Adhesive Interactions in Normal and Transformed Cells
wwwwwwwwwwwwwwww
Yury A. Rovensky
Adhesive Interactions in Normal and Transformed Cells
Yury A. Rovensky, M.D., Ph.D., D.Sci. Former Leading Researcher at Cancer Research Center of the Russian Academy of Medical Sciences Moscow, Russia
[email protected] ISBN 978-1-61779-303-5 e-ISBN 978-1-61779-304-2 DOI 10.1007/978-1-61779-304-2 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011934257 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)
To my wife Tanya with love
wwwwwwwwwwwwwwww
Preface
The ability of tissue cells to be attached to each other and to the surrounding solid substance (extracellular matrix) is a pivotal regulator of major cellular functions such as proliferation, responses to growth-stimulating factors, cell survival, differentiation, and migration of cells in an organism. Therefore, the cellular adhesive interactions play a critical role in basic biological processes such as formation of tissues and organs in embryonic development, maintenance of structural integrity of all tissues in an adult organism, and tissue regeneration and remodeling. The adhesive interactions are also involved in inflammation and degeneration processes, which are at the basis of many diseases. As a result of oncogenic transformation, the adhesive interactions of transformed cells are significantly altered. In the pathological behavior of malignant tumor cells, significant weakening of their ability to adhere to each other, to normal cells, and to the extracellular matrix, plays a key role. Alterations in these adhesive interactions form the basis of invasion and metastasis of malignant tumors. Therefore, the understanding of mechanisms of cellular adhesive interactions and their alterations in malignant tumors is very important in both biological and medical aspects. Adhesive Interactions in Normal and Transformed Cells starts with the description of molecular composition of the extracellular matrix, which tissue cells adhere to. The matrix proteins that are bound with the specific cell surface receptors resulting in the cell-matrix adhesion are also discussed. Several sections are devoted to the cytoskeleton systems. Particular attention is given to the actin filaments and microtubules that play a pivotal role in cell-extracellular matrix and cell–cell adhesive interactions, and also in cell migration. The formation, regulation, and dynamics of these cytoskeleton systems are examined.
vii
viii
Preface
Different types of pseudopodia that are formed and used by cells as “driving organs” during cell spreading and cell migration are described. Various types of specific adhesion structures formed by cells in order to attach to the extracellular matrix are considered. Attention is given to focal adhesions (focal contacts), to their structure, regulation, and dynamics, which play a critical role in cell migration. Several sections are devoted to the intracellular signal transduction pathways. The signaling pathways are triggered by the extracellular molecules (ligands) that bind to specialized cell surface receptor proteins. Different types of cell surface receptors are characterized. Particular attention is given to integrin receptors, which as components of focal adhesions play a key role in cell-matrix attachment and also fulfill functions of transducers of intracellular signals. Different integrin receptormediated signaling pathways that determine and control cell morphology, proliferation, survival, and locomotion are considered. Also, the growth factor receptor-mediated mitogenic and morphogenic signaling pathways are examined. Special attention is given to significant alterations in the integrin mediated cellmatrix adhesion caused by oncogenic transformation of the cells. The consequences of these alterations manifested in such typical traits of transformed cells as weakening of the cell-matrix adhesion, “anchorage independence”, constitutive mitogenic activation, escape from anoikis, and high locomotory activity are considered. The movement of fibroblastic cells and different factors involved in the cell locomotion machinery are considered. These factors include actin cytoskeleton reorganizations and microtubule dynamics, the phenomenon of “contact inhibition of cell locomotion”, and dynamic regulation of focal adhesions during cell locomotion. The morphogenic action of soluble growth factors resulting in cell locomotion is also examined. Several sections are devoted to fundamental alterations in cell locomotion machinery caused by oncogenic transformation of the cells. These alterations apply to the pseudopodial activity and focal adhesion formation in transformed cells, and also their sensitivity to growth factors. The ability of cells to respond to the adhesion heterogeneity or various geometrical configurations (topography) of the extracellular matrix surfaces is discussed in detail. The topographic cell responses to cylindrical surfaces of high curvatures or the surface reliefs of various kinds (such as nanoscale or microscale linear grooves, holes, or vertical rods) are examined. These responses apply to the cell shape, locomotion, and other cellular functions. The mechanisms of these cell responses are discussed. The alterations in the topographic cell responses caused by oncogenic transformation of cells are considered. In particular, alterations of the cell shape, changes in the direction of cell migration, and alterations in the functional activities as a result of oncogenic transformation are described. Last chapter of the book is devoted to the intercellular adhesive interactions. The compositions of several types of the intercellular adhesion structures are described. Particular attention is paid to the adherens junctions, their structure and dynamic regulation, which is the basis of cell rearrangement and tissue integrity maintenance.
Preface
ix
A critical contribution of cadherin receptors and local actin cytoskeleton to the regulation of cell–cell adhesion is examined. Signaling pathways coupling cadherinmediated intercellular contacts to cell proliferation are considered. The cell–cell adhesion alterations caused by oncogenic transformation of the cells are further examined. These alterations result in uncontrolled proliferation of malignant tumor cells, their inability to form orderly tissue structures, cancer invasion, and metastasis. Adhesive Interactions in Normal and Transformed Cells is based on modern scientific data and includes the results of the author’s long-term research. It is intended for researchers, postdocs, undergraduate, and graduate students, whose scientific interests are in the fields of cell biology, cancer biology, cancer research, and developmental biology. West Hollywood, CA
Yury A. Rovensky
wwwwwwwwwwwwwwww
Acknowledgements
I am extremely grateful to Dr. Alexander V. Ljubimov (Cedars-Sinai Medical Center, Los Angeles, CA) for critical reading of the manuscript, exceptionally helpful comments, stimulating discussions, and valuable advices. Generous help of Julia Y. Moers at all stages of the manuscript preparation including processing of the figures and assistance in preparing the References is highly appreciated. I thank Dr. Eugene B. Mechetner (Stonsa Biopharm, Inc., Irvine, CA) for his friendly encouragement and support. I would like to express my gratitude to Dr.Tatyana M. Svitkina (University of Pennsylvania, Philadelphia, PA) for providing her spectacular TEM photos and her valuable comments, and also to all my collaborators, who gave me their figures; they are acknowledged in the legends. I thank Raymond T. Moers for his help. I am much obliged to all my colleagues and friends, with whom I worked for many years. I apologize to all those whose valuable work in this field have not been referenced due to space limits.
xi
wwwwwwwwwwwwwwww
Contents
1 Introduction................................................................................................ References....................................................................................................
1 5
2 The Extracellular Matrix.......................................................................... References....................................................................................................
7 10
3 Cytoskeleton............................................................................................... 3.1 Actin Filaments................................................................................... 3.1.1 Actin-Binding Proteins........................................................... 3.1.2 Actin Filament Dynamics....................................................... 3.2 Microtubules....................................................................................... 3.2.1 Motor Proteins........................................................................ 3.2.2 Nonmotor Proteins.................................................................. 3.3 Intermediate Filaments........................................................................ References....................................................................................................
13 13 16 20 24 27 28 29 31
4 Pseudopodia and Adhesion Structures.................................................... 4.1 The Formation of Pseudopodia........................................................... 4.2 Cell–Extracellular Matrix Adhesion Structures.................................. 4.2.1 Focal Contacts (Focal Adhesions).......................................... 4.2.2 Focal Complexes..................................................................... 4.2.3 Fibrillar Adhesions.................................................................. 4.2.4 Hemidesmosomes................................................................... 4.2.5 Podosomes and Invadopodia................................................... References....................................................................................................
37 37 44 45 50 51 51 51 53
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix.......................................................................................................... 5.1 Cell Spreading on the Extracellular Matrix Surface........................... 5.1.1 Cells in a Suspended State...................................................... 5.1.2 The Morphology of Cell Spreading Process in Normal Cells.......................................................................
57 57 57 64
xiii
xiv
Contents
5.1.3 Morphological Alterations in the Spreading of Transformed Cells.............................................................. 5.2 The Signaling Pathways in the Spread Cells...................................... 5.2.1 Cell Surface Receptors............................................................ 5.2.2 Intracellular Signal Transduction............................................ 5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells.................................................................. 5.3.1 Integrin Receptor-Mediated Mitogenic Signaling Pathways................................................................................. 5.3.2 Growth Factor Receptor-Mediated Mitogenic Signaling Pathways................................................................. 5.3.3 Integrin and Growth Factor Receptor-Mediated Antiapoptotic Signaling Pathways.......................................... 5.3.4 The “Anchorage Dependence”................................................ 5.3.5 Integrin and Growth Factor Receptor-Mediated Morphogenic Signaling Pathways.......................................... 5.3.6 Integrin-Mediated Mechanical Force-Induced Signaling....... 5.4 Alterations in Integrin-Mediated Adhesion and Signaling in Transformed Cells........................................................................... 5.4.1 Defective Adhesive Function.................................................. 5.4.2 Alterations in the Mitogenic Signal Transduction.................. 5.4.3 The “Anchorage Independence”............................................. References.................................................................................................... 6 Cell Migration............................................................................................ 6.1 Factors Involved in Cell Migration..................................................... 6.1.1 Formation of Pseudopodia...................................................... 6.1.2 Polarization of Migrating Cells............................................... 6.1.3 Contact Inhibition of Cell Migration...................................... 6.1.4 Effect of Growth Factors......................................................... 6.1.5 Role of Focal Adhesions in Cell Migration............................ 6.2 Abnormalities of Cell Migration Machinery in Transformed Cells... 6.2.1 Pseudopodial Activity with Actin-Myosin Structure Deficiencies............................................................. 6.2.2 Cell-Matrix Adhesion Alterations........................................... 6.2.3 Hypersensitivity to Mitogens-Motogens................................. References....................................................................................................
74 88 88 91 94 95 96 97 99 100 105 108 108 110 111 113 121 121 123 126 127 128 130 132 133 134 135 137
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”................................................................... 145 References.................................................................................................. 152 8 Topographic Cell Responses.................................................................... 8.1 Cylindrical Substrata......................................................................... 8.1.1 Normal Cell Responses......................................................... 8.1.2 Transformed Cell Responses.................................................
153 154 154 158
Contents
xv
8.2 Grooved Substrata............................................................................. 8.2.1 Normal Cell Responses......................................................... 8.2.2 Transformed Cell Responses................................................. 8.3 Discontinuous Substrata.................................................................... 8.3.1 Lattices.................................................................................. 8.3.2 Multiple Vertvical Rods........................................................ 8.4 Effects of the Substratum Surface Topography on Cell Adhesion, Proliferation, and Synthetic Activities.............................................. 8.5 Mechanisms of Topographic Cell Responses................................... 8.5.1 Cylindrical Substrata............................................................. 8.5.2 Grooved Substrata................................................................. 8.5.3 Lattice Substrata.................................................................... References..................................................................................................
161 161 163 167 167 172
9 Intercellular Adhesive Interactions ....................................................... 9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions...... 9.1.1 Structure of Adherence Junctions......................................... 9.1.2 Dynamic Regulation of Adherens Junctions......................... 9.1.3 Contact Inhibition of Cell Proliferation................................ 9.2 Altered Regulation of Adherens Junctions Caused by Oncogenic Transformation........................................................... 9.2.1 Alterations in Cadherin–Catenin Complex........................... 9.2.2 Loss of Contact Inhibition of Cell Proliferation.................... References..................................................................................................
185 187 187 189 197
174 177 178 179 180 181
200 201 204 205
10 Conclusions .............................................................................................. 213 References.................................................................................................. 215 Index.................................................................................................................. 217
wwwwwwwwwwwwwwww
Abbreviations
ADAM (or MDC) proteins … ADF ADP AMF APC Arp Arp2/3 Asef ATP ATPase
Metalloproteinase-like disintegrin- cystein rich proteins
Bad BAX Bcl2, Bcl-x BCL2, BCL-x BPAG1 (BP1230)
Apoptogenic protein Apoptogenic tumor suppressor gene Antiapoptogenic proteins Antiapoptogenic proto-oncogenes Bullous pemphigoid antigen1
CAM kinases cAMP Cas (p130 Cas) CDH1 c-ErbB1/HER1 CH-ILKBP c-met Cobl Cortactin CTNNB1
Calmodulin dependent protein kinases Cyclic adenosine monophosphate Focal adhesion protein Tumor suppressor gene encoding E-cadherin Proto-oncogene encoding EGF receptor Protein parvin Proto-oncogene encoding HGF/SF receptor Protein cordon-bleu Cortical actin-binding protein Proto-oncogene encoding b-catenin
Actin-depolymerizing factor (cofilin) Adenosine diphosphate Autocrine motility factor Adenomatous polyposis coli protein Actin-related protein Actin-related protein complex Rac1-activating protein Adenosine triphosphate Adenosine triphosphatase
xvii
Abbreviations
xviii
DAG DAP kinase Dsh
1,2-diacylglycerol Death-associated protein kinase Disheveled protein
EB1 EC1, EC5 E-cadherin ECM EGF EMT Ena/VASP ERM
(Microtubule plus-) end-binding protein1 Extracellular cadherin subdomains Epithelial cadherin Extracellular matrix Epidermal growth factor Epithelial-mesenchymal transition Enabled/vasodilator-stimulated phosphoprotein family Ezrin, radixin and moezin protein family
F-actin FAK FGF FH domain FIP 200 FM formins
Filamentous actin Focal adhesion tyrosine protein kinase Fibroblast growth factor Formin homology domain Focal adhesion kinase family interaction protein of 200 kD Fluorescent microscopy Formin homology proteins
G protein G-actin GAP GDI GDP GEF GEF-H1 GF GFAP Girdin GPCR GSK3b GTP GTPase
Guanine nucleotide-binding protein Globular actin GTPase activating protein GDP dissociation inhibitor Guanosine diphosphate Guanine nucleotide exchange factor Rho guanine nucleotide exchange factor Growth factor Glial fibrillary acidic protein Girders of actin filaments protein G protein coupled receptor Glycogen synthase kinase-3 b Guanosine triphosphate Guanosine triphosphatase
HGF/SF
Hepatocyte growth factor, scatter factor
IFs IGF-1 ILK INK4a IP3 IQGAP1
Intermediate filaments Insulin-like growth factor Integrin-linked protein kinase Tumor suppressor gene Inositol 1,4,5-triphosphate IQ motif-containing GTPase activating protein1
JMY
Junction-mediated regulatory protein
Abbreviations
xix
LM Lmod
Light microscopy Protein leiomodin
MAP kinase (ERK) mDia MHC MLC MMP MRTF MSF MT1-MMP MTOC
Mitogen-activated protein kinase (extracellular signal- regulated kinase) Formin homology protein Myosin heavy chain Myosin light chain Matrix metalloproteinase Myocardin-related transcription factor Migration stimulating factor Membrane type 1-matrix metalloproteinase Microtubule-organizing center (centrosome)
N-cadherin Necl NPF
Neural cadherin Nectin-like immunoglobulin-like adhesion molecule Nucleation-promoting factors
p140 Cap p27/KIP1 p53 PAK P-cadherin PDGF PI PI3K PIK PINCH
Cas-associated protein Tumor suppressor gene Tumor suppressor gene, encoding p53 protein p21-activating protein kinase Placental cadherin Platelet-derived growth factor Phosphatidylinositol Phosphatidylinositol 3-kinase Phosphatidylinositol kinase Particularly interesting new cystein- histidine-rich protein Phosphoinositide Phosphatidylinositol biphosphate Phosphatidylinositol triphosphate Phosphatidylinositol phosphate kinase Protein kinase A, cAMP-dependent protein kinase Protein kinase B Protein kinase C Tumor suppressor gene, encoding phosphatase and tensin homolog (PTEN) protein
PIP PIP2 PIP3 PIPK PKA PKB (PKB/Akt) PKC PTEN Rab, Ras, Rho GTPases Rac, Rho, Cdc42 Rap1 R-cadherin RGD ROCK RTK
Families of small GTPases Members of Rho family of small GTPases Member of Ras family of small GTPases Retinal cadherin Arginine-glycine-aspartic acid sequence Rho-associated kinase, Rho kinase Receptor tyrosine kinase
Abbreviations
xx
Scar(WAVE) SEM SMA Small G proteins SNAIL1
Suppressor of cAMP receptor Scanning electron microscopy Alpha-smooth muscle actin Small GTPases, Ras superfamily GTPases Transcription repressor of CDH1 gene
Tcf/Lef TGF-a TIAM1 TIMP +TIP
T-cell factor/lymphocyte- enhancer factor Transforming growth factor Rac-1 specific exchange factor Tissue inhibitor of metalloproteinase Microtubule plus-end tracking protein
VASP VE-cadherin VEGF
A member of Ena/VASP protein family Vascular endothelial cadherin Vascular endothelial growth factor
WAF1 WASP WASP/WAVE (WASP/Scar)
Wnt1 glycoprotein
p21 protein, a cell cycle inhibitor Wiscott-Aldrich syndrome protein family Wiscott-Aldrich syndrome protein (WASP) family that includes WASP family verprolin-homologous (WAVE) proteins WASP family verprolin-homologous proteins WASP-homolog 2 domain WASP homolog associated protein with actin, membranes, and microtubules WASP-interacting protein Protein family. The name “Wnt” is a combination of Wg (“wingless” gene in Drosophila melanogaster) and “Int” (mouse oncogene) Protein encoded by WNT1 proto-oncogene
XMAP215
Microtubule associated protein
WAVE (Scar) WH2 WHAMM WIP Wnt
Chapter 1
Introduction
In multicellular animal organisms, tissue cells exist and function under the conditions of their direct contacts with the extracellular matrix and with each other. The extracellular matrix is a regulated three-dimensional frame, on the surface of and inside which the tissue cells attach, spread, move and interact with each other. The extracellular matrix consists of complex proteins composed of proteins and carbohydrates. These complex proteins are secreted by tissue cells. The protein– carbohydrate complexes are organized in an orderly network, the extracellular matrix. One type of the protein–carbohydrate complexes is represented by proteoglycans that are composed of proteins with covalently bound polysaccharides, glycosaminoglycans. Proteoglycans form strongly hydrated gels creating the tissue resilience that resists compression. Other types of the protein–carbohydrate complexes are glycoproteins and proteoglycans, composed of proteins with bound oligosaccharides. Among the glycoproteins of the extracellular matrix, the most important ones are collagens of at least 29 types, elastin, fibronectin, and laminins. These are fibrillar proteins involved in the formation of specialized structures of the matrix, fibers and basement membranes. The collagen and elastin fibers have mainly structural functions; elastin fibers are capable of tension and compression. Basement membranes include collagens, laminins, nidogens, and sometimes fibronectin. Glycoproteins fibronectin and laminin have mainly adhesion functions. These proteins critically contribute to adhesion of cells to the matrix and exert the regulating influence on cell migration. Animal tissue cells have the ability to attach to and spread on the extracellular matrix, to migrate to new areas of the matrix, and to respond to its physical-chemical characteristics and geometrical configurations. The tissue cells enter into the contacts with each other to form junctions, and these intercellular junctions are dynamically regulated. All these cell activities are, in essence, the “cell–extracellular matrix” or “cell– cell” adhesive interactions.
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_1, © Springer Science+Business Media, LLC 2011
1
2
1 Introduction
The adhesive interactions exert the strong controlling influence on the morphology of tissue cells, their locomotion and proliferation. The contacts of cells with the extracellular matrix or to each other determine the cell shapes, orientation of the cells, their locomotion activity, directions of the cell migrations, the survival of the cells and their ability to proliferate. Adhesive interactions of “cell–extracellular matrix” and “cell–cell” types play the most important role both in embryogenesis and in the adult organism, ensuring the preservation of the structural integrity of all tissues and their normal functioning. Adhesive interactions are the basis of the morphogenesis, when the displacements of cells over the basement membranes and fibers of the extracellular matrix, accompanied by the selective formation of stable intercellular adhesions, lead to the development of specific tissue structures. Adhesive interactions play important role in the regenerative processes, when the migration of connective tissue cells into the wound occurs directionally, being subordinate to the orientation of the matrix fibers. Another example, which indicates the role of adhesive interactions in the organism, is the attachment of platelets to the microvascular endothelium, which is an important step in thrombosis. Adhesive interactions are the regulators of survival and proliferative activity of tissue cells. Most of the normal cell types are capable of surviving and proliferating only being attached and spread on the surface of the solid substratum – the extracellular matrix (this phenomenon is termed substratum dependence of cell proliferation, or anchorage dependence). Losing the connections with the matrix, normal cells lose the ability to respond by proliferation to the soluble growth factors; many types of the unattached cells undergo programmed suicide, termed apoptosis (specifically, its variety, anoikis). The inhibition of proliferation of normal tissue cells after the cell monolayer formation and the establishment of stable adhesions between the cells (this phenomenon is termed contact inhibition of cell proliferation) is another example of the regulating influence of the adhesive interactions. Just as the anchorage dependence, this property plays an important role in the maintenance of the definite numbers of cells in different tissues, the retention of their structural integrity, in regeneration processes, etc. Thus, the behavior of normal tissue cells influenced by the adhesive interactions is controllable in the sense that it is checked and regulated by the extracellular matrix and surrounding cells. Oncogenic (tumorigenic) transformation of tissue cells with the subsequent development of malignant or benign tumors is caused by the alterations of the specific normal genes termed proto-oncogenes and tumor suppressor genes [1, 2]. These genes play key roles in the life of a normal cell. They control cell morphology and cytoskeleton, cell proliferation and the adhesive interactions of the cells with the extracellular matrix and with each other. The proto-oncogenes and tumor suppressor genes also control apoptosis – the active mechanism of cell suicide that protects an organism from the accumulation of the cells with genetic alterations [3]. (a) Proto-oncogenes are the group of normal genes in the genome (there are more than 100 proto-oncogenes in the human genome). The products of these genes, i.e., the proteins encoded by them, are the components of the intracellular signal transduction pathways. In particular, these
1 Introduction
3
proteins participate in the transduction of mitogenic signals initiated by soluble growth factors and (or) cell adhesive bonds to the extracellular matrix. Thus, the products of the proto-oncogenes promote the multiplication of cells, in that way playing the role of the positive regulators of cell proliferation. These proteins take part in the control of cytoskeleton reorganization and cell morphology changes, in adhesive interactions of cells with the extracellular matrix and with each other. These proteins also take part in the regulation of apoptosis. Because of the mutations of the proto-oncogenes or their translocations caused by the chromosomal rearrangements, the proto-oncogenes become permanently activated: their expressions are permanently increased, they are not being “turned off,” and the proteins encoded by them have altered functional activities and/or structures. The proto-oncogenes can also undergo the amplification: the number of copies of the gene considerably increases, and because of that, a quantity of the encoded protein grows. Such altered and permanently activated proto-oncogenes are termed oncogenes. Thus, because of the genetic alterations, the proto-oncogenes are turned into the permanently activated oncogenes encoding the proteins termed oncoproteins. The oncoproteins are responsible for the oncogenic transformation of the tissue cells and for the characteristic features of transformed cells. The oncogenes can induce the development of benign and malignant tumors. (b) Tumor suppressor genes encode the proteins, which act as inhibitors in the intracellular transduction pathways of the mitogenic signals. Thus, whereas protooncogenes carry out positive control of the proliferation of cells, promoting them to multiply, the tumor suppressor genes are negative regulators of cell proliferation, protecting the cells from deregulated multiplication. The tumor suppressor genes induce apoptosis, and favor the maintenance of the cell genome stability. Because of the inactivating mutations of the tumor suppressor genes, they get “knocked out,” which may abolish negative control of the cell proliferation and apoptosis stimulation [3]. Inactivation of the some tumor suppressor genes can cause the oncogenic transformation of the tissue cells and the development of tumors. The chemical carcinogens, ultraviolet or ionized irradiation can induce mutations and other genetic alterations causing the conversion of the protooncogenes to the permanently activated oncogenes, or the inactivation of tumor suppressor genes. The oncogenes can be delivered into cells by oncogenic viruses. Because of all these alterations normal tissue cells become transformed, and finally different kinds of malignant tumors can develop. Transformed cells have altered morphology, permanently stimulated and unregulated proliferative activity, and the block of apoptosis; the regulatory influences of the adhesive interactions of the cells with the extracellular matrix and with each other weaken or totally disappear. These characteristic features of the transformed cells determine their uncontrollable behavior in an organism. This uncontrollable behavior has both high biological and medical significance.
4
1 Introduction
Oncogenic transformation of tissue cells results in serious disturbances of adhesive interactions. These disturbances result in the deregulation of survival and proliferation of tumor cells and are the basis of important manifestations of anomalous, uncontrollable cell behavior. Neither cell detachment from the extracellular matrix nor the intercellular contacts inhibit the growth of tumor cells, which continue to proliferate, crawling over each other and forming the multilayered foci. This leads to the disorganization of the regular tissue structure, which is typical for malignant tumors. Because of altered adhesive interactions of tumor cells with each other, with normal cells and with the extracellular matrix, some tumor cells are easily separated from the malignant tumor. They actively penetrate surrounding healthy tissues, including lymph- and blood microcirculation system. The loss of substratum dependence of cell proliferation (termed anchorage independence) allows for the tumor cells (at least, for some of them), which enter the microcirculation system, to survive in their suspended state and circulate for some time. They can be retained at certain sites of microvasculature and can attach to the vascular endothelium with subsequent vessel penetration and formation of secondary foci of malignant tumor growth – metastases. In the cascade of events in malignant tumor invasion, the changes in morphology and locomotion of tumor cells take essential place. These changes are also related to the abnormalities of adhesive interactions. The study of adhesive interactions “cell–extracellular matrix” and “cell–cell” can be successfully carried out in vitro (out of the organism) under the conditions of the cultivation of cells in an artificial nutrient medium. Under these conditions cells can spread, move, contact with each other and proliferate on the flat bottoms of culture dishes. The extracellular matrix components are secreted by tissue cells not only in the organism, but also in the cell culture conditions. For example, fibroblasts and endothelial cells in culture secrete fibronectin; many types of cells secrete collagens, and epithelial cells secrete laminins. These proteins and secreted proteoglycans being adsorbed on the culture dishes, form thin interlayer, which actually performs the role of solid substratum for the cells. Thus, both in the organism and in the cell culture conditions, tissue cells establish adhesive interactions with the surfaces of the extracellular matrix formed by the cells. The next chapters will describe “cell–extracellular matrix” and “cell–cell” adhesive interactions in normal cells, and also their alterations in the cells as a result of oncogenic transformation. We will mostly analyze data on cultured normal cells of two main tissue types: connective tissue cells (fibroblastic cells) and epithelial cells. We will also discuss adhesive properties of various cultured transformed (by chemical carcinogens, oncogenic viruses or spontaneously transformed) cells of mesenchymal or epithelial origin; those cells in the future will be called “transformed fibroblasts” or “transformed epitheliocytes,” respectively.
References
5
References 1. Kopnin BP (2000) Targets of oncogenes and tumor suppressors: key for understanding basic mechanisms of carcinogenesis. Biochemistry (Mosc) 65(1):2–27 2. Abelev GI, Eraiser TL (2008) On the path to understanding the nature of cancer. Biochemistry (Mosc) 73(5):487–497. doi:10.1134/S0006297908050015 DOI:dx.doi.org 3. Cotter TG (2009) Apoptosis and cancer: the genesis of a research field. Nat Rev Cancer 9(7):501–507. doi:10.1038/nrc2663 DOI:dx.doi.org
wwwwwwwwwwwwwwww
Chapter 2
The Extracellular Matrix
Abstract The extracellular matrix is the ordered macromolecular network, on the surface of which and inside the tissue cells are attached to it and to each other, migrate, proliferate or survive. The matrix is composed of protein–carbohydrate complexes, which, in particular, include the glycoproteins carrying out mainly structural or mainly adhesive functions. The extracellular matrix is not only a mechanical framework but also a regulator of cell behavior. The matrix proteins are bound with the specific cell surface receptors resulting in the cell–matrix adhesion, which exerts effect on cell shapes, migration, proliferation, cell survival, and metabolism. In multicellular animal organisms, the majority of tissue cells are surrounded by the complex orderly network of interconnected extracellular macromolecules termed the extracellular matrix. The matrix consists of secreted complex molecules containing covalently attached protein and carbohydrate moieties; these matrix macromolecules are called protein–carbohydrate complexes. The extracellular matrix also includes highly specialized structures, such as cartilage, tendons, basement membranes, and also (with secondary deposition of calcium phosphate crystals) bones and teeth. The matrix macromolecules are produced and secreted by fibroblasts in connective tissue, chondroblasts in cartilage, osteoblasts in bone, histiocytes (macrophages in connective tissue), mast cells, epithelial cells in parenchymal organs, muscle cells, and endothelial cells of blood vessels. Molecular composition of the matrix is also influenced by white blood cells, which can migrate from blood vessel into the matrix in response to the specific stimuli. The molecular composition of the extracellular matrix includes several classes of the protein–carbohydrate complexes. The carbohydrate component content in these complexes may vary from less than 10% to more than 95%. (a) Proteoglycans are composed of the proteins (called core proteins) covalently attached to long nonbranched chains of polysaccharides, glycosaminoglycans. The polysaccharide content is more than 95% in the proteoglycans.
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_2, © Springer Science+Business Media, LLC 2011
7
8
2 The Extracellular Matrix
Glycosaminoglycans include several families: hyaluronic acid (which is in a free state, not bound to the protein), chondroitin sulfates, dermatan sulfates, keratan sulfates, heparin, and heparan sulfates. Owing to their high hydrophilia, glycosaminoglycans occupy large volumes in tissues, forming strongly hydrated gels that cause tissue turgor (resilience). The turgor gives the tissue an ability to resist compression forces. For example, an articular cartilage can resist mechanical pressures of a hundred atmospheres. Proteoglycans can form huge polymeric complexes in the extracellular matrix. Besides providing tissue turgor, proteoglycans can also be connected to other extracellular matrix proteins forming complex structures, e.g., basement membranes. Proteoglycans, such as heparan sulfate proteoglycans, are able to bind to and interact with a variety of proteins, including growth factors, some extracellular matrix components, and other molecules. Heparan sulfate proteoglycans can be involved in intracellular signaling as cell surface receptors or coreceptors for multiple ligands to modulate the distinct signal transduction pathways [1–4]. For instance, syndecans, which are members of the heparan sulfate proteoglycan family, act as coreceptors for growth factors in conjunction with cell surface integrin receptors and are involved in the regulation of cell–extracellular matrix adhesion and migration [5–9]. (b) Glycoproteins and proteoglycans consist of proteins with attached oligosaccharides. Glycoproteins and proteoglycans are similar in their structures and differ only in their carbohydrate content, which is significantly lower in glycoproteins (less than 10%, in comparison with 10–50% in proteoglycans). In contrast to proteoglycans, the carbohydrate component in glycoproteins is represented by short branched oligosaccharides, often with sialic acid at their ends. The most important glycoproteins of the extracellular matrix are represented by proteins of two functional types: • Collagen and elastin proteins that are mainly structural. • Fibronectin and laminin proteins that are mainly adhesive. Collagens are the main proteins of the extracellular matrix. They account for 25% of total protein content in a human organism. Unlike proteoglycans, collagens provide resistance to the mechanical stretching of a tissue, whereas proteoglycans oppose to its compression. Collagens are secreted by the cells of connective tissue, such as fibroblasts, osteoblasts, chondroblasts, and many other cells [10, 11]. To date, at least 29 types of collagen (they are collagen isoforms) are known. All collagen molecules contain a stiff triple helix structure: three polypeptide chains (named a-chains) are twisted up to the regular helix forming a collagen molecule. Many collagen types also have noncollagenous domains that do not form triple helices. The carbohydrate component in collagens is represented by monosaccharides and disaccharides. Collagens types I–III are the main collagens of connective tissues; type I collagen accounts for 90% of total collagen content in a human organism. After their secretion,
2 The Extracellular Matrix
9
molecules of collagen types I–III self-assemble into orderly polymers called collagen fibrils. The fibrils are further assembled to fibers of several micrometers (mm) in thickness called collagen fibers. Type IV collagen is a main component of basement membranes that also contain type VII collagen and some other collagen types. Types V, IX, and XII collagens provide connections of the collagen fibers with other proteins of the extracellular matrix. Elastin, unlike collagen, does not form the stiff triple helix. An elastin molecule consists of flexible polypeptide chains and has an ability to be reversibly unrolled under the action of mechanical stretching forces. Like collagen, elastin molecules are secreted into the extracellular space, where they are connected with each other to form fibers and sheets. The elastic fibers are coated by microfibrils of 10–20 nanometers (nm) in diameter. The microfibrils contain glycoproteins called fibrillins [12] that are members of the fibronectin family. These microfibrils obviously play an important role in the formation of elastin fibers. There is a striking difference between the mechanical characteristics of the stiff, nontensile collagen fibers and the rubber-like network of elastic fibers. The ability of elastic fibers to be stretched allows the tissues to restore their shapes after mechanical influences. Fibronectin. The extracellular matrix contains several adhesive noncollagenous proteins. Their characteristic features are the domains able to specifically bind with the cell surface receptors. The necessary component of these domains is the amino acid sequence arginine-glycine-aspartic acid (RGD). Fibronectin is one of the adhesive glycoproteins providing the attachment of cells to the extracellular matrix. Fibronectin is secreted by various types of cells, including fibroblasts and epithelial cells. There are at least 20 different fibronectin isoforms in humans. Secreted fibronectin molecules assemble into fibrils in the matrix. The fibronectin fibrillogenesis is initiated by the cell surface integrin receptors [13, 14]. Some part of fibronectin in form of fibrils is connected with the cell surfaces. Fibronectin in soluble state is found in blood and other biological fluids. Fibronectin has several domains, which can specifically bind to the cells and also to other matrix molecules, such as collagens (the strongest binding being with type III collagen) and heparin. Laminins (at least 15 isoforms identified so far) are cross-shaped trimeric adhesive glycoproteins that have different domains to specifically bind to cells, type IV collagen, nidogens, and some glycosaminoglycans. Laminins, just as type IV collagen and fibronectin, are components of basement membranes. Laminins mediate the attachment of parenchymal cells to type IV collagen thereby providing the interaction between cells and basement membranes. Other extracellular matrix glycoproteins are nidogens, tenascins, and fibulins. Nidogens (entactins) bind to both laminin and type IV collagen forming the additional connection between laminins and collagen.
10
2 The Extracellular Matrix
Tenascin family of proteins (tenascin-C, -X, -R, and -W) can bind fibronectin. However, unlike fibronectin, tenascins have both cell adhesive and antiadhesive functions depending on the cell type. These different functions are mediated by different tenascin domains; the number of these domains in a tenascin molecule varies because of alternative splicing [15, 16]. Fibulins can interact with many matrix components, such as some basement membrane proteins, fibronectin, fibrillin, and proteoglycans, to form supramolecular structures within the matrix [17]. The extracellular matrix is not only a mechanical framework that stabilizes a tissue structure. The matrix plays a much more active and complex role in the regulation of cell behavior, influencing the shape, migration, proliferation, survival, and metabolism of cells, which are involved in adhesive interactions with the matrix [18–20]. Migrations of cells during embryogenesis or in regeneration processes depend on the extracellular matrix. The matrix molecules are involved in acute and chronic inflammation in tissues and also in such widespread human diseases as rheumatoid arthritis, osteoarthritis, asthma, and others [21–26]. The collagen diseases (collagenosis) are caused by genetic disturbances in the expression and regulation of extracellular matrix molecules. For instance, mutations in the genes encoding types I, III, or V collagen cause heritable connective tissues disorders, mutations in the gene encoding type VI collagen result in congenital muscular dystrophy or myopathies, and mutations in the genes encoding types II, IX, and XI collagen cause skeletal dysplasias [27–29]. The problem of cancer cell invasion and metastasis is closely related to the extracellular matrix. Adhesive interactions of tissue cells with the extracellular matrix include the following: 1 . Spreading of cells on the extracellular matrix. 2. Active displacement of cells (cell migration). 3. Cell responses to the chemical heterogeneity of the extracellular matrix. 4. Cell responses to the geometric configuration of the extracellular matrix. All these adhesive interactions are accomplished by means of two base cellular functions: formation of the pseudopodia and formation of the special adhesive structures, which ensure the attachment of cells to the extracellular matrix.
References 1. Heinegård D (2009) Proteoglycans and more- from molecules to biology. Int J Exp Pathol 90(6):575–586 2. Kirn-Safran C, Farach-Carson MC, Carson DD (2009) Multifunctionality of extracellular and cell surface heparan sulfate proteoglycans. Cell Mol Life Sci 66(21):3421–3434. doi:10.1007/ s00018-009-0096-1 DOI:dx.doi.org 3. Schaefer L, Schaefer RM (2010) Proteoglycans: from structural compounds to signaling molecules. Cell Tissue Res 339(1):237–246. doi:10.1007/s00441-009-0821-y DOI:dx.doi.org
References
11
4. Mythreye K, Blobe GC (2009) Proteoglycan signaling co-receptors: roles in cell adhesion, migration and invasion. Cell Signal 21(11):1548–1558. doi:10.1016/j.cellsig.2009.05.001 DOI:dx.doi.org 5. Xian X, Gopal S, Couchman JR (2010) Syndecans as receptors and organizers of the extracellular matrix. Cell Tissue Res 339(1):31–46. doi:10.1007/s00441-009-0829-3 DOI:dx. doi.org 6. Bass MD, Morgan MR, Humphries MJ (2009) Syndecans shed their reputation as inert molecules. Sci Signal 2(64): pe18. doi:10.1126/scisignal.264pe18 DOI:dx.doi.org 7. Lambaerts K, Wilcox-Adelman SA, Zimmermann P (2009) The signaling mechanisms of syndecan heparan sulfate proteoglycans. Curr Opin Cell Biol 21(5):662–669. doi:10.1016/j. ceb.2009.05.002 DOI:dx.doi.org 8. Schmidt S, Friedl P (2010) Interstitial cell migration: integrin-dependent and alternative adhesion mechanisms. Cell Tissue Res 339(1):83–92. doi:10.1007/s00441-009-0892-9 DOI:dx.doi.org 9. Streuli CH, Akhtar N (2009) Signal co-operation between integrins and other receptor systems. Biochem J 418(3):491–506. doi:10.1042/BJ20081948 DOI:dx.doi.org 10. Gordon MK, Hahn RA (2010) Collagens. Cell Tissue Res 339(1):247–257. doi:10.1007/ s00441-009-0844-4 DOI:dx.doi.org 11. Shoulders MD, Raines RT (2009) Collagen structure and stability. Annu Rev Biochem 78:929–958. doi:10.1146/annurev.biochem.77.032207.120833 DOI:dx.doi.org 12. Ramirez F, Sakai LY (2010) Biogenesis and function of fibrillin assemblies. Cell Tissue Res 339(1):71–82. doi:10.1007/s00441-009-0822-x DOI:dx.doi.org 13. White ES, Baralle FE, Muro AF (2008) New insights into form and function of fibronectin splice variants. J Pathol 216(1):1–14. doi:10.1002/path.2388 DOI:dx.doi.org 14. Leiss M, Beckmann K, Girós A, Costell M, Fässler R (2008) The role of integrin binding sites in fibronectin matrix assembly in vivo. Curr Opin Cell Biol 20(5):502–507. doi:10.1016/j. ceb.2008.06.001 DOI:dx.doi.org 15. Brellier F, Tucker RP, Chiquet-Ehrismann R (2009) Tenascins and their implications in diseases and tissue mechanics. Scand J Med Sci Sports 19(4):511–519. doi:10.1111/j. 1600-0838.2009.00916.x DOI:dx.doi.org 16. Tucker RP, Chiquet-Ehrismann R (2009) The regulation of tenascin expression by tissue microenvironments. Biochim Biophys Acta 1793(5):888–892. doi:10.1016/j.bbamcr.2008.12.012 DOI:dx.doi.org 17. de Vega S, Iwamoto T, Yamada Y (2009) Fibulins: multiple roles in matrix structures and tissue functions. Cell Mol Life Sci 66(11–12):1890–1902. doi:10.1007/s00018-009-8632-6 DOI:dx.doi.org 18. Hynes RO (2009) The extracellular matrix: not just pretty fibrils. Science 326 (5957):1216–1219. doi:10.1126/science.1176009 DOI:dx.doi.org 19. Tsang KY, Cheung MC, Chan D, Cheah KS (2010) The developmental roles of the extracellular matrix: beyond structure to regulation. Cell Tissue Res 339(1):93–110. doi:10.1007/s00441009-0893-8 DOI:dx.doi.org 20. Rozario T, DeSimone DW (2010) The extracellular matrix in development and morphogenesis: a dynamic view. Dev Biol 341(1):126–140. doi:10.1016/j.ydbio.2009.10.026 DOI:dx.doi.org 2 1. Sofat N (2009) Analysing the role of endogenous matrix molecules in the development of osteoarthritis. Int J Exp Pathol 90(5):463–479. doi:10.1111/j.1365-2613.2009.00676.x DOI:dx.doi.org 22. Järveläinen H, Sainio A, Koulu M, Wight TN, Penttinen R (2009) Extracellular matrix molecules: potential targets in pharmacotherapy. Pharmacol Rev 61(2):198–223. doi:10.1124/ pr.109.001289 DOI:dx.doi.org 23. Salerno FG, Barbaro MP, Toungoussova O, Carpagnano E, Guido P, Spanevello A (2009) The extracellular matrix of the lung and airway responsiveness in asthma. Monaldi Arch Chest Dis 71(1):27–30 24. Yurchenco PD, Patton BL (2009) Developmental and pathogenic mechanisms of basement membrane assembly. Curr Pharm Des 15(12):1277–1294. doi:10.2174/138161209787846766 DOI:dx.doi.org
12
2 The Extracellular Matrix
25. Loeser RF (2009) Aging and osteoarthritis: the role of chondrocyte senescence and aging changes in the cartilage matrix. Osteoarthritis Cartilage 17(8):971–979. doi:10.1016/j. joca.2009.03.002 DOI:dx.doi.org 26. Bateman JF, Boot-Handford RP, Lamandé SR (2009) Genetic diseases of connective tissues: cellular and extracellular effects of ECM mutations. Nat Rev Genet 10(3):173–183. doi:10.1038/nrg2520 DOI:dx.doi.org 27. Malfait F, De Paepe A (2009) Bleeding in the heritable connective tissue disorders: mechanisms, diagnosis and treatment. Blood Rev 23(5):191–197. doi:10.1016/j.blre.2009.06.001 DOI:dx.doi.org 28. Carter EM, Raggio CL (2009) Genetic and orthopedic aspects of collagen disorders. Curr Opin Pediatr 21(1):46–54. doi:10.1097/MOP.0b013e32832185c5 DOI:dx.doi.org 29. Maraldi NM, Sabatelli P, Columbaro M, Zamparelli A, Manzoli FA, Bernardi P, Bonaldo P, Merlini L (2009) Collagen VI myopathies: from the animal model to the clinical trial. Adv Enzyme Regul 49(1):197–211. doi:10.1016/j.advenzreg.2008.12.009 DOI:dx.doi.org
Chapter 3
Cytoskeleton
Abstract Actin microfilaments, microtubules, and intermediate filaments are the cytoskeleton systems that play crucial roles in basic cell functions and behavioral cell responses. Actin cytoskeleton and microtubules participate cooperatively in the formation of pseudopodia and adhesive bonds of cells with the extracellular matrix, thereby determining the capability of the cells to migration. Both these systems play a key role in cell shape determination and intercellular adhesion. Microtubules are critically involved in mitotic cycle. Intermediate filaments carry out both mechanical and some nonmechanical functions in cells. The cytoskeleton actively participates in all adhesive interactions. The cytoskeleton is represented by three types of intracellular structures: actin filaments (actin microfilaments), microtubules, and intermediate filaments (Fig. 3.1). From these structures, the actin cytoskeleton and the system of microtubules play a key role in adhesive interactions.
3.1 Actin Filaments Actin cytoskeleton is the determinant in cell shape and cell migration; it is critically involved in such basic cellular functions as adhesion of cells to the extracellular matrix or to each other, cell proliferation, and survival. Alterations in the actin cytoskeleton formation, organization, and regulation play a key role in cancer invasion and metastasis [1–4]. Actin filaments are polymerized from monomeric protein actin. This protein has a globular form. Globular-actin (G-actin) readily polymerizes under physiological conditions to form filamentous-actin (F-actin) with the concomitant hydrolysis of ATP. Actin monomers are connected with each other and form F-actin having the appearance of the polymeric double-helical threads with a diameter of 6–7 nm that are termed actin filaments (actin microfilaments). The actin polymerization resulting in the formation of actin filaments strongly needs arginylation of actin Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_3, © Springer Science+Business Media, LLC 2011
13
14
3 Cytoskeleton
Fig. 3.1 Rat fibroblastic cell. The cytoskeleton: actin microfilament bundle (mf), microtubules (arrows), intermediate filaments (arrowheads). Electron microscopy (EM). Scale bar, 0.2 mm. Courtesy of T.M. Svitkina, reproduced with permission from the Journal of Structural Biology, 1995; 115(3):290–303
(the transfer of the arginine residue, arginyl, to actin): the inhibition of actin arginylation significantly decreases the level of actin polymerization [5]. Actin filaments are very dynamic: the actin monomers in the cytoplasm constantly join the ends of the filaments faced the cell membrane (termed barbed, or plus-ends), whereas at the opposite ends (termed pointed, or minus-ends) the depolymerization of actin occurs. Thus, filaments possess structural polarity: a lengthening of the thread occurs from the plus-end, a shortening from the minus-end of a filament. There are b- or g-isoforms of cytoplasmic actin in fibroblastic and epithelial cells. The b-actin filaments are located in the basal but not in the dorsal portions of cells, and these filaments are organized in bundles (Fig. 3.1). The b-actin filament bundles are critically involved in the formation and stability of cell– extracellular matrix and cell–cell adhesions, and play the main role in cell contractility. The g-actin filaments in moving cells are mainly organized as networks, which are located under the dorsal cell membrane and in the motile cell parts (e.g., in lamellipodia). In nonmigrating cells g-actin is also recruited into the filament bundles [6]. Actin filaments are organized differently in fibroblastic and epithelial cells. Fibroblastic cells have polygonal or elongated shapes, and they contain in their cytoplasm numerous, predominantly straight, long actin filaments and filament bundles, which are oriented mostly along the cell axis (Fig. 3.2). The single epitheliocytes acquire discoid shapes, and they have a circular actin filament bundle located along the entire cell periphery (Fig. 3.3).
3.1 Actin Filaments
15
Fig. 3.2 Human fibroblastic cells. Linear actin microfilaments and the microfilament bundles (arrow). Staining for actin. Fluorescent microscopy (FM). Scale bar, 24.5 mm. Courtesy of A.Y. Alexandrova
Fig. 3.3 Epithelial mouse cell. Circular actin microfilament bundle (arrow). Staining for actin. FM. Scale bar, 40 mm
16
3 Cytoskeleton
3.1.1 Actin-Binding Proteins Organization and functioning of actin cytoskeleton are ensured by a large family of actin-binding proteins that can directly interact with F- and G-actin. At present, there are over 150 known actin-binding proteins, which account for approximately 25% of cellular protein. Actin-binding proteins regulate the processes of the polymerization– depolymerization of actin filaments, carry out their nucleating, severing and capping, and also connect the filaments with each other and give them the contractile properties. Actin-binding proteins have relatively low affinity for actin. The weak bonds are necessary for dynamic actin cytoskeleton remodeling [7]. In the large group of actin-binding proteins, the actin nucleators play a critical role. These proteins directly nucleate actin filament formation de novo. The group of actin nucleators includes the actin-related proteins (Arp2/3 complex) and nucleation-promoting factors (NPF), and also the proteins Spire, Cordonbleu (Cobl), Leiomodin (Lmod), and formins. Arp2/3 complex. This complex consists of seven proteins, from which the Arp2 and Arp3 proteins are homologous to actin. Arp2/3 is an initiator of actin filament nucleation to form branched filament network [8–14]. The Arp2/3 complex is intrinsically inactive and needs the activating NPFs, such as WASP/WAVE (also called WASP/Scar) protein family, WASP homolog-associated protein with actin, membranes and microtubules (WHAMM), and also junctionmediated regulatory protein (JMY). WASP/WAVE (WASP/Scar). Wiscott–Aldrich syndrome protein (WASP) family includes WASP, N-WASP, and three WASP family verprolin-homologous (WAVE) proteins. WAVE proteins are also known as suppressor of cAMP receptor (Scar) proteins. Therefore, WASP/WAVE is also called WASP/Scar protein family. All WASP family members have a domain through which Arp2/3 complex is activated to nucleate actin polymerization resulting in the formation of new branched actin filaments [8, 13, 15–18]. The functions of WASP and N-WASP in the Arp2/3 complex-mediated actin nucleation are regulated by the WASP-interacting protein (WIP) that interacts with WASP and N-WASP [19, 20]. WHAMM. This nucleation promoting factor (NPF) is associated with actin, membranes, and cytoskeleton system of microtubules. WHAMM activates the Arp2/3mediated actin nucleation along microtubules and also at the Golgi apparatus. Thus, WHAMM functions at the interface of the actin cytoskeleton and microtubules [13]. JMY. It activates the Arp2/3-mediated actin nucleation; however, JMY can induce actin nucleation in the absence of the Arp2/3 complex. JMY can function both as an NPF to activate the Arp2/3 complex and as an actin nucleator, like Spire (see below), that directly nucleates new actin filaments. Thereby, JMY can induce the formation of both branched and unbranched actin filaments [13].
3.1 Actin Filaments
17
It is interesting that JMY is a transcriptional coactivator of p53 gene and is primarily located in the cell nucleus. However, JMY shuttles between cell nucleus and the cytoplasm: in highly motile cells JMY is excluded from the nucleus to the cytoplasm and is colocalized with actin filaments at the leading edge of the migrating cell. Increased JMY expression promotes cell migration. Spire, Cordon-bleu (Cobl), Leiomodin (Lmod) proteins, and formin homology proteins (formins). These are actin nucleators that directly nucleate new actin filaments. Both Spire and Cobl proteins generate nucleation of unbranched actin filaments and remain bound to the pointed end of the new emerging filament. Cobl is mainly expressed in the brain. Lmod isoforms are homologs of tropomodulins that are capping proteins (see Sect. 3.1.1). Lmod binds the actin-binding protein tropomyosin (see Sect. 3.1.1) and acts as nucleator of tropomyosin-decorated actin filaments in muscles [12, 13, 21, 22]. Formins not only directly nucleate new actin filaments but also protect the filaments from their capping, thereby providing the progressive elongation of the filament barbed ends [13, 14, 23–25] (see Sect. 3.1.2). All the above mentioned actin nucleators, except formins, use small actin-binding WASP-homology 2 domains (WH2 domains) that were first identified in the WASP/Scar (WASP/WAVE) protein family. WH2 domain binds actin monomers to directly nucleate actin filaments de novo. For example, the nucleator Spire brings actin monomers together with four tandem WH2 domains to form a linear actin tetramer, to which free monomers then bind. In this way, a new actin filament is formed and further elongated. WH2 domain is involved in various functional interactions between actin monomers and different actin-binding proteins. Owing to the multifunctional character of the WH2 domains, such actin nucleators as Spire or Cobl, can not only nucleate actin filaments but also sever them or cap the filaments at their barbed ends thereby blocking the barbed end growth [21]. Unlike other actin nucleators, formins use the formin-homology 2 domain (FH2 domain) to interact with G-actin (see Sect. 3.1.2). Profilin. It binds to actin monomers and stimulates the replacement of ADP to ATP in them. After the separation of profilin, the monomers rapidly begin polymerization at the barbed ends of the filaments, resulting in their growth [26]. Profilin interacts with multiple proteins to regulate actin cytoskeleton dynamics and membrane trafficking [27]. Gelsolin. It is the most potent member of the gelsolin/villin protein superfamily. Gelsolin regulates both the assembly and disassembly of actin filaments by their severing, capping filament ends (preventing the addition or separation of actin monomers), and actin filament nucleation [28, 29]. Actin-depolymerizing factor (ADF), also called cofilin. This protein accelerates depolymerization of actin and prevents released actin monomers from being polymerized again (until the monomers stay bound with cofilin). Thus, cofilin causes the fragmentation of actin filaments. Through its actin severing activity,
18
3 Cytoskeleton
cofilin increases the number of free microfilament barbed ends to initiate actin polymerization in cell protrusions, such as lamellipodia [29–32]. Cortical actin-binding protein (Cortactin). It facilitates actin filament branching via the Arp2/3 complex. Cortactin also induces cross-linking of actin filaments, promoting the formation of the filament bundles [33–35]. Capping proteins. These proteins are capable of being joined to the barbed or to the pointed end of the actin filaments, preventing the addition or separation of actin monomers and thus regulate filament length. The gelsolin and also protein villin that belongs to the gelsolin/villin superfamily [36] cause not only calcium-dependent fragmentation of actin filaments, but also capable of capping the filament barbed ends. Another family of capping proteins tropomodulins needs their binding to tropomyosins (see below) for the capping function at actin filament pointed ends [37]. Alpha-actinin, filamins, and fimbrins (plastins). The first two proteins form the flexible connections between the actin filaments. Because of that, the threedimensional reticular structure is created. Alpha-actinin is an actin cross-binding protein. In nonmuscle cells, a-actinin is found along actin filaments cross-linking them. Besides binding to actin, a-actinin associates with some proteins (such as vinculin, zyxin, and a-catenin) in cell–matrix and in cell–cell adhesion complexes, organizing actin framework and linking it to these adhesion complexes [38]. Filamins are a family of three actin-binding and cross-binding proteins that organize actin filaments in networks and fibers. Filamin has an actin-binding domain and a rod segment consisting of up to 24 immunoglobulin-like domains. Two hinges in the rod segment result in a V-shaped flexible actin-crosslinker molecule that can generate actin filament networks with the high-angle filament branching. In addition, filamins can bind many transmembrane cell receptors and signaling proteins, thereby promoting their connection to the actin cytoskeleton [39]. Fimbrins (also termed plastins) cause cross-linking of separate actin filaments into the parallel tight bundles [40]. Fascin. It is the actin cross-linker protein that sews together actin filaments with their associated barbed ends into the long bundles. Fascin plays an important role in filopodia formation. Girdin (also reported as GIRDers of actin filaments, APE or GIV). Its alternative names Akt phosphorylation enhancer, Girders of actin filament. Girdin is required for the formation of actin filament bundles and lamellipodia and is involved in both remodeling of actin cytoskeleton and cell motility [41]. The ezrin, radixin, and moezin (ERM) protein family. These proteins have a plasma membrane-binding domain and an actin-binding one. Thus, the ERM proteins mediate plasma membrane–actin cytoskeleton cross-linking. The ERM are involved in cortical actin cytoskeleton remodeling, cell migration, stabilization of cell–cell adhesions, and other cell functions [42, 43].
3.1 Actin Filaments
19
Myosins. The superfamily of myosins includes about 100 members. Myosins are actin-binding proteins having ATPase activity and fulfilling motor function. Myosins bind actin filaments and use ATP hydrolysis to generate force and to move along the filament toward its barbed end. The myosin VI is an exception: it is a unique pointed enddirected myosin motor protein [44]. Myosin molecule has a domain to interact with cargo molecules. Thus, myosins can transport various membranous organelles along actin microfilaments that act as “rails.” Normal actin dynamics is a necessary condition for the myosin-mediated transport of organelles [45–48]. Myosins play an important role not only as intracellular cargo transporters but also as determinants of cell contraction (see below) and are critically involved in cell adhesion and migration. Tropomyosins. These proteins are located along the filaments giving them necessary hardness and stabilizing them. Tropomyosin isoforms collaboratively interact with different actin-binding proteins including cofilin, gelsolin, tropomodulins, Arp2/3, myosin, and caldesmon, thereby conferring different properties to the actin filaments [49, 50]. Caldesmon. It directly interacts with actin as well as with tropomyosins, thereby being involved in the assembly, dynamics, and stability of actin filaments. Caldesmon inhibits the ATPase activity of myosin II, so caldesmon is the negative regulator of cell contractility [51]. Therefore, caldesmon is involved in actin cytoskeleton rearrangement, cell motility, cell shape changes, and exo- or endocytosis. Among the actin-binding proteins, myosins have a particularly important role. Interaction of a member of the myosin superfamily, myosin II, with actin is the basis of muscle contraction. In nonmuscle cells (e.g., in the fibroblasts) myosin II gives actin filaments contractile properties [52]. A molecule of myosin II consists of heavy chains (MHC) and light chains (MLC) and has two “heads” and a “tail.” Phosphorylation of MLC causes assembly of myosin molecules into short bipolar aggregates (of 10–20 molecules) (Fig. 3.4). The myosin aggregates are connected by the “heads” with the lateral sides of two actin filaments of opposite polarity. The myosin “heads” change their conformation and thereby exert the pulling influence on the filaments. As a result, two adjacent filaments slide relative to each other in opposite directions (Fig. 3.5). Sliding of the actomyosin filaments relative to each other is the basis of the actin cytoskeleton contractility. Necessary energy for these motions is freed owing to the ATP hydrolysis caused by the ATPase activity of myosin. The contractility of actomyosin is controlled by regulatory enzymes. Calciumdependent myosin light chain kinase (MLC kinase) phosphorylates MLC, increasing contractility of actomyosin. The opposite action is achieved by MLC phosphatase. These enzymes, in turn, are regulated by special proteins from the Rho family of small GTPases (Rho family of GTPases or Rho GTPases) (see Sect. 5.3.5). The activation of some proteins of the Rho family leads to the increase in the contractility of the actomyosin. The negative regulation of the contractility is achieved by the protein caldesmon, which inhibits ATPase activity of myosin II [51]. The contractility of actin filaments plays a key role in the formation and maintenance of stable cell contacts with the extracellular matrix, in cell spreading and in cell locomotion.
20
3 Cytoskeleton
Fig. 3.4 Fibroblastic mouse cell. Individual myosin II minifilaments have bipolar morphology with globular ends and a bare central region. EM. Scale bar, 0.1 mm. Courtesy of T.M. Svitkina, reproduced with permission from the Journal of Structural Biology, 1995; 115(3):290–303
Fig. 3.5 Diagram showing the interaction of myosin II with actin microfilaments (see the text for explanation)
3.1.2 Actin Filament Dynamics Formation of new actin filaments in the cell proceeds via their offshoot from the preexisting filaments (Fig. 3.6). The actin-binding proteins have a critical role in this process. For the formation of a new filament, the unique “priming” is necessary. Actinbinding proteins of the Arp 2/3 complex play a key role in its formation as the initiators
3.1 Actin Filaments
21
Fig. 3.6 Diagram showing the formation of new actin microfilaments (see the text for explanation)
of actin filament nucleation [8, 9, 11, 17, 18, 30]. The Arp 2/3 complex is intrinsically inactive. It is activated by the WASP/WAVE (WASP/Scar) [13, 15–18]. The actinbinding protein cortactin binds to the Arp2/3 complex and recruits it to a preexisting actin filament [33–35]. WASP/WAVE (WASP/Scar) proteins bring together an actin monomer and Arp2/3 complex on the lateral side of the preexisting filament to initiate the formation of a new one [8, 13, 15–18]. After the Arp 2/3 complex attachment to the lateral side of preexisting filament, the Arp 2/3 changes its configuration and acquires the ability to join one additional monomer of actin to itself. So the “priming” is serving for further actin polymerization and for the growth of a new filament, which in the form of the offshoot goes out from the lateral side of the old filament at an angle of approximately 70° (Figs. 3.6–3.8). This way, a branched network of actin filaments is formed in the cell [8, 9, 11, 18, 30, 53]. Another group of actin-binding proteins, formins, mediates linear growth of actin filaments. Formins directly nucleate the actin polymerization of linear filaments and mediate their elongation by a special mechanism [13, 14, 23–25]. Formin can bind in a stepwise manner to each next attached actin monomer, thereby protecting the growing barbed end of the filament against its binding to capping proteins and
22
3 Cytoskeleton
Fig. 3.7 Multiple branching of actin filaments. Overview of the branched filament network (a) and enlargements of the boxed regions (b–g). EM. Scale bar, 0.3 mm. Courtesy of T.M. Svitkina, reproduced with permission from the Journal of Cell Biology [30]
3.1 Actin Filaments
23
Fig. 3.8 Localization of Arp2/3 complex at actin branching points. The Arp2/3 complex is immunostained using 10-nm gold-conjugated antibody. Gold particles are highlighted in yellow. EM. Scale bar, 40 nm. Courtesy of T.M. Svitkina, reproduced with permission from the Journal of Cell Biology [30]
initiating the filament elongation. Formin is rapidly translocated along the filament. One formin domain (formin homology 2 domain, FH2) initiates actin filament assembly and remains associated with the barbed end of the filament, providing rapid addition of actin monomers and protecting the filament end from capping proteins. The adjacent formin domain (formin homology 1 domain, FH1) influences the FH2 domain function through binding to the actin monomer-binding protein profilin, thereby promoting filament elongation. The growth of actin filaments discontinues because of the capping of their barbed ends. Pointed ends are released and the depolymerization of actin begins at these sites. The filaments are disassembled to separate ADP-containing actin monomers. The binding with cofilin contributes to the filament disassembly (Fig. 3.6) [29–32].
24
3 Cytoskeleton
After the replacement of ADP for ATP (this replacement is catalyzed by the actin-binding protein profilin), actin monomers are newly prepared to enter the polymerization reaction [26]. There are retrograde and anterograde flows of actin filaments in a cell. The filaments are polymerized from actin monomers at the active cell edge, namely at the free edge of lamellar cytoplasm where pseudopodia are formed (see Sect. 4.1). From there, the filaments start to move rapidly backward. These rapidly moving actin filaments are not associated with myosin II molecules. When the filaments appear to be in the more proximal zone of lamellar cytoplasm, their retrograde movement becomes much slower, and the filaments form regular bundles. In these bundles, the actin gets associated with myosin II and with certain other actin-binding proteins. In the course of the retrograde flow, part of filaments is gradually “disbranched” and depolymerized; the depolymerization is promoted by cofilin. The retrograde flow stops in the convergence zone between the lamellar cytoplasm and the central part of the cell. Forming actin monomers rapidly flow anterogradely back to the active cell edge for their repolymerization [1]. The formation and the organization of actin cytoskeleton are controlled by the Rho family of GTPases, Rho GTPases. These proteins control the polymerization of actin, assembly and stabilization of actin filaments, their contractility, and their organization [1] (see Sect. 5.3.5). The actin cytoskeleton dynamics can influence gene activity. In the process of actin polymerization, the transcriptional coactivators termed myocardin-related transcription factors (MRTFs) are liberated. They are involved in the activation of the expression of many genes, including those regulating actin cytoskeletal organization and cell growth [54]. Actin cytoskeleton plays a key role in: (a) the formation of pseudopodia that are cellular outgrowths, which are necessary for cell spreading and movement; (b) formation of special adhesive structures – the molecular complexes, which ensure the attachment of cells to the extracellular matrix; (c) the formation of stable intercellular contacts.
3.2 Microtubules The cytoskeletal system of microtubules, just as actin cytoskeleton, plays a key role in cell shape determination, cell migration, and intercellular and cell–matrix adhesion. Microtubules are involved in the formation of mitotic spindle, the structure used by cells to segregate their chromosomes during cell division. Microtubules, like actin filaments, critically contribute to the specific behavior of cancer cells [3]. Microtubules are self-assembling straight hollow cylinders 25 nm in diameter, variable in length; they are built from ab-tubulin heterodimers (Fig. 3.9). Microtubules have outstanding mechanical properties combining high resilience and stiffness [55]. The ab-tubulin heterodimers are joined end-to-end to form protofilaments with alternating a and b subunits. Therefore, in a protofilament, one end (designated
3.2 Microtubules
25
Fig. 3.9 Epithelial monkey cell. Microtubules that are immunogold labeled with detyrosinated tubulin. Gold-labeling microtubules (arrows). EM. Scale bar, 0.2 mm. Courtesy of T.M. Svitkina, reproduced with permission from the Journal of Structural Biology, 1995; 115(3):290–303
minus-end) has the a-tubulin exposed, while the other end (designated plus-end) has the b-tubulin exposed. The protofilaments then associate parallel to one another to eventually form hollow cylindrical microtubules. Thus, a microtubule has one end (plus-end) with only b-tubulin exposed and another end (minus-end) with only a-tubulin exposed. This structural polarity of microtubules is a very important feature playing a pivotal role in their dynamics and functions [56]. In contrast to actin filaments, the system of microtubules in a cell is centralized: they are nucleated and radiate into the peripheral cytoplasm (Fig. 3.10) from a specific place, called the microtubule-organizing center (MTOC), or centrosome. In the interphase cell, the MTOC is located near the nucleus and surrounds centrioles that are cylindrical structures, usually in pairs, oriented at right angles to one another, formed by nine triplets of microtubules. Proteins that are present in the MTOC or at the surfaces of centrioles include g-tubulin. This tubulin, which is homologous to a- and b-tubulins, combines with several associated proteins to form a circular structure called the g-tubulin ring complex. It acts as a scaffold for ab-tubulin heterodimers to initiate polymerization [55–58]. Microtubules nucleated by the g-tubulin ring complex are capped at their minusends, and the polymerization at these ends is inhibited [56]. Centrioles, similar to chromosomes, can double and serve as the centers of initiation of microtubules of the mitotic spindle. Therefore, minus-ends of most microtubules are anchored to the MTOC; the growth of the microtubules continues away from the MTOC in the plus-end direction up to the approximation of the distal plus-ends to the cell edges.
26
3 Cytoskeleton
Fig. 3.10 Fibroblastic rat cells. Microtubules (arrow). Staining for tubulin. FM. Scale bar, 25 mm. Courtesy of S.N. Rubtsova
The assembly of microtubules from tubulin molecules is similar to the assembly of microfilaments from actin: like actin filaments that exchange actin monomers with dissolved cytoplasmic actin, microtubules exchange ab-tubulin heterodimers with cytoplasmic tubulin. With the minus-ends anchored to the MTOC, microtubules grow or shorten through addition or loss of tubulin heterodimers at the plus-ends. Therefore, microtubules may grow steadily and then shorten (shrink) rapidly. The random transitions from their growth (caused by tubulin polymerization) to the shrinkage (caused by tubulin depolymerization) are called microtubule catastrophes; the transitions from the shrinkage to the growth are called rescues. Thus, microtubules oscillate between their growth and shrinkage. Such behavior is termed dynamic instability [56]. Dynamic instability of microtubules is determined by GTP hydrolysis. A molecule of GTP is bound to both a and b subunit of a tubulin heterodimer; however, only GTP bound to b-tubulin may be hydrolyzed to GDP. During polymerization, a subunit of a tubulin heterodimer from the cytoplasmic pool of tubulin comes into contact with the b-tubulin exposed at the microtubule plus-end. This promotes hydrolysis of GTP bound to the now interior b-tubulin. The GTP hydrolysis causes the tubulin depolymerization leading to the microtubule shrinkage. Until the GTP that is associated with b-tubulin at the microtubule plus-end is not hydrolyzed, tubulin continues to polymerize: the tubulin heterodimers are added faster than the GTP can be hydrolyzed. As a result, the microtubule grows. A rapidly growing microtubule may accumulate a few layers of GTP-bound tubulin at the plus-end. A GTP cap stabilizes the plus-end of a microtubule preventing its depolymerization. However, when the GTP has already hydrolyzed before a new tubulin heterodimer was incorporated, the intense tubulin depolymerization begins leading to the microtubule shrinkage.
3.2 Microtubules
27
Microtubules can be released from the centrosome; besides, long microtubules can break. As a result of that, a subpopulation of microtubules with free minus-ends can be found in some cells. These microtubules have the stable minus-ends unlike the plus-ends that show the dynamic instability [59]. On the released g-tubulin “primings,” the nucleation and growth of new microtubules can be initiated. In addition to the centrosome, the Golgi complex is a microtubule-organizing organelle. A large number of microtubules originate from the Golgi apparatus at its peripheral compartment; the Golgi membranes can directly stimulate the assembly of microtubules. Both centrosomal and Golgi-derived microtubules need g-tubulin for nucleation. In contrast to the high level of dynamics of centrosomal microtubules, the Golgi-based microtubules are stabilized early. Also, the Golgi-derived microtubules, in contrast to radial centrosomal microtubule arrays, are preferentially oriented toward the leading edge in polarized moving cells [60–62]. The dynamic instability of microtubules (the regime of their polymerization– depolymerization) is regulated by multiple microtubule-associated proteins. They include motor proteins and nonmotor proteins.
3.2.1 Motor Proteins The most prominent function of microtubules is an intracellular transport. It requires microtubule-associated molecular motors [45, 63]. Microtubule motor proteins are kinesins and dynein. There is also another motor protein, Eg5, that is active in mitotic spindle assembly. Kinesins and dynein are ATPases, which convert the chemical energy contained in ATP into the mechanical force used for their active movement along microtubules. The motor proteins directionally transport various membrane vesicles, organelles, proteins, and mRNAs by “walking” along microtubules. The molecule of the motor protein is fastened by its one end to the lateral side of microtubule, and by its other end to the “cargo” that needs to be transported, e.g., a specific molecule, an organelle, or an adjacent microtubule. This is similar to myosin connected with actin; the motor protein (in the presence of ATP) develops pulling action, which causes the displacement of the “cargo” along the microtubule or the mutual slip of microtubules relative to each other. The motor protein determines the direction of motion: kinesins transport various “cargos” toward the plus-ends of microtubules, while dynein moves toward their minus-ends. The example of the microtubule-based intracellular transport is the rapid transport of vesicles and mitochondria along the axons of neurons [45, 63–68]. Besides the transport function, some motor proteins can also directly influence the polymerization dynamics of microtubules and regulate their lengths. For example, some kinesins accelerate depolymerization of microtubules inducing their shortening [69, 70]. However, the microtubules themselves, in the absence of the microtubule-associated motor proteins, can also move some intracellular organelles. The growth and shrinkage of microtubules attached with their plus-ends to the organelles generate forces
28
3 Cytoskeleton
that can push and pull these organelles. For example, the microtubules attached to chromosomes (via the kinetochore) cause the movement of the chromosomes during metaphase and anaphase of mitosis.
3.2.2 Nonmotor Proteins Microtubule-associated nonmotor proteins include various proteins and protein families, which control the dynamic properties of microtubules and their interactions with various cellular structures. Protein tau polymerizes tubulin dimers and stabilizes microtubules. Microtubule plus-end tracking proteins (+TIPs) are specifically accumulated at the plus-ends of growing microtubules. +TIPs act as microtubule-stabilizing factors. Besides, they mediate the interactions between microtubule ends and actin cytoskeleton, and participate in intracellular transport [71, 72]. Moreover, +TIP group includes the end-binding protein1 (EB1) that stimulates spontaneous nucleation and growth of microtubules [73]. EB1 binds to another member of the +TIP’s group, the adenomatous polyposis coli (APC) protein that is product of a tumor suppressor gene. APC participates in many cellular processes [74]. It has multiple domains, through which APC binds to various proteins. APC interacts with microtubules and accumulates at the plus-ends of microtubules in cell protrusions. APC may regulate the polymerization dynamics of these microtubules: APC-decorated microtubules have an increased phase of their growth compared with microtubules that are not decorated by APC. Increased time of growth of microtubules mediated by APC in cell protrusions plays a significant role in directional cell migration [75–78]. A microtubule-associated protein, XMAP215, moves with growing microtubule plus-ends, where it catalyzes the addition of tubulin subunits [70]. Septins are proteins that are colocalized with microtubules and probably play an important role in microtubule dynamics regulation by interacting with microtubuleassociated proteins. Septins can be associated with both microtubules and actin filaments to link these cytoskeleton systems. Septins polymerize to form filaments approximately 8 nm in thickness that can assemble along actin bundles [79]. Posttranslational tubulin modifications influence motor- and nonmotor microtubuleassociated proteins; thereby, tubulin modifications are involved in the regulation of microtubule dynamics [80]. The dynamic instability of microtubules (the regime of their assemblydepolymerization) is a necessary condition for normal functioning of cells. The inhibition of the microtubule dynamic instability by special agents leads to cell cycle arrest. Some plant poisons, e.g., colchicines or colcemide, while being joined to the monomers of tubulin, prevent its polymerization and block the growth of microtubules. Since in this case the depolymerization continues, microtubules are gradually destroyed, including microtubules of mitotic spindle, which stops cell division at the stage of mitosis. The antitumor plant alkaloids vincristine and vinblastine
3.3 Intermediate Filaments
29
possess an analogous mechanism of action. Another plant agent, taxol, on the contrary, does not suppress, but activates the polymerization of tubulin, preventing the depolymerization of microtubules: they become stable, and do not get short. However, this stabilization also stops the division of a cell at the stage of mitosis. Many of such microtubule-targeting agents (vincristine, vinblastine, taxol, and others) are used in cancer chemotherapy [81–83]. Intracellular microtubule-based transport of organelles and molecules is necessary for the maintenance of asymmetric polarized cell morphology (cell polarization) and cell migration (see Sect. 6.1.2) [1, 84, 85]. The transported molecules include those locally inhibiting cell contractility, thereby promoting the disassembly of the individual cell–extracellular matrix adhesions (see Sect. 4.2.1) [86]. The proteins that regulate actin cytoskeleton can also be transported by microtubules [86–89]. Therefore, microtubules take part in the regulation of cell contractility, actin cytoskeleton remodeling, cell–extracellular matrix adhesion dynamics, and cell migration. Actin cytoskeleton and the system of microtubules work coordinately. The mechanical and regulatory connections between these two cytoskeletal systems contribute to this coordination. Transverse mechanical connections between actin filaments and microtubules are mediated by several types of cross-linking proteins. The Rho family of GTPases regulates dynamics and organization not only of the actin cytoskeleton but also of microtubules (see Sect. 5.3.5). Both cytoskeletal systems participate cooperatively in the formation of pseudopodia and in the assembly and functioning of special adhesive structures, which connect the cell with the extracellular matrix, determining the capability of cells to active movement – cell migration.
3.3 Intermediate Filaments Intermediate filaments (IFs) are proteinaceous fibrillar structures intermediate in their widths (8–12 nm) between actin filaments and microtubules (Fig. 3.1). Unlike those cytoskeleton elements, IFs are flexible and have a unique ability to withstand substantial deformations: single IFs can stretch to more than 3 times their initial length before breaking [90–92]. IFs are composed of a variety of tissue-specific proteins [91, 93–95]. The human IF protein family includes about 70 members divided into five groups. In various cells and tissues, IFs are formed by proteins of different groups [94]. In epithelial cells IFs consist of proteins called keratins. Acid and basic keratins bind each other to form acidic-basic heterodimers that then associate to make a keratin filament. About 20 keratin isoforms compose epithelial keratin IFs. Different sets of keratins are expressed in different types of epithelia and even in different areas of the same epithelium. The IFs composed of protein vimentin are widely distributed. They are characteristic of fibroblasts and many other cell types.
30
3 Cytoskeleton
In muscle cells, IFs are composed of vimentin, copolymerized with another protein called desmin. Proteins syncoilin and synemin (also called desmuslin) also compose IFs in muscle cells. In glial cells of central nervous system, IFs (named glial filaments) are composed of vimentin copolymerized with the protein called glial fibrillary acidic protein (GFAP). In neurons, IFs (called neurofilaments) are composed of several different proteins including internexin, peripherin, and nestin. (The latter is found in developing neurons and is lost as development proceeds). A distinct type of IFs is located inside a cell nucleus and forms thin “nuclear lamina,” a dense network of interconnected IFs composed of the proteins lamins. Nuclear lamina is associated with the inner face of the nuclear membrane. Lamins and connected nuclear envelope proteins participate in gene expression. Mutations in the genes encoding lamins cause human diseases called laminopathies [96, 97]. Tissue differences in the IF protein composition are used in immunopathological diagnostics of the tumors of unclear genesis. For example, the use of monoclonal antibodies to keratins 8 and 17 in immunohistological assay makes it possible to distinguish between morphologically similar benign and malignant human breast tumors [98]. IFs are dynamic and motile elements of the cytoskeleton. They undergo assembly from their nonfilamentous precursors called “particles.” Molecular motors including those of microtubules, kinesins and dyneins, rapidly translocate the “particles” and short IFs along microtubules to the specific regions of the cytoplasm, where they assemble into mature IFs [91, 99, 100]. IFs have a unique combination of mechanical properties, such as extensibility, flexibility, and resilience. They serve structural functions maintaining mechanical integrity and viscoelastic properties of cells and tissues [92, 95]. IFs apparently can also play a role in transduction of mechanical signals mediated by the extracellular matrix [91, 92, 99]. However, because of the possibility of IF association with some signaling molecules, IFs can apparently have other functions besides supporting mechanical integrity of a cell [99]. IFs are involved in various human diseases of skin, heart, and some other organs. Mutations of genes encoding IF proteins cause more than 80 human tissue-specific diseases, many of which are probably linked to the nonmechanical functions of IFs [91, 101, 102]. IFs interact with each other and with other cytoskeletal systems in a cell via IF-associated proteins. The protein filaggrin is involved in cross-linking of keratin IFs. Cytoskeletal systems of actin filaments, microtubules, and IFs are connected to each other by special proteins named cytoskeleton linkers. A family of the linker proteins, plakins consists of large multidomain proteins that cross-link different cytoskeleton elements. One of the members of the plakin protein family, plectin, mediates the interactions of IFs with actin filaments and microtubules. Plakins also connect IFs to the adhesion structures such as desmosomes and hemidesmosomes in epithelial cells [103, 104].
References
31
References 1. Vasiliev JM (2004) Cytoskeletal mechanisms responsible for invasive migration of neoplastic cells. Int J Dev Biol 48(5–6):425– 439. doi:10.1387/ijdb.041806jv DOI:dx.doi.org 2. Lindberg U, Karlsson R, Lassing I, Schutt CE, Höglund AS (2008) The microfilament system and malignancy. Semin Cancer Biol 18(1):2–11. doi:10.1016/j.semcancer.2007.10.002 DOI:dx.doi.org 3. Hall A (2009) The cytoskeleton and cancer. Cancer Metastasis Rev 28(1–2):5–14. doi:10.1007/ s10555-008-9166-3 DOI:dx.doi.org 4. Pollard TD, Cooper JA (2009) Actin, a central player in cell shape and movement. Science 326(5957):1208–1212. doi:10.1126/science.1175862 DOI:dx.doi.org 5. Saha S, Mundia MM, Zhang F, Demers RW, Korobova F, Svitkina T, Perieteanu AA, Dawson JF, Kashina A (2010) Arginylation regulates intracellular actin polymer level by modulating actin properties and binding of capping and severing proteins. Mol Biol Cell 21(8):1350–1361. doi:10.1091/mbc.E09-09-0829 DOI:dx.doi.org 6. Dugina V, Zwaenepoel I, Gabbiani G, Clément S, Chaponnier C (2009) Beta and gammacytoplasmic actins display distinct distribution and functional diversity. J Cell Sci 122(Pt 16):2980–2988. doi:10.1242/jcs.041970 DOI:dx.doi.org 7. Pollard TD (1999) Introduction to actin and actin-binding proteins. In: Kreis T and Vale R (eds) Guidebook to the Cytokeletal and Motor Proteins. Oxford University Press Inc, New York 8. Higgs HN, Pollard TD (2001) Regulation of actin filament network formation through ARP2/3 complex: activation by a diverse array of proteins. Annu Rev Biochem 70:649–676. doi:10.1146/annurev.biochem.70.1.649 DOI:dx.doi.org 9. Goley ED, Welch MD (2006) The ARP2/3 complex: an actin nucleator comes of age. Nat Rev Mol Cell Biol 7(10):713–726. doi:10.1038/nrm2026 DOI:dx.doi.org 10. Chhabra ES, Higgs HN (2007) The many faces of actin: matching assembly factors with cellular structures. Nat Cell Biol. 9(10):1110–1121. doi:10.1038/ncb1007-1110 DOI:dx.doi.org 11. Cai L, Bear JE (2008) Peering deeply inside the branch. J Cell Biol 180(5):853–855. doi:10.1083/jcb.200802062 DOI:dx.doi.org 12. Dominguez R (2009) Actin filament nucleation and elongation factors--structure-function relationships. Crit Rev Biochem Mol Biol 44(6):351–366. doi:10.3109/10409230903277340 DOI:dx.doi.org 13. Campellone KG, Welch MD (2010) A nucleator arms race: cellular control of actin assembly. Nat Rev Mol Cell Biol 11(4):237–251. doi:10.1038/nrm2867 DOI:dx.doi.org 14. Mellor H (2010) The role of formins in filopodia formation. Biochim Biophys Acta 1803(2):191–200. doi:10.1016/j.bbamcr.2008.12.018 DOI:dx.doi.org 15. Takenawa T, Suetsugu S (2007) The WASP-WAVE protein network: connecting the membrane to the cytoskeleton. Nat Rev Mol Cell Biol 8(1):37–48. doi:10.1038/nrm2069 DOI:dx.doi.org 16. Pollitt AY, Insall RH (2009) WASP and SCAR/WAVE proteins: the drivers of actin assembly. J Cell Sci 122(Pt 15):2575–2578. doi:10.1242/jcs.023879 DOI:dx.doi.org 17. Kurisu S, Takenawa T (2009) The WASP and WAVE family proteins. Genome Biol 10(6):226. doi:10.1186/gb-2009-10-6-226 DOI:dx.doi.org 18. Derivery E, Gautreau A (2010) Generation of branched actin networks: assembly and regulation of the N-WASP and WAVE molecular machines. Bioessays 32(2):119–131. doi:10.1002/bies.200900123 DOI:dx.doi.org 19. Antón IM, Jones GE (2006) WIP: a multifunctional protein involved in actin cytoskeleton regulation. Eur J Cell Biol 85(3–4):295–304 20. Antón IM, Jones GE, Wandosell F, Geha R, Ramesh N (2007) WASP-interacting protein (WIP): working in polymerisation and much more. Trends Cell Biol 17(11):555–562 21. Renault L, Bugyi B, Carlier MF (2008) Spire and Cordon-bleu: multifunctional regulators of actin dynamics. Trends Cell Biol 18(10):494–504. doi:10.1016/j.tcb.2008.07.008 DOI:dx.doi.org
32
3 Cytoskeleton
22. Chesarone MA, Goode BL (2009) Actin nucleation and elongation factors: mechanisms and interplay. Curr Opin Cell Biol 21(1):28–37. doi:10.1016/j.ceb.2008.12.001 DOI:dx.doi.org 23. Goode BL, Eck MJ (2007) Mechanism and function of formins in the control of actin assembly. Annu Rev Biochem 76:593–627. doi:10.1146/annurev.biochem.75.103004.142647 DOI:dx.doi.org 24. Paul AS, Pollard TD (2009) Review of the mechanism of processive actin filament elongation by formins. Cell Motil Cytoskeleton 66(8):606–617. doi:10.1002/cm.20379 DOI:dx.doi.org 25. Young KG, Copeland JW (2010) Formins in cell signaling. Biochim Biophys Acta 1803(2):183–190. doi:10.1016/j.bbamcr.2008.09.017 DOI:dx.doi.org 26. Yarmola EG, Bubb MR (2009) How depolymerization can promote polymerization: the case of actin and profilin. Bioessays 31(11):1150–1160. doi:10.1002/bies.200900049 DOI:dx.doi.org 27. Birbach A (2008) Profilin, a multi-modal regulator of neuronal plasticity. Bioessays 30(10):994–1002. doi:10.1002/bies.20822 DOI:dx.doi.org 28. Silacci P, Mazzolai L, Gauci C, Stergiopulos N, Yin HL, Hayoz D (2004) Gelsolin superfamily proteins: key regulators of cellular functions. Cell Mol Life Sci 61(19–20):2614–23. doi:10.1007/s00018-004–4225-6 DOI:dx.doi.org 29. Ono S (2007) Mechanism of depolymerization and severing of actin filaments and its significance in cytoskeletal dynamics. Int Rev Cytol 258:1–82. doi:10.1016/S0074– 7696(07)58001-0 DOI:dx.doi.org 30. Svitkina TM, Borisy GG (1999) Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol 145(5):1009–1026. doi:10.1083/jcb.145.5.1009 DOI:dx.doi.org 31. Van Troys M, Huyck L, Leyman S, Dhaese S, Vandekerkhove J, Ampe C (2008) Ins and outs of ADF/cofilin activity and regulation. Eur J Cell Biol 87(8–9):649–667 32. Oser M. Condeelis J (2009) The cofilin activity cycle in lamellipodia and invadopodia. J Cell Biochem 108(6):1252–1262. doi:10.1002/jcb.22372 DOI:dx.doi.org 33. Buday L, Downward J (2007) Roles of cortactin in tumor pathogenesis. Biochim Biophys Acta 1775(2):263–273. doi:10.1016/j.bbcan.2006.12.002 DOI:dx.doi.org 34. Ammer AG, Weed SA (2008) Cortactin branches out: roles in regulating protrusive actin dynamics. Cell Motil Cytoskeleton 65(9):687–707. doi:10.1002/cm.20296 DOI:dx.doi.org 35. Ren G, Crampton MS, Yap AS (2009) Cortactin: Coordinating adhesion and the actin cytoskeleton at cellular protrusions. Cell Motil Cytoskeleton 66(10):865–873. doi:10.1002/ cm.20380 DOI:dx.doi.org 36. Khurana S, George SP (2008) Regulation of cell structure and function by actin-binding proteins: villin’s perspective. FEBS Lett 582(14):2128–2139. doi:10.1016/j.febslet.2008.02.040 DOI:dx.doi.org 37. Kostyukova AS (2008) Capping complex formation at the slow-growing end of the actin filament. Biochemistry (Mosc) 73(13):1467–1472. doi:10.1134/S0006297908130075 DOI:dx.doi.org 38. Sjöblom B, Salmazo A, Djinović-Carugo K (2008) Alpha-actinin structure and regulation. Cell Mol Life Sci 65(17):2688–2701 39. Popowicz GM, Schleicher M, Noegel AA, Holak TA (2006) Filamins: promiscuous organizers of the cytoskeleton. Trends Biochem Sci 31(7):411–419. doi:10.1016/j.tibs.2006.05.006 DOI:dx.doi.org 40. Delanote V, Vandekerckhove J, Gettemans J (2005) Plastins:versatile modulators of actin organization in (patho) physiological cellular processes. Acta Pharmacol Sin 26(7):769–779. doi:10.1111/j.1745-7254.2005.00145.x DOI:dx.doi.org 41. Enomoto A, Ping J, Takahashi M (2006) Girdin, a novel actin-binding protein, and its family of proteins possess versatile functions in the Akt and Wnt signaling pathways. Ann N Y Acad Sci 1086:169–184. doi:10.1196/annals.1377.016 DOI:dx.doi.org 42. Niggli V, Rossy J (2008) Ezrin/radixin/moesin: versatile controllers of signaling molecules and of the cortical cytoskeleton. Int J Biochem Cell Biol 40(3):344–349. doi:10.1016/j. biocel.2007.02.012 DOI:dx.doi.org 43. Fehon RG, McClatchey AI, Bretscher A (2010) Organizing the cell cortex: the role of ERM proteins. Nat Rev Mol Cell Biol 11(4):276–287. doi:10.1038/nrm2866 DOI:dx.doi.org
References
33
44. Chibalina MV, Puri C, Kendrick-Jones J, Buss F(2009) Potential roles of myosin VI in cell motility. Biochem Soc Trans 37(Pt 5):966–970. doi:10.1042/BST0370966 DOI:dx.doi.org 45. Ross JL, Ali MY, Warshaw DM (2008) Cargo transport: molecular motors navigate a complex cytoskeleton. Curr Opin Cell Biol 20(1):41–47. doi:10.1016/j.ceb.2007.11.006 DOI:dx.doi.org 46. Semenova I, Burakov A, Berardone N, Zaliapin I, Slepchenko B, Svitkina T, Kashina A, Rodionov V (2008) Actin dynamics is essential for myosin-based transport of membrane organelles. Curr Biol 18(20):1581–1586. doi:10.1016/j.cub.2008.08.070 DOI:dx.doi.org 47. Woolner S, Bement WM (2009) Unconventional myosins acting unconventionally. Trends Cell Biol 19(6):245–252. doi:10.1016/j.tcb.2009.03.003 DOI:dx.doi.org 48. Nambiar R, McConnell RE, Tyska MJ (2010) Myosin motor function: the ins and outs of actin-based membrane protrusions. Cell Mol Life Sci 67(8):1239–1254. doi:10.1007/s00018009-0254-5 DOI:dx.doi.org 49. Gunning P, O’Neill G, Hardeman E (2008) Tropomyosin-based regulation of the actin cytoskeleton in time and space. Physiol Rev 88(1):1–35. doi:10.1152/physrev.00001.2007 DOI:dx.doi.org 50. O’Neill GM, Stehn J, Gunning PW (2008) Tropomyosins as interpreters of the signalling environment to regulate the local cytoskeleton. Semin Cancer Biol 18(1):35–44 51. Lin JJ, Li Y, Eppinga RD, Wang Q, Jin JP (2009) Chapter 1: roles of caldesmon in cell motility and actin cytoskeleton remodeling. Int Rev Cell Mol Biol 274:1–68. doi:10.1016/ S1937–6448(08)02001-7 DOI:dx.doi.org 52. Vicente-Manzanares M, Ma X, Adelstein RS, Horwitz AR (2009) Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat Rev Mol Cell Biol 10(11):778–790. doi:10.1038/nrm2786 DOI:dx.doi.org 53. Borisy GG, Svitkina TM (2000) Actin machinery: pushing the envelope. Curr Opin Cell Biol 12(1):104–112. doi:10.1016/S0955-0674(99)00063-0 DOI:dx.doi.org 54. Olson EN, Nordheim A (2010) Linking actin dynamics and gene transcription to drive cellular motile functions. Nat Rev Mol Cell Biol 11(5):353–365. doi:10.1038/nrm2890 DOI:dx.doi.org 55. Wade RH (2007) Microtubules: an overview. Methods Mol Med 137:1–16. doi:10.1007/ s12033-009-9193-5 DOI:dx.doi.org 56. Wade RH (2009) On and around microtubules: an overview. Mol Biotechnol 43(2):177–191. doi:10.1007/s12033-009-9193-5 DOI:dx.doi.org 57. Wiese C, Zheng Y (2006) Microtubule nucleation: gamma-tubulin and beyond. J Cell Sci 119(Pt 20):4143– 4153. doi:10.1242/jcs.03226 DOI:dx.doi.org 58. Raynaud-Messina B, Merdes A (2007) Gamma-tubulin complexes and microtubule organization. Curr Opin Cell Biol 19(1):24 –30 59. Vorobjev IA, Svitkina TM, Borisy GG (1997) Cytoplasmic assembly of microtubules in cultured cells. J Cell Sci 110 ( Pt 21):2635–2645 60. Efimov A, Kharitonov A, Efimova N, Loncarek J, Miller PM, Andreyeva N, Gleeson P, Galjart N, Maia AR, McLeod IX, Yates JR 3rd, Maiato H, Khodjakov A, Akhmanova A, Kaverina I (2007) Asymmetric CLASP-dependent nucleation of noncentrosomal microtubules at the trans-Golgi network. Dev Cell 12(6):917–930. doi:10.1016/j.devcel.2007.04.002 DOI:dx.doi.org 61. Vinogradova T, Miller PM, Kaverina I (2009) Microtubule network asymmetry in motile cells: role of Golgi-derived array. Cell Cycle 8(14):2168–2174. doi:10.4161/cc.8.14.9074 DOI:dx.doi.org 62. Miller PM, Folkmann AW, Maia AR, Efimova N, Efimov A, Kaverina I (2009) Golgi-derived CLASP-dependent microtubules control Golgi organization and polarized trafficking in motile cells. Nat Cell Biol 11(9):1069–1080. doi:10.1038/ncb1920 DOI:dx.doi.org 63. Gross SP, Vershinin M, Shubeita GT. Cargo transport: two motors are sometimes better than one. Curr Biol. 2007; 17(12):R478–86. doi:10.1016/j.cub.2007.04.025 DOI:dx.doi.org 64. Hirokawa N, Nitta R, Okada Y (2009) The mechanisms of kinesin motor motility: lessons from the monomeric motor KIF1A. Nat Rev Mol Cell Biol 10(12):877–884. doi:10.1038/ nrm2807 DOI:dx.doi.org 65. Marx A, Hoenger A, Mandelkow E (2009) Structures of kinesin motor proteins. Cell Motil Cytoskeleton 66(11):958–966. doi:10.1002/cm.20392 DOI:dx.doi.org
34
3 Cytoskeleton
66. Verhey KJ, Hammond JW (2009) Traffic control: regulation of kinesin motors. Nat Rev Mol Cell Biol 10(11):765–777. doi:10.1038/nrm2782 DOI:dx.doi.org 67. Kardon JR, Vale RD (2009) Regulators of the cytoplasmic dynein motor. Nat Rev Mol Cell Biol 10(12):854–865. doi:10.1038/nrm2804 DOI:dx.doi.org 68. Carter AP, Vale RD (2010) Communication between the AAA+ ring and microtubule-binding domain of dynein. Biochem Cell Biol 88(1):15–21. doi:10.1139/O09-127 DOI:dx.doi.org 69. Hunter AW, Wordeman L (2000) How motor proteins influence microtubule polymerization dynamics. J Cell Sci 113 Pt 24:4379–4389 70. Howard J, Hyman AA (2007) Microtubule polymerases and depolymerases. Curr Opin Cell Biol 19(1):31–35. doi:10.1016/j.ceb.2006.12.009 DOI:dx.doi.org 71. Akhmanova A, Steinmetz MO (2008) Tracking the ends: a dynamic protein network controls the fate of microtubule tips. Nat Rev Mol Cell Biol 9(4):309–322. doi:10.1038/nrm2369 DOI:dx.doi.org 72. Jaworski J, Hoogenraad CC, Akhmanova A (2008) Microtubule plus-end tracking proteins in differentiated mammalian cells. Int J Biochem Cell Biol 40(4):619–637. doi:10.1016/j. biocel.2007.10.015 DOI:dx.doi.org 73. Vitre B, Coquelle FM, Heichette C, Garnier C, Chrétien D, Arnal I (2008) EB1 regulates microtubule dynamics and tubulin sheet closure in vitro. Nat Cell Biol 10(4):415–421. doi:10.1038/ncb1703 DOI:dx.doi.org 74. Brocardo M, Henderson BR (2008) APC shuttling to the membrane, nucleus and beyond. Trends Cell Biol 18(12):587–596. doi:10.1016/j.tcb.2008.09.002 DOI:dx.doi.org 75. Akiyama T, Kawasaki Y (2006) Wnt signalling and the actin cytoskeleton. Oncogene 25(57):7538–7544. doi:10.1038/sj.onc.1210063 DOI:dx.doi.org 76. Aoki K, Taketo MM (2007) Adenomatous polyposis coli (APC): a multi-functional tumor suppressor gene. J Cell Sci 120(Pt 19):3327–3335. doi:10.1242/jcs.03485 DOI:dx.doi.org 77. Barth AI, Caro-Gonzalez HY, Nelson WJ (2008) Role of adenomatous polyposis coli (APC) and microtubules in directional cell migration and neuronal polarization. Semin Cell Dev Biol 19(3):245–251. doi:10.1016/j.semcdb.2008.02.003 DOI:dx.doi.org 78. McCartney BM, Näthke IS (2008) Cell regulation by the Apc protein Apc as master regulator of epithelia. Curr Opin Cell Biol 20(2):186–193. doi:10.1016/j.ceb.2008.02.001 DOI:dx.doi.org 79. Silverman-Gavrila RV, Silverman-Gavrila LB (2008) Septins: new microtubule interacting partners. ScientificWorldJournal 8:611–620. doi:10.1113/jphysiol.2008.151837 DOI:dx.doi.org 80. Hammond JW, Cai D, Verhey KJ (2008) Tubulin modifications and their cellular functions. Curr Opin Cell Biol 20(1):71–76. doi:10.1016/j.ceb.2007.11.010 DOI:dx.doi.org 81. Jordan MA, Kamath K (2007) How do microtubule-targeted drugs work? An overview. Curr Cancer Drug Targets 7(8):730–742. doi:10.2174/156800907783220417 DOI:dx.doi.org 82. Pasquier E, Kavallaris M (2008) Microtubules: a dynamic target in cancer therapy. IUBMB Life 60(3):165–170. doi:10.1002/iub.25 DOI:dx.doi.org 83. Carlson RO (2008) New tubulin targeting agents currently in clinical development. Expert Opin Investig Drugs 17(5):707–722. doi:10.1517/13543784.17.5.707 DOI:dx.doi.org 84. Pletjushkina OJ, Ivanova OJ, Kaverina IN, Vasiliev JM (1994) Taxol-treated fibroblasts acquire an epithelioid shape and a circular pattern of actin bundles. Exp Cell Res 212(2):201–208. doi:10.1006/excr.1994.1135 DOI:dx.doi.org 85. Siegrist SE, Doe CQ (2007) Microtubule-induced cortical cell polarity. Genes Dev 21(5):483–496. doi:10.1101/gad.1511207 DOI:dx.doi.org 86. Kaverina I, Krylyshkina O, Small JV (1999) Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J Cell Biol 146(5):1033–1044. doi:10.1083/ jcb.146.5.1033 DOI:dx.doi.org 87. Elbaum M, Chausovsky A, Levy ET, Shtutman M, Bershadsky AD (1999) Microtubule involvement in regulating cell contractility and adhesion-dependent signalling: a possible mechanism for polarization of cell motility. Biochem Soc Symp 65:147–72 88. Small JV, Geiger B, Kaverina I, Bershadsky A (2002) How do microtubules guide migrating cells? Nat Rev Mol Cell Biol 3(12):957–964. doi:10.1038/nrm971 DOI:dx.doi.org
References
35
89. Basu R, Chang F (2007) Shaping the actin cytoskeleton using microtubule tips. Curr Opin Cell Biol 19(1):88–94. doi:10.1016/j.ceb.2006.12.012 DOI:dx.doi.org 90. Wagner OI, Rammensee S, Korde N, Wen Q, Leterrier JF, Janmey PA (2007) Softness, strength and self-repair in intermediate filament networks. Exp Cell Res 313(10):2228–2235. doi:10.1016/j.yexcr.2007.04.025 DOI:dx.doi.org 91. Kreplak L, Fudge D (2007) Biomechanical properties of intermediate filaments: from tissues to single filaments and back. Bioessays 29(1):26–35. doi:10.1002/bies.20514 DOI:dx.doi.org 92. Qin Z, Buehler MJ, Kreplak L (2010) A multi-scale approach to understand the mechanobiology of intermediate filaments. J Biomech 43(1):15–22. doi:10.1016/j.jbiomech.2009.09.004 DOI:dx.doi.org 93. Oshima RG (2007) Intermediate filaments: a historical perspective. Exp Cell Res 313(10):1981–1994. doi:10.1016/j.yexcr.2007.04.007 DOI:dx.doi.org 94. Minin AA, Moldaver MV (2008) Intermediate vimentin filaments and their role in intracellular organelle distribution. Biochemistry (Mosc). 73(13):1453–1466. doi:10.1134/ S0006297908130063 DOI:dx.doi.org 95. Herrmann H, Strelkov SV, Burkhard P, Aebi U (2009) Intermediate filaments: primary determinants of cell architecture and plasticity. J Clin Invest 119(7):1772–1783. doi:10.1172/ JCI38214 DOI:dx.doi.org 96. Kreplak L, Richter K, Aebi U, Herrmann H (2008) Electron microscopy of intermediate filaments: teaming up with atomic force and confocal laser scanning microscopy. Methods Cell Biol 88:273–297. doi:10.1016/S0091-679X(08)00415-9 DOI:dx.doi.org 97. Andrés V, González JM (2009) Role of A-type lamins in signaling, transcription, and chromatin organization. J Cell Biol 187(7):945–957. Review 98. Guelstein VI, Tchypysheva TA, Ermilova VD, Ljubimov AV (1993) Myoepithelial and basement membrane antigens in benign and malignant human breast tumors. Int J Cancer 53(2):269–277. doi:10.1002/ijc.2910530217 DOI:dx.doi.org 99. Chou YH, Flitney FW, Chang L, Mendez M, Grin B, Goldman RD (2007) The motility and dynamic properties of intermediate filaments and their constituent proteins. Exp Cell Res 313(10):2236–43. doi:10.1016/j.yexcr.2007.04.008 DOI:dx.doi.org 100. Godsel LM, Hobbs RP, Green KJ (2008) Intermediate filament assembly: dynamics to disease. Trends Cell Biol 18(1):28–37. doi:10.1016/j.tcb.2007.11.004 DOI:dx.doi.org 101. Eriksson JE, Dechat T, Grin B, Helfand B, Mendez M, Pallari HM, Goldman RD (2009) Introducing intermediate filaments: from discovery to disease. J Clin Invest 119(7):1763–1771. doi:10.1172/JCI38339 DOI:dx.doi.org 102. Omary MB (2009) “IF-pathies”: a broad spectrum of intermediate filament-associated diseases. J Clin Invest 119(7):1756–1762. doi:10.1172/JCI39894 DOI:dx.doi.org 103. Svitkina TM, Verkhovsky AB, Borisy GG (1996) Plectin sidearms mediate interaction of intermediate filaments with microtubules and other components of the cytoskeleton. J Cell Biol 135(4):991–1007. doi:10.1083/jcb.135.4.991 DOI:dx.doi.org 104. Sonnenberg A, Liem RK (2007) Plakins in development and disease. Exp Cell Res 313(10):2189–2203. doi:10.1016/j.yexcr.2007.03.039 DOI:dx.doi.org
wwwwwwwwwwwwwwww
Chapter 4
Pseudopodia and Adhesion Structures
Abstract Pseudopodia, the cytoplasmic outgrowths of lamellar shape (lamellipodia, ruffles), filamentous shape (filopodia), or spherical shape (blebs), are the “driving organs” formed by cells to spread and migrate, and also can have sensory or metabolic functions. The attachment of lamellipodia or filopodia to the extracellular matrix is achieved by different types of special adhesion structures, such as focal contacts (focal adhesions), focal complexes, fibrillar adhesions, hemidesmosomes, podosomes, and invadopodia. The dynamic properties and regulation of the main adhesion structures, focal contacts, play a pivotal role in cell migration. Other types of adhesion structures, podosomes and invadopodia, can locally degrade the extracellular matrix components, thereby creating spaces for the migration of normal cells or for the invasion of malignant tumor cells.
4.1 The Formation of Pseudopodia In the process of spreading of tissue cells, the change of their shapes, during migration, and in the cell interactions to the extracellular matrix, the cells form pseudopodia, which are cytoplasmic outgrowths confined by the cell membrane. The pseudopodia are of several morphological types: (a) Filopodia are the filamentous outgrowths that can be short or very long (10– 15 mm). Most frequently, they are developed at the cell edge (Figs. 4.1, 4.2, and 4.4); in the beginning of cell spreading, filopodia are formed at the cell base (Fig. 4.3). The distal ends of filopodia attached to the extracellular matrix or to the adjacent cells sometimes branch; they can also end with bulbs (Fig. 4.4). A filopodium contains actin filaments oriented along it. Formations, which are morphologically very similar to the filopodia, but which are not pseudopodia, are called retraction fibrils. They appear at the edges of spread cells while their retraction. The retraction can be observed with the entrance of the cell into mitosis (Fig. 4.5); the cell retraction accompanies the process of gradual separation of the cell from the extracellular matrix (Fig. 4.6a–c). Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_4, © Springer Science+Business Media, LLC 2011
37
38
4 Pseudopodia and Adhesion Structures
Fig. 4.1 Fibroblastic rat cells. Filopodia (arrows) coming out of the cell edges. SEM (scanning electron microscopy*). Scale bar, 36 mm
Fig. 4.2 Transformed mouse fibroblasts. Filopodia (arrows) coming out of the cell edges. SEM. Scale bar, 13 mm
(b) Lamellipodia are comparatively small (2–5 mm in width) lamellar outgrowths abutting the extracellular matrix surface. Lamellipodia are developed at the bases of cells at the early stages of their spreading (Fig. 4.7); they are formed at * All SEM photos were made by the author of this book.
4.1 The Formation of Pseudopodia
39
Fig. 4.3 Fibroblastic mouse cell in the beginning of its spreading. Filopodia (arrow) coming out of the cell base. SEM. Scale bar, 8 mm Fig. 4.4 Transformed hamster fibroblast. Filopodia (arrow) and ruffles (double arrows) are at the cell edge. SEM. Scale bar, 2.5 mm
40
4 Pseudopodia and Adhesion Structures
Fig. 4.5 Transformed hamster fibroblasts. Retraction fibers (arrows) coming out of the retracted mitotic cell. SEM. Scale bar, 6 mm
Fig. 4.6 (a–c) Fibroblastic mouse cells at the different stages of their retraction. Retraction fibers (arrows) coming out of the edges of the progressively retracting cells (a, b) and coming out of the completely retracted cell just before its detachment (c). SEM. Scale bar, 5 mm
the free edge of the lamellar cytoplasm. Lamellar cytoplasm, also called lamella, with lamellipodia at its free edge, surrounds the central prominent part of the spreading cell (Fig. 4.8). In a completely spread cell, the lamellar cytoplasm can have the shape of a wide thin rim along the entire cell perimeter (Fig. 4.9), or it can be divided into a few peripheral lamellas. Lamellipodia contain the branched networks of actin filaments that are oriented in a certain way with respect to the leading cell edge [1].
4.1 The Formation of Pseudopodia
41
Fig. 4.7 Fibroblastic hamster cell at the early stage of its spreading. Lamellipodia (arrows) are in the cell base. SEM. Scale bar, 6 mm
Fig. 4.8 Fibroblastic mouse cell in its spreading process. The lamellar cytoplasm (arrow) with lamellipodia (double arrows) at its free edge surrounds the central prominent part of the cell. SEM. Scale bar, 8.5 mm
42
4 Pseudopodia and Adhesion Structures
Fig. 4.9 Fibroblastic mouse cell on the termination of the spreading process. The entire cell periphery is the wide lamellar cytoplasm (arrow). SEM. Scale bar, 14 mm
Filopodia and lamellipodia possess the ability to attach to the extracellular matrix and then to retract. This ability makes these pseudopodia the “driving organs” with the aid of which the cells can spread and move. Filopodia have also sensory or exploratory functions of “detectors,” with the aid of which the cells select the parts of the matrix optimal for the cell attachment. Filopodia play a role in cell guidance toward a chemoattractant [2]. (c) Blebs are spherical protrusions that can participate in cell migration in special conditions of reduced adhesiveness of the substratum surface [3, 4]. (d) Ruffles are the lamellar outgrowths, which have the form of “frills.” The width of the ruffles varies, frequently it is 6–8 mm, and their extent can reach 10 mm. In contrast to the lamellipodia, ruffles are elevated above the matrix surface. Ruffles are most frequently developed at cell edges (Figs. 4.4 and 4.10); ruffles like lamellipodia are often formed at the free edge of the lamellar cytoplasm (Fig. 4.11). They can also be observed on the spherical surfaces of the cells in their suspended states, and at the bases of cells in the initial stages of their spreading (Fig. 4.12). Ruffles are dynamic formations: the period between the appearance of a ruffle and its collapse is less than 1 min. They have metabolic functions, participating in the phagocytosis or in the capture of drops of the nutrient medium by pinocytosis. Ruffles contain irregularly distributed actin filaments, but they do not contain microtubules. The mechanical forces, source of which is the polymerization of actin filaments, cause formation of pseudopodia. The growing barbed ends of filaments
4.1 The Formation of Pseudopodia
43
Fig. 4.10 Transformed mouse fibroblasts. Ruffles (arrows) are at the cell edges. SEM. Scale bar, 6 mm
Fig. 4.11 Fibroblastic mouse cell in its spreading process. The ruffles (arrows) are at the free edge of the lamellar cytoplasm. SEM. Scale bar, 7.5 mm
44
4 Pseudopodia and Adhesion Structures
Fig. 4.12 Transformed mouse fibroblast at the initial stage of its spreading. Ruffle (arrow) is seen in the cell base. SEM. Scale bar, 3 mm
forming the dendritic network push the cell membrane forward at its relatively wide parts, making the flattened protrusions, lamellipodia. Some of growing actin filaments within the lamellipodial dendritic network acquire a privileged status by binding specific molecules to their barbed ends, which protects them from capping and mediates the association of the barbed ends with each other. These filaments with their associated barbed ends are sewed together into the long bundles that protrude the cell membrane, forming very narrow long outgrowths, filopodia. Thus, the pseudopodium formation is based on the polymerization of actin filaments and subsequent reorganizations of the actin cytoskeleton. These processes are controlled, among other factors, by the proteins of Rho family of GTPases (see Sect. 5.3.5).
4.2 Cell–Extracellular Matrix Adhesion Structures Specific adhesion structures ensure the attachment of tissue cells to the extracellular matrix, and therefore, they play a key role in the cell–matrix adhesive interactions. A variety of these adhesion structures have been identified: focal contacts (also known as focal adhesions), focal complexes, fibrillar adhesions, hemidesmosomes, podosomes, and invadopodia [5].
4.2 Cell–Extracellular Matrix Adhesion Structures
45
4.2.1 Focal Contacts (Focal Adhesions) The main specific adhesion structures are focal contacts (focal adhesions). These structures are discovered in all types of cultured tissue cells. The focal contacts are dynamic molecular complexes that are formed at small discrete sites of the basal cell surface [6]. The focal contacts provide tight attachment of the cells to the extracellular matrix. The focal adhesions are joined with the ends of actin filament bundles of the cytoskeleton (Fig. 4.13a–c). These tension-bearing filament bundles are also called stress fibers. They are formed at the edge of a cell in the zone of lamellipodia formation (Fig. 4.13c) and at the distal ends of filopodia. The focal contacts have two stages of their maturation: focal complexes, which are round spots less than 1–2 mm in diameter (see Sect. 4.2.2), and mature focal contacts. Focal complexes usually disappear soon after formation or are transformed to the mature focal contacts, which are elongated streaks, 2–8 mm in length. The evolution from the focal complexes to the mature focal contacts is associated with the stretch developed by stress fibers (see further).
Focal Adhesion Components The focal contacts consist of two components: the transmembrane component made by clusters of integrin molecules bound to the extracellular matrix, and the submembrane component made by a large complex of specific cytoplasmic proteins connected to actin filaments of the cytoskeleton (Fig. 4.14) [7]. Integrins are large dimeric protein complexes made up of two different types of polypeptide chains termed the a and b subunits that come together to form a heterodimer. Each subunit is a transmembrane glycoprotein that has a relatively large extracellular domain and a short cytoplasmic domain. There are 19 a and 8 b subunits that combine to form at least 24 different heterodimeric integrin receptors [8]. The integrins can be endocytosed from plasma membrane into the cytoplasm; some of them are later exocytosed back to the cell surface. This cellular process is called integrin trafficking [9–11]. Each integrin can bind to specific protein components of the extracellular matrix: collagens, fibronectin, laminins, etc. Integrin receptors bind to those protein matrix components that contain the tripeptide Arginine-Glycine-Aspartic acid (called RGD-sequence); it is the recognition sequence for binding of many integrins to extracellular matrix proteins. Some members of the integrin family recognize the RGD sequence in the native matrix molecules; a number of matrix proteins, such as collagen and certain laminin isoforms, expose their RGD sequences upon proteolytic processing of these proteins by matrix metalloproteinases (MMPs) (see Sect. 4.2.5) [12, 13]. Different combinations of a и b subunits determine the specificity of the integrin binding to an extracellular matrix protein. Some integrins can bind to only with one
46
4 Pseudopodia and Adhesion Structures
Fig. 4.13 (a–c) Fibroblastic rat cells (a, c), human fibroblastic cell (b). Mature focal contacts adjoined the ends of actin microfilaments. The focal contacts stained for paxillin (a, b) or vinculin (c) (red) and actin microfilaments stained for actin (green). FM. Scale bars, 28.5 mm (a), 15 mm (c). Confocal laser scanning microscopy. Scale bar, 12.5 mm (b). Courtesy of V.B. Dugina (b), reproduced with permission from The International Journal of Developmental Biology (ref. [40]) (c)
Fig. 4.14 Diagram of a focal contact (see the text for explanation)
4.2 Cell–Extracellular Matrix Adhesion Structures
47
matrix protein (e.g., a5b1 integrin binds only to fibronectin, and a6b1 integrin binds only to laminin); however, the majority of integrins are capable of binding to a few matrix proteins. On the other hand, various integrins can bind to the same matrix protein (e.g., a few different integrins can bind to laminin) [12, 13]. Some integrin receptors can also mediate the intercellular binding. Integrins are cell surface receptors, which mechanically connect tissue cells with the corresponding ligands, the extracellular matrix proteins. During their movement in the plane of the cell membrane, integrin receptors form clusters (accumulations). The integrin clustering and the conformational changes in the receptors affecting their affinity (see below) determine the stability of focal contacts. The integrin extracellular domains specifically bind to the matrix proteins, whereas the integrin cytoplasmic domains are associated with the large submembrane protein complex that consists of many different proteins: talin, vinculin, focal adhesion protein kinase (FAK), a-actinin, tensin, paxillin, and some others (Fig. 4.14) [7, 14, 15]. This protein complex is bound to the ends of actin filaments (Figs. 4.13a–c and 4.14). Some proteins of the complex are directly linked to the cytoplasmic domain of the b integrin subunit (FAK, paxillin, talin, and a-actinin) or to the filaments (tensin, vinculin, talin, and a-actinin); others are bridges between the cytoplasmic domains and the filaments (talin or a-actinin) (Fig. 4.14). Thus, the integrin receptors, which are bound to the matrix proteins, become anchored from within the cell by actin filaments. This ensures the structural connection between the extracellular matrix and the actin cytoskeleton of the attached cells.
Dynamic Properties and Regulation The ability of tissue cells to dynamically regulate their adhesion to the extracellular matrix is pivotal in multicellular organism. Dynamic regulation of focal contacts is the basis for the maintenance of asymmetric polarized cell morphology (cell polarization) and for cell migration. Various mechanisms are involved in the regulation of integrin-mediated cell– matrix adhesion. (a) The important regulatory mechanism is the integrin activation. It is the controlled increase in the affinity (ligand-binding activity) of integrin receptors for their protein ligands in the extracellular matrix. The integrin activation is caused by “inside-out” intracellular signals that through their action on integrin cytoplasmic domains induce rapid and reversible conformational changes in the integrin extracellular domains increasing their affinity [16–18]. In the integrin activation, the binding of talin and also cytoplasmic proteins termed kindlins to integrin b cytoplasmic domain plays a central role. There is a salt “bridge” between the cell membrane proximal regions of a and b subunit cytoplasmic domains. This “bridge” holds an integrin in an inactive state. One of the small GTPases,
48
4 Pseudopodia and Adhesion Structures
Rap1 protein, being activated, promotes the binding of talin to integrin b cytoplasmic domain. This binding disrupts the salt “bridge” and induces the conformation changes in the integrin extracellular domains increasing their affinity [16, 19]. Kindlins cooperate with talin, and therefore, these proteins play a crucial role in the integrin activation [18, 20–23]. Phosphorylation of integrin b subunit cytoplasmic domain by submembranous Src tyrosine kinase may prevent integrin–talin interactions and inhibit integrin activation. The activation of one integrin receptor can inhibit the activation of other integrins. This cross-talk among different integrin receptors of the same cell is termed transdominant inhibition of integrin activation. The basis of the trans-dominant inhibition is the competition for talin. The b subunit cytoplasmic domain of one integrin competes with the b domain of another integrin for talin. As a result, the ligand binding to one integrin can inhibit the activation of a second integrin. The transdominant inhibition of integrin activation contributes to a number of cellular processes, such as leukocyte transmigration, platelet adhesion, and angiogenesis. Integrin activation can be also caused by the “outside-in” signaling pathway from intercellular contacts. Cells have the transmembrane receptors termed the Notch receptors. Their ligands are also transmembrane proteins, and the receptors are only activated by direct cell-to-cell contacts. The Notch receptor activation results in proteolytic liberation of its intracellular domain, which is translocated into the cell nucleus and effect the expression of many genes including those controlling cell proliferation, survival, and migration. The intercellular contact-dependent Notch receptor activation increases the integrin affinity for the extracellular matrix proteins, such as fibronectin, collagens I and IV, and vitronectin [24]. (b) The integrin-linked kinase (ILK), which is a cytoplasmic serine/threonine kinase, is recruited to the sites of focal adhesion formation, where it interacts with two proteins, CH-ILKBP (abbreviated as parvin) and the PINCH (particularly interesting new cystein-histidine rich) protein. The formation of the ILKparvin-PINCH protein complex precedes the formation of focal contacts and is involved in the control of their assembly. This protein complex plays an important role in the regulation of focal adhesion dynamics [25–27]. (c) Critical contribution to the regulation of integrin-mediated cell adhesion is made by ADAM proteins (they are also being referred to as metalloproteinase-like disintegrin-like cysteine rich or MDC proteins). ADAMs are transmembrane proteins having a disintegrin domain and a metalloproteinase domain. Disintegrin domain of ADAMs can bind to integrin receptor and thus competitively inhibits normal integrin–ligand interaction blocking adhesive functions. Metalloproteinase domain can mediate antiadhesive functions by proteolytic cleavage of extracellular portions of transmembrane adhesion molecules [28, 29]. (d) Another important factor that contributes to dynamic regulation of cell–matrix adhesion is the intracellular calcium-dependent proteinase, calpain. This protein induces the focal contact disassembly by proteolysis of talin. The endogenous inhibitor of calpain, calpastatin, protects focal adhesions from their disassembly [30].
4.2 Cell–Extracellular Matrix Adhesion Structures
49
(e) Cell contractility is a key factor involved in the dynamic regulation of integrin-mediated cell–matrix adhesion. Since the actin filaments anchoring the integrins possess myosin II-driven contractility, the filaments are under the action of centripetal (directed toward a center) stretch forces. These forces organize the anchored actin filaments into the linear tension-bearing bundles (stress fibers), which leads to the increase in the stretch force. There is a hypothesis that focal contacts may act as sites of actin polymerization to generate stress fibers [31]. There are different sorts of stress fibers. Part of them named ventral stress fibers are anchored by both their ends to different focal contacts. Contraction of ventral stress fibers has static nature and critically contributes to the cell’s isometric tension. Their contraction causes lengthening (maturation) of the focal contacts (see below). The stress fibers of another category, dorsal stress fibers, are anchored by their front ends to focal contacts, while their back ends are intertwined with the cortical actin filament network. The dorsal stress fibers are more suited for cell motility (see Sect. 6.1.5) [32, 33]. The increase in the stretch force of stress fibers is the necessary condition for the maturation of the focal contacts and for the maintenance of their structure. The transition of initial weak and transient focal complexes to stronger and longerlived focal contacts is dependent on actomyosin-driven cell contractility. The significant weakening of the stretch (induced by the inhibition of the actin filament contractility) caused by protein caldesmon [34] stops the formation of new focal adhesions and causes the disassembly of the preexisting ones [35, 36]. The development of cell contractility and focal contact formation are controlled by the members of the family of small GTPases, Rho proteins (see Sect. 5.3.5) [37]. The transmission of the contractile forces strongly depends on vinculin, a focal contact-associated protein that is directly linked to actin filaments [38]. Thus, the individual focal contacts function as mechanical sensors: they increase their sizes proportional to the stretch forces applied to them, and they are disassembled when this stretch weakens. (f) Microtubules play an important role in cell contractility regulation: they are involved in the control of forces produced by the cell against focal contacts. Drug-induced depolymerization of microtubules in fibroblastic cells leads to a significant increase in cell contractility and promotes the formation of focal adhesions. This effect is mediated by Rho protein, a member of Rho family of GTPases (see Sect. 5.3.5). Rho is activated by the Rho guanine nucleotide exchange factor (GEF-H1) that is bound to microtubules and gets free while they depolymerize [39]. The responses to the increased cell contractility are the increase in the focal contact areas and significant elevation of the levels of the focal contact-associated proteins, including vinculin, paxillin, and FAK. Sometime later the tyrosine phosphorylation of the focal contact components is activated [35, 36, 40, 41]. Microtubules can also affect individual focal adhesions. When the stretch forces applied to increased focal contacts become excessive, these forces can detach the
50
4 Pseudopodia and Adhesion Structures
focal contacts from the extracellular matrix. In those cases, the cell “switches on” the special mechanism: the plus-ends of the growing microtubules move toward the extensively increased focal contacts and interact with them. This phenomenon is called the microtubule targeting. Direct interaction of microtubules with the focal contacts causes selective suppression of the myosin II-driven contractility at the distal ends of the stress fibers connected with these focal contacts. As a result, the growth of the focal contacts stops and they are frequently disassembled. Evidently, some signal molecules, which inhibit myosin II-driven contractility, are transported along the microtubules to locally suppress cell contractility, and thereby promote the focal contact disassembly [40, 42–45]. The microtubule targeting is induced by mDia1 (also called Dia1) protein that belongs to the formin protein family. The proteins of this family catalyze actin nucleation and polymerization. Under the effect of mDia1, the plus-ends of microtubules are concentrated near growing focal contacts. Microtubule-induced focal contact disassembly is mediated by FAK and a large GTPase protein called dynamin that is responsible for endocytosis. Dynamin interacts with FAK and colocalizes with the focal contacts during their microtubule targeting-induced disassembly. Possibly, microtubules may concentrate dynamin at focal adhesion sites [46]. Thus, the microtubule targeting stimulates focal adhesion disassembly. Focal contacts, in turn, can affect the dynamic instability of microtubules. The frequency of the transitions from the growth of microtubules to their shrinkage (catastrophe frequency) is much higher at focal adhesion sites than elsewhere. Thus, focal contacts can induce microtubule catastrophes. It is supposed that paxillin, a focal adhesion protein, which exhibits affinity for the microtubules, is involved in the catastrophe induction [47]. Therefore, the dynamic regulation of focal adhesions includes both integrin affinity-dependent mechanisms (integrin activation) and integrin affinity-independent ones including the actions of the intracellular proteinase calpain and its inhibitor calpastatin, the action of transmembrane proteins ADAMs, activity of the cytoplasmic ILK, cell contractility, and microtubule targeting. It is extremely important that the connection between the spread cell and the extracellular matrix is not only structural but also functional: the binding of the integrin to its matrix ligand triggers the “outside-in” signal transduction pathways that reach the cell nucleus and induce the expression of the specific genes. These signaling pathways will be described in detail below (see Sect. 5.3).
4.2.2 Focal Complexes Focal complexes are initial dot-like integrin-mediated cell–extracellular matrix adhesions. They are relatively small and transient. Focal complexes are formed underneath the lamellipodia and filopodia in motile cells. Some of focal complexes then turn into longer-lived elongated focal contacts (focal adhesions) associated with actin filament bundles. The formation of focal complexes is independent on
4.2 Cell–Extracellular Matrix Adhesion Structures
51
actomyosin-mediated cell contractility, but their transition into the matured focal contacts is blocked by the inhibition of myosin II. However, even in the conditions when the cell contractility is inhibited, external mechanical tension force applied to focal complexes at the cell edge induces their conversion into matured focal contacts. Therefore, focal complexes play a role of mechanosensors responding to an external force [37, 48].
4.2.3 Fibrillar Adhesions These transient integrin-mediated adhesions arise from medial ends of matured focal contacts but structurally distinct from them. Fibrillar adhesions contain a5b1 fibronectin-binding integrin; they are enriched in tensin but contain very low levels of paxillin, vinculin, and tyrosine-phosphorylated proteins. Fibrillar adhesions are associated with fibronectin fibrils in the extracellular matrix. These adhesions are independent of actomyosin-mediated cell contractility. Low degree of rigidity of the matrix favors the formation of fibrillar adhesions, but not focal contacts, by fibroblastic cells [49, 50].
4.2.4 Hemidesmosomes Hemidesmosomes are very small cell–extracellular matrix adhesion structures, which attach the stratified epithelial cells (such as in skin) to the basement membrane. In a hemidesmosome, a6b4 integrin is bound to the cellular keratin intermediate filaments via the submembrane protein plaque, which includes the proteins plectin and the bullous pemphigoid antigen 1 (BPAG1, also called BP230) [51]. Plectin and BPAG1 are members of the plakin family of proteins acting as cytolinkers to connect intermediate filaments to other cytoskeletal networks and/or adhesion complexes at the plasma membrane [52].
4.2.5 Podosomes and Invadopodia Some types of normal cells, such as osteoblasts, endothelial and epithelial cells, smooth muscle cells, macrophages, myeloid cells, and also invasive cancer cells, can locally degrade the extracellular matrix. During development and in disease states, such as cancer, the cells create increased spaces in the matrix, including penetration of basement membranes favoring cell migration [53]. The local matrix degradation is achieved by means of the specialized shortlived and highly dynamic cell adhesion structures termed podosomes (in normal cells) and invadopodia (in cancer cells), though podosomes are more involved in
52
4 Pseudopodia and Adhesion Structures
cell–matrix adhesion than invadopodia. The matrix degradation appears to be caused by metalloproteinase activity associated with these adhesive structures. The focal extracellular matrix degradation favors cell migration and cancer invasion [5, 54–61]. Podosomes are similar in their composition and structure to invadopodia. They are small cylindrical actin-rich formations. Their molecular components include integrins, actin, vinculin, paxillin, talin, actin-binding and actin-assembling proteins, signaling proteins, and also transmembrane metalloproteinases. Though podosomes/invadopodia share with focal adhesions some proteins (e.g., integrins, vinculin, talin, and paxillin), these adhesion formations clearly differ from focal contacts or focal complexes in the structural organization. Instead of cluster of integrin receptors linked to actin filament bundles by a submembrane protein complex consisting of structural and signaling proteins, podosomes/invadopodia are formed of a domain of integrins and associated proteins circularly surrounding a dense core consisting of actin filaments. This actin core is oriented perpendicularly to the plasma membrane and to the extracellular matrix unlike tangential orientation of actin stress fibers anchored to focal adhesions. Both podosomes and invadopodia are mechanosensitive structures like focal adhesions [62, 63]. In the formation and regulation of podosomes/invadopodia, the actin cytoskeleton regulating proteins are involved. Polymerization of actin filaments and myosin II-driven cell contractility are critical to podosomes/invadopodia formation and functioning. Actin-binding protein cortactin that regulates the Arp2/3 complex is the important component of podosomes/invadopodia [64]. Cortactin depletion inhibits the formation of these adhesion structures. Overexpression of cortactin found in a number of human carcinomas is proposed to contribute to the malignant tumor cell invasion [65, 66]. Depletion of another podosomes/invadopodia protein component, caldesmon, which is a negative regulator of cell contractility, facilitates formation of podosomes/invadopodia and favors invasion of cancer cells [34, 67]. The enzymes that confer proteolytic activity to podosomes and invadopodia are transmembrane matrix metalloproteinases (MT-MMPs), zinc-dependent hydrolytic enzymes. They degrade extracellular matrix components, such as collagens, laminins, and fibronectin, and irreversibly remodel basement membranes creating increased spaces for the cell migration [53]. Besides, MMP-mediated matrix proteolysis can release the matrix-bound growth factors (e.g., fibroblast growth factor, FGF-2), produce specific biologically active molecules, directly regulate epithelial tissue architecture through cleavage of the basement membrane components, and participate in vascular remodeling [68–71]. MMPs are inhibited by their specific natural inhibitors, tissue inhibitors of metalloproteinases (TIMPs) found in most tissues [72]. The dynamic balance between MMPs and their TIMPs plays an important role in the epithelial cell migration to close wounds, in inflammation and angiogenesis. The disturbance of the balance is involved in the development of different human diseases, malignant tumor invasion and metastasis [71, 72].
References
53
References 1. Verkhovsky AB, Chaga OY, Schaub S, Svitkina TM, Meister JJ, Borisy GG (2003) Orientational order of the lamellipodial actin network as demonstrated in living motile cells. Mol Biol Cell 14(11):4667–4675. doi:10.1091/mbc.E02-10-0630 DOI:dx.doi.org 2. Faix J, Rottner K (2006) The making of filopodia. Curr Opin Cell Biol 18(1):18–25. doi:10.1016/j.ceb.2005.11.002 DOI:dx.doi.org 3. Fackler OT, Grosse R (2008) Cell motility through plasma membrane blebbing. J Cell Biol 181(6):879–884. doi:10.1083/jcb.200802081 DOI:dx.doi.org 4. Charras G, Paluch E (2008) Blebs lead the way: how to migrate without lamellipodia. Nat Rev Mol Cell Biol 9(9):730–736. doi:10.1038/nrm2453 DOI:dx.doi.org 5. Block MR, Badowski C, Millon-Fremillon A, Bouvard D, Bouin AP, Faurobert E, GerberScokaert D, Planus E, Albiges-Rizo C (2008) Podosome-type adhesions and focal adhesions, so alike yet so different. Eur J Cell Biol 87(8-9):491–506. doi:10.1016/j.ejcb.2008.02.012 DOI:dx.doi.org 6. Lo SH (2006) Focal adhesions: what’s new inside. Dev Biol 294(2):280–291. doi:10.1016/j. ydbio.2006.03.029 DOI:dx.doi.org 7. Delon I, Brown NH (2007) Integrins and the actin cytoskeleton. Curr Opin Cell Biol 19(1):43–50. doi:10.1016/j.ceb.2006.12.013 DOI:dx.doi.org 8. Humphries MJ (2000) Integrin structure. Biochem Soc Trans 28(4):311–339. doi:10.1042/03005127:0280311 DOI:dx.doi.org 9. Caswell P, Norman J (2008) Endocytic transport of integrins during cell migration and invasion. Trends Cell Biol 18(6):257–263. doi:10.1016/j.tcb.2008.03.004 DOI:dx.doi.org 10. Caswell PT, Vadrevu S, Norman JC (2009) Integrins: masters and slaves of endocytic transport. Nat Rev Mol Cell Biol 10(12):843–853. doi:10.1038/nrm2799 DOI:dx.doi.org 11. Ulrich F, Heisenberg CP (2009) Trafficking and cell migration. Traffic 10(7):811–818. doi:10.1111/j.1600-0854.2009.00929.x DOI:dx.doi.org 12. Heino J, Käpylä J (2009) Cellular receptors of extracellular matrix molecules. Curr Pharm Des 15(12):1309–1317. doi:10.2174/138161209787846720 DOI:dx.doi.org 13. Barczyk M, Carracedo S, Gullberg D (2010) Integrins. Cell Tissue Res 339(1):269–280. doi:10.1007/s00441-009-0834-6 DOI:dx.doi.org 14. Arnaout MA, Goodman SL, Xiong JP (2007) Structure and mechanics of integrin-based cell adhesion. Curr Opin Cell Biol 19(5):495–507. doi:10.1016/j.ceb.2007.08.002 DOI:dx.doi.org 15. Legate KR, Fässler R (2009) Mechanisms that regulate adaptor binding to beta-integrin cytoplasmic tails. J Cell Sci 122(Pt 2):187–198 16. Wegener KL, Campbell ID (2008) Transmembrane and cytoplasmic domains in integrin activation and protein-protein interactions. Mol Membr Biol 25(5):376–387. doi:10.1080/ 09687680802269886 DOI:dx.doi.org 17. Shattil SJ, Kim C, Ginsberg MH (2010) The final steps of integrin activation: the end game. Nat Rev Mol Cell Biol 11(4):288–300. doi:10.1038/nrm2871 DOI:dx.doi.org 18. Wang W, Luo BH (2010) Structural basis of integrin transmembrane activation. J Cell Biochem 109(3):447–452 19. Banno A, Ginsberg MH (2008) Integrin activation. Biochim Soc Trans 36 (Pt2): 229–234. doi:10.1042/BST0360229 DOI:dx.doi.org 20. Larjava H, Plow EF, Wu C (2008) Kindlins: essential regulators of integrin signalling and cellmatrix adhesion. EMBO Rep 9(12):1203–1208. doi:10.1038/embor.2008.202 DOI:dx.doi.org 21. Critchley DR (2009) Biochemical and structural properties of the integrin-associated cytoskeletal protein talin. Annu Rev Biophys 38:235–254. doi:10.1146/annurev.biophys.050708.133744 DOI:dx.doi.org 22. Meves A, Stremmel C, Gottschalk K, Fässler R (2009) The Kindlin protein family: new members to the club of focal adhesion proteins. Trends Cell Biol 19(10):504–513. doi:10.1016/j. tcb.2009.07.006 DOI:dx.doi.org
54
4 Pseudopodia and Adhesion Structures
23. Moser M, Legate KR, Zent R, Fässler R (2009) The tail of integrins, talin, and kindlins. Science. 324(5929):895–899. doi:10.1126/science.1163865 DOI:dx.doi.org 24. Karsan A (2008) Notch and integrin affinity: a sticky situation. Sci Signal 1(2):pe2. doi:10.1126/ stke.12pe2 DOI:dx.doi.org 25. Boulter E, Van Obberghen-Schilling E (2006) Integrin-linked kinase and its partners: a modular platform regulating cell-matrix adhesion dynamics and cytoskeletal organization. Eur J Cell Biol 85(3-4):255–263. doi:10.1016/j.ejcb.2005.09.002 DOI:dx.doi.org 26. Legate KR, Montañez E, Kudlacek O, Fässler R (2006) ILK, PINCH and parvin: the tIPP of integrin signalling. Nat Rev Mol Cell Biol 7(1):20–31 27. Hehlgans S, Haase M, Cordes N (2007) Signalling via integrins: implications for cell survival and anticancer strategies. Biochim Biophys Acta 1775(1):163–180. doi:10.1016/j. bbcan.2006.09.001 DOI:dx.doi.org 28. Reiss K, Ludwig A, Saftig P (2006) Breaking up the tie: disintegrin-like metalloproteinases as regulators of cell migration in inflammation and invasion. Pharmacol Ther 111(3):985–1006. doi:10.1016/j.pharmthera.2006.02.009 DOI:dx.doi.org 29. Reiss K, Saftig P (2009) The “a disintegrin and metalloprotease” (ADAM) family of sheddases: physiological and cellular functions. Semin Cell Dev Biol 20(2):126–137. doi:10.1016/j. semcdb.2008.11.002 DOI:dx.doi.org 30. Franco SJ, Rodgers MA, Perrin BJ, Han J, Bennin DA, Critchley DR, Huttenlocher A (2004) Calpain-mediated proteolysis of talin regulates adhesion dynamic. Nat Cell Biol 6(10):977– 983. doi:10.1038/ncb1175 DOI:dx.doi.org 31. O’Neill GM (2009) The coordination between actin filaments and adhesion in mesenchymal migration. Cell Adh Migr 3(4):355–357. doi:10.4161/cam.3.4.9468 DOI:dx.doi.org 32. Pellegrin S, Mellor H (2007) Actin stress fibres. J Cell Sci 120(Pt 20):3491–3499. doi:10.1242/ jcs.018473 DOI:dx.doi.org 33. Naumanen P, Lappalainen P, Hotulainen P (2008) Mechanisms of actin stress fibre assembly. J Microsc 231(3):446–454. doi:10.1111/j.1365-2818.2008.02057.x DOI:dx.doi.org 34. Lin JJ, Li Y, Eppinga RD, Wang Q, Jin JP (2009) Chapter 1: roles of caldesmon in cell motility and actin cytoskeleton remodeling. Int Rev Cell Mol Biol 274:1–68. doi:10.1016/S19376448(08)02001–7 DOI:dx.doi.org 35. Elbaum M, Chausovsky A, Levy ET, Shtutman M, Bershadsky AD (1999) Microtubule involvement in regulating cell contractility and adhesion-dependent signalling: a possible mechanism for polarization of cell motility. Biochem Soc Symp 65:147–72 36. Grosheva I, Vittitow JL, Goichberg P, Gabelt BT, Kaufman PL, Borrás T, Geiger B, Bershadsky AD (2006) Caldesmon effects on the actin cytoskeleton and cell adhesion in cultured HTM cells. Exp Eye Res 82(6):945–58. doi:10.1016/j.exer.2006.01.006 DOI:dx.doi.org 37. Bershadsky AD, Ballestrem C, Carramusa L, Zilberman Y, Gilquin B, Khochbin S, Alexandrova AY, Verkhovsky AB, Shemesh T, Kozlov MM (2006) Assembly and mechanosensory function of focal adhesions: experiments and models. Eur J Cell Biol 85(3-4):165–173. doi:10.1016/j. ejcb.2005.11.001 DOI:dx.doi.org 38. Mierke CT (2009) The role of vinculin in the regulation of the mechanical properties of cells. Cell Biochem Biophys 53(3):115–126. doi:10.1007/s12013–009-9047-6 DOI:dx.doi.org 39. Krendel M, Zenke FT, Bokoch GM (2002) Nucleotide exchange factor GEF-H1 mediates cross-talk between microtubules and the actin cytoskeleton. Nat Cell Biol 4(4):294–301. doi:10.1038/ncb773 DOI:dx.doi.org 40. Vasiliev JM (2004) Cytoskeletal mechanisms responsible for invasive migration of neoplastic cells. Int J Dev Biol 48(5-6):425–439. doi:10.1387/ijdb.041806jv DOI:dx.doi.org 41. Kirchner J, Kam Z, Tzur G, Bershadsky AD, Geiger B (2003) Live-cell monitoring of tyrosine phosphorylation in focal adhesions following microtubule disruption. J Cell Sci 116(Pt 6):975–986. doi:10.1242/jcs.00284 DOI:dx.doi.org 42. Kaverina I, Krylyshkina O, Small JV (1999) Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J Cell Biol 146(5):1033–1044. doi:10.1083/ jcb.146.5.1033 DOI:dx.doi.org
References
55
43. Small JV, Geiger B, Kaverina I, Bershadsky A (2002) How do microtubules guide migrating cells? Nat Rev Mol Cell Biol 3(12):957–964. doi:10.1038/nrm971 DOI:dx.doi.org 44. Krylyshkina O, Anderson KI, Kaverina I, Upmann I, Manstein DJ, Small JV, Toomre DK (2003) Nanometer targeting of microtubules to focal adhesions. J Cell Biol 161(5):853–859. doi:10.1083/jcb.200301102 DOI:dx.doi.org 45. Small JV, Kaverina I (2003) Microtubules meet substrate adhesions to arrange cell polarity. Curr Opin Cell Biol 15(1):40–47. doi:10.1016/S0955-0674(02)00008-X DOI:dx.doi.org 46. Ezratty EJ, Partridge MA, Gundersen GG (2005) Microtubule-induced focal adhesion disassembly is mediated by dynamin and focal adhesion kinase. Nat Cell Biol 7(6):581–590. doi:10.1038/ncb1262 DOI:dx.doi.org 47. Efimov A, Kaverina I (2009) Significance of microtubule catastrophes at focal adhesion sites. Cell Adh Migr 3(3):285–287. doi:10.4161/cam.3.3.8858 DOI:dx.doi.org 48. Kaverina I, Krylyshkina O, Small JV (2002) Regulation of substrate adhesion dynamics during cell motility. Int J Biochem Cell Biol 34(7):746–761. doi:10.1016/S1357-2725(01)00171–6 DOI:dx.doi.org 49. Katz BZ, Zamir E, Bershadsky A, Kam Z, Yamada KM, Geiger B (2000) Physical state of the extracellular matrix regulates the structure and molecular composition of cell-matrix adhesions. Mol Biol Cell 11(3):1047–1060 50. Zamir E, Katz M, Posen Y, Erez N, Yamada KM, Katz BZ, Lin S, Lin DC, Bershadsky A, Kam Z, Geiger B (2000) Dynamics and segregation of cell-matrix adhesions in cultured fibroblasts. Nat Cell Biol 2(4):191–196. doi:10.1038/35008607 DOI:dx.doi.org 51. de Pereda JM, Ortega E, Alonso-García N, Gómez-Hernández M, Sonnenberg A (2009) Advances and perspectives of the architecture of hemidesmosomes: lessons from structural biology. Cell Adh Migr 3(4):361–364. doi:10.4161/cam.3.4.9525 DOI:dx.doi.org 52. Svitkina TM, Verkhovsky AB, Borisy GG (1996) Plectin sidearms mediate interaction of intermediate filaments with microtubules and other components of the cytoskeleton. J Cell Biol 135(4):991–1007. doi:10.1083/jcb.135.4.991 DOI:dx.doi.org 53. Rowe RG, Weiss SJ (2008) Breaching the basement membrane: who, when and how? Trends Cell Biol 18(11):560–574. doi:10.1016/j.tcb.2008.08.007 DOI:dx.doi.org 54. Gimona M (2008) The microfilament system in the formation of invasive adhesions. Semin Cancer Biol 18(1):23–34. doi:10.1016/j.semcancer.2007.08.005 DOI:dx.doi.org 55. Gimona M, Buccione R, Courtneidge SA, Linder S (2008) Assembly and biological role of podosomes and invadopodia. Curr Opin Cell Biol 20(2):235–241. doi:10.1016/j.ceb.2008.01.005 DOI:dx.doi.org 56. Weaver AM (2006) Invadopodia: specialized cell structures for cancer invasion. Clin Exp Metastasis 23(2):97–105. doi:10.1007/s10585-006-9014-1 DOI:dx.doi.org 57. Yamaguchi H, Pixley F, Condeelis J (2006) Invadopodia and podosomes in tumor invasion. Eur J Cell Biol 85(3-4):213–218. doi:10.1016/j.ejcb.2005.10.004 DOI:dx.doi.org 58. Gimona M, Buccione R (2006) Adhesions that mediate invasion. Int J Biochem Cell Biol 38(11):1875–1892. doi:10.1016/j.biocel.2006.05.003 DOI:dx.doi.org 59. Stylli SS, Kaye AH, Lock P (2008) Invadopodia: at the cutting edge of tumour invasion. J Clin Neurosci 15(7):725–737. doi:10.1016/j.jocn.2008.03.003 DOI:dx.doi.org 60. Friedl P, Wolf K (2009) Proteolytic interstitial cell migration: a five-step process. Cancer Metastasis Rev 28(1-2):129–135. doi:10.1007/s10555-008-9174-3 DOI:dx.doi.org 61. Buccione R, Caldieri G, Ayala I (2009) Invadopodia: specialized tumor cell structures for the focal degradation of the extracellular matrix. Cancer Metastasis Rev 28(1-2):137–149. doi:10.1007/s10555-008-9176-1 DOI:dx.doi.org 62. Albiges-Rizo C, Destaing O, Fourcade B, Planus E, Block MR (2009) Actin machinery and mechanosensitivity in invadopodia, podosomes and focal adhesions. J Cell Sci. Sep 1;122 (Pt 17):3037–3049. doi:10.1242/jcs.052704 DOI:dx.doi.org 63. Caldieri G, Ayala I, Attanasio F, Buccione R (2009) Cell and molecular biology of invadopodia. Int Rev Cell Mol Biol 275:1–34. doi:10.1016/S1937-6448(09)75001-4 DOI:dx.doi.org
56
4 Pseudopodia and Adhesion Structures
64. Clark ES, Weaver AM (2008) A new role for cortactin in invadopodia: Regulation of protease secretion. Eur J Cell Biol 87(8-9):581–590. doi:10.1016/j.ejcb.2008.01.008 DOI:dx.doi.org 65. Buday L, Downward J (2007) Roles of cortactin in tumor pathogenesis. Biochim Biophys Acta 1775(2):263–273. doi:10.1016/j.bbcan.2006.12.002 DOI:dx.doi.org 66. Ammer AG, Weed SA (2008) Cortactin branches out: roles in regulating protrusive actin dynamics. Cell Motil Cytoskeleton 65(9):687–707. doi:10.1002/cm.20296 DOI:dx.doi.org 67. Yoshio T, Morita T, Kimura Y, Tsujii M, Hayashi N, Sobue K (2007) Caldesmon suppresses cancer cell invasion by regulating podosome/invadopodium formation. FEBS Lett 581(20):3777–3782. doi:10.1016/j.febslet.2007.06.073 DOI:dx.doi.org 68. Chen P, Parks WC (2009) Role of matrix metalloproteinases in epithelial migration. J Cell Biochem 108(6):1233–1243. doi:10.1002/jcb.22363 DOI:dx.doi.org 69. Kessenbrock K, Plaks V, Werb Z (2010) Matrix metalloproteinases: regulators of the tumor microenvironment. Cell .141(1):52–67. doi:10.1016/j.cell.2010.03.015 DOI:dx.doi.org 70. Rodríguez D, Morrison CJ, Overall CM (2010) Matrix metalloproteinases: what do they not do? New substrates and biological roles identified by murine models and proteomics. Biochim Biophys Acta 1803(1):39–54. doi:10.1016/j.bbamcr.2009.09.015 DOI:dx.doi.org 71. Fanjul-Fernández M, Folgueras AR, Cabrera S, López-Otín C (2010) Matrix metalloproteinases: evolution, gene regulation and functional analysis in mouse models. Biochim Biophys Acta 1803(1):3–19. doi:10.1016/j.bbamcr.2009.07.004 DOI:dx.doi.org 72. Brew K, Nagase H (2010) The tissue inhibitors of metalloproteinases (TIMPs): an ancient family with structural and functional diversity. Biochim Biophys Acta 1803(1):55–71. doi:10.1016/j. bbamcr.2010.01.003 DOI:dx.doi.org
Chapter 5
Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Abstract Adhesive interactions of cells with the extracellular matrix start soon after they settle on the matrix from suspended state. The matrix-attached spherical cells begin to spread and gradually reach high degree of flattening. The cell spreading process includes defined consecutive changes both in the cell shape and in the cell surface relief. Oncogenic transformation results in a deficient cell spreading. Adhesive interactions include intracellular signal transduction pathways in the cells. The signaling pathways are triggered by the extracellular stimulatory molecules (ligands) following their binding to different types of specialized cell surface receptors. In particular, integrin receptors, which are components of focal adhesions and play a key role in cell-matrix attachment, also function as signal transducers. Different integrin-mediated and growth factor receptor-mediated signaling pathways determine and control cell morphology, proliferation, survival, and migration. The oncogenic transformation results in the weakening of integrin-mediated cellmatrix adhesion and induces serious alterations in the integrin-mediated and growth factor receptor-mediated signaling pathways. The consequences of these alterations are the “anchorage independence” (substratum independence of cell proliferation), permanent mitogenic activation, loss of cell detachment-induced apoptosis (anoikis), and high migratory activity of transformed cells.
5.1 Cell Spreading on the Extracellular Matrix Surface 5.1.1 Cells in a Suspended State Being isolated from the extracellular matrix, normal or transformed tissue cells convert to the suspended state. They are in that state until they settle on the matrix surface and enter into contact interactions with it. In a suspended state, tissue cells acquire a spherical shape. The surfaces of these spherical cells have numerous protrusions of four morphological types: microvilli,
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_5, © Springer Science+Business Media, LLC 2011
57
58
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.1 Transformed mouse fibroblast in a suspended state. Microvillous cell surface relief. SEM. Scale bar, 4 mm
blebs, folds, and ruffles. These are all cytoplasmic protrusions bordered with the cell membrane. (a) Microvilli are cylindrical elongated protrusions about 0.1–0.2 mm in diameter; their lengths can vary considerably from 0.2–0.5 to 5–6 mm (Fig. 5.1), they can be straight or bent. (b) Blebs are spherical or hemispherical protrusions with smooth surfaces and with the diameters varying from less than 1 to 2–3 mm (Fig. 5.2). (c) Folds are protrusions of flattened shapes. Their thickness is usually 0.1–0.4 mm but their widths and lengths can vary considerably (Fig. 5.3). (d) Ruffles are flattened protrusions like folds. Ruffles can have shapes of big and deep folds; however, more often they have characteristic shapes of “frills” (Fig. 5.3). The whole surface of a suspended cell can have the protrusions of one type or several types; the surface relief patterns can be designated, respectively, as homogeneous or mixed. The combination of blebs and microvilli seems to be the most common among mixed cell surface relief patterns (Fig. 5.4). Blebbed, microvillous, and mixed patterns of the cell surface relief are present both in normal and in transformed suspended tissue cells. There are, however, considerable differences in the relative frequencies of the surface relief patterns between normal and transformed cell populations [1]. In suspensions of normal fibroblastic or epithelial cells, the blebbed surface relief pattern significantly prevails over the microvillous pattern. In suspensions of
Fig. 5.2 Fibroblastic mouse cell in a suspended state. Blebbed cell surface relief. SEM. Scale bar, 2.5 mm
Fig. 5.3 Ascites sarcoma mouse cell in a suspended state. Mixed cell surface relief: folds (double arrows) and ruffles (arrows). SEM. Scale bar, 2.8 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
60
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.4 Transformed mouse fibroblast in a suspended state. Mixed cell surface relief: blebs (arrow) and microvilli (double arrow). SEM. Scale bar, 3.5 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
transformed cells, the reverse ratio is typical: there is a very low relative abundance of the blebbed surface relief and high abundance of cells with the microvillous relief [1, 2]. Considerable decreases in the ratios of cells with the blebbed surface relief and increases in ratios of cells with the microvillous surface relief are typical for transformed cell populations in their suspended state. In populations of suspended transformed cells, one can see some variability of morphology and distribution of cell surface protrusions. Some microvilli can be branched and have bulbous ends (Fig. 5.5). Sometimes one can see discrete zones, “foci,” of the altered relief: atypically long microvilli (Fig. 5.6), large folds and (or) peculiar flattened protrusions like frills, or ruffles (Figs. 5.7 and 5.8). In suspended state the cells of some epithelial tumors form two-cellular or multicellular aggregates (Fig. 5.9a, b). The pattern of the surface relief of suspended cells depends on their “previous history.” More specifically, it depends on the degree of the cell spreading on the extracellular matrix before the detachment and the transition of the cells to their suspension state [1]. Well spread normal fibroblastic cells (Fig. 5.10) after their detachment from the matrix acquire mainly blebbed relief (Fig. 5.2). The same fibroblastic cells on the surface of the special adhesion-decreasing polymer [poly (2-hydroxyethyl) metacrylate] coating are much less spread (Fig. 5.11a, b).
5.1 Cell Spreading on the Extracellular Matrix Surface
61
Fig. 5.5 Transformed mouse fibroblast in a suspended state. Some microvilli are branched and have bulbous ends (arrow). SEM. Scale bar, 1.5 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
Fig. 5.6 Ascites sarcoma mouse cell in a suspended state. A “focus” of the altered cell surface relief: atypically long microvilli (arrow). SEM. Scale bar, 3 mm
62
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.7 Ascites epithelial tumor mouse cell in a suspended state. A “focus” of the altered cell surface relief: folds and long microvilli (arrow). SEM. Scale bar, 4.5 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
Fig. 5.8 Ascites epithelial tumor mouse cell in a suspended state. A “focus” of the altered cell surface relief: ruffles (arrow). SEM. Scale bar, 1.5 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
5.1 Cell Spreading on the Extracellular Matrix Surface
63
Fig. 5.9 (a, b) Ascites epithelial tumor rat cells in a suspended state: two-cellular aggregate (a), multicellular aggregate (b). SEM. Scale bars, 3.7 mm (a), 5.7 mm (b)
Fig. 5.10 Fibroblastic mouse cell in a well spread state. SEM. Scale bar, 10.5 mm. Reproduced with permission from the International Review of Cytology (ref. [1])
Being transitioned to the suspended state, this cell population has a lower percentage of the cells with blebbed relief and a higher percentage of the cells with microvillous one compared to the suspended fibroblastic cells, which were well spread before their detachment. It is possible that the relatively high ratios of microvillous surface relief of transformed cells in the suspended state are the result of the previous poor spreading of these cells before their detachment from extracellular matrix (see Sect. 5.1.3).
64
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.11 (a, b) Fibroblastic mouse cells on the adhesion-decreasing polymer surface. The cells are badly spread. SEM. Scale bars, 11.5 mm (a), 5 mm (b). Reproduced with permission from the International Review of Cytology (ref. [1])
5.1.2 The Morphology of Cell Spreading Process in Normal Cells The process of spreading of fibroblastic or epithelial cells begins several minutes after their settling on the extracellular matrix. For a while, the cells maintain their spherical shapes, and then they gradually get flattened. Finally, the cells acquire the shapes typical for their cellular types. The cell spreading is fulfilled by the pseudopodial activity and the formation of focal contacts connecting the pseudopodia with the extracellular matrix. The cell spreading process has two stages. (a) Stage of radial spreading. In the initial stage of cell spreading, filopodia appears at the base of a spherical cell. At first, few filopodia are formed and they are short (Fig. 5.12); then their number and their lengths can be gradually increased (see Fig. 4.3 in Chap. 4). The filopodia are radially extended from the base of the cell, being attached with their distal ends to the extracellular matrix by means of focal contacts. Instead of filopodia, short lamellipodia can appear at the cell base (Fig. 5.13). They are also radially extended and attached to the extracellular matrix. In other cases, lamellipodia are formed simultaneously with filopodia (Fig. 5.14) or they are formed a little bit late. The attached filopodia may act as “rails” moving the lamellipodia in the outward direction. With the development of lamellipodia, the spherical cell begins to flatten gradually [3]. Immediately after their appearance, filopodia or lamellipodia do not contain myosin II that diffuses into them a little later. The interaction of myosin II with the
5.1 Cell Spreading on the Extracellular Matrix Surface
65
Fig. 5.12 Fibroblastic mouse cell at the stage of radial spreading. Short filopodia (arrows) are at the base of the spherical cell. SEM. Scale bar, 3.3 mm
Fig. 5.13 Fibroblastic mouse cell at the stage of radial spreading. Lamellipodia (arrow) are at the base of the spherical cell. SEM. Scale bar, 2.8 mm
66
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.14 Fibroblastic hamster cell at the stage of radial spreading. There are lamellipodia (arrows) and filopodia (double arrow) are at the base of the spherical cell. SEM. Scale bar, 3.5 mm
actin of the filaments results in the retraction of pseudopodia. When pseudopodia do not have time to form the focal adhesions with the extracellular matrix they retract. The retraction of the attached pseudopodia generates their centripetal stretch. This stretch leads to the spreading and gradual flattening of the cell. The cell flattening becomes more obvious when lamellipodia at the cell base are fused into the thin circular plate (called lamellar cytoplasm) adjoining the extracellular matrix. It is concentrically extended around the central convex part of the cell (Fig. 5.15). On the edge of the lamellar cytoplasm, lamellipodia and/or ruffles are observed (Fig. 5.16). The cell is progressively flattened and looks like a thin disc, along the entire perimeter of which lamellipodia and ruffles are continuously formed (Fig. 5.17) [3]. The discoid fibroblastic cell has a circular actin filament bundle located along the entire cell edge (Fig. 5.18). The stage of radial spreading takes 20–30 min after the cell seeding. In epithelial cells after their settling on the extracellular matrix, the spreading begins with a certain delay. At the base of the spherical cell, filopodia do not appear, but lamellipodia are immediately formed. The lamellipodia are soon fused into the concentrically extended lamellar cytoplasm. The cell acquires the shape of thin disc (Fig. 5.19). (b) Polarization stage. The main difference between the spreading of fibroblastic and epithelial cells is related to the fact that in the epithelial cells the radial stage is the final one: completely spread epithelial cells maintain the disc-like shape with circular lamellar cytoplasm and pseudopodial formation along the entire cell edge (Fig. 5.19).
5.1 Cell Spreading on the Extracellular Matrix Surface
67
Fig. 5.15 Fibroblastic mouse cell at the stage of radial spreading. Lamellar cytoplasm (arrow) concentrically located around the semispherical part of the cell. SEM. Scale bar, 5.2 mm
Fig. 5.16 Fibroblastic mouse cell at the stage of radial spreading. There are ruffles (arrows) on the edge of the lamellar cytoplasm. SEM. Scale bar, 3 mm
68
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.17 Fibroblastic mouse cell in the termination of the radial spreading stage. The thin disclike cell. The entire cell periphery is the circular lamellar cytoplasm with lamellipodia (arrow) and ruffles (double arrows) along its edge. SEM. Scale bar, 9 mm
Fig. 5.18 Fibroblastic mouse cell in the termination of the radial spreading stage. Circular actin microfilament bundle. Staining for actin. FM. Scale bar, 24 mm
5.1 Cell Spreading on the Extracellular Matrix Surface
69
Fig. 5.19 Epithelial mouse cell in the termination of the radial spreading stage. The thin disc-like cell. The entire cell periphery is the circular lamellar cytoplasm (arrow) with pseudopodia (double arrows) along its edge. SEM. Scale bar, 13 mm
As to the fibroblastic cells, they move into the final stage of the spreading, the polarization stage. In this stage, fibroblastic cells acquire polarized shape: they become considerably flattened, polygonal, or more elongated (Fig. 5.20), the cells form one or several wide peripheral lamellas (lamellar cytoplasm), at the free edge of which the pseudopodia are formed (see Fig. 4.9 in Chap. 4). The transition of fibroblastic cells to the polarization stage occurs because of the redistribution of their pseudopodial activity: pseudopodia cease to appear evenly at the entire free cell edge; however, they continue being formed at the definite zones of the free edge. Because of this, the fibroblastic cells transition from disc-like shapes to polygonal or more elongated ones. Simultaneously, the circular actin filament bundle (Fig. 5.18) is disintegrated, and the linear actin filament bundles crossing the cell body are formed (Fig. 5.21). The pseudopodial activity redistribution is controlled not only by the actin cytoskeleton that participates in the formation, attachment, and stretch of the pseudopodia, but also by the system of microtubules. Both these cytoskeletal systems acting cooperatively “determine” the cell edge parts where pseudopodia are formed (see Sects. 6.1.1 and 6.1.2). The cooperated action of the cytoskeletal systems is controlled in turn by the proteins of one of the families of small GTPases, Rho-GTPases (see Sect. 5.3.5). In the process of the transition of fibroblastic cells from their suspended state to the spread one, the cell surface relief undergoes progressive smoothing: all protrusions (blebs, folds, microvilli or ruffles) gradually disappear (Figs. 5.14, 5.15, 5.17, and 5.20). In epithelial cells, the surface relief smoothing is less pronounced
70
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.20 Fibroblastic mouse cells at the polarization stage of the spreading. The cells are well spread, flattened, and have polygonal or elongated shapes. SEM. Scale bar, 26 mm
Fig. 5.21 Fibroblastic rat cell at the polarization stage of the spreading. Linear actin microfilaments and the microfilament bundles crossing the cell body. Staining for actin. FM. Scale bar, 16.5 mm. Courtesy of A.Y. Alexandrova
(Fig. 5.19). Evidently, the cell surface is “reserved” in the protrusions that compose the surface relief patterns of suspended cells. Later on, the reserved cell surface is progressively spent in the process of cell spreading.
5.1 Cell Spreading on the Extracellular Matrix Surface
71
Fig. 5.22 Fibroblastic mouse cells. The well spread and considerably flattened cells with well-developed thin lamellar cytoplasm (arrows) and smooth dorsal cell surfaces. SEM. Scale bar, 28 mm
The entire process of cell spreading usually takes several hours after the cell seeding. Completely spread fibroblastic cells are considerably flattened. The cell has a well-developed lamellar cytoplasm in the shapes of one or several peripheral wide thin lamellas (Fig. 5.22). Polarized fibroblastic cells, in contrast to spread epithelial cells, maintain their relatively constant average cell lengths. The mechanism of the cell length control is based on the dynamic balance of two forces acting in the opposite directions: centripetal tension of the actin-myosin filaments and centrifugal force generated by the growth of the microtubules in the fibroblastic cells [4]. A typical feature of completely spread fibroblastic cells is their morphological polarity manifested by division of the cell edge into active, pseudopodiaforming zones, and stable, nonactive zones. Pseudopodia are formed only at the free edge of the lamellar cytoplasm, and pseudopodia-forming cell edge is called the active one (Fig. 5.22). The fibroblastic cell elongates in the direction of its active edge. Irregularly distributed pseudopodial activity is the prerequisite for the directional movement of fibroblastic cells. The dorsal surfaces of completely spread fibroblastic cells are devoid of morphological formations and look smooth (Figs. 5.20 and 5.22). Fibroblastic cells contain linear tension-bearing actin filament bundles (stress fibers) associated with focal contacts (see Fig. 4.13a–c in Chap. 4). The stress fibers are numerous, cross the cell body and are oriented mostly along the cell axis (Fig. 5.21).
72
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.23 Epithelial mouse cell. The thin disc-like cell with the circular lamellar cytoplasm (arrow) at the entire cell periphery. Microvillous cell surface relief (short microvilli). SEM. Scale bar, 10.5 mm
Completely spread single epithelial cells have discoid or rather polygonal shapes (Fig. 5.23). The central part of the cell is slightly convex. The dorsal cell surfaces have a relief in the form of very short microvilli (Figs. 5.19 and 5.23). The microvilli are often less numerous at the peripheral zone that is circular lamellar cytoplasm (Figs. 5.23 and 5.24). Epithelial cells are not polarized: the entire cell perimeter keeps its pseudopodial activity (Figs. 5.19 and 5.24). The cell contains an actin filament bundle, which has a circular shape and is located along the entire cell periphery near the active edge (see Fig. 3.3 in Chap. 3). Isolated spread epitheliocytes are not capable of maintaining their constant average lengths. The lack of the cell length control is caused by another organizing and contractile feature of the actin cytoskeleton in epitheliocytes in comparison with fibroblastic cells [4]. Epithelial cells can be associated with each other into islets (Fig. 5.25) or into continuous cellular layers (Fig. 5.26). Thus, normal tissue cells completely spread on the extracellular matrix reach high degree of cell flattening; the cells have a well-developed lamellar cytoplasm. Fibroblastic cells prove to be polarized and maintain the cell length control, contrary to nonpolarized epithelial cells. The dorsal surfaces of fibroblastic cells are devoid of a surface relief; the smoothing of the cell surface relief is less pronounced in epithelial cells.
5.1 Cell Spreading on the Extracellular Matrix Surface
73
Fig. 5.24 Epithelial mouse cell. A part of the cell periphery: lamellar cytoplasm (arrow) with pseudopodia (double arrows) at its edge; short microvilli less numerous on the lamellar cytoplasm dorsal surface. SEM. Scale bar, 5.8 mm
Fig. 5.25 Epithelial mouse cells: an islet of the cells united with each other. SEM. Scale bar, 22.7 mm
74
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.26 Epithelial bull cells: the continuous cell monolayer. SEM. Scale bar, 14 mm
5.1.3 Morphological Alterations in the Spreading of Transformed Cells Transformed cells of mesenchymal or epithelial origin settling from suspension on the extracellular matrix undergo the process of spreading similar to normal tissue cells. However, the morphology of the cell spreading has essential differences resulting from oncogenic transformation. (a) At the radial stage of spreading of transformed fibroblasts, the excessive development of filopodia is frequently observed (Fig. 5.27). As a rule, the developing lamellar cytoplasm is defective, and its deficiency is typical for the early stage of spreading both the transformed fibroblasts and some types of transformed epitheliocytes. The defective lamellar cytoplasm is thickened, reduced, and/or fragmented (Figs. 5.28 and 5.29): instead of the united circular plate, the isolated lamellar “tongues” are formed (Fig. 5.30). With the development of the defective lamellar cytoplasm, transformed fibroblasts are spread unevenly, and they do not acquire the disc-like shapes (Fig. 5.31), which are observed in the normal fibroblastic or epithelial cells at the stage of radial spreading (Figs. 5.17 and 5.19). In contrast to transformed fibroblasts, transformed epitheliocytes at the stage of radial spreading can acquire disc-like shapes. However, the discoid cells, unlike normal epithelial cells at the radial spreading stage, often do not have the appearance of thin discs (Fig. 5.19) but look thickened, and they do not have a developed circular lamellar cytoplasm (Fig. 5.32).
5.1 Cell Spreading on the Extracellular Matrix Surface
75
Fig. 5.27 Transformed hamster fibroblast at the radial spreading stage. There are multiple filopodia (arrow) at the cell base. SEM. Scale bar, 3 mm
Fig. 5.28 Transformed mouse epitheliocytes at the radial spreading stage. There is the defective (thickened, reduced and fragmented) lamellar cytoplasm (arrows) at the cell base. SEM. Scale bar, 6 mm
76
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.29 Transformed hamster fibroblast at the radial spreading stage. The thickened and fragmented lamellar cytoplasm (arrow). SEM. Scale bar, 4 mm
Fig. 5.30 Transformed mouse fibroblast at the radial spreading stage. The defective lamellar cytoplasm: isolated lamellar “tongues” (arrows). SEM. Scale bar, 4.5 mm
5.1 Cell Spreading on the Extracellular Matrix Surface
77
Fig. 5.31 Transformed mouse fibroblast in the termination of the radial spreading stage. The elongated, poorly spread, and unevenly flattened cell. SEM. Scale bar, 8 mm
Fig. 5.32 Transformed mouse epitheliocyte in the termination of the radial spreading stage. The disc-like cell is thickened and the circular lamellar cytoplasm is strongly reduced. SEM. Scale bar, 7 mm
78
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.33 (a–c) Transformed cells at the polarization stage of the spreading: transformed mouse fibroblasts (a) hamster (b) and mouse (c) transformed epitheliocytes. Poorly spread polygonal or elongated cells. The lamellar cytoplasm is absent (a, c) or reduced and fragmented (arrows) (b). There are lamellipodia and ruffles at the polar cell ends. SEM. Scale bars, 7 mm (a), 9 mm (b, c)
(b) All transformed cells undergo the final stage of spreading, the polarization stage. Moreover, not only the transformed fibroblasts pass through the polarization stage, but also transformed epitheliocytes, in contrast to the normal epithelial cells. As a result, the transformed cells acquire the polygonal, elongated, or spindle-like shapes. However, in contrast to normal fibroblastic cells and epitheliocytes, the transformed cells are poorly spread, they are devoid of the lamellar cytoplasm, or it is strongly reduced (Fig. 5.33a–c). Therefore, transformed cells do not reach the significant degree of their flattening on the extracellular matrix. In the process of the transition from the suspended to the spread state in the transformed cells, in contrast to the normal ones, the progressive smoothing of the cell surface does not occur, and the spread transformed cells preserve their complex surface relief patterns (Fig. 5.34). Thus, deficient cell spreading is a typical feature of transformed cells. Morphological peculiarities of the spreading of transformed cells are associated with significant alterations in the functioning of focal contacts as the adhesive structures, accompanied by the relaxation of the centripetal stretch of actin filaments, and deficiency in the formation of tension-bearing stress fibers (see Sects. 5.4.1 and 6.2.1). These abnormalities cause a decrease in the ability of cellular pseudopodia to attach and centripetally stretch. After the completion of spreading, transformed fibroblasts acquire shapes of poorly spread polygonal or elongated cells, sometimes, hemispherical ones (Fig. 5.34). The mean area occupied by a transformed fibroblast is 2–5 times less than in a normal fibroblast. This decrease may be due to lack, reduction, or fragmentation of the lamellar cytoplasm. The decrease in the transformed fibroblast’s spreading is not even: it is the transversal, not the longitudinal cell spreading that decreases. As a result, the widths but not the lengths of fibroblastic cells decrease following their oncogenic transformation [3, 5]. The transformed fibroblast does not have a wide leading edge; lamellipodia and ruffles are formed at some short sections of the cell edge (Fig. 5.35). This is not accompanied by the decrease in the
5.1 Cell Spreading on the Extracellular Matrix Surface Fig. 5.34 Transformed mouse fibroblasts. The poorly spread polygonal, elongated, or semispherical cells with the microvillous surface relief. SEM. Scale bar, 15 mm
Fig. 5.35 Transformed hamster fibroblast. There are lamellipodia (arrows) and ruffles (double arrow) at short sections of the cell edge. SEM. Scale bar, 9 mm
79
80
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.36 Transformed mouse fibroblasts. Insufficient or no linear actin microfilament bundles in the cells. Staining for actin. FM. Scale bar, 15 mm
pseudopodial activity: the rate of extension and contraction of lamellipodia per unit length of the active cell edge does not decrease. The numbers of stress fibers and their associated focal contacts formed by transformed fibroblasts are significantly decreased (Fig. 5.36), as compared with normal fibroblastic cells (Fig. 5.21). Mature focal contacts often disappear completely, and only small focal complexes are revealed near the cell poles. Transformed epitheliocytes, like transformed fibroblasts, look like poorly spread cells. They have polygonal or elongated shapes, and their lamellar cytoplasm is often significantly reduced (Figs. 5.33b, c, 5.37, and 5.38). Circular actin filament bundles typical for normal epithelial cells disappear; the transformed epitheliocytes can contain straight actin filament bundles (Fig. 5.39) or not contain them at all (Fig. 5.40). Oncogenic transformation may lead to changes that are even more drastic: some transformed cells lose almost completely the ability to spread on the extracellular matrix. These cells retain their spherical or hemispherical shapes and they are attached to the matrix by only a few lamellipodia or filopodia (Fig. 5.41). The complex surface relief patterns are the characteristic feature of spread transformed cells. The surface relief is represented by numerous protrusions of different morphological types: microvilli, folds, blebs, or ruffles. The morphology of these protrusions observed in cells, which are in the suspended state, is described earlier. Similar morphology and the distribution of the protrusions on the cell surfaces are also observed in spread transformed cells.
5.1 Cell Spreading on the Extracellular Matrix Surface
81
Fig. 5.37 Transformed mouse epitheliocyte. Poorly spread cell, the lamellar cytoplasm is absent. There are lamellipodia and ruffles (arrows) at the ends of the cell processes. SEM. Scale bar, 10 mm
Fig. 5.38 Transformed mouse epitheliocytes. Poorly spread polygonal or elongated cells with the significantly reduced lamellar cytoplasm. SEM. Scale bar, 12 mm
82
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.39 Transformed mouse epitheliocytes. There are linear actin microfilament bundles in the cells. Staining for actin. FM. Scale bar, 7.5 mm
Fig. 5.40 Transformed mouse epitheliocytes. Insufficient or no linear actin microfilament bundles in the cells. Staining for actin. FM. Scale bar, 70 mm. Courtesy of S.N.Rubtsova
The whole surface of a spread transformed cell can have the protrusions of one type (Fig. 5.42), or the cells can have mixed surface relief consisting of the protrusions of several types (Fig. 5.43).
5.1 Cell Spreading on the Extracellular Matrix Surface
83
Fig. 5.41 Transformed hamster fibroblasts. The very poor spread cells. Some cells have a semispherical shapes with lamellipodia (arrows) and filopodia (double arrows) at the cell bases. SEM. Scale bar, 8.3 mm
Fig. 5.42 Transformed mouse fibroblasts. Poor spread cells. Microvillous cell surface relief. SEM. Scale bar, 10 mm
84
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.43 Transformed mouse fibroblast. The poor spread cell. The mixed cell surface relief: microvilli and folds. SEM. Scale bar, 3 mm
Microvilli are the most frequent element of the transformed cell surface relief. They can strongly vary in their lengths, can be straight or curved (Fig. 5.44), and can be branched (Fig. 5.45). Their distribution can be uniform or uneven; the density of their arrangement can vary even in the cells of the same type (Fig. 5.46). Side by side with microvilli, numerous folds are frequently observed (Fig. 5.47). Blebs are rarer protrusions on the surfaces of spread transformed cells. The bleb’s sizes can considerably vary from less than 1 mm to several mm in diameter (Fig. 5.48). Filopodia of different lengths and ruffles of various shapes can be often observed at the edges of transformed cells (see Figs. 4.2, 4.4, and 4.10 in Chap. 4, Figs. 5.33c and 5.37). The protrusions are dynamic formations. They can disappear and reappear on the cell surfaces in the process of the transition of cells from their spherical shapes in the suspended state to the spread state and vice versa, and also in the mitotic cycle or as a result of different influences. The protrusions are possible reserves of the cell membrane. For example, one microvillus with the length of 3 mm can reserve 1 mm² of the cell membrane. Since the surface of one transformed cell can contain more than 1,000 microvilli, the reserved cell surface is significant. It is possible that due to the cell surface excess in the microvilli and folds, intensive transport of nutrients (in particular, glucose) into the transformed cells is achieved. Actin cytoskeleton plays a crucial role in the determination of the shapes of the cell surface protrusions. Disorganization of actin cytoskeleton by cytochalasins
5.1 Cell Spreading on the Extracellular Matrix Surface Fig. 5.44 Transformed human epitheliocyte. Microvillous cell surface relief. The curved microvilli. SEM. Scale bar, 1 mm
Fig. 5.45 Transformed mouse fibroblast. Microvillous cell surface relief. Dichotomous branched microvilli (arrow). SEM. Scale bar, 0.5 mm
85
86
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.46 Transformed mouse fibroblasts. Microvillous relief of the cell surfaces. The density of microvilli arrangement varies in the cells. SEM. Scale bar, 8 mm
Fig. 5.47 Transformed rat fibroblast. Folded cell surface relief. The folds. SEM. Scale bar, 2 mm
5.1 Cell Spreading on the Extracellular Matrix Surface
87
Fig. 5.48 Transformed human fibroblast. Blebbed cell surface relief. The sizes of the blebs vary. SEM. Scale bar, 1.5 mm
Fig. 5.49 (a–c) Transformed mouse fibroblasts in the suspension state. The cells before (a) and after (b, c) cytochalasin B treatment. The shortening of the microvilli (b) and the formation of blebs (c). SEM. Scale bars, 3 mm (a), 2.5 (b, c). Reproduced with permission from the International Review of Cytology (ref. [1])
(the agents specifically inhibiting the polymerization of actin filaments and inducing their depolymerization) results in the shortening of microvilli and the formation of blebs (Fig. 5.49a–c) [1]. Apparently, the formation of microvilli involves the local polymerization of actin filaments, which form the core of the microvillus. In contrast, the formation of blebs may be a result of the flow of cytoplasm through the “openings” in the actin filament network [6]. Cytochalasins may favor the extension of blebs by increasing the size and number of the “openings.”
88
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Thus, transformed cells, as compared with normal fibroblastic or epithelial cells, poorly spread on the extracellular matrix. They have decreased spread areas and diminished sizes of their active (lamellipodia-forming) edges, decreased numbers and sizes of tension-bearing actin filament bundles (stress fibers) and of their associated focal contacts. Transformed cells have complex surface relief patterns.
5.2 The Signaling Pathways in the Spread Cells Cellular responses to different extracellular signals include activation of specific genes, stimulation or suppression of cell proliferation, alteration in the regulation of cell survival, changes in cell migration, and in the ability of cells to attach themselves to the extracellular matrix and to other cells. Multiple cell responses are based on the transduction of signals from a cell’s exterior to its interior: once a cell picks up a signal, it must transmit this information from the cell surface to the interior parts of the cell, for example, to the nucleus. Intracellular signal transduction involves a direct interaction of specific proteins in their strictly defined sequence. Basic participants are different types of protein kinases, the enzymes, which catalyze the transfer of phosphate from the ATP molecule to the amino acid (tyrosine, serine or threonine) residues of different proteins. The mechanism of signal transduction is based on the specific association of the proteins and on their phosphorylation (or dephosphorylation) state. Thereby, protein kinases regulate signaling pathways [7]. The phosphorylation of the protein targets leads to instantaneous changes in their conformations and properties. The result is the sequential activation of the proteins in the signal chain, which ensures the sequential signal transduction from the cell surface into the cell to its genetic apparatus in the nucleus. The most common are molecular signals, which include hormones, soluble growth factors (GF), extracellular matrix components, and other specific chemical agents. Stretch, pressure, and other mechanical effects can also act as the signals.
5.2.1 Cell Surface Receptors Extracellular signaling molecules called ligands can be freely soluble or can be present in the extracellular matrix, and also at the surfaces of other cells. Ligands first interact with the specialized proteins called cell receptors. They reside at the cell surface (cell surface receptors), but sometimes within the cell (intracellular receptors). The receptors are mostly very specific to signaling molecules: a given receptor usually binds only a definite ligand or a group of related ligands. The vast majority of extracellular signaling molecules are recognized by cell surface receptors. They are integral transmembrane proteins. Cell surface receptors
5.2 The Signaling Pathways in the Spread Cells
89
span the plasma membrane of the cell, with one part of the receptor on the outside of the cell (the extracellular domain), and the other, on the inside of the cell (the intracellular domain). Signal transduction starts as a result of the binding of a ligand to the extracellular domain of the cell surface receptor. The ligand itself does not pass through the plasma membrane. Upon binding, the ligand initiates the transmission of a signal across the plasma membrane by inducing changes in conformation (shape) of the intracellular domain of the receptor. Such conformational changes either result in the activation of an enzymatic activity contained within the receptor or expose a binding site for other proteins within the cell. Once these proteins bind to the receptor, they themselves are activated and propagate the signal into the cytoplasm. Thus, the binding of ligands to cell surface receptors triggers the sequential biochemical reactions inside the cell, which are carried out by enzymes, resulting in a signal transduction pathway. A given receptor activates only specific sets of downstream signaling molecules inside the cell. However, different incoming signals can activate the same signal transduction pathways. The classes of cell surface receptors include receptor tyrosine kinases (RTKs), integrins, G protein coupled receptors (GPCR), and ion channel receptors (the latter may occur both at the cell surface and intracellularly). Receptor Tyrosine Kinases (RTKs) RTKs include receptors for soluble growth factors (GF) (see Sect. 5.3.2) and hormones such as insulin [8]. A typical RTK has a ligand-binding extracellular domain, a plasma membranespanning domain, and a tyrosine kinase intracellular domain. RTKs form dimers (two receptor molecules) in the plasma membrane. In the absence of its specific ligand, one receptor remains unbound to a second receptor. To begin the signal transduction, the ligand must bind to two receptor molecules and crosslink them. The interaction between two tyrosine kinase intracellular domains stimulates the auto-phosphorylation of the tyrosines causing the conformational changes of the intracellular domains. They may now bind to one or more other cytoplasmic proteins that specifically recognize the phosphorylated tyrosines. Many of these cytoplasmic proteins are enzymes, which are often inactive until they bind to the activated intracellular domains of the RTKs. The products of these enzymes may act on other molecules, thus continuing the signal transduction, which is the basis of various cellular processes, such as control of proliferation, metabolism, and cell migration [8]. Soon after the ligand-induced activation of RTK tyrosine kinase intracellular domains, RTKs are efficiently internalized from the surfaces of GF-stimulated cells and are subsequently degraded [9].
90
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Integrin Receptors These receptors play a key role in the adhesion of cells to the extracellular matrix. The protein components of the matrix, such as collagen, fibronectin, and laminin serve as ligands for the integrin receptors. Integrins do not have kinase activity. The binding of the extracellular matrix protein ligand to the extracellular domain of the receptor induces integrin activation based on the conformational changes and a clustering of the receptor. These events initiate integrin-mediated signal transduction pathways through a variety of specific cytoplasmic proteins. The signaling pathways triggered by activated integrins control cell-matrix adhesion, cell migration, proliferation, and survival [10–12] (about the integrin-mediated signaling, see Sect. 5.3.1, 5.3.3, 5.3.5, and 5.3.6). Some integrins can mediate cell–cell adhesion. They are found at the surfaces of circulating blood cells, such as leukocytes or platelets [13]. The ligands for these integrins belong to another superfamily of intercellular adhesion molecules, containing Ig-like domains, which are expressed on vascular endothelium. Therefore, the integrins provide leukocyte adhesion to the vascular endothelium; this is a necessary condition for the subsequent extravasation (release from a vessel) and migration of leukocytes to the inflammation sites [14]. Integrins can cooperate with other types of cell surface receptors, in particular, with GF receptors, and thereby be involved in various cell activities. For example, there is the interplay between integrins having b1 subunit and the epidermal growth factor (EGF) receptor. Integrins with b3 and b5 subunits collaborate with GF receptors to control angiogenesis. Integrins work together with co-receptors for GF, proteoglycans syndecans, to regulate cell-matrix adhesion and cell migration; integrin a6b4 cooperates with the RTKs in normal functioning of epithelial cells [15, 16]. G Protein Coupled Receptors (GPCRs) GPCRs are cell surface receptors linked to guanine nucleotide-binding proteins (G proteins). G proteins belong to two distinct families: heterotrimeric G proteins (sometimes called “large” G proteins) and “small” G proteins, called small GTPases (or Ras superfamily GTPases). An intracellular domain of a GPCR is associated with a particular type of heterotrimeric G protein at the inner surface of the plasma membrane [17, 18]. G proteins, both heterotrimeric G proteins and small GTPases, act as “molecular switches,” alternating between an inactive guanosine diphosphate (GDP) and active guanosine triphosphate (GTP) bound states. In unstimulated cells, G proteins are in the GDP-bound form. They are activated when GDP is replaced by GTP. GTP is in excess in cells, thus GDP release is rapidly followed by GTP binding. Three groups of factors take part in the regulation of transitions from the active to the inactive states of G proteins: (a) Guanine nucleotide exchange factors (GEFs), which catalyze the exchange of GDP to GTP, i.e., GEFs “turn on” the activity of G proteins.
5.2 The Signaling Pathways in the Spread Cells
91
(b) GTPase activating proteins (GAPs), which stimulate the GTP hydrolysis, i.e., GAPs “turn off” the activity of G proteins. (c) GDP dissociation inhibitors (GDIs), which prevent the changing of GDP for GTP, i.e., GDIs maintain the inactive state of G proteins. GPCRs are cell surface receptors for chemokines (small soluble proteins that induce directional chemotaxis and can be involved in the control of cell migration), hormones, and some other substances. The binding of the specific ligand to extracellular domain of GPCR induces its conformational change that is transmitted to the intracellular domain, which interacts with the heterotrimeric G protein at the inner surface of plasma membrane. This results in the activation of the heterotrimeric G protein and its profound conformational changes including the dissociation of the G protein into its subunits. The activated subunits diffuse away from the GPCR and interact with many other molecules in the cell [17, 18]. Signal transduction mediated by GPCRs is involved in the regulation of actin cytoskeleton remodeling leading to cell migration [19]. Ion Channel Receptors They present a class of cell receptors that may exist both at the cell surface and intracellularly. Ion channel receptors are responsive either to specific ligands or to differences of the electrical charge across the plasma membrane. The activated receptors trigger the receptor structural changes that open a channel in the plasma membrane allowing the passage of ions. The movement of ions changes the plasma membrane potential, which in turn changes cellular functions. Besides cell surface receptors, there are receptors inside a cell. These intracellular receptors bind the cell-permeable ligands, such as steroids, thyroid hormone, and vitamin D. Once the receptors are activated by the ligands, they are translocated to the cell nucleus, where they modulate the expression of specific genes.
5.2.2 Intracellular Signal Transduction Following ligand-receptor interaction, the intracellular signal transduction involves the sequential altering of the conformation (and therefore, the activity) of many intracellular proteins participating in the signaling pathway. In altering the conformation and activity of proteins, turning them on or off like a molecular switch, the phosphorylation–dephosphorylation mechanism has a key role. Phosphorylation may open up an enzyme’s active site, allowing it to catalyze the chemical reactions. Phosphorylation may also generate a binding site allowing a specific interaction with a molecular partner. The addition of phosphate group to other proteins is performed by protein kinases; the proteins, which the kinases act
92
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
on, are called protein kinase substrates. Protein kinases themselves can be turned on by phosphorylation performed by other protein kinases. Therefore, a protein kinase can be both an enzyme and a substrate. Depending on the amino acids to which phosphate groups are added, protein kinases are divided into serine/threonine kinases and tyrosine kinases (there are also a few dual-specificity kinases known, such as protein kinase CK2). The turning off is achieved by specific protein phosphatases, which can remove phosphate groups from protein kinase substrates. Most intracellular proteins sequentially activated by a ligand–receptor interaction are enzymes including tyrosine and serine/threonine kinases, phosphatases, lipid kinases and G proteins, both heterotrimeric G proteins, and small GTPases. G proteins play a major role in the intracellular signaling pathways. Heterotrimeric G proteins and small GTPases are involved in the transduction of signals from activated cell surface receptors. Heterotrimeric G proteins participate in GPCRmediated signal transduction, while small GTPases are involved in RTK-mediated and integrin receptor-mediated signal transduction pathways. Small GTPases are composed of several families, including Ras, Rho, Rab, and some other families. Small GTPases, just as heterotrimeric G proteins, act as “molecular switches,” alternating between an inactive GDP and active GTP states [20]. Upon activation, small GTPases are responsible for the recruitment of definite proteins to the specific cell membrane sites where those proteins participate in intracellular signaling. Second Messengers Many of the enzymes participating in the intracellular signal transduction pathways are activated by the family of special substances called the second messengers. The enzymes possess specialized domains that bind to the specific second messenger molecules. Second messengers have a key role in intracellular signal transduction pathways. They are involved in the control of such fundamental cellular processes as cell survival, proliferation, and cell migration, adhesive interactions of cells with the extracellular matrix and with each other. The group of intracellular second messengers includes: Calcium ions, cyclic nucleotides (in particular, adenosine monophosphate, cAMP), fraction of inositol-containing cell membrane phospholipids called phosphoinositides (PIPs), and also soluble inositol triphosphate and diacylglycerol. Calcium ions are normally kept at very low levels in the cytoplasm. The signaling pathways from activated cell surface receptors lead to Ca2+ ions release from the endoplasmic reticulum into the cytoplasm. This results in the binding of released Ca2+ ions to signaling proteins that are then activated. In particular, calcium ions bind specifically to calmodulin, the protein that can bind to and regulate different protein targets. For example, calcium-bound calmodulin activates a family of serine/threonine
5.2 The Signaling Pathways in the Spread Cells
93
kinases, called calmodulin dependent protein kinases (CAM kinases), which themselves regulate the activity of many important proteins. Increased levels of Ca2+ ions in the cytoplasm plays a very important role in cell proliferation and migration, cytoskeleton dynamics, cell contraction, secretion, neural signal transduction, and gene expression. The cAMP is a small diffusible substance. It exerts much of its effects by activating the serine/threonine protein kinase A (PKA). The activity of PKA is regulated by fluctuating levels of cAMP; PKA is also known as cAMP-dependent protein kinase. PKA influences such important cellular processes, as membrane transport (activation of certain calcium and chloride ion channels), energy metabolism, actin-based cell migration, and muscle contraction. PKA also phosphorylates some transcription factors, whereby PKA can regulate the expression of particular genes. PIPs are powerful second messengers having a key role in intracellular signal transduction pathways [21, 22]. They are inositol-containing cell membrane phospholipids, which are phosphorylated products of phosphatidylinositol (PI), a minor phospholipid component of cell membranes. As a result of sequential phosphorylation carried out by phosphatidylinositol kinases (PIKs), in particular phosphatidylinositol 3-kinases (PI3Ks), and phosphatidylinositol phosphate kinases (PIPKs), and also as a result of dephosphorylations carried out by phosphatases, distinct PIPs are generated on the cytoplasmic side of plasma membrane [23]. The list of PIPs includes phosphatidylinositol monophosphates, phosphatidylinositol biphosphates, PIP2 (PI4,5P2; PI3,5P2; PI3,4P2), and phosphatidylinositol triphosphate, PIP3 (PI3,4,5P3). PIPs function as ligands for PIP-binding proteins: specific lipid-binding protein domains mediate interactions between proteins and specific PIPs. These domains are found in many proteins participating in intracellular signaling. PIPs can cause conformational changes of these proteins, and alter enzymatic activities [24–26]. Through direct interactions with PIPs, many proteins involved in signaling pathways are recruited to cellular membranes and/or are activated [27, 28]. The generation of different PIPs in the cell is strictly localized. By means of the regulated PIKs and PIPKs targeting to appropriate intracellular compartments, PIPs are locally generated at different parts of the inner side of plasma membrane. The targeting is carried out by one of subfamilies of small GTPases, Rho-GTPases, which selectively recruit and activate PIKs and PIPKs in response to ligand-receptor interaction [23, 28–30]. On the other hand, small GTPases themselves can be regulated by PIPs through translocation and conformational changes [31]. Among different phosphoinositides, PIP2 (PI4,5P2) and PIP3 (PI3,4,5P3) play particularly important roles as second messengers [21, 22]. PIP2 is a product of PIPKs, a family of kinases, including types I, II, and III. PIP2 is generated by type I PIPK. PIP3 is a product of phosphatidylinositol 3-kinases (PI3Ks), a family of kinases composed of I, II, and III classes. PIP3 is synthesized by class I PI3K. PIPKs, PI3Ks, and their products PIP2 and PIP3 are important components of various signal transduction pathways, controlling actin cytoskeleton organization,
94
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
ion channel regulation, cell migration, chemotaxis, cell survival, proliferation, and also adhesive interactions of cells with the extracellular matrix and with each other. Both PIP2 and PIP3 are critical regulators of actin cytoskeleton dynamics. Many proteins that are involved in the coordinated reorganization of actin cytoskeleton and in focal adhesion assembly–disassembly, which are the basis of cell migration, are regulated by PIP2 and PIP3. Through direct interaction with PIP2 and PIP3, these proteins are recruited and/or activated [21, 22, 27, 32–34]. Local character of PIP2 and PIP3 generation is essential for regulating cell morphology during locomotion. PIP2 is locally generated at cell ruffles and focal adhesions of the moving cell [32, 35]. The localized production of PIP3 at the cell side facing the highest chemoattractant concentration controls the leading cell edge and regulates directional migration of the cell by controlling its actin cytoskeleton [34]. PIP3 is involved in cell survival control contributing to the suppression of the programmed cell death, or apoptosis [36–38]. Inositol triphosphate and diacylglycerol are soluble second messengers, which are the products of the cleavage of PIP2 by phospholipase C. Inositol 1,4,5-triphosphate (IP3) diffuses to IP3 receptors in the endoplasmic reticulum membranes, binds to them, and then opens the ion channels, releasing stored Ca2+ ions into the cytoplasm. IP3-mediated calcium signaling is involved in the effects of many external stimuli inside the cell. Diacylglycerol (DAG) stays on the cell membrane and binds Ca2+-dependent serine/threonine protein kinase C (PKC) that becomes activated and, in turn, activates other cytoplasmic proteins by phosphorylating them [39]. PKC-mediated signaling is involved in the regulation of cell growth, actin cytoskeleton morphology, ion channel activity, and secretion [40, 41].
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells The adhesive binding of spread cells with the extracellular matrix, achieved by means of focal contacts (focal adhesions), is not only structural, but also has an important functional role. The binding of the integrin receptor to its extracellular matrix ligand “triggers” different signal transduction pathways from focal contacts into the cells. These sequential signals reach the cell nucleus and induce the expression of specific genes. Integrin-mediated signaling pathways determine and control basic cell functions: proliferation, survival, and locomotion [10, 12, 42, 43]. The cytokines termed soluble GF, being bound to their specific cell surface receptors, trigger signal transduction pathways. These pathways, like the integrin-mediated signaling pathways, control proliferation of the cells, their locomotion, and survival. These functions are served by the integrin and GF receptors cooperatively.
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
95
Fig. 5.50 Diagram showing the outside-in signal transduction pathways from integrin receptor (see the text for explanation)
5.3.1 Integrin Receptor-Mediated Mitogenic Signaling Pathways In the integrin bound to its extracellular matrix ligand, the b cytoplasmic domain induces the phosphorylation of a number of proteins of the focal contact: focal adhesion kinase (FAK) directly linked to the cytoplasmic domain of the integrin b subunit, paxillin and tensin (see Fig. 4.14 in Chap. 4). FAK activated through its auto-phosphorylation plays a main role in the initiation of the signal transduction pathways from integrins (Fig. 5.50). The autophosphorylation of FAK is regulated by the endogenous protein FIP200 that can inhibit FAK auto-phosphorylation and thereby FAK activity [44]. The activated FAK gets associated with the submembrane Src protein kinase initiating its tyrosine kinase activity. FAK and Src tyrosine kinases form a dual kinase complex that binds to and rapidly phosphorylates on tyrosine various focal adhesion proteins, such as p130Cas (Cas), paxillin and tensin. Another signaling protein, p140Cap (Cas-associated protein) is also tyrosine phosphorylated soon after cell-matrix adhesion [45]. The signal from FAK is also transferred to the Ras GTPases, which belong to one of families of small GTPases, putting them into the active state. The activated Ras GTPases stimulate the activity of cytoplasmic serine/threonine protein kinases of Raf family, which, in turn, activate the cascade of mitogen activated protein (MAP) serine/ threonine kinases. MAP kinases, also known as extracellular signal-regulated kinases
96
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
(ERKs), are key regulators of cellular mitotic cycle. The final products of the MAP kinase cascade are the serine/threonine kinases that are getting translocated from the cytoplasm to the nucleus, where they phosphorylate and activate a series of transcription factors, which are nuclear proteins regulating the activity of specific genes. This leads to the activation of other transcription factors, which activate the genes whose products initiate the entry of the cell into the phase of DNA replication (Fig. 5.50). The efficiency of the Ras-Raf-MAP kinase mitogenic signaling pathway is modulated by p21-activated serine/threonine kinases (PAKs) [46]. Thus, the result of the Ras–Raf-MAP kinase sequential signal transduction pathway from integrin receptors bound to the extracellular matrix is the expression of specific genes, which stimulate cell proliferation.
5.3.2 Growth Factor Receptor-Mediated Mitogenic Signaling Pathways The Ras-Raf-MAP kinase intracellular signaling pathway mediates the transduction of mitogenic signals not only from integrins, but also from GF receptors that are cell surface receptors to specific cytokines, soluble GF. Soluble GF form a large group of different secreted proteins. This group includes such GF as platelet-derived growth factor (PDGF), epidermal growth factor (EGF), insulin-like growth factor (IGF-1), fibroblast growth factors (FGFs), vascular endothelial growth factor (VEGF), hepatocyte growth factor/scatter factor (HGF/SF), and some others. PDGF is an important mitogen for connective tissues. VEGF is endothelial cellspecific mitogen and angiogenic inducer. HGF/SF is a very interesting GF. It is produced by various cells of mesenchymal origin including fibroblasts, vascular endothelium (activated during hepatic damage), smooth muscle cells of the vascular wall, and macrophages. HGF/SF is a strong mitogen for hepatocytes. It is involved in liver regeneration, and is also called hepatocyte growth factor (HGF). It also stimulates proliferation of epitheliocytes, vascular endothelial cells, and melanocytes [47]. GF bind to their specific cell surface receptors that belong to the group of RTKs. GF receptors form dimers, and two receptor molecules remain unbound to each other in the absence of a specific GF. When GF binds to two receptor molecules, it cross-links them. This causes two tyrosine kinase intracellular domains to come into contact with one another, and each tyrosine kinase phosphorylates the other one. The activation of tyrosine kinase domains initiates the phosphorylation and activation of downstream cytoplasmic proteins including tyrosine kinases of Src family, phosphatidylinositol 3-kinase (PI3K) and Ras GTPases. The subsequent passage of the signal along the Ras-Raf-MAP kinase pathway to the nucleus (see above) initiates cell multiplication (Fig. 5.51). Rapid internalization of RTKs from the surfaces of GF-stimulated cells can be carried out through lamellar outgrowths, ruffles, on the dorsal cell surface [9]. However, the cascade of mitogenic signal transduction from GF receptors is achieved only in cells attached to the extracellular matrix: in the case of cell detachment,
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
97
Fig. 5.51 Diagram showing the signal transduction pathways from growth factor receptors (see the text for explanation)
mitogenic signals from GF receptors are blocked at one of the intermediate kinases of the MAP kinase cascade, and as a result, detached cells cannot proliferate [48]. Thus, binding of integrins of focal contacts to the extracellular matrix not only produces mitogenic signals, but is also the necessary condition for mitogenic action of soluble GF.
5.3.3 Integrin and Growth Factor Receptor-Mediated Antiapoptotic Signaling Pathways Focal contacts not only initiate mitogenic signals, but they also suppress programmed cell death, or apoptosis [49]. One of the mechanisms of apoptosis control is the regulation of the release of special apoptotic molecules from cell mitochondria (Fig. 5.52). These molecules finally cause the activation of cytoplasmic proteinases termed “effector caspases.” The latter cleave a series of key proteins, which leads to DNA fragmentation and to cell destruction. The permeability of mitochondrial membrane for apoptotic molecules
98
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.52 Diagram of the signaling pathways regulating apoptosis (see the text for explanation)
is regulated by the products of BCL2 and BCL-x proto-oncogenes and by the product of BAX tumor suppressor gene. Bcl2 and Bcl-x proteins reduce mitochondrial membrane permeability and in this way, they block apoptosis; Bax protein increases permeability and thus induces apoptosis [49–52]. The binding of integrins of focal contacts to the extracellular matrix and subsequent activation of FAK puts Ras GTPases into the active state. The activated Ras initiates antiapoptotic signaling pathways (Fig. 5.52) [49]: (a) One of the targets of Ras, protein kinases of Raf family, directly phosphorylate and inactivate the apoptotic protein Bad; Raf protein kinases also activate the antiapoptotic Bcl2 and Bcl-x proteins. (b) The other target of Ras is phosphatidylinositol 3-kinase (PI3K). The product of PI3K, PIP3 recruits serine/threonine protein kinase B (PKB, widely known as Akt) to the plasma membrane, where it is activated by other kinases that are also dependent on PIP3. Different PKB/Akt isoforms influence numerous cellular targets involved in actin cytoskeleton remodeling and cell motility [53]. Moreover, PKB/Akt is an important
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
99
signaling element in cell survival pathways: PKB/Akt protects cells from apoptosis in several ways, including inactivation of Bad (Fig. 5.52) [36–38]. Since the activation of Ras and its target PI3K can be induced by signals not only from integrins (Fig. 5.50), but also from GF receptors (Fig. 5.51), the antiapoptotic Ras-PI3K-PKB/Akt signaling pathway can be triggered by GF. The antiapoptotic signaling pathway is negatively regulated by a specific protein, called phosphatase and tensin homolog (PTEN). PTEN is a dual protein/lipid phosphatase that dephosphorylates PIP3. In addition to that, PTEN activates the apoptotic protein Bad. Thereby, PTEN inhibits the antiapoptotic effect of that signaling pathway (Fig. 5.52). Suppression of integrin activity and blockage of integrin-mediated survival signals is affected by a special protein kinase called death-associated protein (DAP) that can induce apoptosis. DAP is a serine/threonine kinase, the product of tumor suppressor gene. DAP is associated with actin filament system. Through an insideout mechanism, DAP suppresses integrin-mediated cell adhesion and signal transduction. Thereby, the p53-dependent apoptotic signaling pathway is activated (see Sect. 5.3.4), and the cell undergoes apoptosis [54].
5.3.4 The “Anchorage Dependence” When normal cells become detached from the extracellular matrix, their proliferation is inhibited and apoptosis is induced. The detachment-induced apoptosis is called anoikis [55]. Several mechanisms are involved in the inhibition of cell proliferation and apoptosis in the detached cells. (a) One of the mechanisms is the caveolin-1 – dependent endocytosis of signal molecules. Key components of signal pathways controlling cell proliferation and survival, such as MAP kinases and PI3K, localize to the special cholesterolenriched and sphingolipid-enriched plasma membrane domains in response to various stimuli. The trafficking of these domains called caveolae and lipid rafts is regulated by integrin-mediated cell-matrix adhesion. Upon cell detachment, these domains undergo rapid endocytosis and clearing from the plasma membrane. Endocytosis is mediated by caveolin-1, a constitutive protein of caveolae, and the protein dynamin-2, a regulator of caveolae dynamics. The caveolin-1dependent endocytosis causes the inhibition of MAP kinases and PI3K, and as a result, the detached cells stop their proliferation and undergo anoikis [56–58]. (b) The block of proliferation of detached cells is also caused by the activity of the tumor suppressor genes, which are negative regulators of proliferation: the proteins encoded by these genes prevent the entry of cells into the phase of DNA replication. For example, cell detachment activates p53 tumor suppressor gene and induces the accumulation of the product of the p27/KIP1 tumor suppressor gene resulting in the block of cell proliferation [59]. (c) GF cannot transduce their mitogenic signals in the absence of focal adhesions: the mitogenic signals from GF get blocked in the nonattached cells (see Sect. 5.3.2).
100
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
(d) A pivotal cause of anoikis in the detached cells is the activation of p53 tumor suppressor gene (Fig. 5.52). This gene is activated as a response to various unfavorable factors (cell DNA damage, hypoxia, etc.) including the release of cells from focal adhesions. The activated p53 simultaneously activates apoptotic BAX tumor suppressor gene and inhibits antiapoptotic BCL2 protooncogene. Thus, the activation of p53 caused by cell detachment generates powerful apoptotic signal. (e) In addition, there can be another mechanism of anoikis: in the cells attached to the matrix, one of the signal transduction chains (from the activated Ras) activates the protein kinase PKB/Akt, which protects cells from apoptosis (see Sect. 5.3.3). The detachment of cells from the matrix blocks this antiapoptotic signaling pathway [52, 60, 61]. The described mechanisms are the basis of the phenomenon, called substratum dependence of cell proliferation or anchorage dependence: most of the normal cell types can survive and multiply only after being attached and sufficiently spread on the extracellular matrix; in liquid medium the suspended tissue cells do not multiply and can undergo apoptosis (anoikis) (Fig. 5.53). Cells can avoid anoikis for some time by autophagy. It is a catabolic process that involves a regulated turnover and elimination of damaged or unnecessary proteins and some cellular organelles through their lysosomal degradation. The resultant molecules then are reutilized. Autophagy maintains homeostasis and survival of the cells that are exposed to various stresses including GF deprivation. Presumably, autophagy protects epithelial cells from the stress of their detachment from the extracellular matrix allowing them to avoid anoikis for some time till their reattachment [62]. In this way, focal contacts are not only the adhesive structures, which mechanically connect tissue cells with the extracellular matrix, but are also the transducers of the intracellular signals, which ensure the survival of cells and their proliferative activity. Therefore, integrins can signal in two directions: “inside-out” and “outside-in.” They mediate inside-out signaling, by which the integrin receptor affinity to the protein ligands in the extracellular matrix is modulated through intracellular events (see Sect. 4.2.1). Integrins also mediate outside-in signaling, which is initiated after binding of the extracellular matrix ligands to integrins. The generated signals finally result in the gene expression that induces the proliferation of the cells and ensures their survival. Both types of integrin signaling, inside-out and outside-in, are mediated by integrin b subunit cytoplasmic domain [63].
5.3.5 Integrin and Growth Factor Receptor-Mediated Morphogenic Signaling Pathways Besides the mitogenic Ras-Raf-MAP kinases signaling pathway, integrins and GF receptors can trigger the “morphogenic” Ras-PI3K-Rho signaling pathways (Figs. 5.50 and 5.51). These pathways that lead to the reorganization of the cytoskeleton
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
101
Fig. 5.53 Diagram showing the substratum dependence or independence of cell proliferation. The cells capable (green) or incapable (pink) of the proliferation (see the text for explanation)
structures, cause the assembly or disassembly of focal adhesions, induce pseudopodia formation, and in this way these signals control cell morphology and migration. In the “morphogenic” signaling pathways, the signals from activated Ras GTPases are transferred through phosphatidylinositol-3 kinase (PI3K) to the Rho GTPases, which like Ras GTPases, are one of families of small GTPases. Rho GTPases play a key role in the “morphogenic” signaling pathway leading to actin cytoskeleton reorganization, plasma membrane protrusions, and cell migration. The family of Rho GTPases includes Rho (A, B, and C isoforms), Rac (1, 2, and 3 isoforms), and the Cdc42. Besides, there are some other members of Rho family of GTPases. These small GTPases are involved in integrin-mediated signaling that regulates actin cytoskeleton remodeling, focal adhesion assembly, cellular pseudopodia formation, and directional cell migration [20, 64, 65].
102
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.54 Diagram of the Rho regulation of actin cytoskeleton (see the text for explanation)
Rho Proteins (Fig. 5.54) These activated GTPases have two main targets: Rho-associated kinase (Rho kinase, ROCK) and the formin homology protein mDia1 [66]. ROCK inactivates MLC phosphatase and probably, directly phosphorylates MLC. As a result, myosin II-dependent contractility of actin filaments is increased. The increase results in the formation of filament bundles and associated mature focal contacts. In addition, ROCK phosphorylates and activates LIM serine/threonine protein kinases, which inhibit cofilin-induced depolymerization of actin filaments promoting their stabilization [67–69]. Actin nucleator mDia1 stimulates the polymerization of actin filaments and induces their linear growth (unlike the Arp 2/3 complex that induces the formation of the branching pattern of the filaments). MDia1 is connected to the actin-binding protein profilin. Together with ROCKs, mDia1 takes part in the actin filament polymerization and in the formation of stress fibers. In addition, mDia1 influences the dynamic of microtubules, providing their “targeting,” the concentration of microtubule plus-ends near growing focal contacts. Rho proteins control the polymerization and stabilization of actin filaments, the formation of stress fibers and associated focal contacts, and initiate the microtubular “targeting.”
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
103
Fig. 5.55 Diagram of the Rac regulation of actin cytoskeleton and lamellipodia formation (see the text for explanation)
Rac Proteins (Fig. 5.55) The targets for activated Rac are PIPKs, serine/threonine kinases (p21-activated kinases, PAKs) and the WAVE (also called Scar) proteins. The activated type I PIPK induces the formation of phosphatidylinositol 4, 5- biphosphate (PIP2). The latter, releasing the barbed ends of actin filaments from the capping protein gelsolin, promotes actin polymerization at these ends and the growth of filaments [33]. The activated PAKs phosphorylate LIM kinase that inhibits cofilin-induced depolymerization of actin filaments promoting their polymerization [67–69]. WAVE/Scar, like WASP, activates the Arp2/3 complex, initiating the growth of new branching actin filaments. Their growing ends face the cell membrane and presumably could protrude from it forming lamellipodia (see Sect. 6.1.1). The result of Rac activation is the stimulation of actin filament branching “tree” growth leading to the formation of lamellipodia at the cell edge. In addition, the activation of Rac1 promotes microtubule growth into advancing lamellipodia of migrating cells.
104
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
Fig. 5.56 Diagram of the Cdc42 regulation of actin cytoskeleton and filopodia formation (see the text for explanation)
Cdc42 Protein (Fig. 5.56) Its targets are WASP/WAVE (the alternative name is WASP/Scar) proteins and PAK kinases. The activated WASP/WAVE (WASP/Scar) in their turn activate Arp2/3 complex initiating the growth of the branched actin filaments [70]. The activated PAK kinases promote filament polymerization. Therefore, Cdc42, like Rac, stimulate actin polymerization and the growth of filaments. The result of Cdc42 activation, as well as Rac activation, is the formation of pseudopodia. However, Cdc42 initiates the formation of very narrow outgrowths, filopodia, but not of wide protrusions, lamellipodia. The reason for this is that some of the growing actin filaments acquire privileged status owing to which they are joined with each other by their barbed ends. Then these filaments are sewn together into long bundles, which presumably could protrude the cell membrane, forming filopodia (see Sect. 6.1.1). In addition, Cdc42, together with cytoplasmic dynein, is involved in the orientation of the microtubule-organizing center (centrosome) toward the leading edge of migrating cells. Thus, in the “morphogenic” Ras-PI3K-Rho signaling pathways from integrin and GF receptors, the activated Rho GTPases initiate the “descending” transfer of the signals. These signals cause alterations in the dynamics and organization of actin cytoskeleton and microtubule system, induce the assembly/disassembly of focal contacts, and also initiate the formation of different pseudopodia.
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
105
Activity of Rho GTPases is regulated by FAK. FAK facilitates the activation of GEFs and GTPase-activating proteins (GAPs), which activate or inactivate Rho GTPases respectively [71]. Therefore, the cooperative integrin and GF receptor-mediated signaling pathways control cell proliferation, cellular survival, apoptosis, and also cell-extracellular matrix adhesion and formation of different pseudopodia that are major mediators of cell migration.
5.3.6 Integrin-Mediated Mechanical Force-Induced Signaling Tissue cells completely spread on the extracellular matrix constantly experience isometric tension. It is the result of the interaction of two forces: the centripetal stretch of actin stress fibers anchored to focal contacts (see Figs. 4.13a–c in Chap. 4) and the counteracting force of cell adhesion to the extracellular matrix. The intracellular “shock absorbers,” microtubules, also resist the stretch of stress fibers. The dynamic equilibrium of the mechanical forces in the cell is the important factor, which controls the shape of cells, the degree of their spreading on the extracellular matrix, and their functional and proliferative activity. These forces render the modifying influence on intracellular signal transduction pathways and on gene expression. Alterations of the balance of mechanical forces supporting the state of the isometric tension of cells, lead not only to changes in cell shapes and cytoskeleton reorganizations, but also to the disturbances of the cell synthetic processes and to the changes in the proliferating activity of the cells [72]. The increase in the isometric tension of spread cells can be caused by external mechanical stress. Fibroblastic cells “sense” external mechanical signals from the extracellular matrix when the matrix undergoes periodic bending, pulling, or stretching. The cells respond to exerted mechanical forces by reinforcement and reorientation of actin stress fibers, formation of mature focal contacts, and also by reorienting cell migration and remodeling of the extracellular matrix. Integrin receptors act as a “mechanosensory switches”: they are bound to the extracellular matrix and after mechanical stimulation trigger force-induced intracellular signals [73–77]. The mechanical force-induced signals affect focal contact formation: upon application of external pulling forces, focal adhesions self-assemble and elongate; when these forces are decreased, the focal adhesions disassemble [78–80]. Talin is critical for the force-dependent reinforcement of initial integrin-actin cytoskeleton bonds: deletion of talin blocks early formation of focal contacts in response to external mechanical forces [81]. Formin mDia1 is involved in force-dependent formation of focal adhesions: expression of active mDia1 allows external pulling force-induced assembly of mature focal contacts even in conditions when myosin II-driven cell contractility is inhibited [73]. The mechanical force-induced signals rapidly stimulate tyrosine phosphorylation of FAK and p130Cas (Cas) protein. Thus, FAK and p130Cas can be regarded as mechanosensory proteins.
106
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
The force-induced phosphorylation of FAK generates integrin-dependent mitogenic signal, which stimulates cell proliferation (Fig. 5.50). The force-induced p130Cas phosphorylation is involved in the activation of one of the small GTPases, Rap1, leading to the inside-out integrin activation (see Sect. 4.2.1). Mechanical stress activates RhoA protein, a member of Rho family of GTPases, and thereby induces the assembly of actin filaments into bundles and increases their contractility (Fig. 5.54) [75, 77, 82]. In external force-induced signal transduction, the protein zyxin plays an important role. Zyxin is a zinc-binding phosphoprotein that co-localizes with integrins at sites of fibroblastic cell-matrix attachments (see Fig. 4.14 in Chap. 4). Zyxin has the ability to associate with a-actinin and the enabled/vasodilator-stimulated phosphoprotein (Ena/VASP) family, which mediate the association of the barbed ends of actin filaments with each other and are involved in the assembly of actin cytoskeleton [83, 84]. In addition, zyxin can shuttle between the sites of cell adhesion and the nucleus in fibroblastic cells to trigger the expression of mechanosensory genes [85]. Mechanical stress induces the mobilization of zyxin and zyxin-dependent mobilization of VASP, a member of the Ena/VASP protein family, from focal contacts to actin filaments. It leads to rapid actin stress fiber thickening and reinforcement [82, 86]. FAK plays an important role in the response of migrating cells to mechanical stress. FAK-null cells do not respond to external force by reorienting their migrations. Thus, FAK is involved in mechanosensing during cell migration [87]. External mechanical forces can influence the production of extracellular matrix components. For example, the expression of type XII collagen and tenascin, and also of matrix metalloproteinases (MMPs) by the cells can be stimulated by mechanical stress. This force-dependent regulation occurs at the level of transcription of specific genes [88]. The slackening of the centripetal stretch is observed in conditions of the deficiency of focal adhesions formation. This happens when there is deficiency or irregular distribution of the isolated very small adhesive “islets” of the extracellular matrix. On such “islet-like” matrix, the cells cannot completely spread (Fig. 5.11a, b), and the centripetal stretch forces, respectively, are weakened. This leads to the inhibition of cell proliferation and even, in certain cases, to apoptosis (anoikis). The synthetic processes also are altered: in fibroblastic cells the synthesis of vimentin is decreased; the synthesis of some proteases is increased in epitheliocytes [89]. Cells respond not only to the deformations of the extracellular matrix but also to its mechanical properties, such as rigidity, pliability, or tension state that determines the stiffness or relaxation of the extracellular matrix [76, 90, 91]. When matrix is under high tension, the spread fibroblastic cells acquire elongated shapes; they develop massive stress fibers and increased focal adhesions. In addition, fibroblastic cells can begin to express an isoform of actin, so-called a-smooth muscle actin (SMA). Exogenous mechanical forces can also increase SMA expression in fibroblastic cells [92]. This actin isoform is normally restricted to vascular smooth muscle cells, but it can also be expressed by special fibroblastic cells, called
5.3 Signaling Pathways from Integrin and Growth Factor Receptors in Normal Cells
107
Fig. 5.57 Myofibroblastic rat cell. Actin microfilaments containing a-smooth muscle actin and supermature focal contacts (long strips). The actin microfilaments stained for a-smooth muscle actin (green) and the focal contacts stained for vinculin (red). FM. Scale bar, 25 mm. Courtesy of V.B. Dugina
myofibroblasts, that are present in healing wounds and scars. Fibroblastic cells that express SMA are characterized by SMA-containing massive stress fibers and enlarged focal contacts. These large (“supermature”) focal contacts of 15–20 mm (Fig. 5.57) depend on high SMA-mediated contractility of stress fibers. SMA of filament bundles gives them strong contractility, and their anchorage to focal contacts induces the growth of these contacts. As a result, cells can generate significant traction forces that remodel the extracellular matrix. Owing to these mechanical forces, myofibroblasts promote the contraction of wounds favoring their healing [92–95]. The fibroblastic cells respond differently to the extracellular matrix that is in low-tension state, i.e., the relaxed matrix. In this case, cells do not form their stress fibers, few focal adhesions are formed, and cells stop synthesizing matrix collagen. In addition, these cells spread atypically forming dendrite-like extensions. The process of fibroblastic cell spreading under the relaxed matrix conditions is strongly dependent on microtubules: their disruption by nocodazole completely inhibits cell spreading, whereas on the high-tension matrix, microtubule disruption inhibits the fibroblastic cell polarization but not cell spreading. Therefore, microtubulemediated signaling may be pivotal for the spreading of fibroblastic cells in response to the relaxed state of the extracellular matrix [96]. Since integrins bound to the extracellular matrix mediate cell responses to it, they are considered as the transducers of not only biochemical signals but also
108
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
mechanical ones from the extracellular matrix. It is possible that the deformations or the mechanical properties of the matrix change the conformations of intracellular domains of integrins, initiating intracellular transfer of the signals. Actin cytoskeleton participates in the intracellular transmission of external force-induced signals influencing cell behavior [97]. Cells, in turn, exert their mechanical influence on the extracellular matrix resulting in its remodeling. The matrix remodeling induced by fibroblastic cells can include matrix contraction, formation of thin folds, and orientation of matrix fibers. The remodeling depends on matrix tension state and also on GF stimulation. Fibroblastic cells remodel highly tense matrix using their acto-myosin activity: the centripetal stretch of stress fibers through focal contacts is transferred to the matrix increasing its stiffness. In the case of relaxed extracellular matrix, fibroblastic cells can possibly utilize their microtubule-dependent dendrite-like extensions (see above) for matrix contraction. In response to GF stimulation, fibroblastic cells can convert matrix tension state from relaxed to stiff [96]. Extracellular matrix remodeling occurs in regenerative processes (healing of wounds), in bone tissue formation, etc. A considerable remodeling influence on the matrix is exerted by myofibroblasts owing to the strong centripetal stretch developed by these cells. Thus, the centripetal stretch, experienced by cells spread on the extracellular matrix, serves as powerful signal regulator of cell shape and functional activity, and also influences the organization of the extracellular matrix itself.
5.4 Alterations in Integrin-Mediated Adhesion and Signaling in Transformed Cells Substantial abnormalities in the functioning of focal contacts both as adhesion structures and as transducers of intracellular signals are typical for transformed cells. Because of these alterations, transformed cells acquire their characteristic morphology, lose substratum dependence of cell proliferation, become less dependent on soluble GF, and avoid apoptosis (anoikis). The basis of all these changes is the action of oncogene products and inactivation of tumor suppressor genes.
5.4.1 Defective Adhesive Function Alterations in cell-extracellular matrix adhesion are the major factors defining invasive and migratory characteristics of cancer cells [98].
5.4 Alterations in Integrin-Mediated Adhesion and Signaling in Transformed Cells
109
The oncogenic transformation of a cell can result in attenuation or augmentation of expression of various integrin receptors and also in changes of their affinity to the ligands [99]. Decreased expression of a5b1 fibronectin-binding integrin is often observed in cancer cells leading to weakening of their adhesion to fibronectincontaining structures of the extracellular matrix. On the other hand, many types of invading cancer cells are characterized by increased expression of a6b1 or a6b4 integrin receptors to laminin isoforms laminin-332 and laminin-411 (previously known as laminin-5 and laminin-8, respectively) [100–103]. The epithelial-specific aVb6 integrin that is expressed only during wound healing and in cancer cells, regulates the expression of MMPs in invadopodia of cancer cells and thereby modulates cancer invasion [104]. In a special situation, tumor cells can express “unusual” integrins. The characteristic of cancer cells that survive in the suspended state, when they circulate in lymphatic or blood vessels, is the expression of leukocyte or platelet integrins. These integrins are responsible for cell–cell adhesion, and mediate leukocyte and platelet adhesion to vascular endothelium or indirect binding of platelets to each other [13, 14]. The expression of these integrins on the surface of circulating cancer cells provides their attachment to the vascular endothelium that is a necessary precondition for subsequent cancer cell penetration of the vascular wall and formation of metastatic cancer foci. Expression of platelet integrin by circulating cancer cells promotes the formation of mixed aggregates of cancer cells with platelets that also favors invasion [14]. Adhesion of circulating cancer cells to the vascular endothelium can be mediated not only by integrins but also by other families of intercellular transmembrane adhesion molecules, such as ones that have the Ig-like domains and also selectins [14, 105]. As a consequence of activating mutations or amplifications of proto-oncogenes encoding protein tyrosine kinases in normal cells, in particular, submembrane Src protein kinase [106, 107], these proto-oncogenes are converted into constitutively activated oncogenes. The oncogene products, oncoproteins, cause hyper-phosphorylation of various components of focal adhesions [108]. The hyper-phosphorylation of integrin b subunit cytoplasmic domain may prevent its binding to talin, inhibiting integrin activation (see Sect. 4.2.1) It leads to a strong reduction in the affinity of some types of integrins to their ligands in the extracellular matrix (e.g., to fibronectin), and to the loss of the capability of some integrins for clustering. This concerns, for example, a5b1 fibronectin receptor. The hyper-phosphorylation of vinculin leads to its altered binding to other proteins of focal contacts. The weakening of the bonds between focal contact proteins is accompanied by a reduction in the capability of actin filaments for their anchoring, which, in turn, weakens their centripetal stretch and prevents the formation of stress fibers. The relaxation of centripetal stretch and the deficiency in stress fibers formation prevent maturation of focal contacts and initiate their disassembly. The described abnormalities lead to the defective focal contact formation and to the weakening of the adhesion of transformed cells to specific components of the extracellular matrix. The reduction in the adhesion is aggravated by the fact that transformed cells partially or completely lose the capability for the synthesis of specific matrix proteins, for example, fibronectin or collagen.
110
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
5.4.2 Alterations in the Mitogenic Signal Transduction The abnormalities of focal contacts in transformed cells concern not only their adhesive functions, but also their functions as intracellular signal transducers. Key proteins participating in the sequential transfer of mitogenic signals from integrin receptors (Fig. 5.50) or from the receptors of soluble GF (Fig. 5.51) are encoded by proto-oncogenes. These proteins mediating all steps of the mitogenic signal transduction cascade include soluble GF (e.g., fibroblast GF, FGF-2, or platelet-derived GF, PDGF-b, etc.), their tyrosine kinase receptors (RTK) (e.g., receptors of EGF, or scatter factor, HGF/SF, etc.), FAK and Src protein kinases, Ras GTPases, phosphatidylinositol 3-kinase (PI3K), Raf protein kinases, MAP kinases, and a number of transcription factors. Because of the mutational conversion of proto-oncogenes into oncogenes, one or several key participants of the mitogenic signal transduction pathways are converted into oncoproteins and become constitutively super active [109]. On account of the constitutive activation of oncogenes encoding a series of soluble GF and their RTK receptors, the transformed cells acquire increased number of these receptors in a constitutively activated state, and the ability to overproduce some GF. Dysregulation of GF and their cell receptors plays a very important role in malignancy [110]. As a result of amplifications or activating mutations of proto-oncogenes encoding receptors of such GF as EGF, FGF, PDGF, or HGF/SF, these receptors are overexpressed in various types of human cancers [110–115]. For example, EGF receptor is the product of proto-oncogene c-ErbB1/HER1; the HGF/SF receptor is the product of proto-oncogene c-met. Proto-oncogenes are turned into permanently activated oncogenes to stimulate cancer cell proliferation. Thus, transformed cells themselves can stimulate their proliferation, and they can proliferate in the presence of very low concentrations of GF. FAK as well FAK-regulating signal pathways play critical roles in tumor initiation and progression [116, 117]. In transformed cells, FAK can be constantly auto-phosphorylated and submembrane Src protein kinase can become constantly activated. As a result, proteins of the Ras family will undergo constitutive phosphorylation and be in the state of permanent activation. In turn, proto-oncogene Ras can be mutated and converted into the oncogene; as a result, Ras oncoproteins will be constantly in the active state, transferring intracellular signals further [118]. Cytoplasmic protein kinases of Raf family or kinases of MAP cascades can become constantly activated without stimulation. Because of the mutations or the amplifications of proto-oncogenes encoding transcription factors, the number and the activity of these factors will increase, including those that stimulate specific genes initiating cell proliferation. Increased FAK expression and activity are observed in multiple human cancers, and correlate with progression and metastasis in cancer patients [117, 119, 120]. Src kinase is also overexpressed or highly activated in many types of human cancers
5.4 Alterations in Integrin-Mediated Adhesion and Signaling in Transformed Cells
111
[106, 107]. Hyperactivated FAK-Src complex permanently generates signals leading to cancer growth and invasion [45]. Ras proto-oncogene mutations are found in more than 20% of human malignant tumors. The inhibitors of hyperactivated mitogenic signaling pathways in cancer cells can have a potential for human cancer therapy. Key components of these mitogenic pathways, such as FAK or Src, are considered as promising targets for inhibition [106, 107, 121–123]. Thus, mitogenic signal transduction pathways are permanently activated in transformed cells. They may function autonomously, independently of integrin receptor binding to the extracellular matrix or binding of soluble GF to their cell surface receptors.
5.4.3 The “Anchorage Independence” The detachment of transformed cells from the extracellular matrix does not inhibit their proliferation and does not induce their apoptosis (anoikis). In contrast to normal cells, in detached transformed cells, mitogenic signals from GF receptors are not blocked, and the cells retain their ability to proliferate in the absence of focal adhesions. Besides, in transformed cells negative regulators of cell proliferation, p53, and p27/ KIP1 tumor suppressor genes are often inactivated because of their mutations or deletions. Therefore, the transformed cell detachment does not cause blocking of cell proliferation. The detached transformed cells avoid apoptosis (anoikis) [50, 124, 125]. This is achieved because of mutagenic inactivation of the pro-apoptotic p53 tumor suppressor gene (Fig. 5.52) and/or because of constant transduction of antiapoptotic signals from integrins (Fig. 5.50) irrespective of their binding to the extracellular matrix. Because of p53 inactivation, detached transformed cells do not undergo anoikis and continue to proliferate. The inactivating mutations or deletions of p53 are characteristic of transformed cells. Inactivation of p53 is revealed in approximately 60% of different human tumors. This inactivation leads to the accumulation of permanently proliferating cells with different DNA damage patterns that is a characteristic feature of tumor cells. Another reason for the suppression of anoikis in transformed cells is the permanent activation of Ras-PI3K-PKB/Akt antiapoptotic signal transduction pathway (Fig. 5.52). This signaling pathway in normal cells protects them from apoptosis, and PTEN protein encoded by the tumor suppressor gene inhibits antiapoptotic effect of that signaling pathway (see Sect. 5.3.3). In transformed cells, because of permanent activation of Ras proteins, PI3K and PKB/Akt, and also because of PTEN gene mutagenic inactivation, the antiapoptotic signaling pathway is permanently activated. Activating mutations of genes encoding PI3K and PKB/Akt were found in many human cancers. The mutagenic inactivation of the apoptotic PTEN tumor suppressor gene is observed in 60% of all forms of human cancers [38, 126, 127].
112
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
The avoidance of anoikis ensures survival of malignant tumor cells under the conditions of their circulation in the lymph or the blood flow. Thus, transformed cells lose substratum dependence of cell proliferation and acquire anchorage independence: these cells can survive and proliferate without being attached to the extracellular matrix (Fig. 5.53). In semiliquid medium, many types of suspended transformed cells can proliferate and do not undergo anoikis. The independence of survival and growth of transformed cells from their attachment to the extracellular matrix (anchorage independence) and from the actions of soluble GF is reached by: (a) Permanent activation of intracellular mitogenic signaling pathways. (b) Abolition of the blocking of mitogenic signal transduction in cells detached from the extracellular matrix. (c) Inactivation of apoptotic tumor suppressor genes. (d) Permanent activation of antiapoptotic signal transduction pathways. Because of the deficient formation and functions of focal adhesions and connected stress fibers, the isometric tension of transformed cells is relaxed, and the degree of their spreading on the extracellular matrix is reduced. Transformed cells acquire the characteristic “compact” morphology (Fig. 5.34). Also, transformed cells have altered regulation of their locomotion (see Sect. 6.2). Thus, as a result of the changes of intracellular signal transduction pathways caused by permanent activity of oncogenes and the inactivation of tumor suppressor genes, transformed cells acquire the ability for permanent uncontrollable proliferation. They become “anchorage independent” and less dependent on the soluble GF. Transformed cells can avoid anoikis. They acquire the characteristic morphology and also high locomotory activity. Specialized adhesion structures, invadopodia, that are characteristic of aggressive cancer cells play an important role in their capability for invasion into surrounding healthy tissues and for metastasis [105, 128, 129]. The local extracellular matrix degradation is apparently achieved by MMP activity associated with invadopodia. This activity creates increased space for cells to migrate and favors cancer invasion; furthermore, MMP activity plays a role in the induction and progression of tumor angiogenesis [130–132]. Key enzyme in the proteolytic activity of invadopodia is membrane type 1-matrix metalloproteinase (MT1-MMP, or MMP-14). It is a transmembrane enzyme that is localized at the leading edges of invading cancer cells [133]. One of the major extracellular matrix targets of MT1-MMP is type I collagen. Besides, MT1-MMP binds to and activates pro-MMP-2 that then matures into the active MMP-2. The latter enzyme can degrade type IV collagen in basement membranes. The rigidity of the extracellular matrix increases both the number and MMP activity of invadopodia in invading cancer cells [134]. In some cases, however, MMPs may have paradoxical effect on cancer progression: for example, inhibiting mutations of the gene encoding MMP-8 stimulates the progression of human melanoma [135].
References
113
References 1. Rovensky YuA, Vasiliev JuM (1984) Surface topography of suspended tissue cells. Int Rev Cytol 90:273–307. doi:10.1016/S0074-7696(08)61492-8 DOI:dx.doi.org 2. Rovensky YA, Komissarova EV, Topol LZ, Kisseljov FL (1992) Changes in surface relief of suspended cells are morphological signs of the initial stage of neoplastic transformation in fibroblastic monolayer cultures. Cell Biol Int Rep 16(6):557–65. doi:10.1016/S03091651(05)80054-2 DOI:dx.doi.org 3. Vasiliev JM (1985) Spreading of non-transformed and transformed cells. Biochim Biophys Acta 780(1):21–65. 4. Kharitonova MA, Vasiliev JM (2008) Controlling cell length. Semin Cell Dev Biol. 19(6): 480–484. doi:10.1016/j.semcdb.2008.07.008 DOI:dx.doi.org 5. Kharitonova MA, Kopnin PB, Vasiliev JM (2007) Transformation by RAS oncogene decreases the width of substrate-spread fibroblasts but not their length. Cell Biol Int 31(3):220–223. doi:10.1016/j.cellbi.2006.10.008 DOI:dx.doi.org 6. Charras GT (2008) A short history of blebbing. J Microsc 231(3):466–478. doi:10.1111/ j.1365-2818.2008.02059.x DOI:dx.doi.org 7. Johnson LN (2009) The regulation of protein phosphorylation. Biochem Soc Trans 37 (Pt 4):627–41. doi:10.1042/BST0370627 DOI:dx.doi.org 8. Li E, Hristova K (2006) Role of receptor tyrosine kinase transmembrane domains in cell signaling and human pathologies. Biochemistry 45(20):6241–6251. doi:10.1021/bi060609y DOI:dx.doi.org 9. Orth JD, McNiven MA (2006) Get off my back! Rapid receptor internalization through circular dorsal ruffles. Cancer Res 66(23):11094–11096. doi:10.1158/0008-5472.CAN-06-3397 DOI:dx.doi.org 10. Lo SH (2006) Focal adhesions: what’s new inside. Dev Biol 294(2):280–291. doi:10.1016/ j.ydbio.2006.03.029 DOI:dx.doi.org 11. Schwartz MA, Ginsberg MH (2002) Networks and crosstalk: integrin signalling spreads. Nat Cell Biol 4(4):E65–8. doi:10.1038/ncb0402-e65 DOI:dx.doi.org 12. Legate KR, Wickström SA, Fässler R (2009) Genetic and cell biological analysis of integrin outside-in signaling. Genes Dev 23(4):397–418. doi:10.1101/gad.1758709 DOI:dx.doi.org 13. Bennett JS, Berger BW, Billings PC (2009) The structure and function of platelet integrins. J Thromb Haemost 7 Suppl 1:200–205. doi:10.1111/j.1538-7836.2009.03378.x DOI:dx.doi.org 14. Rovensky YA (1998) Cellular and molecular mechanisms of tumor invasion. Biochemistry (Mosc) 63(9):1029–1043. 15. Streuli CH, Akhtar N (2009) Signal co-operation between integrins and other receptor systems. Biochem J 418(3):491–506. doi:10.1042/BJ20081948 DOI:dx.doi.org 16. Ivaska J, Heino J (2010) Interplay between cell adhesion and growth factor receptors: from the plasma membrane to the endosomes. Cell Tissue Res 339(1):111–120. doi:10.1007/ s00441-009-0857-z DOI:dx.doi.org 17. Rosenbaum DM, Rasmussen SG, Kobilka BK (2009) The structure and function of G-proteincoupled receptors. Nature 459(7245):356–363. doi:10.1038/nature08144 DOI:dx.doi.org 18. Millar RP, Newton CL (2010) The year in G protein-coupled receptor research. Mol Endocrinol 24(1):261–274 19. Cotton M, Claing A (2009) G protein-coupled receptors stimulation and the control of cell migration. Cell Signal 21(7):1045–1053. doi:10.1016/j.cellsig.2009.02.008 DOI:dx.doi.org 20. Lundquist EA (2006) Small GTPases. WormBook 17: 1–18. doi:10.1895/wormbook.1.67.1 DOI:dx.doi.org 21. Skwarek LC, Boulianne GL (2009) Great expectations for PIP: phosphoinositides as regulators of signaling during development and disease. Dev Cell 16(1):12–20. doi:10.1016/j. devcel.2008.12.006 DOI:dx.doi.org
114
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
22. Saarikangas J, Zhao H, Lappalainen P (2010) Regulation of the actin cytoskeleton-plasma membrane interplay by phosphoinositides. Physiol Rev 90(1):259–289. doi:10.1152/physrev.00036.2009 DOI:dx.doi.org 23. Vicinanza M, D’Angelo G, Di Campli A, De Matteis MA (2008) Function and dysfunction of the PI system in membrane trafficking. EMBO J 27(19):2457–2470. doi:10.1038/ emboj.2008.169 DOI:dx.doi.org 24. Saito K, Tautz L, Mustelin T (2007) The lipid-binding SEC14 domain. Biochim Biophys Acta 1771(6):719–726. doi:10.1016/j.bbalip.2007.02.010 DOI:dx.doi.org 25. Krauss M, Haucke V (2007) Phosphoinositides: regulators of membrane traffic and protein function. FEBS Lett 581(11):2105–2111. doi:10.1016/j.febslet.2007.01.089 DOI:dx.doi.org 26. Sasaki T, Sasaki J, Sakai T, Takasuga S, Suzuki A (2007) The physiology of hosphoinositides. Biol Pharm Bull 30(9):1599–1604. doi:10.1248/bpb.30.1599 DOI:dx.doi.org 27. Oude Weernink PA, Han L, Jakobs KH, Schmidt M (2007) Dynamic phospholipid signaling by G protein-coupled receptors. Biochim Biophys Acta 1768(4):888–900. doi:10.1007/ s00210-007-0131-4 DOI:dx.doi.org 28. Oude Weernink PA, López de Jesús M, Schmidt M (2007) Phospholipase D signaling: orchestration by PIP2 and small GTPases. Naunyn Schmiedebergs Arch Pharmacol 374(5-6): 399–411. doi:10.1007/s00210-007-0131-4 DOI:dx.doi.org 29. Heck JN, Mellman DL, Ling K, Sun Y, Wagoner MP, Schill NJ, Anderson RA (2007) A conspicuous connection: structure defines function for the phosphatidylinositol-phosphate kinase family. Crit Rev Biochem Mol Biol 42(1):15–39. doi:10.1080/10409230601162752 DOI:dx.doi.org 30. Kanaho Y, Kobayashi-Nakano A, Yokozeki T (2007) The phosphoinositide kinase PIP5K that produces the versatile signaling phospholipid PI4,5P(2). Biol Pharm Bull 30(9):1605–1609. doi:10.1248/bpb.30.1605 DOI:dx.doi.org 31. Santarius M, Lee CH, Anderson RA (2006) Supervised membrane swimming: small G-protein lifeguards regulate PIPK signalling and monitor intracellular PtdIns(4,5)P2 pools. Biochem J 398(1):1–13. doi:10.1042/BJ20060565 DOI:dx.doi.org 32. Di Paolo G, De Camilli P (2006) Phosphoinositides in cell regulation and membrane dynamics. Nature 443 (7112):651–657. doi:10.1038/nature05185 DOI:dx.doi.org 33. Mao YS, Yin HL (2007) Regulation of the actin cytoskeleton by phosphatidylinositol 4-phosphate 5 kinases. Pflugers Arch 455(1):5–18. doi:10.1007/s00424-007-0286-3 DOI:dx.doi.org 34. Kölsch V, Charest PG, Firtel RA (2008) The regulation of cell motility and chemotaxis by phospholipid signaling. J Cell Sci 121(Pt 5):551–559 35. Ling K, Schill NJ, Wagoner MP, Sun Y, Anderson RA (2006) Movin’ on up: the role of PtdIns(4,5)P(2) in cell migration. Trends Cell Biol 16(6):276–284. doi:10.1016/ j.tcb.2006.03.007 DOI:dx.doi.org 36. Martelli AM, Cocco L, Capitani S, Miscia S, Papa S, Manzoli FA (2007) Nuclear phosphatidylinositol 3,4,5-trisphosphate, phosphatidylinositol 3-kinase, Akt, and PTen: emerging key regulators of anti-apoptotic signaling and carcinogenesis. Eur J Histochem 51 Suppl 1:125–131 37. Kumar A, Carrera AC (2007) New functions for PI3K in the control of cell division. Cell Cycle 6(14):1696–1698. doi:10.4161/cc.6.14.4492 DOI:dx.doi.org 38. Carnero A, Blanco-Aparicio C, Renner O, Link W, Leal JF (2008) The PTEN/PI3K/AKT signalling pathway in cancer, therapeutic implications. Curr Cancer Drug Targets 8(3): 187–198. doi:10.2174/156800908784293659 DOI:dx.doi.org 39. Goto K, Hozumi Y, Nakano T, Saino-Saito S, Martelli AM (2008) Lipid messenger, diacylglycerol, and its regulator, diacylglycerol kinase, in cells, organs, and animals: history and perspective. Tohoku J Exp Med 214(3):199–212. doi:10.1620/tjem.214.199 DOI:dx.doi.org 40. Larsson C (2006) Protein kinase C and the regulation of the actin cytoskeleton. Cell Signal 18(3):276–284. doi:10.1016/j.cellsig.2005.07.010 DOI:dx.doi.org 41. Rosse C, Linch M, Kermorgant S, Cameron AJ, Boeckeler K, Parker PJ (2010) PKC and the control of localized signal dynamics. Nat Rev Mol Cell Biol 11(2):103–112. doi:10.1038/ nrm2847 DOI:dx.doi.org 42. Arnaout MA, Goodman SL, Xiong JP (2007) Structure and mechanics of integrin-based cell adhesion. Curr Opin Cell Biol 19(5):495–507. doi:10.1016/j.ceb.2007.08.002 DOI:dx.doi.org
References
115
43. Huveneers S, Danen EH (2009) Adhesion signaling – crosstalk between integrins, Src and Rho. J Cell Sci 122(Pt 8):1059–1069. doi:10.1242/jcs.039446 DOI:dx.doi.org 44. Gan B, Guan JL (2008) FIP200, a key signaling node to coordinately regulate various cellular processes. Cell Signal 20(5):787–794. doi:10.1016/j.cellsig.2007.10.021 DOI:dx.doi.org 45. Mitra SK, Schlaepfer DD (2006) Integrin-regulated FAK-Src signaling in normal and cancer cells. Curr Opin Cell Biol 18(5):516–523. doi:10.1016/j.ceb.2006.08.011 DOI:dx.doi.org 46. Juliano RL, Reddig P, Alahari S, Edin M, Howe A, Aplin A (2004) Integrin regulation of cell signalling and motility. Biochem Soc Trans 32(Pt3):443–446. doi:10.1042/BST0320443 DOI:dx.doi.org 47. Vasiliev JM (2004) Cytoskeletal mechanisms responsible for invasive migration of neoplastic cells. Int J Dev Biol 48(5-6):425–439. doi:10.1387/ijdb.041806jv DOI:dx.doi.org 48. Howe AK, Aplin AE, Juliano RL (2002) Anchorage-dependent ERK signaling – mechanisms and consequences. Curr Opin Genet Dev 12(1):30–35. doi:10.1016/S0959-437X(01)00260-X DOI:dx.doi.org 49. Gilmore AP, Owens TW, Foster FM, Lindsay J (2009) How adhesion signals reach a mitochondrial conclusion – ECM regulation of apoptosis. Curr Opin Cell Biol 21(5):654–661. doi:10.1016/j.ceb.2009.05.009 DOI:dx.doi.org 50. Cotter TG (2009) Apoptosis and cancer: the genesis of a research field. Nat Rev Cancer 9(7):501–507. doi:10.1038/nrc2663 DOI:dx.doi.org 51. Teng CS (2000) Protooncogenes as mediators of apoptosis. Int Rev Cytol 197:137–202. doi:10.1016/S0074-7696(00)97004-9 DOI:dx.doi.org 52. Grossmann J (2002) Molecular mechanisms of “detachment-induced apoptosis – Anoikis”. Apoptosis 7(3):247–260. doi:10.1023/A:1015312119693 DOI:dx.doi.org 53. Stambolic V, Woodgett JR (2006) Functional distinctions of protein kinase B/Akt isoforms defined by their influence on cell migration. Trends Cell Biol 16(9):461–466. doi:10.1016/ j.tcb.2006.07.001 DOI:dx.doi.org 54. Chen RH, Wang WJ, Kuo JC (2006) The tumor suppressor DAP-kinase links cell adhesion and cytoskeleton reorganization to cell death regulation. J Biomed Sci 13(2):193–199. doi:10.1007/s11373-005-9063-5 DOI:dx.doi.org 55. Echarri A, Del Pozo MA (2006) Caveolae internalization regulates integrin-dependent signaling pathways. Cell Cycle 5(19):2179–2182. doi:10.4161/cc.5.19.3264 DOI:dx.doi.org 56. Salanueva IJ, Cerezo A, Guadamillas MC, del Pozo MA (2007) Integrin regulation of caveolin function. J Cell Mol Med 11(5):969-980. doi:10.1111/j.1582-4934.2007.00109.x DOI:dx.doi.org 57. Del Pozo MA, Schwartz MA (2007) Rac, membrane heterogeneity, caveolin and regulation of growth by integrins. Trends Cell Biol 17(5):246–250 58. Lee J, Kim SS (2009) The function of p27 KIP1 during tumor development. Exp Mol Med 41(11):765–771 59. Chiarugi P, Giannoni E (2008) Anoikis: a necessary death program for anchorage-dependent cells. Biochem Pharmacol 76(11):1352–1364. doi:10.1016/j.bcp.2008.07.023 DOI:dx.doi.org 60. Gilmore AP (2005) Anoikis. Cell Death Differ 12 Suppl 2:1473–1477. doi:10.1038/ sj.cdd.4401723 DOI:dx.doi.org 61. Reddig PJ, Juliano RL (2005) Clinging to life: cell to matrix adhesion and cell survival. Cancer Metastasis Rev 24(3):425–439. doi:10.1007/s10555-005-5134-3 DOI:dx.doi.org 62. Lock R, Debnath J (2008) Extracellular matrix regulation of autophagy. Curr Opin Cell Biol 20(5):583–588. doi:10.1016/j.ceb.2008.05.002 DOI:dx.doi.org 63. Arias-Salgado EG, Lizano S, Shattil SJ, Ginsberg MH (2005) Specification of the direction of adhesive signaling by the integrin beta cytoplasmic domain. J Biol Chem 280(33):29699– 29707. doi:10.1074/jbc.M503508200 DOI:dx.doi.org 64. Hall A (2005) Rho GTPases and the control of cell behaviour. Biochem Soc Trans 33 (Pt 5):891–895. doi:10.1042/BST20050891 DOI:dx.doi.org 65. Ridley AJ (2006) Rho GTPases and actin dynamics in membrane protrusions and vesicle trafficking. Trends Cell Biol 16(10):522–529. doi:10.1016/j.tcb.2006.08.006 DOI:dx.doi.org 66. Narumiya S, Tanji M, Ishizaki T (2009) Rho signaling, ROCK and mDia1, in transformation, metastasis and invasion. Cancer Metastasis Rev 28(1-2):65–76. doi:10.1007/s10555-0089170-7 DOI:dx.doi.org
116
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
67. Oser M. Condeelis J (2009) The cofilin activity cycle in lamellipodia and invadopodia. J Cell Biochem 108(6):1252–1262. doi:10.1002/jcb.22372 DOI:dx.doi.org 68. Scott RW, Olson MF (2007) LIM kinases: function, regulation and association with human disease. J Mol Med 85(6):555–568. doi:10.1007/s00109-007-0165-6 DOI:dx.doi.org 69. Bernard O (2007) Lim kinases, regulators of actin dynamics. Int J Biochem Cell Biol 39(6):1071–1076. doi:10.1016/j.biocel.2006.11.011 DOI:dx.doi.org 70. Pollitt AY, Insall RH (2009) WASP and SCAR/WAVE proteins: the drivers of actin assembly. J Cell Sci 122(Pt 15):2575–2578. doi:10.1242/jcs.023879 DOI:dx.doi.org 71. Tomar A, Schlaepfer DD (2009) Focal adhesion kinase: switching between GAPs and GEFs in the regulation of cell motility. Curr Opin Cell Biol 21(5):676–683. doi:10.1016/ j.ceb.2009.05.006 DOI:dx.doi.org 72. Mammoto A, Ingber DE (2009) Cytoskeletal control of growth and cell fate switching. Curr Opin Cell Biol 21(6):864–870. doi:10.1016/j.ceb.2009.08.001 DOI:dx.doi.org 73. Bershadsky AD, Ballestrem C, Carramusa L, Zilberman Y, Gilquin B, Khochbin S, Alexandrova AY, Verkhovsky AB, Shemesh T, Kozlov MM (2006) Assembly and mechanosensory function of focal adhesions: experiments and models. Eur J Cell Biol 85(3-4):165– 173. doi:10.1016/j.ejcb.2005.11.001 DOI:dx.doi.org 74. Schwartz MA, DeSimone DW (2008) Cell adhesion receptors in mechanotransduction. Curr Opin Cell Biol 20(5):551–556. doi:10.1016/j.ceb.2008.05.005 DOI:dx.doi.org 75. Chiquet M, Gelman L, Lutz R, Maier S (2009) From mechanotransduction to extracellular matrix gene expression in fibroblasts. Biochim Biophys Acta 1793(5):911–920. doi:10.1016/ j.bbamcr.2009.01.012 DOI:dx.doi.org 76. Harjanto D, Zaman MH (2010) Matrix mechanics and receptor-ligand interactions in cell adhesion. Org Biomol Chem 8(2):299–304. doi:10.1039/b913064k DOI:dx.doi.org 77. Baker EL, Zaman MH (2010) The biomechanical integrin. J Biomech 43(1):38–44. doi:10.1016/j.jbiomech.2009.09.007 DOI:dx.doi.org 78. Bershadsky AD, Balaban NQ, Geiger B (2003) Adhesion-dependent cell mechanosensitivity. Annu Rev Cell Dev Biol 19:677–695. doi:10.1146/annurev.cellbio.19.111301.153011 DOI:dx.doi.org 79. Shemesh T, Geiger B, Bershadsky AD, Kozlov MM (2005) Focal adhesions as mechanosensors: a physical mechanism. Proc Natl Acad Sci U S A 102(35):12383–12388. doi:10.1073/ pnas.0500254102 DOI:dx.doi.org 80. Wolfenson H, Henis YI, Geiger B, Bershadsky AD (2009) The heel and toe of the cell’s foot: a multifaceted approach for understanding the structure and dynamics of focal adhesions. Cell Motil Cytoskeleton 66(11):1017–1029. doi:10.1002/cm.20410 DOI:dx.doi.org 81. Critchley DR (2005) Genetic, biochemical and structural approaches to talin function. Biochem Soc Trans 33(Pt 6):1308–1312. doi:10.1042/BST20051308 DOI:dx.doi.org 82. Hirata H, Tatsumi H, Sokabe M (2008) Mechanical forces facilitate actin polymerization at focal adhesions in a zyxin-dependent manner. J Cell Sci 121(Pt 17):2795–2804. doi:10.1242/ jcs.030320 DOI:dx.doi.org 83. Trichet L, Sykes C, Plastino J (2008) Relaxing the actin cytoskeleton for adhesion and movement with Ena/VASP. J Cell Biol 181(1):19–25. doi:10.1083/jcb.200710168 DOI:dx. doi.org 84. Bear JE, Gertler FB (2009) Ena/VASP: towards resolving a pointed controversy at the barbed end. J Cell Sci 122(Pt 12):1947–1953. doi:10.1242/jcs.038125 DOI:dx.doi.org 85. Wang Y, Gilmore TD (2003) Zyxin and paxillin proteins: focal adhesion plaque LIM domain proteins go nuclear. Biochim Biophys Acta 1593(2-3):115–120. doi:10.1016/S01674889(02)00349-X DOI:dx.doi.org 86. Hirata H, Tatsumi H, Sokabe M (2008) Zyxin emerges as a key player in the mechanotransduction at cell adhesive structures. Commun Integr Biol 1(2):192–195. doi:10.4161/ cib.1.2.7001 DOI:dx.doi.org 87. Tilghman RW, Parsons JT (2008) Focal adhesion kinase as a regulator of cell tension in the progression of cancer. Semin Cancer Biol 18(1):45–52. doi:10.1016/j.semcancer.2007.08.002 DOI:dx.doi.org
References
117
88. Chiquet M, Tunç-Civelek V, Sarasa-Renedo A (2007) Gene regulation by mechanotransduction in fibroblasts. Appl Physiol Nutr Metab 32(5):967–973. doi:10.1139/H07-053 DOI:dx. doi.org 89. Dike LE, Chen CS, Mrksich M, Tien J, Whitesides GM, Ingber DE (1999) Geometric control of switching between growth, apoptosis, and differentiation during angiogenesis using micropatterned substrates. In Vitro Cell Dev Biol Anim 35(8):441–448. doi:10.1007/s11626-9990050-4 DOI:dx.doi.org 90. Plant AL, Bhadriraju K, Spurlin TA, Elliott JT (2009) Cell response to matrix mechanics: focus on collagen. Biochim Biophys Acta 1793(5):893–902. doi:10.1016/j.bbamcr.2008.10.012 DOI:dx.doi.org 91. Geiger B, Spatz JP, Bershadsky AD (2009) Environmental sensing through focal adhesions. Nat Rev Mol Cell Biol 10(1):21–33. doi:10.1038/nrm2593 DOI:dx.doi.org 92. Wang J, Zohar R, McCulloch CA (2006) Multiple roles of alpha-smooth muscle actin in mechanotransduction. Exp Cell Res 312(3):205–214. doi:10.1016/j.yexcr.2005.11.004 DOI:dx.doi.org 93. Hinz B, Celetta G, Tomasek JJ, Gabbiani G, Chaponnier C (2001) Alpha-smooth muscle actin expression upregulates fibroblast contractile activity. Mol Biol Cell 12(9):2730–2741 94. Dugina V, Fontao L, Chaponnier C, Vasiliev J, Gabbiani G (2001) Focal adhesion features during myofibroblastic differentiation are controlled by intracellular and extracellular factors. J Cell Sci 114(Pt18):3285–3296 95. Hinz B, Dugina V, Ballestrem C, Wehrle-Haller B, Chaponnier C (2003) Alpha-smooth muscle actin is crucial for focal adhesion maturation in myofibroblasts. Mol Biol Cell 14(6): 2508–2519. doi:10.1091/mbc.E02-11-0729 DOI:dx.doi.org 96. Rhee S (2009) Fibroblasts in three dimensional matrices: cell migration and matrix remodeling. Exp Mol Med 41(12):858–865. doi:10.3858/emm.2009.41.12.096 DOI:dx.doi.org 97. Fletcher DA, Mullins RD (2010) Cell mechanics and the cytoskeleton. Nature 463(7280): 485–492. doi:10.1038/nature08908 DOI:dx.doi.org 98. Makrilia N, Kollias A, Manolopoulos L, Syrigos K (2009) Cell adhesion molecules: role and clinical significance in cancer. Cancer Invest 27(10):1023–1037. doi:10.3109/07357900902 769749 DOI:dx.doi.org 99. Desgrosellier JS, Cheresh DA (2010) Integrins in cancer: biological implications and therapeutic opportunities. Nat Rev Cancer 10(1):9–22. doi:10.1038/nrc2748 DOI:dx.doi.org 100. Ljubimova JY, Fujita M, Khazenzon NM, Ljubimov AV, Black KL (2006) Changes in laminin isoforms associated with brain tumor invasion and angiogenesis. Front Biosci 11:81–88. doi:10.2741/1781 DOI:dx.doi.org 101. Marinkovich MP (2007) Tumour microenvironment: laminin 332 in squamous-cell carcinoma. Nat Rev Cancer 7(5):370–380. doi:10.1038/nrc2089 DOI:dx.doi.org 102. Fujita M, Khazenzon NM, Ljubimov AV, Lee BS, Virtanen I, Holler E, Black KL, Ljubimova JY (2006) Inhibition of laminin-8 in vivo using a novel poly(malic acid)-based carrier reduces glioma angiogenesis. Angiogenesis 9(4):183–191. doi:10.1007/s10456-006-9046-9 DOI:dx.doi.org 103. Tsuruta D, Kobayashi H, Imanishi H, Sugawara K, Ishii M, Jones JC (2008) Laminin-332integrin interaction: a target for cancer therapy? Curr Med Chem 15(20):1968–1975. doi:10.2174/092986708785132834 DOI:dx.doi.org 104. Bandyopadhyay A, Raghavan S (2009) Defining the role of integrin alphavbeta6 in cancer. Curr Drug Targets 10(7):645–652 105. Brooks SA, Lomax-Browne HJ, Carter TM, Kinch CE, Hall DM (2010) Molecular interactions in cancer cell metastasis. Acta Histochem 112(1):3–25. doi:10.1016/j.acthis.2008.11.022 DOI:dx.doi.org 106. Kim LC, Song L, Haura EB (2009) Src kinases as therapeutic targets for cancer. Nat Rev Clin Oncol 6(10):587–595. doi:10.1038/nrclinonc.2009.129 DOI:dx.doi.org 107. Guarino M (2010) Src signaling in cancer invasion. J Cell Physiol 223(1):14–26. doi:10.1002/ jcp.22011 DOI:dx.doi.org 108. Hood JD, Cheresh DA (2002) Role of integrins in cell invasion and migration. Nat Rev Cancer 2(2):91–100. doi:10.1038/nrc727 DOI:dx.doi.org
118
5 Adhesive Interactions of Tissue Cells with the Extracellular Matrix
109. Blume-Jensen P, Hunter T (2001) Oncogenic kinase signalling. Nature 411(6835):355–365. doi:10.1038/35077225 DOI:dx.doi.org 110. Zhang X, Nie D, Chakrabarty S (2010) Growth factors in tumor microenvironment. Front Biosci 15:151–165. doi:10.2741/3612 DOI:dx.doi.org 111. Hanna JA, Bordeaux J, Rimm DL, Agarwal S (2009) The function, proteolytic processing, and histopathology of Met in cancer. Adv Cancer Res 103:1–23. doi:10.1016/S0065230X(09)03001-2 DOI:dx.doi.org 112. Appelmann I, Liersch R, Kessler T, Mesters RM, Berdel WE (2010) Angiogenesis inhibition in cancer therapy: platelet-derived growth factor (PDGF) and vascular endothelial growth factor (VEGF) and their receptors: biological functions and role in malignancy. Recent Results Cancer Res 180:51–81. doi:10.1007/978-3-540-78281-0_5 DOI:dx.doi.org 113. Mitsudomi T, Yatabe Y (2010) Epidermal growth factor receptor in relation to tumor development: EGFR gene and cancer. FEBS J 277(2):301–308. doi:10.1111/j.1742-4658.2009.07448.x DOI:dx.doi.org 114. Knights V, Cook SJ (2010) De-regulated FGF receptors as therapeutic targets in cancer. Pharmacol Ther 125(1):105–117. doi:10.1016/j.pharmthera.2009.10.001 DOI:dx.doi.org 115. Lu X, Kang Y (2010) Epidermal growth factor signalling and bone metastasis. Br J Cancer 102(3):457–461. doi:10.1038/sj.bjc.6605490 DOI:dx.doi.org 116. Provenzano PP, Keely PJ (2009) The role of focal adhesion kinase in tumor initiation and progression. Cell Adh Migr. 3(4):347–350. doi:10.4161/cam.3.4.9458 DOI:dx.doi.org 117. Zhao J, Guan JL (2009) Signal transduction by focal adhesion kinase in cancer. Cancer Metastasis Rev 28(1-2):35–49. doi:10.1007/s10555-008-9165-4 DOI:dx.doi.org 118. DeNicola GM, Tuveson DA (2009) RAS in cellular transformation and senescence. Eur J Cancer 45 Suppl 1:211–216. doi:10.1016/S0959-8049(09)70036-X DOI:dx.doi.org 119. Chatzizacharias NA, Kouraklis GP, Theocharis SE (2008) Clinical significance of FAK expression in human neoplasia. Histol Histopathol 23(5):629–650 120. Luo M, Guan JL (2010) Focal adhesion kinase: a prominent determinant in breast cancer initiation, progression and metastasis. Cancer Lett 289(2):127–139. doi:10.1016/j.canlet.2009.07.005 DOI:dx.doi.org 121. Hao H, Naomoto Y, Bao X, Watanabe N, Sakurama K, Noma K, Motoki T, Tomono Y, Fukazawa T, Shirakawa Y, Yamatsuji T, Matsuoka J, Wang ZG, Takaoka M (2009) Focal adhesion kinase as potential target for cancer therapy (Review). Oncol Rep 22(5):973–979 122. Schwock J, Dhani N, Hedley DW (2010) Targeting focal adhesion kinase signaling in tumor growth and metastasis. Expert Opin Ther Targets 14(1):77–94. doi:10.1517/14728220903460340 DOI:dx.doi.org 123. Wong KK, Engelman JA, Cantley LC (2010) Targeting the PI3K signaling pathway in cancer. Curr Opin Genet Dev 20(1):87–90. doi:10.1016/j.gde.2009.11.002 DOI:dx.doi.org 124. Simpson CD, Anyiwe K, Schimmer AD (2008) Anoikis resistance and tumor metastasis. Cancer Lett 272(2):177–185. doi:10.1016/j.canlet.2008.05.029 DOI:dx.doi.org 125. Westhoff MA, Fulda S (2009) Adhesion-mediated apoptosis resistance in cancer. Drug Resist Updat. 12(4-5):127–136. doi:10.1016/j.drup.2009.08.001 DOI:dx.doi.org 126. Ming M, He YY (2009) PTEN: new insights into its regulation and function in skin cancer. J Invest Dermatol 129(9):2109–2112. doi:10.1038/jid.2009.79 DOI:dx.doi.org 127. Alexiou GA, Voulgaris S (2010) The role of the PTEN gene in malignant gliomas. Neurol Neurochir Pol 44(1):80–86 128. Weber GF (2008) Molecular mechanisms of metastasis. Cancer Lett 270(2):181–190. doi:10.1016/j.canlet.2008.04.030 DOI:dx.doi.org 129. Geiger TR, Peeper DS (2009) Metastasis mechanisms. Biochim Biophys Acta 1796(2): 293–308. doi:10.1016/j.bbcan.2009.07.006 DOI:dx.doi.org 130. Gimona M, Buccione R, Courtneidge SA, Linder S (2008) Assembly and biological role of podosomes and invadopodia. Curr Opin Cell Biol 20(2):235–241. doi:10.1016/ j.ceb.2008.01.005 DOI:dx.doi.org 131. Weaver AM (2008) Invadopodia. Curr Biol 18(9):R362–4. doi:10.1016/j.cub.2008.02.028 DOI:dx.doi.org
References
119
132. Deryugina EI, Quigley JP (2010) Pleiotropic roles of matrix metalloproteinases in tumor angiogenesis: contrasting, overlapping and compensatory functions. Biochim Biophys Acta 1803(1):103–120. doi:10.1016/j.bbamcr.2009.09.017 DOI:dx.doi.org 133. Poincloux R, Lizárraga F, Chavrier P (2009) Matrix invasion by tumour cells: a focus on MT1-MMP trafficking to invadopodia. J Cell Sci. 122(Pt 17):3015–3024. doi:10.1242/ jcs.034561 DOI:dx.doi.org 134. Alexander NR, Branch KM, Parekh A, Clark ES, Iwueke IC, Guelcher SA, Weaver AM (2008) Extracellular Matrix Rigidity Promotes Invadopodia Activity. Curr Biol 18(17): 1295– 1299. doi:10.1016/j.cub.2008.07.090 DOI:dx.doi.org 135. López-Otín C, Palavalli LH, Samuels Y (2009) Protective roles of matrix metalloproteinases: from mouse models to human cancer. Cell Cycle 8(22):3657–3662. doi:10.4161/cc.8.22.9956 DOI:dx.doi.org
wwwwwwwwwwwwwwww
Chapter 6
Cell Migration
Abstract The fibroblastic cell migration machinery involves such factors as cytoskeleton-regulated formation of pseudopodia, cell morphological polarization, dynamic regulation of focal adhesions during cell migration, integrin- and growth factor receptor-mediated signaling. The inhibition of directional movement of one cell across the surface of another cell caused by intercellular collisions (the phenomenon called “contact inhibition of cell migration”) and also biochemical, physical, and topographic characteristics of the extracellular matrix are also pivotal factors, which determine and control cell migration activity and the direction of migration. Oncogenic transformation causes various abnormalities of cell migration machinery. These alterations apply to adhesion and signaling functions of focal contacts in transformed cells, and also to the sensitivity of the cells to soluble growth factors.
6.1 Factors Involved in Cell Migration Cell migration is controlled by multiple factors, such as cytoskeleton-regulated formation of pseudopodia, cell morphological polarization, integrin- and growth factor receptor-mediated signaling, cell–extracellular matrix and cell–cell adhesions, and also biochemical, physical, and topographic characteristics of the extracellular matrix [1, 2]. In physiological processes, such as embryogenesis or regeneration, and also in tumor invasion, the cells can migrate both individually and collectively as sheets or clusters [3, 4]. Fibroblastic cells, contrary to isolated epitheliocytes, have ability to active locomotion across the extracellular matrix. During active translocations of fibroblastic cells to new areas of the extracellular matrix, the cells migrate individually (Fig. 6.1a). They constantly change their mutual positions, orientations, and shapes. The fibroblastic cells move using a combination of coordinated cyclic processes: formation and extension of pseudopodial protrusions at the active (leading) cell edge, Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_6, © Springer Science+Business Media, LLC 2011
121
122
6 Cell Migration
Fig. 6.1 (a, b) Moving fibroblastic and epithelial mouse cells. The individually migrating fibroblastic cells are polarized (a): the cells have wide leading edges (the anterior cell parts) and narrow “tails” (arrows). The coherently migrating epithelial cells are not polarized (arrow) (b). Phasecontrast microscopy. Scale bar, 45 mm. Reproduced with permission from The International Journal of Developmental Biology (ref. [7])
the attachment of pseudopodia to the extracellular matrix, centripetal stretch of the attached pseudopodia, and the detachment of the posterior (“tail”) part of the cell. The centripetal stretch of pseudopodia is the moving force, which translocates the fibroblastic cell in the direction of its leading edge. Repeated extensions and attachments of pseudopodia in the certain zone of the cell edge will eventually lead to the elongation of the fibroblastic cell in the corresponding direction and to the directional cell movement (Fig. 6.1a). Migration of fibroblastic cells is accompanied by the characteristic changes in their shape [5]. A migrating fibroblast acquires the characteristic “locomotory” phenotype with a polarized cell morphology: its anterior (in the direction of cell movement) part looks like a wide and thin plate called lamellar cytoplasm or lamella, at the free edge of which pseudopodia are constantly formed, attaching to the extracellular matrix or pulling back (it is a so-called active or leading cell edge). The opposite posterior part of the cell is retracted and often looks like a narrow “tail” (Fig. 6.1a). There is an assumption that the boundary between the forming lamellipodia and the lamellar cytoplasm is generated by stretching stresses in the lamellipodial actin gel. These stresses could be caused by the actin gel frictions against the cell–extracellular matrix adhesions underneath the lamellipodia. The stresses generate the actin gel disintegration that forms the lamellipodia-lamellar cytoplasm boundary [6]. Therefore, fibroblastic cell migration is carried out by the pseudopodial activity that is concentrated at the leading edge of the polarized cell. Contrary to fibroblastic cells, isolated epithelial cells are not polarized (see Fig. 5.23 in Chap. 5, Fig. 6.1b) and cannot move directionally, as they have their pseudopodial activity along the entire cell perimeter. Epitheliocytes migrate as a coherent sheet: mutual positions of the cells, locked to one another by their intercellular contacts, remain almost unchanged in the course of the migration. The only cells, which move actively, are those at the foremost contact-free edges of the epithelial cell sheet. These cells expand into the cell-free matrix areas without breaking the contacts with their lateral and posterior neighbors (Fig. 6.1b).
6.1 Factors Involved in Cell Migration
123
The coordinated interactions between actin cytoskeleton, microtubules, and cell adhesion structures play a key role in the formation of pseudopodia, polarization, and locomotion of fibroblastic cells [7–9]. For fibroblastic cell migration, a combination of two conditions is necessary: the influence of soluble growth factors, which are bound to their specific receptors on the cell surfaces, and the binding of integrin receptors to their ligands in the extracellular matrix. The upstream signaling events activated by the binding of growth factor receptors and integrins to their ligands are responsible for the stimulation not only of cell proliferation but also of cell migration (see Figs. 5.50 and 5.51 in Chap. 5).
6.1.1 Formation of Pseudopodia Pseudopodial protrusions formed by motile fibroblastic cells at their free active (leading) cell edges serve as “driving organs,” by means of which cells move across the extracellular matrix. The cells use pseudopodia of different morphological types. The main “driving organs” are lamellar protrusions, lamellipodia and filamentous outgrowths, filopodia. Filopodia can also carry out sensory or exploratory functions. Lamellipodia and filopodia are dynamic cytoskeleton-regulated formations. Their shapes and dynamics are determined by the spatial organization of actin filaments: branched filament networks are characteristic for lamellipodia, whereas filopodia contain parallel bundles of long unbranched filaments (Fig. 6.2) [10–12]. The formation of pseudopodia is caused by the polymerization of actin filaments from monomeric actin at the leading cell edge [8, 13–15]. Various actin-binding proteins, such as Arp2/3 complex, WASP/WAVE (WASP/ Scar), formins, cortactin, and other proteins, are critically involved in the generation of new actin filaments and actin cytoskeleton reorganizations (see Sects. 3.1.1 and 3.1.2), which are necessary for pseudopodial formation, integrin-mediated adhesion regulation, and thus for cell migration [10, 16–19]. Arp2/3 complex has a key role as a nucleator of branched actin filaments leading to the formation of dendritic actin filament networks (see Figs. 3.7 and 3.8 in Chap. 3). The developing branched filament networks presumably initiate the pushing force, which can protrude the plasma membrane forming wide lamellipodia [13]. Although some authors suggest that actin networks in lamellipodia are composed of unbranched actin filaments [20], this opinion has been refuted: lamellipodial branched actin filament networks are distinctly demonstrated [10] by means of significantly improved procedures for electron microscopic visualization of cytoskeletal structures [21, 22]. Arp2/3 complex is activated by the proteins of WASP/WAVE (WASP/Scar) family. All members of WASP family have a domain, through which Arp2/3 complex is activated to nucleate rapid actin polymerization in the leading edge of migrating fibroblastic cells [23]. WAVE2 is activated downstream of Rac and induces the formation of lamellipodia (see Fig. 5.55 in Chap. 5), N-WASP is activated downstream of
124
6 Cell Migration
Fig. 6.2 Fibroblast-like Drosophila cell of neural origin. Structural organization of a lamellipodium. Actin microfilaments form dendritic network in the lamellipodium and bundle of long microfilaments in the filopodium. Circled region is enlarged in the inset to show actin microfilament bundle. EM. Scale bar, 0.5 mm. Reproduced with permission from the Journal of Cell Science (ref. [24])
Cdc42 and induces the formation of filopodia (see Fig. 5.56 in Chap. 5) [17, 24–26]. In addition, N-WASP mediates dynamic attachment between plasma membrane and the growing ends of actin filaments to sustain pseudopodial protrusions [27]. Therefore, the Arp2/3 complex is involved in the formation of both lamellipodia and filopodia [28]. However, other stimulators of actin polymerization, the formin family proteins, particularly formin mDia2, are also critical for the formation of both these pseudopodial types: mDia2 depletion inhibits lamellipodia and filopodia formation [29]. mDia2 plays an important role in generation of long actin filaments in lamellipodia by actin nucleation or by protecting individual filaments from their capping. Long actin filaments subsequently exhibit high tendency to gradually converge into filopodial filament bundles (Fig. 6.2) [17, 26, 29]. Thus, filopodia can arise by reorganization of lamellipodial actin filament network, and this actin reorganization is induced by formin mDia2. Ena/VASP protein family plays a very important role in the alternative mechanism of lamellipodia or filopodia formation. Ena/VASP are actin regulatory proteins associated with barbed ends of filaments. Ena/VASP proteins protect the filaments from capping, thereby promoting their elongation [17, 26, 30, 31]. In addition, Ena/ VASP proteins reduce the density of Arp2/3 complex-dependent filament branches in lamellipodia and induce the clustering of the filament barbed ends [31–33]. Subsequently, the actin cross-linker protein fascin is recruited to the clustered barbed ends. Fascin gets these filaments with their associated barbed ends sewn
6.1 Factors Involved in Cell Migration
125
together into the long bundles, which presumably could protrude the cell membrane, forming very narrow long outgrowths, filopodia (Fig. 6.2) [34, 35]. Therefore, Ena/VASP proteins determine the “choice” of a cell between the formation of lamellipodia or filopodia. Ena/VASP proteins are also involved in the regulation of cell migration by controlling the organization pattern of lamellipodial actin filaments. The cells with Ena/VASP-deficient lamellipodia, which contain short highly branched filaments, migrate with increased speed. Lamellipodia with excess Ena/VASP contain longer, less branched filaments, and the cells migrate slower [36]. The actin-binding protein cortactin also plays a critical role in actin cytoskeleton remodeling in pseudopodia formation and in integrin-mediated adhesion. Cortactin is accumulated in lamellipodia and ruffles. It is activated by tyrosine phosphorylation caused by Src or other tyrosine protein kinases. The activated cortactin recruits Arp2/3 complex to existing actin filaments, promoting the Arp2/3-dependent formation of branched filament networks that cause the generation of lamellipodia. In the presence of activated cortactin, actin filaments are cross-linked into the bundles. Decreased expression of cortactin leads to the weakening of lamellipodial activity and decreased cell migration [19, 37]. The idea that the growth of actin filaments arranged in the branched networks or in the parallel bundle may initiate the “pushing force,” which protrudes the plasma membrane forming wide lamellipodia or thread-like filopodia, respectively [13, 16, 34, 38–40], was conventional up to now. However, recent data show that actin filaments may not directly push and deform plasma membrane to form pseudopodia at the leading cell edge. Specifically, during filopodia formation, plasma membrane can be deformed independently and generate small transient actin-free protrusions. These nascent protrusions are subsequently filled by actin filaments that obviously stabilize these protrusions providing their transition to the long extensions, filopodia [41]. From the plasma membrane at the cell leading edge, the newly formed actin filaments start their retrograde flow. In more proximal zone, in the lamellar cytoplasm, filaments are associated with myosin II and other actin-binding proteins, forming more regular actin-myosin filament bundles (see Sect. 3.1.2). The dynamics of the retrograde flow of filaments plays an important role in the formation of focal adhesions by a migrating cell. The assembly of the contractile actin-myosin filament bundles behind the active cell edge is in essence the fibroblastic cell migration machinery [42, 43]. Not only actin cytoskeleton but also microtubules are involved in the machinery of cell migration. Dynamics of microtubules and mutual influences between microtubules and Rho family of GTPases play a critical role in lamellipodia formation and directional locomotion of fibroblastic cells [44]. The displacement of a fibroblastic cell is accompanied by the orientation of the microtubules: their growing plus-ends are arranged in the proximal zone of the leading cell edge, and the minus-ends are arranged in the central part of the cell. Formation of lamellipodia is stimulated at the cell edge zone that is close to the plus-ends of microtubules. This local stimulation by the microtubule plus-ends may be caused by the activation of Rac1 protein, a member of Rho family of GTPases. Rac1-activating
126
6 Cell Migration
protein, called Asef, binds indirectly to plus-ends of growing microtubules via APC protein. APC that belongs to the group of microtubule-end tracking proteins (+TIPs) accumulates at the plus-ends of microtubules in cell pseudopodia. These APCmarked microtubules enter an increased phase of their growth. Besides, APC is associated with actin in the leading edges of migrating fibroblastic cells and influences actin dynamics in pseudopodia [45, 46]. The microtubule-mediated activation of Rac1 induces the polymerization, growth, and branching of actin filaments (see Fig. 5.55 in Chap. 5) leading to the formation of lamellipodia at the cell’s leading edge. Activation of Rac1 promotes microtubule growth into advancing lamellipodia of migrating cells. Another member of Rho family of GTPases, Rho, is activated by the Rho guanine nucleotide exchange factor (GEF-H1) that is bound to microtubules and gets free during their depolymerization [47]. As a result of depolymerization of microtubules at their minus-ends in the cell body, Rho is activated. It induces the assembly of actin filaments into bundles and increases their contractility (see Fig. 5.54 in Chap. 5), which favors the pulling of the “tail” of the migrating fibroblastic cell. A member of Rho family of GTPases, Cdc42, is involved in the orientation of microtubule-organizing center (centrosome) toward the leading edge of the migrating cell favoring the orientation of the growing microtubule plus-ends in the same direction. In some special cases when cells migrate on the low-adhesive surface, they can form spherical pseudopodia, called blebs [48]. The formation of blebs at the anterior cell edge is caused by transient detachment of plasma membrane from the cortical actin layer. Blebs are formed at sites where cortical actin is locally depolymerized and separated from the plasma membrane. The detached plasma membrane undergoes local outward expansion caused by the cytosol that streams out of the cell body and fills up the forming blebs. Polymerized actin is not initially found within the forming blebs, but it appears later. The actin polymerization stops the expansion of blebs [49, 50].
6.1.2 Polarization of Migrating Cells Irregular distribution of pseudopodia along the free cell edge of a migrating fibroblastic cell, their concentration at a definite zone, called active or leading cell edge, results in the characteristic cell shape changes [51] called cell morphological polarization. Cell polarization is a critical condition for directional fibroblastic cell migration. The integrity of centralized microtubule system provides the basic mechanism of the fibroblastic cell polarization [7, 52]. Microtubules control the elongated shape of a fibroblastic cell and the division of its edge into active (pseudopodia-forming) and nonactive zones. Depolymerization of microtubules by specific drugs, e.g., colcemide or nocodazole, or their decentralization induced by taxol, leads to loss of the elongated fibroblastic cell shape and to
6.1 Factors Involved in Cell Migration
127
the disappearance of nonactive zones of the free cell edge. The cell acquires discoid epitheliocyte-like shape, and the pseudopodial activity is randomly distributed along the cell perimeter [7, 53, 54]. The control of fibroblastic cell polarization is effected by microtubule-dependent stimulation of actin polymerization and lamellipodia formation at the definite zone of the free cell edge, which thereby becomes the active (leading) edge (Fig. 6.1a). This stimulation is caused by microtubule-mediated activation of Rac1 protein. Transport functions of microtubules also play a key role in the maintenance of polarized morphology and directional locomotion of fibroblastic cells. Microtubules deliver localized doses of some contractility relaxing agents to definite focal contacts to retard or reverse their development (microtubule targeting). By modulating the pattern of focal contacts developed by the cell, microtubules establish the asymmetry of traction forces exerted on the extracellular matrix through focal contacts. The asymmetric traction forces determine the polarized fibroblastic cell morphology and are involved in directional locomotion. Inhibition of transport function of a microtubule motor protein kinesin reduces fibroblastic cell polarization [7, 55–58].
6.1.3 Contact Inhibition of Cell Migration When two migrating fibroblastic cells collide, the active (leading) edge of one cell continues to move and begins to overlap the surface of the other cell. However, the pseudopodial activity of the overlapping active cell edge is soon inhibited. This inhibition of the pseudopodial activity by cell–cell contact is called contact paralysis [59]. The overlapping active edge retracts, and pseudopodia start to form at another zone of the free cell edge causing the migration of the upper cell in another direction. Finally both cells move away from each other. Thus, the moving fibroblastic cells disperse after their short-term intercellular contact [7]. In contrast to fibroblastic cells, collision of moving epithelial cells is not followed by their overlapping (see Fig. 5.25 in Chap. 5). The pseudopodial activity at the contacting cell edges is immediately inhibited (contact paralysis), the cell movement stops, and the stable cell–cell adhesion structures are formed [60]. Mechanism of “contact paralysis” is possibly based on the suppressive effect of a-catenin on Arp2/3 protein complex. At the initial stage of cell–cell adhesion formation, Arp2/3 complex, which controls the polymerization of branched actin filament networks in pseudopodia at the active edges of colliding cells, can be inhibited by high level of a-catenin (see Sect. 9.1.2). The Arp2/3 activity suppression results in the inhibition of pseudopodial activity [16, 61]. Therefore, moving normal cells respond to the intercellular collisions by the inhibition of directional movement of one cell across the surface of another cell. This phenomenon, called contact inhibition of cell migration [59], is differently manifested in fibroblastic and epithelial cells: as a consequence of collision, fibroblastic cells change the direction of their migration, while epitheliocytes stop their movement and form stable intercellular contacts.
128
6 Cell Migration
The differences in the realization of contact inhibition of cell migration are based on the cytoskeleton mechanical mechanism. When moving epithelial cells collide with each other, their actin filament bundles in the zone of the intercellular contact are reorganized. As a result, the summary vector of the tension forces generated by the contractile filament bundles is laterally directed along the edges of the contacting cells (see Fig. 9.5 in Chap. 9). This mechanical tension force leads to “contact paralysis” of the pseudopodial activity in the zone of the intercellular contact, and causes the expansion and stabilization of cell–cell adhesion, thereby excluding subsequent epithelial cell migration [60]. Contrary to epitheliocytes, in colliding fibroblastic cells, actin filament bundles are not reorganized, and the summary vector of the tension forces remains directed centripetally along the long axis of each of the contacting cells. This tension leads to retraction of the overlapping active cell edge with subsequent formation of a new active edge and thereby changes the direction of cell migration. In the mechanism of contact inhibition of cell migration, nectins and nectin-like (Necls) proteins play a crucial role [62, 63]. Nectins and Necls are immunoglobulinlike transmembrane intercellular adhesion molecules, which interact with the plateletderived growth factor (PDGF) and aVb3 integrins. Nectins and Necls are involved in the signaling pathways from these cell surface receptors. The interaction of Necl-5 with PDGF and aVb3 receptors stimulates the formation of pseudopodia and focal adhesions at the leading cell edges thereby enhancing directional cell migration. When two migrating cells collide, Necl-5 interacts with nectin-3. This interaction causes a drop in the expression of Necl-5 at the cell surface. Then PDGF and aVb3 integrins interact with nectins that initiate intercellular adhesion (see Sect. 9.1.2). The formation of stable adhesion stops cell locomotion. One of the components of cell–cell adhesion structures, p120-catenin, also plays an important role in the mechanism of contact inhibition of cell migration (see Sect. 9.1.2).
6.1.4 Effect of Growth Factors Many soluble peptide growth factors are mitogens-motogens, i.e., they can influence both cell proliferation and locomotion. The binding of growth factors to their specific cell surface receptors stimulates both cell migration and proliferation; some of the growth factors stimulate locomotion and inhibit multiplication of the cells. The cell reaction to a definite growth factor can vary in different cell types. The growth factor HGF/SF is a potent motogen for various epithelial cells (e.g., for mammary gland epithelium) and endothelial cells. The motogenic effect of HGF/SF on an epithelial layer causes disruption of the cell–cell adhesions, change of the discoid to fibroblast-like cell shape, and reorganization of actin cytoskeleton: circular filament bundles typical of epithelial cells disappear, and straight bundles typical for fibroblastic cells develop. The immobile epitheliocytes are converted
6.1 Factors Involved in Cell Migration
129
into elongated motile fibroblast-like cells with polarized pseudopodial activity; they actively migrate from the epithelial cell layer. The transition from the epithelial to the fibroblast-like cell phenotype is called epithelial-mesenchymal transition (EMT) [64–66]. During EMT the special structures providing intercellular adhesions between epithelial cells get broken. The loosening of the intercellular adhesive bonds leads to the activation of the Wnt signaling (see Sect. 9.1.3), resulting in the stimulation of cell proliferation. Epithelial cells decrease the expression of epithelial-specific genes, including the ones encoding E-cadherin receptors of cell–cell adhesion structures and keratins of intermediate filaments. At the same time, the cells gain the expression of mesenchymal-specific genes, such as genes encoding integrins, collagens, and vimentin of intermediate filaments. Finally, epithelial cells acquire fibroblast-like morphology and high motility [67, 68]. EMT is a crucial event during normal embryonic development and also in cancer cell invasion [66, 69, 70]. HGF/SF can induce not only complete scattering of compact epithelial cell sheets into single migrating fibroblast-like cells but also promote the outgrowth of branching tubules from epithelial cysts [7]. EMT and epithelial tubulogenesis induced by HGF/SF are involved in the formation of many organs at certain stages of embryogenesis and during wound healing. The binding of mitogens-motogens to their cell surface receptors “triggers” both the Ras-Raf-MAP kinase “mitogenic” signaling transduction pathway and the “morphogenic” one: Ras-PI3K-Rho GTPases. In the latter pathway, the signals from activated Ras are transferred to Rho GTPases (see Fig. 5.51 in Chap. 5). Being activated, these proteins (Rho, Rac, and Cdc42) stimulate polymerization and contractility of actin filaments; they regulate the assembly/disassembly of focal contacts and initiate the formation of cell pseudopodia of different shapes (see Sect. 5.3.5) resulting in cell migration [71]. Activated Rho proteins stimulate linear growth of actin filaments, increase in their contractility and their organizing into stress fibers, thereby promoting the formation of mature focal contacts (see Fig. 5.54 in Chap. 5). Rho proteins also initiate microtubule targeting, which leads to the disassembly of focal contacts. Both the formation of mature focal contacts and their disassembly are necessary conditions for active cell migration. The activated Rac1 protein stimulates growth of filament branching “tree.” The growing filament ends facing cell membrane may produce the pushing force, which presumably can protrude the cell membrane and form wide pseudopodia, lamellipodia (see Fig. 5.55 in Chap. 5). In a similar way, Cdc42 protein stimulates growth of actin filaments. Some of them are joined to each other by their barbed ends, and then, under the action of the cross-linker protein fascin, these filaments are sewn together into long bundles. These bundles form very narrow and long pseudopodia, filopodia (see Fig. 5.56 in Chap. 5). The p53 tumor suppressor gene, in addition to its apoptotic effect (see Sect. 5.3.4), is involved in the modulation of fibroblastic cell migration including the production of pseudopodia and cell polarization. Loss of p53 function correlates
130
6 Cell Migration
with decreased cell motility [72]. The pseudopodial activity and directional locomotion of fibroblastic cells are simulated by p53 through the regulation of Rho signaling, thereby controlling actin cytoskeleton dynamics [73]. Besides, p53 can activate the genes encoding cell receptors of some mitogens-motogens (e.g., EGF or HGF/SF); in this way p53 modulates signal transduction from these receptors to PI3K-Rac1. At the same time, p53 can inhibit the fibronectin gene in some types of cells, and because of that, these cells with their high pseudopodial activity cannot migrate: they “skid” on the extracellular matrix surfaces [72, 74, 75].
6.1.5 Role of Focal Adhesions in Cell Migration Adhesive interactions of cells with the extracellular matrix are critically involved in cell translocation. Cell migration requires dynamic regulation of integrin-mediated cell adhesion, assembly and disassembly of focal contacts at various regions of the basal cell surface, and the functioning of integrin- and growth factor-mediated signaling pathways [76, 77]. In a migrating fibroblastic cell, focal contacts are usually found underneath lamellipodia and also at the lamellipodia-lamellar cytoplasm (lamella) boundary (see Fig. 4.13c in Chap. 4). In the formation of stable focal adhesions, the dynamics of the retrograde flow of newly formed actin filaments (see Sect. 3.1.2) plays a crucial role. The new focal contacts start to be formed underneath lamellipodia. Almost immediately after the formation of the nascent focal adhesions, the speed of the retrograde actin filament flow in the lamellipodial zone changes from high to low. This filament flow speed change is a necessary condition for the stabilization of focal adhesions: inhibition of the rapid flow results in immediate dissolution of nascent focal contacts. In conditions when a cell does not form focal adhesions, the change in the filament flow speed is not observed [78, 79]. Therefore, the process of focal contact formation depends on actin filament dynamics which, in turn, affects the transition from rapid to slow retrograde flow of actin filaments. Attachment of pseudopodia to the extracellular matrix through the formation of focal contacts and associated stress fibers creates centripetal contractile tension. Stress fibers participating in cell migration machinery are the dorsal ones. They are anchored with their front ends to focal adhesions, whereas their back ends are intertwined with the cortical actin filament network. Contraction of dorsal stress fibers results in the pull of the cell in the direction of anchored stress fiber front ends. This contraction translocates the cell body in the direction of its active (leading) edge and from time to time causes the detachment of the posterior part of the cell. The contraction of the cell “tail” is followed by the activation of pseudopodia formation at the opposite leading cell edge. This activation is due to the contraction-induced stimulation of the forward flow of actin monomers to the leading cell edge.
6.1 Factors Involved in Cell Migration
131
Among various protein components of the extracellular matrix, laminins have a particular role in promoting cell migration. Laminins are a secreted family of large heterotrimeric glycoproteins. Laminins are localized in basement membranes including those of blood vessels [80–83]. One of the laminin isoforms, laminin-332 (previously known as laminin-5), is the major ligand for keratinocyte adhesion in the epidermis. The integrin receptors for laminin-332 are a3b1 and a6b4 [84, 85]. Laminin-332 promotes cell adhesion and migration activity much more efficiently than other extracellular matrix proteins [86–88]. An important function of laminin-332 is the induction of epithelial cell migration. This function is a result of the proteolytic processing of laminin-332 mediated by the matrix metalloproteinases (MMPs): different products of the laminin-332 processing may bind to distinct integrins, activate them, and thereby trigger epithelial cell migration [86, 89, 90]. The migratory behavior of epithelial cells on laminin-332 is determined by its organization that is regulated by a6b4 integrin: keratinocytes expressing this receptor assemble laminin-332 in linear tracks, over which the cells migrate, whereas b4 integrin-deficient epitheliocytes assemble laminin-332 in circular arrays, and the cells move circularly [56, 91]. Another laminin isoform, laminin-411 (formerly known as laminin-8), is present in basement membranes of blood vessels. Cell adhesion to laminin-411 is mediated by a6b1 and a3b1 integrins. Laminin-411 provides a weak adhesion of endothelial cells; however, it is potent in promoting cell migration during development, wound healing, and angiogenesis [92]. In the regulation of laminin-binding integrins, the proteins tetraspanins play a crucial role. Tetraspanins are associated with laminin-binding integrins and organize them into multiprotein complexes (termed tetraspanin-enriched microdomains). These complexes, depending on their composition in different cell types, can crucially promote rapid migration of the cells. However, the complexes can also be involved in the stabilization of intercellular adhesion and thereby can inhibit cell migration [93]. Endocytic-exocytic transport of integrins (integrin trafficking) facilitates cell migration. The trafficking internalizes integrin receptors into endosomes at the rear of the migrating cell and transports them forward within vesicles for exocytosis at the leading cell edge where new focal contacts form. Integrin trafficking may be controlled by one of the families of small GTPases, Rab GTPases, that are regulators of cellular membrane transport [94–96]. FAK and a large GTPase, dynamin, play an important role in the regulation of cell movement. FAK is involved in mechanosensing during fibroblastic cell migration. The microtubule targeting-induced disassembly of focal contacts during cell migration is mediated by FAK and dynamin. FAK-null cells show a decrease in their migration rate and in directional persistence [97]. Dynamin facilitates cell migration by its involvement in the cycle of cell membrane expansion and retraction that is essential for cell movement [98]. Integrin receptor-mediated signals from FAK cause sequential tyrosine phosphorylation of many proteins that are involved in cell motility. The phosphorylated focal contact protein p130Cas (Cas) initiates the activation of Rac1 and lamellipodia
132
6 Cell Migration
formation [99–101]. Activated kinase PKB/Akt, besides its antiapoptotic effect (see Sect. 5.3.3), is involved in actin cytoskeleton dynamics. PKB/Akt in response to the epidermal growth factor (EGF) phosphorylates actin-binding protein girdin. Phosphorylated girdin accumulates at the leading edge of a migrating fibroblastic cell, suggesting its role in the PKB/Akt-dependent regulation of cell migration [102, 103]. The key participants of the “morphogenic” integrin- and growth factor-mediated signaling pathways, Rho GTPases, which are involved in pseudopodia formation (see Sect. 5.3.5), control directional cell migration [104, 105]. P21-activated kinases (PAKs) that modulate mitogenic signaling pathway from integrins also participate in the control of cell motility. The binding of PAK1 to the cytoplasmic protein nischarin inhibits the ability of PAK1 to phosphorylate and causes inhibition of cell migration [106, 107]. For the displacement to the new areas of the extracellular matrix, migrating fibroblastic cells must have the capability to liquidate their old focal adhesions. This possibility is ensured by microtubule targeting: the plus-ends of microtubules are getting concentrated near growing focal contacts and promote their disassembly in conjunction with the cell edge retraction (see Sect. 4.2.1). Microtubule targeting proves to be necessary for the migration of fibroblastic cells, which form focal contacts. The cells, which form only focal complexes, but do not convert them into mature focal adhesions (neutrophils, epidermal keratinocytes), can successfully migrate with the destroyed system of microtubules. The disassembly of focal contacts during cell migration can be also caused by the large family of proteins, called semaphorins. They are specifically bound to and activate their cell transmembrane receptors, plexins. Plexin receptors are directly associated with several GTPases; besides, plexins themselves can activate small GTPases (acting like GTPase activating proteins, GAPs). Semaphorin-induced activation of plexins leads to the disassembly of focal contacts and to actin cytoskeleton reorganization via specific GTPases, thereby affecting the guidance of cell migration [108–110].
6.2 Abnormalities of Cell Migration Machinery in Transformed Cells The migration of malignant tumor cells is the basis of cancer invasion and metastasis [111–114]. Oncogenic transformation of cells caused by permanent hyperexpression of oncogenes leads to typical morphological alterations of the cells and to an increase in their motility. These special features play significant role in tumor invasion, contributing, together with other factors, to the active invasion of malignant tumor cells into the surrounding healthy tissues.
6.2 Abnormalities of Cell Migration Machinery in Transformed Cells
133
6.2.1 Pseudopodial Activity with Actin-Myosin Structure Deficiencies Activity of lamellipodia and filopodia as “driving organs” of migrating malignant tumor cells plays a crucial role in cancer invasion and metastasis [113]. Ras genes (K-ras, H-ras, and N-ras) are the most frequently mutated oncogenes detected in human cancers. Upon transformation, both fibroblastic cells and epitheliocytes acquire poorly spread shape, and the sizes of pseudopodia-forming zone of the active cell edge get significantly reduced (see Figs. 5.33 and 5.35 in Chap. 5). However, this reduction is not accomplished by diminished pseudopodial activity per unit length of the active cell edge, so the cells have high motility [60]. Ras and its downstream effectors, such as Raf and Rho GTPases, are responsible for the morphological alterations of transformed cells and for the stimulation of their locomotion. In these effects, just as in the motogenic effects of mitogensmotogens on normal cells (see Sect. 5.3.2 and 5.3.5), the activation of “morphogenic” Ras-Raf-MAP kinases and “motogenic” Ras-PI3K-Rho GTPases signal transduction pathways plays a key role. In transformed cells, these signaling pathways are permanently activated. This activation is a result of the conversion of proto-oncogenes, which encode key components of signal transduction cascades into oncogenes [115]. Overexpression of FAK that is the initiator of integrin-dependent mitogenic or motogenic signaling stimulates cancer cell migration. Rho GTPases, such as Rho, Rac, and Cdc42, are critically involved in the mechanics of cell migration (see Sect. 6.1.4), and play an important role in tumorigenesis [116]. They are often overexpressed in transformed cells. Permanently high levels of Rac and/or Cdc42 result in the stimulation of pseudopodial formation and favor increased motility of transformed cells. Overexpression of focal adhesion protein p130Cas, initiating Rac1 activation, stimulates lamellipodial formation, and thereby contributes to increased cancer invasion [117]. The elevated level of activated RhoA is found in many transformed cells; however, this high level does not result in expected stimulation of actin filament bundle and focal adhesion formation. On the contrary, stress fibers and connected mature focal contacts, as a rule, are significantly reduced in transformed cells (see Figs. 5.36 and 5.40 in Chap. 5). The reason for this is that permanently activated mitogenic MAP kinase signaling inhibits one of the main Rho effectors, Rho-associated kinase (Rho kinase, ROCK). This Rho effector is required for the assembly of actin filaments into the bundles, mature focal contact formation, and cell contractility (see Sect. 5.3.5). Inhibition of Rho kinase by its specific inhibitor in normal fibroblastic or epithelial cells leads to the acquisition of the “transformed phenotype”: the fibroblastic cells become poorly spread, their active edges are significantly diminished, and the cells lose stress fibers. Epitheliocytes acquire elongated or spindlelike shapes, their circular actin bundles disappear, and the cells actively migrate [7]. Inhibition of Rho kinase induced by the activated MAP kinase signaling is mediated by p21 (WAF1) protein (a cell cycle inhibitor). Permanent activation of mitogenic
134
6 Cell Migration
signaling causes the localization of p21 protein to the cytoplasm. This protein forms a complex with Rho kinase and inhibits its activity. The phosphorylation level of cofilin is also decreased [118, 119]. Typical morphological changes and the stimulation of mobility of transformed epithelial cells need permanent activation of both “mitogenic” and “morphogenic” signaling pathways, whereas for the transformed fibroblasts the activation of any of these pathways is sufficient. One should also note that the described morphological and locomotory cell features are manifested in transformed cells with an additional condition only, when some tumor suppressor genes, including p53, are inactivated [115]. Actively moving transformed cells are polarized. However, instead of a wide leading edge, which determines the direction of movement of a normal fibroblastic cell, pseudopodia in transformed cells are formed at multiple short sections of the cell edge (see Figs. 5.31, 5.33c, and 5.35 in Chap. 5). Because of this, tumor cells migrate by jerky movements, frequently changing the direction of their displacement. When two migrating transformed cells collide with one another, different cell responses are possible. If active cell edges of both cells collide, the lamellipodial activity at the site of the contact immediately stops (“contact paralysis”), and the cells move away from one another. In this case “contact inhibition of cell migration” is still working in transformed cells. However, if an active edge of one cell collides with a nonactive lateral edge of another cell, the active edge of the first cell continues to move under the body of the second cell. In this case “contact paralysis” and “contact inhibition of cell migration” do not take place. Because of underlaps, transformed cells often form multilayered colonies where the cells crisscross one another (see Fig. 9.8 in Chap. 9).
6.2.2 Cell-Matrix Adhesion Alterations The integrin-mediated cell–extracellular matrix adhesion constitutes the core machinery of cell migration. The deficiency of focal adhesions and significant alterations in their adhesive and signal transduction functions are typical for transformed cells (see Sects. 5.4.1 and 5.4.2) and play a critical role in their locomotion [120, 121]. Endocytic-exocytic transport of integrins, which facilitates cell migration, can be disturbed in transformed cells, and this change plays a role in the altered cancer cell migration and invasion [94–96]. The weakening of adhesive bonds of transformed cells to the extracellular matrix can provoke their increased migratory activity. For instance, the reduction in the affinity of the a5b1 integrin receptor to its matrix ligand, fibronectin, is accompanied by intensified malignant cell migration. Among various protein components of the extracellular matrix, laminins have a particular role in promoting cell migration and cancer invasion. The acquisition of invasive properties by some of the malignant human tumors may be in part related
6.2 Abnormalities of Cell Migration Machinery in Transformed Cells
135
to the expression of laminin-411 by the tumor cells. It is possible that the tumor cells are weakly attached to the matrix, which contains laminin-411, which contributes to an increase of their migratory activity and to the invasive properties. In particular, the invasive ability of prostate cancer is possibly related to the high-level expression of the a6b1 integrin binding to laminin-411. The invasive properties of human brain gliomas and breast cancers are related to overexpression of laminin-411 by tumor cells with a switch from b2 to b1 chain-expressing laminin isoforms. During tumor progression of human gliomas to the much more malignant tumors, glioblastomas, the expression of laminin-421 (formerly, laminin-9) is switched to laminin-411 that may significantly facilitate tumor cell migration and tumor spread. The switches from laminin-421 and laminin-521 to laminin-411 and laminin-511 in tumor vascular basement membranes are found during progression of human breast cancers to the invasive stage [122–124]. Possibly, laminin-411, as a vascular basement membrane component, promotes cancer invasion not only by intensifying cancer cell migration but also by inducing the formation of new microvessels in the tumors. On the other hand, the strengthening of the adhesion of transformed cells to some ligands of the extracellular matrix can also provoke the increased migratory activity of these cells. Abnormal expression of laminin-332 (formerly, laminin-5), which very effectively promotes cellular adhesion and migration, and also of its a3b1 and a6b4 binding integrins, is typical for certain tumor types and is believed to promote cancer cell invasion [84, 125]. Laminin-332 and the products of its processing by MMPs are frequently found at the leading edges of invading cancer cells stimulating their migration and favoring cancer invasion [86–88, 126]. The laminin-binding integrins can both promote and inhibit cell migration depending on their regulation by tetraspanins (see Sect. 6.1.5). Therefore, the lamininbinding integrins can serve both pro- and anti-invasive functions [93]. Some components of the extracellular matrix produced by cancer cells are involved in the induction of EMT during which the cancer cells lose their epithelial characteristics and acquire mesenchymal phenotype suitable for active cell migration. Their intercellular adhesions are reduced; the cells acquire fibroblast-like shapes and enhanced motility. Thereby EMT plays a dramatic role in cancer invasion promoting active penetration of surrounding tissues by migrating cancer cells [69, 127–131]. Such components of the extracellular matrix as types I and III collagen are able to initiate disruption of E-cadherin-mediated intercellular adhesions (see Sect. 9.2.1), which is key in the EMT induction [132–135].
6.2.3 Hypersensitivity to Mitogens-Motogens The deregulation of growth factors and/or their specific cell receptors plays a critical role in such malignant properties of tumor cells as their stimulated proliferation, active migration, tumor angiogenesis, invasion, and metastasis [136–140].
136
6 Cell Migration
The mitogens-motogens initiating sequential signal transduction play the important role in the stimulation of the locomotory activity of transformed cells. These cells have the ability to proliferate and to actively move in the presence of concentrations of exogenous mitogens-motogens that are 10 or 100 times less than those required for these processes in normal cells. This capability may be explained by (a) the fact that transformed cells themselves are able to produce and secrete high levels of different mitogens-motogens. Thus, transformed cells can themselves stimulate their multiplication and locomotion, and (b) the number of mitogen-motogen receptors is considerably increased in many types of transformed cells. Tumor cells can produce and secrete such mitogens-motogens as PDGF, IGF-1, the growth factor bombesin, transforming growth factor-a (TGF-a), FGF-2, and VEGF. IGF-1 and bombesin are secreted by human melanoma and lung carcinoma cells, respectively, and stimulate locomotion and growth of these cancer cells. VEGF is produced and secreted by some human malignant tumors, in particular, by ovarian cancer cells. In cooperation with FGF-2, VEGF stimulates proliferation and directional migration of endothelial cells, promoting the growth of blood vessels into tumors [138, 140]. The ensuing neovascularization significantly contributes to the progressive tumor growth. Therefore, small-molecule inhibitors of VEGF, and/ or its receptor signaling may have a potential for the treatment of a number of human cancers [141]. However, as VEGF is also needed for normal angiogenesis, the inhibitor use may lead to considerable side effects. The expression of proto-oncogenes that encode mitogen-motogen receptors is strongly and constitutively increased in cells of many human malignant tumors. For example, the level of EGF receptor that is the product of the proto-oncogen. c-ErbB1/HER is often strongly increased in cells of various human carcinomas including lung cancer, glioblastomas, breast, ovarian, and stomach cancers [142, 143]. The capability of these tumors for invasion correlates with their overexpression of the EGF receptor. This may be related to a motogenic effect of EGF. Stimulating mobility and proliferation of blood vessel endothelial cells, EGF promotes the attachment of circulating cancer cells to blood vessel wall [144]. The other strong motogen for the tumor cells is the scatter factor (HGF/SF). It is produced by fibroblasts and other stromal cells of many human tumors. This may be related to the fact that the tumor cells can secrete some factors, which stimulate HGF/SF production by the tumor stromal cells. Several such factors are known that are secreted by breast and bladder cancer cells. The HGF/SF receptor that is the product of proto-oncogene c-met is permanently overexpressed in many human carcinoma cells. They become the targets for the motogenic action of HGF/SF that initiates the EMT in these cancer cells promoting their active migration and invasion into the surrounding tissues. In addition to the ligand-dependent activation, HGF/SF receptor activation can be negatively regulated by the intercellular contacts: the formation of stable cell–cell adhesions suppresses c-met-mediated signaling. As a result of c-met mutations in human cancers, its ability for negative regulation by the intercellular adhesion is lost [139]. Drugs targeting GF receptors, e.g., EGF or HGF/SF receptors, can be promising for future treatment of cancer invasion [139, 145–148].
References
137
A separate group includes “clean” motogens, which stimulate locomotion of tumor cells without influencing their proliferation. For example, the autocrine motility factor (AMF), produced by human melanoma cells, can stimulate their migration.
References 1. Petrie RJ, Doyle AD, Yamada KM (2009) Random versus directionally persistent cell migration. Nat Rev Mol Cell Biol 10(8):538–549. doi:10.1038/nrm2729 DOI:dx.doi.org 2. Friedl P, Wolf K (2010) Plasticity of cell migration: a multiscale tuning model. J Cell Biol 188(1):11–19. doi:10.1084/JEM2071OIA4 DOI:dx.doi.org 3. Friedl P, Gilmour D (2009) Collective cell migration in morphogenesis, regeneration and cancer. Nat Rev Mol Cell Biol 10(7):445–457. doi:10.1038/nrm2720 DOI:dx.doi.org 4. Weijer CJ (2009) Collective cell migration in development. J Cell Sci 122(Pt 18):3215–3223. doi:10.1242/jcs.036517 DOI:dx.doi.org 5. Aman A, Piotrowski T (2010) Cell migration during morphogenesis. Dev Biol 341(1):20–33. doi:10.1016/j.ydbio.2009.11.014 DOI:dx.doi.org 6. Shemesh T, Verkhovsky AB, Svitkina TM, Bershadsky AD, Kozlov MM (2009) Role of focal adhesions and mechanical stresses in the formation and progression of the lamellipodiumlamellum interface. Biophys J 97(5):1254–1264. doi:10.1016/j.bpj.2009.05.065 DOI:dx.doi.org 7. Vasiliev JM (2004) Cytoskeletal mechanisms responsible for invasive migration of neoplastic cells. Int J Dev Biol 48(5–6):425–439. doi:10.1387/ijdb.041806jv DOI:dx.doi.org 8. Pollard TD, Cooper JA (2009) Actin, a central player in cell shape and movement. Science 326(5957):1208–1212. doi:10.1126/science.1175862 DOI:dx.doi.org 9. Kodama A, Lechler T, Fuchs E (2004) Coordinating cytoskeletal tracks to polarize cellular movements. J Cell Biol 167(2):203–207. doi:10.1083/jcb.200408047 DOI:dx.doi.org 10. Svitkina TM, Borisy GG (1999) Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol 145(5):1009–1026. doi:10.1083/jcb.145.5.1009 DOI:dx.doi.org 11. Mattila PK, Lappalainen P (2008) Filopodia: molecular architecture and cellular functions. Nat Rev Mol Cell Biol 9(6):446–454. doi:10.1038/nrm2406 DOI:dx.doi.org 12. Le Clainche C, Carlier MF (2008) Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol Rev 88(2):489–513. doi:10.1152/physrev.00021.2007 DOI:dx.doi.org 13. Borisy GG, Svitkina TM (2000) Actin machinery: pushing the envelope. Curr Opin Cell Biol 12(1):104–112. doi:10.1016/S0955-0674(99)00063-0 DOI:dx.doi.org 14. Carlier MF, Pantaloni D (2007) Control of actin assembly dynamics in cell motility. J Biol Chem 282(32):23005–23009. doi:10.1074/jbc.R700020200 DOI:dx.doi.org 15. Insall RH, Machesky LM (2009) Actin dynamics at the leading edge: from simple machinery to complex networks. Dev Cell 17(3):310–322. doi:10.1016/j.devcel.2009.08.012 DOI:dx.doi.org 16. Chhabra ES, Higgs HN (2007) The many faces of actin: matching assembly factors with cellular structures. Nat Cell Biol. 9(10):1110–1121. doi:10.1038/ncb1007-1110 DOI:dx.doi.org 17. Mellor H (2010) The role of formins in filopodia formation. Biochim Biophys Acta 1803(2):191–200. doi:10.1016/j.bbamcr.2008.12.018 DOI:dx.doi.org 18. Paul AS, Pollard TD (2009) Review of the mechanism of processive actin filament elongation by formins. Cell Motil Cytoskeleton 66(8):606–617. doi:10.1002/cm.20379 DOI:dx.doi.org 19. Ren G, Crampton MS, Yap AS (2009) Cortactin: Coordinating adhesion and the actin cytoskeleton at cellular protrusions. Cell Motil Cytoskeleton 66(10):865–873. doi:10.1002/ cm.20380 DOI:dx.doi.org
138
6 Cell Migration
20. Small JV, Auinger S, Nemethova M, Koestler S, Goldie KN, Hoenger A, Resch GP (2008) Unravelling the structure of the lamellipodium. J Microsc 231(3):479–485. doi:10.1111/j.1365-2818.2008.02060.x DOI:dx.doi.org 21. Svitkina TM, Verkhovsky AB, Borisy GG (1995) Improved procedures for electron microscopic visualization of the cytoskeleton of cultured cells. J Struct Biol 115(3):290–303. doi:10.1006/jsbi.1995.1054 DOI:dx.doi.org 22. Svitkina T (2009) Imaging cytoskeleton components by electron microscopy. Methods Mol Biol 586:187–206. doi:10.1007/978-1-60761-376-3_10 DOI:dx.doi.org 23. Samarin SN (2005) WASP family proteins act between cytoskeleton and cellular signaling pathways. Biochemistry (Mosc) 70(12):1305–1309. doi:10.1007/s10541-005-0262-5 DOI:dx.doi.org 24. Biyasheva A, Svitkina T, Kunda P, Baum B, Borisy G (2004) Cascade pathway of filopodia formation downstream of SCAR. J Cell Sci 117(Pt 6):837–848. doi:10.1242/jcs.00921 DOI:dx.doi.org 25. Takenawa T, Miki H (2001) WASP and WAVE family proteins: key molecules for rapid rearrangement of cortical actin filaments and cell movement. J Cell Sci 114 (Pt 10):1801–1809 26. Faix J, Breitsprecher D, Stradal TE, Rottner K (2009) Filopodia: Complex models for simple rods. Int J Biochem Cell Biol 41(8–9):1656-1664. doi:10.1016/j.biocel.2009.02.012 DOI:dx.doi.org 27. Svitkina T (2007) N-WASP generates a buzz at membranes on the move. Cell 128(5):828–830. doi:10.1016/j.cell.2007.02.028 DOI:dx.doi.org 28. Korobova F, Svitkina T (2008) Arp2/3 complex is important for filopodia formation, growth cone motility, and neuritogenesis in neuronal cells. Mol Biol Cell 19(4):1561–1574. doi:10.1091/mbc.E07-09-0964 DOI:dx.doi.org 29. Yang C, Czech L, Gerboth S, Kojima S, Scita G, Svitkina T (2007) Novel roles of formin mDia2 in lamellipodia and filopodia formation in motile cells. PLoS Biol 5(11):e317. doi:10.1371/journal.pbio.0050317 DOI:dx.doi.org 30. Trichet L, Sykes C, Plastino J (2008) Relaxing the actin cytoskeleton for adhesion and movement with Ena/VASP. J Cell Biol 181(1):19–25. doi:10.1083/jcb.200710168 DOI:dx.doi.org 31. Bear JE, Gertler FB (2009) Ena/VASP: towards resolving a pointed controversy at the barbed end. J Cell Sci 122(Pt 12):1947–1953. doi:10.1242/jcs.038125 DOI:dx.doi.org 32. Mejillano MR, Kojima S, Applewhite DA, Gertler FB, Svitkina TM, Borisy GG (2004) Lamellipodial versus filopodial mode of the actin nanomachinery: pivotal role of the filament barbed end. Cell 118(3): 363–373. doi:10.1016/j.cell.2004.07.019 DOI:dx.doi.org 33. Applewhite DA, Barzik M, Kojima S, Svitkina TM, Gertler FB, Borisy GG (2007) Ena/ VASP proteins have an anti-capping independent function in filopodia formation. Mol Biol Cell 18(7):2579–2591. doi:10.1091/mbc.E06-11-0990 DOI:dx.doi.org 34. Svitkina TM, Bulanova EA, Chaga OY, Vignjevic DM,Kojima S, Vasiliev JM, Borisy GG (2003) Mechanism of filopodia initiation by reorganization of a dendritic network. J Cell Biol 160(3):409–421. doi:10.1083/jcb.200210174 DOI:dx.doi.org 35. Vignjevic D, Kojima S, Aratyn Y, Danciu O, Svitkina T, Borisy GG (2006) Role of fascin in filopodial protrusion. J Cell Biol 174(6):863–875. doi:10.1083/jcb.200603013 DOI:dx.doi.org 36. Bear JE, Svitkina TM, Krause M, Schafer DA, Loureiro JJ, Strasser GA, Maly IV, Chaga OY, Cooper JA, Borisy GG, Gertler FB (2002) Antagonism between Ena/VASP proteins and actin filament capping regulates fibroblast motility. Cell 109(4):509–521. doi:10.1016/S00928674(02)00731-6 DOI:dx.doi.org 37. Ammer AG, Weed SA (2008) Cortactin branches out: roles in regulating protrusive actin dynamics. Cell Motil Cytoskeleton 65(9):687–707. doi:10.1002/cm.20296 DOI:dx.doi.org 38. Carlier MF, Le Clainche C, Wiesner S, Pantaloni D (2003) Actin-based motility: from molecules to movement. Bioessays 25(4):336–345. doi:10.1002/bies.10257 DOI:dx.doi.org 39. Pollard TD, Borisy GG (2003) Cellular motility driven by assembly and disassembly of actin filaments. Cell 112(4):453–465. doi:10.1016/S0092-8674(03)00120-X DOI:dx.doi.org
References
139
40. Pollard TD (2003) The cytoskeleton, cellular motility and the reductionist agenda. Nature 422(6933):741–745. doi:10.1038/nature01598 DOI:dx.doi.org 41. Yang C, Hoelzle M, Disanza A, Scita G, Svitkina T (2009) Coordination of membrane and actin cytoskeleton dynamics during filopodia protrusion. PLoS One 4(5):e5678 42. Verkhovsky AB, Svitkina TM, Borisy GG (1999) Network contraction model for cell translocation and retrograde flow. Biochem Soc Symp 65:207–222 43. Small JV, Resch GP (2005) The comings and goings of actin: coupling protrusion and retraction in cell motility. Curr Opin Cell Biol 17(5):517–523. doi:10.1016/j.ceb.2005.08.004 DOI:dx.doi.org 44. Birkenfeld J, Nalbant P, Yoon SH, Bokoch GM (2008) Cellular functions of GEF-H1, a microtubule-regulated Rho-GEF: is altered GEF-H1 activity a crucial determinant of disease pathogenesis? Trends Cell Biol 18(5):210–219. doi:10.1016/j.tcb.2008.02.006 DOI:dx.doi.org 45. Aoki K, Taketo MM (2007) Adenomatous polyposis coli (APC): a multi-functional tumor suppressor gene. J Cell Sci 120(Pt 19):3327–3335. doi:10.1242/jcs.03485 DOI:dx.doi.org 46. McCartney BM, Näthke IS (2008) Cell regulation by the Apc protein Apc as master regulator of epithelia. Curr Opin Cell Biol 20(2):186–193. doi:10.1016/j.ceb.2008.02.001 DOI:dx.doi.org 47. Krendel M, Zenke FT, Bokoch GM (2002) Nucleotide exchange factor GEF-H1 mediates cross-talk between microtubules and the actin cytoskeleton. Nat Cell Biol 4(4):294–301. doi:10.1038/ncb773 DOI:dx.doi.org 48. Fackler OT, Grosse R (2008) Cell motility through plasma membrane blebbing. J Cell Biol 181(6):879–884. doi:10.1083/jcb.200802081 DOI:dx.doi.org 49. Charras G, Paluch E (2008) Blebs lead the way: how to migrate without lamellipodia. Nat Rev Mol Cell Biol 9(9):730–736. doi:10.1038/nrm2453 DOI:dx.doi.org 50. Charras GT (2008) A short history of blebbing. J Microsc 231(3):466–478. doi:10.1111/ j.1365-2818.2008.02059.x DOI:dx.doi.org 51. Mogilner A, Keren K (2009) The shape of motile cells. Curr Biol 19(17):R762–71. doi:10.1016/j.cub.2009.06.053 DOI:dx.doi.org 52. Shutova MS, Alexandrova AY, Vasiliev JM (2008) Regulation of polarity in cells devoid of actin bundle system after treatment with inhibitors of myosin II activity. Cell Motil Cytoskeleton 65(9):734–46. doi:10.1002/cm.20295 DOI:dx.doi.org 53. Siegrist SE, Doe CQ (2007) Microtubule-induced cortical cell polarity. Genes Dev 21(5):483–496. doi:10.1101/gad.1511207 DOI:dx.doi.org 54. Omelchenko T, Vasiliev JM, Gelfand IM, Feder HH, Bonder EM (2002) Mechanisms of polarization of the shape of fibroblasts and epitheliocytes: Separation of the roles of microtubules and Rho-dependent actin-myosin contractility. Proc Natl Acad Sci U S A. 99(16):10452–10457. doi:10.1073/pnas.152339899 DOI:dx.doi.org 55. Kaverina I, Krylyshkina O, Small JV (1999) Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J Cell Biol 146(5):1033–1044. doi:10.1083/ jcb.146.5.1033 DOI:dx.doi.org 56. Small JV, Geiger B, Kaverina I, Bershadsky A (2002) How do microtubules guide migrating cells? Nat Rev Mol Cell Biol 3(12):957–964. doi:10.1038/nrm971 DOI:dx.doi.org 57. Rodionov VI, Gyoeva FK, Tanaka E, Bershadsky AD, Vasiliev JM, Gelfand VI (1993) Microtubule-dependent control of cell shape and pseudopodial activity is inhibited by the antibody to kinesin motor domain. J Cell Biol 123(6 Pt 2):1811–1820. doi:10.1083/ jcb.123.6.1811 DOI:dx.doi.org 58. Kaverina I, Krylyshkina O, Beningo K, Anderson K, Wang YL, Small JV (2002) Tensile stress stimulates microtubule outgrowth in living cells. J Cell Sci 115(Pt 11):2283–2291 59. Abercrombie M (1970) Contact inhibition in tissue culture. In vitro 7:128–142. doi:10.1007/ BF02616114 DOI:dx.doi.org 60. Gloushankova NA, Alieva NA, Krendel MF, Bonder EM, Feder HH, Vasiliev JM, Gelfand IM (1997) Cell-cell contact changes the dynamics of lamellar activity in nontransformed epitheliocytes but not in their ras-transformed descendants. Proc Natl Acad Sci USA 94(3):879–883. doi:10.1073/pnas.94.3.879 DOI:dx.doi.org
140
6 Cell Migration
61. Pokutta S, Drees F, Yamada S, Nelson WJ, Weis WI (2008) Biochemical and structural analysis of alpha-catenin in cell-cell contacts. Biochem Soc Trans 36(Pt 2):141–147. doi:10.1042/BST0360141 DOI:dx.doi.org 62. Ogita H, Ikeda W, Takai Y (2008) Roles of cell adhesion molecules nectin and nectin-like molecule-5 in the regulation of cell movement and proliferation. J Microsc 231(3):455–465. doi:10.1111/j.1365–2818.2008.02058.x DOI:dx.doi.org 63. Takai Y, Miyoshi J, Ikeda W, Ogita H (2008) Nectins and nectin-like molecules: roles in contact inhibition of cell movement and proliferation. Nat Rev Mol Cell Biol 9(8):603–615. doi:10.1038/nrm2457 DOI:dx.doi.org 64. Moustakas A, Heldin CH (2007) Signaling networks guiding epithelial-mesenchymal transitions during embryogenesis and cancer progression. Cancer Sci 98(10):1512–1520. doi:10.1111/j.1349-7006.2007.00550.x DOI:dx.doi.org 65. Yang J, Weinberg RA (2008) Epithelial-mesenchymal transition: at the crossroads of development and tumor metastasis. Dev Cell 14(6):818–829. doi:10.1016/j.devcel.2008.05.009 DOI:dx.doi.org 66. Acloque H, Adams MS, Fishwick K, Bronner-Fraser M, Nieto MA (2009) Epithelialmesenchymal transitions: the importance of changing cell state in development and disease. J Clin Invest 119(6):1438–1449. doi:10.1172/JCI38019 DOI:dx.doi.org 67. Kokkinos MI, Wafai R, Wong MK, Newgreen DF, Thompson EW, Waltham M (2007) Vimentin and epithelial-mesenchymal transition in human breast cancer--observations in vitro and in vivo. Cells Tissues Organs 185(1–3):191–203. doi:10.1159/000101320 DOI:dx.doi.org 68. Polette M, Mestdagt M, Bindels S, Nawrocki-Raby B, Hunziker W, Foidart JM, Birembaut P, Gilles C (2007) Beta-catenin and ZO-1: shuttle molecules involved in tumor invasionassociated epithelial-mesenchymal transition processes. Cells Tissues Organs 185(1–3): 61–65. doi:10.1159/000101304 DOI:dx.doi.org 69. De Wever O, Pauwels P, De Craene B, Sabbah M, Emami S, Redeuilh G, Gespach C, Bracke M, Berx G (2008) Molecular and pathological signatures of epithelial-mesenchymal transitions at the cancer invasion front. Histochem Cell Biol 130(3):481–494. doi:10.1007/ s00418-008-0464-1 DOI:dx.doi.org 70. Vasiliev JM (2008) Reorganization of molecular morphology of epitheliocytes and connectivetissue cells in morphogenesis and carcinogenesis. Biochemistry (Mosc) 73(5):528–531. doi:10.1134/S0006297908050052 DOI:dx.doi.org 71. Kedrin D, van Rheenen J, Hernandez L, Condeelis J, Segall JE (2007) Cell motility and cytoskeletal regulation in invasion and metastasis. J Mammary Gland Biol Neoplasia 12(2–3):143–52. doi:10.1007/s10911-007-9046-4 DOI:dx.doi.org 72. Sablina AA, Chumakov PM, Kopnin BP (2003) Tumor suppressor p53 and its homologue p73alpha affect cell migration. J Biol Chem 278(30):27362–27371. doi:10.1074/jbc. M300547200 DOI:dx.doi.org 73. Roger L, Gadea G, Roux P (2006) Control of cell migration: a tumour suppressor function for p53? Biol Cell 98(3):141–152. doi:10.1042/BC20050058 DOI:dx.doi.org 74. Gloushankova N, Ossovskaya V, Vasiliev J, Chumakov P, Kopnin B (1997) Changes in p53 expression can modify cell shape of ras-transformed fibroblasts and epitheliocytes. Oncogene 15(24):2985–2989. doi:10.1038/sj.onc.1201483 DOI:dx.doi.org 75. Alexandrova A, Ivanov A, Chumakov P, Kopnin B, Vasiliev J (2000) Changes in p53 expression in mouse fibroblasts can modify motility and extracellular matrix organization. Oncogene 19(50):5826–5830. doi:10.1038/sj.onc.1203944 DOI:dx.doi.org 76. Broussard JA, Webb DJ, Kaverina I (2008) Asymmetric focal adhesion disassembly in motile cells. Curr Opin Cell Biol 20(1):85–90. doi:10.1016/j.ceb.2007.10.009 DOI:dx.doi.org 77. Lock JG, Wehrle-Haller B, Strömblad S (2008) Cell-matrix adhesion complexes: master control machinery of cell migration. Semin Cancer Biol 18(1):65–76. doi:10.1016/j. semcancer.2007.10.001 DOI:dx.doi.org 78. Alexandrova AY, Arnold K, Schaub S, Vasiliev JM, Meister JJ, Bershadsky AD, Verkhovsky AB (2008) Comparative dynamics of retrograde actin flow and focal adhesions: formation of nascent adhesions triggers transition from fast to slow flow. PLoS One 3(9):e3234
References
141
79. Giannone G, Mège RM, Thoumine O (2009) Multi-level molecular clutches in motile cell processes. Trends Cell Biol 19(9):475–486. doi:10.1016/j.tcb.2009.07.001 DOI:dx.doi.org 80. Páez MC, González MJ, Serrano NC, Shoenfeld Y, Anaya JM (2007) Physiological and pathological implications of laminins: from the gene to the protein. Autoimmunity 40(2):83–94. doi:10.1080/08916930600911519 DOI:dx.doi.org 81. Tzu J, Marinkovich MP (2008) Bridging structure with function: structural, regulatory, and developmental role of laminins. Int J Biochem Cell Biol 40(2):199–214. doi:10.1016/j. biocel.2007.07.015 DOI:dx.doi.org 82. Miner JH (2008) Laminins and their roles in mammals. Microsc Res Tech 71(5):349–356. doi:10.1002/jemt.20563 DOI:dx.doi.org 83. Durbeej M (2010) Laminins. Cell Tissue Res 339(1):259–268. doi:10.1007/s00441-0090838-2 DOI:dx.doi.org 84. Guess CM, Quaranta V (2009) Defining the role of laminin-332 in carcinoma. Matrix Biol 28(8):445–455. doi:10.1016/j.matbio.2009.07.008 DOI:dx.doi.org 85. Hamill KJ, Paller AS, Jones JC (2010) Adhesion and migration, the diverse functions of the laminin alpha3 subunit. Dermatol Clin 28(1):79–87. doi:10.1016/j.det.2009.10.009 DOI:dx.doi.org 86. Marinkovich MP (2007) Tumour microenvironment: laminin 332 in squamous-cell carcinoma. Nat Rev Cancer 7(5):370–380. doi:10.1038/nrc2089 DOI:dx.doi.org 87. Tsuruta D, Kobayashi H, Imanishi H, Sugawara K, Ishii M, Jones JC (2008) Laminin-332integrin interaction: a target for cancer therapy? Curr Med Chem 15(20):1968–1975. doi:10.2174/092986708785132834 DOI:dx.doi.org 88. Miyazaki K (2006) Laminin-5 (laminin-332): Unique biological activity and role in tumor growth and invasion. Cancer Sci 97(2):91–98. doi:10.1111/j.1349-7006.2006.00150.x DOI:dx.doi.org 89. Chen P, Parks WC (2009) Role of matrix metalloproteinases in epithelial migration. J Cell Biochem 108(6):1233–1243. doi:10.1002/jcb.22363 DOI:dx.doi.org 90. Hintermann E, Quaranta V (2004) Epithelial cell motility on laminin-5: regulation by matrix assembly, proteolysis, integrins and erbB receptors. Matrix Biol 23(2):75–85. doi:10.1016/j. matbio.2004.03.001 DOI:dx.doi.org 91. Sugawara K, Tsuruta D, Ishii M, Jones JC, Kobayashi H (2008) Laminin-332 and -511 in skin. Exp Dermatol 17(6):473–480. doi:10.1111/j.1600-0625.2008.00721.x DOI:dx.doi.org 92. Fujiwara H, Gu J, Sekiguchi K (2004) Rac regulates integrin-mediated endothelial cell adhesion and migration on laminin-8. Exp Cell Res 292(1):67–77. doi:10.1016/j.yexcr.2003.08.010 DOI:dx.doi.org 93. Stipp CS (2010) Laminin-binding integrins and their tetraspanin partners as potential antimetastatic targets. Expert Rev Mol Med 12:e3. doi:10.1017/S1462399409001355 DOI:dx.doi.org 94. Caswell P, Norman J (2008) Endocytic transport of integrins during cell migration and invasion. Trends Cell Biol 18(6):257–263. doi:10.1016/j.tcb.2008.03.004 DOI:dx.doi.org 95. Caswell PT, Vadrevu S, Norman JC (2009) Integrins: masters and slaves of endocytic transport. Nat Rev Mol Cell Biol 10(12):843–853. doi:10.1038/nrm2799 DOI:dx.doi.org 96. Ulrich F, Heisenberg CP (2009) Trafficking and cell migration. Traffic 10(7):811–818. doi:10.1111/j.1600-0854.2009.00929.x DOI:dx.doi.org 97. Schlaepfer DD, Mitra SK (2004) Multiple connections link FAK to cell motility and invasion. Curr Opin Genet Dev 14(1):92–101. doi:10.1016/j.gde.2003.12.002 DOI:dx.doi.org 98. Kruchten AE, McNiven MA (2006) Dynamin as a mover and pincher during cell migration and invasion. J Cell Sci 119(Pt 9):1683–1690. doi:10.1242/jcs.02963 DOI:dx.doi.org 99. Mitra SK, Schlaepfer DD (2006) Integrin-regulated FAK-Src signaling in normal and cancer cells. Curr Opin Cell Biol 18(5):516–523. doi:10.1016/j.ceb.2006.08.011 DOI:dx.doi.org 100. Defilippi P, Di Stefano P, Cabodi S (2006) p130Cas: a versatile scaffold in signaling networks. Trends Cell Biol 16(5):257–63. doi:10.1016/j.tcb.2006.03.003 DOI:dx.doi.org 101. Sharma A, Mayer BJ (2008) Phosphorylation of p130Cas initiates Rac activation and membrane ruffling. BMC Cell Biol 9(1):50. doi:10.1186/1471-2121-9-50 DOI:dx.doi.org 102. Enomoto A, Ping J, Takahashi M (2006) Girdin, a novel actin-binding protein, and its family of proteins possess versatile functions in the Akt and Wnt signaling pathways. Ann N Y Acad Sci 1086:169–184. doi:10.1196/annals.1377.016 DOI:dx.doi.org
142
6 Cell Migration
103. Stambolic V, Woodgett JR (2006) Functional distinctions of protein kinase B/Akt isoforms defined by their influence on cell migration. Trends Cell Biol 16(9):461–466. doi:10.1016/j. tcb.2006.07.001 DOI:dx.doi.org 104. Hall A (2005) Rho GTPases and the control of cell behaviour. Biochem Soc Trans 33 (Pt 5):891–895. doi:10.1042/BST20050891 DOI:dx.doi.org 105. Charest PG, Firtel RA (2007) Big roles for small GTPases in the control of directed cell movement. Biochem J 401(2):377–390. doi:10.1042/BJ20061432 DOI:dx.doi.org 106. Juliano RL, Reddig P, Alahari S, Edin M, Howe A, Aplin A (2004) Integrin regulation of cell signalling and motility. Biochem Soc Trans 32(Pt3):443–446. doi:10.1042/BST0320443 DOI:dx.doi.org 107. Suresh K Alahari, Peter J Reddig, Rudy L Juliano (2004) The integrin-binding protein Nischarin regulates cell migration by inhibiting PAK. The EMBO Journal 23: 2777–2788. doi:10.1038/sj.emboj.7600291 DOI:dx.doi.org 108. Casazza A, Fazzari P, Tamagnone L (2007) Semaphorin signals in cell adhesion and cell migration: functional role and molecular mechanisms. Adv Exp Med Biol 600:90–108. doi:10.1007/978-0-387-70956-7_8 DOI:dx.doi.org 109. Tran TS, Kolodkin AL, Bharadwaj R (2007) Semaphorin regulation of cellular morphology. Annu Rev Cell Dev Biol 23:263–292. doi:10.1146/annurev.cellbio.22.010605.093554 DOI:dx.doi.org 110. Püschel AW (2007) GTPases in semaphorin signaling. Adv Exp Med Biol 600:12–23. doi:10.1007/978-0-387-70956-7_2 DOI:dx.doi.org 111. Brooks SA, Lomax-Browne HJ, Carter TM, Kinch CE, Hall DM (2010) Molecular interactions in cancer cell metastasis. Acta Histochem 112(1):3–25. doi:10.1016/j.acthis.2008.11.022 DOI:dx.doi.org 112. Geiger TR, Peeper DS (2009) Metastasis mechanisms. Biochim Biophys Acta 1796(2): 293–308. doi:10.1016/j.bbcan.2009.07.006 DOI:dx.doi.org 113. Machesky LM (2008) Lamellipodia and filopodia in metastasis and invasion. FEBS Lett 582(14):2102–2111. doi:10.1016/j.febslet.2008.03.039 DOI:dx.doi.org 114. Yamaguchi H, Wyckoff J, Condeelis J (2005) Cell migration in tumors. Curr Opin Cell Biol 17(5):559–564. doi:10.1016/j.ceb.2005.08.002 DOI:dx.doi.org 115. Kopnin BP (2000) Targets of oncogenes and tumor suppressors: key for understanding basic mechanisms of carcinogenesis. Biochemistry (Mosc) 65(1):2–27 116. Karlsson R, Pedersen ED, Wang Z, Brakebusch C (2009) Rho GTPase function in tumorigenesis. Biochim Biophys Acta 1796(2):91–98. doi:10.1016/j.bbcan.2009.03.003 DOI:dx.doi.org 117. Tikhmyanova N, Little JL, Golemis EA (2010) CAS proteins in normal and pathological cell growth control. Cell Mol Life Sci 67(7):1025–1048. doi:10.1007/s00018-009-0213-1 DOI:dx.doi.org 118. Sahai E, Olson MF, Marshall CJ (2001) Cross-talk between Ras and Rho signalling pathways in transformation favours proliferation and increased motility. EMBO J 20(4):755–766. doi:10.1093/emboj/20.4.755 DOI:dx.doi.org 119. Lee S, Helfman DM (2004) Cytoplasmic p21Cip1 is involved in Ras-induced inhibition of the ROCK/LIMK/cofilin pathway. J Biol Chem.279(3):1885–1891. doi:10.1074/jbc. M306968200 DOI:dx.doi.org 120. Desgrosellier JS, Cheresh DA (2010) Integrins in cancer: biological implications and therapeutic opportunities. Nat Rev Cancer 10(1):9–22. doi:10.1038/nrc2748 DOI:dx.doi.org 121. Alexandrova AY (2008) Evolution of cell interactions with extracellular matrix during carcinogenesis. Biochemistry (Mosc) 73(7):733–741 122. Ljubimova JY, Fujita M, Khazenzon NM, Ljubimov AV, Black KL (2006) Changes in laminin isoforms associated with brain tumor invasion and angiogenesis. Front Biosci 11:81–88. doi:10.2741/1781 DOI:dx.doi.org 123. Fujita M, Khazenzon NM, Ljubimov AV, Lee BS, Virtanen I, Holler E, Black KL, Ljubimova JY (2006) Inhibition of laminin-8 in vivo using a novel poly(malic acid)-based carrier reduces glioma angiogenesis. Angiogenesis 9(4):183–191. doi:10.1007/s10456-006-9046-9 DOI:dx.doi.org
References
143
124. Khazenzon NM, Ljubimov AV, Lakhter AJ, Fujita M, Fujiwara H, Sekiguchi K, Sorokin LM, Petäjäniemi N, Virtanen I, Black KL, Ljubimova JY (2003) Antisense inhibition of laminin-8 expression reduces invasion of human gliomas in vitro. Mol Cancer Ther 2(10):985–994 125. Kariya Y, Kariya Y, Gu J (2009) Roles of laminin-332 and alpha6beta4 integrin in tumor progression. Mini Rev Med Chem 9(11):1284–1291. doi:10.2174/138955709789878114 DOI:dx.doi.org 126. Waterman EA, Sakai N, Nguyen NT, Horst BA, Veitch DP, Dey CN, Ortiz-Urda S, Khavari PA, Marinkovich MP (2007) A laminin-collagen complex drives human epidermal carcinogenesis through phosphoinositol-3-kinase activation. Cancer Res 67(9):4264–4270. doi:10.1158/0008-5472.CAN-06-4141 DOI:dx.doi.org 127. Guarino M (2007) Epithelial-mesenchymal transition and tumour invasion. Int J Biochem Cell Biol 39(12):2153–2160. doi:10.1016/j.biocel.2007.07.011 DOI:dx.doi.org 128. Klymkowsky MW, Savagner P (2009) Epithelial-mesenchymal transition: a cancer researcher’s conceptual friend and foe. Am J Pathol 174(5):1588–1593. doi:10.2353/ ajpath.2009.080545 DOI:dx.doi.org 129. Voulgari A, Pintzas A (2009) Epithelial-mesenchymal transition in cancer metastasis: mechanisms, markers and strategies to overcome drug resistance in the clinic. Biochim Biophys Acta 1796(2):75–90. doi:10.1016/j.bbcan.2009.03.002 DOI:dx.doi.org 130. Yilmaz M, Christofori G (2009) EMT, the cytoskeleton, and cancer cell invasion. Cancer Metastasis Rev 28(1–2):15–33. doi:10.1007/s10555-008-9169-0 DOI:dx.doi.org 131. Vergara D, Merlot B, Lucot JP, Collinet P, Vinatier D, Fournier I, Salzet M (2010) Epithelialmesenchymal transition in ovarian cancer. Cancer Lett 291(1):59–66. doi:10.1016/j. canlet.2009.09.017 DOI:dx.doi.org 132. Moustakas A, Heldin CH (2007) Signaling networks guiding epithelial-mesenchymal transitions during embryogenesis and cancer progression. Cancer Sci 98(10):1512–1520. doi:10.1111/j.1349-7006.2007.00550.x DOI:dx.doi.org 133. Imamichi Y, Menke A (2007) Signaling pathways involved in collagen-induced disruption of the E-cadherin complex during epithelial-mesenchymal transition. Cells Tissues Organs 185(1–3):180–190. doi:10.1159/000101319 DOI:dx.doi.org 134. Guarino M, Rubino B, Ballabio G (2007) The role of epithelial-mesenchymal transition in cancer pathology. Pathology 39(3):305–318. doi:10.1080/00313020701329914 DOI:dx.doi.org 135. Giehl K, Menke A (2008) Microenvironmental regulation of E-cadherin-mediated adherens junctions. Front Biosci 13:3975–85. doi:10.2741/2985 DOI:dx.doi.org 136. Zhang X, Nie D, Chakrabarty S (2010) Growth factors in tumor microenvironment. Front Biosci 15:151–165. doi:10.2741/3612 DOI:dx.doi.org 137. Hanna JA, Bordeaux J, Rimm DL, Agarwal S (2009) The function, proteolytic processing, and histopathology of Met in cancer. Adv Cancer Res 103:1–23. doi:10.1016/S0065230X(09)03001-2 DOI:dx.doi.org 138. Appelmann I, Liersch R, Kessler T, Mesters RM, Berdel WE (2010) Angiogenesis inhibition in cancer therapy: platelet-derived growth factor (PDGF) and vascular endothelial growth factor (VEGF) and their receptors: biological functions and role in malignancy. Recent Results Cancer Res 180:51–81. doi:10.1007/978-3-540-78281-0_5 DOI:dx.doi.org 139. Matsumoto K, Nakamura T, Sakai K, Nakamura T (2008) Hepatocyte growth factor and Met in tumor biology and therapeutic approach with NK4. Proteomics 8(16):3360–3370. doi:10.1002/pmic.200800156 DOI:dx.doi.org 140. Korc M, Friesel RE (2009) The role of fibroblast growth factors in tumor growth. Curr Cancer Drug Targets 9(5):639–651. doi:10.2174/156800909789057006 DOI:dx.doi.org 141. Ivy SP, Wick JY, Kaufman BM (2009) An overview of small-molecule inhibitors of VEGFR signaling. Nat Rev Clin Oncol 6(10):569–579. doi:10.1038/nrclinonc.2009.130 DOI:dx.doi.org 142. Mitsudomi T, Yatabe Y (2010) Epidermal growth factor receptor in relation to tumor development: EGFR gene and cancer. FEBS J 277(2):301–308. doi:10.1111/j.1742-4658. 2009.07448.x DOI:dx.doi.org 143. Lu X, Kang Y (2010) Epidermal growth factor signalling and bone metastasis. Br J Cancer 102(3):457–461. doi:10.1038/sj.bjc.6605490 DOI:dx.doi.org
144
6 Cell Migration
144. Feldner JC, Brandt BH (2002) Cancer cell motility--on the road from c-erbB-2 receptor steered signaling to actin reorganization. Exp Cell Res 272(2):93–108. doi:10.1006/ excr.2001.5385 DOI:dx.doi.org 145. Govindan R (2010) A review of epidermal growth factor receptor/HER2 inhibitors in the treatment of patients with non-small-cell lung cancer. Clin Lung Cancer 11(1):8–12. doi:10.3816/CLC.2010.n.001 DOI:dx.doi.org 146. Halper J (2010) Growth factors as active participants in carcinogenesis: a perspective. Vet Pathol 47(1):77–97. doi:10.1177/0300985809352981 DOI:dx.doi.org 147. Rosell R, Viteri S, Molina MA, Benlloch S, Taron M (2010) Epidermal growth factor receptor tyrosine kinase inhibitors as first-line treatment in advanced nonsmall-cell lung cancer. Curr Opin Oncol 22(2):112–120. doi:10.1097/CCO.0b013e32833500d2 DOI:dx.doi.org 148. Okamoto I (2010) Epidermal growth factor receptor in relation to tumor development: EGFRtargeted anticancer therapy. FEBS J 277(2):309–315. doi:10.1111/j.1742-4658.2009. 07449.x DOI:dx.doi.org
Chapter 7
Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
Abstract The cells are capable of selecting between substratum surfaces that are poorly adhesive and highly adhesive for these cells: they spread on and migrate over the surfaces that are relatively more adhesive. As a result of this active selection, the cells at the substratum surfaces that have heterogeneous adhesiveness are accumulated at the isolated adhesive “islets” or are strongly elongated along narrow adhesive “paths.” The selection is based on the polarization of pseudopodial activity of the cells. In an organism, various tissue surfaces with which fibroblastic or epithelial cells enter into contact can have different degrees of adhesiveness for these cells. With the possibility to select between the surfaces with low or high adhesiveness, the cells always “prefer” to attach to, to spread on, and to migrate over the surfaces that are relatively more adhesive for these cells. For example, fibroblastic cells demonstrate active selection between the surface of mesothelium, which is poorly adhesive for them, and highly adhesive sub-mesothelium structures of the extracellular matrix: basement membranes and subjacent fibrous connective tissue [1]. Mesothelium is a continuous dense layer of epithelial cells covering tunicae serosae of the peritoneal, pleural or pericardial cavities (Fig. 7.1). Mesothelium is poorly adhesive for many types of cells: they do not attach to the upper surface of the mesothelium covering. However, in the areas of mesothelium regeneration its cells become retracted (Fig. 7.2), and the “islets” of the subjacent basement membrane surface are exposed between the retracted mesothelial cells (Fig. 7.3). The basement membrane is highly adhesive for the attachment and spreading of many types of normal and tumor cells (Fig. 7.4). Thus, the mosaic consisting of low-adhesive (upper surface of mesothelium cells) and high-adhesive “islets” (surface of the basement membrane) is formed on the regenerating mesothelium. Fibroblastic cells settled to the surface of the regenerating mesothelium make active selection: the cell processes are attached to the exposed small sections of the basement membrane, creeping between and under the retracted mesothelium cell (Fig. 7.5a, b). On the surface of the peritoneum, one can see minute openings with the diameters from 5 to 30 mm. In these holes, called “stomata,” the mesothelium and the basement membrane are absent, and only the fibers of the subjacent connective tissue are Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_7, © Springer Science+Business Media, LLC 2011
145
146
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
Fig. 7.1 Mouse parietal peritoneum. The continuous dense layer of mesothelial cells with microvillous surface relief. SEM. Scale bar, 41.5 mm
Fig. 7.2 Mouse parietal peritoneum. The regeneration of the mesothelium: the retracted cells. SEM. Scale bar, 30 mm
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
147
Fig. 7.3 Mouse parietal peritoneum. The regeneration of the mesothelium: the retracted mesotheliocytes with retraction fibers (arrows) coming out of the cell edges; the subjacent basement membrane (double arrow) between the retracted cells is exposed. SEM. Scale bar, 10 mm
Fig. 7.4 Fibroblastic mouse cell attached to and spreading on the basement membrane surface of mouse parietal peritoneum. The cell is at the stage of radial spreading. SEM. Scale bar, 8 mm
148
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
Fig. 7.5 (a, b) Fibroblastic mouse cells spread on the regenerating mesothelium of mouse parietal peritoneum. The fibroblastic cells (arrows) are attached to the subjacent basement membrane between the retracted mesotheliocytes (double arrows) (a); the fibroblastic cell process (arrow) creeps between and under the mesotheliocytes (b). SEM. Scale bars, 20 mm (a), 10 mm (b)
Fig. 7.6 Mouse visceral peritoneum. “Stoma”: the hole where the mesothelium and the basement membrane are absent (arrow). There are the subjacent connective tissue fibers in the depth of the “stoma” depth. SEM. Scale bar, 4.5 mm
exposed (Fig. 7.6). These fibers like basement membrane are high-adhesive for many types of normal and tumor cells (Figs. 7.7–7.9). Therefore, peritoneum stomata are high-adhesive “islets” for the selective cell attachment (Fig. 7.10). It is obvious that if necessary components of the extracellular matrix are distributed in the form of “islets” or narrow “paths,” then cells can spread and move only in the range of such adhesive sections. This picture is actually observed in the
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths” Fig. 7.7 Fibroblastic mouse cell attached to the connective tissue fibers of mouse parietal peritoneum. The cell is in the beginning of spreading: filopodia (arrows) coming out of the cell base. SEM. Scale bar, 6 mm
Fig. 7.8 Fibroblastic mouse cells completely spread on the connective tissue fibers of mouse parietal peritoneum. SEM. Scale bar, 22.5 mm
149
150
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
Fig. 7.9 Ascites epithelial tumor mouse cells attached to the connective tissue fibers of mouse parietal peritoneum. SEM. Scale bar, 10 mm Fig. 7.10 Ascites epithelial tumor mouse cells attached to the connective tissue fibers inside the “stoma.” SEM. Scale bar, 7 mm
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
151
Fig. 7.11 Fibroblastic mouse cells on the special nonadhesive substratum with narrow adhesive “paths.” The cells acquire strongly elongated shapes and are oriented along the “paths.” LM (light microscopy). Scale bar, 50 mm
o rganism during embryogenesis or wound healing: cells migrate directionally along the linear sections of the matrix in accordance with the presence of matrix protein components, necessary for the attachment of cells of this type. The influence of adhesive “paths” or “islets” on cell behavior can be modeled by means of cell cultivation on special artificial substrata, which possess plain, chemically heterogeneous, surfaces. At first, the substratum surfaces are made nonadhesive for cell attachment using special chemical methods. Afterward, adhesive extracellular matrix protein coatings in a form of narrow “paths” or isolated “islets” of different sizes are created on these nonadhesive surfaces [2]. Fibroblastic cells on narrow (25–35 mm wide) adhesive “paths” acquire elongated fusiform shapes and are getting oriented along the “paths” (Fig. 7.11). The active selection of more adhesive surfaces is made by fibroblastic cells owing to the polarization of their pseudopodial activity: repeated formation of pseudopodia continues only in those parts of the cell edge where this process leads to the formation of new stable cell contacts with the extracellular matrix. On the narrow adhesive “paths,” the cell spreading proceeds without constraints only along the “path”; the nonadhesive “path” borders limit the spreading in the perpendicular direction. The longitudinal spreading leads to the longitudinal cell stretching; this causes the corresponding alignment of actin filament bundles in the cells [2]. The maximal length of a fibroblastic cell on a narrow adhesive “path” is not changed as compared with the maximal length of the cell spread on the evenly adhesive surface.
152
7 Cell Responses to Chemical Heterogeneity of Substrata: Adhesive “Islets” or “Paths”
It is due to the balance of two opposite forces: elongation-promoting action of microtubules, which induce continuous extensions of lamellipodia at the anterior cell edge (see Sect. 6.1.1) promoting elongation of the cell along the “path,” and the opposite contractile action of actin-myosin cortex [2, 3]. Therefore, fibroblastic cells on narrow adhesive “paths” can strongly diminish their transversal spreading not changing their longitudinal spreading. If cell-adhesive “islets” have too small sizes, then the multiplication of cells, which are attached on such islets, is reduced; the cells can even undergo apoptosis. The cells cannot sufficiently spread on too small adhesive areas (see Fig. 5.11a, b in Chap. 5); therefore, the number of focal adhesions formed by the cell is diminished. As a result, the transduction of integrin-mediated intracellular signals, which regulate the functional activity of the cell, including its proliferation (see Sect. 5.3), is significantly weakened.
References 1. Rovensky YA, Gvichiya AS, Vasiliev JM (1980) SEM study of the attachment of mouse ascitic hepatoma cells to various substrata. Scan Electron Microsc (3):71–78 2. Levina EM, Kharitonova MA, Rovensky YA, Vasiliev JM (2001) Cytoskeletal control of fibroblast length: experiments with linear strips of substrate. J Cell Sci 114(Pt 23):4335–4341 3. Kharitonova MA, Vasiliev JM (2008) Controlling cell length. Semin Cell Dev Biol. 19(6):480–484. doi:10.1016/j.semcdb.2008.07.008 DOI:dx.doi.org
Chapter 8
Topographic Cell Responses
Abstract Tissue cells have an ability to respond not only to biochemical characteristics of the extracellular matrix, but also to its geometric configuration, the matrix surface topography. Such topographic features as curvature, orderly grooves and ridges, discontinuities, or some other microscale or nanoscale geometries affect cell adhesion and spreading, cell shapes and orientation, direction of migration, and also proliferation and synthetic activities of the cells. The cell responses to the geometric configuration of the substratum surface, which the cells are attached to and spread on so-called “topographic cell responses,” are altered as a result of oncogenic transformation. As long ago as the beginning of last century, it was shown that if the isolated nerve cells are cultured on the cobweb, these cells get elongated and oriented along the cobweb threads. Later, a similar cell response was described in fibroblastic cells, cultured on protein fibers from the blood plasma clot. More recently, such an exotic substratum as fish scale, which possesses a folded surface, was used for cell culture. Fibroblastic cells on the fish scale got elongated and oriented along its folds. The described phenomenon was termed contact guidance of cells [1]. Tissue cells in their natural environment interact with the extracellular matrix structures that have various geometric features, such as a surface curvature, grooves, ridges, pits, and also discontinuities (e.g., holes). Various geometric features of micrometer or nanometer size comprise the surface topography of the extracellular matrix. For example, the surface topography of basement membranes includes cylindrical formations (fibers), ridges, and nanometer size pores [2, 3]. Tissue cells have the ability to “feel” the geometric characteristics of the extracellular matrix and respond to them. The matrix surface topography is a powerful modulator of cell behavior. Micro and nano-topography of the matrix, like its chemical characteristics, determines the possibility for the cells to attach selectively to certain areas of the matrix, induces cytoskeletal reorganizations, and affects cell shape, orientation, migration, and proliferation, and also gene expression.
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_8, © Springer Science+Business Media, LLC 2011
153
154
8 Topographic Cell Responses
Such cell responses to the surface topography of the extracellular matrix are termed topographic cell responses [4, 5]. To model topographic cell responses, the special artificial substrata are used for cell culture. These solid substrata models are chemically homogeneous and have various controlled surface topography patterns of micrometer or nanometer size. The microscale or nanoscale geometries, such as orderly linear grooves and ridges, pits and pillars, and other surface topography patterns, are produced by different methods including micromachining, photo- or electron beam lithography, chemical etching, and some others [6–11]. The model substrata can also constitute artificial cylindrical threads with different degree of their surface curvature and artificial discontinuous substrata containing large substratum-free zones. Aside from its biological role, the production of topographically patterned artificial solid substrata also has a goal of their possible usage as tissue implants.
8.1 Cylindrical Substrata The complex geometric configuration of the extracellular matrix includes various matrix structures having cylindrical shapes. The cylindrical shape of the extracellular matrix surfaces may influence cell behavior during embryogenesis, repair processes, and in cancer cell invasion. To study cell responses to the curvature of a cylindrical surface, the artificial cylindrical substrata, such as quartz glass threads with sufficiently high degrees of surface curvatures can be used for cell culture [12–14].
8.1.1 Normal Cell Responses The spreading of fibroblastic cells attached on cylindrical threads (Fig. 8.1) depends on the degree of surface curvature. On threads with high degree of surface curvature (cylinders of 24–32 mm in diameter), fibroblastic cell spreading is not the same as that on the plane surface (see Sect. 5.1.2). The stage of radial spreading is absent: circular lamellar cytoplasm is not formed, and the cells do not acquire discoid shapes. Instead of that, fibroblastic cells immediately become polarized and get elongated along the cylinders (Fig. 8.2). On the cylindrical threads with lesser degree of surface curvature (cylinders of 50 mm in diameter), fibroblastic cells at the stage of radial spreading acquire typical disc-like shapes similar to the ones observed on plane substrata. The radially spread disc-like cells are getting bent across the cylinders (Figs. 8.3 and 8.4). The cells do not undergo any significant elongation. At the subsequent stage of spreading, polarized fibroblastic cells get elongated and oriented along the cylinders (Fig. 8.5). These cells contain numerous straight actin filaments bundles, which are longitudinally aligned. Epithelial cells spread on the cylindrical threads retain their discoid shapes like the ones on plane substrata; the cells are bent across the cylinders (Fig. 8.6). Elongation
8.1 Cylindrical Substrata
155
Fig. 8.1 Fibroblastic mouse cell attached to the cylindrical substratum surface. The cell is at the initial stage of its spreading. SEM. Scale bar, 4 mm
Fig. 8.2 Fibroblastic mouse cell spreading on the cylindrical substratum with the high surface curvature. The cell “misses” the disc-like shape and is being elongated along the cylinder. SEM. Scale bar, 7.5 mm
156
8 Topographic Cell Responses
Fig. 8.3 Fibroblastic mouse cell at the stage of radial spreading on the cylindrical substratum with the low surface curvature. SEM. Scale bar, 6.7 mm
Fig. 8.4 Fibroblastic mouse cell in the termination of the radial spreading stage on the cylindrical substratum surface. The discoid cell is bent across the cylinder. SEM. Scale bar, 12 mm
8.1 Cylindrical Substrata
157
Fig. 8.5 Fibroblastic mouse cells in the polarization stage of their spreading on the cylindrical substratum surface. The cells are elongated and oriented along the cylinder. SEM. Scale bar, 10 mm
Fig. 8.6 Epithelial mouse cell spread on the cylindrical substratum surface. The cell is bent across the cylinder. SEM. Scale bar, 25 mm
158
8 Topographic Cell Responses
Fig. 8.7 Epithelial mouse cell spread on the cylindrical substratum surface. Straight actin microfilament bundles (arrow) oriented transversely to the cylinder’s axis. Staining for actin. FM. Scale bar, 32 mm
of these cells and their orientation along the cylinders are weak. Their actin cytoskeleton undergoes reorganization: in addition to circular filament bundle along the entire cell periphery, which is typical for single epithelial cells spread on plane substrata (see Fig. 3.3 in Chap. 3), the epithelial cells on cylindrical substrata acquire numerous straight bundles oriented mainly transversely to the cylinder axis (Fig. 8.7).
8.1.2 Transformed Cell Responses Transformed fibroblasts on cylindrical threads do not undergo significant elongation compared to the ones on plane substrata; there is no cell orientation along the cylinder (Fig. 8.8). Relatively more flattened transformed fibroblasts are getting bent across the cylinders (Fig. 8.9). In transformed epithelial cells spread on the cylindrical threads, elongation is considerably increased compared to the ones on plane substrata; the cells get orientated along the cylinders (Fig. 8.10) and develop a longitudinal pattern of their actin filament bundles (Fig. 8.11). Therefore, the cylindrical form of substratum with a sufficiently high degree of curvature can affect cell shapes, cell orientation and actin cytoskeleton organization. Normal fibroblastic cells are elongated and oriented along the cylinders, whereas single epithelial cells are prone to wrap around the cylinders and reorganize their circular filament bundles into straight transversely oriented ones. Transformed epitheliocytes respond the same way as fibroblastic cells.
8.1 Cylindrical Substrata
159
Fig. 8.8 Transformed mouse fibroblasts spread on the cylindrical substratum surface. No cell elongation or orientation along the cylinder. SEM. Scale bar, 8.5 mm
Fig. 8.9 Transformed mouse fibroblasts spread on the cylindrical substratum surface. The cells are bent across the cylinder. SEM. Scale bar, 9.5 mm
Fig. 8.10 Transformed mouse epitheliocytes spread on the cylindrical substratum surface. The cells are elongated and oriented along the cylinder. SEM. Scale bar, 18 mm
Fig. 8.11 Transformed mouse epitheliocyte elongated and oriented along the cylindrical substratum. The longitudinal pattern of the straight actin microfilament bundles (arrow). Staining for actin. FM. Scale bar, 15 mm. Reproduced with permission from the Experimental Cell Research (ref. [14])
8.2 Grooved Substrata
161
Thus, in contrast to normal epithelial cells, motile transformed epitheliocytes can be oriented and guided by the cylindrical structures of the extracellular matrix. These structures may serve as shaped substrata guiding the invasion of cancer cells.
8.2 Grooved Substrata The grooved surface topography of the extracellular matrix can be modeled by the special artificial substrata for cultured cells carrying the fabricated orderly linear grooves. The substratum surface topography can be nanoscale if the depths and widths of the grooves, and also the intervals between them are less than one micrometer. The grooved topography is microscale if these features range from one to several micrometers.
8.2.1 Normal Cell Responses Both nanoscale and microscale grooved substratum topography influence the shape, alignment, and locomotion of normal cultured cells. Fibroblastic cells respond to the groove depths and widths ranging from a few tenths of a micrometer to several micrometers: the cells become elongated and show orientation parallel to the grooves [15–17]. The orientation of fibroblastic cells is preceded by the alignment of their microtubules parallel to the direction of the grooves; thus, microtubules may determine the groove-dependent cell orientation [18]. The microscale grooves also affect the direction and velocity of fibroblastic cell migration [19]. Epithelial cells are also influenced by nanoscale grooved topography: the groove depths of hundredths or thousandths of a micrometer affect cell alignment in the direction parallel to the grooves. Epitheliocytes respond to the grooves as shallow as 0.014 mm [20, 21]. Special types of microscale grooved topography are grooves of large depths and having triangle cross-sections [22, 23]. Some of such grooved topographies are linear grooves, the depths of which are tens of micrometers (from 25 to 65 mm), and distributed parallel to each other or hexagonally. The spaces between neighboring parallel grooves are also large, from one hundred to a few hundred micrometers, and the surfaces in these spaces have cylindrical shapes (Fig. 8.12). Hexagonally distributed grooves form hexagons; the grooves are sides of the hexagons. The distances between two opposite sides of a hexagon are several hundred micrometers, and the surface between grooves has a spherical shape (Fig. 8.13b). Both fibroblastic and epithelial cells respond to deep grooves having triangle cross-sections [22–24]. The cells immediately after their settling are mostly accumulated at the groove bottoms (Figs. 8.13a–c and 8.14). After the spreading is complete, fibroblastic cells begin to directionally migrate from the bottoms to the side slopes of the grooves
162
8 Topographic Cell Responses
Fig. 8.12 The profile of the grooved substratum: the parallel grooves of triangle cross-sections (arrow) and the cylindrical substratum surfaces (double arrow) between the neighbor grooves. LM. Scale bar, 70 mm
Fig. 8.13 (a–c) Fibroblastic mouse cells settled onto the grooved substrata. The cells (arrows) are accumulated at the bottoms of the parallel grooves (a) or of the grooves distributed hexagonally (b, c). LM. Scale bars, 160 mm (a, b). SEM. Scale bar, 180 mm (c). Reproduced with permission from the Experimental Cell Research (refs. [22, 23])
(Fig. 8.15a, b). The migration is fast and directional: already in 2–3 h, the 20–25 mm zones free of the cells are formed on both sides of the groove bottoms. Finally in 24–36 h, almost all fibroblastic cells are accumulated in the areas between the grooves: on cylindrical substratum surfaces between parallel grooves (Fig. 8.16a) or on spherical surfaces of the hexagons (Fig. 8.16b, c).
8.2 Grooved Substrata
163
Fig. 8.14 Fibroblastic mouse cell at the groove bottom. The cell is at the initial stage of its spreading: filopodia (arrows) at the base of the spherical cell are attached to the side slopes of the groove. SEM. Scale bar, 4.5 mm. Reproduced with permission from the Experimental Cell Research (ref. [23])
Most of the cells located on cylindrical surfaces are oriented to the direction of the grooves (Fig. 8.16a). The capability to migrate from deep grooves of triangle cross-sections is different in fibroblastic cells of various species: this capability is high in mouse, rat, and chicken cells, it is less pronounced in hamster and human ones. Unlike fibroblastic cells, the migration of epithelial cells is very weak, and in 24–36 h, almost all of settled epitheliocytes remain in the deep grooves being spread on the groove bottoms.
8.2.2 Transformed Cell Responses Transformed epithelial cells keep the ability to respond to nanoscale grooves: the cells are oriented and migrate along the grooves; for instance, the neuroblastoma cells are sensitive to nanoscale pores of less than 0.02 mm [25–27]. On the microscale grooves of large depths and having triangle cross-sections, transformed fibroblasts, like normal fibroblastic cells, are initially found mostly at the bottoms of deep grooves. In 2–3 h, part of the spreading cells begins to migrate to the side slopes of the grooves. In contrast to normal fibroblastic cells, the
164
8 Topographic Cell Responses
Fig. 8.15 (a, b) Fibroblastic mouse cells migrating from the grooves. The cells (arrows) migrate from the bottoms of the parallel grooves (a) or from the grooves distributed hexagonally (b). LM. Scale bar, 160 mm. Reproduced with permission from the Experimental Cell Research (ref. [22])
igration of transformed fibroblasts from the grooves is not directional: cell m movements are irregular, many cells return from the slopes to the bottoms of the grooves and cross them [24]. Finally, many migrating cells occupy the areas between the grooves; however, numerous spread transformed cells remain in the depths of the grooves. These cells make “bridges” over the groove bottoms attaching to the opposite slopes (but not to the
8.2 Grooved Substrata
165
Fig. 8.16 (a–c) Fibroblastic mouse cells after their migration from the grooves. The cells are distributed unevenly: they are mostly accumulated on the cylindrical substratum surfaces between the parallel grooves (a) or on the convex surfaces of the hexagons (b, c). LM. Scale bars, 150 mm (a, b). SEM. Scale bar, 150 mm (c)
bottoms themselves) by means of cell processes or the reduced lamellar cytoplasm; some cells “line” the grooves attaching both to their slopes and bottoms (Fig. 8.17a, b). In 24–36 h, cell densities at the areas between the grooves and in the immediate proximity to the groove bottoms do not significantly differ: transformed fibroblasts are distributed more or less evenly, irrespective of the presence of the grooves (Fig. 8.18a–c).
166
8 Topographic Cell Responses
Fig. 8.17 (a, b) Transformed mouse fibroblasts spread in the depths of the grooves. The cell makes a “bridge” over the groove bottom (a) or the cell “lines” the groove (b). SEM. Scale bar, 7 mm. Reproduced with permission from the Experimental Cell Research (ref. [23])
Fig. 8.18 (a–c) Transformed mouse fibroblasts on the grooved substrata. The cells are distributed evenly on the substrata with the parallel grooves (a, b) or with the hexagonal ones (c). LM. Scale bar, 100 mm (a). SEM. Scale bars, 50 mm (b), 60 mm (c)
8.3 Discontinuous Substrata
167
In contrast to normal fibroblastic cells, transformed fibroblasts located on the cylindrical substratum surfaces in the areas between parallel deep grooves do not demonstrate definite orientation to the direction of the grooves (Fig. 8.18a). Thus, normal fibroblastic cells actively and directionally migrate from the bottoms of the deep grooves of triangle cross-sections to the cylindrical surface areas between the grooves, where the cells orient themselves along the grooves. Epithelial cells do not migrate from the grooves. Migration of transformed fibroblasts is not directional and it is considerably weakened. Moreover, cells do not orient themselves along the grooves.
8.3 Discontinuous Substrata The ability of cells to spread and move in the presence of large substratum-free zones is an important reaction that allows cells to “overcome” extracellular matrixfree gaps during migrations in an organism. Special artificial substrata, called discontinuous ones, are used for cell culture to model the discontinuous character of the extracellular matrix, in particular, the presence of different kinds of slits, lacunae, etc., which occur on the ways of cell migrations. Discontinuous substrata have surfaces adhesive for cells alternating with large substrate-free zones.
8.3.1 Lattices Lattices can be the specimens of artificial discontinuous substrata. They have both substratum surfaces (lattice bars) and large substratum-free zones (lattice holes). For cell culture, it is possible to use metal lattices, in which the lattice bars have cylindrical forms with a diameter of 5–8 mm, and the lattice holes have square forms with the area of a hole of 2,000–2,500 mm² [28, 29]. Suspended cells attach to the lattice bars and begin to spread (Fig. 8.19a).
Normal Cell Responses The early stages of spreading of fibroblastic and epithelial cells are similar. The attached cell spreads along the bar (Fig. 8.19b, c), and then it begins to expand along the crossbar (Fig. 8.20a). Therefore, during spreading a cell is expanding simultaneously along two bars perpendicular to each other. In this position, the cell forms a small lamellar cytoplasm in the corner between two crossing bars over the void space of the hole (Fig. 8.20b, c). Somewhat later the cell-specific differences become apparent. Fibroblastic cells retract their central parts that are being detached from the bars (Fig. 8.21). The cells acquire fusiform or elongated shapes, and they remain attached
168
8 Topographic Cell Responses
Fig. 8.19 (a–c) Fibroblastic human cells in their spreading process on the lattice bars. The spherical cell attached to the bar by the lamellipodia (arrows) (a); the lamellar cytoplasm (arrow) surrounding the prominent part of the cell (b); the cell spread along the bar: the lamellar cytoplasm at the opposite cell ends (arrows) expands on two crossbars (c). SEM. Scale bars, 7 mm (a), 8 mm (b), 11.5 mm (c)
to the bars only by the cell processes, while the unattached cell bodies are located over the holes (Fig. 8.21). Eventually, most fibroblastic cells acquire elongated shapes, move actively in different directions and “cross” the lattice openings with elongated cell bodies and long thin processes (Fig. 8.22a, b), thus forming only a few discrete short contacts with the bars. In contrast to fibroblastic cells, the retraction of the central parts of epithelial cells does not take place. The epitheliocyte continues its spreading along the bar and the crossbar with the lamellar cytoplasm, which is progressively expanded over the lattice
8.3 Discontinuous Substrata
169
Fig. 8.20 (a–c) Epithelial mouse cells (a, b) and human fibroblastic cell (c) spread on the lattice bars. One cell (arrow) spread along the bar, another cell (double arrows) spread simultaneously along two bars perpendicular to each other (bar and crossbar) (a). Triangular lamellar cytoplasm (arrow) in the corner between two crossing bars stretching over the lattice hole (b, c). SEM. Scale bars, 12 mm (a), 12.5 mm (b), 10.5 mm (c)
opening. Eventually, the epitheliocytes as thin wide plates “cover” considerable parts of the holes, and even can “cover” them completely (Fig. 8.23a, b). The cell forms numerous focal adhesions at all cell edges attached to the bars and remains immobile. HGF/SF is potent motogen for epitheliocytes and causes an alteration of their discoid shapes towards fibroblast-like ones and reorganization of actin cytoskeleton (see Sect. 6.1.4). Upon treatment of epithelial cells with HGF/SF, their behavior on the lattices becomes similar to that of fibroblastic cells: the treated epithelial cells are getting elongated and “cross” the lattice holes (Fig. 8.24).
170
8 Topographic Cell Responses
Fig. 8.21 Fibroblastic human cell spread on the lattice. The partial retraction of the lamellar cytoplasm (arrow) and its detachment from the right crossbar. SEM. Scale bar, 11 mm
Fig. 8.22 (a, b) Fibroblastic human cells spread on the lattice. The cells “cross” the lattice holes by the elongated cell bodies (a, b) and long thin cell processes (arrows) (b). SEM. Scale bars, 150 mm (a), 30 mm (b)
Thus, both normal fibroblastic and epithelial cells on the lattices have the ability to spread over the void spaces of lattice openings, retaining the contacts with the lattice bars only by the cell edges (epithelial cells) or by the cell processes (fibroblastic cells), whereas most of the cell bodies are not attached to any substratum surface.
8.3 Discontinuous Substrata
171
Fig. 8.23 (a, b) Epithelial mouse cells spread on the lattice. The cells widely stretched over the lattice holes: the cells “cover” the considerable parts of the holes (a, b) and can “cover” them completely (arrow) (a). SEM. Scale bars, 13 mm (a), 60 mm (b)
Fig. 8.24 Epithelial mouse cells spread on the lattice after the HGF/SF treatment. The elongated fibroblast-like cells “cross” the lattice holes. SEM. Scale bar, 20 mm
172
8 Topographic Cell Responses
Fig. 8.25 (a, b) Transformed mouse epitheliocytes on the lattice. The cells do not expand to the lattice holes (a, b), some of the cells “cross” the holes (b). SEM. Scale bars, 20 mm (a), 65 mm (b)
Transformed Cell Responses Transformed epithelial cells seeded from suspension attach to the lattice bars and begin to spread along the bars. In contrast to normal cells, in most of the transformed epitheliocytes, the lamellar cytoplasm in the corner between two bars perpendicular to each other does not form at all. Most of the spread transformed epitheliocytes remain on the bars and do not expand to the lattice holes (Fig. 8.25a). Only some of the cells “cross” the holes (Fig. 8.25b).
8.3.2 Multiple Vertvical Rods Other types of discontinuous substratum surface topography are multiple vertical rods separated from one another by void spaces [30]. The examples of such artificial substrata are plates with multiple vertical rod-like silicon microcrystals: the rods’ heights are 70–90 mm, their diameters are 5–10 mm, and the distances between the nearest neighbor rods vary from 10 to 30 mm (Fig. 8.26). The cells seeded on these substrata can attach only to the tops or side surfaces of the rods separated from one another by void spaces, which are often greater than the diameters of nonspread spherical cell. Normal fibroblastic cells seeded on these discontinuous substrata attach to the rods. Single spherical cells attach to the tops of single rods (Fig. 8.27a) or to their lateral surfaces. At the early stage of spreading, fibroblastic cells extend straight filopodia (Fig. 8.27b, d). They are often unusually long and can reach 30–40 mm; their ends attach to the tops and/or to the bodies of the rods located at various distances from the cell (Fig. 8.27c).
8.3 Discontinuous Substrata
173
Fig. 8.26 Multiple vertical rod-like silicon microcrystals on the silicon plate. SEM. Scale bar, 28 mm. Reproduced with permission from the Experimental Cell Research (ref. [30])
Eventually, fibroblastic cells completely spread and acquire flattened, triangular or polygonal shapes (Fig. 8.27e). Most of them attach only to the tops of the rods or to the upper parts of the rod bodies; the well spread cells look like “roofs” on the rods (Fig. 8.28). Sometimes one can see the rods “passing through” the cell bodies (Fig. 8.29). Few cells attach to the lateral sides of two or more rods (Fig. 8.30). The cells expanded on the rods or between them become morphologically similar to the fibroblastic cells spread on usual flat substrata with continuous surfaces. The contacts of cell edges with the rods are usually established by the lamellar cytoplasm, which “envelops” one or both sides of a rod (Fig. 8.31). Thus, fibroblastic cells can spread on discontinuous rod surfaces being attached only to the tops of the rods or to the rod bodies; whereas most parts of the spread cells appear to “hang” in wide void spaces. How do the cells spread on the rods? At the early stage of the spreading, the spherical cells attached to the rod tops extend long filopodia, which attach to the distant rods. These filopodia are apparently not extended gradually along a substrate but they are “shot” by the cells across the void spaces. Occasionally the “lucky” filopodia reach the distant rods and attach to them; all unattached filopodia, probably, contract and disappear. Later, the attached filopodia may act as guidelines directing the outward extension of a lamellar cytoplasm, promoting the subsequent progressive cell spreading.
174
8 Topographic Cell Responses
Fig. 8.27 (a–e) The spreading of fibroblastic mouse cells on the multiple vertical rods. The spherical cell attached to the top of the single rod (arrow) (a); the long straight filopodia (arrows) coming out of the spherical cell (b) or not completely spread one (d): the filopodia extend through void spaces and attach to the neighbor or distant rods (c); the cell completely spread on the rods (arrow) (e). SEM. Scale bars, 8 mm (a), 12 mm (b), 4 mm (c), 8 mm (d), 7 mm (e). Reproduced with permission from the Experimental Cell Research (ref. [30])
Thus, the behavior of normal fibroblastic or epithelial cells on the artificial discontinuous substrates shows that the continuity of the extracellular matrix is not a necessary condition for advance cell spreading. Normal cells have the ability to stretch over wide void spaces and to successfully achieve complete spreading.
8.4 Effects of the Substratum Surface Topography on Cell Adhesion, Proliferation…
175
Fig. 8.28 Well-spread fibroblastic mouse cells on the multiple vertical rods. The cells attached only to the tops of the rods or to upper parts of the rod bodies: the most parts of the cells “hang” in void spaces. SEM. Scale bar, 60 mm
Fig. 8.29 Spread fibroblastic mouse cell “passed through” by the rod (arrow). SEM. Scale bar, 13 mm
176
8 Topographic Cell Responses
Fig. 8.30 Spread fibroblastic mouse cell attached to the bodies of two rods (arrows). SEM. Scale bar, 8 mm
Fig. 8.31 The contact of spread fibroblastic mouse cell edge with the rod: the lamellar cytoplasm (arrows) “envelops” the rod body. SEM. Scale bar, 5 mm
8.5 Mechanisms of Topographic Cell Responses
177
8.4 Effects of the Substratum Surface Topography on Cell Adhesion, Proliferation, and Synthetic Activities The cells respond to the micro- or nanoscale substratum surface topography by not only changing their shape, orientation, and migration, but also by altering cell adhesion, proliferation, differentiation, and the ability to produce some important molecules [7, 9, 11, 31]. The substratum topography affects intracellular signaling pathways, in particular, mitogenic and morphogenic signaling from integrin receptors (see Sect. 5.3). A grooved topography stimulates FAK and MAP kinases in fibroblastic cells. Besides, MAP kinase activation in the cells on the grooves needs FAK-Src complex phosphorylation, whereas in the cells on smooth substratum surfaces, MAP kinase activation is Src independent [32]. Rho- (but not Rac or Cdc42) mediated intracellular signaling is responsible for the alignment of epithelial cells on nanoscale grooved substrata [21]. Micro- and nanoscale topographic features affect cell adhesion. For example, nanoscale pillars or pits, as compared with smooth substratum surface, diminish cell adhesion, whereas micro- or nanoscale pores enhance fibroblastic cell attachment [33–35]. Substratum surface nanoscale topography influences the proliferative activity and differentiation of cultured cells. Fibroblastic cells increase their proliferation on nanoscale fiber-like substrata. The proliferation of epithelial cells is reduced in response to the grooved topography scale decrease from a few micrometers to tenths of micrometers [35, 36]. The increased nanoscale substratum surface roughness, such as randomly distributed “islands” of 0.011 mm height, stimulates bone cell differentiation [37, 38]. Grooved substratum surface topography can influence the production of some extracellular matrix proteins, such as fibronectin and tenascin, as well as matrix metalloproteinase-2 by fibroblastic cells [39, 40]. Therefore, tissue cells have the ability to respond to the geometry of a chemically homogeneous solid substratum. Depending on the geometric configuration and its quantitative parameters, the cell’s responses vary. Topographic cell responses can include not only morphological alterations, but also changes in the intracellular signal transduction chains, which lead to the shifts in synthetic and functional activity of the cells.
8.5 Mechanisms of Topographic Cell Responses The capability of cells to bend can be the basis of many topographic responses. This capability is different in fibroblastic and epithelial cells, and it is determined by different actin cytoskeleton organization in these cells. Fibroblastic cells cultured on a side of a glass prism cannot crawl over the rib onto another side if the sides meet at an angle of less than 164° (Fig. 8.32). The
178
8 Topographic Cell Responses
Fig. 8.32 Diagram showing the resistance of fibroblastic cells to their mechanical bending: a “critical angle” of the bending (see the text for explanation). Reproduced with permission from the Experimental Cell Research (ref. [41])
crawling over is apparently prevented by relatively high resistance to the mechanical bending of linear straight actin filament bundles in the fibroblastic cells. Apparently, stress fibers and their associated focal contacts cannot be formed in the situations where actin bundles would be bent at an angle of less than the “critical” one [41, 42]. Hence, on nonplanar substrata, the efficiency with which the fibroblastic cell can exert the centripetal stretch and move is considerably limited in definite directions. This would lead to changes in cell shape, orientation, and direction of migration. Relatively high resistance to bending is apparently typical only for straight linear bundles of actin filaments. When the bundles have a circular shape (as in epithelial cells), their resistance to bending weakens considerably. As a result, the responses of fibroblastic and epithelial cells to the substratum surface topography are different.
8.5.1 Cylindrical Substrata The cell spreading on cylinders with a sufficiently high convex surface curvature proceeds without constraints along the cylinders, whereas the spreading in the perpendicular direction (across the cylinder) would be accompanied by cell bending. On plane substrata, spread fibroblastic cells have polygonal or more elongated shapes and contain well-developed system of straight linear actin filament bundles (see Fig. 5.21 in Chap. 5). Many types of transformed epithelial cells, in contrast to normal discoid epithelial cells, acquire polarized fibroblastic cell-like morphology, and they often reorganize the circular filament bundle pattern into predominantly straight bundles (see Fig. 5.39 in Chap. 5). On cylindrical substrata linear filament bundles restrict cell bending and “force” the normal fibroblastic cells or transformed epithelial cells to expand longitudi-
8.5 Mechanisms of Topographic Cell Responses
179
nally, with the cells getting elongated and oriented along the cylinders (Figs. 8.5 and 8.10). Their linear actin bundles get oriented in the same direction (Fig. 8.11). Another, predominantly circular pattern of actin filament bundles is typical for single spread epithelial cells (see Fig. 3.3 in Chap. 3) and for fibroblastic cells at the early (radial) stage of their spreading (see Fig. 5.18 in Chap. 5). On plane substrata, cells have discoid shapes and contain circular filament bundles located along the cell periphery. Circular filament bundles in these nonpolarized cells apparently do not restrict their bending, and the cells are getting bent across the cylinders without any significant elongation and orientation (Figs. 8.4 and 8.6). However, with a very high convex curvature of cylindrical surfaces (the cylinders’ diameters of 24–26 mm) the bending apparently becomes excessive even for the cells with the circular pattern of their filament bundles. The attached fibroblastic cells immediately enter the second (polarization) stage of their spreading, and they are being elongated along the cylinders (Fig. 8.2). The formation of transverse actin filament bundles in nonpolarized cells, which are bent around the cylinders (Fig. 8.7), is probably due to the transverse tension forces. It is known from mechanics that a bending of a deformable body is accompanied by the development of tension forces, resisting the bending. The resisting forces should be maximal in the direction transverse to the cylinder axis. The development of transverse deformation tension should cause the formation of transverse actin bundles in cells bending around the cylinders. Typically, transformed fibroblasts are practically devoid of the stress fibers; cells almost do not form focal contacts. Therefore, these cells do not respond to the curvature of cylindrical substrata: they are not getting elongated and not oriented along the cylinders. Transformed fibroblasts are not resistant to their bending around the cylinders (Fig. 8.9).
8.5.2 Grooved Substrata The migration of fibroblastic cells from deep grooves of triangle cross-sections apparently occurs because the cells cannot effectively spread at the bottoms of the grooves with angles between the groove side slopes of about 90°. On the bottoms of deep triangular grooves, the spread cells will be forced to bend at almost right angle. Since fibroblastic cells containing straight linearly organized actin filament bundles are resistant to the mechanical bending at an angle less than “critical” one (see above), the cells will select the position, which would prevent their excessive bending. As a result, fibroblastic cells leave the geometrically “inconvenient” groove bottoms and migrate to the side slopes of the grooves (Fig. 8.15a, b). Subsequently, the capability of fibroblastic cells for directional migration ensures their subsequent migration to and accumulation at the areas between the grooves (Fig. 8.16a, b). Epithelial cells, which have circular filament bundles, possess relatively weak resistance to their bending (see above). Therefore, epitheliocytes can successfully
180
8 Topographic Cell Responses
spread at the bottoms of deep grooves of triangle cross-sections. Since single epithelial cells, unlike fibroblastic cells, are not polarized, they are deprived of the ability to directional locomotion. Because of that, epithelial cells almost do not migrate from deep grooves. Similarly, transformed fibroblasts, whose actin filament bundles are usually strongly reduced, are weakly resistant to mechanical bending. Because of that, transformed fibroblasts can spread at the bottoms of deep grooves of triangular cross-sections (Figs. 8.17b and 8.18a, c). The migration of transformed fibroblasts from the grooves is also weakened by the chaotic, nondirectional locomotion of these cells: many transformed fibroblasts, which have migrated to the groove slopes, return to the bottoms of the grooves. The cylindrical substrata with a sufficiently high degree of curvature can affect cell shapes and cell orientation (see Sect. 8.1.1). Surfaces of the areas between parallel deep grooves have cylindrical forms with their curvature radii from 60 to 150 mm. Many of the fibroblastic cells, which have migrated from the grooves and which are concentrated in the areas between the grooves, respond to cylindrical forms of these areas by the orientation of the cells along the cylinders (Fig. 8.16a). The most pronounced orientation is seen in fibroblastic cells located between parallel grooves on cylindrical surfaces with maximal curvature (radius of 60 mm). Transformed fibroblasts weakly respond to the curvature of cylindrical substrata (see Sect. 8.1.2). Transformed cells located on the cylindrical surfaces between parallel deep grooves do not orient themselves along the cylinders (Fig. 8.18a, b). The surprising ability of cells to respond to the nanoscale substratum surface topography, including nanoscale grooves, is not completely explained at the present time [7, 9, 11, 31]. Some cell types respond to “steps” as shallow as 0.011 mm [43, 44]. It was suggested that cells have specific tension receptors responding to nanoscale topographic features. These hypothetical cell membrane receptors could “trigger” intracellular signaling pathways leading to the reorganization of actin cytoskeleton, alterations in cell adhesion, and redistribution of tension forces in the cell. This, in turn, could cause cell elongation and orientation in a certain direction (e.g., along the nanoscale grooves and ridges). It has been shown that the nanoscale grooves induce the alignment of epithelial cells via Rho-mediated intracellular signaling that controls actin cytoskeleton organization, cell contractility, and focal adhesion formation [21]. The “tension receptors”-dependent signaling could also cause alterations in cell proliferation and gene expression. The ion chloride channels in the cell membrane can be possible candidates for the role of “tension receptors”: in liquid medium with the deficiency of chlorides or when chloride channel inhibitors are present in the medium, the orientation of cells along the shallow grooves considerably weakens [45]. Possibly, topographic responses of tumor cells weaken considerably due to the structural or functional alterations in the “tension receptors” caused by oncogenic transformation.
8.5 Mechanisms of Topographic Cell Responses
181
8.5.3 Lattice Substrata The gradual displacement of cells from the bars to the openings is probably due to the bending of a cell while it is spreading along two bars perpendicular to each other. During this spreading, the cell body expands in two directions perpendicular to each other. The resultant of these expanding forces is directed along the diagonal of the square hole, and it causes cell expansion in the same direction: the bent cell begins to form lamellar cytoplasm at the corner of the hole (Fig. 8.20a–c). The capability for bending of fibroblastic cells is restricted because of the relatively high bending resistance of their straight actin filament bundles. Being bent at an angle that is less than the “critical” one, the cell cannot establish focal contacts (see above). As a result, the fibroblastic cell retracts, and remains attached to the lattice bars only in a few discrete points; the cell acquires “straightened” position crossing the hole (Fig. 8.22a). In contrast to fibroblastic cells, epitheliocytes contain mainly circular actin but not linear filament bundles. Therefore, epithelial cells are not resistant to their bending and they are capable to be in a bent state (see above). As a result, an epithelial cell continues to spread along two crossing lattice bars and progressively develops its lamellar cytoplasm over the lattice hole (Fig. 8.23a, b). The conversion of circular filament bundles into the straight ones in the epithelial cells treated with HGF/SF is the probable cause of altered, fibroblast-like, behavior of the treated cells on the lattices (Fig. 8.24). Transformed fibroblasts contain few straight actin filament bundles (or do not contain them at all), which could restrict cell body bending. Therefore, a transformed fibroblast is able to spread along two crossing lattice bars. However, because of the defectiveness of focal contacts accompanied by the deficiency in filament bundle formation and by the significant weakening of the centripetal stretch (see Sect. 5.4.1), transformed fibroblasts cannot expand as effectively as normal fibroblastic cells can. The expanding forces in the bent transformed fibroblast while it is spreading along two crossing bars are significantly weakened. The resultant of these forces is also weakened, and it is unable to cause the displacement of the transformed fibroblasts from the bars into the lattice holes (Fig. 8.25a, b). Thus, tissue cells in a multicellular organism submit their form and functional activity to the requirements dictated by chemical properties and geometric configuration of the extracellular matrix. The heterogeneity of chemical adhesive properties of the matrix, and its geometric characteristics, including the curvature and topography of its surface, or disruptions in the matrix continuity are potent regulating factors. They determine and control the directions of cellular migrations, the concentration of cells of specific types in the areas of tissue regeneration (e.g., ample, in wound healing) or in embryogenesis; these factors also regulate the proliferation and synthetic activities of the cells. The regulatory influences of both chemical and topographic characteristics of the extracellular matrix on the cells are seriously disturbed as a result of oncogenic transformation.
182
8 Topographic Cell Responses
References 1. Weiss P (1941) Nerve pattern: The mechanics of nerve growth. Growth (suppl.5): 163–203 2. Flemming RG, Murphy CJ, Abrams GA, Goodman SL, Nealey PF (1999) Effects of synthetic micro- and nano-structured surfaces on cell behavior. Biomaterials 20(6):573–588. doi:10.1016/ S0142-9612(98)00209-9 DOI:dx.doi.org 3. Abrams GA, Goodman SL, Nealey PF, Franco M, Murphy CJ (2000) Nanoscale topography of the basement membrane underlying the corneal epithelium of the rhesus macaque. Cell Tissue Res 299(1):39–46. doi:10.1007/s004410050004 DOI:dx.doi.org 4. Curtis A, Wilkinson C (1997) Topographical control of cells. Biomaterials 18(24):1573–1583. doi:10.1016/S0142-9612(97)00144-0 DOI:dx.doi.org 5. Curtis ASG, Clark P (1990) The effects of topographic and mechanical properties of materials on cell behavior. Crit. Rev Biocompat 5: 343–362. 6. Norman JJ, Desai TA (2006) Methods for fabrication of nanoscale topography for tissue engineering scaffolds. Ann Biomed Eng 34(1):89–101. doi:10.1007/s10439-005-9005-4 DOI:dx.doi.org 7. Kriparamanan R, Aswath P, Zhou A, Tang L, Nguyen KT (2006) Nanotopography: cellular responses to nanostructured materials. J Nanosci Nanotechnol 6(7):1905–1919. doi:10.1166/ jnn.2006.330 DOI:dx.doi.org 8. Yap FL, Zhang Y (2007) Protein and cell micropatterning and its integration with micro/nanoparticles assembly. Biosens Bioelectron 22(6):775–788. doi:10.1016/j.bios.2006.03.016 DOI:dx.doi.org 9. Bettinger CJ, Langer R, Borenstein JT (2009) Engineering substrate topography at the microand nanoscale to control cell function. Angew Chem Int Ed Engl 48(30):5406–5415. doi:10.1002/anie.200805179 DOI:dx.doi.org 10. Schmidt RC, Healy KE (2009) Controlling biological interfaces on the nanometer length scale. J Biomed Mater Res A 90(4):1252–1261. doi:10.1002/jbm.a.32501 DOI:dx.doi.org 11. von der Mark K, Park J, Bauer S, Schmuki P (2010) Nanoscale engineering of biomimetic surfaces: cues from the extracellular matrix. Cell Tissue Res 339(1):131–153. doi:10.1007/ s00441-009-0896-5 DOI:dx.doi.org 12. Svitkina TM, Rovensky YA, Bershadsky AD, Vasiliev JM (1995) Transverse pattern of microfilament bundles induced in epitheliocytes by cylindrical substrata. J Cell Sci 108 (Pt 2):735–745. 13. Rovensky YuA, Samoilov VI (1994) Morphogenetic response of cultured normal and transformed fibroblasts, and epitheliocytes, to a cylindrical substratum surface. Possible role for the actin filament bundle pattern. J Cell Sci 107 (Pt 5):1255–1263. 14. Levina EM, Domnina LV, Rovensky YA, Vasiliev JM (1996) Cylindrical substratum induces different patterns of actin microfilament bundles in nontransformed and in ras-transformed epitheliocytes. Exp Cell Res 229(1):159–165. doi:10.1006/excr.1996.0354 DOI:dx.doi.org 15. Clark P, Connolly P, Curtis AS, Dow JA, Wilkinson CD (1990) Topographical control of cell behaviour: II. Multiple grooved substrata. Development 108(4):635–644. 16. Clark P, Connolly P, Curtis AS, Dow JA, Wilkinson CD (1991) Cell guidance by ultrafine topography in vitro. J Cell Sci 99 (Pt 1):73–77. 17. Alaerts JA, De Cupere VM, Moser S, Van den Bosh de Aguilar P, Rouxhet PG (2001) Surface characterization of poly(methyl methacrylate) microgrooved for contact guidance of mammalian cells. Biomaterials 22(12):1635–1642. doi:10.1016/S0142-9612(00)00321-5 DOI:dx.doi.org 18. Oakley C, Brunette DM (1993) The sequence of alignment of microtubules, focal contacts and actin filaments in fibroblasts spreading on smooth and grooved titanium substrata. J Cell Sci 106 (Pt 1):343–354. 19. Kaiser JP, Reinmann A, Bruinink A (2006) The effect of topographic characteristics on cell migration velocity. Biomaterials 27(30):5230–5241. doi:10.1016/j.biomaterials.2006.06.002 DOI:dx.doi.org 20. Teixeira AI, McKie GA, Foley JD, Bertics PJ, Nealey PF, Murphy CJ (2006) The effect of environmental factors on the response of human corneal epithelial cells to nanoscale substrate topography. Biomaterials 27(21):3945–3954. doi:10.1016/j.biomaterials.2006.01.044 DOI:dx.doi.org
References
183
21. Rajnicek AM, Foubister LE, McCaig CD (2008) Alignment of corneal and lens epithelial cells by co-operative effects of substratum topography and DC electric fields. Biomaterials 29(13):2082–2095. doi:10.1016/j.biomaterials.2008.01.015 DOI:dx.doi.org 22. Rovensky YA, Slavnaja IL, Vasiliev JM (1971) Behaviour of fibroblast-like cells on grooved surfaces. Exp Cell Res 65(1):193–201. doi:10.1016/S0014-4827(71)80066-6 DOI:dx.doi.org 23. Rovensky YA, Slavnaya IL (1974) Spreading of fibroblast-like cells on grooved surfaces: a study by scanning electron microscopy. Exp Cell Res 84(1):199–206. doi:10.1016/00144827(74)90397-8 DOI:dx.doi.org 24. Veselý P, Rovensky YuA, Slavnaya IL (1981) Cinemicrographic analysis of migration of normal and neoplastic rat cells seeded upon the grooved surface. Folia Biol (Praha) 27(2):81–90. 25. Karuri NW, Liliensiek S, Teixeira AI, Abrams G, Campbell S, Nealey PF, Murphy CJ (2004) Biological length scale topography enhances cell-substratum adhesion of human corneal epithelial cells. J Cell Sci 117(Pt 15):3153–3164. doi:10.1242/jcs.01146 DOI:dx.doi.org 26. Diehl KA, Foley JD, Nealey PF, Murphy CJ (2005) Nanoscale topography modulates corneal epithelial cell migration. J Biomed Mater Res A 75(3):603–611. doi:10.1002/jbm.a.30467 DOI:dx.doi.org 27. Khung YL, Barritt G, Voelcker NH (2008) Using continuous porous silicon gradients to study the influence of surface topography on the behaviour of neuroblastoma cells. Exp Cell Res 314(4):789–800. doi:10.1016/j.yexcr.2007.10.015 DOI:dx.doi.org 28. Rovensky YA, Domnina LV, Ivanova OY, Vasiliev JM (1999) Locomotory behaviour of epitheliocytes and fibroblasts on metallic grids. J Cell Sci 112 (Pt 8):1273–1282. 29. Rovensky YuA, Domnina LV, Ivanova OYu, Vasiliev JM (2001) Responses of epithelial and fibroblast-like cells to discontinuous configuration of the culture substrate. Membr Cell Biol 14(5):617–627. 30. Rovensky YA, Bershadsky AD, Givargizov EI, Obolenskaya LN, Vasiliev JM (1991) Spreading of mouse fibroblasts on the substrate with multiple spikes. Exp Cell Res 197(1):107–112. doi:10.1016/0014-4827(91)90486-E DOI:dx.doi.org 31. Lim JY, Donahue HJ (2007) Cell sensing and response to micro- and nanostructured surfaces produced by chemical and topographic patterning. Tissue Eng 13(8):1879–1891. doi:10.1089/ ten.2006.0154 DOI:dx.doi.org 32. Hamilton DW, Brunette DM (2007) The effect of substratum topography on osteoblast adhesion mediated signal transduction and phosphorylation. Biomaterials 28(10):1806–1819. doi:10.1016/j.biomaterials.2006.11.041 DOI:dx.doi.org 33. Curtis AS, Casey B, Gallagher JO, Pasqui D, Wood MA, Wilkinson CD (2001) Substratum nanotopography and the adhesion of biological cells. Are symmetry or regularity of nanotopography important? Biophys Chem 94(3):275–283. doi:10.1016/S0301-4622(01)00247-2 DOI:dx.doi.org 34. Zhu X, Chen J, Scheideler L, Altebaeumer T, Geis-Gerstorfer J, Kern D (2004) Cellular reactions of osteoblasts to micron- and submicron-scale porous structures of titanium surfaces. Cells Tissues Organs 178(1):13–22. doi:10.1159/000081089 DOI:dx.doi.org 35. Zheng Z, Zhang L, Kong L, Wang A, Gong Y, Zhang X (2009) The behavior of MC3T3-E1 cells on chitosan/poly-L-lysine composite films: Effect of nanotopography, surface chemistry, and wettability. J Biomed Mater Res A 89(2):453–465. doi:10.1002/jbm.a.31979 DOI:dx. doi.org 36. Liliensiek SJ, Campbell S, Nealey PF, Murphy CJ (2006) The scale of substratum topographic features modulates proliferation of corneal epithelial cells and corneal fibroblasts. J Biomed Mater Res A 79(1):185–192. doi:10.1002/jbm.a.30744 DOI:dx.doi.org 37. Kawahara H, Soeda Y, Niwa K, Takahashi M, Kawahara D, Araki N (2004) In vitro study on bone formation and surface topography from the standpoint of biomechanics. J Mater Sci Mater Med 15(12):1297–1307. doi:10.1007/s10856-004-5738-0 DOI:dx.doi.org 38. Lim JY, Hansen JC, Siedlecki CA, Runt J, Donahue HJ (2005) Human foetal osteoblastic cell response to polymer-demixed nanotopographic interfaces. J R Soc Interface 2(2):97–108. doi:10.1098/rsif.2004.0019 DOI:dx.doi.org
184
8 Topographic Cell Responses
39. Goto T, Brunette DM (1998) Surface topography and serum concentration affect the appearance of tenascin in human gingival fibroblasts in vitro. Exp Cell Res 244(2):474–480. doi:10.1006/excr.1998.4196 DOI:dx.doi.org 40. Chou L, Firth JD, Uitto VJ, Brunette DM (1998) Effects of titanium substratum and grooved surface topography on metalloproteinase-2 expression in human fibroblasts. J Biomed Mater Res 39(3):437–445. doi:10.1002/(SICI)1097-4636(19980305)39:33.0.CO;2-7 DOI:dx.doi.org 41. Dunn GA, Heath JP (1976) A new hypothesis of contact guidance in tissue cells. Exp Cell Res 101(1):1–14. doi:10.1016/0014-4827(76)90405-5 DOI:dx.doi.org 42. Dunn GA (1991) How do cells respond to ultrafine surface contours? Bioessays 13(10):541–543. doi:10.1002/bies.950131008 DOI:dx.doi.org 43. Curtis A, Wilkinson C (1999) New depths in cell behaviour: reactions of cells to nanotopography. Biochem Soc Symp 65:15–26. 44. Yim EK, Leong KW (2005) Significance of synthetic nanostructures in dictating cellular response. Nanomedicine 1(1):10–21. doi:10.1016/j.nano.2004.11.008 DOI:dx.doi.org 45. Tobasnick G, Curtis AS (2001) Chloride channels and the reactions of cells to topography. Eur Cell Mater 2:49–61.
Chapter 9
Intercellular Adhesive Interactions
Abstract Tissue cells form several types of intercellular adhesions. Some of them carry out barrier functions (tight junctions) or metabolic cell cooperation (gap junctions). The main type of intercellular adhesions is cadherin receptor-mediated adherence junction that plays a key role in joining cells into orderly tissue structures. Dynamic regulation of adherence junctions is the basis of cell rearrangements and tissue integrity maintenance. This regulation includes control of the adhesive function of cadherin receptors and the regulation of local actin cytoskeleton assembly. Adherence junctions carry out not only cell–cell adhesive but also signal transduction functions, being involved in signaling pathways that control cell migration (“contact inhibition of cell migration”), and proliferation(“contact inhibition of cell proliferation”). Both adhesive and signaling functions of adherence junctions are significantly altered as a result of oncogenic transformation. These alterations pivotally contribute to malignant tumor anaplasia, unrestricted proliferation, and tumor cell invasion. Intercellular adhesive interactions play a crucial role in embryogenesis and maintenance of tissue integrity. They regulate proliferation of tissue cells, their locomotion and synthetic processes in the cells. Changes in intercellular adhesiveness are critically involved in cancer invasion and metastasis [1–3]. There are several types of adhesive contacts formed by cells with each other: tight junctions, gap junctions, adherens junctions, and desmosomal junctions (desmosomes). Tight junctions (also referred to as zonulae occludens or occluding junctions) are beltlike intercellular junctions between epithelial or endothelial cells. In tight junctions plasma membranes of two cells come very close together, and outer leaflets of the membranes appear to be fused in sites of the junctions. Thereby an impermeable barrier to fluid is formed, and the passage of molecules and ions through the intercellular space (that is not available) is prevented. Therefore, tight junctions serve the barrier functions: they create a highly effective diffusion barrier between epithelial or endothelial cells by preventing free paracellular passage of molecules and ions [4–6].
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_9, © Springer Science+Business Media, LLC 2011
185
186
9 Intercellular Adhesive Interactions
Tight junctions are composed of transmembrane proteins called occludin and claudins. Their extracellular domains join one another directly and connect plasma membranes of two adjacent epithelial or endothelial cells with each other. Claudins and occludin associate with proteins concentrated at the cytoplasmic surfaces of tight junctions. These proteins, such as ZO-1, ZO-2, and ZO-3, link claudins and occludin to actin cytoskeletons of the contacting cells [4, 5, 7, 8]. ZO-1, ZO-2, and ZO-3 proteins are critical determinants of barrier functions of tight junctions. ZO-1 and ZO-2 can also be components of another type of intercellular adhesive contacts, adherens junctions [9] (see further). The tight junction barrier is regulated. The regulation is based on the ability of claudins to interact with multiple proteins. Thereby, claudins can be involved in bidirectional signal transduction to regulate paracellular ion flux in epithelial and endothelial cell sheets [8, 10], and also suppress cell proliferation [11]. The Rho and Rab GTPase families of small GTPases are involved in the dynamic regulation of assembly, disassembly, and functions of epithelial tight junctions. The regulation effected by Rho GTPases is obviously mediated by controlling the contraction of actin filaments [12–14]. Gap junctions are intercellular channels between plasma membranes of two adjoining cells. Gap junction contains a narrow gap between the cells. The plasma membranes of adjoining cells and the gap are run through by the cylindrical structures termed connexons that contain channels. A connexon is composed by a family of six transmembrane proteins termed connexins. The connexon channels can be opened or closed depending on the changes in the connexin conformation. These channels participate in the metabolic cell cooperation enabling contacting cells to directly exchange ions and small molecules without their release into the intercellular space. Connexins interact with diverse cellular proteins including cytoskeleton proteins, phosphatases, and protein kinases; these interactions can regulate gap junctionmediated intercellular communication [15]. Recently a new protein family that forms gap junctions has been described. These proteins called pannexins are structurally similar to connexins [16]. Gap junctions play critical roles in electrical coupling of cardiomyocytes and in the electrical synaptic transmission in the central nervous system [17, 18]. Adherens junctions and desmosomal junctions (desmosomes, also known as maculae adherens), like focal adhesions (focal contacts), are composed of the transmembrane adhesion receptors, which via the submembrane protein complex, are linked to the cytoskeleton of contacting cells [19]. The principal components of adherens junctions and desmosomes are the transmembrane adhesion receptors, cadherins. They form a superfamily of glycoproteins that mediate calcium-dependent intercellular adhesion. The so-called classical cadherins are components of adherens junctions; desmosomal cadherins are called desmocollins and desmoglein. Cadherin-based intercellular adhesion is established by direct interaction between cadherin molecules extending from the adjacent cells. Classical cadherins initiate cell–cell adhesion bonds and promote cell sorting and cell rearrangements. Desmosomal cadherins supply added strength to intercellular connections producing very strong cell–cell adhesion that provides mechanical stability in epithelial and
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
187
also in a few nonepithelial tissues, e.g., in cardiac muscle [19–21]. Though desmosomal junctions are highly stable, they have dynamic activity and lateral motility [22]. Cadherin transmembrane receptor exists as a parallel homodimer. Cadherins have extracellular domains consisting of repeats (EC subdomains, extracellular cadherin subdomains), which mediate adhesive binding of cells to each other. Like integrins, cadherins have intracellular domains bound to cytoskeletal structures of contacting cells via the submembranous complex of interrelated cytoplasmic proteins called catenins in adherens junctions or plakoglobin (also known as g-catenin), desmoplakin, and plakophilins in desmosomes [19, 23]. These protein complexes, in turn, are bound to actin filaments in adherens junctions or to intermediate filaments in desmosomal junctions.
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions 9.1.1 Structure of Adherence Junctions Adherens junctions play a key role in joining up the cells into tissue structures and in regulating tissue architecture. An adherens junction is a molecular formation including transmembrane cadherin receptor, submembrane catenin complex, and actin cytoskeleton. This adhesive structure, like focal adhesion, is dynamic and carries out not only adhesive but also intracellular signal transduction functions [24–26]. Classical cadherin receptors of adherens junctions include E (epithelial), N (neural), P (placental), R (retinal), VE (vascular endothelial), and some other cadherins [24, 27]. Their extracellular domains consist of five EC1–EC5 subdomains that are bound together by Ca2+ ions to form rod-like proteins (Fig. 9.1). Classical cadherins are particularly important for the formation and dynamic regulation of adhesive intercellular contacts [28]. E- or VE-cadherins form adherens junctions in epithelial or endothelial cells, respectively, resulting in the formation of tight cell layers (Fig. 9.2). E-cadherin is the epithelial-specific adhesion molecule. It plays a pivotal role in the formation, maintenance, and dynamic regulation of adherens junctions, and it is also involved in the control of proliferation of epithelial cells [29, 30]. VE-cadherin is important for the formation, remodeling, and permeability of the vascular wall [31–34]. N-cadherin is normally found only in the mesenchymal tissues; it is present in neural cells and participates in the formation of neuron connections [30]. P-cadherin is present in epidermal basal cells and in the myoepithelial cells of mammary glands. P-cadherin plays an important role in the morphogenesis of epidermis and skin appendages. R-cadherin is expressed in the nervous system. Alhough classical cadherins were named after the tissues, in which they are mostly expressed, they can be expressed not only in the specific cell type but also in many different ones. Cadherin-mediated intercellular adhesion can be homophilic or heterophilic: both identical classical cadherins and different ones in two adjacent cells can be bound to each other.
188
9 Intercellular Adhesive Interactions
Fig. 9.1 Diagram of the adherens junction: classic cadherin–catenin complex (see the text for explanation)
Fig. 9.2 The intercellular contacts between epithelial cells. Staining for E-cadherin (green). The adherens junctions are oriented linearly along the contacting cell edges. Confocal laser scanning microscopy. Scale bar, 25 mm. Reproduced with permission from the Proceedings of the National Academy of Sciences USA, 1999; 96(17):9666–9670. Copyright 1999 National Academy of Sciences, USA
In embryogenesis, cadherins control cell sorting and cell rearrangements d etermining the separation of distinct tissue layers and formation of tissue boundaries. These adhesion molecules control the formation of connections between neurons in the developing nervous system. In adult tissues, cadherins are involved in the turnover of rapidly growing tissues, e.g., of the intestinal lining or the epidermis. Cadherins are involved in cell migrations in regeneration processes, and they maintain the stability of tissue organization [35].
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
189
Isoform switching between classical cadherins allows cell types to segregate from one another [36]. Although the interaction of cadherin extracellular domains can provide the contacts between the adjacent cells, their cytoplasmic domains play essential role in the establishment of stable cell–cell adhesion. The cytoplasmic cadherin domain interacts with the submembranous catenin complex consisting of a-catenin, b-catenin, and p120-catenin (Fig. 9.1) [37, 38]. b-Catenin and p120-catenin are directly bound to the cytoplasmic cadherin domains: p120-catenin is bound to the juxtamembrane region of the domain, whereas b-catenin is bound to the domain’s distal region. b-Catenin, in turn, is bound to a-catenin. The b-catenin’s place can be taken by plakoglobin (also called g-catenin) that is typical for desmosomes; however, plakoglobin can be also found in the adherens junctions. a-Catenin binds to actin cytoskeleton, either by directly binding actin filaments or through actin-binding proteins, vinculin, a-actinin or formin1 (Fig. 9.1). Catenins have different important functions in adherens junctions [38]. Catenins are structural components physically linking cadherins to actin cytoskeleton (Figs. 9.1 and 9.3). This linkage is essential for the maintenance of the integrity of adherens junctions. Catenins are not only structural components, but they are also components of different signal transduction pathways that carry out the dynamic regulation of the cadherin-mediated intercellular adhesions. This regulation is necessary for the moving cells to be able to continually break and remake their adhesive bonds to change cell neighbors. The dynamic regulation is the basis of cell rearrangements; it maintains the integrity of tissue organization. Adherens junctions can also include proteins ZO-1 and ZO-2 that are characteristic components of tight junctions [9].
9.1.2 Dynamic Regulation of Adherens Junctions The process of adherens junction formation includes the initial stage, the stage of contact stabilization, and the maturation stage. At the initial stage, the bonds between the contacting cells are mechanically weak. These bonds are based on the interactions of individual cadherins, cytoplasmic domains of which are bound to b-catenin molecules. An important role in the stabilization of the initial cadherin-based adhesion belongs to other adhesion molecules, nectins. The family of nectins consists of four members. Nectins are immunoglobulin-like Ca2+-independent adhesion molecules, which are involved in the formation of adherens and tight junctions, cooperating with cadherins or claudins, respectively. They are transmembrane molecules, and their extracellular parts are homophilically or heterophilically bound forming cell–cell adhesions of epithelial or fibroblastic cells. The cytoplasmic tails of nectins are connected to the actin cytoskeleton in the contacting cells through an actin-binding protein, afadin [39]. Nectin-mediated adhesion is the initial step in adherens junction formation. In the nascent adherens junctions, nectins recruit cadherins to the adhesion sites; nectins
190
9 Intercellular Adhesive Interactions
Fig. 9.3 The intercellular contact between two fibroblastic cells. Double staining for b-catenin (green) and for actin (red). The adherens junctions are oriented perpendicular to the contacting cell edges and associated with the linear actin microfilament bundles. Confocal laser scanning microscopy. Scale bar, 16.5 mm. Reproduced with permission from the Proceedings of the National Academy of Sciences USA [76]. Copyright 1998 National Academy of Sciences, USA
also induce Rac and Cdc42 activation through the nectin cytoplasmic tails causing reorganization of actin cytoskeleton and thereby promoting the stabilization of cadherin-based adherens junctions. Besides, nectins interact with aVb3 integrin (vitronectin receptor) and PDGF receptor and are involved in their signaling [39–43]. At the stage of contact stabilization, the components of the submembrane protein complex are recruited linking the cadherin to actin cytoskeleton. The interactions of cadherins with actin filaments are crucially important for the intercellular contact stabilization. Then cadherin clusters are formed. Subsequent maturation of the adherens junctions is due to further actin cytoskeleton assembly at their sites [26, 42]. The Regulation of Adhesive State of Cadherin Receptors The ability of tissue cells to dynamically regulate the formation and maintenance of their intercellular adhesions is a necessary condition for tissue integrity and reorganizations of tissue architecture. The dynamic regulation of cadherin-mediated intercellular adhesions is a critical factor in embryonic development and in adult tissues. The formation of tissue boundaries, changes in shapes of tissues caused by cell rearrangements, formation of synapses between neurons, and the maintenance of stable tissue organization require the dynamic regulation of the stability and strength of intercellular cadherin-mediated bonds [42].
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
191
The control of the adhesive function of cadherin receptors is a crucial contribution to the dynamic regulation of cell–cell adhesion [42]. E-cadherin dimers can be arranged parallel to each other (so-called lateral dimers), or they can be aligned in an antiparallel fashion. In the latter case, the dimers are formed by E-cadherin molecules that are derived from both contacting epithelial cells to mediate intercellular adhesion, and so these structures are called adhesive dimers. The cadherin dimers in adherens junctions are structurally unstable and have a short lifetime: the continuous cadherin dimerization is counterbalanced by the dimer dissociation. Individual cadherin molecules shuttle between their monomeric and adhesive dimeric forms. Because of this instability, the dynamic regulation of adherens junctions, their strength and plasticity, can be carried out by shifts in the equilibrium between two opposing processes: cadherin adhesive dimer assembly and disassembly [19]. Adhesive dimers are continuously removed from adherens junctions by cadherin endocytosis and are continuously recruited into the adherens junctions. This cadherin endocytic and exocytic trafficking critically contributes to the intercellular adhesive potential [44–46]. The endocytosis is the main way of the disassembly of cadherin adhesive dimers. The amount of cadherin dimers is increased dramatically soon after the inhibition of endocytosis. Besides, cadherin dimers can be degraded by some metalloproteinases that cleave E-cadherin at the cell surface [42, 46–48]. p120 Catenin prevents cadherin endocytosis and thereby regulates adhesive functions of cadherin receptors [26, 42, 48]. The loss of p120-catenin stimulates internalization of adhesive cadherin dimers with their following proteolytic degradation in lysosomes and causes the weakening of cadherin-based intercellular adhesion. Mechanisms of p120-catenin-mediated regulation of cadherin endocytosis are not clear yet. It is possible, that p120-catenin connected with the cytoplasmic domain of a cadherin receptor functions as a capping molecule preventing cadherin from endocytosis. In this case, some factors causing p120-catenin dissociation from cadherin could stimulate its internalization [26]. The internalization of cadherins may be stimulated in response to growth factors (GFs). Phosphoinositides, specifically phosphatidylinositol biphosphate (PIP2), which is the product of phosphatidylinositol phosphate kinases (PIPKs) (see Sect. 5.2.2), is an important component of signaling pathways involved in the regulation of E-cadherin exocytosis and endocytosis [49]. Therefore, the stability and strength of cell–cell adhesion can be mediated by the regulation of the equilibrium between cadherin dimer assembly–disassembly and the lifetime of adhesive cadherin dimers. Presumably, this regulation is inside-out. Recently, some new data on cadherin trafficking and adherens junction dynamics regulation were obtained. As it turned out, E-cadherin molecules can be recruited into the adherens junctions spontaneously, independently on the suppressed cadherin endocytosis and on the disassociation of E-cadherin with b-catenin and p120catenin. E-cadherin molecules reside in the adherens junctions only for several minutes. They are associated into clusters (see further), and then cadherin molecules are actively released from these clusters. This active release of E-cadherin from the
192
9 Intercellular Adhesive Interactions
adherens junctions is unaffected by suppressed cadherin endocytosis; the mechanism of this process remains unclear [50]. Another critical factor contributing to the regulation of cadherin adhesive function is cadherin clustering [42]. Cadherin molecules residing in adherens junctions are laterally associated into clusters. The cadherin clustering increases the strength of cell–cell adhesion. b-catenin and p120-catenin binding to cadherin cytoplasmic domains affects the extent of cadherin clustering at the cell membrane and also cadherin dimerization. This effect could be presumably based on the phosphorylation status of b-catenin or p120-catenin. Their tyrosine phosphorylation is caused by the “outside-in” signals from receptors of soluble GFs or by the “inside-out” signals triggered by nonreceptor tyrosine kinases (e.g., submembraneous Src kinase). Tyrosine phosphorylation of b-catenin, p120-catenin, or some other tyrosine kinase targets weakens cadherinmediated intercellular adhesion [51]. The important factor in the maintenance of the integrity of cadherin-mediated adhesions is the microtubule system. Microtubules interact not only with integrin-based focal contacts (see Sect. 4.2.1) but also with adherens junctions, thereby affecting the tntegrity of these adhesions. Potential links between adherens junctions and microtubules may include p120-catenin, which has affinity for microtubules and also for the microtubule motor protein dynein that can bind to b-catenin. These direct links favor the interactions between microtubules and the adhesion sites. Microtubule-based transport provides delivery of newly synthesized or recycled cadherin molecules to the adherens junctions. In addition, microtubule plus-ends keep intact cortical myosin II that controls local concentration of E-cadherin at the adherens junctions. The integrity of adherens junctions is significantly disturbed when the microtubule plus-ends dynamics is damaged [52]. Therefore, in contrast to the negative influence of the microtubule targeting on focal contacts, microtubules have a positive influence on cadherin-mediated intercellular adhesions. Small GTPase Rap1 is not only a critical regulator of the inside-out integrin activation (see Sect. 4.2.1) but also plays a key role in the outside-in regulation of cadherin-mediated adhesion [26, 53]. Rap1 is activated by many extracellular stimuli. The guanine nucleotide exchange factors (GEFs) that “turn on” the Rap1 activity are directly linked to E-cadherin or to other proteins of adherens junctions [54]. The activity of Rap1 is necessary for the targeting of E-cadherin molecules to maturing cell–cell adhesions. Inhibition of Rap1 results in the formation of immature adherens junctions. Activation of Rap1 is critically required for the formation of the mature adherens junctions, and also for their even distribution around the circumferences of the cells within a continuous epithelial layer [53–55]. The Regulation of Actin Cytoskeleton in Adherens Junctions The local actin cytoskeleton rearrangements play a crucial role in the formation, stabilization, and dynamic regulation of adherens junctions [26, 42, 56].
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
193
Extensive actin dynamics is required to assemble the mature adhesive junctions. The key actin nucleating and filament assembling proteins, such as Arp2/3 protein complex, cortactin, and formin1 (see Sect. 3.1.1), are recruited to nascent cell–cell adhesions. These proteins are bound to cadherin cytoplasmic domains and trigger the assembly of actin filaments from the monomers at adherens junction sites [57– 59]. The members of WASP family, including N-WASP and WAVE2, which activate Arp2/3 complex, and also actin regulatory proteins of Ena/VASP family, co-localize with cadherins at the sites of new adherens junctions and are involved in the local assembly of actin filaments [26, 60, 61]. Inhibition of cortactin or proteins of WASP family blocks the assembly of new adherens junctions and disorganizes the already existing ones [26]. Control of actin cytoskeleton dynamics at adherens junctions is carried out by the “outside-in” signaling pathways from cadherin receptors through catenins (Fig. 9.4). In these signaling pathways, small GTPases of the Rho family (Rho, Rac, and Cdc42) play an important role [12, 26]. E-cadherin-mediated intercellular interaction activates submembraneous Src tyrosine kinase that facilitates the activation of phosphatidylinositol 3-kinase (PI3K) and its recruitment into the nascent E-cadherin-mediated adhesions. The E-cadherinproduced signals activating PI3K are possibly transferred through b-catenin that interacts with PI3K (Fig. 9.4). The PI3K product, phosphatidylinositol triphosphate PIP3 (PI3,4,5P3) (see Sect. 5.2.2) in the nascent cell–cell adhesions induces the recruitment of Rac1-specific activators (in particular, Rac-1-specific exchange factor, Tiam1) and protein kinase PKB/Akt. Cdc42 is also recruited into the nascent E-cadherin-mediated intercellular adhesions [62]. One of the targets of the activated Rac is the WASP/WAVE (WASP/Scar) protein family that, in turn, activates the Arp2/3 protein complex. The latter initiates actin polymerization and growth of new branching actin filaments (see Fig. 5.55 in Chap. 5, Fig. 9.4) (see Sect. 3.1.2). Rho protein is also involved in the control of local actin cytoskeleton assembly at the adherens junction sites. One of Rho targets, formin protein mDia1, induces linear growth of actin filaments (see Fig. 5.54 in Chap. 5) (see Sect. 3.1.2), and it is necessary for the stabilization of cadherin-mediated intercellular adhesions. Downregulation of mDia1 results in the disassembly of adherens junctions in epithelial cells [63]. Activities of Rho GTPases involved in adherens junction formation and maintenance are regulated by p120-catenin that activates Rac or Cdc42 and inhibits Rho (Fig. 9.4) [64]. Thus, both b-catenin and p120-catenin can influence local actin cytoskeleton organization. It has been suggested that the activation of Rac1 and Cdc42 inducing growth and assembly of actin filaments in nascent adherens junctions depends on Rap1 activity. Thereby, Rap1 activation favors the maturation of adherens junctions [26]. Additionally, the members of ABL family of tyrosine kinases, Abl and Arg kinases, play an important role in the regulation of Rho GTPases Rac and Rho. Their inhibition results in Rho activation that stimulates the formation and contractility of actin filament bundles. This action causes the disruption of adherens junctions [65].
194
9 Intercellular Adhesive Interactions
Fig. 9.4 Diagram of the outside-in signal transduction pathways from cadherin receptors (see the text for explanation)
In response to the activation of Rho GTPases, formin1 interacts with a-catenin and induces local assembly of linear actin filaments at the adherens junctions (Fig. 9.4). Protein kinase PKB/Akt, an important element in the cell survival signaling pathways (see Fig. 5.52 in Chap. 5) and that may be involved in actin cytoskeleton remodeling [66], is also recruited into the nascent E-cadherin-mediated adhesions. In the local cytoskeleton dynamics and associated adherens junction assembly, a-catenin plays a special role [37, 67, 68]. a-Catenin can change its conformation and activity depending on whether it is bound or not bound to b-catenin (Fig. 9.1). a-Catenin cannot bind b-catenin and actin filaments simultaneously: binding to b-catenin prevents a-catenin from its binding to actin filaments. a-Catenin binding to actin filaments suppresses the Arp2/3 protein complex activity and thereby prevents the formation of branched actin filament networks. Therefore, a-catenin directly regulates local actin filament assembly [37, 57, 69]. It is recently shown that cytosolic aE-catenin (one of two forms of a-catenin), independently on the cadherin– catenin complex and cell–cell adhesion, can regulate cellular actin dynamics, thereby influencing membrane dynamics and cell motility [70]. The a-catenin-mediated inhibition of Arp2/3 occurs at the initial stage of adherens junction assembly between two contacting cells. The Arp2/3 inhibition can be the mechanism of the “contact paralysis” (see Sect. 6.1.3) that favors the subsequent formation of mature cell–cell adhesions [57, 71]. The dissociation of a-catenin from the b-catenin–a-catenin complex is induced by a GTPase activating protein, IQGAP1 (IQ motif-containing GTPase activating
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
195
protein1). IQGAP1 is often associated with adherens junctions and can bind to many proteins including actin, E-cadherin, b-catenin, APC, and also Rho GTPases Rac1 and Cdc42. Interacting directly with b-catenin, IQGAP1 induces the separation of the latter from a-catenin, thereby negatively regulating E-cadherin-mediated intercellular adhesion. Rac1 and Cdc42 inhibit the interaction of IQGAP1 with b-catenin, and thus negatively regulate the function of IQGAP1, stabilizing the catenin complex and increasing cell–cell adhesion stability [72, 73]. Besides catenins, the proteins ZO-1 and ZO-2, which can be components of not only tight junctions but also adherens junctions, may play a role in the regulation of local assembly and organization of actin cytoskeleton [9]. The contractility of actin filaments is a key factor involved in the maintenance of adherens junctions. The negative regulator of myosin II-driven cell contractility, caldesmon, induces destabilization of E-cadherin-mediated intercellular contacts [74]. The contractile actin filaments generate tension forces, which affect the spatial organization of the intercellular adhesive structures. Collision of two moving fibroblastic cells is followed by the formation of unstable adhesions between the overlapping leading edge of one cell and lower leading edge of another one. The adherens junctions in fibroblastic cells, typically N-cadherin-based [75], are organized into strands oriented radially, that is, perpendicular to the boundary of the intercellular contact. Their orientation is caused by the centripetal (directed toward a center) tension forces generated by the contractile actin filament bundles that are associated with adhesive structures (Fig. 9.3) [76]. Circular actin filament bundle in an isolated epithelial cell generates tension forces, summary vector of which is directed centripetally (Fig. 9.5a). After collision of epithelial cells, their contacting edges immediately form very stable E-cadherinbased adherens junctions. In the zone of the intercellular contact circular actin bundles are disassembled, and new arch-like bundles are formed [77]. The contraction of these bundles generates tension forces, with the summary vector directed along the boundary of the intercellular contact from the center to the periphery (Fig. 9.5b). This tension vector determines the spatial organization of the adherens junctions, which are oriented linearly along the boundary of the intercellular contact (Fig. 9.2). Furthermore, the summary tension vector produces lateral expansion of the intercellular contact. The linear orientation of the adherens junctions along contacting epithelial cell edges is regulated by mDia and Rac1 [77, 78]. The collision of a fibroblastic cell and an epithelial one does not lead to the formation of stable heterotypical cell–cell adhesion. The epithelial cell reorganizes its actin filament system into the pattern more typical of a fibroblast. The spatial organization of these heterotypical adhesion structures is fairly similar to the organization of adherens junctions between fibroblastic cells [79]. Therefore, different signaling pathways can dynamically regulate adherens junctions. Local assembly and organization of actin cytoskeleton, controlled by catenins and small GTPases affects the formation and integrity of adherens junctions, which can regulate cell–cell adhesion. Changes in cadherin clustering caused by catenin tyrosine phosphorylation or by shifts in cadherin dimerization/dissociation equilibrium can directly affect the adhesive function of cadherin receptors.
196
9 Intercellular Adhesive Interactions
a
b
X X X X X X X X X
X X X X X X X X X
Fig. 9.5 The model of epithelial intercellular contact formation: two epithelial cells before their collision (a) or after the contact formation (b). Actin microfilament bundles (yellow lines): circular bundles (a) or arch-like bundles (b); tension forces (black arrows); summary vectors (red arrows) of the tension forces: the vectors directed centripetally (a) or along the boundary of the intercellular contact (b) (see the text for explanation). Reproduced with permission from the Proceedings of the National Academy of Sciences USA [77]. Copyright 1997 National Academy of Sciences, USA
Adherens junctions, like focal adhesions, are mechanosensitive: the mechanical stimulation strengthens them. The external physical forces can be transmitted along the actin filaments into the cytoplasm and nucleus, thereby influencing gene activities [80, 81]. Besides providing dynamic mechanical bonds between cells, adherens junctions are critically involved in the regulation of cell migration and proliferation [82]. Contact inhibition of cell migration (see Sect. 6.1.3) and contact inhibition of cell proliferation (see further) are phenomena that demonstrate these important adherens junction-mediated regulations. In the regulation of contact inhibition of cell migration (see Sect. 6.1.3), one of catenins, p120-catenin, plays a role. This catenin is concentrated in cadherin-based intercellular adhesions, whereas in isolated moving cells, p120-catenin is distributed throughout the cytoplasm. The increased level of p120-catenin in the cytoplasm causes significant elevation of activities of Rac and Cdc42, which in turn, results in the increased pseudopodial activity of the cells and in the stimulation of their locomotion. Thus, p120-catenin may couple the formation and disruption of cadherinbased intercellular adhesions with the regulation of cell motility via modulation of Rho GTPase activity [64, 83, 84].
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
197
Fig. 9.6 Diagram showing the “contact inhibition of cell proliferation” phenomenon (see the text for explanation)
9.1.3 Contact Inhibition of Cell Proliferation Striking examples of the “outside-in” signaling pathways triggered by cadherins are the pathways controlling cell proliferation. This control is demonstrated by the phenomenon of contact inhibition of cell proliferation (Fig. 9.6): normal cells stop their multiplication with the achievement of such density of the cell monolayer, when there are no more cell-free matrix surface areas. If a part of the confluent monolayer is mechanically removed (a “wound” in the cell monolayer), then the cells, which were at the “wound” edge, begin to proliferate and to migrate into the free surface of the “wound.” Thus, it seems very likely that steady intercellular contacts inhibit cell proliferation. There are different “outside-in” signaling pathways coupling cadherin-mediated intercellular contacts to cell proliferation. One of them is the Wnt signal transduction pathway (see below). In this pathway, b-catenin is a key component [85, 86]. In a cell, b-catenin is present in two different states: bound to cadherin cytoplasmic domain state (b-catenin at adherens junctions) and in the free state (soluble b-catenin in the cytoplasm). Cytoplasmic b-catenin functions as a transcriptional co-activator, promoting gene transcription: from the cytoplasm, b-catenin is translocated into the cell nucleus where it promotes transcription of specific genes, including the ones that initiate cell proliferation (Fig. 9.4) [85]. The signal activity of free b-catenin and its binding with the cadherin cytoplasmic domain compete with each other: the lessening of the binding increases a fraction of free cytoplasmic b-catenin thereby stimulating cell proliferation. The dynamic equilibrium between the bound and free states of b-catenin in cells is controlled by the system regulating the level of free b-catenin in the cytoplasm.
198
9 Intercellular Adhesive Interactions
The proteins APC and Axin/conductin play a crucial role in the regulation of the lifetime and the transcriptional activity of free cytoplasmic b-catenin [87]. These proteins are products of tumor suppression genes. The gene coding for axin is called Conductin (or Axil), and so the name Axin/conductin is commonly used. APC binds directly to various proteins including b-catenin and Axin/conductin forming the cytoplasmic multiprotein complex. Two serine/threonine protein kinases, glycogen synthase kinase-3b (GSK3b) and its priming kinase, casein kinase1, are recruited to this complex causing multiple phosphorylation of free b-catenin (Fig. 9.7). As a result, free b-catenin finally undergoes proteasomal degradation, and its level in the cytoplasm drops [88, 89]. Therefore, APC and axin inducing the degradation of free cytoplasmic b-catenin are negative regulators of the Wnt signaling pathway (see further). The integrin-linked kinase (ILK) is another important regulator of free b-catenin level in the cytoplasm: ILK inhibits GSK3b activity and thereby induces the increase in the free b-catenin [90]. Thus, ILK as a regulating element is involved in the Wnt signaling (see below). The increase in the content of free b-catenin in the cytoplasm is mediated by Wnt signaling pathway (Fig. 9.7): the Wnt proteins (products of WNT proto-oncogene) bind to the transmembrane receptors of the Frizzled family. This binding causes phosphorylation and activation of the Dishevelled (Dsh) cytoplasmic protein, which translocates to the cell membrane and inhibits GSK3b. As a result, the level of free unphosphorylated (“activated”) b-catenin increases in the cytoplasm. This activated b-catenin penetrates into the cell nucleus where it functions as a transcription co-activator: b-catenin gets associated with the Tcf/Lef (T-cell factor/lymphocyteenhancer factor) family of transcription factors to activate the expression of target genes, including those that stimulate cell proliferation [85]. The participation of b-catenin as a structural component in adherens junction formation and its involvement in the Wnt signaling pathway compete with each other. Therefore, the Wnt signaling activation is coupled with a loosening of cadherin adhesive bonds between epithelial cells. This is observed during epithelialmesenchymal transitions (EMTs), regeneration, and other developmental processes, as well as during cancer progression. Thus, b-catenin couples the loss of cell–cell adhesion to increased Wnt signaling. In the switch between b-catenin’s adhesive and signal functions, Bcl 9-2 protein plays essential role. Bcl 9-2 binds to the amino-terminal portion of b-catenin. This portion overlaps the a-catenin-binding region of b-catenin. There is a tyrosine residue within this region that needs to be phosphorylated for efficient interaction between b-catenin and Bcl 9-2. Therefore, this tyrosine determines whether b-catenin binds to a-catenin, or its phosphorylation diverts b-catenin from the intercellular adhesion to Bcl 9-2, that is, to the Wnt signaling pathway (Fig. 9.7). Thus, the “choice” of b-catenin between its adhesive and transcriptional functions is determined by the competitive binding to a-catenin vs. Bcl 9-2. Besides, Bcl 9-2 is involved in the translocation of free b-catenin from the cytoplasm into the cell nucleus [26]. Another protein, APC, also influences that “choice” of b-catenin: APC can compete with E-cadherin and the Tcf/Lef transcription factors for binding to b-catenin (Fig. 9.7).
9.1 Cadherin-Mediated Intercellular Contacts: Adherens Junctions
199
Fig. 9.7 Diagram of the b-catenin and Wnt-signaling pathway (see the text for explanation)
In addition, there can be another factor influencing the “choice” of b-catenin between its adhesive and transcriptional functions. This factor is the presence of distinct b-catenin molecular forms having different binding properties. A closed conformation of b-catenin binds Tcf/Lef transcriptional factors alone, whereas an open conformation binds both cadherins and Tcf/Lef factors. Therefore, the Wnt signaling pathway coupling cadherin-mediated intercellular contacts to cell proliferation can be the basis of “contact inhibition of cell proliferation” phenomenon. Another cause of this phenomenon is the activation of some tumor suppressor genes.
200
9 Intercellular Adhesive Interactions
The increased E-cadherin expression and the formation of intercellular adhesive junctions initiate signal transduction pathways (their components are still not known in detail), which activate INK4a and p27/KIP1 tumor suppressor genes in epithelial cells. These genes encode proteins, which block the entry of cells into the phase of DNA replication, and this way, these proteins inhibit cell proliferation [91]. The product of p53 tumor suppressor gene negatively regulates cell proliferation, initiating its blocking in special situations. The formation of confluent epithelial cell monolayer causes p53 protein accumulation in the cells and inhibition of their proliferation. The disconnecting of the epithelial cells in the monolayer and/or inactivating mutations of the CDH1 tumor suppressor gene that encodes E-cadherin cause the destabilization of p53 and consequently, cell proliferation is resumed. Thus, E-cadherin apparently initiates intracellular signals that lead to p27/KIP1 and p53 gene activation and to the block of proliferation of the epithelial cells when they establish stable intercellular contacts. One of ways of p53-dependent inhibition of cell proliferation is the effect of p53 on the dynamic equilibrium between the bound and free states of b-catenin. P53 stimulates the phosphorylation of free cytoplasmic b-catenin thereby favoring its degradation. The level of free b-catenin in the cytoplasm and Wnt signaling are downregulated, and as a result, cell proliferation is inhibited [92].
9.2 Altered Regulation of Adherens Junctions Caused by Oncogenic Transformation Oncogenic transformation causes serious alterations of cell–cell adhesion. These changes endow cancer cells with invasive and metastatic abilities [26, 93]. When moving transformed cells, both epithelial cells and fibroblasts, collide with each other, their behavior is similar to that of normal fibroblastic cells (see Sect. 6.1.3). After the collision, transformed cells do not form stable intercellular contacts, and the cells continue to move and underlap each other. These underlappings are common in dense populations of transformed cells (Fig. 9.8). If the cell-free extracellular matrix surface is available, the transformed cells move away from one another after their collisions. The changes of cell–cell adhesion lead to significant weakening of mechanical bonds between transformed tissue cells, the acquisition of invasion ability by the cells, and permanent stimulation of their proliferation. Mutations of the genes, which encode proteins participating in the formation and regulation of cadherinmediated intercellular bonds, are the basis of these changes. The development and progression of many sporadic or heritable human tumors are associated with those gene mutations. The proteins that are key participants in the formation and dynamic regulation of cadherin-based intercellular contacts can be the products of both proto-oncogenes (e.g., CTNNB1 encoding b-catenin or Wnt1 proto-oncogene) and tumor suppressor genes (e.g., CDH1 gene that encodes E-cadherin, APC or Conductin genes).
9.2 Altered Regulation of Adherens Junctions Caused by Oncogenic Transformation
201
Fig. 9.8 Transformed hamster fibroblasts. The cells overlap each other. SEM. Scale bar, 14 mm
Activating mutations of proto-oncogenes and/or inactivating mutations of tumor suppressor genes cause steady destabilization of cadherin-based adhesions and alterations in the intracellular signaling pathways [26]. These events can finally contribute to the development of many sporadic or heritable epithelial tumors.
9.2.1 Alterations in Cadherin–Catenin Complex Cadherins Loss or dysfunctions of E-cadherin significantly contribute to the transition of human benign adenomas to carcinomas and are critically involved in human cancer progression, stimulating tumor cell proliferation and invasion [29, 30, 94, 95]. Inactivation of E-cadherin encoding CDH1 tumor suppressor gene causes the loss of this intercellular adhesive receptor [26]. The deficiency of E-cadherin causes serious attenuation of cell–cell adhesion bonds. Besides, because of mutagenic loss of E-cadherin expression, the level of cytoplasmic free b-catenin is increased, and it is translocated into the cell nucleus to carry out the co-transcriptional functions (see Sect. 9.1.3). The loss of E-cadherin expression is a typical feature of many human carcinomas [30, 94]. Even a single mutation of the CDH1 gene can be responsible for neoplastic transformation of adenoma to malignant invasive carcinoma. CDH1 gene mutations or its deletions in germ cells are the causal genetic defects for inherited stomach cancer. The inactivating CDH1 mutations in somatic cells can cause many forms of
202
9 Intercellular Adhesive Interactions
sporadic cancers: lobular breast carcinoma, stomach cancer, and many others. The loss of E-cadherin expression is directly related to invasiveness and progression of many human malignant epithelial tumors. Inhibition of E-cadherin expression in malignant tumors can be achieved by not only CDH1 gene mutations but also by its transcriptional repression [26]. The CDH1 transcription repressors are expressed in many types of human cancers, including breast and ovarian cancers, melanomas, and other cancer types. The expression of some CDH1 transcription repressors often correlates with poor clinical prognosis. CDH1 transcription repressors inhibit not only E-cadherin gene expression but also inhibit many different genes in epithelial cells, and thereby decrease the expression of some proteins of tight or desmosomal junctions, and actin-binding protein gelsolin. CDH1 transcription repressors can influence proliferation and inhibit apoptosis in tumor cells. The loss of E-cadherin expression in cancer cells can be also caused by its cleavage by lysosomal proteinases cathepsins that are often translocated to the cell surfaces during cancer progression [96]. The deficiency of E-cadherin adhesive dimers resulting in the weakening of cell– cell adhesion can also be caused by enhanced cadherin endocytosis in cancers. Altered regulation of E-cadherin internalization is typical for the oncogenic transformation of cells. Since cadherin endocytosis is stimulated in response to soluble GFs, permanent activation of the oncogenes encoding some GF receptors causes increased cadherin endocytosis in cancer cells. The expression of p120-catenin, which prevents cadherin endocytosis and thereby regulates the stability of cadherin-mediated adhesion, is decreased up to its complete disappearance in many human malignant tumors. The product of the Mdm2 oncogene, which can trigger E-cadherin endocytosis through its binding to E-cadherin, is overexpressed in several human tumor types including soft tissue sarcomas, osteosarcomas, and breast cancers. High levels of Mdm2 protein contribute to the acquisition of invasive ability by the tumor cells [26]. Therefore, altered regulation of E-cadherin endocytosis in tumor cells resulting in the breaking of the intercellular adhesive bonds and enhancement of the migration ability of the cells contributes to tumor progression. A small GTPase Rap1 required for maturation of adherens junctions is inactive in some human cancers as a result of the inactivating mutations of the guanine nucleotide exchange factor (GEF) that “turns on” Rap1 activity [54]. In human tumor cells, the switching from one cadherin isoform to another one is often observed. For example, aberrant P-cadherin expression occurs during adenomatous and hyperplastic colon polyp development prior to changes in E-cadherin, b-catenin, or APC expression [97]. In normal mammary gland, P-cadherin is expressed in myoepithelial cells but not in the epithelial ones. However, many breast ductal carcinomas express P-cadherin, though these malignant tumors are thought to be epithelial origin. P-cadherin expression in breast carcinomas strongly correlates with poor cell differentiation, lack of estrogen receptors, and increased cell proliferation [98]. N-cadherin that is typical for mesenchymal tissues, such as connective or nerve tissues, is aberrantly expressed in many human carcinomas. N-cadherin expression induces the EMT and stimulates migratory activity of cancer cells, thereby contributing to cancer invasion [30, 36, 98].
9.2 Altered Regulation of Adherens Junctions Caused by Oncogenic Transformation
203
Thus, cadherin isoform switching apparently allows cancer cells to leave the primary tumor and disseminate. The invasion of cancer cells into the surrounding normal tissues is associated with EMT [99–102]. The transition from epithelial to fibroblast-like phenotype is accompanied by the disruption of E-cadherin-mediated intercellular adhesions in epithelial cells and acquisition of fibroblast-like phenotype suitable for active cell migration (see Sect. 6.1.4). The repression of E-cadherin gene CDH1 caused by its transcription repressors is key regulator of EMT in cancers [26]. Cancer cells in the center of the tumor and at its invasive front are different: the latter undergo EMT [103]. In a result of EMT, invading cancer cells acquire fibroblast-like morphology, high motility, and invasive properties. EMT allows cancer cells to disseminate from the primary tumor and metastasize. The acquisition of these characteristics is caused by the loss of expression of epithelial-specific genes and by the acquisition of expression of mesenchymal genes (e.g., CDH2 gene encoding N-cadherin). It is interesting that even in cases, when after the oncogenic transformation, epithelial cells retain their E-cadherin-based adhesions and the ability to form confluent epithelial cell monolayers, the arrangement of their adherens junctions changes from the linear orientation along the contacting cell edges (characteristic for normal epithelial cells, Fig. 9.2) to the radial orientation, that is, perpendicular to the boundary of the intercellular contact (characteristic for fibroblastic cells, Fig. 9.3). The actin cytoskeletal control of the arrangement of E-cadherin-based adherens junctions is different in normal and transformed epithelial cells. In normal epitheliocytes, the linear orientation of adherens junctions is regulated by mDia and Rac1, whereas in transformed epithelial cells, similar to fibroblastic cells, the radial orientation of the adherens junctions is critically determined by myosin II-driven contractility of actin filaments [78]. Some components of the extracellular matrix in epithelial tumors, in particular, types I and III collagen, have the ability to reduce E-cadherin expression and to initiate the disruption of the adhesions between the tumor cells, promoting their EMT and proliferation. The molecular mechanism of this matrix collagen-induced effect includes signaling pathways from activated integrin receptors, integrin-mediated activation of FAK, its recruitment to the E-cadherin complex, and increased tyrosine phosphorylation of b-catenin [104–106]. Catenins Alterations of catenins are also implicated in malignant tumor formation and progression. The phosphorylation status of b-catenin is an important factor in the maintenance of stable cadherin-based intercellular adhesion. The signals triggered by nonreceptor tyrosine kinases (e.g., submembraneous Src kinase) or receptor tyrosine kinases, such as receptors of soluble GFs, cause the disassembly of cadherin-based adhesions via the tyrosine phosphorylation of b-catenin. In malignant tumor cells, because of the permanent activation of oncogenes, such as Src or genes encoding GF receptors (e.g., c-ErbB1/HER1 or c-met), b-catenin is maintained in tyrosine
204
9 Intercellular Adhesive Interactions
phosphorylated state, causing the disassembly of cadherin-based bonds between cancer cells, and their EMT [26]. Separation of b-catenin from a-catenin induced by a GTPase activating protein IQGAP1 contributes to the weakening of E-cadherin-mediated intercellular adhesion in malignant tumors; IQGAP1 is overexpressed in a range of human cancers [107, 108]. b-catenin as a signaling molecule plays an especially important role in tumorigenesis. Mutations of the CTNNB1 proto-oncogene that encodes b-catenin increase its stability in the cytoplasm of cancer cells. These mutations prevent free cytoplasmic b-catenin degradation and favor its accumulation in the cytoplasm with subsequent translocation into the cell nucleus to function as transcriptional co-activator inducing cancer cell proliferation (see below). p120-Catenin is implicated in cancer progression. P120-catenin expression, which inhibits E-cadherin endocytosis and promotes actin cytoskeleton assembly in adherens junctions, is strongly decreased in many human malignant tumors. Loss of p120-catenin causes marked E-cadherin deficiency, disassembly of cell–cell adhesions, and EMT of epithelial tumor cells favoring their dissemination [26].
9.2.2 Loss of Contact Inhibition of Cell Proliferation (Fig. 9.6) As a result of oncogenic transformation, the dynamic balance between b-catenin bound to cadherin in adherens junctions and free cytoplasmic b-catenin is disturbed. This change leads to significant accumulation of free b-catenin in the cytoplasm of cancer cells. There are several causes of free b-catenin accumulation. As a result of the loss of E-cadherin expression, the level of bound b-catenin is decreased; this leads to the significant increase of free cytoplasmic b-catenin. Mutations of the CTNNB1 proto-oncogene deprive free cytoplasmic b-catenin of its ability to be phosphorylated by GSK3b, thereby preventing free b-catenin from its degradation and favoring its accumulation in the cytoplasm. The inactivating mutations in the APC or Axin/conductin tumor suppressor genes, which encode key molecules to target b-catenin for its degradation (see Sect. 9.1.3), have the same effect. These mutations prevent APC from binding to cytoplasmic b-catenin and Axin/conductin from binding to APC or GSK3b. Following the inhibition of free b-catenin degradation, its level in the cytoplasm is increased. The mutagenic activation of the proto-oncogene encoding Wnt glycoprotein causes permanent inhibition of GSK3b and, as a result, increases the content of free b-catenin in the cytoplasm. GSK3b is involved not only in the degradation of free b-catenin and thereby in the negative regulation of the Wnt signaling pathway, but also in the regulation of E-cadherin expression [109]. One of the transcription repressors of the gene encoding E-cadherin, SNAIL1 protein, is inhibited by its phosphorylation caused by GSK3b. In cancer cells, permanent activation of the antiapoptotic signaling pathway (see Sect. 5.4.3), and, specifically, the hyperactivation of PKB/Akt kinase, leads to phosphorylation of GSK3b causing its proteasomal degradation. As a result of the GSK3b degradation, the stability of SNAIL1 is increased, leading to the inhibition
References
205
of E-cadherin expression [110]. Because of E-cadherin deficiency, the level of free b-catenin in the cancer cell cytoplasm significantly increases. The accumulating free b-catenin is translocated from the cytoplasm of cancer cells into their nuclei to co-activate Wnt-responsive genes including those initiating cell proliferation. Since APC, besides its suppressing effect on Wnt signaling, is also involved in the regulation of actin cytoskeleton and microtubules, in lamellipodia formation and cell migration (see Sects. 3.2 and 6.1.1), the inactivating mutations in the APC gene lead not only to sustained cell proliferation but also to the changes in cytoskeleton organization and altered migration of cancer cells [111]. The CTNNB1 or APC gene mutations in human germ cells can cause inherited precancerous disease, familial adenomatous polyposis coli that often transits to invasive colon cancer. The same gene mutations in somatic cells can cause sporadic colon carcinoma and other types of human cancer. The Axin/conductin gene inactivating mutations cause sporadic human hepatocellular and colon carcinoma . Overexpression of Bcl 9-2 protein, which is involved in the translocation of free cytoplasmic b-catenin into the cell nucleus, is found in human colon carcinomas [26]. Therefore, changes in the Wnt signaling pathway regulation are typical for many human cancers. Inhibitors of free b-catenin-mediated transcription can thus be considered for cancer therapy [112]. Defective intercellular contacts cannot initiate signals activating INK4a, p27/ KIP1, or p53 tumor suppressor genes that inhibit cell proliferation. The inactivating mutations of these suppressor genes also result in the loss of contact inhibition. Because of that, tumor cells weakly bound to each other continue to proliferate. The activation of some proto-oncogenes (e.g., Ras) that causes the degradation of p27/KIP1 protein can also lead to the loss of contact inhibition [91]. Therefore, the proliferation of the adherens junction-defective cancer cells gets permanently stimulated. It is possible, that not only adherens junctions but also desmosomal junctions are implicated in tumor progression. The loss of desmoplakin in breast cancers correlates with enhanced tumor cell proliferation, suggesting that desmosomal proteins might play a role in suppressing breast cancer progression. Together with the loss of substrate dependence of cell proliferation by tumor cells, their loss of contact inhibition of cell proliferation plays a key role in such basic manifestations of malignant tumor growth as uncontrolled cell proliferation, inability of tumor cells to form orderly tissue structures (tissue anaplasia), tumor invasion and metastasis.
References 1. Brooks SA, Lomax-Browne HJ, Carter TM, Kinch CE, Hall DM (2010) Molecular interactions in cancer cell metastasis. Acta Histochem 112(1):3–25. doi:10.1016/j.acthis.2008.11.022 DOI:dx.doi.org 2. Weber GF (2008) Molecular mechanisms of metastasis. Cancer Lett 270(2):181–190. doi:10.1016/j.canlet.2008.04.030 DOI:dx.doi.org
206
9 Intercellular Adhesive Interactions
3. Geiger TR, Peeper DS (2009) Metastasis mechanisms. Biochim Biophys Acta 1796(2): 293–308. doi:10.1016/j.bbcan.2009.07.006 DOI:dx.doi.org 4. Niessen CM (2007) Tight junctions/adherens junctions: basic structure and function. J Invest Dermatol. 127(11):2525–2532. doi:10.1038/sj.jid.5700865 DOI:dx.doi.org 5. Hartsock A, Nelson WJ (2008) Adherens and tight junctions: structure, function and connections to the actin cytoskeleton. Biochim Biophys Acta 1778(3):660–669. doi:10.1016/ j.bbamem.2007.07.012 DOI:dx.doi.org 6. Marchiando AM, Graham WV, Turner JR (2010) Epithelial barriers in homeostasis and disease. Annu Rev Pathol 5:119–144. doi:10.1146/annurev.pathol.4.110807.092135 DOI:dx.doi.org 7. Ouban A, Ahmed AA (2010) Claudins in human cancer: a review. Histol Histopathol 25(1):83–90. 8. Lal-Nag M, Morin PJ (2009) The claudins. Genome Biol 10(8):235. doi:10.1186/gb-200910-8-235 DOI:dx.doi.org 9. Fanning AS, Anderson JM (2009) Zonula occludens-1 and −2 are cytosolic scaffolds that regulate the assembly of cellular junctions. Ann N Y Acad Sci 1165:113–120. doi:10.1111/j.1749-6632.2009.04440.x DOI:dx.doi.org 10. Terry S, Nie M, Matter K, Balda MS (2010) Rho signaling and tight junction functions. Physiology (Bethesda) 25(1):16–26. doi:10.1152/physiol.00034.2009 DOI:dx.doi.org 11. Balda MS, Matter K (2009) Tight junctions and the regulation of gene expression. Biochim Biophys Acta 1788(4):761–767. doi:10.1016/j.bbamem.2008.11.024 DOI:dx.doi.org 12. Popoff MR, Geny B (2009) Multifaceted role of Rho, Rac, Cdc42 and Ras in intercellular junctions, lessons from toxins. Biochim Biophys Acta 1788(4):797–812. doi:10.1016/ j.bbamem.2009.01.011 DOI:dx.doi.org 13. Nishimura N, Sasaki T (2009) Rab family small G proteins in regulation of epithelial apical junctions. Front Biosci.; 14:2115–2129. doi:10.2741/3366 DOI:dx.doi.org 14. Samarin S, Nusrat A (2009) Regulation of epithelial apical junctional complex by Rho family GTPases. Front Biosci 14:1129–1142. doi:10.2741/3298 DOI:dx.doi.org 15. Olk S, Zoidl G, Dermietzel R (2009) Connexins, cell motility, and the cytoskeleton. Cell Motil Cytoskeleton 66(11):1000–1016. doi:10.1002/cm.20404 DOI:dx.doi.org 16. D’hondt C, Ponsaerts R, De Smedt H, Bultynck G, Himpens B (2009) Pannexins, distant relatives of the connexin family with specific cellular functions? Bioessays 31(9):953–974. doi:10.1002/bies.200800236 DOI:dx.doi.org 17. Hesketh GG, Van Eyk JE, Tomaselli GF (2009) Mechanisms of gap junction traffic in health and disease. J Cardiovasc Pharmacol 54(4):263–272. doi:10.1097/FJC.0b013e3181ba0811 DOI:dx.doi.org 18. Jansen JA, van Veen TA, de Bakker JM, van Rijen HV (2010) Cardiac connexins and impulse propagation. J Mol Cell Cardiol 48(1):76–82. doi:10.1016/j.yjmcc.2009.08.018 DOI:dx.doi.org 19. Green KJ, Getsios S, Troyanovsky S, Godsel LM (2010) Intercellular junction assembly, dynamics, and homeostasis. Cold Spring Harb Perspect Biol 2(2):a000125. doi:10.1101/cshperspect.a000125 DOI:dx.doi.org 20. Holthöfer B, Windoffer R, Troyanovsky S, Leube RE (2007) Structure and function of desmosomes. Int Rev Cytol 264:65–163. doi:10.1016/S0074-7696(07)64003-0 DOI:dx.doi.org 21. Garrod D, Kimura TE (2008) Hyper-adhesion: a new concept in cell-cell adhesion. Biochem Soc Trans 36(Pt 2):195–201. doi:10.1042/BST0360195 DOI:dx.doi.org 22. Gloushankova NA, Wakatsuki T, Troyanovsky RB, Elson E, Troyanovsky SM (2003) Continual assembly of desmosomes within stable intercellular contacts of epithelial A-431 cells. Cell Tissue Res 314(3):399–410. doi:10.1007/s00441-003-0812-3 DOI:dx.doi.org 23. Hatzfeld M (2007) Plakophilins: Multifunctional proteins or just regulators of desmosomal adhesion? Biochim Biophys Acta 1773(1):69–77. doi:10.1016/j.bbamcr.2006.04.009 DOI:dx. doi.org 24. Pokutta S, Weis WI (2007) Structure and mechanism of cadherins and catenins in cell-cell contacts. Annu Rev Cell Dev Biol 23:237-261. doi:10.1146/annurev.cellbio.22.010305.104241 DOI:dx.doi.org 25. Niessen CM, Gottardi CJ (2008) Molecular components of the adherens junction. Biochim Biophys Acta 1778(3):562–571. doi:10.1016/j.bbamem.2007.12.015 DOI:dx.doi.org
References
207
26. Gloushankova NA (2008) Changes in regulation of cell-cell adhesion during tumor transformation. Biochemistry (Mosc) 73(7):742–750. doi:10.1134/S000629790807002X DOI:dx.doi.org 27. Vestweber D (2008) VE-cadherin: the major endothelial adhesion molecule controlling cellular junctions and blood vessel formation. Arterioscler Thromb Vasc Biol 28(2):223–232. doi:10.1161/ATVBAHA.107.158014 DOI:dx.doi.org 28. Goodwin M, Yap AS (2004) Classical cadherin adhesion molecules: coordinating cell adhesion, signaling and the cytoskeleton. J Mol Histol 35(8-9):839–844. doi:10.1007/s10735-0041833-2 DOI:dx.doi.org 29. van Roy F, Berx G (2008) The cell-cell adhesion molecule E-cadherin. Cell Mol Life Sci 65(23):3756–3788. doi:10.1007/s00018-008-8281-1 DOI:dx.doi.org 30. Stemmler MP (2008) Cadherins in development and cancer. Mol Biosyst 4(8):835–850. doi:10.1039/b719215k DOI:dx.doi.org 31. Mochizuki N (2009) Vascular integrity mediated by vascular endothelial cadherin and regulated by sphingosine 1-phosphate and angiopoietin-1. Circ J 73(12):2183–2191. doi:10.1253/ circj.CJ-09-0666 DOI:dx.doi.org 32. Dejana E, Tournier-Lasserve E, Weinstein BM (2009) The control of vascular integrity by endothelial cell junctions: molecular basis and pathological implications. Dev Cell 16(2):209–221. doi:10.1016/j.devcel.2009.01.004 DOI:dx.doi.org 33. Gavard J (2009) Breaking the VE-cadherin bonds. FEBS Lett 583(1):1–6. doi:10.1016/j.febslet.2008.11.032 DOI:dx.doi.org 34. Vestweber D, Winderlich M, Cagna G, Nottebaum AF (2009) Cell adhesion dynamics at endothelial junctions: VE-cadherin as a major player. Trends Cell Biol 19(1):8–15. doi:10.1016/j.tcb.2008.10.001 DOI:dx.doi.org 35. Halbleib JM, Nelson WJ (2006) Cadherins in development: cell adhesion, sorting, and tissue morphogenesis. Genes Dev 20(23):3199–3214. doi:10.1101/gad.1486806 DOI:dx.doi.org 36. Wheelock MJ, Shintani Y, Maeda M, Fukumoto Y, Johnson KR (2008) Cadherin switching. J Cell Sci 121(Pt 6):727–735. doi:10.1242/jcs.000455 DOI:dx.doi.org 37. Perez-Moreno M, Fuchs E (2006) Catenins: keeping cells from getting their signals crossed. Dev Cell 11(5):601–612. doi:10.1016/j.devcel.2006.10.010 DOI:dx.doi.org 38. McCrea PD, Gu D (2010) The catenin family at a glance. J Cell Sci. 123(Pt 5):637–642. doi:10.1242/jcs.039842 DOI:dx.doi.org 39. Takai Y, Ikeda W, Ogita H, Rikitake Y (2008) The immunoglobulin-like cell adhesion molecule nectin and its associated protein afadin Annu Rev Cell Dev Biol 24:309–342. doi:10.1146/ annurev.cellbio.24.110707.175339 DOI:dx.doi.org 40. Ogita H, Ikeda W, Takai Y (2008) Roles of cell adhesion molecules nectin and nectin-like molecule-5 in the regulation of cell movement and proliferation. J Microsc 231(3):455–465. doi:10.1111/j.1365-2818.2008.02058.x DOI:dx.doi.org 41. Rikitake Y, Takai Y (2008) Interactions of the cell adhesion molecule nectin with transmembrane and peripheral membrane proteins for pleiotropic functions. Cell Mol Life Sci 65(2):253–263. doi:10.1007/s00018-007-7290-9 DOI:dx.doi.org 42. Nishimura T, Takeichi M (2009) Remodeling of the adherens junctions during morphogenesis. Curr Top Dev Biol 89:33–54. doi:10.1016/S0070-2153(09)89002-9 DOI:dx.doi.org 43. Ogita H, Takai Y (2008) Cross-talk among integrin, cadherin, and growth factor receptor: roles of nectin and nectin-like molecule. Int Rev Cytol 265:1–54. doi:10.1016/S00747696(07)65001-3 DOI:dx.doi.org 44. Delva E, Kowalczyk AP (2009) Regulation of cadherin trafficking. Traffic. 10(3):259–267. doi:10.1111/j.1600-0854.2008.00862.x DOI:dx.doi.org 45. Wirtz-Peitz F, Zallen JA (2009) Junctional trafficking and epithelial morphogenesis. Curr Opin Genet Dev 19(4):350–356. doi:10.1016/j.gde.2009.04.011 DOI:dx.doi.org 46. Troyanovsky RB, Sokolov EP, Troyanovsky SM (2006) Endocytosis of cadherin from intracellular junctions is the driving force for cadherin adhesive dimer disassembly. Mol Biol Cell 17(8):3484–3493. doi:10.1091/mbc.E06-03-0190 DOI:dx.doi.org 47. Yap AS, Crampton MS, Hardin J (2007) Making and breaking contacts: the cellular biology of cadherin regulation. Curr Opin Cell Biol 19(5):508–514. doi:10.1016/j.ceb.2007.09.008 DOI:dx.doi.org
208
9 Intercellular Adhesive Interactions
48. Troyanovsky SM (2009) Regulation of cadherin-based epithelial cell adhesion by endocytosis. Front Biosci (Schol Ed) 1:61–67. doi:10.2741/s7 DOI:dx.doi.org 49. Schill NJ, Anderson RA (2009) Out, in and back again: PtdIns(4,5)P(2) regulates cadherin trafficking in epithelial morphogenesis. Biochem J 418(2):247–260. doi:10.1042/BJ20081844 DOI:dx.doi.org 50. Hong S, Troyanovsky RB, Troyanovsky SM (2010) Spontaneous assembly and active disassembly balance adherens junction homeostasis. Proc Natl Acad Sci U S A 107(8):3528–3533. doi:10.1073/pnas.0911027107 DOI:dx.doi.org 51. Yasmeen A, Bismar TA, Al Moustafa AE (2006) ErbB receptors and epithelial-cadherincatenin complex in human carcinomas. Future Oncol 2(6):765–781. doi:10.2217/ 14796694.2.6.765 DOI:dx.doi.org 52. Stehbens SJ, Akhmanova A, Yap AS (2009) Microtubules and cadherins: a neglected partnership. Front Biosci.; 14:3159–3167. doi:10.2741/3442 DOI:dx.doi.org 53. Boettner B, Van Aelst L (2009) Control of cell adhesion dynamics by Rap1 signaling. Curr Opin Cell Biol 21(5):684–693. doi:10.1016/j.ceb.2009.06.004 DOI:dx.doi.org 54. Kooistra MR, Dubé N, Bos JL (2007) Rap1: a key regulator in cell-cell junction formation. J Cell Sci 120(Pt 1):17–22. doi:10.1242/jcs.03306 DOI:dx.doi.org 55. Pannekoek WJ, Kooistra MR, Zwartkruis FJ, Bos JL (2009) Cell-cell junction formation: the role of Rap1 ans Rap1 guanine nucleotide exchange factors. Biochim Biophys Acta 1788 (4): 790–796. doi:10.1016/j.bbamem.2008.12.010 DOI:dx.doi.org 56. Mège RM, Gavard J, Lambert M (2006) Regulation of cell-cell junctions by the cytoskeleton. Curr Opin Cell Biol 18(5):541–548. doi:10.1016/j.ceb.2006.08.004 DOI:dx.doi.org 57. Chhabra ES, Higgs HN (2007) The many faces of actin: matching assembly factors with cellular structures. Nat Cell Biol. 9(10):1110–1121. doi:10.1038/ncb1007-1110 DOI:dx.doi.org 58. Ren G, Crampton MS, Yap AS (2009) Cortactin: Coordinating adhesion and the actin cytoskeleton at cellular protrusions. Cell Motil Cytoskeleton 66(10):865–873. doi:10.1002/ cm.20380 DOI:dx.doi.org 59. Bershadsky A (2004) Magic touch: how does cell-cell adhesion trigger actin assembly? Trends Cell Biol 14(11):589–593. doi:10.1016/j.tcb.2004.09.009 DOI:dx.doi.org 60. Vasiliev JM (2004) Cytoskeletal mechanisms responsible for invasive migration of neoplastic cells. Int J Dev Biol 48(5-6):425–439. doi:10.1387/ijdb.041806jv DOI:dx.doi.org 61. Trichet L, Sykes C, Plastino J (2008) Relaxing the actin cytoskeleton for adhesion and movement with Ena/VASP. J Cell Biol 181(1):19–25. doi:10.1083/jcb.200710168 DOI:dx.doi.org 62. Rivard N (2009) Phosphatidylinositol 3-kinase: a key regulator in adherens junction formation and function. Front Biosci 14:510–22. doi:10.2741/3259 DOI:dx.doi.org 63. Carramusa L, Ballestrem C, Zilberman Y, Bershadsky AD (2007) Mammalian diaphanousrelated formin Dia1 controls the organization of E-cadherin-mediated cell-cell junctions. J Cell Sci 120(Pt 21):3870–3882. doi:10.1242/jcs.014365 DOI:dx.doi.org 64. Anastasiadis PZ (2007) p120-ctn: A nexus for contextual signaling via Rho GTPases. Biochim Biophys Acta 1773(1):34–46. doi:10.1016/j.bbamcr.2006.08.040 DOI:dx.doi.org 65. Zandy NL, Pendergast AM (2008) Abl tyrosine kinases modulate cadherin-dependent adhesion upstream and downstream of Rho family GTPases. Cell Cycle 7(4):444–448. doi:10.4161/ cc.7.4.5452 DOI:dx.doi.org 66. Stambolic V, Woodgett JR (2006) Functional distinctions of protein kinase B/Akt isoforms defined by their influence on cell migration. Trends Cell Biol 16(9):461–466. doi:10.1016/j. tcb.2006.07.001 DOI:dx.doi.org 67. Scott JA, Yap AS (2006) Cinderella no longer: alpha-catenin steps out of cadherin’s shadow. J Cell Sci 119(Pt 22):4599–4605. doi:10.1242/jcs.03267 DOI:dx.doi.org 68. Benjamin JM, Nelson WJ (2008) Bench to bedside and back again: molecular mechanisms of alpha-catenin function and roles in tumorigenesis. Semin Cancer Biol 18(1):53–64. doi:10.1016/j.semcancer.2007.08.003 DOI:dx.doi.org 69. Drees F, Pokutta S, Yamada S, Nelson WJ, Weis WI (2005) Alpha-catenin is a molecular switch that binds E-cadherin-beta-catenin and regulates actin-filament assembly. Cell 123(5):903–915. doi:10.1016/j.cell.2005.09.021 DOI:dx.doi.org
References
209
70. Benjamin JM, Kwiatkowski AV, Yang C, Korobova F, Pokutta S, Svitkina T, Weis WI, Nelson WJ (2010) AlphaE-catenin regulates actin dynamics independently of cadherin-mediated cell-cell adhesion. J Cell Biol 189(2):339–352. doi:10.1083/jcb.200910041 DOI:dx.doi.org 71. Pokutta S, Drees F, Yamada S, Nelson WJ, Weis WI (2008) Biochemical and structural analysis of alpha-catenin in cell-cell contacts. Biochem Soc Trans 36(Pt 2):141–147. doi:10.1042/ BST0360141 DOI:dx.doi.org 72. Kuroda S, Fukata M, Nakagawa M, Fujii K, Nakamura T, Ookubo T, Izawa I, Nagase T, Nomura N, Tani H, Shoji I, Matsuura Y, Yonehara S, Kaibuchi K (1998) Role of IQGAP1, a target of the small GTPases Cdc42 and Rac1, in regulation of E-cadherin- mediated cell-cell adhesion. Science 281(5378):832–835. doi:10.1126/science.281.5378.832 DOI:dx.doi.org 73. Fukata M, Kuroda S, Nakagawa M, Kawajiri A, Itoh N, Shoji I, Matsuura Y, Yonehara S, Fujisawa H, Kikuchi A, Kaibuchi K (1999) Cdc42 and Rac1 regulate the interaction of IQGAP1 with beta-catenin. J Biol Chem 274(37):26044–26050. doi:10.1074/jbc.274.37.26044 DOI:dx.doi.org 74. Grosheva I, Vittitow JL, Goichberg P, Gabelt BT, Kaufman PL, Borrás T, Geiger B, Bershadsky AD (2006) Caldesmon effects on the actin cytoskeleton and cell adhesion in cultured HTM cells. Exp Eye Res 82(6):945–58. doi:10.1016/j.exer.2006.01.006 DOI:dx.doi.org 75. El Sayegh TY, Kapus A, McCulloch CA (2007) Beyond the epithelium: cadherin function in fibrous connective tissues. FEBS Lett 581(2):167–174. doi:10.1016/j.febslet.2006.12.029 DOI:dx.doi.org 76. Gloushankova NA, Krendel MF, Alieva NO, Bonder EM, Feder HH, Vasiliev JM, Gelfand IM (1998) Dynamics of contacts between lamellae of fibroblasts: essential role of the actin cytoskeleton. Proc Natl Acad Sci U S A 95(8):4362–4367. doi:10.1073/pnas.95.8.4362 DOI:dx.doi.org 77. Gloushankova NA, Alieva NA, Krendel MF, Bonder EM, Feder HH, Vasiliev JM, Gelfand IM (1997) Cell-cell contact changes the dynamics of lamellar activity in nontransformed epitheliocytes but not in their ras-transformed descendants. Proc Natl Acad Sci U S A 94(3):879– 883. doi:10.1073/pnas.94.3.879 DOI:dx.doi.org 78. Ayollo DV, Zhitnyak IY, Vasiliev JM, Gloushankova NA (2009) Rearrangements of the actin cytoskeleton and e-cadherin-based adherens junctions caused by neoplasic transformation change cell-cell interactions. PLoS One 4(11):e8027. 79. Omelchenko T, Fetisova E, Ivanova O, Bonder EM, Feder H, Vasiliev JM, Gelfand IM (2001) Adhesive interactions between epitheliocytes and fibroblasts: formation of heterotypic cadherin-containing adhesion sites is accompanied by local cytoskeletal reorganization. Proc Natl Acad Sci USA 98(15):8632–8637. doi:10.1073/pnas.151247698 DOI:dx.doi.org 80. Schwartz MA, DeSimone DW (2008) Cell adhesion receptors in mechanotransduction. Curr Opin Cell Biol 20(5):551–556. doi:10.1016/j.ceb.2008.05.005 DOI:dx.doi.org 81. Wang N, Tytell JD, Ingber DE (2009) Mechanotransduction at a distance: mechanically coupling the extracellular matrix with the nucleus. Nat Rev Mol Cell Biol 10(1):75–82. doi:10.1038/nrm2594 DOI:dx.doi.org 82. Rudini N, Dejana E (2008) Adherens junctions. Curr Biol 18(23):R1080-2. doi:10.1016/ j.cub.2008.09.018 DOI:dx.doi.org 83. Grosheva I, Shtutman M, Elbaum M, Bershadsky AD (2001) p120 catenin affects cell motility via modulation of activity of Rho-family GTPases: a link between cell-cell contact formation and regulation of cell migration. J Cell Sci 114(Pt 4):695–707. 84. Boguslavsky S, Grosheva I, Landau E, Shtutman M, Cohen M, Arnold K, Feinstein E, Geiger B, Bershadsky A (2007) p120 catenin regulates lamellipodial dynamics and cell adhesion in cooperation with cortactin. Proc Natl Acad Sci U S A 104(26):10882–10887. doi:10.1073/ pnas.0702731104 DOI:dx.doi.org 85. Jin T, George Fantus I, Sun J (2008) Wnt and beyond Wnt: multiple mechanisms control the transcriptional property of beta-catenin. Cell Signal 20(10):1697–1704. doi:10.1016/j.cellsig.2008.04.014 DOI:dx.doi.org 86. MacDonald BT, Tamai K, He X (2009) Wnt/beta-catenin signaling: components, mechanisms, and diseases. Dev Cell 17(1):9–26. doi:10.1016/j.devcel.2009.06.016 DOI:dx.doi.org
210
9 Intercellular Adhesive Interactions
87. Brocardo M, Henderson BR (2008) APC shuttling to the membrane, nucleus and beyond. Trends Cell Biol 18(12):587–596. doi:10.1016/j.tcb.2008.09.002 DOI:dx.doi.org 88. Aoki K, Taketo MM (2007) Adenomatous polyposis coli (APC): a multi-functional tumor suppressor gene. J Cell Sci 120(Pt 19):3327–3335. doi:10.1242/jcs.03485 DOI:dx.doi.org 89. McCartney BM, Näthke IS (2008) Cell regulation by the Apc protein Apc as master regulator of epithelia. Curr Opin Cell Biol 20(2):186–193. doi:10.1016/j.ceb.2008.02.001 DOI:dx.doi.org 90. Boulter E, Van Obberghen-Schilling E (2006) Integrin-linked kinase and its partners: a modular platform regulating cell-matrix adhesion dynamics and cytoskeletal organization. Eur J Cell Biol 85(3-4):255–263. doi:10.1016/j.ejcb.2005.09.002 DOI:dx.doi.org 91. Del Pozo MA, Schwartz MA (2007) Rac, membrane heterogeneity, caveolin and regulation of growth by integrins. Trends Cell Biol 17(5):246–250. 92. Levina E, Oren M, Ben-Ze’ev A (2004) Downregulation of beta-catenin by p53 involves changes in the rate of beta-catenin phosphorylation and Axin dynamics. Oncogene 23(25):4444–4453. doi:10.1038/sj.onc.1207587 DOI:dx.doi.org 93. Makrilia N, Kollias A, Manolopoulos L, Syrigos K (2009) Cell adhesion molecules: role and clinical significance in cancer. Cancer Invest 27(10):1023–1037. doi:10.3109/0735790090 2769749 DOI:dx.doi.org 94. Tsanou E, Peschos D, Batistatou A, Charalabopoulos A, Charalabopoulos K (2008) The E-cadherin adhesion molecule and colorectal cancer. A global literature approach. Anticancer Res 28(6A):3815–3826. 95. Jeanes A, Gottardi CJ, Yap AS (2008) Cadherins and cancer: how does cadherin dysfunction promote tumor progression? Oncogene 27(55):6920–6929. doi:10.1038/onc.2008.343 DOI:dx.doi.org 96. Gocheva V, Joyce JA (2007) Cysteine cathepsins and the cutting edge of cancer invasion. Cell Cycle 6(1):60–64. doi:10.4161/cc.6.1.3669 DOI:dx.doi.org 97. Hardy RG, Tselepis C, Hoyland J, Wallis Y, Pretlow TP, Talbot I, Sanders DS, Matthews G, Morton D, Jankowski JA (2002) Aberrant P-cadherin expression is an early event in hyperplastic and dysplastic transformation in the colon. Gut 50(4):513–519. doi:10.1136/ gut.50.4.513 DOI:dx.doi.org 98. Knudsen KA, Wheelock MJ. (2005) Cadherins and the mammary gland. J Cell Biochem 95(3):488–496. doi:10.1002/jcb.20419 DOI:dx.doi.org 99. Klymkowsky MW, Savagner P (2009) Epithelial-mesenchymal transition: a cancer researcher’s conceptual friend and foe. Am J Pathol 174(5):1588–1593. doi:10.2353/ajpath.2009.080545 DOI:dx.doi.org 100. Voulgari A, Pintzas A (2009) Epithelial-mesenchymal transition in cancer metastasis: mechanisms, markers and strategies to overcome drug resistance in the clinic. Biochim Biophys Acta 1796(2):75–90. doi:10.1016/j.bbcan.2009.03.002 DOI:dx.doi.org 101. Yilmaz M, Christofori G (2009) EMT, the cytoskeleton, and cancer cell invasion. Cancer Metastasis Rev 28(1-2):15–33. doi:10.1007/s10555-008-9169-0 DOI:dx.doi.org 102. Vergara D, Merlot B, Lucot JP, Collinet P, Vinatier D, Fournier I, Salzet M (2010) Epithelialmesenchymal transition in ovarian cancer. Cancer Lett 291(1):59-66. doi:10.1016/j. canlet.2009.09.017 DOI:dx.doi.org 103. De Wever O, Pauwels P, De Craene B, Sabbah M, Emami S, Redeuilh G, Gespach C, Bracke M, Berx G (2008) Molecular and pathological signatures of epithelial-mesenchymal transitions at the cancer invasion front. Histochem Cell Biol 130(3):481–494. doi:10.1007/s00418008-0464-1 DOI:dx.doi.org 104. Moustakas A, Heldin CH (2007) Signaling networks guiding epithelial-mesenchymal transitions during embryogenesis and cancer progression. Cancer Sci 98(10):1512–1520. doi:10.1111/j.1349-7006.2007.00550.x DOI:dx.doi.org 105. Imamichi Y, Menke A (2007) Signaling pathways involved in collagen-induced disruption of the E-cadherin complex during epithelial-mesenchymal transition. Cells Tissues Organs 185(1-3):180–190. doi:10.1159/000101319 DOI:dx.doi.org 106. Guarino M, Rubino B, Ballabio G (2007) The role of epithelial-mesenchymal transition in cancer pathology. Pathology 39(3):305–318. doi:10.1080/00313020701329914 DOI:dx.doi.org
References
211
107. Johnson M, Sharma M, Henderson BR (2009) IQGAP1 regulation and roles in cancer. Cell Signal 21(10):1471–1478. doi:10.1016/j.cellsig.2009.02.023 DOI:dx.doi.org 108. White CD, Brown MD, Sacks DB (2009) IQGAPs in cancer: a family of scaffold proteins underlying tumorigenesis. FEBS Lett 583(12):1817–1824. doi:10.1016/j.febslet.2009.05.007 DOI:dx.doi.org 109. Wu D, Pan W (2010) GSK3: a multifaceted kinase in Wnt signaling. Trends Biochem Sci 35(3):161-168. doi:10.1016/j.tibs.2009.10.002 DOI:dx.doi.org 110. Qiao M, Sheng S, Pardee AB (2008) Metastasis and AKT activation. Cell Cycle 7(19):2991–2996. 111. Näthke I (2005) Relationship between the role of the adenomatous polyposis coli protein in colon cancer and its contribution to cytoskeletal regulation. Biochem Soc Trans 33(Pt 4): 694–697. doi:10.1042/BST0330694 DOI:dx.doi.org 112. Huang SM, Mishina YM, Liu S, Cheung A, Stegmeier F, Michaud GA, Charlat O, Wiellette E, Zhang Y, Wiessner S, Hild M, Shi X, Wilson CJ, Mickanin C, Myer V, Fazal A, Tomlinson R, Serluca F, Shao W, Cheng H, Shultz M, Rau C, Schirle M, Schlegl J, Ghidelli S, Fawell S, Lu C, Curtis D, Kirschner MW, Lengauer C, Finan PM, Tallarico JA, Bouwmeester T, Porter JA, Bauer A, Cong F (2009) Tankyrase inhibition stabilizes axin and antagonizes Wnt signalling. Nature 461(7264):614-620. doi:10.1038/nature08356 DOI:dx.doi.org
wwwwwwwwwwwwwwww
Chapter 10
Conclusions
The alterations to specific genes, proto-oncogenes, and/or tumor suppressor genes, can arise spontaneously or be induced by chemical carcinogens, ultraviolet and ionizing radiation, or oncogenic viruses. These gene alterations result in the oncogenic transformation of tissue cells that leads to subsequent development of a malignant tumor. The oncogenic transformation is accompanied by serious abnormalities in cell–extracellular matrix and cell–cell adhesive interactions. As a result of the deficiency of adhesive bonds of transformed cells with the matrix and with each other, dysregulation of intracellular signal transduction pathways initiated by these adhesive bonds takes place. In normal cells, these signaling pathways control cell survival, proliferation, morphological characteristics of the cells, and their migratory activity. Survival and proliferation of normal tissue cells critically depend on their adhesion to the extracellular matrix, but transformed cells lose this dependence. Whereas proliferation of normal cells is negatively controlled by the intercellular adhesion, transformed cells lose this control. Unlike normal cells, transformed ones are “ascetic”: they can proliferate well and migrate in the conditions of considerable deficiency of soluble growth factors. The migration of transformed cells, in comparison with normal ones, is much less regulated by the adhesive interactions with adjacent cells or the geometric configuration of the extracellular matrix. The impairment of adhesive interactions strongly changes the behavior of transformed cells giving them such anomalous characteristics as the ability to get detached from the extracellular matrix and from each other relatively easily, the ability to survive in suspended state, the tolerance for the deficiency of growth factors, the ability to actively migrate and penetrate surrounding healthy tissues, the loss of the ability to form normal tissue structures, and the maintenance of sustained high proliferative activity. The inability of transformed cells to abide by the local regulating influences allows defining their behavior as an “antisocial” one [1]. This genetically determined cell behavior is the basis of the principal features of malignant tumor progression, including such characteristics as tissue anaplasia (loss of normal tissue structure), tumor invasion, and metastasis. The phenotypical characteristics of malignant tumor cells that determine their ability for invasion and metastasis include: dysregulation of adhesive interactions of Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2_10, © Springer Science+Business Media, LLC 2011
213
214
10 Conclusions
tumor cells with each other, with normal cells of the microenvironment, and with the extracellular matrix; overproduction of matrix metalloproteinases; “epithelialmesenchymal transition” and acquisition of locomotory phenotype that includes alterations of cell morphology and cytoskeleton; induction of angiogenesis that provides additional routes for spread of primary tumor cells. These phenotypical signs are obviously determined by the expression of different molecules encoded by the genes, which are either activators or suppressors of malignant tumor invasion and/or metastasis. It is necessary to say that both activators and suppressors code for specific molecules necessary for normal cell physiology; however, they activate or suppress tumor growth, invasion, and metastasis depending on their expression changes. Usually, genes-activators have increased expression in tumors, whereas genes-suppressors become repressed. The genes encoding some of the intercellular heterotypical adhesion molecules, which provide tight attachment of circulating tumor cells to the microvascular endothelium (specifically, integrin receptors typical for leukocytes or platelets and expressed by tumor cells), can be examples of the activators of invasion and metastasis. This tumor cell attachment is an important precondition for subsequent invasion of the vascular wall and the formation of secondary (metastatic) foci of malignant tumor growth. The other genes activating tumor invasion include those encoding specific extracellular matrix glycoproteins (such as laminin-411 or laminin-332), and integrin receptors providing the adhesion of tumor cells to these matrix components. Malignant tumor invasion may be activated by matrix metalloproteinases, and also soluble growth factors and their cell surface receptors. The group of the genes-activators also includes the genes encoding key proteins involved in the mitogenic, morphogenic, or antiapoptotic signal transduction pathways triggered by cell–extracellular matrix or cell–cell adhesive interactions and also by growth factors. Sustained activation of these genes, i.e., their conversion into oncogenes, favors acquisition of invasive and metastatic properties by tumor cells. Various genes encoding intercellular homotypic adhesion molecules (e.g., E-cadherin) or certain molecules of cell–extracellular matrix adhesion (e.g., fibronectin receptor a5b1 integrin) can be considered as suppressors of tumor invasion. The pivotal role of the abnormalities of cell adhesive interactions in the acquisition of invasive and metastatic activities by tumor cells are considered as a basis for the development of new perspective drugs aiming at counteracting malignant tumor invasion and metastasis. Such drugs could be used in the treatment of human malignant tumors. These drugs could include different groups. One group includes integrin antagonists, such as RGD-based synthetic peptides, which compete with the extracellular matrix ligands for the binding to integrin receptors, as well as integrin blocking antibodies. These agents could prevent the attachment of circulating malignant tumor cells to microvascular endothelium and thereby prevent their invasion. Besides, some integrin antagonists are antiangiogenic agents, e.g., such as RGD-based peptides specific for aV-containing integrins, antibodies to aVb3 or a5b1 integrins, small nonpeptide inhibitors, or peptidomimetic
References
215
signaling inhibitors of aV or a5b1 integrins, are recognized as perspective agents for the treatment of pathological ocular neovascularization [2]. These antiangiogenic integrin antagonists could also emerge as promising agents for inhibiting neoangiogenesis in cancers, thereby preventing progressive tumor growth. Small molecule inhibitors of VEGF receptor signaling are also promising for prevention of neovascularization in malignant tumors. The drugs that restore the functions of E-cadherin-catenin complex could be potentially effective in cancer invasion treatment. Inhibitors of key signaling proteins, such as protein tyrosine kinases involved in the hyperactivated mitogenic, morphogenic, or antiapoptotic pathways that are triggered by adhesive interactions and/or growth factors could be promising agents for human cancer therapy.
References 1. Vasiliev JM (1997) Social behavior of normal cells and anti-social behavior of tumor cells. Soros Educational Journal 5:20–25. 2. Rangasamy S, Ljubimov AV, Roy S, McGuire P, Das A (2010) Proteinases and integrins in ocular angiogenesis. In: Das A, Friberg T (eds) Therapy of ocular angiogenesis: Principles and Practice. Lippincott Williams & Wilkins, Baltimore
wwwwwwwwwwwwwwww
Index
A ABL tyrosine kinases, 193 Actin-binding proteins, 16–21, 24, 52, 102, 123, 125, 132, 189, 202 Actin cytoskeleton, 13, 16–19, 24, 28, 29, 44, 47, 52, 69, 72, 84, 91, 93, 94, 98, 101–106, 108, 123, 125, 128, 130, 132, 158, 169, 177, 180, 185–187, 189, 190, 192–197, 203–205 Actin-depolymerizing factor(ADF), 17–18. See also Cofilin Actin dynamics, 19, 126, 193, 194 Actin filaments, 13–26, 28–30, 37, 40, 42, 44, 45, 47, 49, 50, 52, 66, 69, 71, 72, 78, 80, 87, 88, 99, 102–104, 106, 109, 123–130, 133, 151, 154, 158, 178–181, 186, 187, 190, 193–196, 203 Actin polymerization, 13, 14, 16, 18, 21, 24, 49, 103, 104, 123, 124, 126, 127, 193 Actin-related proteins (Arp2/3), 16, 18, 19, 21, 23, 52, 103, 104, 123–125, 127, 193, 194 Activation, 19, 24, 47, 48, 50, 57, 88–93, 96–100, 103–106, 109–112, 125–127, 129–134, 136, 177, 190, 192–194, 198–205, 214 Active cell edge, 24, 80, 125, 127, 128, 133, 134 Actomyosin, 19, 49, 51, 108 ADAM proteins, 48, 50 Adenomatous polyposis coli (APC) protein, 28, 126, 195, 198, 204 ADF. See Actin-depolymerizing factor (ADF) Adherens junctions, 185–205
Adhesions, 1, 2, 7, 13, 14, 18, 19, 24, 29, 30, 57, 60, 64, 66, 90, 94, 95, 99–101, 105–112, 121–123, 125, 127–136, 153, 169, 177, 180, 185–196, 198, 200, 203, 204, 213, 214 Adhesion structures, 30, 37–52, 108, 112, 123, 127–129, 195 Adhesive functions, 7, 48, 108–110, 185, 191, 192, 195 Adhesive interactions, 1–4, 10, 13, 44, 57–112, 130, 185–205, 213–215 Afadin, 189 Affinity, 16, 47, 48, 50, 100, 109, 134, 192 Alpha-actinin, 18, 47, 106, 189 Alpha-catenin, 18, 127, 189, 194, 195, 198, 204 Alpha-tubulin, 25 Alterations, 2–4, 13, 57, 74–88, 104, 105, 108–112, 121, 132–135, 169, 177, 180, 185, 200–204, 213, 214 AMF. See Autocrine motility factor (AMF) Anaplasia, 185, 205, 213 Anchorage dependence, 2, 99–100 Anchorage independence, 4, 57, 111–112 Angiogenesis, 48, 52, 90, 112, 131, 135, 136, 214 Anoikis, 2, 57, 99, 100, 106, 108, 111, 112 Anti-apoptotic signaling, 97–100, 111, 204 APC. See Adenomatous polyposis coli (APC) protein Apoptosis, 2, 3, 57, 94, 97–100, 105, 106, 108, 111, 152, 202 Arginylation, 13, 14 Arg kinase, 193 Arp2/3. See Actin-related proteins (Arp2/3) Asef. See Rac1-activating protein (Asef) Assembling, 24, 52, 193
Y.A. Rovensky, Adhesive Interactions in Normal and Transformed Cells, DOI 10.1007/978-1-61779-304-2, © Springer Science+Business Media, LLC 2011
217
218 ATP hydrolysis, 19 Attachment, 2, 9, 10, 21, 24, 37, 42, 44, 57, 69, 106, 109, 122, 124, 130, 136, 145, 148, 151, 177, 214 Autocrine motility factor (AMF), 137 Autophagy, 100 Axin, 198, 204, 205 Axin/conductin, 198, 204, 205 B Bad protein, 98, 99 Barbed ends, 17–19, 21, 23, 42, 44, 103, 104, 106, 124, 129 Basement membranes, 1, 2, 7–10, 51, 52, 112, 131, 135, 145, 147, 148, 153 Bax protein, 98 BAX tumor suppressor gene, 98, 100 Bcl2 protein, 98 BCL2 proto-oncogene, 98, 100 Bcl 9–2 protein, 198, 205 BCL-x, proto-oncogene, 98 Bcl-x protein, 98 Beta-catenin, 189–195, 197–205 Beta-tubulin, 25, 26 Blebs, 37, 42, 58, 60, 69, 80, 84, 87, 126 Bombesin, 136 BPAG1, BP230. See Bullous pemphigoid antigen1 (BPAG1, BP230) Bullous pemphigoid antigen1 (BPAG1, BP230), 51 Bundles, 14, 15, 18, 24, 28, 44, 45, 48, 51, 52, 66, 68, 70–72, 80, 82, 88, 102, 104, 106, 107, 123–125, 128, 133, 151, 154, 158, 178–181, 190, 193, 195, 196 C Cadherin isoform switching, 203 Cadherin receptor, 129, 185, 187, 190–195 Calcium ions, 92 Caldesmon, 19, 49, 52, 195 Calmodulin, 92 Calmodulin dependent protein kinases (CAM), 93 Calpain, 48, 50 Calpastatin, 48, 50 CAM. See Calmodulin dependent protein kinases (CAM) cAMP. See Cyclic adenosine monophosphate (cAMP) cAMP-dependent protein kinase, 93 Cancer cells, 10, 24, 51, 52, 108–112, 129, 133–136, 154, 161, 200–205
Index Cancer progression, 112, 198, 201, 202, 204, 205 Capping, 16–18, 21, 103, 124, 191, 2344 Cas. See P130 Cas (Cas) protein Cas-associated protein, 95 Casein kinase1, 198 Catastrophe frequency, 50 Catenin complex, 187–189, 194, 195, 201–204, 215 Catenins, 187, 189, 193, 195, 196, 203–204 Caveolae, 99 Caveolin–1, 99 Cdc42, 101, 104–105, 124, 126, 129, 133, 176, 190, 193, 195, 196 CDH1 tumor suppressor gene, 200, 201 Cell cultivation, 151 Cell detachment, 97, 99, 100, 111 Cell-extracellular matrix adhesion, 105, 108 Cell length control, 71, 72 Cell motility, 18, 19, 49, 98, 130–132, 194, 196 Cell receptors, 18, 88, 91, 110, 130, 135 Cell spreading, 19, 24, 37, 57–88, 107, 108, 148, 151, 154, 155, 157, 158, 167–171, 173, 174, 178 Cell surface receptors, 7–9, 47, 57, 88–92, 94, 96, 111, 128, 129, 214 Cell surface relief, 57–62, 69, 72, 82–87 Centrioles, 25 Centripetal stretch, 49, 66, 78, 105, 106, 108, 109, 122, 178, 181 Centrosomal microtubules, 27 Centrosome, 25, 27, 104, 126 c-ErbB1/HER1 proto-oncogene, 110 Chondroitin sulfates, 8 Claudins, 185, 186, 189 Clustering, 47, 90, 109, 124, 192, 195 c-met proto-oncogene, 110, 136 Cobl. See Cordon-bleu (Cobl) Cofilin, 17–19, 23, 24, 102, 103, 134. See also Actin-depolymerizing factor(ADF) Colcemide, 28, 126 Colchicines, 28 Collagen fibers, 9 Collagens, 1, 4, 8–10, 45, 48, 52, 90, 106, 107, 109, 112, 129, 135, 203 Collision, 121, 127, 195, 196, 200 Conductin. See Axin/conductin Conductin (Axil) tumor suppressor gene. See Axin/conductin Conformation, 19, 48, 88, 89, 91, 108, 186, 194, 199 Connective tissue, 2, 4, 7, 8, 10, 96, 145, 148–150 Connexins, 186 Connexon channels, 186 Connexons, 186
Index Contact guidance, 153 Contact inhibition, 2, 121, 127–128, 134, 185, 196–200, 204–205 Contact paralysis, 127, 128, 134, 194 Contractility, 14, 19, 24, 29, 49–52, 105–107, 126, 127, 129, 133, 180, 193, 195, 203 Contraction, 19, 49, 80, 93, 107, 108, 130, 186, 195 Cordon-bleu (Cobl), 16, 17 Cortactin, 18, 21, 52, 123, 125, 193 Cortical actin, 18, 49, 126, 130, 192 Curvature, 153–156, 158, 178–181 Cyclic adenosine monophosphate (cAMP), 16, 92, 93 Cyclic nucleotides, 92 Cylindrical surface, 154, 163, 167, 179, 180 Cylindrical threads, 154, 158 Cytochalasins, 84, 87 Cytoskeleton, 2, 3, 13–30, 44, 45, 47, 52, 69, 72, 84, 91, 93, 94, 98, 100–106, 108, 121, 123, 125, 128, 130, 132, 158, 169, 177, 180, 185–187, 189, 190, 192–197, 204, 205, 214 Cytoskeleton linkers, 30 D DAG. See Diacylglycerol (DAG) DAP. See Death-associated protein (DAP) Death-associated protein (DAP), 99 Deletions, 105, 111, 201 Depolymerization, 14, 16, 17, 23, 24, 26–29, 49, 87, 102, 103, 126 Dermatan sulfates, 8 Desmocollins, 186 Desmoglein, 186 Desmoplakin, 187, 205 Desmosomal cadherins, 186 Desmosomal junctions. See Desmosomes (desmosomal junctions) Desmosomes (desmosomal junctions), 30, 185–187, 189, 202, 205 Desmuslin, 30 Detachment, 4, 40, 57, 60, 63, 97, 99, 100, 111, 122, 126, 130, 170 Diacylglycerol (DAG), 92, 94 Disassembly, 17, 23, 29, 48–50, 94, 101, 104, 109, 129–132, 186, 191, 193, 203, 204 Discontinuous substrata, 154, 167–176 Dishevelled (Dsh) protein, 198 Disintegrin domain, 48 Domain, 8–10, 16–19, 23, 45, 47, 48, 52, 89–93, 95, 96, 99, 100, 108, 109, 123, 185, 187, 189, 191–193, 197
219 Dsh. See Dishevelled (Dsh) protein Dynamic instability, 26–28, 50 Dynamin, 50, 99, 131 Dynein, 27, 30, 104, 192 E EB1. See End-binding protein1 (EB1) E-cadherin. See Epithelial cadherin (E-cadherin) EGF. See Epidermal growth factor (EGF) Elastin, 1, 8, 9 Elongation, 17, 21, 23, 122, 124, 152, 154, 158, 159, 179, 180 Embryogenesis, 2, 10, 121, 129, 151, 154, 181, 185, 188 EMT. See Epithelial-mesenchymal transition (EMT) Enabled/vasodilator-stimulated phosphoprotein (Ena/VASP), 106 Ena/VASP. See Enabled/vasodilator-stimulated phosphoprotein (Ena/VASP) End-binding protein1 (EB1), 28 Endocytic-exocytic transport, 131, 134 Endocytosis, 19, 50, 99, 191, 192, 202, 204 Endothelium, 2, 4, 90, 96, 109, 214 Entactins, 9 Epidermal growth factor (EGF), 90, 96, 110, 130, 132, 136 Epithelial cadherin (E-cadherin), 129, 135, 187, 188, 191–195, 198, 200–205, 214, 215 Epithelial cells, 4, 7, 9, 14, 29, 30, 51, 58, 64, 66, 69, 71, 72, 74, 78, 80, 88, 90, 100, 122, 127–129, 131, 133, 134, 145, 154, 158, 161, 163, 167–170, 172, 174, 177–181, 187, 188, 191, 193, 195, 196, 198, 200, 202, 203 Epithelial-mesenchymal transition (EMT), 129, 135, 136, 198, 202–204, 214 Epitheliocytes. See Epithelial cells Epithelium, 29, 128 ERK. See Extracellular signal-regulated kinases (ERK) ERM. See Ezrin, radixin, moezin (ERM) Exocytosis, 131, 191 Expression, 3, 10, 17, 24, 30, 48, 50, 91, 93, 94, 96, 100, 105, 106, 109, 110, 125, 128, 129, 135, 136, 153, 180, 198, 200–205, 214 Extracellular matrix, 1, 7–10, 13, 37, 57–112, 121, 145, 153, 200, 213 Extracellular signal-regulated kinases (ERK), 96 Ezrin, radixin, moezin (ERM), 18
220 F F-actin, 13 FAK. See Focal adhesion kinase (FAK) Fascin, 18, 124, 129 FGF. See Fibroblast growth factor (FGF) Fibers, 1, 2, 9, 18, 40, 145, 147–150, 153 Fibrillar adhesions, 37, 44, 51 Fibrillin, 9, 10 Fibroblast growth factor (FGF), 52, 96, 110, 136 Fibroblastic cells, 4, 14, 15, 49, 51, 60, 63, 69, 71, 72, 78, 80, 105–108, 121–123, 125–128, 130, 132, 133, 145, 148, 153, 154, 158, 161–163, 167–170, 172, 173, 175, 177–181, 189, 190, 195, 200, 203 Fibroblasts, 4, 7–9, 19, 29, 38–40, 43, 44, 52, 58, 60, 61, 74–80, 83–87, 96, 110, 124, 128–130, 134–136, 158, 159, 163–167, 179–181, 195, 200, 201, 203 Fibronectin, 1, 4, 8–10, 45, 47, 48, 51, 52, 90, 109, 130, 134, 177, 214 Fibulins, 9, 10 Filament bundles, 14, 18, 45, 50, 52, 66, 68, 69, 71, 72, 80, 88, 102, 107, 124, 125, 128, 133, 151, 158, 178–181, 193, 195 Filament nucleation, 16, 17, 21 Filamins, 18 Filopodia, 18, 37–39, 42, 44, 45, 50, 64–66, 74, 75, 80, 83, 84, 104, 123–125, 129, 133, 149, 163, 172–174 Fimbrins, 18 FIP200 protein, 95 Focal adhesion kinase (FAK), 47, 49, 50, 95, 98, 105, 106, 110, 111, 131, 133, 177, 203 Focal adhesions. See Focal contacts (focal adhesions) Focal complexes, 37, 44, 45, 49–52, 80, 132 Focal contacts (focal adhesions), 37, 44–52, 57, 64, 66, 71, 78, 80, 88, 94, 95, 97–102, 104–112, 121, 125, 127–134, 152, 169, 178, 179, 181 Folds, 58–60, 62, 69, 80, 84, 86, 108, 153 Force-induced signals, 105, 108 Formin homology protein mDia1, 50, 102, 105, 193 Formin homology protein mDia2, 124 Formins, 16, 17, 21, 23, 50, 105, 123, 124, 193, 194 Forward flow, 130 Frizzled, 198
Index G G-actin. See Globular actin (G-actin) Gamma-catenin (g-catenin ), 187, 189 Gamma-tubulin (g-tubulin ), 25, 27 Gap junctions, 185, 186 GAPs. See GTPase activating proteins (GAPs) GDI. See GDP dissociation inhibitor (GDI) GDP. See Guanosine diphosphate (GDP) GDP dissociation inhibitor (GDI), 91 GEF-H1. See Rho guanine nucleotide exchange factor (GEF-H1) GEFs. See Guanine nucleotide exchange factors (GEFs) Gelsolin, 17–19, 103, 202 Gene, 3, 10, 17, 24, 28, 30, 93, 98–100, 105, 111, 112, 129, 130, 153, 180, 196–198, 200–205, 213 Geometric configuration, 10, 153, 154, 177, 181, 213 GFAP. See Glial fibrillary acidic protein (GFAP) Girdin, 18, 132 Glial fibrillary acidic protein (GFAP), 30 Glial filaments, 30 Globular actin (G-actin), 13, 16, 17 Glycogen synthase kinase–3b (GSK3b), 198 Glycoproteins, 1, 7–9, 45, 131, 186, 204, 214 Glycosaminoglycans, 1, 7–9 Golgi complex, 27 GPCRs. See G protein coupled receptors (GPCRs) G protein coupled receptors (GPCRs), 89–91 G proteins, 90, 91 Grooved topography, 161, 177 Grooves, 153, 154, 161–167, 177, 179–180 Growth factor receptors, 57, 94–108, 121, 123 Growth factors (GFs), 2, 3, 8, 52, 88, 121, 123, 128–130, 135, 191, 202, 203, 213–215 GSK3b. See Glycogen synthase kinase–3b GTP. See Guanosine triphosphate (GTP) GTPase. See Guanosine triphosphatase (GTPase) GTPase activating proteins (GAPs), 91, 105, 132 GTP hydrolysis, 26, 91 Guanine nucleotide exchange factors (GEFs), 49, 90, 105, 126, 192 Guanosine diphosphate (GDP), 26, 90–92 Guanosine triphosphatase (GTPase), 19, 24, 29, 44, 47, 49, 50, 69, 90–93, 95, 96, 98, 101, 102, 104–106, 110, 125, 126, 129, 131–133, 186, 192–196, 202, 204 Guanosine triphosphate (GTP), 26, 90–92 Guidance, 42, 132, 153
Index H Hemidesmosomes, 51 Heparan sulfates, 8 Heparin, 8, 9 Hepatocyte growth factor, 96, 110, 128–130, 136, 169, 171, 181 Heterotrimeric G proteins, 90–92 HGF/SF. See Hepatic growth factor Human cancers, 110, 111, 133, 136, 201, 202, 204, 205, 215 I IFs. See Intermediate filaments (IFs) IGF–1. See Insulin-like growth factor (IGF–1) ILK. See Integrin-linked kinase (ILK) Inactivation, 3, 99, 108, 111, 112, 201 INK4a tumor suppressor gene, 200, 205 Inositol triphosphate (IP3), 92–94, 193 Inside-out signaling, 100 Insulin-like growth factor (IGF–1), 96, 136 Integrin activation, 47, 48, 50, 90, 106, 109, 192 Integrin affinity, 48, 50 Integrin cytoplasmic domains, 47 Integrin extracellular domains, 47, 48 Integrin-linked kinase (ILK), 48, 50, 198 Integrin-mediated cell adhesion, 48, 99, 130 Integrin-mediated cell-matrix adhesion, 47, 49, 57, 99, 134 Integrin-mediated signaling, 90, 94, 101 Integrin receptors, 8, 9, 45, 47, 48, 52, 57, 90, 92, 94–96, 100, 109–111, 123, 131, 134, 177, 203, 214 Integrins, 8, 45, 57, 121, 152, 177, 187, 214 Integrin trafficking, 45, 131 Intercellular adhesion, 2, 13, 90, 128, 129, 131, 135, 136, 185–187, 189–193, 195, 196, 198, 203, 204, 213 Intermediate filaments (IFs), 13, 14, 29–30, 51, 129, 187 Invadopodia, 37, 44, 51–52, 109, 112 Invasion, 4, 10, 13, 37, 52, 109, 111, 112, 121, 129, 132–136, 154, 161, 185, 200–203, 205, 213–215 Ion channel receptors, 89, 91 IP3. See Inositol triphosphate (IP3) IQGAP1. See IQ motif-containing GTPase activating protein1 (IQGAP1) IQ motif-containing GTPase activating protein1 (IQGAP1), 194, 195, 204 Isometric tension, 49, 105, 112
221 J JMY. See Junction-mediated regulatory protein (JMY) Junction-mediated regulatory protein (JMY), 16–17 Junctions, 1, 185–205 K Keratan sulfates, 8 Keratins, 29–30, 51, 129 Kindlins, 47–48 Kinesins, 27, 30, 127 L Lamellar cytoplasm, 24, 38, 40–43, 66–69, 71–78, 80–81, 122, 125, 130, 154, 165, 167–170, 172, 173, 176, 181 Lamellipodia, 14, 18, 37, 38, 40–42, 44, 45, 50, 64–66, 78–81, 83, 103, 104, 122–127, 129, 130, 133–134, 152, 205 Laminin(s), 1, 4, 8, 9, 45, 47, 52, 90, 109, 131, 134, 135, 214 Laminin isoforms, 45, 109, 131, 135 Lamins, 30 Leading cell edge, 40, 94, 121–123, 125, 126, 128, 130, 131 Leiomodin (Lmod), 16, 17 Ligand-receptor interaction, 91–93 Ligands, 8, 47, 48, 57, 88–91, 93–95, 100, 109, 123, 131, 134, 135, 214 LIM protein kinases, 102, 103 Lmod. See Leiomodin (Lmod) Locomotion, 2, 4, 94, 112, 121, 123, 125, 127, 128, 130, 133, 134, 136, 137, 161, 180, 185, 196 “Locomotory phenotype,” 122, 214 M Maculae adherens. See Desmosomes (desmosomal junctions) MAP kinases. See Mitogen activated protein kinases (MAP kinases) Matrix metalloproteinases (MMP), 45, 52, 106, 109, 112, 131, 135, 214 Matrix tension state, 108 Mdm2 protein, 202 Mechanical forces, 27, 42, 105–108 Mechanical sensors, 49 Mechanosensory switches, 105 Melanoma, 112, 136, 137, 202 Membrane transport, 93, 131
222 Membrane type 1-matrix metalloproteinase (MT1-MMP), 112 Mesothelium, 145–148 Metastasis, 10, 13, 52, 110, 112, 132, 133, 135, 185, 213, 214 Microcirculation system, 4 Microfibrils, 9 Microtubule associated protein (XMAP215), 27, 28 Microtubule catastrophes, 26, 50 Microtubule dynamics, 28 Microtubule growth, 26, 103, 126 Microtubule-organizing center. See Centrosome Microtubule plus-end tracking proteins, 28, 126 Microtubules, 13, 14, 16, 24–30, 42, 49–50, 69, 71, 102–105, 107, 123, 125–127, 129, 131, 132, 152, 161, 192, 205 Microtubule shrinkage, 26 Microtubule system, 104, 126, 192 Microtubule targeting, 29, 50, 127, 129, 131, 132, 192 Microvilli, 57, 58, 60, 61, 69, 72, 80, 84, 86, 87 Migration, 1, 2, 7, 8, 10, 13, 17–19, 24, 28, 29, 37, 42, 47, 48, 51, 52, 57, 88–94, 101, 105, 106, 121–137, 153, 161–165, 167, 178–180, 185, 188, 196, 202, 203, 205, 213 Mitogen activated protein kinases (MAP kinases), 96, 97, 99, 110, 133, 177 Mitogens, 3, 57, 95–97, 99, 106, 110–112, 132–134, 177, 214, 215 Mitogens-motogens, 128–130, 133, 135–137 Mitosis, 28–29, 37 Mitotic cycle, 13, 84, 96 Mitotic spindle, 24, 25, 27, 28 MLC. See Myosin light chain (MLC) MLC phosphatase, 19, 102 MMP. See Matrix metalloproteinases (MMP) Mobility, 134, 136 Morphogenic signaling, 100–105, 129, 132–134, 177, 214, 215 Motogenic signaling, 128, 133 Motogens, 128–130, 133, 135–137, 169 Motor proteins, 19, 27–28, 127, 192 Movement, 24, 27, 28, 47, 71, 91, 121, 122, 127, 131, 134, 164 MT1-MMP. See Membrane type 1-matrix metalloproteinase (MT1-MMP) MTOC. See Centrosome Mutations, 3, 10, 30, 109–112, 136, 200–202, 204–205 Myofibroblasts, 106–108 Myosin II, 19, 20, 24, 51, 64, 125, 192
Index Myosin II-driven contractility, 49, 50, 52, 102, 105, 195, 203 Myosin light chain (MLC), 19, 102 Myosins, 19, 20, 24, 27, 49–52, 64, 71, 102, 105, 125, 133–134, 192, 195, 203 N Nascent focal adhesions, 130 Necl. See Nectin-like (Necl) protein Nectin-like (Necl) protein, 128 Nectins, 128, 189–190 Neovascularization, 136, 215 Nestin, 30 Neurofilaments, 30 Nidogens, 1, 9 Nischarin, 132 Nocodazole, 107, 126 Nonmotor proteins, 27–29 Notch receptors, 48 NPF. See Nucleation-promoting factors (NPF) Nucleation, 16, 17, 21, 27, 28, 50, 124 Nucleation-promoting factors (NPF), 16 Nucleators, 16–17, 123 Nucleus, 17, 25, 30, 48, 50, 91, 94, 96, 106, 196–198, 201, 204, 205 N-WASP. See Wiscott-Aldrich syndrome protein (WASP) family O Occludin, 185, 186 Occluding junctions. See Zonula occludens (occluding junctions) Oligosaccharides, 1, 7 Oncoproteins, 3, 109, 110 Orientation, 2, 52, 104, 108, 121, 125, 126, 153, 158, 161, 167, 177–180, 195, 203 Outside-in signaling, 48, 50, 95, 100, 192–194, 197 Overexpression, 52, 133, 135, 136, 205 P P21-activated kinase (PAK), 96, 103, 104, 132 P21 protein, 133–134 P27/KIP1 tumor suppressor gene, 99, 200, 205 P53 tumor suppressor gene, 17, 99, 100, 111, 129–130, 134, 200, 205 p120-Catenin, 128, 189, 191–193, 196, 202, 204 P130 Cas (Cas) protein, 95, 105, 106, 131, 133 P140 Cas. See Cas-associated protein PAK. See P21-activated kinase (PAK)
Index Pannexins, 186 Particularly interesting new cystein-histidine rich (PINCH) protein, 48 Parvin, 48 Paxillin, 47, 49–52, 95 P-cadherin. See Placental cadherin (P-cadherin) PDGF. See Platelet-derived growth factor (PDGF) Peripherin, 30 Phagocytosis, 42 Phosphatase and tensin homolog (PTEN) protein/lipid phosphatase, 99, 111 Phosphatidylinositol (PI), 93 Phosphatidylinositol 3-kinase (PI3K), 93, 96, 98, 99, 101, 110, 111, 193 Phosphatidylinositol biphosphate (PIP2), 93–94, 103, 191 Phosphatidylinositol kinase (PIK), 93 Phosphatidylinositol phosphate kinase (PIPK), 93–94, 103, 191 Phosphatidylinositol triphosphate (PIP3), 93–94, 98, 99, 193 Phosphoinositides (PIPs), 92, 93, 191 Phospholipase C, 94 Phosphorylation, 19, 48, 91–92 PI. See Phosphatidylinositol (PI) PI3K. See Phosphatidylinositol 3-kinase (PI3K) PIK. See Phosphatidylinositol kinase (PIK) PINCH. See Particularly interesting new cystein-histidine rich (PINCH) protein Pinocytosis, 42 PIP2. See Phosphatidylinositol biphosphate (PIP2) PIP3. See Phosphatidylinositol triphosphate (PIP3) PIPK. See Phosphatidylinositol phosphate kinase (PIPK) PKA. See Protein kinase A (PKA) PKB. See Protein kinase B (PKB) PKB/Akt. See Protein kinase B (PKB/Akt) PKC. See Protein kinase C (PKC) Placental cadherin (P-cadherin), 187, 202 Plakins, 30, 51 Plakoglobin. See Gamma-catenin (g-catenin ) Plakophilins, 187 Plasma membrane, 18, 45, 51, 52, 89–91, 93, 99, 101, 123–126, 185, 186 Platelet-derived growth factor (PDGF), 96, 110, 128, 136, 190 Plectin, 30, 51 Plexin receptors, 132 Plexins, 132 Plus-end, 14, 25–28, 50, 102, 125–126, 132, 192
223 Podosomes, 37, 44, 51–52 Pointed ends, 17–19, 23 Polarization, 29, 47, 66, 69–74, 78, 107, 121, 123, 126–127, 129, 145, 151, 157, 179 Polymerization, 13, 14, 16–18, 21, 24–29, 42, 44, 49, 50, 52, 87, 102–104, 123, 124, 126, 127, 129, 193 Profilin, 17, 23, 24, 102 Proliferation, 2–4, 7, 10, 13, 48, 57, 88–90, 92–94, 96, 99–101, 105, 106, 108, 110–112, 123, 128, 129, 135–137, 152, 153, 177, 180, 185–187, 196–205, 213 Protein-carbohydrate complexes, 1, 7 Protein kinase A (PKA), 93 Protein kinase B (PKB, PKB/Akt), 98–100, 111, 132, 193, 194, 204 Protein kinase C (PKC), 94 Protein kinases, 88, 91–92, 98, 110, 125, 186, 198 Protein phosphatases, 92 Proteoglycans, 1, 4, 7–8, 10, 90 Proteolysis, 48, 52 Protofilaments, 24–25 Proto-oncogenes, 2–3, 109, 110, 133, 136, 198, 200, 201, 204, 205, 213 Protrusions, 18, 28, 42, 44, 57, 58, 60, 69, 70, 80, 82, 84, 101, 104, 121, 123–125 Pseudopodia, 10, 13, 24, 29, 37–52, 64, 66, 69, 71, 78, 101, 104, 105, 121–123, 125–130, 132–134, 151 Pseudopodial activity, 64, 69, 71, 72, 80, 122, 127–130, 133–134, 145, 151, 196 PTEN. See Phosphatase and tensin homolog (PTEN) protein/lipid phosphatase Pushing force, 123, 125, 129 R Rab. See Rab GTPases Rab GTPases, 131, 186 Rac1-activating protein (Asef), 103, 125–127, 129, 131, 133, 193, 195 Rac–1-specific exchange factor (Tiam1), 193 Rac GTPase, 101, 103, 129, 133, 193 Radial spreading, 64–69, 74–77, 154, 156 Raf. See Raf protein kinases Raf protein kinases, 98, 110 Rap1 GTPase, 48, 106, 192, 193, 202 Ras. See Ras GTPases Ras GTPases, 90, 92, 95–96, 98, 101, 110, 129, 133 R-cadherin. See Retinal cadherin (R-cadherin)
224 Receptors, 7–9, 18, 45, 47, 48, 52, 57, 88–91, 93–111, 121, 123, 128–132, 134–136, 177, 180, 185–187, 190–194, 198, 201–203, 214–215 Receptor tyrosine kinase (RTK), 89, 90, 96, 110, 203 Retinal cadherin (R-cadherin), 187 Retraction, 37, 40, 66, 128, 131, 132, 168, 170 Retrograde flow, 24, 125, 130 RGD sequence, 9, 45 Rho. See Rho GTPases Rho-associated kinase (ROCK). See Rho kinase Rho family of GTPases. See Rho GTPases Rho GTPases, 19, 24, 29, 44, 47, 69, 92, 93, 101, 105, 106, 125, 126, 129, 132, 133, 186, 193–196 Rho guanine nucleotide exchange factor (GEF-H1), 49, 126 Rho kinase, 102, 133–134 ROCK. See Rho kinase RTK. See Receptor tyrosine kinase (RTK) Ruffles, 37, 39, 42–44, 58–60, 62, 66–69, 78–81, 84, 94, 97, 125 S Scar. See Wiscott-Aldrich syndrome protein (WASP) family Scatter factor. See Hepatic growth factor Second messengers, 92–94 Semaphorins, 132 Septins, 28 Serine/threonine kinases, 48, 88, 92–96, 98, 99, 102, 103, 198 Signaling pathways, 48, 57, 88–108, 111, 112, 128, 130, 132–134, 177, 180, 185, 191, 193–195, 197–199, 201, 203–205, 213 Signal transduction, 2, 8, 57, 88–97, 99, 100, 106, 110–112, 130, 133, 134, 136, 185–187, 194, 199, 200, 213, 214 Small GTPases, 19, 47, 49, 69, 90, 92, 93, 95, 101, 106, 131, 132, 186, 192, 193, 195, 202 Smooth muscle actin (SMA), 106–107 SNAIL1. See Transcription repressor of CDH1 gene (SNAIL1) Spire, 16–17 Src protein kinase, 48, 95, 96, 109, 110, 125, 192, 193, 203 Stomata, 145, 148 Stress fibers, 45, 49, 50, 52, 71, 78, 80, 102, 105–109, 112, 129, 130, 133, 178, 179
Index Stretch force, 49, 106 Substratum, 2, 4, 42, 145, 151, 153, 155–162, 165, 167, 172, 177, 180 Substratum dependence, 2, 4, 100, 101, 108, 112 Surface relief, 57–63, 69, 70, 72, 80, 82–87 Surface topography, 153–154, 161, 172, 177, 180 Survival, 2, 4, 7, 10, 13, 48, 57, 88, 90, 92, 94, 99, 100, 112 Suspended state, 4, 42, 57–64, 69, 80, 84, 109, 213 Syncoilin. See Desmuslin Syndecans, 8, 90 Synemin. See Desmuslin T Talin, 47–48, 52, 105, 109 Taxol, 29, 126 T-cell factor/lymphocyte-enhancer factor (Tcf/Lef), 198, 199 Tcf/Lef. See T-cell factor/lymphocyteenhancer factor (Tcf/Lef) Tenascin, 9–10, 106, 177 Tensin, 47, 51, 95 Tension forces, 51, 128, 179, 180, 195 Tetraspanin-enriched microdomains, 131 Tetraspanins, 131, 135 TGF-a. See Transforming growth factor-a (TGF-a) Tiam1. See Rac–1-specific exchange factor (Tiam1) Tight junctions, 185–186, 189, 195 TIMP. See Tissue inhibitor of metalloproteinase (TIMP) +TIP. See Microtubule plus-end tracking proteins Tissue cells, 1–4, 7, 10, 37, 44, 45, 47, 57–112, 153, 177, 181, 184, 190, 200, 213 Tissue inhibitor of metalloproteinase (TIMP), 52 Topography, 153–181 Trafficking, 17, 99, 131, 191 Transcription factors, 93, 96, 110, 198 Transcription repressor of CDH1 gene (SNAIL1), 202–204 Transcription repressors, 202–204 Trans-dominant inhibition, 48 Transducers, 57, 100, 107, 108, 110 Transformation, 2–4, 57, 74, 78, 80, 109, 121, 132, 133, 153, 180, 181, 185, 200–205, 213
Index Transformed cells, 3, 4, 58, 60, 63, 74–88, 108–112, 132–137, 158–161, 163–167, 172, 180, 200, 213 Transforming growth factor-a (TGF-a), 136 Translocations, 3, 93, 121, 130, 198, 204, 205 Transport, 19, 27–29, 50, 84, 93, 127, 131, 134, 192 Tropomodulins, 17–19 Tropomyosin, 17–19 Tumor cells, 4, 52, 109, 111, 112, 132–137, 145, 148, 180, 185, 201–205, 213–214 Tumor growth, 4, 136, 205, 214, 215 Tumor suppressor genes, 2, 3, 28, 99, 100, 108, 111, 112, 134 Tyrosine kinases, 48, 89, 92, 95, 96, 109, 110, 192, 193, 203, 215 V Vascular-endothelial cadherin (VE-cadherin), 187 Vascular endothelial growth factor (VEGF), 96, 136, 215 VE-cadherin. See Vascular-endothelial cadherin (VE-cadherin) VEGF. See Vascular endothelial growth factor (VEGF) Villin, 17, 18 Vimentin, 29, 30, 106, 129 Vinculin, 18, 47, 49, 51, 52, 109, 189 Vitronectin, 48, 190 Void spaces, 167, 170, 172–175
225 W WAF1. See P21 protein WASP homolog-associated protein with actin, membranes and microtubules (WHAMM), 16 WASP-interacting protein (WIP), 16 WASP/Scar. See Wiscott-Aldrich syndrome protein (WASP) family WASP/WAVE. See Wiscott-Aldrich syndrome protein (WASP) family WAVE. See Wiscott-Aldrich syndrome protein (WASP) family WHAMM. See WASP homolog-associated protein with actin, membranes and microtubules (WHAMM) WIP. See WASP-interacting protein (WIP) Wiscott-Aldrich syndrome protein (WASP) family, 16, 21 Wnt1 protein, 200 Wnt signaling, 129, 197–200, 204, 205 X XMAP215. See Microtubule associated protein (XMAP215) Z ZO–1, ZO–2, ZO–3 proteins, 186, 189, 195 Zonula occludens (occluding junctions), 185 Zyxin, 18, 106
