for

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the localisation and maturation of glutamate receptors and the coupling of these receptors to a complex of proteins involved in cytoplasmic signalling pathways.

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Table 2. Interacting partners of PSD-95 and SAP97 Scaffolding Protein PSD-95

Protein Domain PDZ1, 2 and 3

Interacting Protein SynGAP

PSD-95

GK

SPAR

PSD-95

GK

PSD-95

PDZ

PSD-95

Role of interacting protein

Reference Rockliffe & Gawler, 2006

GKAP

Regulates Ras activation in response to NMDA receptor stimulation A GTPase for Rap. Regulates the size and shape of dendritic spines. Couples PSD-95 to Shank

PDZ3

Neuronal nitrate synthase (nNOS) Neuroligins

Facilitates activation of nNOS by NMDA receptor mediated Ca2+ influx Synaptic surface proteins

SAP97

PDZ

NrCAM

A cell adhesion molecule

SAP97

N-terminal domain

CASK

Another MAGUK

Sheng, 2001

Sala et al., 2001 Ishii et al., 2006 Levinson et al., 2005 Dirks et al., 2006 Lee et al., 2002b

1.3.2. Cortactin There are several isoforms of cortactin, a ubiquitous protein present in the majority of cell types, including neurons where it is present in the PSDs [Weed et al., 1998]. Overexpression of cortactin has been linked to several human diseases including breast and bladder cancer [Lua & Low, 2005]. Cortactin consists of numerous protein interaction domains including an N-terminal acidic region which has been shown to bind to the Arp2/3 complex and activate it. This is followed by a region containing six complete copies and one incomplete copy of a 37 amino acid tandem repeat responsible for binding F-actin. The next domain is an alpha helical region which is rich in proline, threonine, serine and tyrosine residues that harbour sites of phosphorylation [Lua & Low, 2005]. The kinase Fyn, a protein involved in postsynaptic signalling pathways phosphorylates cortactin in this region [Iki et al., 2005]. The C-terminal region contains an SH3 domain that interacts with numerous proteins including cortactin-binding protein 1 and CBP90, two brain-specific proteins and ZO1, a cellcell adhesion complex protein [Webb et al., 2006; Campbell et al., 1999]. The SH3 domain of cortactin also interacts with regulators of the actin cytoskeleton including dynamin2 [Mizutani et al., 2002], GTPase regulators including BPGAP1 which is a Rho GTPase activating protein [Lua & Low, 2005], N-WASP [Kinley et al., 2003], and WASP interacting protein (WIP) [McNiven et al., 2000] as well as an additional PSD scaffolding protein, Shank [Naisbitt et al., 1999].

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Cortactin does not bind any glutamate receptors directly but does so indirectly through binding additional scaffolding proteins such as Shank. The PDZ domain of Shank binds to GKAPs (guanylate kinase associated proteins) which in turn bind to the GK domain of PSD95. As mentioned earlier PSD-95 interacts directly with transmembrane receptors in the PSD such as the NMDA and AMPA receptors [Daly, 2004]. Cortactin is able to promote the nucleation of actin in response to glutamate binding to various receptors in the PSD. Thereby cortactin acts as a scaffold linking postsynaptic signalling in the PSD to the actin cytoskeleton. 1.3.3. Homer Homer proteins are scaffolding molecules that regulate the localisation of group 1 metabotropic glutamate receptors (mGluR) [Irie et al., 2002]. Three Homer genes have been identified, Homer 1, 2 and 3 and these genes are alternatively spliced to form 15 long and short isoforms [Duncan et al., 2005]. The long forms are tetramers which oligomerise due to their C-terminal coiled coil domains. This oligomerisation allows Homer to interact with four ligand proteins at the same time [Hayashi et al., 2006]. This coiled coil domain is absent in the short forms of Homer. The other domain present in all Homer isoforms is the N-terminal EVH1 domain. The EVH1 domains are highly conserved and bind to a proline rich motif present in mGluRs [Xiao et al., 2000]. The long forms are constitutively expressed, whereas the short forms are expressed in an activity dependent manner. The long forms due to their ability to oligomerise are able to couple postsynaptic proteins to the receptor, whereas the short forms which are unable to oligomerise, act to compete with the long form of Homer and block its function [Hayashi et al., 2006]. Homer is able to bind many proteins apart from the mGluR through its EVH1 domain. These proteins include amongst others the receptors for ryanodine [Hwang et al., 2003] and IP3 (inositol [1,4,5]-triphosphate) [Yuan et al., 2003], members of the Rho family of small GTPases [Shiraishi et al., 1999], Shank [Tu et al., 1999] an additional PSD scaffolding protein (discussed in section 1.2.4.4), TRPC [transient receptor potential canonical] channels [Yuan et al., 2003], dynamin III [Gray et al., 2003], syntakin 13 [Minakami et al., 2000] and actin [Shiraishi et al., 1999]. Homer is able to regulate the trafficking and clustering of mGluRs through these various interactions. For example dynamin III, a protein involved in endocytosis enables Homer to mediate the trafficking of mGluRs. Additionally, the Homer complex can promote the accumulation of F-actin in synapses and as it binds actin directly may link the mGluRs to the actin cytoskeleton. Homer aids in the targeting and clustering of mGluRs, and signalling proteins to the plasma membrane. 1.3.4. Shank Shank proteins are relatively large scaffolding proteins (>200 kDa in molecular mass) and three isoforms have been identified named Shank 1 to Shank 3 [Sheng & Kim, 2000]. These isoforms are generated by alternative splicing events and their presence and level of expression varies amongst different tissues. All three isoforms however are found in the PSD [Lim et al., 1999]. The Shank members share a similar domain structure consisting of several protein interaction domains. The structure begins with 5-6 N-terminal ankyrin repeats, followed by an SH3 domain, a PDZ domain, a long proline rich region and a sterile alpha

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motif (SAM) domain [Gundelfinger et al., 2006]. Numerous Shank binding partners have been identified these include α-fodrin which binds to the ankyrin repeats. α-fodrin interacts with actin and calmodulin and thereby links Shank to the actin cytoskeleton [Boeckers et al., 2001]. The ankyrin repeats are also responsible for Shank binding to sharpin, a PSD protein which contains a region homologous to a protein kinase C binding protein [Lim et al., 2001]. The SH3 domain of Shank binds Densin 180, a membrane targeted PSD protein [Quitsch et al., 2005]. The PDZ domain is able to facilitate the interactions with numerous proteins including GKAP (guanylate kinase-associated protein) via its C-terminus which aids in recruitment of Shank to postsynapses [Naisbitt et al., 1999]. GKAP interacts with the PSD scaffolding protein PSD-95 and thereby this interaction links Shank to NMDA and AMPA receptors [Boeckers et al., 2002]. The PDZ domain is also responsible for the interaction of Shank with numerous transmembrane receptors such as the somatostatin receptor, the α-latrotoxin receptor, the cystic fibrosis transmembrane conductance regulator and the Cav1.3 L-type Ca2+ channel [Gundelfinger et al., 2006].

[Sheng & Kim, 2000]. Figure 11. Scaffolding proteins in the PSD. The three transmembrane glutamate receptors of the PSD are bound to different PSD scaffolding proteins. The AMPA receptors (AMPAr) are bound directly by GRIP (Not discussed in the text, also bound by MAGUK family members either directly or indirectly and not shown on this figure) The NMDA receptors [NMDAr] interact with the MAGUK family of scaffolding proteins represented by PSD-95 in this figure, and the mGluR interact with Homer. Cortactin is a scaffolding protein which interacts with F-actin. Shank is able to interact with all of the scaffolding proteins depicted in this figure and as such acts as a master scaffold linking all the glutamate receptors to the actin cytoskeleton. (Reproduced with permission of the Company of Biologists).

The proline rich region contains proline rich motifs which are binding sites for many interaction domains such as SH3, EVH1 and WW domains. Homer, a scaffolding protein discussed earlier, is able to interact with the proline rich region of Shank via its EVH1

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domain. This interaction links Shank proteins to the mGluRs through their interaction with Homer [Du et al., 1998]. Shank is able to interact with an additional scaffolding protein cortactin (discussed in Section 1.2.4.2) [Naisbitt et al., 1999]. This interaction links Shank to the regulation of the actin cytoskeleton. Shank is found in a deeper part of the PSD than other scaffolding proteins such as the MAGUK family members which are located close to the membrane [Sheng & Kim, 2000]. Shank is able to interact with numerous additional PSD scaffolding proteins and indirectly to all three glutamate receptors and as such may act as a master scaffold linking all receptor complexes in the PSD as illustrated in Figure 11 [Sheng & Kim, 2000].

1.4. Actin Binding Protein Scaffolds

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The dynamic reorganisation of the cytoskeleton which occurs within seconds after chemotactic stimulation is facilitated by a large family of proteins called actin binding proteins (ABP). ABP’s have been identified as the major regulators of these dynamic rearrangements that occur when a cell moves, divides or takes up particles and liquid. They control the polymerisation of the G-actin monomers into filaments, the steady state equilibrium between F and G-actin and the 3D organisation of the filamentous network. Many of these actin binding proteins act as scaffolds in signalling pathways interacting with many proteins in addition to actin. Two such actin binding scaffolds will be described here, IQGAP and filamin. 1.4.1. Filamin (ABP280) Filamin belongs to a large family of actin binding proteins which are responsible for forming and stabilizing 3D cortical actin networks. Homologs of filamin are present in all eukaryotes and in mammals three filamin isoforms have been identified, FLNA, B and C. The three isoforms share strong sequence homology over the entire sequence with the exception of the two hinge regions which show greater divergence [Feng & Walsh, 2004]. FLNA and B are ubiquitously expressed in all tissues whereas FLNC is largely restricted to skeletal and cardiac muscle. Filamins have been shown to be essential for normal development, with mutations leading to developmental defects of many organs including the brain, bone and cardiovascular system [Feng & Walsh, 2004]. Filamin A is a 280 kDa protein and as it is the most abundant of the three isoforms will be the focus of this discussion. Filamin is comprised of two major domains and forms a homodimer. At the N-terminus resides an F-actin binding domain consisting of two calponin homology (CH) domains of approximately 110 residues named after the first protein in which they were identified, calponin. This actin binding domain is shared amongst many actin binding proteins including α-actinin, β-spectrin and dystrophin. Following the actin binding domain is the rod domain which is comprised of a series of 24 repeat elements that form sandwiches of β-sheets resembling the structure of immunoglobulin domains. Sequence insertions immediately before repeats 16 and 24 predict two hinges which are susceptible to cleavage by the protease calpain. The last repeat (24th) is responsible for the dimerisation of the protein which enables filamin to crosslink two F-actin filaments. As with other immunoglobulin-like domains these

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domains are responsible for protein-protein interactions. Filamin has been reported to interact directly with more than 50 proteins [Nakamura et al., 2006], and further research will no doubt identify more interacting partners. Figure 12 provides an illustration of some of the filamin interacting proteins and the locations of their binding sites. These binding partners include several receptors including members of the GPCR (G-protein coupled receptors) such as dopamine D2/D3 and the calcium sensing receptor, the Fcg receptor and the insulin receptor, platelet glycoproteins Ibα, β1 and β2 integrins, many signalling proteins including Ras-related small GTPases such as Rac, Rho, Cdc42, RalA and FilGAP, GEFs including Trio, Lbc, Pak1 and ROCK, kinases such as PKC, PKA and members of the MAPK family and other structural proteins such as caveolin-1 [Scott et al., 2006; Ohta et al., 2006; Awata et al., 2001; Hjalm et al., 2001]. Most of these binding partners interact with the C-terminal region of filamin from repeats 14-24. Only three proteins have been shown to interact with filamin between repeats 1-14. These include PKC illustrated in Figure 12 which has been shown to bind filamin between repeats 1-4 and repeats 22-24, however the significance of binding between repeats 1-4 has not been established [Tigges et al., 2003]. The other two proteins are furin which binds filamin between repeats 13 and 14 [Liu et al., 1997] and Cyclin B1 which binds filamin repeat 9 [Cukier et al., 2007].

Figure 12. Filamin interacting proteins. This figure is a schematic representation of some of the known filamin interacting proteins and the region of filamin to which they bind. As evident in this figure most proteins tend to interact with the C-terminal repeats in the rod domain except for actin which is bound by the N-terminal actin binding domain. (Reprinted by permission from Macmillan Publishers Ltd. (Nature Cell Biology), Feng & Walsh, 2004, copyright 2004).

Filamin contains many phosphorylation consensus sequences most of which are located in or near the hinge regions. Protein kinases which phosphorylate filamin include PKA [Jay et al., 2000], PKC [Tigges et al., 2003], p21-activated kinase [Vadlamudi et al., 2002], Cdk1 [Cukier et al., 2007] and CaM kinase II (Ca2+/calmodulin-dependent protein kinase II) [Ohta & Hartwig, 1995]. Phosphorylation by these kinases provides a mechanism of regulation of filamin. PKA-mediated phosphorylation provides increased resistance to the calcium

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activated protease calpain and can alter filamin’s interaction with Rho GTPases. Phosphorylation by CaM kinase II or Cdk1 decreases filamin’s actin crosslinking activity [Cukier et al., 2007; Ohta & Hartwig, 1996]. Filamin acts as a scaffold in diverse signal transduction cascades by binding various receptors and intracellular signalling molecules such as protein kinases and G proteins and physically linking these signalling complexes to the actin cytoskeleton. Several examples of signalling pathways in which filamin acts as a scaffold are described below. Filamin is involved in a MAPK cascade involving MEKK1, SEK-1 and SAPK (a member of the ERK family). Filamin binds constitutively to SEK-1 (a MKK) between repeats 21 to 23. Bound SEK1 is then able to be phosphorylated by the upstream kinase MEKK1. Phosphorylated SEK1 is then able to phosphorylate and activate SAPK which phosphorylates c-Jun and initiates programmed cell death. Although filamin does not interact directly with the other two members of the cascade, mutational analysis has revealed that filamin is required for efficient signalling between the three components [Marti et al., 1997]. Filamin has been shown to directly bind other MKK’s including MEK1 and MKK4 which activate ERK and JNK respectively. Filamin does not bind the other members of the MAPK cascade in these signalling pathways either, but again is essential for efficient signalling [Scott et al., 2006]. Activation of SAPK was shown to be dependent on the expression of filamin, with filamin-deficient cell lines unable to activate SAPK in response to the cellular stress TNF-α (Tumour Necrosis Factor). These cells also exhibited an 80 % reduction in stimulation via LPA (Lysophosphophatidic acid) [Marti et al., 1997]. Direct regulation of filamin may occur through its association with receptors and GTPase’s, however the specific components were not identified for this pathway. Likewise filamin may be regulated via phosphorylation by the protein kinases listed earlier-filamin has been shown to be phosphorylated in response to LPA and other stimuli by kinases such as Ribosomal S6 kinase (RSK) [Woo et al., 2004; Marti et al., 1997]. The exact mechanisms of this pathway are still to be elucidated, however it is clear that filamin plays a direct and essential role in the MAPK cascade leading to activation of SAPK in response to both LPA and TNF-α. Filamin is able to interact with other scaffolding proteins such as β-arrestin to form an additional level of complexity through a double scaffold. Filamin binds β-arrestin via its 22nd repeat which is in close proximity to the binding sites for the MAP2K members MEK1 and MKK4 [Scott et al., 2006]. β-arrestin has been shown in other MAPK signalling pathways to bind MAP3K members including ASK1 and also to the MAPK JNK, but it does not bind directly to the MAP2K [Mc Donald et al., 2000]. Likewise in the ERK signalling pathway βarrestin may bind the MAP3K members and ERK and interact indirectly through filamin with the MAP2K. As filamin is concentrated to the cell periphery where it binds the cytoskeleton, filamin may localise β-arrestin and the other MAPK members to this cell compartment [Scott et al., 2006]. Filamin is able to bind furin, a protein responsible for the proteolytic maturation of proproteins within the trans-Golgi and endosomal network. This interaction occurs between repeats 13 and 14 of filamin and the cytosolic domain of furin. Furin is primarily located in the trans-Golgi network but cycles to the cell surface where it is bound by filamin. Filamin can tether furin to the surface and regulate the rate of furin internalization. Filamin deficient

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cell lines internalized furin at a 2.3 fold higher rate, than the wild type. The efficient sorting of furin from the early endosomes to the Golgi network was also altered in filamin deficient cell lines suggesting that filamin not only controls the rate of furin internalization but also aids in the efficient trafficking of furin from the early endosome stage onwards [Liu et al., 1997]. Filamin can concentrate furin to regions of the cell surface that contain potential substrates including MT-MMP1 (membrane type 1 matrix metalloprotease) which localises to the cell surface due to its cytosolic domain. MMP are proteins which are involved in the degradation of extracellular matrix proteins [Zhu et al., 2007]. Colocalised furin may cleave this substrate at the cell surface. The internalization motif of furin is masked when bound to filamin and hence the rate of internalization is reduced. However disruption of this interaction through unknown regulatory mechanisms would enable the motifs to interact with the internalization machinery and relocate furin to intracellular compartments. Filamin is also important for appropriate localisation of late endosomes and lysosomes in the cell as demonstrated by fluorescence microscopy with filamin-deficient cell lines showing incorrect localisation of vesicles [Liu et al., 1997]. Other mutational experiments have demonstrated the essential role of filamin in many signalling processes. Glogauer et al. [1998] have shown that filamin binds to β1-integrin and this interaction localised filamin to focal adhesions in response to shear stress. This resulted in modulation of the cytoskeleton to stiffen the cells which made them more resistant to physical strain. Cells lacking filamin were unable to produce a stiffening response to shear stress. Filamin has also been shown to be involved in the regulation of cell polarity with filamin null cells exhibiting impaired locomotion which is partially caused by their reduced ability to polarise [Ohta et al., 2006]. Filamin can crosslink actin filaments and also interacts with many other proteins involved in the regulation of the actin cytoskeleton. These proteins include Rho GTPases, GEFs (guanine nucleotide exchange factors), Pak1 which promotes actin assembly and many transmembrane proteins involved in locomotion and adhesion. Filamin is able to localise these proteins to promote retraction and suppress leading lamellae formation which aids in the regulation of cell polarity [Ohta et al., 2006]. 1.4.2. IQGAP IQGAP proteins belong to a family of conserved actin binding proteins which have been identified in many eukaryotic organisms from yeasts to mammals [Briggs & Sacks, 2003]. They can form dimers with each monomer binding F-actin and crosslinking these filaments [Bashour et al., 1997; Fukata et al., 1997]. In mammals three isoforms have been identified IQGAP1, 2 and 3. All three isoforms share a high degree of sequence similarity and domain structure yet differ in tissue distribution. IQGAP1 is expressed in all tissues, IQGAP3 is enriched in the brain and the lung and IQGAP2 is predominantly expressed in the liver and stomach but has been identified in other tissues including platelets [Brandt & Grosse, 2007]. IQGAP1 has been the most extensively studied and will be the focus of this section. IQGAP1 is a 189 kDa protein that contains a conserved domain structure consisting of an N terminal CH (Calponin homology) domain, a WW domain, an IQ domain, a GAP-related domain and a RasGAP domain, shown schematically in Figure 12. The CH domain binds to

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F-actin (as described for filamin, Section 1.4.1) [Mateer et al., 2004], the WW domain is a known protein interaction domain (refer to Table 1) however no binding partners for this domain in IQGAPs have been identified to date. The next two domains, the IQ and GAPrelated domains lend the protein its name. The IQ domain in IQGAP1 is constructed from four tandem IQ motifs which are involved in binding numerous proteins including calmodulin [Hart et al., 1996; Ho et al., 1999; Joyal et al., 1997], myosin essential light chain [Weissbach et al., 1998], and a Zn2+ and Ca2+ binding protein, S100B [Mbele et al., 2002]. The GAP-related domain shares high sequence similarity to Ras GTPase activating proteins yet does not contain any GAP activity [Briggs & Sacks, 2003]. This domain mediates the binding of Rho GTPases, Cdc42 and Rac1 but not RhoA or Ras [Hart et al., 1996; Ho et al., 1999; Kuroda et al., 1996]. Finally, the Cterminal RasGAP domain interacts with the microtubule binding protein CLIP70 (cytoplasmic linker protein 70) [Fukata et al., 2002], E-cadherin [Kuroda et al., 1998] and βcatenin [Briggs et al., 2002].

Figure 13. IQGAP interacting proteins. This figure provides a schematic representation of the known binding partners of IQGAP and the protein interaction domain to which each binds. No binding partners have been identified as yet for the WW domain, but in other proteins this domain has been shown to mediate proteinprotein interactions.

The multiple partners of IQGAP indicate that it is an important protein involved in cell movement. In support of this IQGAP has been shown to be necessary for cell-cell adhesion [Noritake et al., 2005] and cytokinesis [Machesky, 1998], is overexpressed in cancer cells [Clark et al., 2000; Nabeshima et al., 2002] and is localised to lamellipodia of motile cells [Kuroda et al., 1996]. IQGAP binds Cdc42 and inhibits its GTPase activity, which stabilises the active GTP form of Cdc42. In turn, active Cdc42 enhances the ability of IQGAP to crosslink F-actin filaments. Active Cdc42 also stimulates N-WASp, a nucleation promoting factor which stimulates the Arp2/3 complex to generate branched actin meshworks from preexisting filaments [LeClainche et al., 2007]. N-WASp contains several domains, two of which are the CRIB (Cdc42-Rac interactive binding domain) and the VCA or C-terminal catalytic domain. These two domains interact with each other and hold the N-WASp protein in an inactive state. Binding of Cdc42 to the CRIB domain abolishes its interaction with the VCA domain which is then free to interact with Arp2/3, G-actin and an actin filament barbed end [Kim et al, 2000]. The C-terminal region of IQGAP can also interact with the CRIB

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domain of N-WASp and appears to act in synergy with Cdc42 to activate N-WASp [LeClainche et al., 2007]. IQGAP also stimulates actin polymerisation through formins. Formins are actin nucleating proteins which initiate the formation of linear actin filaments through processive elongation at the growing (barbed) end. IQGAP1 binds to the Rho-activated form of Dia1 (a formin protein) and recruits it to sites of actin assembly such as the leading edge of migrating cells. It is thought that IQGAP1 binding to Dia1 stabilises its active conformation [Brandt et al., 2007]. IQGAP1 is involved in cell-cell adhesion. Cell to cell adhesion is achieved through the interaction of cadherins present on adjacent cells. Cadherins are a family of cell surface adhesion molecules which are regulated by Ca2+. The cytoplasmic domain of E-cadherin binds to β-catenin or γ-catenin and this complex is coupled to the actin cytoskeleton through α-catenin [Bracke et al., 1996]. IQGAP1 binds directly to E-cadherin and β-catenin and reduces the ability of E-cadherin to interact with the cytoskeleton. This weakens the cell-cell attachment which is necessary for cells to detach from each other and move [Kuroda et al., 1998; Li et al., 1999]. In addition to interaction with the actin cytoskeleton IQGAP1 also interacts with the microtubule cytoskeleton. Microtubules are a main element of the cytoskeleton and are essential for cell division, migration and vesicle transport. IQGAP1 interacts with a microtubule tip protein CLIP70 at the leading edge of migrating cells. This interaction localises IQGAP and its binding partners to microtubules linking the plus ends of microtubules to the actin meshwork and promotes cell polarisation during migration [Fukata et al., 2002]. Because it interacts with both the actin cytoskeleton and the microtubular cytoskeleton, IQGAP provides a mechanism for crosstalk between them and the pathways described above. IQGAP’s role in the processes is regulated by its interactions with Cdc42 and Ca2+/calmodulin and through phosphorylation. Activated Cdc42 enhances both the actin crosslinking activity of IQGAP [Fukata et al., 1997] and its interaction with the microtubule cytoskeleton through stimulation of binding to CLIP70 [Fukata et al., 2002]. Active Cdc42 and Rac1 additionally inhibit the interaction of IQGAP with E-cadherin and β-catenin. This prevents IQGAP from inhibiting cell-cell adhesions [Kuroda et al., 1998]. Like many actin binding proteins IQGAP is regulated by Ca2+/calmodulin. Calmodulin interacts with IQGAP1 through the four tandem IQ motifs and this interaction is mediated by Ca2+. Calmodulin can interact with IQ1 and IQ2 only when Ca2+ is present whereas calmodulin interacts with IQ3 and 4 regardless of the presence or absence of Ca2+. The binding of Ca2+/calmodulin prevents IQGAP from stabilising active Cdc42 and inhibits actin polymerisation. Conversely, the displacement of active Cdc42 enables the interaction of IQGAP with E-cadherin and β-catenin and cell adhesion is inhibited [Li & Sacks, 2003]. Phosphorylation of IQGAP provides an additional level of regulation. IQGAP is held in an inactive configuration through interaction of its C-terminal and N-terminal domains. Phosphorylation of Serine1443 relieves this inhibition allowing IQGAP to interact with its binding partners such as Cdc42 and N-WASp to promote actin polymerisation [LeClainche et al., 2007]. IQGAP has been shown to be phosphorylated by PKC, and this phosphorylation

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event was shown to alter the conformation of IQGAP so that the C-terminus is free to interact with Cdc42 [Brandt & Grosse, 2007]. Most of the proteins with which IQGAP interacts are involved in regulation of the actin or microtubule cytoskeleton. These interactions enable IQGAP to act as a scaffold in many processes which involve the regulation of the cytoskeleton such as in cytokinesis, cell invasion and metastasis, cell migration and cell polarity.

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1.5. Conclusion The cytoskeleton is a highly dynamic structure which can rapidly reorganise in response to various stimuli. This reorganisation is important for a cell to respond to various stimuli, for intracellular and intercellular transport, for maintenance of cell shape, cell division, growth and development. In order for a cell to reorganise its cytoskeleton so rapidly and in response to various stimulation cues, numerous proteins are employed in signal transduction pathways to aid in the cytoskeletal rearrangement. These proteins range from receptors, G proteins, signalling molecules and second messengers to kinases, phosphatases and cytoskeletal components. These proteins are often involved in many pathways and located in different compartments of the cell. Scaffolding proteins are important proteins in the regulation of the cytoskeleton as they are able to localise many of the components of a particular pathway to a particular location within the cell and assist in the efficient relaying of the response. Scaffolding proteins can tether members of a complex through protein interaction domains which abound on these proteins. They can also sequester binding partners to a particular pathway, thereby avoiding crosstalk between components involved in multiple pathways. Scaffolding proteins can also stabilise weak interactions between particular proteins in a cascade and catalyse activation of the pathway components. Due to these roles scaffolding proteins are essential members of signalling pathways that regulate the cytoskeleton and without them the efficiency, selectivity and rapidity of responses would not be possible. Many scaffolding proteins have been identified to date, however growing interest will no doubt lead to the identification of many more.

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[174] Scott, M. G. H., Pierotti, V., Storez, H., Lindberg, E., Thuret, A., Muntaner, O., LabbeJullie, C., Pitcher, J.A. & Marullo, S. (2006). Cooperative regulation of extracellular signal-regulated kinase activation and cell shape change by filamin A and β-arrestins. Mol Cell Biol 26, 3432-3445. [175] Ohta, Y., Hartwig, J. H. & Stossel, T. P. (2006). FilGAP, a Rho- and ROCK-regulated GAP for Rac binds filamin A to control actin remodelling. Nature Cell Biol 8, 803-814. [176] Awata, H, Huang, C, Handlogten, M. E. & Miller, R. T. (2001). Interaction of the calcium-sensing receptor and filamin, a potential scaffolding protein. J Biol Chem 276, 34871-34879. [177] Hjalm, G., MacLeod, R. J., Kifor, O., Chattopadhyay, N. & Brown, E. M. (2001). Filamin-A binds to the carboxyl-terminal tail of the calcium-sensing receptor, an interaction that participates in CaR-mediated activation of mitogen-activated protein kinase. Am Soc Biochem Mol Biol 276, 34880-34887. [178] Tigges, U., Koch, B., Wissing, J., Jockusch, B. M. & Ziegler, W. H. (2003). The Factin cross-linking and focal adhesion protein filamin A is a ligand and in vivo substrate for protein kinase Cα. J Biol Chem 278, 23561-23569. [179] Liu, G., Thomas, L., Warren, R. A., Enns, C. A., Cunningham, C. C., Hartwig, J. H. & Thomas, G. (1997). Cytoskeletal protein ABP-280 directs the intracellular trafficking of furin and modulates proproteins processing in the endocytic pathway. J Cell Biol 139, 1719-1733. [180] Cukier, I. H., Li, Y. & Lee, J. M. (2007). Cyclin B1/Cdk1 binds and phosphorylates Filamin A and regulates its ability to cross-link actin. FEBS Letters 581, 1661-1672. [181] Jay, D., Garcia, E. J., Lara, J. E., Medina, M. A. & de la Luz Ibarra, M. (2000). Determination of a cAMP-dependent protein kinase phosphorylation site in the Cterminal region of human endothelial actin-binding protein. Arch Biochem Biophys 377, 80-84. [182] Vadlamudi, R. K., Li, F., Adam, L., Nguyen, D., Ohta, Y., Stossel, T. P. & Kumar, R. (2002). Filamin is essential in actin cytoskeletal assembly mediated by p21-activated kinase. Nature Cell Biol 4, 681-690. [183] Ohta, Y. & Hartwig, J. H. (1995). Actin filament cross-linking by chicken gizzard filamin is regulated by phosphorylation in vitro. Biochem 34, 6745-6754. [184] Marti, A., Luo, Z., Cunningham, C., Ohta, Y., Hartwig, J., Stossel, T. P., Kyriakis, J. M. & Avruch, J. (1997). Actin-binding protein-280 binds the stress-activated protein kinase (SAPK) activator SEK-1 and is required for tumor necrosis factor-α activation of SAPK in melanoma cells. J Biol Chem 272, 2620-2628. [185] Woo, M. S., Ohta, Y., Rabinovitz, I., Stossel, T. P. & Blenis, J. (2004). Ribosomal S6 kinase (RSK) regulates phosphorylation of Filamin A on an important regulatory site. Mol Cell Biol 24, 3025-3035. [186] Zhu, T-N., He, H-J., Kole, S., D’Souza, T., Agarwal, R., Morin, P. J. & Bernier, M. (2007). Filamin A-mediated down-regulation of the exchange factor Ras-GRF1 correlates with decreased matrix metalloproteinase-9 expression in human melanoma cells. J Biol Chem 282, 14816-14826.

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ISBN: 978-1-60692-570-6 © 2009 Nova Science Publishers, Inc.

Chapter II

Systems Dynamics Behind Cell Movement Mikiya Otsuji1 and Shinya Kuroda2 1

Department of Anesthesiology, Faculty of Medicine, University of Tokyo, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, University of Tokyo, Tokyo, Japan

2

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Abstract Recent studies have shown that the interactions of various molecules are responsible for cell movement. Despite a consensus that systems of molecular interactions mediate the various dynamic properties of cells, it is still intuitively difficult to understand the ‘systems dynamics’ underlying cell movement. One way to visualize these dynamics is to use a systems-biological approach. The cause-result relationship between molecular interactions and cell movement can be understood as differential equations and their solution. Conceptual models, rather than detailed models, are often of greater use for extracting the essence of systems dynamics. In the first section, we review several mathematical conceptual models proposed for the analysis of cell polarity in migrating cells. The models are classified into four groups: the ‘LEGI + MSNL system’, the ‘CTSP system’, the ‘MCRD system’ and the ‘Meinhardt system.’ Chemotactic cells, even under normal conditions, do not always migrate with a stable front-back axis, but rather exhibit dynamic behaviors, depending on the conditions. In the second section, we show that MCRD systems composed of multiple species generate dynamic properties, which, in fact, can be observed in chemotaxing cells. The MCRD models with multiple species of molecules shed light on systems dynamics behind cell movement.

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Mikiya Otsuji and Shinya Kuroda

Introduction

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There have been a number of reports of molecules and their interactions that are involved in cell movements [1-18]. These molecules and their interactions, referred to as a signal transduction network, are shown schematically in the upper panel of Figure 1

Figure 1. The signal transduction network that regulates cell behavior. The upper panel is an overview of the signal transduction pathways involved in cell movements. The lower panel shows a Jurkat cell (human acute T-cell leukemia cell line) migrating towards a source of chemoattractant (SDF-1).

Although we know that cell behaviors are regulated by signal transduction networks such as these, it is difficult to visualize the dynamics of cell movement from such schematic representations of molecular interactions (Figure 1). One way to visualize such systems dynamics is to use a systems-biological approach, whereby the signal transduction network and the cell behavior are represented by the differential equations and the solutions of the equations, respectively. In particular, conceptual models that limit the number of variables and equations renders such analyses more simple and clear. In this chapter, we first classify the earlier mathematical models proposed to explain front-back polarities of migrating cells into four groups. Rather than presenting many detailed models, we make use of some

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Systems Dynamics Behind Cell Movement

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conceptual models that are representative of each of these groups. Second, we show that one of these four groups, the MCRD system, can exhibit not only stable polarity but also a characteristic dynamic behavior, both of which we can observe in living cells.

1. Conceptual Models for Stable Front-Back Axis of Migrating Cells Eukaryotic cells, such as neutrophils and Dictyostelium cells respond to slight gradients of extracellular signals with directional movement [19-21]. Such migrating cells usually exhibit a stable front-back axis, with various specific intracellular molecules localized at either the front or the back edge of the cell. This phenomenon is known as cell polarity. The lower panel of Figure 1 shows a migrating Jurkat cell, a leukocytic cell line, responding to a slight gradient of the chemoattractant, stromal cell–derived factor (SDF)-1. Many mathematical models that account for the front-back axis have been proposed. By describing the molecular interactions as a set of differential equations, the transitions of local concentrations of molecules can be obtained as the solutions. Some models consist of many variables and many equations. We can simplify these detailed models, keeping their core structures and main properties, into conceptual models. We classify these conceptual models into the following four groups:

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a) b) c) d)

LEGI (local excitation global inhibition) + MSNL (monostable nonlinearity) system, CTSP (classic Turing pattern with single peak) system, MCRD (mass conserved reaction diffusion) system, Meinhardt system.

Here, we introduce one or two conceptual models from each group. We consider onedimensional circular systems with circumference L: and we applied periodic boundary conditions. In each model, A, B, C and D denote the concentrations of molecules ‘A’, ‘B’, ‘C’ and ‘D’, respectively, at time t and at position x ( 0 ≤ x ≤ L ). kxxs (kA1, kA2, ..) denote the parameters of the systems, and Dxs (DA, DB, ..) represent the diffusion coefficients. S represents the extracellular stimulation (‘S’) derived from the equation S = S m {1 + S d cos[2π (x − xc ) / L]}, where Sm and Sd denote the average and gradient (0.005 – 0.05) of S, respectively, and xc denotes the highest point of stimulation. The initial conditions are homogenous states with small perturbations, unless specified otherwise.

(A) LEGI + MSNL System An easily understood model, called the ‘local excitation global inhibition (LEGI) model’, has been proposed to explain the sensing of stimulation gradients [22,23]. Because the model alone cannot explain the amplification of the signal, other mechanisms including positive

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feedbacks have been combined. The first of the four groups is the LEGI model combined with a monostable nonlinearity (MSNL) mechanism. There are several models that are classified within this group [24,25]. Figure 2 illustrates this system. The system can be divided into two parts: LEGI and MSNL. In the LEGI part, a stimulation ‘S’ activates molecules ‘A’ and ‘B’, which respectively activates and inhibit molecule ‘C’. Given that the diffusivity of ’B’ is much larger than ‘A’, the spatial profile of ‘C’ reflects that of ‘S’ without amplification.

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Figure 2. LEGI + MSNL system. Illustration of ‘LEGI + MSNL system.’ ‘A’, ‘B’, ‘C’, ‘D’ represent signaling molecules, and ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity. Arrow-headed and bar-headed lines indicate activation and inhibition, respectively.

In the MSNL part, the spatial gradient of ‘C’ is nonlinearly enhanced. As a result, the combined system can detect and amplify a slight gradient of stimulation. We introduce one conceptual model belonging to this system represented as follows: Model-a1

∂A ∂2 A = D A 2 + k A1 + k AS S − k A2 A , ∂t ∂x

(1)

∂B ∂2B = DB 2 + k B1 + k BS S − k B 2 B , ∂t ∂x

(2)

∂C ∂ 2C = DC + k CA A − k CB BC , ∂t ∂x 2

(3)

∂D ∂2D = DD + k DC C kn − k D 2 D . 2 ∂t ∂x

(4)

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The details of the parameter values are described in the APPENDIX. Application of the stimulations shown in the left top panel of Figure 3 generates the profiles of ‘C’ (left middle panel) and ‘D’ (left bottom panel). A high average of ‘S’ leads to high concentrations of ‘C’ and ‘D’. Here we can point out two things. First, despite the variations of the average of ‘S’, the averages of ‘C’ and ‘D’ are relatively constant. Second, the spatial gradients of ‘D’ are larger than those of ‘S’.

Figure 3. Simulation of the LEGI + MSNL system. Left panels show the results of simulations (Model-a1). The given profile of ‘S’ and the final profiles of ‘C’ and ‘D’ are indicated. Horizontal axis: position x. Right panels show the role of the LEGI part and the MSNL part.

The important role of the LEGI part, which converts the profile of ‘S’ into ‘C’, is to fix the average of ‘C’ to a certain value without a loss of gradient (right top panel of Figure 3). The MSNL part, which converts the profile of ‘C’ into ‘D’, nonlinearly amplifies the gradient of ‘C’, especially at the fixed value (right middle panel). Thus, the gradient of ‘S’, independently of its average (if not too small), gives ‘D’ a larger gradient with a relatively fixed average (right bottom panel). In the LEGI + MSNL system, ‘D’ faithfully reflects the spatial profile of ‘S’. For instance, when ‘S’ has two local maximums, ‘D’ also has two maximums. This system is reversible and responds to the directional change in stimulation (change in xc) (data not shown). In models that lack a molecule corresponding to ‘C’, the LEGI part and the MSNL part are inseparable. The MSNL part can be achieved by either a

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feedforward or feedback mechanism. Indeed, in one model, a high-order reaction represented by multiple LEGI parts combined in parallel works as an MSNL part [25], while in another model, multiple positive feedbacks toward substrates functions as an MSNL part [24]. These models share many features.

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(B) CTSP System Diffusion induced instability, presented by Turing in 1952 [26], is known to generate various dynamic and static spatial patterns. This instability is very attractive to one who considers intracellular spatial patterns including cell polarity. A classic Turing pattern in onedimensional space is stripes with an intrinsic scale length. This means that the numbers of stripes increases as the size of the system becomes larger. However, the number of stripes should be only one for a model of cell polarity, such as a front-back axis. There are at least two ways to achieve a single stripe: one is the CTSP (classic Turing pattern with single peak) system, where the system happens to have only one stripe for a given system size, and the other is the MCRD (mass conserved reaction diffusion) system, where the instability of the multiple-peak state leads to a one-peak state regardless of the system size. We categorize the former and the latter as the second and the third groups, respectively, and here we present the second group. There are some models belonging to this group, such as the mutual inhibition model [27] and a model introduced as the activator substrate – depletion model (ASDM) [28]. Consider a classic Turing model composed of two molecules ‘A’ and ‘B’. The diffusivity of ‘B’ is much larger than that of ‘A’. There are two well-known types of reaction terms, i.e. the activator – inhibitor system and the activator – substrate system. There are also several forms of positive feedback of activator ‘A’, i.e. the autocatalysis and mutual activation or inhibition with another molecule ‘C’. We can consider either types of reaction terms and either forms of positive feedback. Figure 4 illustrates this system. We introduce two conceptual models. Both are activator – substrate types, but one (Model-b1) includes positive feedback by autocatalysis of ‘A’ (left panel) [28] and the other (Model-b2) includes positive feedback by mutual inhibition between ‘A’ and ‘C’ (right panel) [27]. In Model-b1, where ‘A’ and ‘B’ generate a Turing pattern and ‘A’ unilaterally inhibits ‘C’, ‘C’ does not contribute to system behavior. These models are represented as follows: Model-b1

∂A ∂2 A = D A 2 + k AB + k AA SA 2 B − k A 2 A , ∂t ∂x

(5)

∂B ∂2B = DB 2 + k B1 − k BA A − k B 2 B , ∂t ∂x

(6)

(

)

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∂C ∂ 2C C = DC + k C1 − k CA A − kC 2C . 2 ∂t km + C ∂x

57

(7)

Figure 4. CTSP system. Illustration of the ‘CTSP system.’ ‘A’, ‘B’, ‘C’ represent signaling molecules, and ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity. Arrow-headed and bar-headed lines indicate activation and inhibition, respectively. Left and right panels correspond to Model-b1 and –b2, respectively.

Model-b2

∂A ∂2 A = D A 2 + k AB S − k AC AC 2 B − k A 2 A , ∂t ∂x

(8)

∂B ∂2B = DB 2 + k B1 − k BA A − k B 2 B , ∂t ∂x

(9)

∂C C ∂ 2C = DC + k C1 − k CA A − kC 2C . 2 ∂t km + C ∂x

(10)

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(

)

The details of parameter values are described in the APPENDIX. Introduction of the stimulations shown in the top panel of Figure 5, generate the profiles of ‘A’ and ‘C’ (Modelb1: middle panel, Model-b2: bottom panel of Figure 5). These models generate a Turing pattern with a single peak. As shown in Figure 6, they are reversible and respond to the directional change in stimulation (change in xc) (Model-b1: upper panel, Model-b2: lower panel of Figure 6). Here, two questions arise. (1) How severe is the condition for generating a single peak by chance? (2) While the Turing pattern is considered to be stable and less sensitive to changes in the parameters’ position-dependency [29], what determines the sensitivity of the CTSP system? First, to address the question (1), we consider the following reaction-diffusion system, which is composed of two components, ‘A’ and ‘B’:

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∂A ∂2 A = D A 2 + f A ( A, B ) , ∂t ∂x

(11)

∂B ∂2B = DB 2 + f B ( A, B ) , ∂t ∂x

(12)

where fA and fB are the reaction terms. The homogenous solution of this system, (A, B) = (A0, B0), is obtained as a solution of fA = fB = 0. In the homogenous state, the Jacobian matrix for the reaction terms is given by

J 12 ⎞ ⎛ ∂f A / ∂A ∂f A / ∂B ⎞ ⎟=⎜ ⎟ . J 22 ⎟⎠ ⎜⎝ ∂f B / ∂A ∂f B / ∂B ⎟⎠

⎛J J = ⎜⎜ 11 ⎝ J 21

(13)

When we defined the following:

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J *ij =

J ij

4π (DB − D A ) / L2 2

,

(14)

the number of peaks which arise from the initial homogenous state depends on J*11–J*22 and –J*12 J*21, as shown in Figure 7 (under DA/DB = 0.02). Note that the condition on J*11+J*22, which is required for instability, has little influence on the number of peaks. Figure 7 suggests that the condition for ‘homogenous or just one peak’ is less severe than those for others such as ‘just three peaks.’ Next, we consider the question (2). The Neumann boundary condition, which is often applied to reaction diffusion systems, clearly renders the pattern difficult to move. The periodic boundary condition is advantageous from this aspect. We consider the following reaction-diffusion system: Model-b3

∂A ∂2 A = D A 2 + k AA A 1 − A 2 S − k AB B , ∂t ∂x

(15)

∂B ∂2B = DB 2 + k BA ( A − A0 ) − k BB (B − B0 ) , ∂t ∂x

(16)

(

)

where (A0, B0) is the homogenous solution that satisfies kAA A0(1-A02) - kAB B0 = 0. As we can arbitrarily set A0, we investigated the case where A0 = 0 (left panels of Figure 8) and the case where A0 = 0.2 (right panels of Figure 8). The top panels of Figure 8 indicate the reaction terms of ‘A’ and ‘B’, i.e. kAA A (1-A2) - kAB B = 0 (solid lines), and kBA (A-A0) - kBB (B-B0) = 0 (dashed lines), respectively. The intersection points represent the homogenous solutions. For these analyses, first, we generate stable spatial profiles (Turing pattern).

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Figure 5. Simulation of the CTSP system. The results of simulations (Model-b1, -b2). The given profile of ‘S’ (upper panel) and the final profiles of ‘A’ and ‘C’ are indicated. The results of Model-b1 and –b2 are shown in the middle and the bottom panels, respectively. Horizontal axis: position x.

Figure 6. Simulation of the CTSP system. The results of simulations from the initial homogenous states (Model-b1, -b2). The given profile of ‘S’ are indicated at the top of the figure. The results of Model-b1 and – b2 are shown in the upper and the lower panels, respectively. Horizontal axis: time t, vertical axis: position x, color: A.

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Figure 7. Number of peaks in the Turing model. The number of peaks that arises from the initial homogenous state in the Turing model is determined by J*11–J*22 and –J*12 J*21.

Figure 8. Sensitivity of the CTSP system (influence of reaction terms). The sensitivity of Model-b3 to the directional change in stimulation (left: model with A0 = 0, right: model with A0 = 0.2). The top panels indicate the properties of reaction terms (solid lines: reaction terms of ‘A’, dashed line: reaction terms of ‘B’). We examine whether the Turing pattern can respond to the stimulation indicated in each panel. The middle panels represent the case where L = 6, and the bottom panels represents the case where L = 10.

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Then, by giving the stimulation shown in each panel of Figure 8, we obtained the profiles of ‘A’. Here, the middle panels represent a situation where L = 6, and the bottom panels represent a situation where L = 10. The peaks respond to ‘S’ only when A0 = 0.2 (right panels). These results suggest that the reaction terms affect the sensitivities of systems in some way. Next, we set A0 = 0.2 and L = 12, leading to the generation of stable profiles with three peaks (Figure 9). Then, we give stimulation with 1-5 local maxima as shown in each panel. Only in the case of stimulation with three local maxima, which corresponds to the number of peaks, do the peaks become sensitive and easy to move (Figure 9, middle left panel). Thus the peaks respond only when the number of maxima of ‘S’ is identical to the number of peaks. This result suggests that the one-peak state is easier to move than multiplepeak states. Thus, we can conclude that the condition for the one-peak state is not so severe and that the one-peak state is relatively easy to move.

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(C) MCRD System The uniqueness of the axis in the Turing pattern can also be achieved by the MCRD system, where the total quantity of some molecules is conserved. We can consider a reaction diffusion system composed of two molecules, ‘A’ and ‘B’, where the average of A+B in the space is constant. There are many models belonging to this group [28,30-37]. Periodic boundary conditions in one-dimensional space are applied to some models [31-33,35,36], and Neumann conditions in one-dimensional space to some [28], and two-dimensional space to others [34,37]. Despite the variations in equations or spatial conditions, these models have, at least partially, similar properties: they generate one distinct peak, which can respond to the directional change in stimulation. The most important point is that they exhibit the instability of the multiple-peak state, which eventually leads to a one-peak state. Figure 10 illustrates this system. ‘A’ and ‘B’ represent two states of one molecule whose total quantity in a cell is conserved. There is a positive feedback through which an increase of ‘A’ accelerates the conversion of ‘B’ to ‘A’. The diffusivity of ‘B’ is much larger than that of ‘A’. We introduce two conceptual models given as follows: Model-c1

∂A ∂2 A = D A 2 + k1 S A 2 + k a B − k i A , ∂t ∂x

(17)

∂B ∂2B = D B 2 − k1 S A 2 + k a B + k i A , ∂t ∂x

(18)

(

(

)

)

Model-c2

∂A ∂2 A = D A 2 + k1 S A 2 + k a B − (k i + c m ( A))A , ∂t ∂x

(

)

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Figure 9. Sensitivity of the CTSP system (influence of stimulation profile). The sensitivity of Model-b3 to a directional change in stimulation. We examined whether the Turing pattern can respond to the stimulation indicated in each panel. The number of maxima of the stimulation profiles, n, vary from one to five as indicated in the panels.

Figure 10. MCRD system.Illustration of the ‘MCRD system.’ ‘A’ and ‘B’ represent two states of one signaling molecule: the total quantity of ‘A’ and ‘B’ in the whole space is conserved. ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity.

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∂B ∂2B = DB 2 − k1 S A 2 + k a B + (k i + c m ( A))A , ∂t ∂x

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(

)

63

(20)

where the function cm(A) prevents ‘A’ from an excessive increase (a certain degree of increase in ‘A’ prompts the conversion of ‘A’ to ‘B’). The details of the parameter values and cm(A) are described in the APPENDIX. The stimulations shown in Figure 11 generate the profiles of ‘A’ (Model-c1: upper panel of Figure 11, Model-c2: lower panel). Multiple peaks emerge at an early phase, but only one persists, resulting in a stable one-peak state, in each model. The MCRD system, unlike the CTSP system, generates a one-peak state regardless of the system size. The final one-peak states are indicated in the upper panels of Figure 12 (Model-c1: left upper panel, Model-c2: right upper panel). The shapes of the peaks depend upon the reaction terms, e.g. with or without the upper limit of ‘A’, cm(A). The peaks can respond to the directional change in stimulation (change in xc) (Model-c1: left lower panel, Model-c2: right lower panel of Figure 12). The MCRD system with two components, ‘A’ and ‘B’, appears to be one of the simplest models for cell polarity. The models described above are assumed to consist of one species (i.e. two components because the species has two states), so that we refer to such models as single-species (SS)MCRD systems. Here, we can expand these conceptual models to multiple-species (MS)MCRD systems, e.g. a 2S-MCRD system composed of two species (four components). Because a ‘behavior’ is a sequence of states (e.g. homogenous state, one-peak state) and because the destiny of each state is determined by its ‘stability’ (e.g. stable, collapse of the peak), analyzing the stability of the various states is useful for predicting the system’s behaviors. The MCRD system has several instabilities, including a Turing instability that destabilizes the homogenous state. We briefly introduce three remarkable instabilities of nonhomogenous states (Figure 13). The first is the ‘instability of the multiple-peak state’ (Instability I) (top panel of Figure 13). This instability can be observed even in the SSMCRD system as shown above. When two peaks exist, one of them becomes larger and the other becomes smaller. The second is an ‘instability that deforms (split) the peak’ (Instability II) (middle panel of Figure 13). This instability can be observed only in the MS-MCRD system, depending upon the reaction terms (interactions between species are important). When some conditions are satisfied, the peak splits into two. The third is ‘instability that leads to spontaneous movement of the peak’ (Instability III) (bottom panel of Figure 13). This instability can also be observed only in the MS-MCRD system, depending upon the reaction terms. When some conditions are satisfied, the peak spontaneously starts to move laterally as shown in the bottom panel of Figure 13. The combinations of these instabilities can generate various dynamic behaviors, which are described in the next section.

(D) Meinhardt System The model proposed by Meinhardt in 1999 for front-back polarity of migrating cells is so interesting that we define the fourth group for this model, although there are few models belonging to this group. This system responds to extracellular stimulations by producing hot spots in a dynamic way.

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Figure 11. Simulation of the MCRD system.The results of simulations from the initial homogenous states (Model-c1, -c2). The given profiles of ‘S’ are indicated at the top of the figure. The results of Model-c1 and –c2 are shown in the upper and the lower panels, respectively. Horizontal axis: time t, vertical axis: position x, color: A.

Figure 12. Simulation of the MCRD system. The results of simulations (left: Model-c1, right: Model-c2). The given profiles of ‘S’ are indicated in each panel. The upper panels show the final stable profiles of ‘A’, and the lower panels show their responses to a directional change in stimulations. Horizontal axis: position x.

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Figure 13. Three instabilities of the MCRD system. Three instabilities of non-homogenous solutions. Instability I: instability of the multiple-peak solution. Instability II: instability that splits the peak. Instability III: instability that leads to spontaneous movement of the peak.

Figure 14 illustrates this system. Here, we introduce two conceptual models. They share the same equations given as follows, but they have different parameter values: Model-d1, -d2

(

)

A 2 / B + k AB S ∂A ∂2 A = DA 2 + −k A , ∂t (k AC + C ) 1 + k AA A 2 A2 ∂x

(21)

∂B ∂2B = DB 2 + k BA A − k B 2 B , ∂t ∂x

(22)

∂C ∂ 2C = DC + k CA A − k C 2 B , ∂t ∂x 2

(23)

(

)

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The details of the parameter values are described in the APPENDIX. The stimulations shown at the top of Figure 15, generates the profiles of ‘A’. The upper and lower panels of Figure 15 show the profiles of Model-d1 and –d2, respectively. While Model-d1 generates waves of multiple transient spots, Modeld-d2 generates slowly moving patterns. Although these models exhibit apparently different dynamic patterns, which are caused by different values of one parameter, kC2, both models are reversible and respond to a directional change in stimulation. As the leading edges of migrating cells have been suggested to be dynamic rather than static [15], this Meinhardt system deserves further study, including from the standpoint of the molecular interpretations.

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Figure 14. Meinhardt system.Illustration of the ‘Meinhardt system.’ ‘A’, ‘B’, ‘C’ represent signaling molecules, and ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity. Arrow-headed and barheaded lines indicate activation and inhibition, respectively.

Figure 15. Simulation of the Meinhardt system. The results of simulations from the initial homogenous states (Model-d1, -d2). The given profiles of ‘S’ are indicated at the top of the figure. The results of Model-d1 and –d2 are shown in the upper and the lower panels, respectively. Horizontal axis: time t, vertical axis: position x, color: A.

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Distinctions among the Groups One may wonder whether the four groups are mutually exclusive. Here, we consider some individual cases.

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1. Because only the ‘LEGI + MSNL system’ does not depend on any instability, this system can be distinguished from the others. The ‘LEGI + MSNL system’ responds tightly to the profiles of stimulation. When homogenous stimulations are given, the LEGI + MSNL system can only respond homogenously, while the other systems can generate, although randomly, distinct spatial profiles that are induced by their instability. The role of the LEGI part is essential for the system where LEGI is combined with the MSNL mechanism, but is limited for the system where it is combined with other mechanisms depending on instabilities. For instance, properties of a system where LEGI is combined with the MCRD system originate largely from those of the MCRD system. 2. Although both the CTSP system and the MCRD system are based on the Turing pattern, they achieve the uniqueness of axis by different mechanisms and thus have different features. For instance, while the one-peak state of the CTSP system depends upon the system size, that of the MCRD system does not. But having stated this, these two models can be compatible. In such a system, one peak emerges from the initial homogenous state, and most properties of the MCRD system might be inherited. 3. Because only the Meinhardt system exhibits polarity in a dynamic manner, this system can be distinguished from the other systems. Note that other systems, such as the MS-MCRD system, can generate various dynamic behaviors. In the future, there may be a need for further groups.

Variables in Models and Corresponding Molecules Although each variable in the models is assumed to represent the local concentration of one molecule, it is not always feasible to establish correspondence between variables and molecules. In the case of the MCRD system, it seems relatively easy to find candidate molecules, i.e. molecule that exhibit two states with distinct diffusivities, accompanied by positive feedback. For example, the Rho family of small GTPases, including RhoA, Rac, Cdc42, exist in two states, i.e. a GTP-bound form on the membrane and a GDP-bound form in the cytosol [38], and Rac and phosphatidilinositol 3,4,5-triphosphate constitute a positive feedback loop [1,39] (Figure 16). In addition, Rho GTPases have also been shown to play a central role in cell polarity and movements through their direct regulation of actin polymerization [18,38]. Thus, Rho GTPases seems to be compatible with the MCRD system. In a migrating cell, many molecules on the membrane and in the cytosol are involved in the signal transduction network. Because conceptual models represent the core structures of such networks in extremely simplified forms, each variable implies abstraction of one or more pathways rather than one specific molecule. This leads to the following considerations:

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Mikiya Otsuji and Shinya Kuroda 1. ‘Local excitation and global inhibition’ does not necessarily require the large diffusivity of a molecule that directly inhibits some other molecules. In the case of Figure 2, we can consider the situation where ‘B’, which exhibits small diffusivity, is activated through a cytosolic second messenger that exhibits greater diffusivity. 2. The LEGI + MSNL system does not necessarily require the large diffusivity of the inhibitory molecule. In the case of Figure 2, we can consider the situation where the excitatory molecule, ‘A’, exhibits a large diffusivity and the inhibitory molecule, ‘B’, exhibits a small diffusivity. In such a situation, the inhibition of ‘D’ by ‘C’ leads to the same relationship between ‘S’ and ‘D’ as Figure 3. 3. That ‘A’ activates ‘B’ does not necessarily imply that ‘A’ directly activates some other molecules. We can consider the situation where ‘A’ inhibits another molecule, ‘E’, which inhibits ‘B’. 4. The MCRD system can consist of molecules that form dimers. In this case, the molecule exists as monomers, ‘A’, or dimers, ‘B’, with distinct diffusivities, and the spatial average of A+2B is conserved.

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These considerations mean that ‘activate/inhibit’ and diffusivity in the conceptual model do not indicate the specific activity and diffusivity of the corresponding molecule, respectively, and that permissive extended interpretations may facilitate applying concrete molecules or pathways.

Figure 16. Rho-GTPases.Rho GTPases cycle between an active GTP-bound form and an inactive GDPbound form. This cycle is regulated by guanine exchange factors (GEF), GTPase activating proteins (GAP) and guanine dissociation inhibitors (GDI). In the active state, they interact with effector molecules, through which they can activate or inactivate other molecules.

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2. Dynamic Behaviors in MCRD System A Characteristic Dynamic Behavior: ‘Split and Choice’ Dictyostelium cells in a shallow gradient of chemoattractant (cAMP) exhibit a characteristic behavior, i.e. repeated split of a leading edge of a migrating cell into two and the choice of one of the two [40] (‘split and choice’). Figure 17 shows Jurkat cells within a gradient of the chemoattractant SDF-1. The cell, migrating upward, exhibits this ‘split and choice’ behavior: it chooses the left pseudopod after the first split and then the right pseudopod after the second split. Note that cells migrating in a straight line with a stable front-back axis (lower panel of Figure 1) and cells exhibiting the ‘split and choice’ behavior (Figure 17) co-exist under the same conditions. Thus, the ‘split and choice’ behavior may be a key criterion for evaluating models that are proposed in an attempt to explain the behaviors of migrating cells. ‘Split and choice’ behavior can be reproduced by at least two types of MCRD system. Here, we introduce two conceptual models, corresponding to these two systems. One is a SSMCRD system with oscillator, where Turing instability, Instability I (instability of multiple peak), and reversibility are involved in the generation of ‘split and choice’ (Model-e1, Figure 18). The other is a 2S-MCRD system, where Instability I and Instability II are involved in the emergence of spontaneous ‘split and choice’ behavior (Model-e2, Figure 21). Model-e1

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⎛ k os (t ) + k 2 ∂ 2 Am ∂Am k4 ⎜ + + − k SA = Dm 1 c 2 2 2 ⎜ ∂t ∂x 1 + k 5 Am ⎝ 1 + k 3 Am ⎛ k (t ) + k 2 ∂ 2 Ac ∂Ac k4 + − k1 SAc + ⎜⎜ os = Dc 2 2 2 ∂t ∂x 1 + k 5 Am ⎝ 1 + k 3 Am

⎞ ⎟ Am ⎟ ⎠

⎞ ⎟ Am ⎟ ⎠

,

(24)

,

(25)

where kos(t) is a function of t and acts as an oscillator, given by

k os (t ) = k 0 [1 − cos(2πt / k T )] .

(26)

Model-e2

(

)

∂ 2 Am ∂Am 2 + k1 A SAm + k 2 A + k AB Bm Ac − (k 3 A + c mA ( Am ))Am = Dm 2 ∂t ∂x

(

)

∂ 2 Ac ∂Ac 2 − k1 A SAm + k 2 A + k AB Bm Ac + (k 3 A + c mA ( Am ))Am = Dc 2 ∂t ∂x

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, (27)

,

(28)

Mikiya Otsuji and Shinya Kuroda

70

(

)

∂ 2 Bm ∂Bm 2 + k1B Bm + k 2 B Bc − (k 3 B + k BA Am + c mB (Bm ))Bm = Dm ∂t ∂x 2

(

)

∂ 2 Bc ∂Bc 2 − k1B Bm + k 2 B Bc + (k 3 B + k BA Am + c mB (Bm ))Bm = Dc ∂t ∂x 2

,

,

(29)

(30)

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Figure 17. Characteristic behavior of chemotaxing cell.A Jurkat cell (human acute T-cell leukemia cell line) migrating towards a source of chemoattractant (SDF-1).

Figure 18. SS-MCRD system with oscillator. Illustration of the ‘SS-MCRD system with an oscillator.’ ‘Am’ and ‘Ac’ represent two states of one signaling molecule. ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity.

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The details of the parameter values are described in the APPENDIX. Figure 19 shows ‘split and choice’ generated by Model-e1. The upper panel indicates the time course of the oscillator, which is exogenously given. The lower panel indicates the profiles of Am. We can see the repetitions of a sequence of split of a peak into two and choice of one of these peaks. The spatial profiles of Am at six points (six arrows in Figure 19) are shown in Figure 20. From the semi-homogenous state (a), two peaks emerge by Turing instability in association with an increase in kos (b,c). However, one of the peaks eventually disappears, and only one peak remains, which is caused by Instability I (d,e). The one-peak state returns to a semihomogenous state in association with a decrease in kos (f). Figure 22 indicates ‘split and choice’ generated by Model-e2. Similarly to Figure 19, we can see the repetitions of a sequence of splits and choices of peak. However, this behavior occurs spontaneously without any exogenous pace maker. The spatial profiles are shown in Figure 23. One peak (a) splits into two by Instability II (b,c), but only one peak is chosen by Instability I (d,e,f). Model-e1 requires an oscillator that is not required to be localized. Intracellular oscillations have been studied extensively [41]. Because the inhibition of phosphoinositide 3kinase (PI3K) in Dictyostelium cells, which exhibit ‘split and choice’ in a shallow gradient of chemoattractant, have been reported not to impede this process, but to slow the rate [40], PI3K might be involved in such an oscillation mechanism. Model-e2 requires interactions between multiple species. Rho GTPases have been shown to interact indirectly with one another, in addition to participating in positive feedback loops (upper panel of Figure 1, Figure 15). In the upper panel of Figure 1, we can find a similar structure to Figure 21, when we consider that ‘A’ and ‘B’ correspond to Rac and RhoA, respectively.

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Other Dynamic Behaviors Chemotactic cells, when they are treated with inhibitors of signaling molecules, exhibit various dynamic behaviors [39,42,43]. It should be noted that the MS-MCRD system can generate various dynamic behaviors, depending on the parameter values. The upper panel of Figure 24 indicates the profiles of Am generated by Model-e3, which is obtained by varying the parameter values of Model-e2 (details are described in APPENDIX). A split of a peak by Instability II occurs at an early phase, but once a peak starts to move by Instability III, a traveling wave persists. The lower panel of Figure 24 indicates the profiles generated by Model-e4, which is also obtained by varying the parameter values of Model-e2. In this model, the instabilities, such as Instability I, II and III, generate a more complex dynamic pattern. In addition to these behaviors, many other behaviors, which cannot always be explained by Instability I-III, are observed in the MS-MCRD system (data not shown). These may correspond to the dynamic behaviors seen in the inhibitor-treated cells.

Conclusion Many mathematical models have been proposed to associate the overview of a signal transduction network with the observable behaviors of chemotactic cells. We have classified

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the existing models for front-back polarity into four groups, termed the ‘LEGI + MSNL system’, the ‘CTSP system’, the ‘MCRD system’ and the ‘Meinhardt system’, and have introduced several conceptual models for each group. These models comprise no more than four components. They are sufficiently simple to allow examination of their mathematics in detail. Chemotactic cells exhibit various dynamic behaviors that depend at least in part on the conditions, such as the gradient of chemoattractant and the presence of inhibitors. ‘Split and choice’ is a characteristic behavior that can be observed even in normal cells. Importantly, this behavior is reproduced by two types of the MCRD system. Although the mechanisms that generate this behavior are different, the ‘instability of the multiple-peak state’ plays a key role in both systems. Other dynamic behaviors can be observed in inhibitor-treated cells, and also in the MCRD system. Taken together, MCRD systems have good potential as a central mechanism of cell movement in the broader sense.

Figure 19. ‘Split and choice’ by ‘SS-MCRD system with oscillator. The results of simulations from the initial homogenous states (Model-e1). The upper panel indicates the time course of the oscillator function, which is exogenously given. The result is shown in the lower panel. Horizontal axis: time t, vertical axis: position x, color: Am.

Figure 20. ‘Split and choice’ by SS-MCRD system with oscillator.The spatial profiles of ‘Am’ at six points in the lower panel of Figure 19 (six arrows). Horizontal axis: position x.

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Figure 21. 2S-MCRD system.Illustration of the ‘2S-MCRD system.’ ‘Am’, ‘Ac’ and ‘Bm’, ‘Bc’ represent two states of two signaling molecules. ‘S’ denotes stimulation. The suffix ‘D’ implies large diffusivity.

Figure 22. ‘Split and choice’ by 2S-MCRD system.The results of simulations from the initial homogenous states (Model-e2). Horizontal axis: time t, vertical axis: position x, color: Am.

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Figure 23. ‘Split and choice’ by 2S-MCRD system.The spatial profiles of ‘Am’ at six points in Figure 22 (six arrows). Horizontal axis: position x.

Figure 24. Dynamic behaviors by the 2S-MCRD system. The results of simulations from the initial homogenous states (Model-e3, -e4). The results of Model-e3 and –e4 are shown in the upper and the lower panels, respectively. Horizontal axis: time t, vertical axis: position x, color: Am.

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Appendix Parameter values of models used in simulations. Model-a1 kA1 = 0.02, kAS = 0.3, kA2 = 0.3, kB1 = 0.01, kBS = 0.1, kB2 = 0.1, kCA = 0.5, kCB = 0.5, kDC = 1, kD2 = 1, kn = 5, DA = DC = DD = 0.001, DB = 1, L = 10, initial (A, B, C, D) = (0.5, 0.5, 0.5, 0.5). Model-b1 kAB = 0.05, kAA = 0.1, kA2 = 0.15, kB1 = 0.2, kBA = 0.1, kB2 = 0.1, kC1 = 0.4, kCA = 0.3, km = 1, kC2 = 0.1, DA = 0.02, DB = 1, DC = 0.02, L = 10, initial (A, B, C) = (0.5, 0.5, 0.5). Model-b2 kAB = 0.8, kAC = 0.15, kA2 = 0.25, kB1 = 0.4, kBA = 0.1, kB2 = 0.2, kC1 = 0.3, kCA = 0.1, km = 0.01, kC2 = 0.1, DA = 0.02, DB = 1, DC = 0.02, L = 10, initial (A, B, C) = (0.5, 0.5, 0.5). Model-b3 kAA = 0.2, kAB = 0.2, kBA = 0.6, kBB = 0.4, DA = 0.02, DB = 1, Sm = 1.

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Model-c1 k1 = 2, ka = 0.3, ki = 0.5, DA = 0.02, DB = 1, L = 10, initial (A, B) = (0.5, 0.5). Model-c2 k1 = 2, ka = 0.3, ki = 0.5, DA = 0.02, DB = 1, L = 10, initial (A, B) = (0.5, 0.5).

cm ( A ) =

30 1 + e[− 6( A − 3 )]

.

Model-d1 kAB = 0.1, kAC = 0.2, kAA = 0.001, kA2 = 0.06, kBA = 0.1, kB2 = 0.1, kCA = 0.05, kC2 = 0.02, DA = 0.001, DB = 1, DC = 0.001, L = 10, initial (A, B, C) = (0.5, 0.5, 0.5). Model-d2 kAB = 0.1, kAC = 0.2, kAA = 0.001, kA2 = 0.06, kBA = 0.1, kB2 = 0.1, kCA = 0.05, kC2 = 0.1, DA = 0.001, DB = 1, DC = 0.001, L = 10, initial (A, B, C) = (0.5, 0.5, 0.5).

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76 Model-e1

k1 = 10, k2 = 1, k3 = 20, k4 = 10, k5 = 10, k0 = 15, kT = 800, Dm = 0.01, Dc = 2, L = 10, Sm = 1, xc = 5, initial (Am, Ac) = (0.5, 0.5). Model-e2 k1A = 4, k2A = 0.2, k3A = 1.2, k1B = 4, k2B = 0.1, k3B = 0.3, kAB = 0.8, kBA = 0.3, Dm = 0.06, Dc = 2, L = 10, Sm = 1, xc = 5, initial (Am, Ac, Bm, Bc) = (0.5, 0.5, 0.5, 0.5).

c mA ( Am ) =

30

1+ e

[−6 ( Am − 2 )]

, c mB (Bm ) =

30

1+ e

[− 6 ( Bm −1.5 )]

.

Model-e3 k1A = 4, k2A = 0.2, k3A = 1.5, k1B = 4, k2B = 0.2, k3B = 0.5, kAB = 1, kBA = 0.2, Dm = 0.06, Dc = 2, L = 10, Sm = 1, xc = 5, initial (Am, Ac, Bm, Bc) = (0.5, 0.5, 0.5, 0.5).

c mA ( Am ) =

30

1+ e

[−6 ( Am − 2 )]

, c mB (Bm ) =

30

1+ e

[− 6 ( Bm −1.5 )]

.

Model-e4 k1A = 4, k2A = 0.2, k3A = 1.2, k1B = 4, k2B = 0.1, k3B = 0.3, kAB = 0.8, kBA = 0.4, Dm = 0.06, Dc = 2, L = 10, Sm = 1, xc = 5, initial (Am, Ac, Bm, Bc) = (0.5, 0.5, 0.5, 0.5).

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c mA ( Am ) =

30

1+ e

[−6 ( Am − 2 )]

, c mB (Bm ) =

30

1+ e

[− 6 ( Bm −1.5 )]

.

References [1]

[2]

[3]

[4]

[5]

Weiner OD, Neilsen PO, Prestwich GD, Kirschner MW, Cantley LC, et al. (2002) A PtdInsP(3)- and Rho GTPase-mediated positive feedback loop regulates neutrophil polarity. Nat. Cell Biol. 4: 509-513. Welch HCE, Coadwell WJ, Ellson CD, Ferguson GJ, Andrews SR, et al. (2002) PRex1, a PtdIns(3,4,5)P3- and Gbetagamma-regulated guanine-nucleotide exchange factor for Rac. Cell 108: 809-821. Kunisaki Y, Nishikimi A, Tanaka Y, Takii R, Noda M, et al. (2006) DOCK2 is a Rac activator that regulates motility and polarity during neutrophil chemotaxis. J. Cell Biol. 174: 647-652. Vicente-Manzanares M, Cruz-Adalia A, Martin-Cofreces NB, Cabrero JR, Dosil M, et al. (2005) Control of lymphocyte shape and the chemotactic response by the GTP exchange factor Vav. Blood 105: 3026-3034. Wong K, Van Keymeulen A, Bourne HR (2007) PDZRhoGEF and myosin II localize RhoA activity to the back of polarizing neutrophil-like cells. J. Cell Biol. 179: 11411148.

Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

Systems Dynamics Behind Cell Movement [6]

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[14]

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Kitzing TM, Sahadevan AS, Brandt DT, Knieling H, Hannemann S, et al. (2007) Positive feedback between Dia1, LARG, and RhoA regulates cell morphology and invasion. Genes Dev. 21: 1478-1483. Chikumi H, Fukuhara S, Gutkind JS (2002) Regulation of G protein-linked guanine nucleotide exchange factors for Rho, PDZ-RhoGEF, and LARG by tyrosine phosphorylation: evidence of a role for focal adhesion kinase. J. Biol. Chem. 277: 12463-12473. Francis SA, Shen X, Young JB, Kaul P, Lerner DJ (2006) Rho GEF Lsc is required for normal polarization, migration, and adhesion of formyl-peptide-stimulated neutrophils. Blood 107: 1627-1635. Nimnual AS, Taylor LJ, Bar-Sagi D (2003) Redox-dependent downregulation of Rho by Rac. Nat. Cell Biol. 5: 236-241. Ohta Y, Hartwig JH, Stossel TP (2006) FilGAP, a Rho- and ROCK-regulated GAP for Rac binds filamin A to control actin remodelling. Nat. Cell Biol. 8: 803-814. Tsuji T, Ishizaki T, Okamoto M, Higashida C, Kimura K, et al. (2002) ROCK and mDia1 antagonize in Rho-dependent Rac activation in Swiss 3T3 fibroblasts. J. Cell Biol. 157: 819-830. Li Z, Hannigan M, Mo Z, Liu B, Lu W, et al. (2003) Directional sensing requires G beta gamma-mediated PAK1 and PIX alpha-dependent activation of Cdc42. Cell 114: 215-227. Nishimura T, Yamaguchi T, Kato K, Yoshizawa M, Nabeshima Y, et al. (2005) PAR-6PAR-3 mediates Cdc42-induced Rac activation through the Rac GEFs STEF/Tiam1. Nat. Cell Biol. 7: 270-277. Gerard A, Mertens AE, van der Kammen RA, Collard JG (2007) The Par polarity complex regulates Rap1- and chemokine-induced T cell polarization. J. Cell Biol. 176: 863-875. Nakayama M, Goto TM, Sugimoto M, Nishimura T, Shinagawa T, et al. (2008) Rhokinase phosphorylates PAR-3 and disrupts PAR complex formation. Dev. Cell 14: 205215. Li Z, Dong X, Wang Z, Liu W, Deng N, et al. (2005) Regulation of PTEN by Rho small GTPases. Nat. Cell Biol. 7: 399-404. Stephens L, Ellson C, Hawkins P (2002) Roles of PI3Ks in leukocyte chemotaxis and phagocytosis. Curr. Opin. Cell Biol. 14: 203-213. Ridley AJ (2001) Rho GTPases and cell migration. J. Cell Sci. 114: 2713-2722. Chung CY, Funamoto S, Firtel RA (2001) Signaling pathways controlling cell polarity and chemotaxis. Trends Biochem. Sci. 26: 557-566. Iijima M, Huang YE, Devreotes P (2002) Temporal and spatial regulation of chemotaxis. Dev. Cell 3: 469-478. Affolter M, Weijer CJ (2005) Signaling to cytoskeletal dynamics during chemotaxis. Dev. Cell 9: 19-34. Kutscher B, Devreotes P, Iglesias PA (2004) Local excitation, global inhibition mechanism for gradient sensing: an interactive applet. Sci STKE 2004: pl3. Parent CA, Devreotes PN (1999) A cell's sense of direction. Science 284: 765-770.

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[24] Levchenko A, Iglesias PA (2002) Models of eukaryotic gradient sensing: application to chemotaxis of amoebae and neutrophils. Biophys. J. 82: 50-63. [25] Ma L, Janetopoulos C, Yang L, Devreotes PN, Iglesias PA (2004) Two complementary, local excitation, global inhibition mechanisms acting in parallel can explain the chemoattractant-induced regulation of PI(3,4,5)P3 response in Dictyostelium cells. Biophys. J. 87: 3764-3774. [26] Turing AM (1952) The chemical basis of morphogenesis. Philos. Trans. R. Soc. Lond B 237: 37-72. [27] Narang A (2006) Spontaneous polarization in eukaryotic gradient sensing: a mathematical model based on mutual inhibition of frontness and backness pathways. J. Theor. Biol. 240: 538-553. [28] Mori Y, Jilkine A, Edelstein-Keshet L (2008) Wave-pinning and cell polarity from a bistable reaction-diffusion system. Biophys. J. 94: 3684-3697. [29] Meinhardt H (1999) Orientation of chemotactic cells and growth cones: models and mechanisms. J. Cell Sci. 112: 2867-2874. [30] Postma M, Van Haastert PJ (2001) A diffusion-translocation model for gradient sensing by chemotactic cells. Biophys. J 81: 1314-1323. [31] Narang A, Subramanian KK, Lauffenburger DA (2001) A mathematical model for chemoattractant gradient sensing based on receptor-regulated membrane phospholipid signaling dynamics. Ann. Biomed. Eng. 29: 677-691. [32] Subramanian KK, Narang A (2004) A mechanistic model for eukaryotic gradient sensing: spontaneous and induced phosphoinositide polarization. J. Theor. Biol. 231: 49-67. [33] Skupsky R, Losert W, Nossal RJ (2005) Distinguishing modes of eukaryotic gradient sensing. Biophys. J. 89: 2806-2823. [34] Gamba A, de Candia A, Di Talia S, Coniglio A, Bussolino F, et al. (2005) Diffusionlimited phase separation in eukaryotic chemotaxis. Proc. Natl. Acad. Sci. U S A 102: 16927-16932. [35] Onsum M, Rao CV (2007) A mathematical model for neutrophil gradient sensing and polarization. PLoS Comput Biol. 3: e36. [36] Otsuji M, Ishihara S, Co C, Kaibuchi K, Mochizuki A, et al. (2007) A mass conserved reaction-diffusion system captures properties of cell polarity. PLoS Comput. Biol. 3: e108. [37] Goryachev AB, Pokhilko AV (2008) Dynamics of Cdc42 network embodies a Turingtype mechanism of yeast cell polarity. FEBS Lett 582: 1437-1443. [38] Etienne-Manneville S, Hall A (2002) Rho GTPases in cell biology. Nature 420: 629635. [39] Srinivasan S, Wang F, Glavas S, Ott A, Hofmann F, et al. (2003) Rac and Cdc42 play distinct roles in regulating PI(3,4,5)P3 and polarity during neutrophil chemotaxis. J. Cell Biol. 160: 375-385. [40] Andrew N, Insall RH (2007) Chemotaxis in shallow gradients is mediated independently of PtdIns 3-kinase by biased choices between random protrusions. Nat. Cell Biol. 9: 193-200.

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[41] Kholodenko BN (2006) Cell-signalling dynamics in time and space. Nat. Rev. Mol. Cell Biol. 7: 165-176. [42] Xu J, Wang F, Van Keymeulen A, Herzmark P, Straight A, et al. (2003) Divergent signals and cytoskeletal assemblies regulate self-organizing polarity in neutrophils. Cell 114: 201-214. [43] Van Keymeulen A, Wong K, Knight ZA, Govaerts C, Hahn KM, et al. (2006) To stabilize neutrophil polarity, PIP3 and Cdc42 augment RhoA activity at the back as well as signals at the front. J. Cell Biol. 174: 437-445.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 81-121

ISBN: 978-1-60692-570-6 © 2009 Nova Science Publishers, Inc.

Chapter III

Mast Cells in Injury Response Stefano Bacci*, Aurelio Bonelli and Paolo Romagnoli Department of Anatomy, Histology and Forensic Medicine; University of Florence, Italy

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Abstract Mast cells are bone marrow derived cells capable of secreting many active molecules, ranging from the mediators stored in specific granules, some of which are known since several decades (histamine, heparin), to small molecules produced immediately upon stimulation (membrane lipid derivatives, nitric oxide), to a host of constitutively secreted, multifunctional cytokines. With the aid of a wide array of mediators the activated mast cells control the key events of inflammation and healing and participate to the regulation of local immune response. On the basis of the structure, origin, principal subtypes, localization and function of these cells, their involvement in injury repair is therefore to be considered in acute and chronic conditions respectively. The importance of mast cells in regulating healing processes is underscored by the proposed roles of a surplus or a deficit of their mediators in the formation of exuberant granulation tissue (such as keloids and hypertrophic scars), the delayed closure or dehiscence of wounds and the transition of acute to chronic inflammation.

Abbreviations Alpha chemokine receptor beta chemokine receptor endothelin

CXCR CCR ET

* Correspondence to: Dr. Stefano Bacci, Department of Anatomy, Histology and Forensic Medicine, Section “E. Allara”, Viale Pieraccini 6, 50134 Florence, Italy, Tel: xx-39-55-4271389, Fax: xx-39-55-4271385, mail: [email protected] Cell Movement : New Research Trends, Nova Science Publishers, Incorporated, 2008. ProQuest Ebook Central,

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Stefano Bacci, Aurelio Bonelli and Paolo Romagnoli enterochromaffin-like fibroblast growth factor glycosaminoglycan granulocyte monocyte-colony stimulatory function high-affinity IgG receptor interferon interleukin inducible NOS immunoglobulin high affinity receptor leukotrienes macrophage inflammation protein mast cells mast cells containing only tryptase mast cells containing both tryptase and chymase mast cells containing only chymase matrix metalloproteinase melanocortin receptor monocyte chemoattractant protein nerve growth factor neuronal nitric oxide synthase neurokinin receptor nitric oxide peroxisome proliferator activated receptor prostaglandin E receptor protease activated receptor platelet activating factor platelet-derived endothelial cell growth factor plateled derived growth factor prostaglandin peroxisome proliferator activated receptor regulated on activation, normal T cell expressed and secreted molecule smooth muscle cells of the synthetic phenotype stem cell factor TNF-related apoptosis inducing ligand Toll-like receptor transforming growth factor tumour necrosis factor vascular cell adhesion molecule vascular endothelial growth factor very late (activation) antigen vaso active intestinal polypeptide VIP receptor-2

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ECL FGF GAG GM-CSF FcγRI IFN IL iNOS FcεRI LT MIP MC MCT MCTC MCC MMP MCR MCP NGF nNOs NKR NO PPAR EP PARs PAF PDEGF PDGF PGD PPAR RANTES s-SMCs SCF TRAIL TLR TGF TNF VCAM VEGF VLA VIP VPAC2

Mast Cells in Injury Response

83

Introduction Mast cells (MC) had long been elusive as to their functional role. The name itself tells that they were first interpreted as nutrient storing cells, before being recognized as secretory cells. Later on and for a long time, emphasis on MC as secreting histamine and heparin and as effector cells of immediate type hypersensitivity had almost completely distracted from other possible roles of MC in health and disease. The expanded knowledge on the structure, origin, and function of these cells has brought them on the front stage of the injury response and repair processes through the release of histamine, glycosaminoglycans, enzymes, cytokines, arachidonic acid derivatives and nitric oxide and perhaps through direct, membrane molecule mediated cell interactions.

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Structure of Mast Cells Von Recklinghausen (1863) probably observed what later came to be called MC, however his description did not lead to their identification as a specific cell type. Ehrlich (1878), while still a medical student, first identified MC in the connective tissue on the basis of the metachromatic staining of their cytoplasmic granules, which are not demonstrated by haematoxylin and eosin. These cells are found most often close to blood and lymph vessels, nerves, smooth muscle cells, exocrine glands, hair follicles and epithelial surfaces that are exposed to environmental antigens, such as those of the respiratory and digestive systems and skin. MC are oval, spindle, or spider shaped; the nucleus is oval or round, with peripheral clumps of chromatin. The chromotropic (i.e., metachromatically stained) secretion granules are 0.3-1 μm in diameter (Fig. 1). These cells are easily recognized by electron microscopy (Figs. 2, 3). The specific granules are delimited by a membrane and contain either a paracrystalline matrix with lamellar arrays (Fig. 2) or whorls and scrolls (Fig. 3). The latter pattern reveals lack of chymase, one of the serine protease products of MC. Significant granule heterogeneity can be found in any particular tissue and even between granules of a single MC (reviewed by Bacci et al., 1992; Metcalfe et al., 1997; Crivellato et al., 2004). Preformed mediators stored in the specific secretory granules can be released by morphologically distinct secretion types, referred to as compound exocytosis, piecemeal degranulation (Dvorak, 1997) and focal exocytosis, i.e. the exocytosis of individual granules (Crivellato et al., 2002). Compound exocytosis occurs characteristically during IgE stimulated anaphylactic degranulation and consists of the fusion of cytoplasmic granules with each other and of the most superficial ones with the plasma membrane, giving rise to open channels with the quick and massive release of granule content (Dvorak, 1997). In piecemeal degranulation there is a progressive loss of granule contents by exocytosis of shuttle vesicles between single granules and the cell surface (Crivellato et al., 2004; 2006), which allows for the independently regulated secretion of different molecules stored in a same granule (Melo et al., 2008).

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Figure 1. Rat mast cells with metachromatically stained granules; Toluidine blue; bar = 20 μm.

Figure 2. Granules of a human skin mast cell with paracrystalline matrix. Electron microscopy, bar = 1 μm.

Figure 3. Granules of a human skin mast cell with discrete scrolls. Electron microscopy, bar = 1 μm.

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Many molecules secreted by MC are presumably transported to the cell surface through small vesicles since they undergo constitutive secretion. The release of cytokines, chemokines, growth factor and neuroendocrine and antimicrobial peptides secreted constitutively by MC (Cutz et al., 1978; Goetzl et al., 1988; Bacci et al., 1995; Mekori and Metkalfe, 1999) is regulated through variations in their synthesis rate, with a lag of minutes to hours between MC stimulation and increase in cytokine secretion (Castells, 2006). This regulation may affect each cytokine independently of the others (Theoharides et al., 2007). Human MC also contain a few, highly osmiophilic lipid bodies which store arachidonic acid together with enzymes for the metabolism of this acid and possibly with tumour necrosis factor (TNF)alpha and basic fibroblast growth factor (FGF) (Dvorak et al., 2003). Mast cells secrete lipid products which are synthetized from the phospholipids of plasma membrane and of lipid bodies upon stimulation and include prostaglandins, leukotrienes, thromboxanes and platelet activating factor (Dvorak et al., 1997; Castells, 2006). Another secretory product of MC is nitric oxide (NO) (Masini et al., 1991); this molecule is synthesized in the cytosol and is directly secreted through the plasma membrane (Forsythe et al., 2001), moreover a granular form of the synthesizing enzyme, NO synthase, is located in MC granules, whose exact role is yet unclear (Bacci et al., 1994).

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Origin and Survival of Mast Cells Mature MC are the progeny of multipotent haematopoietic stem cells that commit to the MC lineage already in the bone marrow. The committed progenitors are positive for CD13, ckit, and CD34 but negative for FcεRI (Kirshenbaum et al., 1999). The secretion of TNFalpha by other, CD11b+ bone marrow cells is critical to the development and expansion of the MC lineage (Wright et al., 2006) and some yet unidentified action exerted by bone marrow dendritic cells on MC precursors is needed to prime these precursors for appropriate homing in peripheral tissues, at least in mice (Alcaide et al., 2007). Committed progenitors pass into the circulation (Kitamura and Ito, 2005) to complete their maturation within peripheral tissues (Galli, 2000; Crivellato et al., 2004). The adhesion to endothelial cells is mediated by α-4 integrins, vascular cell adhesion molecule (VCAM)1 and E-selectin (Boyce et al., 2002). The migration of MC precursors into tissues is stimulated by stem cell factor (SCF) and by eotaxin (Haley et al., 2000), the latter interacts with the chemokine receptors CXCR2, CCR3, CXCR4 and CCR5 (Ochi et al., 1999). In the tissues, further proliferation and differentiation of MC precursors depend on the presence of the local growth factors and cytokines, SCF, IL3, IL4, IL6, IL9 and nerve growth factor (NGF) (Saito et al., 1996; Kinoshita et al., 1999; Kanbe et al., 2000; Matsuzawa et al., 2003). Mast cell numbers in tissues are relatively constant, but hyperplasia occurs in the inflammatory and repair/remodeling stages of inflammatory/fibrotic disorders (Bischoff and Sellge, 2002). Growth factors and chemokines mediate the accumulation of MC to sites of inflammation (Meininger et al., 1992; Nillson et al., 1994; Nillson et al., 1999; Ochi et al., 1999; Romagnani et al., 1999; Zhao et al., 2002). A functional SCF and c-kit signaling system is crucial for MC growth and development and the lack of c-kit signaling in mice results in severe MC deficiency (Kitamura et al., 1978; Nakahata and Toru, 2002), whereas an elevated expression of c-kit in patients induces

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mastocytosis (Castells et al., 1996; Worobec et al., 1998) and the injection of SCF into the skin of humans results in local accumulation of MC (Costa et al., 1996). MC can quickly move within connective tissue and even transfer into epithelia and backwards (Enerback et al., 1986, Laitinen et al., 1993). Histamine itself promotes MC migration (Thurmond et al., 2004). Nerve growth factor has been reported to be a survival factor for rat peritoneal MC (Kawamoto et al., 1995) and for human cord blood-derived MC in the presence of SCF (Kanbe et al., 2000). Interlukin3 induces mouse MC growth and enhances their development in response to SCF in vitro; in humans, IL3 receptor is expressed by MC progenitors and intestinal MC but not by MC from lung, uterus, kidney, tonsils and skin (Bischoff and Sellge, 2002). The survival, adhesion to fibronectin and cytokine production of human umbilical cord blood-derived MC are stimulated by IL33 (Iikura et al., 2007); a possible role of adhesion receptors in MC survival has also been proposed (Ra et al., 1994; Okayama and Kawakami, 2006). Human MC express functional TNF-related apoptosis inducing ligand (TRAIL) receptors and SCF prevents their apoptosis induced by TRAIL receptor ligation (Puxeddu et al. 2003, Berent-Maoz et al, 2006); the survival of human mature MC in tissues depends on the local production of SCF since its withdrawal results in apoptosis (Iemura et al., 1994). Murine MC express the death receptor Fas (Apo-1/CD95), but only the C57 mast cell line is known to be susceptible to Fas dependent apoptosis; possible mechanisms of survival correlated with Fas inhibition are yet to be established (Hartmann et al., 1997).

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Heterogeneity of Mast Cells Fully differentiated MC display histochemical, electron microscopical, biochemical and functional heterogeneity. Different MC types are preferentially located in different tissues (Enerbäck, 1986; Galli, 1990). So-called connective tissue MC and mucosal MC have been identified in rodents (reviewed by Metcalfe et al., 1997). In humans, MC containing only tryptase (MCT) are the human equivalent of rat mucosal MC and are typically present in the lungs and intestinal mucosa, while MC containing both tryptase and chymase (MCTC) are the human equivalent of rat connective tissue MC and are typically found in the skin, synovial membrane and around blood vessels (Irani et al., 1989; Bacci et al., 1992; Bradding et al., 1995a; Ghannadan et al., 1998, Puxeddu et al., 2003; Table I). Another, less frequent subpopulation of human MC containing only chymase (MCC) has been observed in tissues (Weidner and Austen, 1993, Bacci et al., 1995) and can be differentiated in vitro (Li et al., 1996); its features have not yet been fully elucidated. The heterogeneity of MC is even more complex since, for instance, human lung MCTC but not skin derived MCTC release leukotriene (LT)C4 upon receptor mediated and receptorindependent stimulation (Oskeritzian et al., 2005). Mast cell types can interchange with each other, their ultimate phenotype (MCT or MCTC) being determined by the microenvironment. Human MC of different phenotypes may be obtained in vitro by varying the culture conditions.

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Table 1. Differences in some properties of T and TC mast cells

Morphology Formaldehyde sensitivity of granule metachromasia Staining properties of granules Electron microscopy of granules Granule glycosaminoglycan Monoamine content Enzyme content of granules

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Response to compound 48/80 Sensitivity to disodiumchromoglycate Life span Main location in humans Synonyms

Characteristic antigens Distinctive cytokines secreted Complement receptor Behavior in pathology

T mast cells Small, sparsely granulated Sensitive

TC mast cells Large, densely granulated Resistant

Sensitive Smaller granules, more variable in shape, containing discrete scrolls Chondroitin sulphate Low Tryptase (perhaps carboxypeptidase B, elastase)

Resistant Larger granules, variable in shape

Proliferative, non secretory Insensitive Short Lung, bowel mucosa So called “mucosal”, “typical”, mouse E (for chondroitinsulphate- E) CD50 GM-CSF, TGF-β None Increased around sites of Th –activation and in allergic and parasitic diseases. Decreased in AIDS and chronic immunodeficiency diseases

less

Heparin High Chymase, tryptase, carboxypeptidase B, elastase, cathepsin G Secretory Sensitive Long Skin, bowel submucosa So called “connective tissue, “atypical”, mouse H (for heparin) CD32 IL3 CD88 (C5aR) Increased in fibrotic diseases. Unchanged in allergic and parasitic diseases, AIDS and chronic immunodeficiency diseases

Stimulation with SCF alone or plus IL4 leads to early expression of tryptase and only after several weeks to that of chymase (Xia et al., 1997; Kinoshita et al., 1999). Expression of chymase seems to be inhibited by culture with IL6 (Toru et al., 1998a) and on the contrary is promoted since early in culture with medium conditioned by a human mastocytosis cell strain (Li et al., 1996). Compound 48/80 tested in the skin of patients with scleroderma caused the release of histamine in the skin at the edge of lesions to a much higher extent than in healthy skin, suggesting that in a same tissue MC in disease can differ in their functional properties from MC in health (Pearson et al., 1988). The numbers of intestinal mucosal and submucosal MCT, but not MCTC, are reduced in patients with congenital combined immunodeficiency or acquired immunodeficiency syndrome, and a mixture of MCTC and MCT cells is seen in the synovial membrane of subjects with rheumatoid arthritis in areas with heavy lymphocytic infiltration, whereas only

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MCTC cells are found in normal synovium (Irani et al., 1987b). These findings are consistent with a T cell requirement for the generation or maintenance of MCT (Irani et al., 1987a). The influence of microenvironment is indicated also by the fact that MC can vary their protease content in a tissue specific pattern (Friend et al., 1996).

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Mast Cell Receptors As already mentioned, MC express c-Kit, which drives their differentiation in response to SCF (Zhang et al., 1998; Hide et al., 2007). Mast cells express the immunoglobulin high affinity receptor (FcεRI); cross linking of these receptors upon binding of multivalent antigens initiates MC activation. Besides FcεRI receptors, human MC express a large array of immunoglobulins and chemokine receptors and of adhesion molecules which may play roles in the control of MC distribution within tissues and of MC activation (Vliagolftis and Metcalfe, 1997). The high-affinity IgG receptor (FcγRI) is expressed on human MC surface after interferon (IFN) gamma treatment. Its signaling pathway resembles that observed following FcεRI aggregation, with phosphorylation of Src kinases and p72 and subsequent activation of multiple substrates, including phosphatidylinositol-3-kinase; this enzyme is phosphorylated also after FcεRI stimulation, but only FcγRI signaling requires this molecule for degranulation. The induction of MC activation through FcγRI in addition to FcεRI in an IFNgamma rich environment appears to be an additional mechanism by which MC can be induced to produce a diversity of inflammatory mediators (Woolhiser et al., 2001; Okayama et al., 2003; Tkaczyk et al., 2004 ). Mast cells seem to express beta-2 adrenoceptors since beta-2 adrenoceptor agonists inhibit histamine secretion by lung MC (Scola et al., 2004a; 2004b). Mast cells express the chemokine receptor CCR3, which binds eotaxin, eotaxin-2 and eotaxin-3. This receptor can also bind other chemokines such as monocyte chemoattractant protein (MCP)3, MCP4 and the regulated on activation RANTES (Nickel et al., 1999; Romagnani et al., 1999). Most, perhaps all these chemokines cause MC migration in vitro. Another receptor expressed by MC is the C3a receptor. Known primarily for its secretagogue activity on skin MC, C3a is also a potent chemoattractant for these cells (Hartmann et al., 1997). Human MCTC, but not MCT, also express C5aR, i.e. CD88 (Oskeritzian et al., 2005); the effect of CD88 binding is still to be addressed. Mature MC express IL3 receptor (Dahl et al., 2004) which is expressed also by human basophils and blood monocytes (Valent and Bettelheim, 1992), IL4 (Hide et al., 2007) and IL5 (Dahl et al. 2004) receptors and c-kit, the receptor for SCF (Zhang et al., 1998). As already seen, c-kit stimulation drives the terminal differentiation of MC and plays other important roles in regulating MC biology, such as survival, activation and degranulation of mature MC. Mast cells express the prostaglandin E receptor (EP)-2, EP3 and EP4 (Feng et al., 2006; Kay et al., 2006; Wang and Lau, 2006). Prostaglandin E-receptor mediated stimulation leads to the secretion of peptide factors independent of degranulation (Abdel-Majid et al., 2004; Nakayama et al., 2006); it also potentiates the immunologically stimulated histamine

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secretion by human peripheral blood derived MC (Wang and Lau, 2006) and on the contrary inhibits the degranulation of human lung MC (Kay et al., 2006). Human MC express the peroxisome proliferator activated receptor (PPAR)-gamma, it seems to be involved in MC proliferation and differentiation (Bagga et al., 2004) and prostaglandin D2 generation (Paruchuri et al., 2008). Mast cells express the urokinase plasminogen activator receptor, CD87 (Sillaber et al., 1997; Hildenbrand et al., 2008): this receptor leads to tight localization of its ligand at the cell surface and hence regulates extracellular matrix turnover, cell migration, local fibrinolysis and invasion, and probably also cell signalling, proliferation and apoptosis in physiological and pathological conditions (Hildenbrand et al., 2008). Human MC express several receptors for neuropeptides namely neurokinin receptor (NKR)1, NKR2, NKR3 and VIP receptor-2 (VPAC2); activation of MC through FCεRI upregulates the expression of NK2R, NK3R and VPAC2 (Kulka et al., 2008). Skin MC express melanocortin receptors (MCR), mainly MCR5 (Böhm et al., 2006). Angiogenic factors, such as FGF2, vascular endothelial growth factor (VEGF) and platelet-derived endothelial cell growth factor (PDEGF) stimulate MC migration (Gruber et al., 1995) but the mechanisms mediating this response remain to be elucidated. In humans, pathogenic bacteria and fungi are able to induce a highly selective production of MC mediators [IL1ß, granulocyte monocyte-colony stimulatory function (GM-CSF), leukotrienes] through Toll-like receptor (TLR) stimulation (Mc Curdy et al., 2003). Mast cells, at least in rodents, also express protease activated receptor (PAR)1 and PAR2 by which they respond to thrombin and perhaps other proteases; this response may be mediated also by PAR independent mechanisms (Stenton et al., 2002; Dugina et al., 2004). The ligand receptor interaction is peculiar in that the enzyme cleaves the receptor, a part of which behaves as a tethered ligand for the other part thus activating a cell response (Barnes et al., 2004). Mast cells express receptors for several molecules which they secrete themselves. They respond to histamine by active migration (Thurmond et al., 2004), mediated by the G-proteincoupled H4 histamine receptor (Varga et al., 2005). Mouse MC respond to prostaglandin (PGD)2 with a block in histamine release (Hashimoto et al., 2005). Human cord bloodderived CD34-expressing MC progenitors respond to exogenous LT with calcium influx and phosphorylation of extracellular signal related protein kinase, which is enhanced when the cells are primed with IL4 (Mellor et al., 2001). LT and uridine diphosphate induce the generation of IL5, TNFalpha, macrophage inflammation protein (MIP)1beta and IL8 in IL4– primed human MC, and LT inhibitors depress the production of IL5 and TNFalpha after FcεRI stimulation, implying autocrine signaling by LT in MC (Mellor et al., 2002). Platelet activating factor (PAF) induces histamine release from human skin MC in vivo (Petersen et al., 1997), and calcium mobilization leading to migration of a human MC line in vitro (Nilsson et al., 2000); human nasal mucosa MC express immunohistochemically detectable PAF receptor (Shirasaki et al., 2005). Interleukin 6 inhibits the SCF-dependent development of human MC from CD34+ precursor (Kinoshita et al., 1999) implying that the latter have a receptor for this cytokine. GM-CSF controls mouse and human MC differentiation (Brambilla et al., 1993; Du et al., 1997; Welker et al., 2001; Dahl et al., 2004). Dahl et al. (2004), have described the presence of the receptor for this substance in human MC. Human

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and rodent MC respond to NGF, which stimulates their differentiation and survival (Matsuda et al., 1991; Horigome et al., 1994; Welker et al., 1998, 2000; Kulka et al., 2008). Cathelicidin-like antimicrobial peptides also elicit responses from MC, implying the presence of receptors; more precisely, MC respond to peptide LL-37 with positive chemotaxis, degranulation and PGD2 production (Niyonsaba et al., 2001; 2002; 2003). Nitric oxide elicits responses by MC modulating histamine release (Sekar et al., 2005). Studies on MC obtained from human tissues have demonstrated surface expression of the ß1 integrins very late (activation) antigen (VLA)3, VLA4, and VLA5, and of αvβ3 integrin (Valent and Bettelheim, 1992; Columbo et al., 1995). The natural ligands of VLA3 are laminin, type I collagen and fibronectin; those of VLA4 are fibronectin and VCAM1; those of VLA-5 are fibronectin and vitronectin, and those of αvβ3 are fibronectin, thrombospondin and fibrinogen. It has been reported that β1 integrins are not simple binding sites for adhesion, but also receptors involved in MC activation, up-regulation of cytokine expression, and survival (Ra et al., 1994; Zwart et al., 2004; Okayama and Kawakami, 2006). Additional adhesion molecules expressed by MC are CD44, a hyaluronic acid receptor, and siglec-8, a molecule which binds to sialic acid receptor (Bochner and Schleimer, 2001). Mast cells degranulate in response to several non-immunological agents, such as compound 48/80, basic polypeptides (polylysine, polyarginine), exogenous opioids and cannabinoids, substance P, anaphylatoxin C5a, and the calcium ionophore A23187 (Selye, 1966; Lowman et al., 1988; Church et al., 1991). Most of these agents apparently can penetrate the cell membrane and interact with G-proteins through a receptor independent mechanism (Klinker and Seifert, 1997; Bueb et al., 2001; Lau et al., 2001; Jonsson et al., 2006; Sheen et al., 2007; Shefler et al., 2008). Compound 48/80 may also interact with a phospholipase (Palomäki and Laitinen, 2006). However, basic neuropeptides may act through specific receptors (Kulka et al., 2008) and basic molecules in general may activate MC through a Mas-related gene receptor, namely MrgX2 (Tatemoto et al., 2006).

Mast Cell Mediators Mast cells release a heterogeous group of mediators that differ in their synthesis, storage and secretion sites and processes.

Preformed Mediators in the Specific Granules Histamine is stored in the secretory granules of all mature MC (Schwartz et al., 1987). The amounts of stored histamine and presumably the mechanisms for synthesis and storage are similar among different types of MC (Schwartz et al., 1987; Leino et al., 1990). The effects of histamine are mediated through H1, H2, H3, and H4 receptors (Harvey and Shocket, 1980; Thurmond et al., 2004; Varga et al., 2005) which influence inflammatory and immune processes (Meretey et al., 1989; Falus and Meretey, 1992) and the function of epithelial cells (Lim et al., 2005), myocardiocytes (Meretey et al., 1989; Falus and Meretey, 1992) and neurons (Chrush et al., 1999; Lim et al., 2005). Mast cells themselves express H4

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receptor (Varga et al., 2005), therefore, MC both secrete and respond to histamine, activating possible feed-back loops by this way. In the gastric mucosa, where also epithelial enterochromaffin-like (ECL) cells store and release histamine; the numbers of MC and ECL cells are differently regulated: MC increase in inflammatory processes and ECL cells upon gastrin hyperstimulation (Bechi et al., 1995; 1996). Mast cell granules are rich in proteoglycans, the glycosaminoglycan (GAG) composition of which depends on species, tissue distribution, and maturity of the cell. Human MC contain heparin. In rodents, heparin is restricted to MC of connective tissue and serosa (peritoneal MC) and is absent from mucosal MC. Chondroitin sulfate E is also found within the granules of human MC (Stevens et al., 1988) and of mucosal MC of mouse (Razin et al., 1982) and rat (Stevens et al., 1986). Heparin and chondroitin sulfate are present in mature human lung MC in a ratio of 2:1 (Stevens et al., 1988). Heparin and chondroitin sulfate proteoglycans stabilize and regulate secretory granule proteases (Serafin et al., 1986). Upon secretion, heparin binds to, and activates antithrombin III, which prevents the activation of the clotting cascade; heparin may also interact with and inhibit the activity of cytokines, chemokines, and growth factors, including FGF2 which is involved in angiogenesis and wound healing (Macri et al., 2007). Therefore, heparin is a negative regulator of angiogenesis (Presta et al., 2003). The major MC protease is tryptase, a 130 kd serine protease, which is stored in a fully active form in the specific granules. It represents the most abundant constituent of human MC. About 10 pg/cell has been detected in lung MC and up to 35 pg in skin MC (Schwartz et al., 1987). Tryptase is present in all human MC but is absent or in negligible quantities in other cell types (Craig et al., 1986; Walls et al., 1990) and is relatively resistant to most fixation methods (Walls et al., 1990; Bacci et al., 1995), therefore it is considered the most specific and sensitive target for the immunohistochemical detection of MC in tissues, even when using fixatives that are not suitable for conventional histochemical procedures. Tryptase is secreted from human MC in complexes with proteoglycans which have molecular weights of 200 to 250 kDa. The large size of these active complexes severely limits diffusion away from sites of MC activation and helps to explain why an increase in circulating levels of tryptase seems to occur only upon massive MC activation such as anaphylactic shock. A consequence of this is that the most likely targets of tryptase are in the immediate vicinity of MC (Goldstein et al., 1992). The action of tryptase is to cleave various neuropeptides and matrix components. In addition, it is emerging as a potent growth factor for fibroblasts, endothelial cells and muscle cells (Blair et al., 1997; Gruber et al., 1997), and therefore can stimulate angogenesis (Blair et al., 1997; Somasundaram et al., 2005). The actions of tryptase on cells or tissues seem in each case to be dependent on an intact catalytic site, the effects being markedly reduced when tryptase is added or administered in the presence of a protease inhibitor. Tryptase appears capable of interacting with and activating the G-protein coupled proteases activated receptor (PARs) which are also receptors for thrombin (Molino et al., 1997; Ui et al., 2006, Palmer et al., 2007). Since PARs are expressed by MC (Stenton et al., 2002; Dugina et al., 2004), this molecule may also be part of a feeback loop. Human chymase is associated to the MCTC and MCC populations (Irani et al., 1989). This 30 kD protease is present within the granules of MCTC, in an estimated concentration of 4.5 pg/cell. It is stored in the same secretory granules that contain tryptase and is released in a macromolecular complex with carboxypeptidase and proteoglycans, distinct from that of

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tryptase. Like tryptase, chymase is present in a catalytically active form in the granules but appears to have little activity in the acidic conditions therein (reviewed by Church et al., 2003). Chymase may modulate inflammatory processes by controlling the bioavailability of certain cytokines at sites of MC activation. IL1 is converted into an active form by this protease (Mizutani et al., 1991), whereas IL4 is degraded. This enzyme also may be responsible for tissue destruction and matrix remodeling, by acting directly on different collagen molecules and activating matrix proteases (Caughey, 2007). Also, it can release transforming growth factor (TGF)beta, a multifunctional cytokine which can stimulate the formation of connective tissue, from the extracellular matrix (Taipale et al., 1995, Lindstet et al., 2001). Chymase catalyzes the formation of a 31 amino acid form of endothelin (ET), called ET1 (1-31), from pro ET1 (D'Orléans-Juste et al., 2003). ET1 (1-31), similar to whole ET1, elicits a vasoconstrictor effect and is mitogenic for vascular smooth muscle cells (Nagata et al., 2000). The addition of chymase, isolated and purified from rat serosal MC, to cultured rat aortic smooth muscle cells of the synthetic phenotype (sSMCs) inhibits their proliferation by blocking the G0/G1-S transition in the cell (Wang et al., 2001). Rat chymase and recombinant human chymase inhibit the expression of collagen type I and type III mRNA in sSMCs and in human coronary arterial SMCs. The mechanism of these actions remains to be elucidated, it may be in part mediated by growth factors since the growth-inhibitory effect of chymase on s-SMCs was partially reversed by the addition of a neutralizing antibody against TGFbeta (Wang et al., 2001). Chymase is also able to convert angiotensin I to angiotensin II independent of plasma angiotensin converting enzyme, which may be relevant to stimulate both intimal proliferation at sites of vascular injury and angiogenesis, with the intermediate of angiotensin II-induced secretion of VEGF (Ibaraki et al., 2005; Kondo et al. 2006; Miyazaki et al., 2006); indeed chymase it is a potent mediator of MC-induced angiogenesis (Muramatsu et al., 2000). Chymase can also interact with PAR but the possible effects of this interaction are not yet established (Doggrell and Wanstall, 2004; Takai et al., 2004). Carboxypeptidase A is a zinc – containing metalloexopeptidase, stored in human MC specific granules along with tryptase and chymase and released upon activation: its presence in the plasma has not been reported. Human MC carboxypeptidase A is not expressed by other cells of hematopoietic lineages (Castells, 2006). By immunohistochemistry, the enzyme can be detected in MCTC subset (Irani et al., 1991; Weidner et al., 1992). At least in the rat, carboxypeptidase converts angiotensin I to angiotensin II and can also inactivate bradykinin and substance P (Cole et al., 1991). In humans, the expression of cathepsin G is confined largely to MCC and MCTC subsets while it is expressed little or not at all in the subset of MCT (Schechter et al., 1990). Compared with most proteases, cathepsin G is generally weaker but with broader specificity. It exhibits tryptic and chymotryptic activity (Caughey, 2007). Since cathepsin G, and chymase as well, are likely to be inactivated quickly by natural inhibitors and diffuse easily into the tissue, they probably degrade cytokines already present in the extracellular space and deposited by other cells, but not those produced by their own cell of origin (Zhao et al., 2005). Cathepsin G can exert some effects also directly on cells including MC themselves by interacting with PARs (Steinhoff et al., 2005; Ramachandran et al., 2007).

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The proteases secreted by MC include also several matrix metalloproteinases (MMP), however not stored in the specific granules (Di Girolamo and Wakefield, 2000).

Lipid Mediators Prostaglandin D2 is generated after the immunologic activation of human MC. It has been detected in human pulmonary MC after IgE-mediated activation (Lewis et al., 1982). Two receptors, DP1 and DP2, both G-protein coupled, mediate the activity on several cell types, including leukocyte and dendritic cells (Hirai et al., 2001, Boyce, 2007). Murine MC also respond to PGD2 which inhibits IgE-mediated scratching by suppressing histamine release from MC (Hashimoto et al., 2005). Typically, upon stimulation, MC release large amounts of leukotrienes (Malaviya et al, 1993), and are also a target of these autocoids, therefore MC secreted LT may be involved in autocrine loops (Mellor et al., 2001; 2002). Platelet activating factor is produced from arachidonate in membrane phospholipids and is found, among several cell types, in MC (reviewed by Castells et al., 2006). Human MC also respond to PAF, in a possible autocrine loop (Petersen et al., 1997; Nilsson et al., 2000; Shirasaki et al., 2005)

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Cytokines and Chemokines Human mature MC secrete several cytokines and chemokines. Tumour Necrosis factor alpha plays roles in regulating many biological processes, including inflammation, immune responses, cytokine production, apoptosis, and cell activation (for an extensive review see Cheng et al., 2003). TNFalpha was the first cytokine localized in MC (Gordon and Galli, 1990, 1991). It is stored in part in the secretory granules (Gordon and Galli, 1990; Bradding et al., 1995b; Bacci et al., 1998; 2006; 2008; RiccardiArbi et al., 2004), and in part is released by constitutive secretion; its transcription is further induced after MC activation (Gordon and Galli, 1991). The release of TNFalpha by MC may be a key factor in the ultraviolet radiation-induced inhibition of cell mediated immune reactions (Alard et al., 1999). Interleukin-4 is a key cytokine in the development of allergic inflammation through its ability to drive the differentiation of naive Th cells into Th2 lymphocytes, leading to the production of effector cytokines such as IL4, IL5, IL9, and IL13. It is associated with the induction of the ε isotype switch and the secretion of IgE by B lymphocytes and enhances IgE-mediated responses by up-regulating IgE receptors on B lymphocytes, MC, and basophils. IL4 also induces VCAM-1 on vascular endothelium and thus directs the migration of T lymphocytes, monocytes, basophils, and eosinophils to the inflammation site (Reviewed by Weiss and Brown, 2001). Interleukin-4 is immunocytochemically detectable in MC of the bronchial and nasal mucosa. Unlike IL5, it appears to be present in both MCT and MCTC (Bradding et al., 1995c). Immuno electron microscopy has shown that IL4 is localized to the specific secretion granules (Bradding et al., 1992). This cytokine regulates the expression of

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FcεRI, LT4 synthase and cysteinyl-LT receptors by MC themselves (Hsieh et al., 2001; Mellor et al., 2002). By immunohistochemistry (Bradding et al., 1993; 1995c) and in situ hybridization (Kay et al., 1995; Ying et al., 1995) approximately 10% of MC appear to be interleukin-5 positive; positivity appears to be restricted to MCT (Bradding et al., 1995c). In vitro studies have shown that IgE-dependent stimulation of human lung MC induces IL5 production (Okayama et al., 1995; Jaffe et al., 1996), which may be suppressed by dexamethasone (Glaum et al., 1995). Dahl et al. (2004), have described receptors for IL5 in human MC. Interleukin-6 has been shown to be present in human MC (Bradding et al., 1993; 1994; Krüger-Krasagakes et al., 1996) and to be restricted to the MCT subset in both bronchial and nasal mucosa, where this cytokine promotes mucus production (Bradding et al., 1995a). IL6 inhibits the SCF-dependent development of human MC derived from CD34+ cord blood cells (Kinoshita et al., 1999). Interleukin-8 mRNA is expressed by HMC-1 after activation (Möller et al., 1993; Buckley et al., 1995) and immunoreactive IL8 has been found upon activation in the specific granules of skin MC and the HMC-1 cell line (Möller et al., 1993; Grutzkau et al., 1997). Interleukin-13, a cytokine with many properties in common with IL4, has been demonstrated to be associated with MC in conjunctival biopsies from patients with seasonal allergic conjunctivitis (Anderson et al., 2001). IL13 is produced by MC in response to FcεRI receptor cross-linking (Burd et al., 1995; Jaffe et al., 1996; Kobayashi et al., 1998; Toru et al., 1998b). The gene expression of IL13 by HMC-1 cells and human lung MC, which was detected at a low level in an unstimulated condition, was increased by phorbol-myristoylacetate (PMA)/ionomycin and suppressed by dexamethasone. (Fushimi et al., 1998). Mast cells secrete IL13 also in the presence of IL4 (Toru et al., 1998b) and of SCF (Kanbe et al., 1999a). Mast cells are a source of many chemokines (Nakajima et al., 2002), i.e. of chemotactic cytokines that, among several functions, regulate the migration of hematopoietic cells including MC (Okayama and Kawakami, 2006). These substances are able to recruit effector cells and regulate immune response (Marshall, 2004). Mast cells themselves express many chemokine receptors and migrate in response to their stimulation by several ligands (Juremalm and Nillson, 2005); in particular, chemokines induce human MC migration with no exocytosis or degranulation. Chemokine receptors are differentially expressed during MC maturation and on human MC of different phenotypes (Castells et al., 2006).

Growth Factors Rat MC synthesize, store and release nerve growth factor (Leon et al., 1994) and immunoreactivity to NGF has been also found in human MC (Marinova et al., 2007). NGF has also MC among the target. It induces in vitro the development of MC from mouse bone marrow cells in the presence of IL3 (Matsuda et al., 1991), inhibits the apoptosis of rat peritoneal MC (Horigome et al., 1994), and promotes in vitro the maturation of human cord blood-derived MC and of the immature human HMC-1 line (Welker et al., 1998, 1999, 2000; Kulka et al., 2008). Treatment of newborn rats with NGF induces an increase in the number

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of MC in several locations (Aloe et al., 1977), while the administration of neutralizing antiNGF antibodies reduces it (Aloe, 1988). In these activities NGF can replace SCF that otherwise, as mentioned earlier, is the key growth factor responsible for MC differentiation, survival and activation. Stem cell factor is produced by human MCT and MCTC (Zhang et al., 1998). Human skin and lung MC as well as cord blood and peripheral blood derived MC and mast/basophil cell line (HMC-1 and KU-812) store and secrete SCF upon activation (De Paulis et al., 1999; Welker et al., 1999). Besides MC precursors proliferation and differentiation, SCF regulates MC survival, chemotaxis, adhesion and release of inflammatory mediators (see Nillson et al., 1994, Grabbe et al., 1996, Da Silva et al., 2006), in particular of chemokines (Oliveira and Lukacs, 2001) and proinflammatory cytokines (Bishoff and Sellge, 2002). Injection of SCF into the skin causes MC hyperplasia (Costa et al., 1996). Transforming growth factor-beta is released by mouse bone marrow derived MC after IgE stimulation (Gordon, 2000) and in turn stimulates MC recruitment (Gruber et al., 1994). In the mouse, the production of TGFbeta is stimulated by the Th2 cytokine IL9 (Mesples et al., 2005). Since chymase catalyzed proteolysis activates TGFbeta (Lindstedt et al., 2001) and this factor and chymase are co-stored in secretory granules, degranulation might coordinately trigger secretion and activation of TGFbeta (Lindstedt et al., 2001). Human MC can secrete granulocyte macrophage-colony stimulating factor (Lippert et al., 2000); this secretion is stimulated by neuropeptides (Kulka et al., 2008). Mast cells also secrete platelet derived growth factor (PDGF), VEGF and FGF (Artuc et al., 1999).

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Neuropeptides Mast cells contain vaso active intestinal polypeptide (VIP) (Cutz et al., 1978; Bacci et al., 1995). Goetzl et al. (1988) have demonstrated that MCs release both VIP10-28 free acid and, to a lesser extent two amino terminally extended forms of VIP1-28 as well as a somatostatin-like peptide. It is yet unclear whether and to which extent these molecules are contained within chromotropic granules.

Gas Immunoreactivity with polyclonal antibodies specific for neuronal (n) nitric oxide synthase (NOs) (Bacci et al., 1994; 1995; 1998) was first described in the granules of the MC of human normal mucosa of nose and digestive system. The possible biological role of this enzyme within granules destined to exocytosis is still obscure. More recently, some subsets of MCs in the cirrhotic liver were shown to synthesize inducible (i) NOS mRNA and protein (Koda et al., 2000), and iNOs expression was found in MCs in testicular tissue (Sezer et al., 2005), and in the normal and atheromatous human arterial wall (Bacci et al., 2008). Taken together, these results seem to suggest the hypothesis that human MC utilize both nNOs for

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constitutive NO production and iNOs for an additional role in inflammatory processes. These cells can also responde to this gas by modulating histamine release (Sekar et al., 2005).

Antimicrobial Peptides Human and mouse MC have been shown to produce cathelicidin and cathelicidin-related antimicrobial peptides (Di Nardo et al., 2003, Eckmann, 2005), through which they contribute to antimicrobial host defence (Niyonsaba et al., 2001, 2002, 2003; Di Nardo et al., 2003; Zanetti, 2004). However, the presence of the cathelicidin peptide has not been confirmed in human skin MC by light and electron immunocytochemistry (Nelson et al., 2007). In turn, MC chemotaxis and secretion are stimulated by peptide LL-37 (Niyonsaba, 2001; 2002); in particular LL-37 stimulates the expression of TLR4 and induces the release of IL4, IL5 and IL1beta from MC (Yoshioka et al., 2008).

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Mast Cell Relationships to Basophils Mast cells and basophils are two distint cell populations, which exhibit some common features including histamine and heparin secretion and share a common precursor up to an advanced stage of differentiation. Basophils mature entirely in the bone marrow whereas elements of MC lineage circulate in the blood as precursors and complete their differentiation in peripheral tissues (Galli, 2000; Crivellato, 2004). Under physiological conditions, basophils have a life-span of several days. IL3 can promote the production and survival of human basophils in vitro and can induce basophilia in vivo (Valent, 1989). On the contrary, MC can be very long-living and even MC that are apparently mature can proliferate under certain conditions (Wedemeyer et al., 2000). Many aspects of MC development and survival are critically regulated by SCF; this is the ligand for the c-kit receptor, a member of the receptor tyrosine kinase III family of growth factor receptors, which is expressed on the MC surface (Zhang et al., 1998). Basophils on the contrary do not express this receptor (De Paulis et al., 1999). Several cytokines stimulate the activation and degranulation of basophils, while they induce MC to migrate but not to degranulate (Castells et al., 2006). The spectrum of basophil-derived cytokines appears to be more limited than that of MC (Redrup et al., 1998). The main differences between MC and basophils are summarized in Table II (Bacci et al., 1992; Galli et al., 2000; Galli et al., 2005).

Immune Function of Mast Cells During evolution MC have preceded lymphocytes and other cells of the immune system (Kinet, 2007). MC exist in lower vertebrates, such as fish and amphibians (Monteforte et al., 2001; Silphaduang and Noga, 2001; Baccari et al., 2003; Reite and Evensen, 2006), suggesting a function in primitive immune systems that lacked an adaptive branch. Thus,

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evolution has built up a useful immune network of redundant mechanisms, MC probably being one of the first (Kinet, 2007). Table 2. Differential features of basophils and mast cells in human

Average diameter Nucleus Granules

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Aggregates of glycogen Compound exocytosis Tryptase in granules Differentiation of precursor under fibroblast stimulation Expression of c-kit

Basophils 9 micron Segmented Relatively few, large and uniform in appearance Present Rare Absent No

Mast cells 14 micron Non Segmented Relatively small, numerous and variable in appearance Absent Common Present Yes

No

Yes

The role of MC in IgE mediated reactions is well known and does not need to be analyzed here. Experimental evidence shows that MC not only exert critical effector functions in classic IgE-associated allergic disorders, but also in a variety of IgE-independent processes, which lead to cell activation and initiation of inflammatory responses (Stassen et al., 2002) and can contribute to host immune defence in the context of both acquired (adaptive) and innate responses (Mekori and Metcalfe, 1999). At least in rodents, MC have even the capacity to recognize and phagocytose bacteria and subsequently serve as antigen presenting cells to T lymphocytes in an MHC class I restricted fashion (Malaviya and Abraham, 2001). Mouse bone marrow derived MC can express MHC class II antigens upon stimulation with IFNgamma (Frandji et al., 1993; Raposo et al., 1997) or without activation (Vincent-Schneider et al., 2001); the expression of MHC class II by MCs is upregulated by lipopolysaccharide and downregulated by IL3 (Castells et al., 1999). However, recent studies have failed to detect MHC class II on either resting or activared mouse MC (Nakae et al. 2006; Kambayashi et al., 2008). MC can express the costimulatory molecules needed to make an MHC mediated signal immunogenic: lung or bone marrow derived human MC express the costimulatory molecule CD54 (Ghannadan et al., 1998; Shimizu et al., 2002), mouse bone marrow-derived MC express B7 (Nakae et al., 2006); human bone marrow derived MC express CD86 (B7-2) when stimulated with GMCSF (Frandji et al., 1996) and cultured human MC express CD80 (B7-1) after internalization of E. coli (Kulka et al., 2006). Mast cells are necessary for inducing local lymph node hypertrophy following infection (McLachlan et al., 2003). This role may depend in part by TNFalpha mediated DC migration (Wang et al., 1997; McLachlan et al., 2003; Bryce et al. 2004; Suto et al., 2006). Mast celldeficient mice fail to display DC migration out of the epidermis, an observation that correlates with decreased production of mRNAs encoding IL1beta, IL6, MCP1, mouse mast cell protease-6, and TNFalpha (Bryce et al. 2004). Mast cell-deficient mice and TNFalpha deficient mice have similar deficits in contact hypersensitivity responses following

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sensitization and challenge, including reduced numbers of DCs in draining lymph nodes at early times after antigen administration, which may be overcome in mast cell-deficient mice by reconstitution with bone marrow from wild type mouse but not with TNFalpha deficient mouse bone marrow (Suto et al., 2006). In humans, MC release of CCL1, a chemokine, recruits CCR8 (CCL1 receptor)-expressing Langerhans cells to sites of atopic inflammation (Gombert et al., 2005). These interactions may explain the relationship of MC to dendritic cells during immune responses in the skin (Mori et al., 1994) and in secondary lymphoid organs (Marcenaro et al., 2006). The LT receptor pathway is essential in the recruitment of CD4 and CD8 T-effector cells to sites of inflammation (Goodarzi et al., 2003; Ott et al., 2003; Tager et al., 2003; Taube et al., 2006). MC produce LT4 and are able to induce T-cell chemotaxis through this pathway in vitro and in vivo (Goodarzi et al., 2003; Ott et al., 2003; Tager et al., 2003). A role for MC in helminth immunity has been described in T. spiralis; studies using c-kit defective, MC deficient W/W(v) mice or SCF specific antibodies that impair mast-cell function showed that SCF is required for mucosal mastocytosis during helminth infection, and is important for effective helminth expulsion. Furthermore, mice deficient in mast cell protease 1 are unable to expel T. spiralis indicating the antihelminth properties of mast-cell cytoplasmic granules; however this protease has little effect on N. brasiliensis host resistance (Antony et al., 2007). Human MC can be activated by different HIV-1 proteins (gp120 and Tat) and thereby represent a potentially important source of Th2 cytokines during HIV-1 infection (Marone et al., 2001).

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Mast Cells and Wound Repair Mast cells are able to control the key events of wound healing: inflammatory response, revascularization of damaged tissue, re-epithelialization, and deposition and subsequent remodeling of connective tissue (Artuc et al., 1999, Baum and Arpey, 2005). Once activated by tissue injury, MC release mediators which induce vasodilation and increase vascular permeability (Weller et al., 2006). The endothelial cells, in turn, influence the functional state of MC by releasing SCF, IL3 and thrombin which enhance migration, proliferation and local differentiation of MC (Baghestanian et al., 1997; Metcalfe et al., 1997). At the edge of wound, keratinocytes can secrete several cytokines and LL37, which influence MC recruitment and function. Mast cell numbers increase at the border of a wound (Bonelli et al., 2003) up to a maximum within 1–3 h from trauma, and decrease thereafter becoming less than baseline values after 6 h (Bonelli et al., 2003) (Fig. 3). The quick variation in MC numbers and location within connective tissue (15 min for trauma, in the skin) speaks against recruitment of precursors and differentiation of new MC as a relevant mechanism in this phase of the response to injury. Migration of already present cells can be proposed as an alternative explanation. These cells increase again in number later on, to a maximum within 10 days, and decrease thereafter returning to control values after 21 days from wounding (Trautmann et al., 2000; Fig. 4). The late increase correlates with the upregulation of MCP1 (Trautmann et

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al., 2000) and with the production of TGFbeta, which is also a potent chemoattractant for MC (Gruber et al., 1997).

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Figure 4. Time course of mast cell numbers in skin wounds. The figure was constructed from the data of Bacci et al. (2006) and Trautmann et al. (2000) with the permission of the authors and publishers.

Molecules of MC origin such as TNFalpha, which is released already in the first 15 minutes after wounding (Bacci et al., 2006), and histamine, which is released even earlier, favor the adhesion of leukocytes to vessel by increasing the expression of several endothelial adhesion molecules (Walsh et al., 1991; Christofidou-Solomidou et al., 1996). Leukotrienes, proteases and cytokines, especially IL8, released by MC, represent chemotactic signals for neutrophils, basophils and eosinophils (Ribeiro et al., 1997). Also tryptase and cathepsin G regulate endothelial-leukocyte interactions and leukocyte behavior at sites of inflammation (Steinhoff et al., 2005). Chymase can induce eosinophils to express chemokines for neutrophils (Terakawa et al., 2006). As the inflammatory process develops, circulating leukocytes are recruited within 3 hours from trauma (Bonelli et al., 2003), first granulocytes and shortly after monocytes; the latter cells are activated and subsequently transform into macrophages, which peak in number at later time points (48–72 hours), remain longer (days to weeks), and participate in a much more complex role to wound healing than granulocytes. Chemoattractants for monocytes and macrophages include fibronectin, elastin, complement factors such as C3a and C5a, thrombin and many factors secreted by MC: TGFbeta, platelet derived growth factor, TGFalpha, VEGF, insuline like growth factor1, NGF, MCP1, and MIP1. Once activated, macrophages reside in the provisional fibrin-based extracellular matrix and secrete a number of growth factors and cytokines which play roles in the regulation of the late phases of inflammation and tissue repair (Singer and Clark, 1999; Baum and Arpey, 2005). Mast cells themselves can contribute directly to regulating these phases of injury response. The responses of fibroblasts may be regulated by TGFbeta, TNFalpha and proteases (Ramachandran et al., 2007; Sharma et al., 2007).

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Angiogenesis is stimulated by TGFbeta, VEGF (Abdel-Majid and Marshall, 2004), chymase (Muramatsu et al., 2000; Doggrell and Wanstall, 2004), and tryptase (Somasundaram et al., 2005). On the contrary, heparin may inhibit angiogenesis by interacting with, and inhibiting pro-angiogenetic factors (Presta et al., 2003). In the late phases of repair, MC derived growth factors (i.e. FGF, TGF) and cytokines (IL1, IL4, IL6) may influence the phenotype of the activated fibroblasts inducing the appearance of particular cells with contractile activity (the myofibroblasts), which ensure the changeover from fibroplasia to contraction and final healing of the wound (Hebda et al., 1993; Moulin et al., 1998). As anticipated, MC can promote the conversion of fibroblasts to a myofibroblast phenotype which facilitates wound contraction and closure (Gailit et al., 2001). Mast cell extracts can activate fibroblasts, promoting collagen synthesis and activation of gelatinase A, an enzyme involved in matrix remodeling (Berton et al., 2000). This effect may be partly due to tryptase, which has been shown to stimulate the synthesis of type I collagen in human dermal fibroblast (Abe et al., 1998). Indeed MC may be involved in tissue remodeling, the key feature of late wound healing response but which is also relevant to physiological conditions. Skin MC produce and release potent proteolytic enzymes, such as matrix metalloproteinases, which facilitate tissue remodeling by initiating the degradation of extracellular matrix (Kanbe et al., 1999b). It has been demonstrated that cutaneous remodeling, e.g. during hair follicle growth and regression, is modulated by skin MCs and MC-derived histamine (Maurer et al., 1997a). Inhibition of MC histamine synthesis in wounded rats has been shown to decrease the hydroxyproline content of granulation tissue, to delay epithelization, and to reduce wound breaking strength (Bairy et al., 1991). The participation of MC to the response to injury is not restricted to trauma, but takes place upon ischemia, as shown in myocardial infarction. In this case, in the early phases of the response to temporary occlusion of a coronary artery in the dog, there is an increase of TNFalpha and histamine suggestive of massive degranulation of MC. This phenomen is not followed by an increase in the number of these cells. On the contrary, an increase in MC number has been shown to occur late, as a preliminary step to reparative fibrosis (Frangogiannis et al., 1998; 2002). The questions arises as to how are MC stimulated to move into and within tissue. Possible sources of stimuli early upon injury are extravasated platelets and autochthonous cells of damaged tissue itself, in particular endothelial cells (Diegelman and Evans, 2004, Yukami et al., 2007), nerve fibers (Steinhoff et al., 2003) and the epidermis, as indicated by the ordered approach of MC to this tissue upon selective photoinduced damage (Prignano et al., 2003). The late, new increase in number is presumably dependent on influx of precursors and their differentiation under stimuli provided by macrophages and possibly other cells in the site of inflammation (Trautmann et al., 2000). The final reduction to baseline numbers and activity may simply derive from the lack of further stimulation as repair proceeds and macrophages themselves disappear, however the possible existence of active mechanisms to switch off the inflammatory process, instead of simple lack of stimulation, has yet to be addressed. Impairment of such a switch off, or persistent activation of stimulating mechanisms may be responsible for the persistence and protracted activation of high numbers

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of MC, leading to fibrosis (Garzubenko et al., 2002; Ozbilgin and Inan, 2003; Garzubenko et al., 2004; Riekki et al., 2004).

Conclusion

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Mast cells can be proposed for a double role in injury response. Very early they release different types of mediators activating the vascular phase of inflammation and the recruitment of leukocytes which provide for the cellular phase of inflammation itself. To these aims, MC most probably concur with other ready-to-fire local control systems, i.e. platelets (which are activated immediately upon endothelial damage) and peripheral nerve fibers, in particular sensitive fibers which are involved in axo-axonal reflexes and secrete peptide mediators. In the skin, but not necessarily in other organs, MC gather near the site of injury to perform their roles. In the intermediate phase of the response MC decrease in number, probably in part because they degranulate and so become undetectable by the usual histochemical methods, in part because they die in response to hyperstimulation and to toxic substances (e.g. high concentrations of NO and other oxidants). The new, late increase in number and the late activity of MC can concur to drive definitive tissue repair by stimulating angiogenesis and inducing fibroblasts to secrete extracellular matrix and to differentiate into myofibroblasts to contract the collagen matrix. Mast cells promote the proliferation of fibroblasts, endothelial cells, and keratinocytes during the proliferative phase of wound healing. In the mouse, histamine and serotonin (which is also secreted by rodent MC) exert mitogenic effects on murine epidermal keratinocytes in situ and therefore influence re-epithelialization (Katayama et al., 1992; Maurer et al., 1997b).

Acknowledgement The authors are grateful to Prof. M. Serio for precious encouragement and support. Financial support was granted by the Italian Ministry of Education, University and Research (PRIN protocol n. 2007LTAJMA_003), the University of Florence and Tuscany region (project TRESOR). Thanks are given to Mr. T. and P. Venturi, and Mr. W. Calugi for technical assistance.

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Welker, P; Grabbe, J; Gibbs, B; Zuberbier, T; Henz, BM. Human mast cells produce and differentially express both soluble and membrane bound stem cell factor. Scand J Immunol, 1999 49, 495-500. Welker, P; Grabbe, J; Gibbs, B; Zuberbier, T; Henz, BM. Nerve growth factor-beta induces mast-cell marker expression during in vitro culture of human umbilical cord blood cells. Immunology, 2000 99, 418-426. Welker, P; Grabbe, J; Zuberbier, T; Grützkau, A; Henz, BM. GM-CSF downmodulates c-kit, Fc(epsilon)RI(alpha) and GM-CSF receptor expression as well as histamine and tryptase levels in cultured human mast cells. Arch Dermatol Res, 2001 293, 249-258. Weller, K; Foitzik, K; Paus, R; Syska, W; Maurer, M. Mast cells are required for normal healing of skin wounds in mice. FASEB J, 2006 20, 2366-2368. Woolhiser, MR; Okayama, Y; Gilfillan, AM; Metcalfe, DD. IgG-dependent activation of human mast cells following up-regulation of FcgammaRI by IFN-gamma. Eur J Immunol, 2001 31, 3298-3307. Worobec, AS; Semere, T; Nagata, H; Metcalfe, DD. Clinical correlates of the presence of the Asp816Val ckit mutation in the peripheral blood mononuclear cells of patients with mastocytosis. Cancer, 1998 83: 2120–2129. Wright, HV; Bailey, D; Kashyap, M; Kepley, CL; Drutskaya, MS; Nedospasov, SA; Ryan, JJ. IL-3-mediated TNF production is necessary for mast cell development. J Immunol, 2006 176: 2114-2121. Xia, HZ; Du, Z; Craig, S; Klisch, G; Noben-Trauth, N; Kochan, JP; Huff, TH; Irani, AM; Schwartz, LB. Effect of recombinant human IL-4 on tryptase, chymase, and Fc epsilon receptor type I expression in recombinant human stem cell factor-dependent fetal liver derived human mast cells. J Immunol, 1997 159: 2911–2921. Ying, S; Durham, SR; Corrigan, CJ; Hamid, Q; Kay, AB. Phenotype of cells expressing mRNA for TH2-type (interleukin 4 and interleukin 5) and TH1-type (interleukin 2 and interferon gamma) cytokines in bronchoalveolar lavage and bronchial biopsies from atopic asthmatic and normal control subjects. Am J Respir Cell Mol Biol, 1995 12, 477487. Yoshioka, M; Fukuishi, N; Kubo, Y; Yamanobe, H; Ohsaki, K; Kawasoe, Y; Murata, M; Ishizumi, A; Nishii, Y; Matsui, N; Akagi, M. Human cathelicidin CAP18/LL-37 changes mast cell function toward innate immunity. Biol Pharm Bull, 2008 31, 212-216. Yukami T, Hasegawa M, Matsushita Y, Fujita T, Matsushita T, Horikawa M, Komura K, Yanaba K, Hamaguchi Y, Nagaoka T, Ogawa F, Fujimoto M, Steeber DA, Tedder TF, Takehara K, Sato S. Endothelial selectins regulate skin wound healing in cooperation with L-selectin and ICAM-1. J Leukoc Biol 2007 82, 519-531. Zanetti; M. Cathelicidins, multifunctional peptides of the innate immunity. J Leukoc Biol, 2004 75, 39-48. Zehnder, JL; Galli, SJ. Mast-cell heparin demystified. Nature, 1999 400, 714-715. Zhang, S; Anderson, DF; Bradding, P; Coward, WR; Baddeley, SM; MacLeod, JD; McGill, JI; Church, MK; Holgate, ST; Roche, WR. Human mast cells express stem cell factor. J Pathol, 1998 186, 59-66.

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Zhao, W; Oskeritzian, CA; Pozez, AL; Schwartz, LB. Cytokine production by skin-derived mast cells: endogenous proteases are responsible for degradation of cytokines. J Immunol, 2005, 15 175, 2635-2642. Zhao, ZZ; Sugerman, PB; Walsh, LJ; Savane, NW. Expression of RANTES and CCR1 in oral lichen planus and association with mast cell migration. J Oral Pathol Med; 2002 31, 158-162. Zwartz, G; Chigaev, A; Foutz, T; Larson, RS; Posner, R; Sklar, LA. Relationship between molecular and cellular dissociation rates for VLA-4/VCAM-1 interaction in the absence of shear stress. Biophys J; 2004 86, 1243-1252.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 123-157

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Chapter IV

Role of Rho GTPases in Tumor Cell Migration and Metastasis Yong Tang and Daotai Nie† Department of Medical Microbiology, Immunology and Cell Biology, Southern Illinois University, School of Medicine and Simmons Cooper Cancer Institute, Springfield, IL, USA

1. Abstract

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Small GTPase Rho signaling pathways regulate the growth, motility, invasion and metastasis of a variety of cancer cells. Aberrant Rho signaling, as results from alterations in the levels of Rho GTPase proteins, the status of activation, or the abundance of effector proteins, endows cancer cells with elevated invasive and metastasis capability. Alterations of Rho signaling particularly impact the cytoskeleton, whose organization and reorganization underpin the motility of cancer cells during the invasive growth and metastasis process. Progress is being made to elucidate the underlying mechanisms by which Rho GTPases activate the downstream signaling effectors. Further investigations are required for development of novel tumor therapeutic strategies targeting the Rho GTPase signaling pathways to inhibit invasion and metastasis of cancer cells.

†

Correspondence to: Dr. Daotai Nie. Department of Medical Microbiology, Immunology and Cell Biology, Southern Illinois University School of Medicine and SimmonsCooper Cancer Institute, Springfield, IL 627949626. Phone: 217-545-9702; Fax: 217-545-3227; E-mail: [email protected]

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2. Introduction

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2.1. Cell Migration and Tumor Metastasis In metazoan organisms, migration of cells, either as individuals or as groups, is essential for morphogenesis during embryonic development (1). In the adult, it is crucial for tissue integrity maintenance and immune surveillance, in which leukocytes migrate from the lymph or blood circulation into the surrounding tissues to destroy exogenous microorganisms as well as infected somatic cells. Cell migration also contributes to several severe human diseases, including tumor formation and metastasis, atherosclerosis, osteoporosis, mental retardation and chronic inflammatory diseases, such as rheumatoid arthritis. In cancer, metastasis is the principal cause of death for most cancer patients, accounting for approximately 90% of all cancer related deaths (2-4). For dissemination and metastasis to occur, tumor cells must invade the tissue surrounding the primary tumor, intravasate into the lymphatic system or blood supply system, extravasate and attach to target tissue at the distant site, and then form macrometastases (5). Acquisition of cell motility has been implicated in the spreading of cancer cells (6) and is essential for metastasis (7). Since cancer mortality has been mainly associated with metastatic disease rather than with the primary tumor, better understanding of the factors leading to elevated motility of tumor cells is of vital importance for development of effective therapeutic approaches to inhibit invasion and metastasis of cancer cells. Cell migration is a coordinated multi-step process involving directed protrusion of the cell membrane, adhesion of the cell membrane to surrounding substrates, contraction of the cell body, and detachment of the cell at the trailing edge (8). The notion that Rho family GTPases could regulate cell migration originates from their ability to mediate the formation of specific actin containing structures (9, 10). Subsequent findings, which show the Rho family to regulate focal adhesion complex assembly, cell polarity, vesicle trafficking and gene transcription, reinforce its important roles relevant to cell migration. Overexpression and hyperactivation of Rho GTPases (particularly overexpression of RhoA and RhoC) or their corresponding signaling effector proteins ROCK I and ROCK II are found in a spectrum of human cancers and are often associated with more invasive and metastatic phenotypes (4, 11-19). Therefore, Rho GTPases are involved in cancer metastasis, proliferation, and progression which contribute substantially to patient morbidity and mortality. Herein, we first make a brief summarization of the general established conceptual framework of the Rho GTPases family, including their family members, their active-inactive regulators, and their biochemical and biological functions. Next, we focus on how Rho proteins and their signaling partners contribute to cell migration and address the alterations in GTPase signaling in cancer and their roles in metastasis and growth of cancer in general particularly in breast cancer. Finally therapeutic approaches targeting Rho GTPases are discussed as potential treatment for invasion and metastasis of cancer.

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2.2. Rho Subfamilies and Their Regulation

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Rho was first identified in 1985 due to its homology to Ras (Rho stands for Ras homologous) from the sea slug, Aplysia Californica (20). Later it was reported that Rho GTPases constitute a distinct subgroup within the superfamily of Ras-related small GTPases and are found in all eukaryotic cells. Five and seven Rho family members have been found in Drosophila melanogaster and Caenorhabditis elegans, respectively, most of which have been implicated in cell migration and morphogenesis. Plant-specific Rho subfamily, the Rop proteins, has also been reported (21). So far, twenty-two mammalian Rho family proteins have been identified and grouped into subfamilies: Rho-like (RhoA–C), RhoD (RhoD and Rif), RhoH/TTF, RhoBTB (RhoBTB1–3), Rac-like (Rac1–3, RhoG), Cdc42-like (Cdc42, TC10, TCL, Chp/Wrch-2, Wrch-1), and Rnd (Rnd1–2, Rnd3/RhoE) (22). Like all members of the Ras superfamily, Rho GTPases function as molecular switches, cycling between an active GTP-bound conformation and an inactive GDP-bound conformation (Figure 1). In the GTP-bound active form, they interact with downstream target proteins to trigger cellular response events. The switch between active GTP-bound conformation to inactive GDP-bound conformation is regulated by the opposing effects of guanine nucleotide exchange factors (GEFs), which promote the active GTP-bound state by facilitating the exchange of GDP by GTP, and by the GTPase-activating proteins (GAPs), which inactivate the Rho GTPases by promoting GTP hydrolysis. In addition, Rho proteins are also regulated by guanine nucleotide dissociation inhibitors (GDIs), which can inhibit both the exchange of GTP and the hydrolysis of bound GTP (23). In most cases, Rho proteins are posttranslationally modified at their C-termini by prenylation at a conserved cysteine, which is required for the attachment of Rho proteins to cell membranes (24).

Figure 1. Regulation of Rho family proteins. Solid arrows stand for activating signals; while red bars stand for inhibitory signals. GEFs, guanine nucleotide exchange factors; GAPs, GTPase-activating proteins; GDIs, guanine nucleotide dissociation inhibitors.

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2.2.1. GEFs As regulators of Rho GTPases, GEFs for Rho-like GTPases belong to a protein family sharing a Dbl-homology (DH) domain whose prototype is Dbl oncogene product (25). Originally discovered by its capability to induce focal complex formation and tumorigenicity when expressed in NIH-3T3 cells, Dbl proteins represent the GDP/GTP exchange protein (GEP) activity on all the Rho GTPases family members, including RhoA and Rac1 as well as Cdc42 (26). In addition to the DH domain, many of GEFs of Rho GTPases possess other domains that are commonly found in signaling molecules, such as Src homology 3 (SH3) domain and a diacylglycerol-binding zinc butterfly motif, suggesting their involvement in other cellular regulation events (25). A case in point here is a Rho GTPases GEF named Vav, whose prototype is the product of proto-oncogene Vav. Vav was first identified by a mutation that enables it to transform fibroblasts (27, 28). Vav contains a domain similar to the protooncogene Dbl. In addition, Vav contains a pleckstrin homology domain, a single SH2 domain, and two SH3 domains, which suggest that Vav can interact with several components of signal transduction events (28). Moreover, biochemical and genetic analyses in yeast have shown that the tyrosine phosphorylated form of Vav could promote Rac1 and other Rhofamily proteins to the active GTP-bound active conformation (29).

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2.2.2. GAPs The first GAP protein specific to the Rho family was purified based on its biochemical interaction with recombinant Rho. This protein, named p50Rho–GAP, possesses GAP activity toward Rho, Cdc42, and Rac in vitro (30, 31). Since then, additional proteins that exhibit GAP activity toward Rho GTPases have been identified, all of which share a GAP domain that bears no significant resemblance to Ras GAP. In addition to accelerating the hydrolysis of GTP Rho, GAPs may function as an effector of the Rho proteins to mediate other downstream functions in mammalian systems, such as cytoskeletal rearrangements (32). 2.2.3. GDIs Rho GDI was first characterized by its association with the GDP-bound form of the Rho family members, such as RhoA, Cdc42 and Rac to inhibit the dissociation of GDP (33, 34). Further studies demonstrated that Rho GDI is also associated weakly with the GTP-bound form of Rho, Cdc42, and Rac (35, 36). This weak interaction leads to inhibitory effects on the intrinsic and GAP-stimulated GTPase activity of the Rho GTPases. Thus, Rho GDI functions as a molecule capable of blocking the GTP-GDP binding cycling of GTPase through two approaches: at the GDP/GTP exchange step and/or at the GTP hydrolytic step. In addition, a significant role of Rho GDIs during Rho GTPases circulation lies in regulation of the translocation of the Rho GTPases between membranes and the cytoplasm. In resting cells, the Rho proteins exist as a complex with Rho GDIs in the cytosol to inhibit their GTP/GDP exchange ratio. During cellular activation events, Rho proteins are released from the GDI and translocated to the membranes (37).

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3. Upstream Signaling Pathways of Rho GTPases Rho family proteins are involved in translating the cellular signals sent from plasma membrane receptors into the assembly and organization of the actin cytoskeleton. In fibroblasts, diverse extracellular stimuli have been shown to result in Rho GTPase activation. Addition of lysophosphatidic acid (LPA) to quiescent fibroblasts induces the assembly of focal adhesions and actin stress fibers, which can be blocked with C3 transferase, a bacterial coenzyme that ribosylates ADP and inactivates Rho proteins (38). A Rac-regulated signaling pathway linking growth factor receptors to the polymerization of actin at the plasma membrane has also been established. Growth factors, such as platelet-derived growth factor (PDGF), insulin, and bombesin, stimulate polymerization of actin at the plasma membrane to induce lamellipodia formation and membrane ruffling, which can be inhibited by the dominant-negative mutant of Rac, RacN17 (39, 40). Moreover, the activation of Cdc42 by bradykinin promotes the formation of peripheral actin microspikes and filopodia, with the subsequent formation of lamellipodia, which can be inhibited by a dominant-negative mutant of Cdc42, Cdc42N17 (41). Our recently published data indicated that functional TP(s), the G protein–coupled receptor (GPCR) for Thromboxane A2 (TxA2), is expressed in prostate tumor cells as well as in tumor specimens. Further, activation of this receptor by TxA2 mimetics (U46619) has profound effects on tumor cell cytoskeleton organization and causes contraction through the small GTPase RhoA. The constitutively active mutant of RhoA was sufficient to cause cell contraction, whereas the dominant-negative mutant of RhoA blocked cell contraction induced by U46619. The data affirm that activation of RhoA is required for TxA2-TP to induce cell contraction and that RhoA may serve as intermediary for TxA2-TP signaling axis to regulate tumor cell motility (42). Since the bradykinin, LPA, bombesin and TP receptors belong to the seven-transmembrane-domain heterotrimeric G protein coupled receptor family, the trimeric G proteins are likely to play a role in the activation of the respective GTPases. Several lines of evidence suggest the involvement of phosphoinositide 3 kinase (PI3 kinase) in PDGF and insulin induced lamellipodia formation and membrane ruffling. PDGF could stimulate an increase in GTP-Rac by enhancing GEF activity in a PI3 kinase activation dependent manner (43). Furthermore, treatment of fibroblasts with PI3 kinase inhibitor wortmannin inhibits Rho and Rac mediated membrane ruffling induced by PDGF, epidermal growth factor (EGF), and insulin or insulin-like growth factor 1 (IGF-1) (40, 44, 45). These data suggest that PI3 kinase acts on the upstream of Rac to induce membrane ruffling when exposed to extracellular growth stimuli. In contrast with these findings, studies showed that activities of p42/44 MAPK, p38 MAPK, protein kinase G phosphatidylinositol 3-kinase, or pertussis toxin-sensitive Gi subunit were not required for U46619 to induce cell cytoskeleton reorganization and cell contraction through the small GTPase RhoA by binding to TP receptors (42). Another upstream signaling pathway of Rho family proteins is the tyrosine phosphorylation of the exchange factor Vav, which is crucial for its ability to activate members of the Rho family (29, 46). As described previously, Vav is guanine nucleotide exchange factor for Rho, Rac, and CDC42 family. When exposed to Lck, a member of the Src family, Vav increases the GDP/GTP exchange activity. Furthermore, coexpression of Lck

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with Vav enhances the Vav-transforming activity and its capability to induce Jun N-terminal kinase (JNK) activation. The mechanism by which LPA activates Rho appears also attributable to the involvement of tyrosine kinase because the LPA-induced stress fiber formation can be abolished by tyrphostin, a tyrosine kinase inhibitor (47, 48). In summary, extracellular stimuli, such as LPA, PDGF, EGF, and insulin, trigger Rhofamily GTPases mediated assembly and organization of the actin cytoskeleton in a PI3 kinase dependent manner. Moreover, our recent data indicate that the induction of cell cytoskeleton reorganization and cell contraction by Thromboxane A2 receptor relies on small GTPase RhoA, but in a PI3 kinase independent manner. In addition, tyrosine phosphorylation is also involved in the activation of Rho. Further elucidation of upstream signaling molecules will provide a more thorough insight into the signaling pathways that lead to the activation of the Rho-like GTPases in response to extracellular stimuli.

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4. Biochemical and Biological Functions of Rho GTPases in Cell Migration Eukaryotic cellular morphology and motility are underlain by the organization and reorganization of cytoskeleton. Cell migration and axonal path finding during development, immune cell patrolling in normal adults, phagocytosis, as well as pathological processes such as metastasis, all rely on coordinated regulation of the actin network to produce directed cell movement. The actin cytoskeleton is part of the cytoskeleton (the internal framework of a cell) composed of actin filaments and many specialized actin-binding proteins (48, 49). Although actin filaments are created by the simple polymerization of actin monomers, regulation of the dynamics of the filament network requires participation of many interacting protein members of the Src family. Vav increases the GDP/GTP exchange activity. Furthermore, coexpression of Lck with Vav enhances the Vav-transforming activity and its capability to induce JNK activation. The mechanism by which LPA activates Rho appears also to be attributable to the involvement of tyrosine and is regulated by numerous upstream signaling pathways. This regulation of actin polymerization, for the most part, is orchestrated by Rho GTPases. Separate members of Rho GTPases proteins regulate the organization of the actin cytoskeleton in their own manners. Rho activation in fibroblasts results in the assembly of contractile actin/myosin filaments, the formation of stress fibers, and the clustering of integrins involved in the formation of focal adhesion complexes. Activation of Rac facilitates actin polymerization at the cell periphery to generate protrusive actin-rich lamellipodia and membrane ruffling. Activation of Cdc42 results in actin polymerization to form peripheral actin microspikes and filopodia. In addition, cellular signaling cross-talk between the members of the Rho GTPases proteins has been observed. For example, Cdc42 is a potent activator of Rac, leading to observations that filopodial extensions are usually accompanied with lamellipodial protrusions (38, 39, 41, 47) (Table 1).

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Table 1. Roles of characterized Rho protein Rho Protein RhoA

Rac1

Cdc42

• • • • • • • • • • •

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•

Function Promotes the rearrangement of cell cytoskeleton Initiates the assembly of focal adhesion Induces the contraction of cell body Stimulates the movement of the rear of the cell to induce forward migration of cell body and nucleus Promotes the detachment of the rear of the cell from the ecm Promotes actin-mediated formation of the lamellipodia Stimulates the attachment of lamellipodia to ecm at the front of the cell body Allows for new adhesion contacts to ecm Required for cell polarization Promotes directionality of cell movement Plays role in the rate of cell migration, by enhancing rac-mediated membrane extension Causes the formation of filopodia

4.1.1. Rho and Cytoskeletion Reorganization A number of proteins have been identified as targets of Rho by the yeast two-hybrid selection system and affinity chromatography purification. These targets include Rhokinase/ROK/ROCK, myosin-binding subunit (MBS) of myosin phosphatase, protein kinase N (PKN)/PRK1, p140mDia, rhophilin, rhotekin and citron and so on. Most of them are involved in Rho mediated cytoskeletal rearrangements (Figure 2). The involvement of Rho in cytoskeleton reorganization is firstly implicated by the observations that the stress fibers and focal adhesion complexes were induced in fibroblasts with ectopic expression of an activated mutant of Rho, RhoV14 (38). The discoveries and characterization of numerous proteins that bind Rho in a GTP-dependent manner have shed light on the molecular mechanisms by which Rho affects the cytoskeleton (Figure 2). Among these proteins, serine/threonine kinase, ROK alpha (also known as Rho kinase) and its close relative (52% homology), p160Rho kinase (also known as ROCKII or ROKβ) have been extensively studied to date (50-54). The kinase activity of p160Rho kinase is enhanced (though not by much) after binding to Rho-GTP. Clues to their functions in the reorganization of actin filaments and in myosin contractility come from two lines of evidence. The expression of full-length ROKα and its amino-terminal portion induces stress fibers and focal adhesion complexes formation (50), which is abolished by the expression of a kinase-dead mutant of the protein (51). Furthermore, two substrates for the p160Rho kinase have been identified, the MBS of myosin light-chain (MLC) phosphatase and myosin light chain itself (Figure 3).

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Rho

PIP5-Kinase

Rhophilin

PKN

Rhotekin

PRK2

citron

Rho kinase

MBS

p140mDia

Actin polymerization Ptdlns(4,5)P2

Actin-binding proteins recruitment and Actin rearrangement

LIMK

Cofilin

Actin filament depolymerization

MLC Phosphatase

MLC-P

MLC

Stress fiber formation Focal adhesion formation Smooth muscle contraction Cell contraction and mobility

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Figure 2. Mammalian downstream targets of Rho signaling pathway for cytoskeleton reorganization. Solid arrows stand for activating signals; while red bars stand for inhibitory signals. Abbreviations: PIP5-kinase, Phosphatidylinositol 4-phosphate 5-kinase; Ptdlns(4,5)P2, phosphatidylinositol 4,5-bisphosphate (PIP2); LIMK, LIM kinase; MLC-P, phosphorylated myosin II regulatory light chain; MLC phosphatase, myosin light chain phosphatase; MBS, myosin-binding Subunit; PKN, protein kinase N; PRK2, protein kinase C-like 2.

It was subsequently shown that phosphorylation of MBS downregulates MLC phosphatase activity, resulting in an accumulation of the phosphorylated form of MLC (55). Phosphorylation of MLC induces a conformational change in myosin, thereby increasing its binding to actin filaments to drive the formation of stress fibers and focal adhesions (56, 57). Together, phosphorylation of these two substrates would be expected to lead to an increase in myosin light chain phosphorylation, myosin filament assembly, F-actin bundling, and stress fiber formation. Both Rho and Rac have been shown to stimulate the synthesis of phosphatidylinositol 4,5-bisphosphate (PIP2), the product of phosphatidylinositol 4-phosphate 5-kinase (PIP 5kinase) (58, 59). The observation that PIP2 can regulate the interactions of a number of actinbinding proteins, including profilin, α-actinin, gelsolin, and p39CapZ in vitro (60), led to the hypothesis that Rho-stimulated PIP2 synthesis may induce actin rearrangements. A case in point here is the protein vinculin. Vinculin is a membrane-cytoskeletal protein in focal adhesion plaques that is involved in linkage of integrin adhesion molecules to the actin cytoskeleton. More specifically, the amino-terminus of vinculin binds to talin which, in turn, binds to β-integrins, and the carboxy-terminus binds to actin, phospholipids and paxillin forming homodimers. The binding of vinculin to talin and actin is regulated by PIP2 and inhibited by acidic phospholipids (61). Furthermore, injection of anti-PIP2 antibodies into fibroblasts inhibits LPA-induced formation of stress fiber and focal adhesions, suggesting a crucial role for PIP2 in focal adhesion and stress fiber assembly (62).

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Rho-GTP Active GDP

PI(4)P 5-K Rho-kinase (ROCK)

Myosin Phosphatase

Inactive MLC

Active

MBS p MLC

Active

•Contraction

Myosin Phosphatase

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•Actin-Myosin interaction

p Inactive MBS

Figure 3. Model for regulation of myosin light chain phosphorylation by Rho-kinase and myosin phosphatase. Solid arrows stand for activating signals; while red bars stand for inhibitory signals. Abbreviations: MLC, myosin light chain; MBS, myosin-binding Subunit; PI(4)P 5-K, Phosphatidylinositol 4Phosphate 5-Kinase.

Recently, a downstream effector of the Rho protein, p140mDia, was shown to selectively interact with mammalian Rho in a GTP-dependent manner and also interact with profilin (63). The NH2-terminal portion of p140mDia ensures that p140mDia binds selectively to the GTP-bound form of Rho. p140mDia also contains repetitive polyproline stretches that can bind profilin. Thus, p140mDia provides a direct molecular linkage between Rho and profilin. RhoA, p140mDia and profilin are colocalized in the spreading lamellae of cultured fibroblasts and also can be recruited around phagocytic cups induced by fibronectin (FN)coated beads. Overexpression of p140mDia in COS cells enhances actin filament assembly, while colocalization of RhoA, p140mDia and profilin is abolished after Rho is inactivated by microinjection of botulinum C3 exoenzyme. Based on these findings, a model was proposed in which Rho regulates actin polymerization by targeting profiling to a specific site beneath the plasma membrane via p140mDia, which results in a locally increased profilin concentration (63, 64). The functions and subcellular localization of the three molecules

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suggest a role for p140mDia in mediating the effects of Rho on actin reorganization beneath the dynamic plasma membranes in mammalian cells. In addition to Rho kinase, MBS, and p140mDia, other mammalian proteins have been identified as potential Rho targets (Figure 2). PKN is the first identified serine/threonine protein kinase that can bind to and be activated by a small GTPase Rho. It is a protein kinase that has a catalytic domain homologous to protein kinase C (PKC) family members and a unique regulatory region containing antiparallel coiled-coil (ACC) domains previously called the “CZ region” (charged amino acid and Leu-zipper-like sequence) or “HR1” (65). There are at least three different isoforms of PKN (PKN-α/PAK-1/PRK-1, PKN-β, and PRK2/PAK2/PKN-γ) in mammals, each of which shows different enzymological properties, tissue distributions, and varied functions. GTP-bound Rho interacts with the first leucine zipper-like motif in the NH2-terminal portion and activates the catalytic activity of PKN. The expression of a kinase-deficient form of PRK2 disrupts actin stress fibers, implicating PRK2 in actin cytoskeleton reorganization. Whereas rophilin and Rhotekin share homology with both MBS and PKN/PRK1 in their Rho binding domain (66, 67), overall domain structure of citron is similar in sequence to Rho-kinase. However, citron lacks a kinase domain (68). Citron-kinase is localized at the cleavage furrow and mid-body during cytokinesis. Citron is implicated in cytokinesis (69). So far, however, there is still no well established evidence to clarify the role of Rho-mediated cytoskeletal rearrangements in cytokinesis. 4.1.2. RAC, CDC42 and Cytoskeleton Reorganization In fibroblasts, Rac has been shown to be a crucial regulator in the reorganization of the actin cytoskeleton in growth factor-induced membrane ruffling (39). Later, a role for yet another member of the Rho subfamily, Cdc42, in actin remodeling was established. Both Rac and Cdc42 have also been shown to induce the assembly of multimolecular focal adhesion complexes (FACs) at the plasma membrane of fibroblasts (47). These focal complexes display morphologies like punctate spots of vinculin and phosphotyrosine around the leading edge of the lamellipodia and at the tips of filopodia, which are distinct from the Rho regulated focal adhesions, in spite of the similarity in their protein components. To date, several potential targets of Cdc42 and Rac have been identified, including p21-activated kinases (PAKs), WASP/N-WASP, IQGAP1, MRCK, Por1, p140Sra-1, and Posh (Figure 4), which help to elucidate the molecular basis for the involvement of Rac and Cdc42 in cytoskeleton organization. Although Rac and Cdc42 each have seven or eight known target proteins, some are common to both (Figure 4). For example, the first Rac target identified, p65PAK, also interacts with Cdc42. To date, at least three isoforms of p21-activated kinase (PAK1–3) have been identified (70-74), all of which bind Rac and Cdc42 in a GTP-dependent manner. PAKs comprise an NH2-terminal regulatory domain and a COOH-terminal catalytic domain (70). GTPase binding disrupts an inhibitory interaction between the PAK kinase domain and the PAK auto-inhibitory domain, which results in PAK phosphorylation and activation (75, 76).

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Rho GTPases in Cell Migration Rac

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Actin polymerization filopodia formation

Actin filament depolymerization and Stress fibers disassociation

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Figure 4. Mammalian targets of Rac and Cdc42. Solid arrows stand for activating signals; while red bars stand for inhibitory signals. Abbreviations: PIP5-kinase, Phosphatidylinositol 4-phosphate 5-kinase; PI3Kinase, phosphatidylinositol-3-kinase; PAK, p21-activated kinase; LIMK, LIM kinase; MLK3, mixed lineage kinase-3; MEKK4, mitogen-activated protein kinase kinase kinase 4; MRCKs, myotonic dystrophy kinase-related Cdc42 binding kinases; WASp, Wiskott-Aldrich syndrome protein; N-WASp, neural WASp; ACK-1, activated Cdc42-associated kinase-1;LIMK, LIM kinase.

In addition, all PAK proteins identified to date share a conserved domain of approximately 20 residues that is responsible for interaction with Rac or Cdc42 and is referred to as CRIB (Cdc42/Rac interactive binding) site. Several other mammalian proteins of potential targets for Cdc42 and Rac, like MSE55, MLK2/3, and WASP, share this motif (77). PAKs have been shown to be the key components of pathways that regulate cell morphology, including formation of lamellipodia and disassembly of stress fibers and focal adhesions in response to stimuli that activate Rac and Cdc42 and promote cell migration (78, 79). Overexpression of constitutively activated mutant PAK1 triggers dissolution of actin stress fiber and focal adhesions, accompanied by increased cell polarization, membrane protrusions and cell motility (80-82). PAKs are also upstream elements in the JNK and p38 kinase pathways that control gene expression (74). The N terminus of PAK1 contains multiple proline-rich sites that bind to Src homology (SH) 3 domain-containing proteins, including Pix, a GEF for Rac (83), adaptor proteins NCK (84) and Grb2 (85). The binding of PIX and NCK to PAK seems required for the localization of PAK to focal adhesions. Despite the fact that Rac and Cdc42 trigger morphologically distinct lamellipodia and filopodia at the plasma membrane, the formation of both protrusions rely on the Arp2/3 complex to initiate peripheral actin polymerization. Arp2/3 complex is a heptameric, actinnucleation machine associating with the sides and perhaps the ends of existing actin filaments to initiate new branched filaments formation (86). Both Rac and Cdc42 activate Arp2/3 indirectly through members of the Wiskott-Aldrich syndrome protein (WASP) family. NWASP, or the closely related hemopoietic-specific WASP, binds to Cdc42 in a GTPdependent manner. The binding generates an intra-molecular, auto-inhibitory interaction and

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exposes a C-terminal Arp2/3 binding/activation site, which contributes to Cdc42 mediated cytoskeletal rearrangements and actin clustering (87). Activation of Arp2/3 by Rac is mediated by WASP family Verprolin-homologous proteins (WAVE), which although structurally related to N-WASP, do not interact directly with the GTPase (88, 89). Another protein with a potential role in cytoskeletal organization is IQGAP (90-92). The IQGAP1 gene was originally isolated as a member of the Ras GAP family (93). At least two isoforms of IQGAP have been identified, IQGAP1 and IQGAP2. Both isoforms directly interact with GTP-bound Cdc42 and Rac, but not with the GDP-bound forms. IQGAPs contain some interesting motifs found in signaling molecules, such as a WW domain, an SH3-binding domain, and a calmodulin-binding domain. However unlike PAKs and WASP, IQGAPs do not contain a CRIB site. IQGAP interacts with both Rac and Cdc42 and localizes to membrane ruffles. In addition, both IQGAP1 and IQGAP2 have been shown to interact directly with actin filaments. It is not known if they play any role in filopodia formation, but there are some data linking them to the assembly of actin filaments during cytokinesis (94). In addition to its role in actin filament organization, IQGAP1 appears to play a pivotal role in cell-cell adhesion through the cadherin-catenins pathway (95). As described previously, cytoskeleton rearrangement is potentially linked to phospholipid metabolism. In fact, both Rac and Rho bind to PIP5 kinases and recruit them to the plasma membrane (96), while ADP-ribosylation factor 6 (Arf6) acts downstream of Rac to activate PIP5 kinases (97). It is likely that PIP5 kinase may be the responsible kinase mediating Rac-stimulated PIP2 generation. Although a number of potential targets or effectors of Rho family members have been identified and characterized, there is still not sufficient evidence for the physiological relevance of these proteins in Rho-mediated cytoskeletal remodeling, which is likely to be a focus of attention in the future. 4.1.3. Other Functions of Rho Family of GTPases Rho GTPases have been reported to regulate various other cell functions, including membrane trafficking, transcriptional activation, cell growth control and development in mammals and lower eukaryotes (9, 10). Because this review focuses on the role of these GTPases proteins on cytoskeletons rearrangement and cell adhesions involved in cell migration, the above functions regulated by the Rho family GTPases are not described here.

4.2. Rho GTPases in Cell Migration Cell migration can be mechanistically divided into separate steps: cell polarization, lamellipodium protrusion and adhesion formation, followed by cell body contraction, and tail detachment (8) (Figure 5). The initiation of cell migration requires polarization and generation of protrusions in the direction of migration. These protrusions can be large, broad lamellipodia or spike-like filopodia, which are usually driven by actin polymerization. To achieve efficient migration, these protrusions must be stabilized by anchoring to the extracellular matrix (ECM) or adjacent cells via specific transmembrane receptors of the integrin protein family linked to the actin cytoskeleton. This in itself is not sufficient, however, because cell contractility is

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required to allow the body and rear of the cell to follow the extending front (98). As the front protrusions (mostly in form of lamellipodia or filopodia) advances, the cell becomes stretched and elongated. Subsequent detachment of the trailing edge requires acto-myosin-driven tension that results in release of adhesions within the trailing edge and allows cell body to advance whereupon the cell becomes more rounded (symmetrical) (99, 100). All aspects of cellular motility and invasion, including cellular polarity, cytoskeletal organization, and transduction of signals from the outside environment involved in adhesion formation are controlled through the interplays between the Rho-GTPases. Herein, we will discuss the evidence linking specific Rho proteins to each of these steps and the ways in which they contribute to cell movement. 3. Cell body contraction (Rho) through actin stress fibers MLC-P

1. Protrusion (Lamellipodium) extension (Rac) through actin filaments

4. Rear retraction (Rho) through actin stress fibers MLC-P

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Ta

il r

e tr ac t

Mic rotubules i on

Direction of Movement

2. New adhesions formation (Rac)

Figure 5. A model for the steps of cell migration. MTOC, microtubule-organizing center. MLC-P, phosphorylated myosin II regulatory light chain.

4.2.1. Cell Polarization: A Keystone of Cell Migration The ability of a cell to move depends on an asymmetrical organization of intracellular activities, which means the molecular processes at the front and the back of a moving cell must be different. Triggered by extracellular stimuli such as chemoattractants, the reorientation of the intracellular signaling machinery starts and drive the cells to a given direction. Establishing and maintaining cell polarity in response to extracellular stimuli appear to be mediated by a set of interlinked positive feedback loops involving Rho family GTPases, PI3Ks, integrins, microtubules, and vesicular transport. Among the Rho family GTPases involved in cell polarity regulation, Cdc42 is a master regulator in eukaryotic organisms ranging from yeast to humans. Genetic analysis of budding

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yeast provides the first piece of evidence for the involvement of Cdc42 in cell polarity. During the cell cycle, yeast cells adopt alternative states of polarized growth, ranging from tightly focused apical growth to non-focused isotropic growth. In the absence of Cdc42, Saccharomyces cerevisiae fail to establish focused apical growth and, as a consequence, cells expand isotropically (101). One way in which Cdc42 influences polarity is to direct where lamellipodia will form (102). Cdc42 can also regulate cell polarity by directing the localization of the microtubule-organizing center (MTOC) and Golgi apparatus in front of the nucleus, oriented toward the direction of cell migration. In this way, the direction of MTOC orientation at the front of the leading edge may contribute to cell migration by facilitating delivery of Golgi derived vesicles to the leading edge, wherein microtubules bundle into the lamellipodium (103). The effects of Cdc42 on MTOC position seem to be mainly exerted through its downstream effecters, such as kinase PAK1. PAK1 itself can mediate Cdc42 activation and therefore positive feedback loops between Cdc42 and PAK1 ensure the maintenance of high Cdc42 activity at the leading edge (104). As targets of Rac, PIP3 and phosphatidylinositol (3,4) bisphosphate (PI(3,4)P2) are key signaling molecules that contribute to Cdc42 accumulation at the front of moving cells. This mediation involves both localized accumulation and activation of PI3Ks, which produce PIP3/PI (3,4) P2, and phosphatase PTEN, which removes them. To be more specific, PI3Ks rapidly accumulate at the leading edge of cells, whereas PTEN becomes restricted to the sides and the rear (105, 106). Altered PI3K or PTEN activity significantly compromises the ability of the cell to directionally move up a chemoattractant gradient. Together these results indicate that there exist positive feedback loops between Cdc42, PI3K products, and PTEN that coordinate together to ensure the initiation and maintenance of the polarity of migrating cells. 4.2.2. The Protrusive Machinery Actin filaments are polarized with fast growing plus (+) or barbed ends and slow growing minus (-) or pointed ends. This inherent polarity is used to drive membrane protrusion. However, the organization of filaments depends on the types of protrusion: as for lamellipodia, actin filaments form a branching dendritic network, whereas as for filopodia they are organized into long parallel bundles (107). The distinct molecular attributes of lamellipodia and filopodia account for their capacity to perform distinct functions. Biophysically, the dendritic organization of lamellipodia provides a tight brush-like structure that is able to push along a broad range of plasma membrane (108). Through localization of activated Arp2/3 complex the lamellipodium can grow in a particular direction. In contrast, filopodia are particularly well designed to serve as sensors to explore the local environment with parallel bundles. Rac is required for lamellipodium extension induced by extracellular stimuli, such as growth factors, cytokines and extracellular matrix components (109, 110). Reports about cells derived from Rac1- and Rac2-null mice support a pivotal role for Rac in cell migration. Effects of constitutively active Rac1 on cell migration vary in terms of cell types, stimuli, expression levels and time courses of expression. For example, constitutively active Rac1 inhibits macrophage migration induced by growth factors, because the lamellipodia extend all around the cells and fail to polarize.

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How does activated Rac coordinate lamellipodium extension? Several Rac target proteins are likely to be involved in this process. As shown in Figure 4, Rac stimulates new actin polymerization relying on the Arp2/3 complex, which binds to pre-existing filaments. Activation of Arp2/3 complex by Rac is carried out by its target IRSp53 (111). After activation by Rac, IRSp53 interacts through its SH3 domain with WAVE, which finally binds to and activates the Arp2/3 complex. IRSp53 can also bind to Cdc42 with a separate domain (112). Therefore, IRSp53 can serve as a direct link between Cdc42 and Rac, which sheds light on the observation that Cdc42 can induce Rac-involved lamellipodium formation. IRSp53 can also bind to a Rho target, Dia1 (113), which might underlie the capability of Rho to facilitate lamellipodium extension. In addition to activating the Arp2/3 complex, Rac can promote the uncapping of actin filaments at the plasma membrane to facilitate actin polymerization. In platelets, Rac acts via a PIP5-kinase to generate PIP2, which then binds to capping proteins and removes them from the barbed ends of actin filaments for actin polymerization (114, 115). Apart from facilitating actin polymerization, Rac has been reported to downregulate the rate of actin depolymerization. Through its downstream target PAKs, Rac is capable of stimulating the activity of LIM-kinase (116, 117). As well as inducing actin polymerization, Rac may affect the rate of actin depolymerization. LIM-kinase can catalyze the phosphorylation and inactivation of cofilin, an actin-regulatory protein that will promote Factin depolymerization. This suggests that Rac would inhibit cofilin-induced depolymerization. 4.2.3. Formation and Turnover of New Cell-Substrate Adhesions For migration to occur, a protrusion must form and attach to the extracellular matrix by specific receptors of the integrin protein family, forming sites of cell–substrate focal complexes, i.e., focal adhesions (FA) and focal contacts (FC). Adhesion assembly in migrating cells begins with small-scale clustering, dependent on the multivalent nature of ECM to which the cell is adhering. The speed of cell migration is dependent on ECM composition, which is decisive for the relative activation levels of Rho, Rac and Cdc42 (118, 119). Small focal complex structures can be observed at the leading edge and are localized in the lamellipodia of most migrating cells. Early studies reported that cells plated on ECM proteins rapidly developed extended filopodia and lamellipodia, indicative of Cdc42 and Rac activity, respectively (120). These protrusive structures contain focal complexes, which are localized clusters of integrins, and cytoskeletal and signaling proteins that are smaller than, but similar in composition to focal adhesions. Therefore, the continuous formation of new interactions between ECM and integrins at the leading edge of cells help to maintain the level of active Rac there, which means there exists a positive feedback loop to allow cells to carry on migration (109, 121). In this way, the continuous crosstalk between integrins and Rac allows cells to respond to changing ECM composition. These adhesions stabilize the lamellipodium by attaching to ECM (8). To the contrary, if the formation of these adhesions is interrupted, cell protrusions, i.e., lamellipodia and filopodia are retracted and therefore the efficiency of migration is markedly reduced (122). The formation of small focal complexes involves the aggregation of integrins triggering the activation of RhoA-ROCK pathway, which promotes the formation of contractile acto-

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myosin bundles and generation of the larger focal adhesions (123). The formation of small focal adhesions at the leading edge of protrusions is thought to be critical for cell migration in that they function as anchorage sites for generation of acto-myosin tension. In contrast, the larger focal contacts seem to inhibit cell migration by forming stronger anchorage (124-126). The formation of focal complexes is an integral step for lamellipodia attachment to the ECM of most migrating cells. However, cells with a high level of integrin-mediated adhesion are typically either nonmigratory or move very slowly due to the intense strength of attachment to the ECM, which is correlated with high levels of Rho activity (126). In addition to being involved in focal complex formation in lamellipodia, Rac can induce focal complex turnover directly through PAK (127). PAK interacts with a complex of exchange factors, such as paxillin, to localize focal adhesion complexes. Paxillin binds to many proteins, such as vinculin and actopaxin, which are involved in inducing changes in the organization of the actin cytoskeleton during cell migration. 4.2.4. Cell Body Contraction Cell body contraction is dependent on actomyosin contractility and the force transmitted to sites of adhesion derives from the interaction of myosin II with actin filaments that attach to these sites. Myosin II activity is regulated by MLC phosphorylation. The phosphorylation of MLS results in an increased contractility and transmission of tension to sites of adhesion, which is either positively regulated by ROCKs or negatively regulated by MLC phosphatase, which in turn is phosphorylated and inhibited by ROCK. MLC phosphorylation is also regulated by MLC kinase (MLCK), which is regulated by intracellular calcium concentration as well as stimulated by the ERK MAPKs (Figure 3). In addition to mediating MLC phosphorylation, Rho exerts other effects on the actin cytoskeleton organization that are related to cell migration (Figure 2). Cofilin is an actin binding protein that is essential for the depolymerization of actin filaments. Either through ROCK or LIM-kinases (LIMK), Rho abolishes the actin-binding activity of cofilin by inducing cofilin phosphorylation, thus enhancing the polymerization of actin filaments (128). As described previously, another way for Rho to induce F-actin accumulation is via PIP 5kinases. Rho enlists Dia proteins for the actin cytoskeleton organization. Through interacting with either Src kinases or IRSp53 (103), Dia proteins induce actin polymerization and stress fibers formation (129). 4.2.5. Adhesion Disassembly and Retraction at the Rear At the rear of migrating cells, adhesions must disassemble to ensure the fulfillment of cell movement. In most cases, tail detachment is the rate-limiting step during cell migration (125). The mechanisms regulating tail detachment depend on the cell types and their corresponding adhesion strength to the extracellular matrix. On one hand, in slowly moving cells, tail detachment depends on calpain, which is a member of calcium-dependent, nonlysosomal cysteine proteases family. Calpain has been implicated as a critical calciumdependent regulator of the actin cytoskeleton and cell migration by degrading focal adhesion components at the rear of cells. Inhibition of calpain reduces cell migration rates and invasiveness (125, 130). On the other hand, in cells such as fibroblasts that have relatively

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large focal adhesion complexes, Rho assumes a role to reduce adhesion and promote tail detachment. In fibroblasts, the tight rearmost adhesions often tether the cell strongly to ECM. Strong tension forces exerted across the cells on the rear adhesions is required to physically break the link between integrin and the actin cytoskeleton. Several lines of evidence support a role for myosin II and Rho in this event. Dictyostelium cells deficient in myosin II or its regulator PAK alpha show impaired retraction (131) and similar phenotypes can be observed in monocytes or neutrophils by blocking myosin II assembly with Rho or Rho kinase inhibitors (132, 133). FAK, Src, and the other regulators that can induce focal adhesion disassembly also contribute to tail detachment (134). Rho mediated adhesion disassembly and retraction at the rear is a critical step to accomplish cell migration. However, the migration ability of cells with a constant high level of activated Rho-ROCK pathway is comprised. Out recent findings showed that on one hand, blockade of the Thromboxane A2 receptor (TP) by its antagonists (SQ29548 or PTA2) was found inhibitory for cell migration, suggesting that TP activation is required for tumor cells (PC-3) to migrate; on the other hand, sustained activation of TP with high levels of U46619 significantly compromised tumor cell motility (42). Taken together, the data suggest that Rho-ROCK pathway activation has to be controlled in a spatial and temporal manner in its regulation of cell migration.

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5. Altered Rho GTPase Signaling Pathways in Breast Cancer Cells Metastases, not the primary tumor, are the cause of death in 90% of cancer patients because of the difficulties in detecting and eradicating numerous small secondary tumors disseminated throughout the body, particularly those with breast cancer. Metastases from the breast commonly occur in the sentinal and axillary lymph nodes, lungs, spinal cord, brain and bones (135). To successfully metastasize, tumor cells must perform an ordered series of steps that constitute the metastatic cascade. This includes invading the tissue surrounding the primary tumor, intravasating into the lymphatic or the blood supply system, avoiding host immune defenses and surviving in circulation, extravasating and arresting at the distant site for final macrometastasis formation. Two key components in obtaining metastatic competence are the acquisition of a motile and invasive phenotype, both of which are mainly controlled by members of the Ras-superfamily of small GTP-binding proteins. In following part of this review, we will discuss evidence for altered Rho GTPase signaling events in breast cancer and their role in tumor development, metastasis, and invasiveness.

5.1. Overexpression of Rho Proteins In Human Breast Tumors Approximately 30% of human tumors carry an identifiable specific Ras mutation, accounting for its protein level overexpression or constitutive activation (135), while only approximately 5% of breast tumors harbor an activating Ras (136). In contrast to Ras, no mutation in any of the Rho GTPases has been identified in breast or other human cancers.

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Rather, these GTPases, particularly RhoA and RhoC, are often either significantly overexpressed or hyperactive in a variety of tumors, especially during development to more a invasive and malignant stage (4, 11-17). In vitro, overexpression of RhoA results in transformation of mouse fibroblasts (137), supporting the notion that increased Rho A is a crucial event for malignant transformation and development of human tumor cells. Comparing the expression of proteins of the Rho family (RhoA, Rac, Cdc42) in tumorigenic tissue and the corresponding normal tissue obtained from the same patient, it was found that Rho GTPases, in particular RhoA, are overexpressed in different types of human tumors, including colon, breast and lung tumors (2). The higher levels of RhoA protein are correlated with more advanced grades of breast carcinoma (2). Microarray analysis identified that RhoC is one of the metastasis-inducing genes in an experimental mouse melanoma model) (138). Increased RhoC expression was found in inflammatory breast cancer (IBC) (2) and in invasive ductal carcinomas (32%) (4), particularly those with distant metastasis. Inflammatory breast cancer is a phenotypically distinct form of localized breast cancer due to its high metastatic capability. In an attempt to identify specific genes responsible for the unique IBC phenotype, Merajver’s laboratory identified RhoC GTPase as being overexpressed in the SUM149 IBC cell line. Further studies indicate that RhoC seems to be a marker of metastatic potential in breast cancer, since 47% of the invasive ductal carcinomas that develop metastases express RhoC, in contrast to 12% of the invasive carcinomas without metastases (4). Subsequently, RhoC overexpression has been found to be a potential prognostic marker for tumors with the ability and propensity to metastasize. These results are in concordance with previous data showing that forced expression of RhoC GTPase induces the malignant transformation of immortalized human mammary epithelial (HME) cells by generating an aggressive, highly motile, and invasive phenotype, while a dominant-negative Rho inhibits metastasis (138). In addition to Rho, increased protein levels of Cdc42 (2, 3) and Rac (3) also occur in breast cancer. The reports that Rac1 is required for Ras-induced malignant transformation implicate the involvement of Rac1 in breast cancer (139). Compared to benign breast tissue, increased Rac1 mRNA expression and elevated Rac1 GTPase proteins were seen in malignant breast tissues, suggesting that constitutive activation of Rac1 signaling may be present in more aggressive breast cancers (140). Endogenous, hyperactive Rac3 is present in highly proliferative human breast cancer cell lines such as MCF-7, T47D, and MDA-MB435, and is associated with persistent kinase activity of two isoforms of the Rac effectors PAK and of JNK (141). The level of interactions between Rho GTPase and downstream effector proteins is determined by the absolute levels of the GTP-bound GTPases. By p21 binding domain (PBD) affinity assay, Mira et al., found that consistently active Rac3 GTPases lead to hyperactivity of its effector protein kinase PAK in human breast cancerderived epithelial cell lines (142). Collectively, these findings suggest that Rho proteins are overexpressed in breast cancer. In addition, it appears that the level of these proteins correlates directly to the advancement of breast cancer. If a breast cancer is highly metastatic, Rho protein expression is likely to be high.

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5.2. Role of Rho GTPases Regulatory Proteins in Breast Metastasis Dominant inhibitory approaches have demonstrated a role of Rho GTPases in focal contact formation, motility, primary tumor growth, and macro-metastasis. Each dominant inhibitory Rho protein shows a decrease in intravasation and metastasis, suggesting that Rho, Cdc42, and Rac1 play a role in the formation of breast metastasis (143). As mentioned previously, no mutations have been observed in Rho proteins in tumors. However, overexpression of the Rho GTPases has been seen in human tumors. The examination of Rho regulatory proteins may provide an explanation for these observations. For example, mutation or oncogene expression that disrupts the Rho regulatory proteins can cause dysfunctional regulation of Rho proteins in breast cancer cells, leading to cellular motility, invasion and metastasis (144).

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5.2.1. Rho GDP Dissociation Inhibitor (Rhogdi) As previously mentioned, RhoGDI inhibits Rho proteins and consequently cellular motility. GDI inhibits Rho proteins by binding to the protein’s prenylation group, preventing the localization of the GTPase to the inner plasma membrane via inhibiting the release of GDP. The introduction of RhoGDI to immortal fibroblasts has been shown to disrupt motility. However, the overexpression of RhoGDI in human keratinocytes leads to inhibition of cellular motility and the disturbance of the actin cytoskeleton (144). In addition, the overexpression of RhoGDI-α explicitly increases ERα and ERβ activation (145). 5.2.2. Rho GTPase Activating Protein (Rhogaps) Rho GTPase activating proteins (RhoGAPs) catalyze the hydrolysis of GTP to GDP, inactivating Rho proteins. p190RhoGAP is a well studied GAP protein that exists as two isoforms: p190-B and p190-A, a tumor suppressor gene. Previous results reveal that p190 inhibition leads to cytoskeleton reorganization. Introduction of p190 middle domain into motile cells disturbs heterodimer formation leading to inhibition of cell motility and migration, while the reintroduction of p120 into cells re-established directional cellular motility. The treatment of fibroblasts with sodium fluoride and LPA caused the inhibition of GAP activity while triggering GEF activity and amplifying Rho protein activation. Taken together, it appears that p190RhoGAP obstructs constitutive Rho activity (146). p190RhoGAP and p120RasGAP have also been implicated in the development of inflammatory breast cancer. Amplification of EGF-R prompts the tyrosine phosphorylation of p190RhoGAP and p120RasGAP. This causes the association of p190 and p120, leading to the formation of a heterodimer which hinders the inactivation of Rho proteins, allowing for extended GTPase activation that leads to cellular invasion. Overexpression of both p190 and p120 has been seen in HME and SUM149 inflammatory breast cancer cells, consistent with aggressive mouse mammary tumors (144). 5.2.3. Rho Guanine Exchange Factors (Rhogefs) Rho guanine exchange factors (RhoGEFs) activates Rho proteins by exchanging bound GDP for GTP, upon its phosphorylation. Once activated Rho proteins are able to activate downstream effectors that can impact the cellular motility. Activating mutations of RhoGEF

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can induce the constitutive activation of Rho proteins, suggesting a link between RhoGEF and the abnormal GTPase activity found in breast cancer. Proto-oncogene Dbl is the first identified RhoGEF. Mutations within Dbl can cause the constitutive activation of cellular transformation, possibly contributing to the metastatic phenotype (144).

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5.3. Rho Proteins in Proliferation and Progression of Breast Cancer Cells In addition to mediating breast cancer metastasis, Rho proteins can also promote breast cancer cell proliferation to facilitate their colonization. Several lines of evidence suggest an important role for Rho proteins in normal and cancerous cell growth, including G1 phase progression, mitogenesis, and Ras transformation (10, 147). The transforming potential and cell growth stimulating activity of three GTPases correlate with elevated cyclin D1 transcription due to its promoter activation. The neu (cerbB-2, HER-2) proto-oncogene encodes a receptor tyrosine kinase that is overexpressed in 20 to 30% of human breast tumors (148). ErbB-2 induced transformation of breast cells requires cyclin D1, which is induced through an E2F-dependent signaling pathway (149). In fact, Cyclin D1 is overexpressed in more than 50% of breast cancers. The mechanisms by which Rho GTPases induce cyclin D1 expression have been supported by several lines of evidence. Typically, cyclin D1 abundance is induced transcriptionally. Upon the withdrawal of growth factors, cyclin D1 undergoes rapid degradation via the ubiquitin-proteasome pathway (150). By a similar approach, the transcription regulation of cyclin D1 by Rac1 occurs via the transcription factors NF-κB and ATF-2, which directly bind to multiple sites in the cyclin D1 promoter for its activation (151, 152). Thus, activating mutants of Rac1 (RacLeu-61, RacVal-12) induced NF-κB abundance and DNA-binding to the cyclin D1 NFκB promoter, and expression of NF-κB trans-dominant inhibitors inhibited Rac induction of cyclin D1. In addition, RhoA overexpression can also inhibit p21 (CIP1) expression (153), suggesting multiple cell cycle controllers are interrupted by Rho proteins.

5.4. Potential Roles of Rho GTPase Effectors in Breast Cancer Rho GTPases stimulate different downstream signaling pathways through a spectrum of effectors. The serine/threonine kinase PAKs, for example, are important downstream effectors of both Cdc42 and Rac, and have been shown to modulate dynamics of the actin and microtubule cytoskeletons to regulate cell migration. In some breast cancers, the protein levels and kinase activity of PAK1 are elevated (154). Increased PAK kinase activity and expression are correlated with enhanced motility and invasiveness of human breast cancer cell lines (155). Ectopic expression of constitutively activated PAK1 in non-metastatic MCF7 breast carcinoma cells increases their motility (155). In addition to increased motility and invasiveness of breast cancer cells, enhanced PAK levels and PAK kinase activity can be also linked to a series of functions like DNA synthesis (142), anchorageindependent growth, and abnormal mitotic spindles (155). PAKs have been shown to physically interact with and directly phosphorylate Raf kinase, which binds to

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retinoblastoma protein (156), and interact with cyclin dependent kinases to upregulate cyclin D1 expression (157). Targeted overexpression of PAK1 in mice leads to mammary gland hyperplasia (158), while inhibiting PAK kinase activity in multiple human breast carcinoma cell lines decreases cell proliferation (142). Finally, it seems that PAK contributes greatly to the completion of the last stage of tumor progression, the formation of macrometastases. Once the tumor cells have reached a proper environment during metastatic cascades, the formation of large focal contacts could firmly attach the cell to the tissues and result in constitutive hyperactivation of PAK. Upon activation, PAK has been demonstrated to promote angiogenesis, elevate resistance to apoptosis treatment and thereby promote tumor cell growth (159). As a downstream effector of Rho, ROCK I and/or ROCK II were reported to be expressed in human testicular germ cell tumour (160), pancreatic cancer (19), and esophageal squamous cell carcinoma (18). Rho/ROCK pathway was also reported to be associated with occurrence and progression of human bladder cancer (12). In rat, elevated ROCK II expression was found in a metastatic variant of a nonmetastatic rat mammary adenocarcinoma cell line (161). Expression of conditionally activated ROCK led to endowment of colon cancer cells with epithelial morphology in vitro and augmented the aggressiveness of colon carcinoma cells’ invasion into stroma surrounding blood vessels in vivo. This suggests that increased Rho–ROCK signaling is sufficient to trigger invasion from solid tumors. ROCK disrupts cell–cell contacts by blocking the formation of adherens junctions (AJs) to increase invasiveness of tumor cells, particularly during the progression to invasive and metastatic phenotypes. Accordingly, activation of the Rho–ROCK pathway may be an essential step during the transcellular migration of tumor cells (162) and may be a valuable prognostic marker. Hence, ROCK inhibitors would be adopted as antimetastatic chemotherapeutic agents in tumors induced by elevated Rho–ROCK pathway activation. Cortactin, another downstream effector of Rho GTPases, is overexpressed in breast cancer and usually correlated with poor prognosis, presumably because of enhanced metastasis. Cortactin binds directly to the Arp2/3 complex and activates it to promote nucleation of actin filaments, an integral event in Rho GTPases regulated actin rearrangement (163). Thus, multiple Rho GTPase effectors have been implicated in the breast cancer phenotype.

5.5. Rho GTPase Pathways as Anti-Cancer Targets Since Rho GTPases play a pivotal role in the development and progression of breast cancer, they have become targets for several anti-cancer treatments. So far, several drugs have been shown to abrogate or diminish Rho GTPases triggered signaling events. These drugs either target Rho GTPases themselves or directly inhibit their downstream effecters. 5.5.1. Farnesyltransferase Inhibitors (FTIs) and Statins The activation of Rho proteins relies heavily on the post-translational modification of GTPase at the CAAX domain of the carboxyl terminus of the protein. C20 isoprenoid lipid moiety is attached to the C-terminus allowing for the localization of GTPase to the inner

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plasma membrane where it becomes activated by GAPs (164). Geranylgeranyl transferase I (GGTase I) and farnesyltransferase (FTase) are responsible for catalyzing this modification. If the localization of Rho GTPases is prevented, then Rho activity could successfully be inhibited (165). This detail has made it an attractive target for several anti-cancer treatments. Originally designed to prevent Ras attachment to inner cell membrane by abrogating Ras isoprenylation, farnesyltransferase inhibitors (FTIs) can exert similar effects on Rho protein modification. Treatment of tumor cells with a FTI leads to a decrease in farnesylated RhoB levels and a corresponding increase in geranylgeranylated RhoB levels, which promotes apoptosis and selectively inhibits cell cycle transit in malignant epithelial cells. FTI treatment of RhoC overexpressing HME or SUM149 IBC cells results in a marked reduction in their motility and invasiveness (166). Inhibition of Rho GTPases can also be achieved by hindering the activity of hydroxymethylglutaryl coenzyme A reductase (HMG-CoA), a protein required for the cholesterol biosynthesis of farnesyl pyrophosphate (FPP) and geranylgeranyl pyrophosphate (GGPP) (167). Statins are effective 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitors. Treatment of MDA-MB-231 breast cancer cells with cerivastatin obstructs the invasive phenotype of tumor cells, thus stimulating the disassembly of the actin cytoskeleton and focal adhesions (168). Cerivastatin was also shown to cause the delocalization of RhoA and decrease the activity of proteases associated with cell migration, further explaining the loss in cell motility and invasion observed in aggressive breast cancer cells (168). In MDAMB-231 cells, an aggressive breast cancer cell line, cerivastatin induces inhibition of both cell proliferation and invasion by p21Waf1/Cip1-induced G1/S arrest (168). Atorvastatin also inhibits Rho geranylgeranylation; it represses Rho activation and reverses the metastatic phenotype of human melanoma cells in vitro. In addition, it inhibits in vivo metastasis, but not proliferation of melanoma cells that overexpress RhoC (169). 5.5.2. Strongylophorine-26 Strongylophorine-26 is an alternative anti-metastatic drug that also targets Rho activity (170). The anti-cancer treatment inhibits the invasive phenotype of breast tumor cells. Treatment of MDA-MB-231 breast cancer cells with Strongylophorine-26 inhibits tumor cell motility, suggesting that Strongylophorine-26 targets aspects of Rho-mediated motility. This treatment causes a decrease in stress fiber formation, while causing an increase in the formation of focal adhesions and actin filaments that stabilize the cellular body. Strongylophorine-26 also induces the transient activation of Rho GTPases, possibly reestablishing more regulated Rho activity (170). 5.5.3. RHO siRNA Another innovative approach to treatment of aggressive breast cancer is the use of siRNA against RhoA and RhoC. Previous studies demonstrate that the overexpression of RhoA contributes to the proliferative and invasive properties of human breast cancer (3, 11), suggesting that if RhoA synthesis is inhibited, breast metastasis can be prevented. In this approach, siRNA of anti-RhoA and anti-RhoC is chemically synthesized and used to obstruct the synthesis of the respective Rho GTPases (167). Pillé and colleagues transfected the siRNA of Rho into MDA-MB-231 breast cancer cells and demonstrated the inhibition of cell

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invasion and proliferation in vitro and the abolishment of tumor cell growth and angiogenesis in vivo (167). Therefore, siRNA may be employed in the future to inhibit the synthesis and activation of respective Rho GTPases. 5.5.4 Y-27632 Other promising approaches involve selective inhibition of certain downstream effecters of Rho GTPase signaling pathways. A specific ROCK inhibitor, Y-27632, capable of blocking both Rho-mediated activation of actomyosin and invasive activity, can abolish Rhomediated activation of actomyosin and invasive activity of MM1 hepatoma cells. Studies by Robert Torka et al. (171) showed that simultaneous inhibition of ROCK and MMP reduced the invasion ability of MDA-MB231 cells by 60%. The observation that the treatment of rats with Y-27632 for 11 days substantially inhibits tumor cell dissemination without apparent adverse side effects suggests Y-27632 as a valuable drug for breast tumor cell treatment (172).

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6. Conclusion It is apparent that Rho GTPases play a pivotal role in cell motility and invasion. The aberrant activity of Rho proteins in cancer can now be explained by the identification of several factors that influence Rho activity. Rho GTPases are found to be overexpressed in metastatic cancer cells. Manipulation of Rho GTPases’ regulatory proteins and their effectors can also cause abnormal Rho activation. Abnormal Rho activation leads to the aberrant activity of transcription factors like NF-κB that can enhance the invasive phenotype of these tumor cells. Although several studies on Rho GTPases have provided a plethora of information, more studies are needed to examine the mechanisms that are driving Rho overexpression in cancer cells.

Conflict of Interest We have no conflict of interest to present.

Acknowledgements We acknowledge the support from Southern Illinois University School of Medicine and SimmonsCooper Cancer Institute, Southern Illinois University School of Medicine Central Research Committee, Illinois Department of Public Health Ticket-for-Cure Breast Cancer Research Program, Department of Defense Breast Cancer Research Program (BC074897), and National Institute of Health (R15CA133776 and R01CA13445).

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phosphorylation of GH receptor, JAK2, and STAT5B. Mol Endocrinol, 14: 1338-1350, 2000. [142] Mira, J. P., Benard, V., Groffen, J., Sanders, L. C., and Knaus, U. G. Endogenous, hyperactive Rac3 controls proliferation of breast cancer cells by a p21-activated kinasedependent pathway. Proc Natl Acad Sci U S A, 97: 185-189, 2000. [143] Bouzahzah, B., Albanese, C., Ahmed, F., Pixley, F., Lisanti, M. P., Segall, J. D., Condeelis, J., Joyce, D., Minden, A., Der, C. J., Chan, A., Symons, M., and Pestell, R. G. Rho family GTPases regulate mammary epithelium cell growth and metastasis through distinguishable pathways. Mol Med, 7: 816-830, 2001. [144] Lin, M. and van Golen, K. L. Rho-regulatory proteins in breast cancer cell motility and invasion. Breast Cancer Res Treat, 84: 49-60, 2004. [145] Su, L. F., Knoblauch, R., and Garabedian, M. J. Rho GTPases as modulators of the estrogen receptor transcriptional response. J Biol Chem, 276: 3231-3237, 2001. [146] Vincent, S. and Settleman, J. Inhibition of RhoGAP activity is sufficient for the induction of Rho-mediated actin reorganization. Eur J Cell Biol, 78: 539-548, 1999. [147] Khosravi-Far, R., Solski, P. A., Clark, G. J., Kinch, M. S., and Der, C. J. Activation of Rac1, RhoA, and mitogen-activated protein kinases is required for Ras transformation. Mol Cell Biol, 15: 6443-6453, 1995. [148] Slamon, D. J., Clark, G. M., Wong, S. G., Levin, W. J., Ullrich, A., and McGuire, W. L. Human breast cancer: correlation of relapse and survival with amplification of the HER-2/neu oncogene. Science, 235: 177-182, 1987. [149] Lee, R. J., Albanese, C., Fu, M., D'Amico, M., Lin, B., Watanabe, G., Haines, G. K., 3rd, Siegel, P. M., Hung, M. C., Yarden, Y., Horowitz, J. M., Muller, W. J., and Pestell, R. G. Cyclin D1 is required for transformation by activated Neu and is induced through an E2F-dependent signaling pathway. Mol Cell Biol, 20: 672-683, 2000. [150] Diehl, J. A., Zindy, F., and Sherr, C. J. Inhibition of cyclin D1 phosphorylation on threonine-286 prevents its rapid degradation via the ubiquitin-proteasome pathway. Genes Dev, 11: 957-972, 1997. [151] Joyce, D., Bouzahzah, B., Fu, M., Albanese, C., D'Amico, M., Steer, J., Klein, J. U., Lee, R. J., Segall, J. E., Westwick, J. K., Der, C. J., and Pestell, R. G. Integration of Rac-dependent regulation of cyclin D1 transcription through a nuclear factor-kappaBdependent pathway. J Biol Chem, 274: 25245-25249, 1999. [152] Guttridge, D. C., Albanese, C., Reuther, J. Y., Pestell, R. G., and Baldwin, A. S., Jr. NF-kappaB controls cell growth and differentiation through transcriptional regulation of cyclin D1. Mol Cell Biol, 19: 5785-5799, 1999. [153] Liberto, M., Cobrinik, D., and Minden, A. Rho regulates p21(CIP1), cyclin D1, and checkpoint control in mammary epithelial cells. Oncogene, 21: 1590-1599, 2002. [154] Salh, B., Marotta, A., Wagey, R., Sayed, M., and Pelech, S. Dysregulation of phosphatidylinositol 3-kinase and downstream effectors in human breast cancer. Int J Cancer, 98: 148-154, 2002. [155] Vadlamudi, R. K., Adam, L., Wang, R. A., Mandal, M., Nguyen, D., Sahin, A., Chernoff, J., Hung, M. C., and Kumar, R. Regulatable expression of p21-activated kinase-1 promotes anchorage-independent growth and abnormal organization of mitotic spindles in human epithelial breast cancer cells. J Biol Chem, 275: 36238-36244, 2000.

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[156] Wang, S., Ghosh, R. N., and Chellappan, S. P. Raf-1 physically interacts with Rb and regulates its function: a link between mitogenic signaling and cell cycle regulation. Mol Cell Biol, 18: 7487-7498, 1998. [157] Tjandra, H., Compton, J., and Kellogg, D. Control of mitotic events by the Cdc42 GTPase, the Clb2 cyclin and a member of the PAK kinase family. Curr Biol, 8: 9911000, 1998. [158] Wang, R. A., Mazumdar, A., Vadlamudi, R. K., and Kumar, R. P21-activated kinase-1 phosphorylates and transactivates estrogen receptor-alpha and promotes hyperplasia in mammary epithelium. Embo J, 21: 5437-5447, 2002. [159] Bokoch, G. M. Biology of the p21-activated kinases. Annu Rev Biochem, 72: 743-781, 2003. [160] Kamai, T., Arai, K., Sumi, S., Tsujii, T., Honda, M., Yamanishi, T., and Yoshida, K. I. The rho/rho-kinase pathway is involved in the progression of testicular germ cell tumour. BJU Int, 89: 449-453, 2002. [161] Wang, W., Wyckoff, J. B., Frohlich, V. C., Oleynikov, Y., Huttelmaier, S., Zavadil, J., Cermak, L., Bottinger, E. P., Singer, R. H., White, J. G., Segall, J. E., and Condeelis, J. S. Single cell behavior in metastatic primary mammary tumors correlated with gene expression patterns revealed by molecular profiling. Cancer Res, 62: 6278-6288, 2002. [162] Sahai, E. and Marshall, C. J. ROCK and Dia have opposing effects on adherens junctions downstream of Rho. Nat Cell Biol, 4: 408-415, 2002. [163] Uruno, T., Liu, J., Zhang, P., Fan, Y., Egile, C., Li, R., Mueller, S. C., and Zhan, X. Activation of Arp2/3 complex-mediated actin polymerization by cortactin. Nat Cell Biol, 3: 259-266, 2001. [164] Mondal, M. S., Wang, Z., Seeds, A. M., and Rando, R. R. The specific binding of small molecule isoprenoids to rhoGDP dissociation inhibitor (rhoGDI). Biochemistry, 39: 406-412, 2000. [165] van Golen, K. L., Bao, L., DiVito, M. M., Wu, Z., Prendergast, G. C., and Merajver, S. D. Reversion of RhoC GTPase-induced inflammatory breast cancer phenotype by treatment with a farnesyl transferase inhibitor. Mol Cancer Ther, 1: 575-583, 2002. [166] Du, W. and Prendergast, G. C. Geranylgeranylated RhoB mediates suppression of human tumor cell growth by farnesyltransferase inhibitors. Cancer Res, 59: 5492-5496, 1999. [167] Pille, J. Y., Denoyelle, C., Varet, J., Bertrand, J. R., Soria, J., Opolon, P., Lu, H., Pritchard, L. L., Vannier, J. P., Malvy, C., Soria, C., and Li, H. Anti-RhoA and antiRhoC siRNAs inhibit the proliferation and invasiveness of MDA-MB-231 breast cancer cells in vitro and in vivo. Mol Ther, 11: 267-274, 2005. [168] Denoyelle, C., Vasse, M., Korner, M., Mishal, Z., Ganne, F., Vannier, J. P., Soria, J., and Soria, C. Cerivastatin, an inhibitor of HMG-CoA reductase, inhibits the signaling pathways involved in the invasiveness and metastatic properties of highly invasive breast cancer cell lines: an in vitro study. Carcinogenesis, 22: 1139-1148, 2001. [169] Collisson, E. A., Kleer, C., Wu, M., De, A., Gambhir, S. S., Merajver, S. D., and Kolodney, M. S. Atorvastatin prevents RhoC isoprenylation, invasion, and metastasis in human melanoma cells. Mol Cancer Ther, 2: 941-948, 2003.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 159-186

ISBN: 978-1-60692-570-6 © 2009 Nova Science Publishers, Inc.

Chapter V

Role of Serine Proteases and Their Receptors in Cellular Motility V.M. Shpacovitch1, M.D. Hollenberg2 and M.Steinhoff 1 1

Department of Dermatology and Ludwig Boltzmann Institute for Immunobiology of the Skin, University of Münster, Münster, Germany 2 Canadian Institutes of Health Research Proteinases and Inflammation Network, Department of Pharmacology and Therapeutics, University of Calgary, Calgary, Alberta, Canada

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Abstract Serine proteinases constitute a family of proteolytic enzymes with diverse biological functions ranging from the digestion of extracellular matrix and dietary proteins to complement formation. However, recent groundbreaking studies revealed a role of certain serine proteinases as signalling molecules which are able to regulate cell function via cell surface receptors. Urokinase plasminogen activator receptor (uPAR) and proteinase-activated receptors (PARs) are two receptor types via which serine proteinases can trigger intracellular signalling cascades. Currently, the role of these receptors in serine proteinase-mediated signalling is under intensive investigation. Both types of receptors have been reported to influence the migratory behaviour of different cells under physiological and pathophysiological conditions. Among these cells are human leukocytes, smooth muscle cells, sperm, and invading cancer cells. Moreover, activation of uPAR as well as PARs is known to affect the expression of cell adhesion molecules, actin polymerisation and to induce cell shape changes, subsequently acting on cell movement. Thus, serine proteinases are capable of regulating cell motility via two major pathways. By one pathway, the enzymes induce changes in cell microenvironment (for example, via digestion of extracellular matrix proteins). By the other pathway, serine proteases may directly trigger the signalling cascades resulting in changes of cell migratory behaviour. Understanding of mechanisms involved in both pathways could lead to new therapeutic approaches in inflammatory, allergic, and neoplastic diseases.

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Introduction Proteinases (also termed proteases or endopeptidases) can be organised into catalytic types according to the amino acid residue or metal ion that plays the primary role in catalysis. Thus, serine, cysteine, aspartic acid and metalloproteinases can be distinguished by their catalytic mechanisms. Serine proteases are widespread and very numerous. These enzymes are found in viruses, bacteria and eukaryotes. In addition to serine (nucleophile), the catalytic machinery of serine proteases involves a proton donor. Usually, a histidine residue plays the role of proton donor (trypsin, mast cell tryptase, chymotrypsin, for example). The essential third residue of the serine protease ‘catalytic triad’ is aspartic acid, which may on occasion be substituted by a second histidine. Thus, the “classical catalytic triad” of serine proteases consists of Ser, His and Asp. However, in some serine proteases (signal peptidase I from E. coli; signalase from Saccharomyces cerevisiae, for example) a lysine residue plays the role of proton donor and in these enzymes the third residue of the classical ‘catalytic triad ’ is not required [1]. Serine proteases are produced as inactive precursors or zymogens. Subsequent zymogen conversion into a mature physiologically active enzyme is mediated via a process called “limited proteolysis” or zymogen activation [1]. In mammals, serine proteases are involved in many important physiological processes ranging from digestion of dietary proteins to blood coagulation and leukocyte migration [1]. In addition, serine proteases are thought to be involved in diseases such as pulmonary emphysema, arthritis, inflammation, and tumour metastasis. Altogether, these facts single out serine proteases as excellent targets for drug design. Three different types of endogenous serine protease inhibitors can be distinguished, based on their mechanism of action: canonical, non-canonical and serpins [2]. Recently, many synthetic serine protease inhibitors have been developed. Some of the inhibitors demonstrate a potential not only as tools for understanding the roles of the enzymes in biological processes, but also as promising therapeutic compounds for a variety of diseases. For example, synthetic inhibitors of the serine protease, urokinase plasminogen activator (uPA), show promise as anti-metastatic drugs [3]. It is important to point out that an imbalance between the endogenous inhibitors of serine proteases and their targeted enzymes can affect immune/inflammatory responses and may trigger disease development. The presence of serine protease inhibitors not only limits direct proteolytic activity of enzymes, which could be potentially hazardous and lead to tissue damage, but also regulates interactions between serine proteases and their receptors. Recent groundbreaking studies have revealed a role of certain serine proteases as signalling molecules which are able to regulate cell function via cell surface receptors. Protease-activated receptors (PARs) and urokinase plasminogen activator receptor (uPAR) are triggered by serine proteinases to generate signalling cascades in cells [4][5].

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Role of Serine Proteases and Their Receptors in Cellular Motility

(A)

NH

Tethered ligand 2

161

Protease-activated receptor

membrane

COOH

(B)

Serine protease Activating peptide

COOH

COOH

Intracellular signaling cascades

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(C)

COOH

COOH

Figure 1. Activation of protease-activated receptors (PARs). (A) N-terminal part of PARs contains “tethered ligand” sequence, which remains hidden during inactive status of the receptor. Under these conditions, “tethered ligand” sequence is not able to interact with the second extracellular loop of the receptor and can not trigger signaling cascades. (B) Upon activation, accessible serine protease cleaves the N-terminal domain of the receptor and unmasks a “tethered ligand” sequence. Further, the “tethered ligand“ interacts with the second extracellular loop of the same receptor and thus, triggers signaling events. However, synthetic activating peptides (PAR-APs) are capable of PAR activation without any cleavage. These peptides can directly interact with the extracellular domain of the receptor and induce identical downstream signaling events. (C) The mechanism of intermolecular trans-activation is also described for PARs. For example, thrombin cleaved N-terminal domain (unmasked “tethered ligand “sequence) of PAR1 is able to interact with the second extracellular loop of untouched PAR2 and, subsequently, activate the PAR2 receptor.

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PARs represent a novel subfamily of G protein-coupled receptors with seven transmembrane domains [6][4]. PARs can account for a significant proportion of signalling generated by serine proteases. To date, four PARs have been cloned and characterized. Classically, PAR1,3, and 4 are considered primarily as targets for thrombin, but can also be regulated by trypsin, cathepsin G or by tissue kallikreins (e.g. KLK14). In contrast, PAR2 is resistant to thrombin but can be activated by trypsin, mast cell tryptase, tissue kallikreins (KLKs 5, 6 and 14) and by enzymes produced by immune cells and pathogens [4][7][8]. The unique mechanism whereby serine proteases signal via PARs involves the cleavage of the receptor N-terminal domain, to expose a new previously cryptic sequence. The exposed sequence remains tethered to the receptor and acts further as a receptor-activating ligand, referred to as a “tethered ligand” [6][4] (Fig.1). It is also important to note that some proteases are able to cleave PARs downstream of the tethered ligand sequence, making further proteolytic activation of PARs impossible. The PARs can be activated not only by proteolytic unmasking of their tethered ligand but also by synthetic peptides with sequences based on the revealed tethered ligands. The PAR-selective receptor-activating peptides (PARAPs: Table 1) can activate the PARs without the need for proteolysis. The PAR-APs have proved to serve as important tools in the PAR research field because they can eliminate the need to trigger the PARs with serine proteases that can cause both PAR-dependent as well as PAR-independent responses in various cells. Although signaling by PARs 1, 2, and 4 can be triggered by either proteases or receptor-specific PAR-APs, the role of PAR3, which cannot be activated by its ‘tethered ligand’, remains elusive. Currently, PAR3 is considered as an accessory receptor for PAR1 or PAR4 [9][10][11]. PARs are known to be expressed by different cell types and are involved in a wide spectrum of cellular responses under physiological or pathophysiological conditions. The effects of PAR activation on the motility of different cell types have been demonstrated both in vitro and in vivo. PAR activation affects leukocyte motility, adhesion molecule expression, and invasiveness of several types of cancer cells [6][4]. These effects of PAR activation on cell motility will be discussed further in the sections that follow. The other receptor involving serine protease for activation of cell signalling events is uPAR. The urokinase plasminogen activator (uPA) is a serine protease that catalyzes the conversion of plasminogen to plasmin. The binding of uPA to uPAR leads to the proteolysis on cell surface. uPAR is a glycosylphosphatydilinositol (GPI)-anchored protein consisting of three domains [12]. This receptor binds with high affinity to uPA, pro-uPA and the aminoterminal fragment of uPA (ATF), as well as the extracellular matrix (ECM) protein vitronectin. Despite the lack of a cytosolic domain, uPAR is able to transmit intracellular signals by the means of its association with at least three types of trans-membrane proteins: integrins, G Protein-coupled receptors (FPRL1, for example) and caveolin [5]. uPAR is expressed on a variety of cells, including endothelial cells and hematopoietic cells. Recent studies indicate that uPA-uPAR complex affects cell migration, shape changes, motility, adhesion, and differentiation [5]. This complex also affects the invasive potential of some tumours. The data concerning the role of uPAR in cell motility will be discussed in sections that follow. It is now clear that serine proteases are able to affect cell motility not only directly via digestion of extracellular matrix (ECM) proteins, but also via signalling events triggered after

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interaction of enzymes with either PARs or uPAR. Thus, both the direct as well as the receptor-mediated effects of serine proteases on the motility of different cell types will be discussed below. Table 1 Protease-activated receptors (PARs), urokinase plasminogen activator receptor (uPAR) and their effects at the motility of human cells

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Receptor type

PAR1 GPCR

Protease-activated receptors (PARs) PAR2 PAR3 GPCR (GPCR)a

Classical activating or interacting proteases

Thrombin

Trypsin Tryptase Tissue Kallikreins (KLK14)

None known

Classical activating peptides

TFLLRN-NH2 (m,h,r)

SLIGKV-NH2 (h) SLIGRL-NH2 (m,r)

None known

G-proteins associated with receptor

Gi, Gq/11 and G12/13

Gi and Gq/11

Human cells, which change their motility behaviour after receptor activation

Eosinophil migration; invasiveness of breast, gastric, colon, kidney and prostate cancer cells, invasiveness of melanoma cells

Trans-endothelial migration and motility in collagen gels of neutrophils; invasiveness of breast, colorectal and pancreatic cancer cells; spermatozoa motility; migration of VSMCs

Does not signal on its own; serves as a co-receptor for PAR1 or PAR4 None known

PAR4 GPCR

Thrombin Trypsin Tissue Kallikreins (KLK14) GYPGQV-NH2 (h) GYPGKV-NH2 (m) AYPGKF-NH2 (r) G12/13 and Gq

Migration and invasion of human hepatocellular carcinoma cells

uPAR GPI-anchored membrane protein (urokinase plasminogen activator)b

None known

Does not signal on its own; could transmit signal via association with integrins, GPCRs or calveolin Leukocyte motility via uPAR-integrin interactions; leukocyte chemotaxis; uPAuPAR complex promotes cancer cell invasiveness; migration of VSMCs

To date four different PARs are known. Thrombin, trypsin and mast cell-derived tryptase are considered as classical PAR activators. Apart from these enzymes, tissue kallikreins, pathogen- and immune cell-derived proteases appear to activate or inactivate PARs. Another protease-associated receptor is uPAR. The binding of urokinase plasminogen activator (uPA) to its receptor (uPAR) leads to cell surface proteolysis associated with the conversion of plasminogen into enzymatically active plasmin. The following abbreviations were used: h-human; m-mouse; r-rat. a The ability of PAR3 to signal on its own remains in doubt, because this receptor appears to lack the cytoplasmic tail domain shown in other PARs to couple with G-proteins. b In uPA-uPAR complex, some signalling effects triggered by uPA do not require its proteolytic activity since chemically blocked or catalytically inactive uPA derivatives are equally effective. However, the expression of uPAR and its ability to bind uPA represent necessary conditions for the triggering of signalling events.

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Role of Serine Proteases and Their Receptors in the Motility of Hematopoietic Cells The migration of immune cells towards the site of tissue injury caused by physical means or by invading pathogens plays a crucial role in host immune responses. The central event of leukocyte and lymphocyte migration is the translocation of these cells from the vascular system, through endothelium, and into the extracellular matrix surrounding the injured tissue. The process of immune cell transmigration through the endothelium includes several main steps: initial leukocyte “capture” by the vessel wall followed by the immune cell “rolling” along the vessel wall. Then, the process of firm adhesion of immune cells to endothelial cells, followed finally by immune cell homing and transmigration [13]. Different types of cell adhesion molecules (selectins, integrins etc.) expressed by immune as well as endothelial cells actively participate in the transmigration process. It is also important to note that the translocation of immune cells across the vascular wall into the interstitial space is normally governed by different types of chemoattractive molecules. The migration of different cell types through the extracellular matrix is achieved via both proteolytic and non-proteolytic strategies. Proteolytic migration is associated with the matrixdegrading activity of enzymes such as matrix metalloproteinases, serine proteases and cathepsins at the cell surface of moving cells (“pericellular proteolysis”). This strategy is usually used by slow-moving cells such as tumour cells or fibroblasts. Other cells, especially rapidly moving cells like T lymphocytes, often use non-proteolytic strategy to migrate through the extracellular matrix [14]. Additionally, recent studies have revealed that some neutrophil serine proteases as well as mast cell-derived chymase possess a chemotactic activity for leukocytes [15] [16]. Human chymase demonstrates a potent concentration-dependent chemotactic activity for monocytes and neutrophils. Chymase also stimulates migratory activity of lymphocytes and purified T cells, but this effect appears to be chemokinetic rather than chemotactic [17]. Interestingly, the inhibition of proteolytic activity of chymase also significantly reduced the chemotactic activity of this enzyme. This fact indicates that the proteolytic activity of chymase is required for its chemotactic function [17]. Moreover, it is important to note that proteolytic cleavage by chymase of certain molecules (endothelins, for example) results in release of biologically active peptides with chemotactic activity for neutrophils and monocytes [18]. A study performed by Sun and colleagues with formyl peptide receptorexpressing basophilic leukemia cells indicates that the chemotactic activity of the neutrophil serine protease, cathepsin G, is linked to the activity of the formyl peptide receptor (FPR) [19]. Therefore, extracellular cathepsin G might contribute to leukocyte recruitment in vivo using FPR-mediated responses of neutrophils and monocytes [20]. Neutrophil serine proteases are also able to induce proteolytic modification of chemokines, small molecules, which involved in the recruitment of leukocytes [16].Additionally, neutrophil serine proteases could cleave cell surface adhesion molecules, such as intercellular adhesion molecule-1 (ICAM-1), vascular adhesion molecule 1 (VCAM1) and epithelial (E)-cadherin [21][22][23]. Acting this way, neutrophil-derived serine proteases are able to induce shedding of adhesion molecules from the cell surface of immune and endothelial cells. Since immune cell migration is associated with a series of attachment and detachment events,

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shedding of adhesion molecules might limit cell-cell and cell-extracellular matrix contacts, thereby affecting leukocyte migration. Thus, immune cell-derived serine proteases are able to affect leukocyte recruitment serving as chemotactic molecules, utilizing proteolytic activity to generate chemoattractive peptide fragments, and to modify cell adhesion molecules. These effects of serine proteases on leukocyte motility via agonist peptide generation and extracellular matrix digestion are not associated with intracellular signalling events mediated via membrane receptors like the PARs, for example, which are proteolytically activated. That said, a significant proportion of the motility effects triggered by serine proteases (especially those of the coagulation cascade) in leukocytes and/or endothelial cells can now be seen to be due to the cleavage and activation of one or more members of protease-activated receptors family (PARs). PAR-mediated effects of serine proteases on leukocyte motility in vitro and in vivo under normal as well as pathophysiological conditions are just beginning to be appreciated due to recent studies. In these studies, specific synthetic receptor-selective PAR-activating peptides (so-called PAR-APs) have played very important roles to distinguish the PAR-dependent effects of serine proteases from PAR-independent mechanisms whereby proteases can regulate cell function [24] Isolated human neutrophils from healthy and non-pregnant donors express PAR2 and PAR3 mRNA [25]. However, the functional role of PAR3, apart from its ability to act as a coreceptor for other PARs, remains elusive. Howells and colleagues demonstrated that PAR2 agonists affect neutrophil shape changes and, if applied along with fMLP, enhance the effect of the latter on neutrophil surface expression of integrins (Mac-1) [26]. In this study, the classical PAR2-AP (SLIGKV-NH2) was used. However, this peptide is sensitive to peptidase activity and potentially also to enzymes derived from human neutrophils. Thus, it seemed of value to use aminopeptidase resistant synthetic PAR2-APs (trans-cinnamoyl-LIGRLO-NH2, furoyl-LIGRLO-NH2 for example) for studies employing human neutrophils exposed to a PAR2 agonist over prolonged time periods. In such studies, Shpacovitch and colleagues [27] demonstrated that stimulation of human neutrophils with PAR2 agonists (trypsin and transcinnamoyl-LIGRLO-NH2) leads to a significantly enhanced motility of PMNs in threedemensional (3-D) collagen lattices in vitro, suggesting a role for PAR2 in regulating neutrophil migration through extracellular matrix towards the site of inflammation and/or infection. Moreover, PAR2 activation enhanced the shedding of L-selectin from the cell surface of human neutrophils [27]. The effect of PAR2 agonists on neutrophil motility in an in vitro extracellular matrix model system was significantly reduced by the co-application of the L-selectin shedding inhibitor, KD-IX-73-4 [27]. This finding indicated a link between PAR2 agonist-induced shedding of L-selectin and the enhanced motility of neutrophils through the extracellular matrix in vitro. In the same study, the authors demonstrated that PAR2 activation with PAR2-AP increases the expression of VLA-4, which is involved in leukocyte-endothelial interactions (the firm adhesion and extravasation of leukocytes) and in leukocyte-leukocyte interactions. However, a potential effect of PAR2 agonists on the transmigration of human neutrophils via the endothelium was not evaluated. In recent studies, Shpacovitch and colleagues reported that simultaneous stimulation of human endothelial cells (HMEC-1) and neutrophils with proteases (trypsin, tryptase) or PAR2-AP (tc-LIGRLO-NH2) reduces trans-endothelial migration of granulocytes in vitro [28]. Together with previous

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findings, this fact indicates the ability of serine proteases to regulate neutrophil motility via PARs depending on the compartmentalisation of protease-receptor interactions (inside blood vessel or in the extravascular compartment). PAR2 also appears to be the predominant functional member of the PAR family expressed by another type of human granulocytes: human eosinophils. However, it is still not known whether activation of PAR2 on human eosinophils affects the migratory ability of these cells in a 3-dimensional collagen gel assay as well as in a trans-endothelial migration assay. The presence and functionality of another member of PAR family -PAR1- on human eosinophils cannot be completely excluded [25]. This receptor seems to be involved in the thrombin-induced effects on eosinophil migration [29]. The authors studied cell migration using a micropore filter assay. The involvement of PAR1 in thrombin-dependent migration of isolated human eosinophils was tested using a selective PAR1 peptide agonist (TFLLRNPNDK) and an antibody that blocks the enzymatic cleavage and activation of PAR1 [29]. Thrombin significantly stimulated eosinophil chemotaxis in vitro in a concenrationdependent manner. This effect was mimicked by the PAR1-activating synthetic peptide. The receptor cleavage-blocking antibody reversed the effect of thrombin on eosinophil chemotaxis. A specific PAR2-activating peptide did not affect the chemotaxis of human eosinophils [29]. Thus, serine proteases are able to affect migratory behaviour of human granulocytes via members of PAR family, PAR1 and PAR2, in vitro. The ability of PARs to mediate effects of serine proteases on the motility of other human immune cells such as monocytes, macrophages, dendritic cells, mast cells and lymphocytes merits further investigation. Threedimensional assays with extracellular matrix proteins (for example, a collagen gel assay), chemotaxis and transmigration assays could serve as important tools for in vitro studies of migratory behaviour triggered by proteases and PARs in human immune cells. The role of PARs in the motility of immune cells was demonstrated not only in different in vitro assays mentioned above, but also in in vivo model systems. Zimmerman and colleagues used a non-selective PAR1/PAR2 agonist (SFLLRNPNDKYEPF: activates both of PARs 1 and 2) to assess a potential involvement of PARs in leukocyte trafficking in vivo [30]. In this work a method of superfusion of the rat mesenteric venule was used. However, the use of a non-selective PAR1/PAR2 agonist in the study did not allow a conclusion as to which of the PARs (PAR1, PAR2, or both) were responsible for causing the effects on leukocyte trafficking in vivo. In a subsequent work, Vergnolle and colleagues investigated further, the involvement of different thrombin receptors (PAR1 and PAR4) in leukocyte rolling and adherence in vivo in the same animal model [31]. Topical application of the selective PAR1 agonist (TFLLR-NH2) to rat mesenteric venules failed to reproduce the increased leukocyte rolling and adhesion observed after topical thrombin application. Moreover, topical addition of PAR2-APs in the rat preparation resulted in increased leukocyte rolling and adhesion [32]. Thus, first of all, these data demonstrated the involvement of PAR2 activation in leukocyte trafficking in vivo. Further, these data indicated that thrombin-induced effects on leukocyte rolling and adherence are not associated with PAR1, and, thus are either mediated via PAR4 (another thrombin receptor) or are PAR-independent. To clarify the role of PAR4 in leukocyte rolling and adhesion, Vergnolle and colleagues [31] used a selective PAR4-activating peptide (AYPGKF-NH2) in the same superfusion animal model. In these

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experiments, the application of the PAR4-activating peptide significantly increased the number of rolling leukocytes and also enhanced the number of leukocytes adherent to the vessel wall [31]. Moreover, the authors demonstrated that the intraperitoneal injection of the PAR4-activating peptide in rats enhances PMN recruitment into the peritoneal cavity [31]. The role of PARs in the regulation of leukocyte motility in vivo was also investigated in further studies using PAR null mice. In comparison with wild-type animals, PAR2-deficient mice showed significantly decreased leukocyte rolling in a model of acute inflammation induced by surgical trauma [33]. Moreover, disruption of the PAR2 gene abolishes trypsininduced increases in lymphocyte adhesion and reactive oxygen species (ROS) production [34]. This finding indicates the involvement of PAR2 activation in trypsin-stimulated lymphocyte activation in vivo. Taken together, the findings mentioned above indicate that, among the members of PAR family, PAR2 and PAR4 play an important role in the receptormediated effects of endogenous proteases such as trypsin and thrombin on leukocyte motility in vivo. The role of urokinase plasminogen activator and its receptor uPAR in immune cell motility in vitro and in vivo is also under intensive investigation. Integrins are cell surface receptors, which are involved in many aspects of cell motility: interactions with extracellular matrix, leukocyte adhesion to endothelium, tumour cell invasion and some other effects. uPAR-integrin interactions on different cell types have been summarized recently in a review [35]. However, several major findings concerning uPAR-integrin interactions on leukocytes as well as new data will be discussed below. Leukocytes express both uPA and uPAR. On leukocytes, uPAR (CD87) has been reported to form complex with β2 integrins, which are known to play a crucial role in the firm adhesion of leukocytes to the endothelium [36][37]. This finding indicated possible functional interactions between uPAR and β2 integrins. Indeed, in further studies, May and colleagues have demonstrated that β2 integrin-dependent leukocyte-endothelial cell interactions in vitro as well as leukocyte recruitment into inflamed areas in vivo require the presence of uPAR on the leukocyte surface [38]. The data of this study also suggest that the initial phase of leukocyte interaction with the vessel wall is associated with proteolysisindependent cross talk between uPAR and β2 integrins [38]. This observation is also consistent with the data of more recent work of Gyetko and colleagues, where the authors demonstrated that uPAR participates in neutrophil recruitment in response to Pseudomonas aeruginosa infection via uPA-independent manner [39]. P. aeruginosa infection is known to recruit neutrophils to pulmonary parenchyma by a β2 integrin-dependent mechanism. It was demonstrated that uPAR deficient mice have profoundly diminished neutrophil recruitment in response to P. aeruginosa pneumonia compared with wild-type animals [39]. As expected, uPAR-β2 integrin interactions are strongly involved in this neutrophil migration [39]. In further work, Gyetko and colleagues showed that uPAR also participates in T lymphocyte migration [40]. It is also interesting to note that αM β2 integrin (CD11b/18, Mac-1) possesses uPA binding activity and it seems that the kringle as well as the proteolytic domains of uPA can be recognized by Mac-1 [41]. These findings are consistent with earlier work demonstrating that uPAR-dependent proteolytic activity does not seem to be critical for leukocyte recruitment [42]. Thus, it seems that uPA-independent mechanisms of uPAR function play the major role in the initial phase of leukocyte transmigration in vivo.

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It is also important to mention that the uPA-uPAR complex plays a significant role in leukocyte chemotaxis. Gradients of uPA, pro-uPA and amino-terminal fragment of uPA are chemotactic for a variety of cells, all of which express uPAR [43]. Gyetko and colleagues have provided the first in vitro data indicating a potential involvement of the uPA-uPAR complex in chemotaxis of human monocytes and neutrophils [44][45]. Further, Resnati and colleagues shed some light on the signalling events triggered by uPA-uPAR complex in leukocytes and leading to the changes of chemotactic activity of these cells [43]. The authors demonstrated that intracellular tyrosine kinase(s) are required for such signalling events. It is also important to note that the soluble form of uPAR (which lacks a GPI anchor) promotes the adhesion of THP-1 cells (monocytic cell line) to fibronectin and vitronectin, indicating that soluble uPAR (suPAR) can modify cell adhesion as well as migration [46]. Remarkably, the contact of endothelial cells with peripheral blood cells has been shown to enhance the production of suPAR by endothelial cells [47]. The D2D3 domain of suPAR can induce chemotaxis in uPAR-deficient cells, thereby bypassing the requirement for uPA to bind to uPAR for signalling [48]. Further studies have demonstrated the ability of pertussis toxin to block chemotaxis response to uPA or the D2D3 fragment, thereby pointing to the involvement of G protein coupled receptor (GPCR) in uPA-uPAR triggered chemotactic responses [5]. One of the possible candidates among GPCRs is FPRL1, which has been found to transduce the chemotactic activity of uPA-uPAR. FPRL1 was demonstrated to interact with suPAR. Subsequently it was found that the association of FPRL1 with suPAR induces chemotaxis of isolated human peripheral blood monocytes as well as THP-1 cells [49]. Taken together, these findings indicated that suPAR could participate in chemotaxis and transendothelial migration of human leukocytes not only in vitro but also potentially in vivo. FPRL1 triggered signalling events could play an important role in uPA-uPAR chemotactic responses. In this context it is important to mention that uPAR could have signalling partnership interactions with other receptors, among which are cell adhesion molecules such as selectins. For example, L-selectin expressed by human neutrophils seems to have a direct physical association with uPAR and thus may form a functional signalling complex [50]. The role of such a complex between L-selectin and uPAR in neutrophil motility merits further investigation. According to recent publications, leukocytes and lymphocytes can move through the extracellular matrix using a non-proteolytic strategy [14][51][52]. Thus, the proteolytic function of uPA-uPAR complex may play only a minor role for the motility of leukocytes through the extracellular matrix. However, Al-Atrash and colleagues have reported that uPA contributes to the degradation of extracellular matrix by NK cells. This degradation may affect the ability of NK cells to accumulate at sites of tumour metastasis [53]. Taken together, the data from studies done to date clearly demonstrate that serine proteases utilize their proteolytic activity not only to degrade matrix proteins and generate chemotactic peptides, but also to function as signalling molecules for the regulation of leukocyte motility. The function of serine proteases as signalling molecules is especially important for leukocyte movement since playing this role the enzymes could trigger several signalling cascades in parallel. Such parallel signalling events could result not only in direct effects on leukocyte motility, but also in indirect effects mediated via enhanced or reduced production of chemoattractive molecules by activated leukocytes. Thus, the receptor-

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mediated interactions between serine proteases and leukocytes represent a complex system with a potential opportunity for an amplification of a direct signalling effect triggered by proteases. On the other side, receptor-independent effects of serine proteases on leukocyte motility appear to be more limited. It is also important to note, that apart from endogenous serine proteases, some pathogen-derived proteases could affect leukocyte functions via receptor-mediated signalling events [8]. The ability of pathogen-derived proteases to affect leukocyte recruitment by PAR-dependent mechanism is an area that is of interest for further investigation that may represent an attractive opportunity for the design of new antiinflammatory therapeutic strategies.

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Role of Serine Proteases and Their Receptors in The Motility of Tumour Cells A role for proteases in the oncogenic process was proposed some time ago with the hypothesis that tumour-associated proteolytic activity could degrade the extracellular matrix to promote the invasion of tumour cells into the surrounding healthy tissue [54]. This concept focused on extracellular and pericellular proteolysis, which helps to degrade extracellular matrix components in order to facilitate tumour invasion and metastasis. In further studies, individual proteases with pro-metastatic activities were identified among serine, cysteine and metalloproteinases. Currently, it is well recognized that proteolytic enzymes are capable of contributing to carcinogenesis not only during tumour invasion (considered as a late stage process of tumorigenesis) but at all stages of tumour progression [55][56][57]. Despite the well established role of proteases as pro-metastatic factors, recent data shed some light on their potential function as tumour suppressor molecules [58]. According to the classical point of view, it appears that intracellular proteases may function as tumour suppressors, whereas extracellular proteases are considered as enzymes that can promote tumour progression. However, there are some exemptions from this rule [58].The data concerning a role of proteases in tumour progression support the idea that the enzymes may play a dual function, either promoting or suppressing tumorigenesis, depending on the type of tumour, the localisation of the protease (intracellular or extracellular), the tumour microenvironment and other, as yet unidentified, factors. Amongst the serine proteases, there are enzymes with pro-metastatic as well as antiinvasive activity. For example, a membrane serine protease, hepsin, is reported to be one of the most strongly up-regulated genes in prostate cancer. This enhanced hepsin expression appears to correlate with disease progression [59]. A role for neutrophil elastase to enhance the progression of skin tumours is supported by data obtained by Starcher and colleagues in mice [60]. The study found that elastase-deficient mice had significantly fewer tumours as compared with wild type animals. Additionally, treatment with elastase inhibitors significantly reduced the incidence of ultraviolet radiation-induced tumours. Application of an elastase inhibitor was also beneficial in ischaemia-reperfusion-assisted hepatic metastasis of a colon adenocarcinoma in rodent model of disease [61]. In clinical reports, the local production of neutrophil elastase serves as an important indicator of poor prognosis in nonsmall cell lung cancer. The patients with a higher concentration of neutrophil elastase had a

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shorter survival than those with a low-level of enzyme [62]. Furthermore, neutrophil serine proteases could potentially promote tumour cell invasion acting via cleavage and activation of matrix metalloproteinases such as gelatinase A (MMP-2) [63]. MMP-2 is known to be involved in tumour invasion and in neoangiogenesis. Apart from neutrophil proteases, some other serine proteases from immune cells also appear to be involved in tumour invasion. Recently, Kankkunen and collegues have reported that tryptase-containing mast cells were concentrated at the tumour edge in malignant lesions of breast cancer, whereas the number of chymase-containing mast cells did not increase in that area [64]. Among non-immune cell proteases, human trypsins are known to be involved in tumour metastasis. For example, trypsins appear to promote tumour cell proliferation and metastasis in the setting of colorectal carcinogenesis [65]. Additionally, tumour-associated trypsins appear to increase tumour aggressiveness [66]. However, it is also important to point out that not all of pro-metastatic effects of trypsins are mediated via PARs, some of these effects are PAR-independent. Thrombin can also promote metastasis by activating PARs. Thrombin is known to be produced either by tumour cells or by tumour- associated platelets and to induce invasion and metastasis of various cancers. This pro-metastatic action of thrombin, that involves the migration of cells through matrix, appears to be mediated in large part by signalling events triggered by PAR1 [67][4][6]. Altogether, the data summarized above suggest a potential pro-metastatic activity for a number of well-known serine proteases. However, as already mentioned recent data indicate that some serine proteases appear to suppress tumour proliferation as well as invasion [58]. For example, dipeptidyl peptidase 4 (DPP4/CD26), a cell surface serine protease, is reported to suppress the malignant phenotype of melanocytic cells [68]. Kajiyama and colleagues demonstrated that over-expression of DPP4 upregulates E-cadherin and tissue inhibitors of metalloproteinases leading to reduction of the metastatic potential of ovarian carcinoma cells (SKOV3) [69]. Moreover, DPP4 is able to block the FGF2 signalling cascade and thus affect cell adhesion of prostate cancer cells [70]. However, the mechanisms underlying these effects of DPP4 are not yet fully understood. The glycosylphosphatidylinositol (GPI)-anchored serine protease, prostasin, also seems to suppress tumour invasion. This enzyme suppresses the invasion of breast cancer cells [71]. As yet, the mechanisms of this effect of prostasin also remain elusive. The kallikrein-related peptidase family of serine proteases (KLKs or “tissue kallikreins “), that includes both tryptic and chymotryptic-like enzymes, is thought to play a “dual“ role in tumour progression [57]. Although most reports have associated high kallikrein expression with a poor prognosis for patients [72][73][74], a few recent studies indicate that some KLK family members may serve as favourable prognostic indicators and may be able to suppress tumour invasion. For example, human kallikrein 3 (KLK3), commonly known as prostate-specific antigen (PSA) and long used as a marker for prostate cancer, may suppress tumour growth under certain circumstances. KLK3 may suppress tumour growth by activating transforming growth factor β (TGFβ) [75]. Furthermore, human KLKs might potentially affect tumour cell migration by regulating the activity of PARs since these enzymes are capable of either activating or dis-arming PARs 1, 2 and 4 [7][76]. KLK8 seems to suppress tumour cell invasiveness through its ability to modify the extracellular microenvironment by cleaving fibronectin. This cleavage suppresses integrin signalling and impairs cancer cell motility by inhibiting actin polymerization [77]. Altogether, kallikreins

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could potentially exert either cancer-promoting or cancer-inhibiting activities, depending on the type of expressed KLK, the tissue type and the tumour microenvironment. However to date, data supporting a tumour suppressor role for KLKs have come from studies done in vitro. Thus, further investigation using in vivo models will be required to assess the potential anti-tumour activity of the KLKs. Taken together, data obtained so far illustrate that serine proteases posses both prometastatic and tumour suppressive potential. Thus, the impact of these enzymes on tumour invasion is quite complex. Much work remains to be done to unravel the mechanisms whereby proteases can play this dual role. In this context, it is especially interesting to investigate the receptor-dependent effects of serine proteases at tumour cell invasiveness. The role of PARs in tumour cell motility is described in several recent publications. The level of PAR expression may potentially represent a useful prognostic factor in certain types of cancer. For, example the magnitude of PAR1 expression in samples of gastric cancers appears to correlate with the depth of wall invasion and peritoneal dissemination [78]. Additionally, PAR1 seems to be a marker (in immunohistochemical studies) of the recurrence risk in human malignant melanoma [79]. It is also important to mention that PAR1 may be particularly useful prognostic marker in melanoma studies because there is a gradation of the expression of this receptor that correlates with tumour thickness [79]. Furthermore, PAR1 is proposed to be involved in mediating the invasiveness of several types of cancer, including melanoma [80], breast cancer [67], colon cancer [81], kidney cancer [82], and prostate cancer [83]. It is also important to mention that thrombin is not the only enzyme that is capable of activating PAR1. Activated protein C appears to induce its effects on the invasiveness of breast cancer cell line cells (MDA-MB-231 and MDA-MB-435) via PAR1 activation [84]. Another PAR1 activating enzyme, that is also associated with tumour invasion, is matrix metalloproteinase 1 (MMP-1). This enzyme has for some time been linked to cancer progression due to its role in degrading of extracellular matrix. In addition, more recent data indicate that the invasiveness of human MDA and MCF-7 breast cancer-derived cells is triggered by MMP1-mediated activation of PAR1 [85]. Both stromal-derived and tumour cellderived MMP-1 could, in principle, activate PAR1 expressed on tumour cells [86] and on endothelial cells [87]. Activation of endothelial PAR1 can regulate vascular tension and could also promote endothelial cell activation potentially leading to enhanced adhesion of tumour cells to endothelium. Furthermore, PAR1 activation can stimulate mitogen-activated protein kinase kinase kinase (MAP3K) and via this mechanism promote actin cytoskeleton reorganization and cell migration [88]. In prostate-derived PC-3 cancer cells, PAR1 activation also initiates cytoskeletal reorganization. Stimulation of PAR1 by thrombin or a PAR1activating peptide induces the formation of a migratory phenotype in PC-3 cells [89]. The receptor-selective PAR1-activating peptide, TFLLRN-NH2, as well as thrombin is also able to stimulate cell migration of hepatocellular carcinoma cell lines (HEP-3B and SK-HEP1) as well as primary hepatocellular carcinoma cells [90]. PAR1 is reported to mediate the tyrosine phosphorylation of the epidermal growth factor receptor (EGFR) in human renal carcinoma cells via a mechamism that involved the trans-activation of a matrix metalloproteinase [82]. Apart from PAR1, similar cross-talk mechanism involving matrix metalloproteinase activity has been observed previously for other G-protein-coupled receptors [91]. The PAR1-EGF receptor cross-talk is involved in the regulation of renal carcinoma cell migration across a

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collagen barrier [82]. Thrombin-induced migration and adhesion of human colon carcinoma cells also appears to be associated with an activation of PAR1 expressed by tumour cells. PKCepsilon is involved in this effect as an essential signal transducer [92]. The examples discussed above illustrate the role of the best investigated receptor among PARs -PAR1- in the process of tumour cell migration/invasion and also shed some light on the signalling events underlying these effects. Apart from PAR1, other PARs are also reported to be involved in tumour invasiveness, but up to now, significantly less is known about their role in this process. Morris and co-authors demonstrated that factor VIIa and Xa-induced migration and invasion of breast cancer cells requires PAR2 as a mediator [93]. Enhanced expression of PAR2 by ovarian cancer cells serves as an indicator of poor prognosis for patients [94]. Trypsin, a classical enzymatic activator of PAR2, also serves as a prognostic factor in certain types of cancer. For example, patients with trypsin-positive colorectal cancers (CRCs) are reported to have shorter overall and disease-free survival than patients with trypsin-negative CRCs [65]. Trypsin-induced effects on proliferation and invasiveness of CRC appear to be associated, at least in a part, with PAR2 activation [95][65]. Moreover, PAR2 seems to be involved in cancer cell invasion in human pancreatic cancer [96]. On the other hand, it is important to mention recent work describing a tumour-protective role for PAR2 in epidermal skin tumours [97]. The role of PAR4 in tumour progression is just at the beginning of investigation. PAR4 expression appears to be a negative prognostic indicator for a patient survival in the case of non-small-cell lung cancers [98]. Recently, PAR4 selective agonist, AYPGKF-NH2, was demonstrated to stimulate the migration of hepatocellular carcinoma cells (HCC). This PAR4-triggered HCC cell migration in cooperation with PAR1 mediated effect on HCC cell migration contributes to thrombin enhanced migration of HCC cells [90]. However, the potential role that PAR4 may play in tumour progression and tumour cell invasiveness needs further investigation. Thus, although PAR1 activation appears to play a pro-metastatic role for the majority of tumour types, the role of PAR2 may be dual. PAR2 stimulation may trigger tumour promoting or tumour-suppressive activity, depending on the type of cancer. The value of PAR4 expression as a potential prognostic factor for patient survival and tumour progression and the potential effects of PAR4 activation on cancer cell motility and invasion remain very interesting topics for future investigation. Numerous reports have implicated another serine protease-associated receptor- uPAR- in the progression of different types of tumours. The uPA-uPAR system is known to be involved in the degradation of extracellular matrix, leading to tumour cell invasion and metastasis. Binding of enzymatically inactive uPA (pro-uPA) to its receptor, uPAR, converts the enzyme into its active form- active uPA. Active uPA is able to cleave the zymogen plasminogen to convert it into enzymatically active plasmin. Further, plasmin, though its own proteolytic function degrades extracellular matrix components and also activates matrix metalloproteinases, subsequently facilitating cancer cell migration [99][5]. Independent of uPA’s catalytic activity, uPAR is also involved in signalling cascades affecting cancer cell motility [5]. In several types of tumour (breast cancer, colorectal cancer, prostate cancer), increased levels of uPA and uPAR strongly correlate with a poor prognosis for patients [100][101][102]. On the other hand, soluble uPAR (suPAR) appears to antagonize cancer progression in some types of tumours. In animal models, suPAR was reported to reduce the

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growth and metastasis of MDA-MB 231 breast cancer cells and OV-MZ-6#8 ovarian cancer cells [103][104]. One possible mechanism explaining these effects may be that suPAR can function as a scavenger for uPA, like a high affinity antibody. According to another hypothesis, suPAR could inhibit tumour cell invasion and growth via a direct regulation of cell signalling events (urokinase-independent mechanism) [105]. The above examples illustrate a pro-metastatic as well as a tumour growth-promoting activity of uPA-uPAR in different types of cancers, despite the tumour-suppressive properties of suPAR. These tumour promoting properties of the uPA-uPAR complex allowed hypothesizing that naturally occurring uPA inhibitors might possess anti-tumour activity. Surprisingly, however, the effects of natural uPA inhibitors on the progression of certain types of tumours were the opposite of what was expected. The specific plasminogen activator inhibitor 1 (PAI-1) is one of the well-known naturally occurring inhibitors of uPA. Perhaps surprisingly, it was observed that the long-term survival of cancer patients with high levels of the PAI-1 was much worse than for individuals with lower PAI-1 levels [106]. Furthermore, endogenous protease inhibitors such as PAI-1 and TIMP-1 appear to promote tumour invasiveness rather than suppress this process [107]. There are some hypotheses concerning potential mechanism of the pro-metastatic effect of PAI-1 [106]. The ability of PAI-1 to modulate cell adhesion and migration, at least partially, may be related to its ability to detach cells from extracellular matrix [108]. Subsequently, this property of PAI-1 as a “de-adhesive” protein may promote the invasiveness of cancer cells. However, a more detailed investigation will be required to understand the link between high levels of PAI-1 and a poor prognosis for cancer patient survival. In summary, a role for serine proteases as regulators the motility of tumour cells is now well supported by data in the literature. The mechanisms involve both a direct targeting of specific substrates for either activation (e.g. chemokines; cytokines) or degradation (e.g. extracellular matrix) and an ability to trigger signalling via cell surface receptors (PARs or uPAR). Surprisingly, the impact of proteases appears to be “ bidirectional ”, either promoting or inhibiting tumour cell invasion. This complex dual role represents a challenge in targeting proteases for therapeutic purposes, since enzyme inhibitors may have effects that promote rather than block tumour growth in certain settings. Nonetheless, the selective inhibition of specific proteases and their receptors (e.g. thrombin, acting via PAR1) in certain settings may be of therapeutic benefit. Alternatively, in other circumstances it may be possible to take advantage of the anti-metastatic action of certain extracellular proteases. Clearly this area of focus is one that merits considerable attention in the future for developing novel anti-cancer therapies.

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Role of uPA-uPAR and PARs in the Motility of Neurons, Spermatozoa and Vascular Smooth Muscle Cells

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Actions of Proteases on Neurons Studies have just begun to evaluate the potential roles of serine protease-associated receptors, such as PARs and uPAR, in the motility of neuronal and glial cells during mammalian neonatal development or during post-traumatic tissue regeneration. In cell culture, growth cones of regenerating axons secret plasminogen activators (PAs), which are capable of activating plasmin that in turn, can facilitate growth cone movement due to the degradation of extracellular matrix components. Indeed, Nakajima and colleagues have reported that uPA-dependent activity is significantly increased in the facial nucleus of the rat facial nerve after its transaction [109]. Moreover, cells from dissociated 2d postnatal murine dorsal root ganglia neurons (DRGs) in culture show a 75- to 165 fold increase in tissue PA (tPA), uPA and uPAR while regenerating their axons. These mRNA increases are coincident with the period of maximal axonal outgrowth [110]. Further, Siconolfi and colleagues have demonstrated that after a surgical injury of sensory neurons in vivo, the PA system is rapidly induced and could play an important role in nerve regeneration [111]. Rather than promoting neurite outgrowth, thrombin for almost two decades now has been recognized as an agent that can cause neurite retraction, presumably acting via PAR1 [112] [113]. Debeir and colleagues have subsequently provided evidence that rat septal neurons express functional PAR1 [114][115]. In further studies, it was demonstrated that neuronal axons retract in response to PAR1 agonists, regulating the changes in actin-related cell motility [116]. Furthermore, several in vitro studies indicate a significant role for PAR1 in function of motor neurons. PAR1 agonists (thrombin as well as PAR1-AP) are reported to interrupt neurite outgrowth and induce apoptosis and degeneration in motor neuron cultures [117][118]. The activation of PAR1 also appears to inhibit neurite outgrowth from isolated DRGs after nerve damage [119]. Thus, a variety of data indicate that serine proteases could affect the neurite out growth and neuron migration acting not only directly (degradation of ECM compounds, which facilitates neurite or cell movement), but also as triggers of PAR-mediated signalling events. In the context of current knowledge, it is worth to investigate further the role played by PARs in the migration of neurons during brain development.

Receptors of Serine Proteases and Their Action on Sperm Both uPAR and PARs also appear to participate in motility of human spermatozoa. Functional PAR2 is reported to be expressed by human spermatozoa. The activation of this receptor by human recombinant tryptase or PAR2-AP (SLIGKV) results in a significant decrease of spermatozoa motility [120]. Mast cell tryptase-induced reduction the motility of human spermatozoa appears to be associated with activation of the mitogen-activated protein kinase (MAPK) pathway [121]. Altogether, these findings represent a very interesting and

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important research field since human mast cells are found in the male and female genital tract. Whether any other members of PAR family could affect the motility of human sperm remains unknown. A potential role of uPA-uPAR system in the motility of human spermatozoa is also still unclear despite a recent report about a positive correlation between the level of uPA in the seminal plasma and sperm motility [122].

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Proteases and the Regulation of Vascular Smooth Muscle Under normal physiological conditions, vascular smooth muscle cells (VSMCs) are differentiated cells specialized for constriction. However, in the case of vessel wall damage, contractile VSMCs can undergo a transition to a less-differentiated phenotype with the potential ability to migrate into the subendothelial space. Both the uPA-uPAR complex and PARs appear to be involved in vascular smooth muscle cell migration. Thrombin stimulation was reported to increase the mobility of VSMCs and also to enhance the cell surface expression of uPAR [123]. Moreover, both uPA and tPA have been demonstrated to augment smooth muscle cell migration and invasion in a plasmin-dependent manner [124]. It has been assumed that the binding of uPA to uPAR facilitates the migration of VSMCs. However, the signalling events underlying this effect of uPA has so far not been elucidated in detail. Work by Dumler and colleagues suggests that the uPAR-signaling complex utilizes at least two distinct signaling pathways, one employing a Jak/Stat cascade, possibly involved in cell migration and Src-kinase driven process that regulates other as yet undetermined cell responses [125]. Furthermore, it is important to mention that monocyte-produced uPA appears to stimulate the migration of human VSMCs in a co-culture model in vitro [126]. On the other hand, monocyte-expressed uPA is capable of inhibiting human VSMC proliferation potentially via Stat1 activation [127]. In addition to stimulating vascular smooth muscle cell migration and replication, proteases like thrombin that activate PAR1 can regulate vascular tension either by activating smooth muscle contraction or by promoting relaxation via the endothelial release of nitric oxide [128]. Altogether, these findings implicate proteasetriggered mechanisms in the regulation of vascular function in a variety of settings, which are potentially important for the development of different diseases in humans (atherosclerosis, for example). PAR activation can also stimulate the migration of smooth muscle cells. PAR2 contributes to VSMC migration, as demonstrated by the work of Marutsuka and colleagues who showed that both trypsin as well as a PAR2 activating peptide (SLIGKV) can promote VSMC migration in a concentration-dependent manner [129]. Moreover, the same study showed that the ability of the coagulation factor complex TF/FVIIa to induced migration of VSMC depends on PAR2 activation. In that study neither PAR1 nor PAR4 activating peptides were able to affect VSMC migration [129]. It is thus possible that PAR2 activation may play an important role in the migration of vascular smooth muscle cells in the setting of vascular injury. On the other hand, there is an increase in the production of prostaglandins PGE2 and PGI2 during vascular injury. Both of these prostaglandins are reported to reduce the expression of PAR1 on VSMCs [130]. Thus, the prostaglandins may play a counter-

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regulatory role in the process of VSMC migration by diminishing the potential effects of PAR1. In summary, the uPA-uPAR complex as well as PARs could potentially be involved in pathological processes associated with the motility of neurons, spermatozoa and vascular smooth muscle cells in vivo. These pathologies may include neurodegeneration, infertility, and atherosclerosis. Further studies are thus warranted to explore furthering details of the potential mechanisms whereby the uPA-uPAR complex and the PARs may participate in the pathophysiology of the three targets discussed in this section.

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Conclusion Abundant evidence now clearly demonstrates a complex role of serine proteases as well as protease associated receptors in regulating cellular motility. For some time, the degradation of extracellular matrix components was considered as of the principal mechanism whereby serine proteases promote cellular migration. This mechanism is linked to the ability of serine proteases to cleave and thereby activate matrix-degenerating enzymes (for example, plasminogen and matrix metalloproteases). However, recent studies have revealed novel mechanisms whereby serine proteases can affect the migration and matrix invasion of different cell types. For instance, these enzymes can act on polypeptide precursors to generate biologically active peptides with chemotactic activity (cleavage of proendothelins by chymase, for example). Moreover, serine proteases by themselves can serve as chemoattractant molecules for migrating leukocytes. Additionally, serine proteases are reported to cleave cell adhesion molecules, and, via such a cleavage to affect leukocyte recruitment. These are examples of receptor-independent effects of serine proteases on cellular motility. However, recent studies have shown that serine proteases are also capable of affecting leukocyte recruitment, tumour cell invasiveness, vascular smooth muscle cell migration and the motility of other cells by activating cell surface receptors, like the PARs or uPAR, to generate intracellular signals that drive the process. The role of PARs in the different aspects of cellular motility is just beginning to be fully appreciated. Despite the demonstrated effects of PARs on leukocyte motility in vitro and in vivo, the signalling events triggered after receptor activation that stimulate the process of migration require further investigation. Such investigation seems to be especially interesting since some bacterial proteases appear to affect leukocyte function via PARs [8]. Thus, the ability of pathogens to affect leukocyte recruitment and, subsequently, host response to infection via protease-PAR interactions merits serious consideration. If pathogen-derived proteases are indeed found to act via PARs to enhance pathogenicity, the enzymes and their receptors may prove to be novel therapeutic targets. In recent works it also has been demonstrated that tumour cell invasiveness involves activation of PARs. Although PAR1 activation appears to play a pro-metastatic role for the majority of tumour types, the role of PAR2 may be dual [67][131] [84][85][86][87][97][93][65]. The ability of non-classical proteases such as activated protein C and MMP-1 to activate PAR1 appears to play a significant role in cancer cell motility. Thus, stromal- and/or tumour-derived proteases (MMP-1, for example) may promote tumour

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cell invasiveness, at least in part acting via PAR1. Identification of other proteases with such properties will be of considerable therapeutic interest since this information would allow targeting the pro-metastatic process by inhibiting the both receptors and their activating enzymes. Altogether, serine proteases are able to utilize their proteolytic activity to affect cellular motility by degrading matrix proteins or by generating chemotactic peptides. On the other hand, serine proteases can act on motility behaviour of different cell types via receptordependent pathway, which involves cell surface receptors PARs and uPAR. These multiple actions of proteases (degradative and signal generating) could take place simultaneously in the tissue microenvironment in order to regulate many of the complex processes involved in cellular motility.

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[127] Kunigal, S., Kusch, A., Tkachuk, N., Tkachuk, S., Jerke, U., Haller, H., Dumler, I. (2003) Monocyte-expressed urokinase inhibits vascular smooth muscle cell growth by activating Stat1. Blood 102, 4377-83. [128] Laniyonu, A.A., Hollenberg, M.D. (1995) Vascular actions of thrombin receptorderived polypeptides: structure-activity profiles for contractile and relaxant effects in rat aorta. Br J Pharmacol 114, 1680-6. [129] Marutsuka, K., Hatakeyama, K., Sato, Y., Yamashita, A., Sumiyoshi, A., Asada, Y. (2002) Protease-activated receptor 2 (PAR2) mediates vascular smooth muscle cell migration induced by tissue factor/factor VIIa complex. Thromb Res 107, 271-6. [130] Pape, R., Rauch, B.H., Rosenkranz, A.C., Kaber, G., Schror, K. (2008) Transcriptional inhibition of protease-activated receptor-1 expression by prostacyclin in human vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 28, 534-40. [131] Even-Ram, S.C., Maoz, M., Pokroy, E., Reich, R., Katz, B.Z., Gutwein, P., Altevogt, P., Bar-Shavit, R. (2001) Tumor cell invasion is promoted by activation of protease activated receptor-1 in cooperation with the alpha v beta 5 integrin. J Biol Chem 276, 10952-62.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 187-207

ISBN: 978-1-60692-570-6 © 2009 Nova Science Publishers, Inc.

Chapter VI

Tuberin and Hamartin in Moving Breast Cancer Cells: Expression, Localization, and Function Marina A. Guvakova1,2‡ and William S.Y. Lee1 1

Department of Microbiology, 2Department of Surgery, School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA

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While the role of hamartin and tuberin complex as a negative regulator of protein translation and cell growth is well established, little information is available about the role of these proteins in regulating cell migration [1]. Hamartin, the protein encoded by the tuberous sclerosis complex (TSC) 1 gene, has been implicated in cell-matrix adhesion through its interaction with the ezrin-radixin-moesin (ERM) family of actin-binding proteins and regulation of the GTPase Rho [2]. Tuberin, encoded by the TSC2 gene, shows homology with Rap1 GTPase activating protein (GAP) and acts as a Rap1GAP in vitro [3]. Rap1, a member of the Ras oncogene superfamily, plays a role in the control of cell adhesion and movement [4]. Here, by laser-scanning confocal microscopy, we revealed that hamartin is present, whereas tuberin is excluded from lamellipodia of moving breast cancer cells stimulated with insulin-like growth factor I (IGF-I). The dissimilar intracellular patterns suggest that hamartin and tuberin may have independent roles in the moving cells. Hence we tested the hypothesis that tuberin regulates intracellular activity of Rap1 and thus might contribute to cell motile behavior. We showed that a partial knockdown of tuberin expression by TSC2 siRNA increased GTPloading of Rap1 in response to IGF-I. We then utilized a set of the tuberin mutants, containing or lacking a putative GAP domain, to investigate if tuberin decreases levels of active GTP-bound Rap1. As expected, tuberins with a truncated GAP domain had no ‡

Correspondence to: M.A.G. at the Division of Endocrine and Oncologic Surgery, Department of Surgery, School of Medicine, University of Pennsylvania, Philadelphia, PA 19104, Tel: 215-662-2449. Fax: 215-614-0765. email: [email protected]

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Marina A. Guvakova and William S.Y. Lee effect on Rap1 activation by IGF-I. Surprisingly, over-expression of the full-length tuberin had no effect on Rap1 either, whereas expression of a deletion mutant of tuberin, lacking the hamartin-binding domain (HBD) spanning a leucine-zipper (amino acids 8198) and a coiled-coil (amino acids 346-371), displayed Rap1GAP function. Further deletion of HBD and the central region of tuberin, including the coiled-coil (amino acids 1008-1021) failed to promote hydrolysis of Rap1-bound GTP. These data reveal that the region spanning amino acid residues 1-460 inhibits, whereas both the central regions of tuberin (amino acids 461- 1114) and the GAP domain (amino acids 1531-1758) promote Rap1 GTPase function of tuberin in vivo. We propose that tuberin may act as a Rap1GAP in the moving breast cancer cells; however, the interaction of hamartin with tuberin inhibits tuberin’s GAP activity towards Rap1.

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Introduction Cell adhesion is essential for maintenance of tissue integrity, whereas reduced adhesion is associated with tissue disorganization and cell migration. Loss of normal tissue organization is the hallmark of cancer [5]. It has been known for a while that mutations in the tuberous sclerosis complex 1 and 2 (TSC1 and TSC2) genes are responsible for an autosomal dominant disorder associated with the formation of benign tumors, hamartomas [6,7]. The development of hamartomas has been attributed to abnormalities in cellular proliferation, migration and adhesion in mesodermal- and ectodermal–derived tissues [6]. Genetic and biochemical studies revealed that the TSC2 and TSC1 genes encode the proteins tuberin and hamartin, which form a heterodimeric complex [8,9]. Hamartin, a protein of predicted molecular weight of 130kDa is highly expressed in skeletal muscle and to lesser extent in the heart, brain, placenta, pancreas, lung, liver and kidney. The highest expression of tuberin, a protein of approximately 200kDa, was found in tissue extracts from the brain, heart, and kidney [10]. The tuberin-hamartin protein complex is best known for suppression of cell growth and proliferation; however, ability of tuberin and hamartin to influence cell adhesion is probably just as significant. The overexpression of tuberin and hamartin in HEK293 human kidney epithelial cells resulted in the increased expression of E-cadherin and cell-cell aggregation [11]. In contrast, re-expression of hamartin in the transitional cell carcinoma (TCC) line with mutation in TSC1 gene appeared to induce migration in these epithelial cells [12]. Several studies suggested requirements for tuberin and hamartin in cell-matrix adhesion based on a role of these proteins in organization of the cytoskeleton, focal adhesions and modulation activity of the Rho family small guanosine triphosphatases (GTPases) [2,13,14]. Inhibition of hamartin expression by microinjection of antisense hamartin cDNA caused loss of stress fibers and disruption of cell-matrix adhesion, whereas re-expression of hamartin in cells lacking focal adhesions resulted in activation of RhoA, assembly of stress fibers and formation of focal adhesions. Since hamartin binds to the ezrin-radixin-moesin (ERM) family of actin-binding proteins [2], its role in cell-matrix adhesion is thought to be mediated through these interactions. Furthermore, in cultured cortical neurons, hamartin physically anchors the neuronal intermediate filaments to the actin cytoskeleton by binding with both neurofilament light chains and the ERM [15]. Studies produce conflicting reports on the functional outcome of re-expression of tuberin in TSC2 deficient (TSC2 -/-) cells [13,14].

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Overexpression of tuberin in MDCK epithelial cells and in tuberin-deficient ELT3 uterine leiomyoma cells increased cell attachment to the substratum in conjunction with reduced cell migration in ELT3 cells [13]. This morphological change was associated with a marked increase in RhoGTP, without any effect on Rac1GTP and Cdc42GTP levels. Another study reported a modest decrease in Rho activity, an increase in Rac1 activation, and loss of stress fiber formation in ELT3 cells overexpressing human tuberin chimeric protein tagged with green fluorescent protein (GFP) [14]. The latter study did not assess the effects of tuberin reexpression on either cell adhesion or migration. Thus, the mechanisms by which tuberin controls cell adhesive and motile behavior remains understood. Based on a sequence homology to Rap1 GTPase activating protein (GAP) [16], p130Spa1 [17] and Drosophila RapGAP1 [18], tuberin has been proposed to function as a GAP for the Ras-related small GTPases. Tuberin has been shown to have GAP activity in vivo toward Rheb and in vitro towards Rab5 and Rap1A. [19] The latter idea is based on the original finding of the DeClue group that the product of the TSC2 gene, tuberin, has Rap1GAP activity in vitro [3] . Tuberin and Rap1 have been shown to co-localize in the Golgi complex in vivo [20]; in accord with Golgi localization, a perinuclear and punctuate pattern of tuberin staining was detected in human tissues [10]. Rap1 acts as a switch that is activated by guanine nucleotide exchange factors (GEFs), such as C3G (CRK SH3-binding guanine nucleotide releasing protein), EPAC 1 and 2 (cAMP-regulated GEFI and II), DOCK4, PDZ (PDZ domain containing)-GEF1 and 2, CalDAG (calcium and diacylglycerol-regulated)GEFI and II. Rap1 inactivation is promoted by GAPs, including RapGAP (RapGAP-I, RapGAP-II) and Spa1 (Spa1, E6TP1/SPAR, SPA-L) family members [21]. In the inactive state, Rap1 binds guanosine diphosphate (GDP); in the active state, GDP is released while guanosine triphosphate (GTP) is incorporated. The function of GAPs is to promote hydrolysis of GTPase-bound GTP to GDP and return GTPase to the inactive state. Since Rap1 has a 10fold lower intrinsic GTPase activity than Ras [22], the action of Rap1GAP in regulating GTP-GDP cycle is essential to maintain the intracellular equilibrium of Rap1 activity. At the cellular level, activated Rap1 has emerged as a key regulatory switch linking signaling by hormones, growth factors, and cytokines to actin dynamics and adhesion receptors [4]. Specifically, Rap1 has a role in the regulation of cell-matrix adhesion and migration [4,2325]. Recent work provided data indicating that basal levels of Rap1 are essential for maintenance of mammary tissue integrity; whereas activated Rap1 may promote tissue disintegration and tumorigenesis in the breast [24]. Although both tuberin and hamartin are expressed in the breast epithelial cells, their functional roles in the breast have not been investigated. There is evidence that the TSC genes and their protein products are aberrantly expressed in human breast cancer and the expression of hamartin is associated with an unfavorable clinical outcome [26]. Furthermore, loss of heterozygosity in the TSC2 region on chromosome 16p13 in papillary breast tumors [27] and the reduced expression of tuberin in invasive versus noninvasive breast carcinomas [26] implicated tuberin in the pathogenesis of breast cancer in humans. Thus, findings on in vitro activity of tuberin as a Rap1GAP and the clinical relevance of reduced tuberin expression to the aggressive nature of breast cancer led us to hypothesize that tuberin may function as a GAP for Rap1 and hence control motile behavior of mammary epithelial cells. To test this hypothesis, we used three approaches: over-expression of the full-length tuberin, siRNA silencing of endogenous tuberin, and over-

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expression of tuberin mutants that have potential to be dominant-negative towards endogenous tuberin. Specifically, we investigated if tuberin decreases levels of active GTPbound Rap1 in vivo in resting and moving breast cancer cells.

Materials and Methods Cells and Reagents Human breast carcinoma MCF-7 cell line and its stable clones overexpressing the wild type (WT) and dead-kinase (DK) human IGF-IR were maintained in culture as described [28]. In the latter, point mutations of the Tyr 1131, Tyr 1135, Tyr 1136 in the receptor beta subunit render the receptor catalytically inactive [28,29]. Human non-tumorigenic MCF-10A cell line, obtained from Charles V. Clevenger (University of Pennsylvania) was cultured as suggested [30]. Human breast cancer cell lines T47 (HTB-133), BT-20 (HTB-19), MDAMB-231 (HTB-26) were purchased from the American Type Culture Collection (ATCC) and maintained according to its protocol.

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DNA Plasmids, siRNAs and Transfections The plasmids expressing EGFP-tagged constructs of full-length tuberin (pEGFP-TSC2), hamartin binding domain (HBD) of tuberin (pEGFP-TSC2-HBD), and tuberin lacking HBD (pEGFP-TSC2 -∆HBD) were kindly provided by Vera P. Krymskaya (University of Pennsylvania). The pEGFP-N-TSC2 and pEGFP-C-TSC2 encoding truncated N- and Cterminal tuberins were a gift from Daniel J. Noonan (University of Kentucky). The pEGFP control vector encoding EGFP was from Clontech Laboratories Inc. Transient transfections with EGFP-TSC2 constructs were performed with FuGENE 6 (Roche). For targeted siRNA delivery, cells were co-transfected with double stranded RNAs, Signal Silence TM Control siRNA, Tuberin/TSC2 siRNA (Cell Signaling Technology, Beverly, MA, CST), which specifically inhibits human tuberin expression, and pEGFP to control for efficiency of transfection. Transfections with siRNAs were performed with X-tremeGENE siRNA transfection reagent (Roche) according to the manufacturer’s protocol.

Western Blotting (WB) and Immunoprecipitation (IP) Cells were serum-starved in phenol red and serum-free DMEM/F12 containing 0.5 mg/ml bovine serum albumin (BSA). Serum-starved cells were stimulated with 100 ng/ml recombinant human IGF-I (BACHEM, Torrance, CA) and analyzed by immunoblotting. Cells lysed in a 1% Triton X-100 containing buffer previously described [28] were subjected to SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and proteins were transferred to Hybond ECL nitrocellulose (Amersham Pharmacia Biotech). The blots were probed with antibodies against hamartin/TSC1, phospho-Akt-Ser 473 and Akt (polyclonal; CST); Rap1

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(polyclonal 121), tuberin (polyclonal, c-20), IGF-IR β subunit (polyclonal, c-20) and extracellular signal-regulated kinase (ERK)1/2 (polyclonal k23) (Santa Cruz Biotechnology, Inc., Santa Cruz, CA); phospho-mitogen-activated protein kinase (MAPK) (clone E10, New England Biolabs, Beverly, MA), and BD Living Colors TM A.v. (monoclonal JL-8; Clontech Laboratories Inc.) to GFP. Proteins were precipitated from 500 μg of total protein extracts in 500 μl of HNTG buffer (20 mM HEPES pH 7.5, 150 mM NaCl, 0.1% Triton X-100, and 10% glycerol) overnight at 4°C. For IP of tuberin and hamartin, extracts were incubated with 50 μl of protein A-agarose (Santa Cruz Biotechnology, Inc) and 5 μg of tuberin (c-20) and hamartin/TSC1 antibodies, respectively. The immunoprecipitated proteins were washed three times in HNTG and resolved by SDS-PAGE. To visualize primary antibody-bound proteins, secondary antibodies linked to horseradish peroxidase (Oncogene Research) and ECL detection reagents (Amersham Biosciences, UK) were applied. The chemiluminescent intensity of bands was digitized and quantified using the Image Analyzer LAS-1000 Plus system and the Image Reader LAS-1000 Lite version 1.0 software (Fuji, Tokio, Japan).

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Immunofluorescence Microscopy Cells were fixed in methanol or 3.7% formaldehyde, blocked in 5% BSA, stained with primary antibodies against tuberin (c-20), hamatrin/TSC1, and actin (monoclonal, AC-40; Sigma), followed by anti-rabbit Rhodamine Red TM -conjugated secondary IgG (Jackson ImmunoResearch Laboratories, Inc.) and anti-mouse fluorescein isothiocyanate (FITC)conjugated secondary IgG (Vector Laboratories, Burlingame, CA). F-actin was visualized in formaldehyde fixed cells by staining with 1μg/ml tetramethylrhodamine B isothiocyanate (TRITC)-phalloidin (Sigma-Aldrich) for 30 minutes. Immunostaining was analyzed using a Nikon Eclipse E600 MRC 1024 confocal laser-scanning microscope (Bio-Rad) with a Plan Apo 60x/1.4 oil objective lens (Nikon) and the Bio-Rad LaserSharp 2000 software. When primary antibody was omitted, no specific staining was detected. Fluorescence intensities were quantified along radial lines drawn from an approximate geometric center of the nucleus to the cell periphery using the processing function of the Bio-Rad LaserSharp 2000 software. The representative graphs of pixel intensities plotted as a function of distance are shown.

Rap1-GTPase Pull-Down Assay The active GTP-bound form of Rap1 (Rap1GTP) was purified from live cells by affinity precipitation using a glutathione S-transferase (GST) fusion protein corresponding to amino acids 788-884 of human RalGDS Rap1-binding domain (RBD-RalGDS). Clarified cell lysates were immediately incubated with 20 μg of GST fusion protein of RBD- RalGDS bound to glutathione agarose beads (Upstate Cell Signaling Solutions) for 45 min at 4°C. The bead-bound complex was washed three times in HNTG buffer and then the complex was removed from the beads in reducing Laemmli buffer. Protein samples were analyzed by 12% SDS-PAGE and immunoblotting with Rap1 antibody. To determine the total quantities of Rap1 in samples, 50 μg of total cellular protein was analyzed in parallel. The relative activity

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of Rap1 is the ratio of GTP-bound Rap1 to the amount of total Rap1 detected in the same sample. The statistical evaluation of the results was performed using a single-factor analysis of variance (ANOVA) with a significance level of p450 kDa) was larger then a sum of molecular weights of one molecule of tuberin and one molecule of hamartin [37]. In summary, although hamartin forms a stable complex with tuberin, there is a tuberinfree pool of hamartin in breast cancer cells. Furthermore, in moving breast cancer cells hamartin, independently of tuberin, translocates to actin-enriched lamellipodia. We also found that a deletion mutant of tuberin, lacking HBD, inhibited IGF-I-induced Rap1 activation and lamellipodia formation. We propose that tuberin may act as a Rap1GAP in lamella of moving breast cancer cells. However, interaction of hamartin with tuberin is likely to inhibit, whereas interaction of the putative protein with the central part of tuberin may promote tuberin’s GAP activity towards Rap1 in vivo. Together, findings in this chapter provide a novel functional link between tuberin, hamartin, and migration of breast cancer cells.

Acknowledgement Supported by the University of Pennsylvania Research Foundation and the American Cancer Society IRG# 78-002026 to M.A.G.

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[46] Manning, B. D. (2004). Balancing Akt with S6K: implications for both metabolic diseases and tumorigenesis. J.Cell Biol. 167, 399-403. [47] Inoki, K., Li, Y., Xu, T., and Guan, K. L. (2003). Rheb GTPase is a direct target of TSC2 GAP activity and regulates mTOR signaling. Genes Dev. 17, 1829-1834. [48] Potter, C. J., Pedraza, L. G., Huang, H., and Xu, T. (2003). The tuberous sclerosis complex (TSC) pathway and mechanism of size control. Biochem.Soc.Trans. 31, 584586. [49] Tee, A. R., Manning, B. D., Roux, P. P., Cantley, L. C., and Blenis, J. (2003). Tuberous sclerosis complex gene products, Tuberin and Hamartin, control mTOR signaling by acting as a GTPase-activating protein complex toward Rheb. Curr.Biol. 13, 1259-1268. [50] Ma, L., Chen, Z., Erdjument-Bromage, H., Tempst, P., and Pandolfi, P. P. (2005). Phosphorylation and functional inactivation of TSC2 by Erk implications for tuberous sclerosis and cancer pathogenesis. Cell 121, 179-193. [51] Guvakova, M. A. (2007). Insulin-like growth factors control cell migration in health and disease. Int.J.Biochem.Cell Biol. 39, 890-909. [52] Daumke, O., Weyand, M., Chakrabarti, P. P., Vetter, I. R., and Wittinghofer, A. (2004). The GTPase-activating protein Rap1GAP uses a catalytic asparagine. Nature 429, 197201. [53] Manning, B. D. and Cantley, L. C. (2003). Rheb fills a GAP between TSC and TOR. Trends Biochem.Sci. 28, 573-576. [54] Liu, L., Schwartz, B. R., Tupper, J., Lin, N., Winn, R. K., and Harlan, J. M. (2002). The GTPase Rap1 regulates phorbol 12-myristate 13-acetate-stimulated but not ligandinduced beta 1 integrin-dependent leukocyte adhesion. J.Biol.Chem. 277, 40893-40900.

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In: Cell Movement: New Research Trends Editors: T. Abreu and G. Silva, pp. 209-226

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Chapter VII

Role of Chemokines in Colorectal Cancer and Metastasis Kathrin Rupertus1, Otto Kollmar1, Michael D. Menger2 and Martin K. Schilling1 1

Department of General, Visceral, Vascular and Pediatric Surgery and 2 Institute for Clinical and Experimental Surgery, University of Saarland, Homburg/Saar, Germany

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Summary Background: Chemotactic cytokines (chemokines) are inflammatory cytokines that stimulate the migration of distinct subsets of cells, including leukocytes, lymphocytes and tumor cells. The chemokines MIP (macrophage inflammatory protein)-2 and SDF (stromal cell-derived factor)-1, which are both members of the CXC-chemokine superfamily, are supposed to have an important impact on tumor progression and metastasis. In our recent studies we investigated the chemotactic response of CT26.WT colorectal cancer cells on MIP-2 and SDF-1 in vitro and the influence of these chemokines on tumor growth in vivo. Material and methods: Using flow cytometry and chemotaxis chambers, we investigated the expression of the chemokine receptors CXCR2 and CXCR4 in vitro and the chemotactic response of CT26.WT colon carcinoma cells. To determine the influence of chemokines on tumor growth in vivo, we analyzed the growth of CT26.WT-tumors after exposure to MIP-2 and SDF-1. Additionally, we investigated the growth characteristics of CT26.WT tumors which were implanted into the left liver lobe of BALB/c mice after exposure to MIP-2 or after treatment with a neutralizing anti-MIP-2 antibody. MIP-2 and tumor growth: Our studies revealed that in vitro 98.8% of the CT26.WT cells were CXCR-2 receptor positive and showed a dose-dependent migration along a MIP-2 gradient. MIP-2 also promoted angiogenesis, hepatic engraftment and tumor growth of liver metastases, as well as tumor growth of established extrahepatic tumors. Blockade of MIP-2

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inhibited the augmentation of angiogenesis and metastatic tumor growth of intrahepatic tumors after liver resection and suppressed engraftment of CT26.WT tumor cells at extrahepatic sites. These results indicate a significant role of the chemokine MIP-2 during tumor engraftment, progression and metastasis and point out to the promising approach of targeting the MIP-2/CXCR2 pathway for anti-tumor therapy. SDF-1 and tumor growth: Despite a lower surface expression of the SDF-1 receptor CXCR4 (31.5%), CT26.WT cells showed a significant and dose-dependent migration in response to a SDF-1 gradient. Exogenously applied SDF-1 was capable to stimulate the growth of already established extrahepatic CT26.WT tumors. It also induced an angiogenic response with persistent changes in tumor vasculature. These results also encourage for further investigation on the influence of chemotactic cytokines in tumor growth and may also be useful to elucidate the pathophysiology of other diseases which are related to angiogenesis and cell migration. Conclusion: The results of our studies demonstrate that chemotactic signaling does not only contribute to the metastatic spread of tumor cells but is also essential for tumor engraftment and progression. Therefore, chemotactic cytokines which are a part of the tissuespecific environment could be promising targets for anti-tumor therapy.

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Directed Tumor Metastasis Colorectal cancer is one of the leading causes of cancer-related mortality among men and women worldwide. The death of the patients is usually the result of the uncontrolled metastatic spread of the tumor. Therapeutic options in the case of unresectable metastases are confined to non-curative chemotherapeutic treatment. As patients often present with advanced primary tumors, more than 50% of them suffer from synchronous or metachronous metastases1-4. The metastatic process consists of several steps which all must be successfully completed until a metastatic tumor becomes clinically detectable5. Therefore, only a small subset of metastatic tumor cells will survive, proliferate and finally form a solid tumor6. Although much research efforts have been made during the last decades, we are far from understanding the metastatic process as a whole. In the case of colorectal malignancies, the liver is the most common site of hematogenous metastasis. There is some evidence that this metastatic pattern is not only a result of hemodynamic conditions given by the portal circulation but is also directed by “homing” mechanisms7. This hypothesis is not new. It is mainly based on the so called “seed and soil”-theory which was first published by Stephen Paget in 1889. Paget postulated that metastatic tumor growth (“seed”) in host tissues (“soil”) can only occur under appropriate, tissue specific conditions8. Among the tissue-specific factors, influencing tumor growth characteristics, chemokines are currently under intensive investigation. Chemokines are chemotactic cytokines of small molecular weight that stimulate migration of a variety of cells including leukocytes, lymphocytes and tumor cells. They are the ligands of the chemokine receptors which represent a familiy of seven transmembrane domain, G-protein-coupled receptors on the cell surface 7,9,10. Chemokines and their receptors can be further divided into subfamilies

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according to their protein structure. One major chemokine subfamiliy consists of the socalled CXC-chemokines which are of special interest in cancer research9. As tumor metastasis is dependent on cell motility, chemokines as factors influencing cell migration have soon become candidate targets for research upon tumor metastasis11. Several studies already displayed a closed correlation between the expression patterns of chemokine receptors on neoplastic cells and their corresponding ligands in organs to which these tumors commonly metastasize12.

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Macrophage Inflammatory Protein (MIP)-2 The prototypic CXC chemokine is Macrophage inflammatory protein (MIP)-2 as the murine functional analogue of human IL-813. It acts as a potent neutrophil chemoattractant, contributes to wound healing and mediates inflammatory reactions through interaction with its receptor, CXCR214,15. The results of certain experimental work indicate that human IL-8 also stimulates tumor cell motility and migration16. To further investigate the influence of MIP-2 on tumor growth and metastasis in colorectal cancer, we used a well established colorectal cancer cell line – CT26.WT – for our in vitro and in vivo experiments. The CT26.WT cell line is a chemically induced, undifferentiated adenocarcinoma of the colon, syngeneic with the BALB/c mouse. When implanted into BALB/c mice, almost 100% of the animals will develop solid tumors with highly aggressive and metastatic growth characteristics17. Using flow cytomety, we found that 98.8% of the CT26.WT cells derived from cell culture expressed the CXCR2 receptor on the cell surface. Therefore, it was not surprising that these cells also migrated along a MIP-2 gradient in an in vitro chemotactic assay. In this assay, we used 24-well chemotaxis chambers. The wells were separated by polyvinylpyrrolidone-coated polycarbonate filters with an 8µm pore size. MIP-2 solutions with different concentrations were added to the lower wells, whereas a suspension of CT26.WT cells was put into the upper wells. Cells that had migrated into the lower wells after 24 hours of incubation were collected and counted by flow cytometry or, if they were adherent to the lower surface of the filters, they were stained and quantified under the microscope. We found that MIP-2 induced an increase of cell adhesion at the polycarbonate filters already at a dose of 0.1nM with a most pronounced adhesion at a dose of 10 and 100nM as compared to controls (4-fold and 6-fold increase respectively). Interestingly, a further increase of the MIP-2 concentration (200 and 400nM) markedly attenuated the adhesive response. In accordance with these results, MIP-2 stimulation also increased the number of cells that had migrated through the filter into the lower wells of the chemotaxis chamber, indicated by a 2 to 4-fold increase after low dose stimulation (0.1 to 100nM). Challenge with higher doses (200 to 400nM) even resulted in an exponential increase (9 and 18-fold) when compared with PBS controls. Under control conditions only a few adherent and migrated cells could be detected18.

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MIP-2 and Colorectal Liver Metastasis Because of the strong chemotactic response of the tumor cells after MIP-2 exposure, we further investigated the influence of MIP-2 on metastatic colorectal tumors. Elucidating the impact of MIP-2/CXCR2 signaling in colorectal malignancy, seems to be clinically important because previous studies on colorectal diseases have revealed an association between an increase of the expression of human IL-8 – the functional analogue of the murine MIP-2 – and the induction and progression of colorectal carcinoma19. As the liver is the most common site of colorectal metastasis, we used an in vivo model that had previously been standardized by our group20. For that purpose, BALB/c mice were anesthesized and laparotomized through a midline incision. The left lateral liver lobe was gently mobilized and a cell suspension containing 1x105 CT26.WT cells was implanted under the capsule of the lower surface. MIP-2 at different concentrations was applied under the subcapsular space with a distance of 2 to 5mm close to the tumor cell implants. The puncture sites were sealed with acrylic glue as to avoid puncture site seeding. The left liver lobe was repositioned anatomically into the peritoneal cavity and the abdominal wall was sutured. Seven days after tumor cell implantation, intravital fluorescence microscopy of the liver tumors was performed. For that purpose, animals were again anesthetized and relaparotomized. The left liver lobe was exteriorized and placed on an adjustable stage as to position the lower surface of the lobe horizontally to the microscope. The surface of the lobe was covered with a glass coverslip. After fluorescent labeling it was possible to study angioarchitecture and microvascular perfusion of the tumors, as well as leukocyte adhesion and apoptotic cell death. After injection of MIP-2 at a concentration of 10nM, tumor volume significantly increased by 14-fold when compared to control animals. At 100nM, tumor volume was further increased up to 27-fold. Interestingly, higher doses of MIP-2 (1000nM) were less effective and led to tumor volumes comparable in size to those of animals treated with 10nM of MIP-2. Because it was previously shown that high doses of MIP-2 were capable of inducing programmed cell death we concluded that the delayed tumor growth might be due to an increase of tumor cell apoptosis21. Indeed, we found a 4-fold increase of nuclear condensation and fragmentation after 1000nM MIP-2 using intravital microscopy. Tumor cell proliferation, as determined by immunhistochemical staining of the proliferation marker PCNA, was not influenced in seven days old tumors after MIP-2 exposure. CXCR2 expression, especially at the tumor margin, significantly increased after exposure to MIP-2 when compared to control tumors. This effect was most pronounced after treatment with 10nM MIP-2. In the context of other studies, we therefore assume an association between the magnitude of CXCR2 receptor expression and the metastastic potential of the colon carcinoma cells19,22. The fact that almost 100% of the CT26.WT cells were CXCR2 positive would therefore indicate that this cell line has to be considered as highly aggressive by the criteria of growth rate and metastatic spread. In vivo, approximately 40% of the cells of non-treated CT26.WT tumors showed positive CXCR2 staining 7 days after tumor cell implantation. This reduction can probably be explained by the fact that solid tumors in vivo consist of a variety of non-malignant cells including endothelia, fibroblasts and stomal cells

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that do not necessarily express CXCR2. Our study indicates that over a 7-day period, in vivo exposure to MIP-2 increases the expression of CXCR2. This can be due to receptor recycling which may provide the basis for efficient MIP-2 signaling. Because receptor expression seems to be required for the proliferative action of chemokines on tumors23, the increased in vivo expression of CXCR2 may represent the mechanism of the MIP-2-associated increase of tumor growth. As tumor progression and increased local tumor growth are usually associated with an angiogenic response, we analyzed the angioarchitecture of the intrahepatic tumors using intravital microscopy. We found that tumors were surrounded by an area, where the fluorescent marker sodium fluorescein did not extravasate into tissue and which therefore appeared dark in comparison with the adjacent tissue. In this area, called angiogenic front, we found a network of chaotically arranged, newly developed capillaries and large draining venules. MIP-2 at concentrations of 100 and 1000nM, but not 10nM, significantly increased the size of the angiogenic front and the density of draining venules (2-fold and 4-fold respectively) when compared to controls. Interestingly, MIP-2 at any concentration significantly increased capillary density within the tumors (5 to 6-fold). The stimulated angiogenic activity in MIP-2-treated tumors was also associated with a significant enlargement of tumor capillaries within the tumor margin after exposure to 100 and 1000nM MIP-2 when compared to controls and to animals treated with lower doses of MIP-2. In all tumor margins, the observed capillary diameters were higher than in the adjacent normal liver tissue including the area where the MIP-2 solution was injected 7 days before. All these observations concerning tumor angiogenesis clearly show that local exposure to MIP-2 induced persistent changes in angioarchitecture with overall stimulation of angiogenic activity. Of interest, these effects were observed after only one single application of MIP-2. These findings were highly interesting because there is still little information concerning the proangiogenic activity of MIP-2 in tumor growth. Because MIP-2 as an inflammatory cytokine is known to induce a chemotactic response of leukocytes, we analyzed the leukocyte adherence within the newly formed tumor capillaries. We found significantly increased numbers of adherent leukocytes after treatment with MIP-2, particularly at concentrations of 10 and 100nM. Challenge with higher doses again attenuated the observed effect. Leukocytes are known to express vascular endothelial growth factor (VEGF) which is one of the most important angiogenic factors in malignant and non-malignant diseases. Therefore the angiogenic effect of MIP-2 we detected in our study is highly likely to be the associated with an increased release of VEGF by the accumulated leukocytes within the tumor vasculature24. Vasodilatation, which is also known to be associated with VEGF, was also found increased after MIP-2 exposure. This finding further supported the hypothesis, that MIP-2 induced tumor angiogenesis is mediated by VEGF. Finally, considering the direct chemotactic effect on CT26 cells in vitro, we analyzed whether satellite metastases could be detected within the area where the chemokine solution had been applicated. Interestingly, no satellite metastases could be observed18.

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Studies on MIP-2 in a Model of Extrahepatic Colorectal Metastasis Even if our studies on colorectal liver metastasis had already revealed interesting aspects concerning the putative mechanisms by which MIP-2 may contribute to metastatic tumor growth, we intended to transfer the setting to a chronic tumor model. With the use of the dorsal skinfold chamber assay, we achieved studying the early stages of tumor development including early angiogenesis repetitively over a period of 14 days using intravital microscopy. The chamber consists of two symmetrical titanium frames, which were positioned to sandwich the extended double layer of the dorsal skin. One layer was completely removed in a circular area of 15mm in diameter. The remaining layers, consisting of the epidermis, subcutaneous tissue and striated skin muscle, were covered with a glass coverslip incorporated into one of the titanium frames25. After a 48-hour recovery period, the coverslip of the chamber was temporarily removed and 1 x 105 tumor cells were implanted onto the surface of the striated muscle tissue within the chamber26. To allow repetitive intravital microscopic analyses without additional application of fluorescent markers, CT26.WT cells had been transfected to express green fluorescent protein (GFP). Because of our finding, that MIP-2 at a concentration of 100nM had the most pronounced effect on tumor growth and angiogenesis of hepatic colorectal metastases, we used this concentration for our studies on extrahepatic tumors. Local application of MIP-2 was done at 5 days after tumor cell implantation, where a small established tumor had grown in the dorsal skinfold chamber. As we had already observed in intrahepatic tumors, MIP-2 treatment provoked a significant acceleration of tumor growth, which was found most pronounced 7 and 9 days after chemokine application. This increased tumor growth was due to a stimulation of tumor cell proliferation that was shown after immunhistochemical staining of tumors which were harvested 9 days after MIP-2 application. Whereas control tumors without MIP-2 treatment displayed a proliferation rate of ~40% at this time point, the amount of proliferating tumor cells increased up to ~80% after exposure to MIP-2. Concerning CXCR2 expression, we found that, within the tumor margin, the majority of the cells were CXCR2 positive. In contrast, within the tumor center only a minor part of the cells could be found to express CXCR2. Local treatment with 100nM MIP-2 did not affect CXCR2 expression in the tumor margin, but significantly increased the number of CXCR2positive cells within the tumor center. Because growth activity of tumors is usually highly pronounced in the tumor margin but not in the tumor center, the significantly higher CXCR2 expression in the tumor margin may underline the important role of this chemokine receptor in tumor growth. The finding that CXCR2 expression was not affected after local application of MIP-2 may be due to the fact that it was already maximally upregulated without MIP-2 stimulation. On the contrary, local exposure to MIP-2 was associated with an upregulation of CXCR2 in the tumor center as we had already observed in intrahepatic tumors. Underlining the hypothesis that an increasing CXCR2 expression is associated with invasiveness and malignancy, histological examinations revealed a more pronounced muscule infiltration by tumors which had been exposed to MIP-2.

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Taken together, all these results demonstrate, that MIP-2 functions as an autonomic determinant for tumor growth, because it is capable of upregulating its binding receptor which is required for adequate signaling. Contrary to our observations in hepatic metastases, MIP-2 application did not stimulate angiogenesis of the established extrahepatic metastases. The onset of the angiogenic response was not significantly different between the two groups whereas the capillary density of newly formed tumor vessels was even lower in the MIP-2 than in the control group. As we ascribe enhanced angiogenic activity after MIP-2 exposure to a release of VEGF, these results indicate that, in contrast to developing tumors, MIP-2 in established tumors does not provoke an increased release of VEGF. Accordingly, neither an increase of microvessel diameter nor an elevation of microvascular permeability that would have pointed towards a VEGF effect could be detected27.

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Liver Resection and Tumor Cell Migration Based on our findings concerning the importance of the chemokine MIP-2 on the outgrowth and progression of colorectal metastases, we made efforts to find possible clinical situations in which targeting chemotactic signaling could provide a therapeutic strategy. In the case of patients with colorectal liver metastases, surgical resection can currently be considered as the only curative treatment1,28. Modern surgical strategies from major hepatobiliary centers have demonstrated that hepatectomy of as much as 70% of the liver can be performed with a mortality rate of less than 5%29-32. Although it is well recognized that the liver completely regenerates after major hepatectomy, the effect of hepatic regeneration on intra- and extrahepatic tumor growth is still controversially discussed with a considerable amount of studies demonstrating an acceleration of intra- and extrahepatic tumor growth after liver resection33-37. These results are contrasted by other studies, which have reported no effects or even regression of tumor metastases after hepatectomy34,38,39. Because of these controversial results, we analyzed the effect of minor (30%) and major (70%) hepatectomy on tumor engraftment, neovascularization and tumor cell migration using the dorsal skinfold chamber assay. We found that tumor growth of CT26.WT-GFP tumors was stimulated dependent on the extend of liver resection. Whereas after minor hepatectomy only a slight increase of extrahepatic tumor growth could be observed, major hepatectomy resulted in a rapid acceleration of tumor growth. Accordingly, we found a massive increase of the angiogenic activity after major hepatectomy. These findings were not surprising because a considerable number of different hepatotrophic factors, which regulate the restoration of the liver cell mass after hepatectomy display angiogenic properties, such as VEGF and hepatocyte growth factor (HGF)40-42. But besides the massive stimulation of tumor growth and angiogenesis, we also observed a significant increase of tumor cell migration, especially after major hepatectomy. After minor hepatectomy, only a slight increase of tumor cell migration was observed. Taken together with the results obtained from the analysis of the increase of tumor size, we assumed a closed correlation between the stimulation of tumor cell migration and the stimulation of solid tumor growth. As it has been shown that chemotactic cytokines, such as ENA-78 and

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MIP-2 also participate in liver regeneration and that blockade with antibodies or deletion in gene-targeted animals significantly impaired liver regeneration43,44, it seems very likely that increased chemotactic signaling after liver resection contributes to an acceleration of tumor growth. The clinical implication of this observations is, that patients with colorectal liver metastases who have undergone surgical resection often present with recurrent intra- and extrahepatic metastases45. One possible source of recurrent colorectal metastasis are nests of tumor cells or dormant micrometastases with diameters