Mechanosensitivity and Mechanotransduction (Mechanosensitivity in Cells and Tissues)

Mechanosensitivity and Mechanotransduction Mechanosensitivity in Cells and Tissues Volume 4 Series Editors A. Kamkin...

Author: Andre Kamkin | Irina Kiseleva

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Mechanosensitivity and Mechanotransduction

Mechanosensitivity in Cells and Tissues Volume 4

Series Editors A. Kamkin Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrovitjanova Str. 1, 117997 Moscow, Russia I. Kiseleva Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrovitjanova Str.1, 117997 Moscow, Russia

For further volumes: http://www.springer.com/series/7878

Andre Kamkin · Irina Kiseleva Editors

Mechanosensitivity and Mechanotransduction

Foreword by Holger Scholz

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Editors Prof. Andre Kamkin Department of Fundamental and Applied Physiology Russian State Medical University Ostrovitjanova Str. 1 117997 Moscow Russia [email protected]

Prof. Irina Kiseleva Department of Fundamental and Applied Physiology Russian State Medical University Ostrovitjanova Str. 1 117997 Moscow Russia [email protected]

Editorial Assistant Dr. Natalia E. Lapina Division of Neurosurgical Research Medical Faculty Mannheim Ruprecht-Karls-University Heidelberg Theodor-Kutzer-Ufer 1-3 D-68167 Mannheim Germany [email protected]

ISBN 978-90-481-9880-1 e-ISBN 978-90-481-9881-8 DOI 10.1007/978-90-481-9881-8 Springer Dordrecht Heidelberg London New York © Springer Science+Business Media B.V. 2011 No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Foreword

“Mechanosensitivity and Mechanotransduction” edited by André Kamkin and Irina Kiseleva describes and discusses the latest research findings in the field of cellular biomechanics. Internationally recognized experts contributed to this comprehensive and visionary work, which represents the accomplishment of a series of four related books. Mechanosensitivity describes the capacity of cells and tissues to perceive mechanical stimuli and translate them into biological signals. As such, mechanosensing and mechanotransduction are fundamental physiological mechanisms allowing cells to react to physical forces. Considering the importance and widespread distribution of mechanosensitive cells in many tissues of the body one can anticipate that a failure of normal mechanosensation may have severe consequences for various disease processes. The book is composed of seven coherent and well-arranged parts covering the multiple aspects of cellular mechanosensation. Part I highlights the role of the cytoskeleton in mechanotransduction. Special attention is given to the function of integrin receptors as mechanotransducers in the myogenic response of blood vessels. Importantly, signalling of integrins in response to enhanced mechanical forces has been implicated in vascular remodelling and arterial hypertension. Another focus is on the significance of the cytoskeleton for mechanosensing in the bone and articular cartilage. As underlined, the integrity of the cytoskeleton is essential for the modulation of gene expression in osteoblasts by mechanical stimuli. The topic is resumed in Chapters 12 and 13 discussing that mechanical loading of bone causes movements of interstitial fluid. This in turn generates fluid shear stress that stimulates anabolic activity in bone cells. Thus, a failure of normal mechanosensing by osteoblasts and chondrocytes may impair the ability of bone tissue and joint cartilage to adjust to the dynamics of biomechanical forces. Such maladaptation bears the risk of mechanical instability and onset of diseases of the skeletal system, i.e. osteoarthritis. The second part of this book discusses recent findings regarding the molecular control of mechanically gated ion channels (MGCs). This work includes the regulation of MGCs in the heart by nitric oxide and the signalling pathways involved in the myocardial response to mechanical stimulation. Substantial progress in the analysis of molecular MGC gating has recently been made with the use of GsMTx4, a gating-modifier peptide obtained from spider venom. v

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Mechanical stretch resulting from pulsatile blood flow can modulate vascular cell differentiation and function. Proteoglycans have a crucial role in mechanobiological responses of the arterial system. The issue of mechanosensing and mechanotransduction in vascular cells is discussed in Chapters 8 and 9. Emphasis is put on the importance of haemodynamic forces for the maintenance of homeostasis in the cardiovascular system. It is pointed out that a detailed knowledge of the molecular mechanisms underlying mechanosensing and mechanotransduction in vascular cells is essential to develop novel concepts for the prevention and therapy of cardiovascular diseases. Mechanotransduction in the lung, the topic highlighted in part IV, is established through the transient receptor potential vanilloid-4 (TRPV4) non-selective cation channel. Activation of TRPV4 initiates increased vascular permeability in response to high airway pressure or vascular pressure in the lung. Understanding the molecular regulation of this ion channel may therefore provide novel insights into various pulmonary disease processes including ventilator induced lung injury and pulmonary oedema. Primary cilia are organelles that have recently been recognized as mechanosensors. Cilia can sense the movement of body fluids in various tissues including endolymph in the inner ear, urine in renal tubules, bile in the hepatic biliary system, and digestive fluid in the pancreatic duct. Mechanosensing by primary cilia is discussed in Chapter 14. As outlined here genetic defects of the molecular components of primary cilia, i.e. polycystins and other interacting proteins, can give rise to various disorders such as polycystic kidney disease. The final chapter addresses the role of mechanosensing and mechanotransduction in blood cells. Blood cells encounter substantial mechanical stimuli due to variable haemodynamics. Large-conductance background K+ channels (LKbg ) have been identified as mechanosensors in mouse B lymphocytes and their molecular regulation by phospholipase C-dependent signalling pathways is discussed. In summary, “Mechanosensitivity and Mechanotransduction” compiles in-depth reviews of the current knowledge of mechanosensing and mechanical signal transduction in various tissues. The reader’s attention is attracted by the profound and clearly written contributions. Articles cover the many facets of cellular mechanosensation ranging from the molecular gating of mechanosensitive ion channels to perspectives for the future treatment of diseases. Institut für Vegetative Physiologie Charité - Universit¨stsmedizin Berlin Hessische Strasse 3-4 10115 Berlin

Holger Scholz

Editorial

Basic Principles of Mechanosensing and Mechanotransduction in Cells Andre Kamkin and Irina Kiseleva

Both mechanosensitivity and mechanotransduction are fundamental physiological processes which are responsible for sensing of mechanical forces and their transformation into electrical or (and) biochemical signals. The cell membrane interfaces with the external medium or with neighbouring cells and gets mechanical stress mainly in the form of stretch or compression due to general tissue deformation. Mechanical stimuli trigger different electrophysiological and biochemical responses. Mechanically gated ion channels (MGCs), reacting to the membrane tension, were shown to play the key role in one of the mechanisms, through which the cell responds to mechanical stimuli. Sachs and Morris (1998) noted that MGCs (originally called MSCs: mechanosensitive channels) are channels that recognize mechanical deformation as a proper physiological signal and react to mechanical stimulation with changes in kinetics. But MGCs are only a link (probably final from the point of view of changed ion transport and electrophysiological responses of the cell) in big chains of reception and transmission of mechanical stress. In living tissues, mechanical sensing and stress-induced responses in cells are defined by several cellular components including extracellular matrix, integrins, the cytoskeleton, and MGCs. The linkage between these cellular components undoubtedly plays a critical role. Mechanical forces are transmitted to cells, obviously, through the physical interactions of the cell with the surrounding extracellular matrix (Geiger et al., 2001; Matthews et al., 2006; Janmey and Weitz, 2004). The extracellular matrix is physically linked to the cytoskeleton via cell surface receptors primarily of the integrin family (Carver and Fuseler, 2010). It was shown, that alterations in the mechanical environment can be transmitted from the extracellular matrix to the cell via integrin receptors (Bershadsky et al., 2006; Sanchez-Esteban et al., 2006). Integrins are the main receptors for extracellular matrix proteins like collagen, fibronectin and laminin. Mechanical deformation, applied to extracellular matrix proteins, which are ligands for integrins, triggers the assembly and growth of focal contacts (Bershadsky et al., 2003) and activation of several downstream second messenger systems, including Rho GTPases, serine/threonine kinases, phosphatases, MAP kinases, Akt,

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and PKC (Lal et al., 2010). For example, in fibroblasts (Kumar et al., 2006; Zhao et al., 2007; Zhang X et al., 2007), endothelial (Mathur et al., 2007), or vascular smooth muscle cells (Na et al., 2007) stress-stiffening has been attributed to an integrin-Rho-dependent activation of stress fibers. Stretch-induced conformational changes in the ECM may alter integrin structure, resulting in activation of liganded integrin receptors and focal contact-associated secondary messenger pathways in the cell, such as FAK, Src family kinases, Abl and integrin-linked kinase – ILK (Li et al., 1999; Liu et al., 2000). Many studies have demonstrated that the physical network involving the extracellular matrix, integrins and the cytoskeleton is critical for the response of cells to mechanical forces (see for review Carver and Fuseler, 2010). Thus, during the deformation of the whole cell the possibility of the transmission of membrane forces into the cell is realized from the extracellular matrix through integrin receptors to the cytoskeleton and then to channel proteins (see for review Boriek and Kumar, 2008). It is beyond any doubt that integrins are major mechanosensors or mechanotransducers in many different cell types because of their unique ability to transduce extracellular mechanical signals exerted by extracellular matrix into intracellular signals. Integrins and cytoskeletal proteins act together as mechanosensors (see for review Thampatty and Wang, 2008). Despite of the achieved progress in addressing a number of questions, in many respects the mechanisms by which different cells sense, transduce and convert mechanical stimuli into cellular signals remain to be completely revealed. For this reason the first part of this volume is devoted to a network involving the extracellular matrix, integrins and the cytoskeleton. The first part of the volume is beginning with an article devoted to the role of integrin-mediated mechanotransduction by Yip et al. (2011). Vascular smooth muscle cells are used as a model for a discussion of the main issues concerned with integrins: Integrins and their associated kinases, integrins as mechanotransducers, mechanisms of coupling integrin activation to myogenic constriction. Depending on the types of integrin involved and the specific vascular beds, integrin dependent myogenic vasoconstriction can be mediated through Ca2+ dependent mechanisms of L-type Ca2+ channel potentiation, activation of global and local Ca2+ release from ryanodine sensitive Ca2+ stores; as well as Ca2+ independent mechanisms of FAK, c-Src, MAPK, and ILK activation and reactive oxygen species release (Yip et al., 2011). The second article is devoted to the role of the actin cytoskeleton in mechanosensation (Luo and Robinson, 2011). The actin cytoskeleton composed of actin filaments, myosin motors, and actin crosslinking proteins plays a critical role in force propagation and in response to deformation. The chapter discusses the microstructures and deformations of the actin cytoskeleton, functions of the actin cytoskeleton in mechanosensation. There are numerous overlaps between the mechanotransduction and “traditional” chemical signal transduction pathways. Because of the overlap between these pathways, mechanical-chemical coupling and feedback loops are a natural consequence of this system integration. Because the actin cytoskeleton is structurally integrated with nearly every aspect of the cell, mechanical inputs can be transmitted quickly throughout the cell. Challenges for understanding mechanosensation through the actin cytoskeleton include revealing how proteins

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function cooperatively over short nanometer-length-scales and fast sub-second time-scales. The following two articles discuss the role of the cytoskeleton in the mechanosensitivity of genes in osteoblasts (Fu et al., 2011) and the involvement of the cytoskeletal elements in articular cartilage mechanotransduction (Blain, 2011). The osteoblast is an important mechanosensitive cell in bone tissue (Fu et al., 2011). This article reviewed the roles of the cytoskeleton in the mechanotransduction of genes in osteoblasts, summarizes how cytoskeleton integrity is essential for the expression of bone formation–related genes in osteoblasts, and concludes that cytoskeleton reorganization inhibition can enhance the mechanosensitivity of some genes in osteoblasts. The article of Blain (2011) focuses on the organization and function of the three major cytoskeletal networks in articular cartilage chondrocytes. Articular cartilage is a major load-bearing tissue of the synovial joint; it is well known that the cytoskeleton acts as a physical interface between the chondrocytes and the extracellular matrix in “sensing” mechanical stimuli. We are yet far away from a clear understanding of how mechanical deformation of a cell can modulate MGCs. One model postulates that the mechanical energy is transferred though changes in lipid bilayer tension. However, the seal of a patch pipette isolates most of the stress in the patch from that in the cell, making a direct energy transfer through the lipid bilayer rather unlikely (Ursell et al., 2008). The second model postulates that mechanical sensing and stress-induced response in cells are defined by several cellular components including extracellular matrix, integrins, the cytoskeleton, and MGCs. The issues related to discussing this model are briefly mentioned above and are discussed in detail in the first Part of the Volume. The discussions presented in this Part of the Volume are devoted to the role, structure and function of the extracellular matrix itself. Extracellular matrix can be connected with certain structural qualities of integrins and here their interaction with extracellular matrix is discussed. The integrin receptor has a long N-ending extracellular transmembrane and very short S-ending cytopasmatic domains. Various combinations of α- and β-subunits define the specific properties of the binding of the extracellular receptor’s domain to certain ligands of the extracellular matrix. The majority of integrin receptors can bind several ligands. On the other hand one and the same ligand, e.g., laminin can bind to several integrins. Cellular integrin receptors bind to those glycoprotein parts of the extracellular matrix that contain the aminoacid sequence Arg-Gly-Asp. The integrin receptor’s intracellular domain binds with cytoskeleton microfilaments through a chain of various interconnected cytoplasmatic proteins. This is the modern vision of how the structural connection between the extracellular matrix and the cytoskeleton is formed. Interactions with extracellular matrix, cytoskeletal and intracellular signaling cascades enables integrins to mediate both “outside-in” and “inside-out” signaling (Hynes, 2002; Ross, 2004). Binding of integrins to extracellular ligands produces intracellular signals (outside-in signals) such as changes in intracellular signaling events and cytoskeletal reorganization that critically influences cell shape, migration, growth, and survival (Hynes, 2002). Inside-out signaling occurs when specific

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intracellular signals impinge on integrin cytoplasmic domains, triggering changes in conformation and ligand-binding affinity in the extracellular domain. Although the first Part is formally devoted to the interaction of extracellular matrix, integrins and the cytoskeleton, practically all parts of the Volume contain discussions of physical networks which are mobilized by mechanosensitivity and mechanotransduction in cells from different organs and tissues like the cardiovascular system (Kazanski et al., 2011; Guo et al., 2011; Shyu, 2011; Baker, 2011), the lung (Parker and Townsley, 2011; Rubacha and Liu, 2011), bone and joint tissues (Young and Pavalko, 2011; Sanchez et al., 2011), sensor systems (Nauli, 2011), and blood cells (Kim and Nam, 2011). The Second Part of the Volume is devoted to molecular mechanisms of mechanotransduction and ion channel modulation. This part of the volume begins with an article devoted to the role of nitric oxide in the regulation of mechanically gated channels in the heart (Kazanski et al., 2010a, b; Kazanski et al., 2011). The article discusses experimental data recorded from isolated ventricular myocytes of mouse, rat and guinea pig by means of patch-clamp in the whole-cell configuration about the role of NO in the regulation of MGCs. The presented data demonstrate that NO donors lead to MGCs activation and the appearance of MG-like currents in undeformed ventricular myocytes while in stretched cells with activated MGCs the NO donors lead to their inactivation and inhibition of the conductivity of these channels. The NO scavenger PTIO causes inactivation of all MGCs. Application of non selective inhibitors of NO syntases, L-NAME or L-NMMA, resulted in complete blockade of MGCs. In ventricular myocytes of wild-type mice, NOS1–/– and NOS2–/–, stretching of cells results in the activation of typical MG-currents. On the contrary, in cells from NOS3–/– mice stretch does not activate MG-currents. The results suggest that NO plays an important role in activation and inactivation of MGCs in cardiomyocytes and demonstrate that NOS3 dominate as the NO origin (Kazanski et al., 2010a, b; Kazanski et al., 2011). Different studies report that besides direct activation of MGCs by mechanical forces (via cytoskeleton or bilayer) fast indirect modulation of MGCs by pharmacological compounds is also possible. Moreover some of those compounds are capable of activating MGCs in the absence of actual cellular stretch, others of MGCs inactivation despite the continuous presence of cellular stretch. If this is true for compounds beside nitric oxide, this provides an excellent opportunity for the development of new drugs for treatment of mechano-induced arrhythmias. The role of signaling pathways in the myocardial response to biomechanical stress and in mechanotransduction in the heart is discussed in the review by Guo et al. (2011). The biochemical signals derived from mechanical stimuli activate both acute phosphorylation of signaling cascades, such as in the PI3K, FAK, and ILK pathways, and long-term morphological modifications via intracellular cytoskeletal reorganization and extracellular matrix remodelling. Mechanotransduction plays a fundamental role in cardiac (and vascular) function and involves interaction between extracellular matrix and intracellular cytoskeletal proteins via cell adhesion complexes, which are modulated by PI3Ks. Loss of PI3K signaling enhances the susceptibility to biomechanical stress while the loss of its

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negative regulator, PTEN, is associated with a wide variety of adaptive mechanisms necessary to resist the progression of maladaptive ventricular remodelling and heart failure. The third review in this part is “Atomistic molecular simulation of gating modifier venom peptides: Two binding modes and effects of lipid structure” and is devoted to GsMTx4 (Nishizawa, 2011). This drug is a gating-modifier peptide obtained from tarantula venom. It is a valuable tool for the investigation of gating mechanisms of mechanosensitive channels. Free energy profile analyses suggests that these toxins exhibit two modes of binding to lipid membranes, namely, the shallow mode and the deep mode. These toxins favor the deep mode, especially in membranes rich in saturated lipid acyl chains, which make the headgroup layer tight. It is hypothesized that in the case of HaTx the deep mode is the action mode, while for GsMTx4 the two modes can explain the concentration-dependent (biphasic) effect of GsMTx4 that has recently been reported. The possibility that such toxins seek out specific types of lipid molecules is discussed. Simulation results support the view that the channel/GsMTx4 (or HaTx)/lipids make a tertiary complex crucial to the effectiveness of the toxin and therefore binding of the toxin to channels occurs only in the presence of lipid molecules with appropriate structures (Nishizawa, 2011). The following parts of the Volume are devoted to the mechanosensitivity and mechanotransduction in vascular cells (Part III), in the lung (Part IV), in bone and joint tissues (Part V), in sensor systems (Part VI) and in blood cells (Part VII). Part III begins with a review devoted to molecular effects of mechanical stretch on vascular cells (Shyu, 2010). The vascular endothelium plays an important role by sensing alterations in biological, chemical, and physical properties of blood flow to maintain homeostasis. Mechanical stretch can modulate cell alignment and differentiation, migration, survival or apoptosis, vascular remodeling, and autocrine and paracrine functions in smooth muscle cells. Laminar shear stress exerts antiapoptotic, anti-atherosclerotic, and anti-thrombotic effects on endothelial cells. Knowledge of the impact of mechanical stretch on the cardiovascular system is vital to the understanding of the pathogenesis of cardiovascular diseases and is also crucial to provide new insights in the prevention and therapy of cardiovascular diseases. The role of proteoglycans in vascular mechanotransduction is discussed in the next chapter (Baker, 2011). This review focuses on the role of proteoglycans in vascular mechanobiological responses of the arterial system. Proteoglycans are proteins that are post-translationally modified with polysaccharide glycosaminoglycan chains. These molecules are intimately involved in controlling cellular organization, proliferation and migration. Several studies have demonstrated that the expression of syndecan-1 and syndecan-4 are increased in mechanically stimulated cells. The syndecans are good candidates for mechanotransduction pathways as they have multiple interactions with integrins, focal adhesion, cytoskeletal elements and growth factor signaling. While the last decade has yielded many new insights into the basic biology of proteoglycans, future studies will hopefully be able to shed additional light on the roles these molecules play in mechanotransduction and vascular remodeling.

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Two chapters are devoted to the mechanosensitivity and mechanotransduction in the lung. The transient receptor potential vanilloid-4 (TRPV4) non-selective cation channel and its effect on the lung is discussed in the first review of Part IV (Parker and Townsley, 2011). Increased pulmonary venous pressure and ventilation with high peak inflation pressures increase endothelial calcium influx, nitric oxide production, and vascular permeability in a TRPV4 dependent fashion in intact lungs. Calcium influx occurs with relatively modest increases in pulmonary vascular or airway pressures, and the channel is essential for initiating the increased vascular permeability induced by high vascular and airway pressures. Mechanical gating is mediated by epoxyeicosatrienoic acids, whereas heat and phorbol ester activation occur through different mechanisms. TRPV4 is a major mediator of injury to pulmonary microvascular endothelium, which lacks the store operated TRPC channels of macrovascular endothelium. Several potential regulatory sites on the TRPV4 molecule have been identified for phosphorylation by Src family kinases, PKA, and PKC. However, the mechanisms for epoxyeicosatrienoic acid dependent regulation of TRPV4 or amplification of TRPV4 by phosphorylation in intact lungs subjected to mechanical stress have not been clarified (Parker and Townsley, 2011). The second review of this part deals with the role of protein-protein interactions in mechanotransduction, and with particular implications in ventilator induced lung injury (Rubacha and Liu, 2011). Under discussion are the issues of basic lung mechanics vs. mechanical ventilation, ventilator induced inflammatory response, protein-protein interactions, unfolding of p130Cas as a mechanosensor, AFAP as an activator of Src PTK, Src PKT activation by multiple physical forces, blocking Src PTK as a potential therapy for VILI. VILI is a complex problem that will likely require a multifaceted approach to solve including work from basic scientists, clinicians, and even engineers. In this chapter the authors discuss this novel mechanism for mechanosensation and mechanotransduction, and propose to inhibit Src protein tyrosine kinase activation as a potential therapy for ventilator induced lung injury. The next Part of the Volume is devoted to bone and joint tissues. Two chapters presented in this Part describe cellular and molecular mechanisms of mechanosensitivity and mechanotransduction. The cells within bone, i.e. osteocytes and osteoblasts, are responsible for detecting and responding to mechanical loading. The first review discusses several of the key mechanisms used by bone cells to convert mechanical signals into altered biochemical responses (Young and Pavalko, 2011). Understanding how bone cells sense and respond to mechanical signals has been the focus of considerable research. The review deals with a number of important topics. It includes detection of mechanical stimuli and in this context focal adhesions with the mechanosome hypothesis, ion channel and purinergic signaling in bone. The propagation of mechanical signals in bone cells is discussed in detail. In this context are considered focal adhesion kinase (FAK), Wnt/β-catenin/sclerostin, Gap junctions, NFAT, Nitric oxide cGMP-dependent kinases, and Nmp4/CIZ (Young and Pavalko, 2011). Recent research has resulted in significant progress toward the identification of the key molecular components used by cells to detect mechanical stimuli and to propagate those signals through the cytoplasm and into the nucleus.

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The second chapter also demonstrates that joint tissues, including cartilage, bone, meniscus, tendon, ligament and synovial membrane, are exposed to high levels of mechanical stimulation. This review is devoted to the mechanosensitivity of cells in joint tissues. Under discussion is the role of the mechanosensitivity in the pathogenesis of joint diseases (Sanchez et al., 2011). The chapter reviews the issues of mechanical stimuli and cartilage matrix remodeling and chondrocyte mechanotransduction. The discussion covers mechanical stimuli and subchondral bone and other joint tissues mechanosensitivity. The following Part presents the review devoted to primary cilia as mechanosensory organelles in vestibular tissues (Nauli, 2011). In this chapter the author describes cilia as newly recognized mechanosensory organelles. The author discusses ciliary classification, function and disease relevance in mammals. Besides, structural and functional ciliary proteins based on their ciliary domains are presented. The importance of sensory cilia in other organ systems is yet to be discovered, and many more cilia-related diseases are still to be identified. The final Part of the Volume presents the review – Mechanosensitive K+ channels in mouse B lymphocytes: PLC-mediated release of TREK-2 from Inhibition by PIP2 (Kim and Nam, 2011). There is a limited amount of data regarding mechanosensitivity and mechanotransduction in blood cells, therefore this chapter is of special interest to our readers. Blood cells can encounter significant mechanical stimuli in variable flow. In mouse B lymphocytes and their cell line WEHI-231, the authors found large-conductance background K+ channels (LKbg ) that show significant mechanosensitivity. The biophysical characteristics of LKbg were similar to those of TREK-2. The activity of TREK-2 is under tonic inhibition by PIP2 , and a relatively mild membrane stretch relieves this inhibition via activation of PLC hydrolysing PIP2 . Apart from the mechano-biochemical signaling mechanism, more direct regulation by stronger membrane stretch is also suggested. Thus, this book is a unique collection of reviews outlining current knowledge and future developments in this rapidly growing field. Currently, investigations of the molecular mechanisms of mechanosensitivity and mechanotransduction are focused on several issues. The majority of studies investigate intracellular signaling pathways. Knowledge of the mechanisms which underlie these processes is necessary for the understanding of the normal functioning of different living organs and tissues and allows to predict changes, which arise due to alterations of their environment, and possibly will allow to develop new methods of artificial intervention. The book brings up the problem closer to the experts in related medical and biological sciences as well as practicing doctors besides just presenting the latest achievements in the field.

References Baker AB (2011) Role of proteoglycans in vascular mechanotransduction. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 219–236

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Bershadsky A, Kozlov M, Geiger B (2006) Adhesion-mediated mechanosensitivity: a time to experiment, and a time to theorize. Curr Opin Cell Biol 18:472–481 Bershadsky AD, Balaban NQ, Geiger B (2003) Adhesion-dependent cell mechanosensitivity. Annu Rev Cell Dev Biol 19:677–695 Blain EJ (2011) Involvement of the cytoskeletal elements in articular cartilage mechanotransduction. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 77–106 Boriek AM, Kumar A (2008) Regulation of intracellular signal transduction pathways by mechanosensitive ion channels. In: Kamkin A, Kiseleva I (eds) mechanosensitivity in cells and tissues 1. Mechanosensitive ion channels. Springer, pp 303–327 Carver W, Fuseler JW (2010) Mechanical stretch-induced reorganization of the cytoskeleton and the small GTPase Rac-1 in cardiac fibroblasts. In: Kamkin A, Kiseleva I (eds.) Mechanosensitivity in cells and tissues 3. Mechanosensitivity of the heart. Springer, pp 35–54 Fu Q, Zhang Y, Xu Y, Li Y, Guo L, Shao M (2011) Effect of cytoskeleton on the mechanosensitivity of genes in osteoblasts. In: Kamkin A, Kiseleva I (eds.) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 67–76 Geiger B, Bershadsky A, Pankov R, Yamada KM (2001) Transmembrane crosstalk between the extracellular matrix- cytoskeleton crosstalk. Nat Rev Mol Cell Biol 2:793–805 Guo D, Kassiri Z, Oudit GY (2011) Role of signaling pathways in the myocardial response to biomechanical stress and in mechanotransduction in the heart. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 141–166 Hynes RO (2002) Integrins: bidirectional, allosteric signaling machines. Cell 110:673–687 Janmey PA and Weitz DA (2004) Dealing with mechanics: mechanisms of force transduction in cells. Trends Biochem Sci 29:364–370 Kazanski VE, Kamkin A, Makarenko EYu, Lysenko NN, Sutiagin PV, Tian B, Kiseleva I (2010a) The role of the Nitric Oxide in regulation of mechanically gated channels activity in cardiomyocytes: Investigation by means of the application of NO-donors. Bulletin of Experimental Biology and Medicine 7:4–9. English, Russian. (See PubMed for details of English version pages) Kazanski VE, Kamkin A, Makarenko EYu, Lysenko NN, Sutiagin PV, Kiseleva I (2010b) The role of the Nitric Oxide in regulation of mechanically gated channels activity in cardiomyocytes: Investigation of NO-synthatases contribution. Bulletin of Experimental Biology and Medicine 8:228–232. English, Russian. (See PubMed for details of English version pages) Kazanski V, Kamkin A, Makarenko E, Lysenko N, Lapina N, Kiseleva I (2011) The role of nitric oxide in regulation of mechanically gated channels in the heart. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 109–140 Kim SJ, Nam JH (2011) Mechanosensitive K+ channels in mouse B lymphocytes; PLC-mediated release of TREK-2 from the membrane-delimited inhibition by PIP2 . In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 353–368 Kumar S, Maxwell IZ, Heisterkamp A, Polte TR, Lele TP, Salanga M, Mazur E, Ingber DE, (2006) Viscoelastic retraction of single living stress fibers and its impact on cell shape, cytoskeletal organization, and extracellular matrix mechanics. Biophys J 90:3762–3773 Lal H, Verma SK, Golden HB, Foster DM, Holt AM, Dostal DE (2010) Molecular signaling mechanisms of myocardial stretch: implications for heart disease. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 3. Mechanosensitivity of the heart. Springer, pp 55–81

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Li F, Zhang Y, Wu C (1999) Integrin-linked kinase is localized to cell-matrix focal adhesions but not cell-cell adhesion sites and the focal adhesion localization of integrin-linked kinase is regulated by the PINCH-binding ANK repeats. J Cell Sci 112 (Pt 24):4589–4599 Liu S, Calderwood DA, Ginsberg MH (2000) Integrin cytoplasmic domain-binding proteins. J Cell Sci 113( Pt 20):3563–3571 Luo T, Robinson DN (2011) The role of actin cytoskeleton in mechanosensation. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 25–65 Mathur AB, Reichert WM, Truskey GA (2007) Flow and high affinity binding affect the elastic modulus of the nucleus, cell body and the stress fibers of endothelial cells. Ann Biomed Eng 35:1120–1130 Matthews BD, Overby DR, Mannix R, Ingber DE (2006) Cellular adaptation to mechanical stress: role of integrins, Rho, cytoskeletal tension and mechanosensitive ion channels. J Cell Sci 119:508–518 Na S, Meininger GA, Humphrey JD (2007) A theoretical model for F-actin remodeling in vascular smooth muscle cells subjected to cyclic stretch. J Theor Biol 246:87–99 Nauli SM, Haymour HS, Aboualaiwi WA, Lo ST, Nauli AM (2011) Primary Cilia are Mechanosensory Organelles in Vestibular Tissues. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 317–350 Nishizawa K (2011) Atomistic molecular simulation of gating modifier venom peptides – two binding modes and effects of lipid structure. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 167–190 Parker JC, Townsley MI (2011) Control of TRPV4 and its effect on the lung. In: Mechanosensitivity in Cells and Tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 239–254 Ross RS (2004) Molecular and mechanical synergy: cross-talk between integrins and growth factor receptors. Cardiovasc Res 63:381–390 Rubacha M, Liu M (2011) The role of protein-protein interactions in mechanotransduction: Implications in ventilator induced lung injury. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 255–273 Sachs F, Morris CE (1998) Mechanosensitive ion channels in nonspecialized cells. Rev Physiol Biochem Pharmacol 132:1–77 Sanchez Ch, Mathy-Hartert M, Henrotin Y (2011) The mechanosensitivity of cells in joint tissues: Role in the pathogenesis of joint diseases. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 297–313 Sanchez-Esteban J, Wang Y, Filardo EJ, Rubin LP, Ingber DE (2006) Integrins beta1, alpha6, and alpha3 contribute to mechanical strain-induced differentiation of fetal lung type II epithelial cells via distinct mechanisms. Am J Physiol Lung Cell Mol Physiol 290:L343–L350 Shyu K-G (2011) Cellular and molecular effects of mechanical stretch on vascular cells. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 193–217 Thampatty BP, Wang JH-C (2008) Mechanobiology of fibroblasts. In: Kamkin A, Kiseleva I (eds) mechanosensitivity in cells and tissues 1. Mechanosensitive ion channels. Springer, pp 351–378 Ursell T, Kondev J, Reeves D, Wiggins PA, Phillips R (2008) Role of lipid bilayer mechanics in mechanosensation. In: Kamkin A, Kiseleva I (eds) mechanosensitivity in cells and tissues 1. Mechanosensitive ion channels. Springer, pp 37–70 Yip K-P, Balasubramanian L, Sham JSK (2011) Integrin-mediated mechanotransduction in vascular smooth muscle cells. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 3–24

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Young SRL and Pavalko FM (2011) Cellular mechanisms of mechanotransduction in bone. In: Kamkin A, Kiseleva I (eds) Mechanosensitivity in cells and tissues 4. Mechanosensitivity and mechanotransduction. Springer, pp 277–296 Zhang X, Stewart JA Jr, Kane ID, Massey EP, Cashatt DO, Carver WE (2007) Effects of elevated glucose levels on interactions of cardiac fibroblasts with the extracellular matrix in vitro. Cell Dev Biol Anim 43:297–305 Zhao XH, Laschinger C, Arora P, Szarszi K, Kapus A, McCulloch CA (2007) Force activates smooth muscle alpha-actin promoter activity through the Rho signaling pathway. Cell Sci 120:1801–1809

Contents

Foreword by Holger Scholz . . . . . . . . . . . . . . . . . . . . . . . . .

v

Editorial . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andre Kamkin and Irina Kiseleva

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Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Part I

The Role of Cytoskeleton in Mechanosensitivity and Mechanotransduction

1 Integrin-Mediated Mechanotransduction in Vascular Smooth Muscle Cells . . . . . . . . . . . . . . . . . . . . . . . . . . Kay-Pong Yip, Lavanya Balasubramanian, and James S.K. Sham 2 The Role of the Actin Cytoskeleton in Mechanosensation . . . . . . Tianzhi Luo and Douglas N. Robinson 3 Effect of Cytoskeleton on the Mechanosensitivity of Genes in Osteoblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Qiang Fu, Yiping Zhang, Yajuan Xu, Yourui Li, Ling Guo, and Minfeng Shao 4 Involvement of the Cytoskeletal Elements in Articular Cartilage Mechanotransduction . . . . . . . . . . . . . . . . . . . . Emma J. Blain Part II

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25

67

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Molecular Mechanisms of Mechanotransduction and Ion Channels Modulation

5 The Role of Nitric Oxide in the Regulation of Mechanically Gated Channels in the Heart . . . . . . . . . . . . . . . . . . . . . Victor Kazanski, Andre Kamkin, Ekaterina Makarenko, Natalia Lysenko, Natalia Lapina, and Irina Kiseleva

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Contents

6 Role of Signaling Pathways in the Myocardial Response to Biomechanical Stress and in Mechanotransduction in the Heart . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Danny Guo, Zamaneh Kassiri, and Gavin Y. Oudit 7 Atomistic Molecular Simulation of Gating Modifier Venom Peptides – Two Binding Modes and Effects of Lipid Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kazuhisa Nishizawa Part III

9 Role of Proteoglycans in Vascular Mechanotransduction . . . . . . Aaron B. Baker

Control of TRPV4 and Its Effect on the Lung . . . . . . . . . . . . James C. Parker and Mary I. Townsley

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The Role of Protein-protein Interactions in Mechanotransduction: Implications in Ventilator Induced Lung Injury . . . . . . . . . . . . . . . . . . . . . . . . . . Matthew Rubacha and Mingyao Liu

Cellular Mechanisms of Mechanotransduction in Bone . . . . . . . Suzanne R.L. Young and Fredrick M. Pavalko

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The Mechanosensitivity of Cells in Joint Tissues: Role in the Pathogenesis of Joint Diseases . . . . . . . . . . . . . . . . . Christelle Sanchez, Marianne Mathy-Hartert, and Yves Henrotin

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Mechanosensing and Mechanotransduction in Bone and Joint Tissues

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Part VI

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Mechanotransduction in the Lung

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Part V

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Mechanosensitivity and Mechanotransduction in Vascular Cells

8 Cellular and Molecular Effects of Mechanical Stretch on Vascular Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kou-Gi Shyu

Part IV

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Mechanotransduction of Sensor System

Primary Cilia are Mechanosensory Organelles in Vestibular Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Surya M. Nauli, Hanan S. Haymour, Wissam A. Aboualaiwi, Shao T. Lo, and Andromeda M. Nauli

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Contents

Part VII

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Mechanosensitivity and Mechanotransduction in Blood Cells

Mechanosensitive K+ Channels in Mouse B Lymphocytes: PLC-Mediated Release of TREK-2 from Inhibition by PIP2 . . . . Sung Joon Kim and Joo Hyun Nam

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

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Contributors

Wissam A. Aboualaiwi Department of Pharmacology, MS 1015; The University of Toledo; Health Science Campus, HEB 274; 3000 Arlington Ave., Toledo, OH 43614, USA Aaron B. Baker Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, 77 Massachusetts Avenue, E25-442, Cambridge, MA 02139, USA, [email protected] Lavanya Balasubramanian Department of Molecular Pharmacology and Physiology, University of South Florida, Tampa, FL 33612, USA Emma J. Blain Connective Tissue Biology Laboratories, Biomedical Sciences Building, School of Biosciences, Cardiff University, Museum Avenue, Cardiff, CF10 3AX, UK, [email protected] Qiang Fu Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China, [email protected]; [email protected] Ling Guo Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China Danny Guo Division of Cardiology, Department of Medicine, Mazankowski Alberta Heart Institute, University of Alberta, Edmonton, AB, T6G 2S2, Canada Hanan S. Haymour Department of Pharmacology, MS 1015; The University of Toledo; Health Science Campus, HEB 274; 3000 Arlington Ave., Toledo, OH 43614, USA Yves Henrotin Bone and Cartilage Research Unit, University of Liège, Institute of pathology level 5, CHU Sart-Tilman, 4000, Liège, Belgium, [email protected] Andre Kamkin Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia, [email protected]

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Zamaneh Kassiri Mazanokowski Alberta Heart Institute, and Department of Physiology, University of Alberta, Edmonton, AB T6G 2S2, Canada Victor Kazanski Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia Sung Joon Kim Department of Physiology, Ischemic/Hypoxic Disease Institute, Kidney Research Institute, Seoul National University College of Medicine, 103 Daehangno, Jongno-gu, Seoul 110-799, Korea, [email protected]; [email protected] Irina Kiseleva Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia, [email protected] Natalia Lapina Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia; Division of Neurosurgical Research Medical Faculty Mannheim, Ruprecht-Karls-University Heidelberg, Theodor-Kutzer-Ufer 1-3, D-68167, Mannheim, Germany, [email protected]; [email protected] Yourui Li Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China Mingyao Liu Faculty of Medicine, University of Toronto, 101 College Street Toronto Medical Discovery Tower, Room 2-814, Toronto, ON M5G 1L7, Canada, [email protected] Shao T. Lo Department of Pharmacology, MS 1015; The University of Toledo; Health Science Campus, HEB 274; 3000 Arlington Ave., Toledo, OH 43614, USA Tianzhi Luo Department of Cell Biology and Department of Pharmacology and Molecular Science, Johns Hopkins University School of Medicine, 725 N. Wolfe Street, Baltimore, MD 21205, USA Natalia Lysenko Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia, [email protected] Ekaterina Makarenko Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia, [email protected] Marianne Mathy-Hartert Bone and Cartilage Research Unit, University of Liège, Institute of pathology level 5, CHU Sart-Tilman, 4000 Liège, Belgium Joo Hyun Nam Department of Pharmacology and Research Center for Human Natural Defense System, College of Medicine, Yonsei University, Seoul, Korea and Department of Physiology, Dongguk University College of Medicine, Gyeongju, 780-714, Korea

Contributors

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Andromeda M. Nauli Department of Health Sciences; East Tennessee State University; College of Public Health; Johnson City, TN 37614, USA Surya M. Nauli Department of Pharmacology, MS 1015; The University of Toledo; Health Science Campus, HEB 274; 3000 Arlington Ave., Toledo, OH 43614, USA, [email protected] Kazuhisa Nishizawa Department of Laboratory Medicine, Teikyo University School of Medical Technology, Kaga, Itabashi, Tokyo 173-8605, Japan, [email protected] Gavin Y. Oudit Division of Cardiology, Department of Medicine, and Mazankowski Alberta Heart Institute, University of Alberta, Edmonton, AB, T6G 2S2, Canada, [email protected] James C. Parker Department of Physiology and Center for Lung Biology, MSB 3074, University of South Alabama, Mobile, AL, 36688, USA, [email protected] Fredrick M. Pavalko Department of Cellular and Integrative Physiology, Indiana University School of Medicine, 635 Barnhill Drive, MS 346A, Indianapolis, IN 46202, USA, [email protected] Douglas N. Robinson Department of Cell Biology and Department of Pharmacology and Molecular Science, Johns Hopkins University School of Medicine, 725 N. Wolfe Street, Baltimore, MD 21205, USA; Department of Chemical and Biomolecular Engineering, Johns Hopkins University Whiting School of Engineering, 3400 N. Charles St, Baltimore, MD 21218, USA, [email protected] Matthew Rubacha Faculty of Medicine, University of Toronto, 101 College Street Toronto Medical Discovery Tower, Room 2-814, Toronto, ON, M5G 1L7, Canada, [email protected] Christelle Sanchez Bone and Cartilage Research Unit, University of Liège, Institute of pathology level 5, CHU Sart-Tilman, 4000, Liège, Belgium Holger Scholz Institut für Vegetative Physiologie, Charité – Universitätsmedizin Berlin, Hessische Strasse 3-4, 10115, Berlin, Germany, [email protected] James S. K. Sham Division of Pulmonary and Critical Care Medicine, Johns Hopkins Medical Institutions, 5501 Hopkins Bayview Circle, Baltimore, MD 21224, USA, [email protected] Minfeng Shao Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China Kou-Gi Shyu Division of Cardiology, Shin Kong Wu Ho-Su Memorial Hospital, Taipei, Taiwan; Graduate Institute of Clinical Medicine, College of Medicine, Taipei Medical University, Taipei, Taiwan, [email protected]

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Contributors

Mary I. Townsley Department of Physiology and Center for Lung Biology, MSB 3074, University of South Alabama, Mobile, AL, 36688, USA, [email protected] Yajuan Xu Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China Kay-Pong Yip Department of Molecular Pharmacology and Physiology, University of South Florida, Tampa, FL 33612, USA, [email protected] Suzanne R.L. Young Department of Cellular and Integrative Physiology, Indiana University School of Medicine, 635 Barnhill Drive, MS 346A, Indianapolis, IN 46202, USA Yiping Zhang Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, PR China

Part I

The Role of Cytoskeleton in Mechanosensitivity and Mechanotransduction

Chapter 1

Integrin-Mediated Mechanotransduction in Vascular Smooth Muscle Cells Kay-Pong Yip, Lavanya Balasubramanian, and James S.K. Sham

Abstract Myogenic response is an intrinsic property of vascular smooth muscle cells (VSMCs) by which VSMCs contract when transmural pressure is increased, and dilates when transmural pressure is decreased. Myogenic response in resistance arterioles is one primary mechanisms for blood flow autoregulation. There is emerging evidence that the dynamic interactions between specific extracellular matrix (ECM) and integrins are required for mechanotransduction in myogenic response. Studies in VSMCs and microvessels show that myogenic response is linked to Ca2+ influx as well as local and global Ca2+ release from ryanodine-gated stores. Electrophysiological evidences suggest that Ca2+ -activated channels might provide the link between activation of integrins and membrane potential modulation in myogenic response. Aside from myogenic vasoconstriction, the elevated mechanical stress imposed on the integrin signaling complex might play important roles in vascular remodeling and hypertension. Keywords Myogenic response · Calcium spark channels · Integrin-linked kinase · Vascular remodeling

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Calcium-activated

1.1 Introduction Mammalian blood pressure has been shown to spontaneously fluctuate up to 40% of the mean arterial blood pressure (Marsh et al., 1990). Blood vessels are therefore constantly receiving mechanical stimuli from the ebb and flow of blood through these conduits. A steady blood flow is vital for the physiological functions in many organs, like the glomerular filtration rate in kidney and cerebral blood flow. One of K.-P. Yip (B) Department of Molecular Pharmacology and Physiology, University of South Florida, Tampa, FL 33612, USA e-mail: [email protected]

A. Kamkin, I. Kiseleva (eds.), Mechanosensitivity and Mechanotransduction, Mechanosensitivity in Cells and Tissues 4, DOI 10.1007/978-90-481-9881-8_1, C Springer Science+Business Media B.V. 2011

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the major mechanisms preventing the conversion of pressure fluctuations into fluctuations in blood flow is the myogenic response of vascular smooth muscle cells (VSMCs). Myogenic response is an intrinsic mechanism of VSMCs by which the vasculature constricts on elevation and dilates on reduction of perfusion pressure (Bayliss, 1902). It is necessary for maintaining constant blood flow and capillary hydrostatic pressure (Davis and Hill, 1999). End organ damage in hypertension has been attributed to the transmission of increased blood pressure into the kidney (Bidani et al., 1987; Karlsen et al., 1997; Abu-Amarah et al., 2005). Impaired myogenic response in fawn-hooded hypertensive rats showed glomerular damage (Simons et al., 1993; van Dokkum et al., 1999) highlighting the importance of normal myogenic response in maintaining a constant GFR and hence renal integrity. However, the mechanisms of how VSMCs sense the changes in the perfusion pressure are not well defined (Davis and Hill, 1999). Mechanotransduction of VSMCs is not only involved in acute adaptation of vascular resistance in myogenic response, but also in chronic adaptation such as vascular remodeling in hypertension. Vascular remodeling is closely related the VSMCs proliferation and extracellular matrix (ECM) accumulation. Genetic hypertension and angiotensin II induced hypertension are associated with changes in integrins and ECM expression in resistance arterioles (Intengan et al., 1999; Intengan and Schiffrin, 2000; Mori and Cowley, 2004; Seubert et al., 2005).

1.2 Potential Candidates for the Sensor of Mechanotransduction In order to alter the vascular resistance in response to changes in perfusion pressure, mechanisms for sensing vascular wall stress/tension should be present (Johnson, 1980; Davis and Hill, 1999). Such mechanosensors should be conveniently seated on the plasma membrane to transduce the external force into the intracellular milieu. Their proximity to other signaling molecules like kinases and local Ca2+ stores is essential to cause an immediate local response. The potential candidates for mechanosensing fall into three major categories. They include ion channels which modulate membrane potential; transmembrane proteins which link the ECM to the cytoskeleton; and kinases which phosphorylate proteins involved in contractility and proliferation.

1.2.1 Stretch-Sensitive Ion Channels It is believed that the non-selective stretch-sensitive cation channels (NSCCs) act as the sensors in mechanotransduction. An increase in transmural pressure stretches the vascular wall and opens the stretch-sensitive channels, which initiates contraction by depolarizing the VSMCs leading to the activation of voltage-gated Ca2+ channels (VGCCs) and hence an increase in the intracellular calcium concentration ([Ca2+ ]i ) (Kirber et al., 1988; Davis et al., 1992; Meininger and Davis, 1992). Stretch-activated whole cells currents have been recorded in many types of VSMCs

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(Wellner and Isenberg, 1994; Wellner and Isenberg, 1995). The stretch-induced depolarization could be explained by the activation of mechanosensitive ion channels promoting Na+ or Ca2+ influx, Cl– efflux, or inhibiting K+ efflux (Davis and Hill, 1999). Ion transport pumps in the plasma membrane have also been suggested as a plausible sensor. Stretching the VSMCs showed an increase in the expression and translocation of the α-subunit of the Na+ , K+ ATPase to the plasma membrane (Sevieux et al., 2003). Another group of NSCCs that has been implicated in myogenic response is the transient receptor potential (TRP) superfamily of channels (Inoue et al., 2006). Most TRPC channels can be activated by PLC pathway, which can likely be triggered by the increase in intravascular pressure. In frog oocytes, TRPC1 forms stretch sensitive cation channel (Maroto et al., 2005), and TRPC6 has been shown to regulate myogenic response by their involvement in pressure-induced membrane depolarization in rat cerebral arteries (Welsh et al., 2002; Spassova et al., 2006). Increasing evidence suggests that TRPM4 channels play a major role in myogenic response. Suppression of TRPM4 expression using antisense oligodeoxynucleotide decreased pressure-induced depolarization and myogenic response in vitro, and altered autoregulation in vivo in rat cerebral arteries (Earley et al., 2004; Reading and Brayden, 2007). Another study proposed that TRPM4 may actually affect myogenic response by stretch-induced Ca2+ release via ryanodine receptors (Morita et al., 2007). Besides cerebral arteries, TRPC3 and TRPC6 are highly expressed in rat renal resistance vessels (Facemire et al., 2004; Inoue et al., 2006); TRPC1, TRPC6, TRPM4, TRPM8, TRPV2, and TRPV4 are also abundant in pulmonary arterial smooth muscle cells (Lin et al., 2004; Yang et al., 2006). However, their roles in myogenic response in these vascular beds are unclear. Moreover, TRPV1 and TRPV2 have been implicated as mechanosensitive channels in aortic smooth muscle cells; but their involvement in myogenic response is debatable (Roman, 2002; Muraki et al., 2003). It is noteworthy that even though stretch-sensitive membrane bound channels may operate as mechanosensors accounting for the initiation of myogenic vasoconstriction, they could not provide an error signal to sustain myogenic constriction because of the abrogation of stretch after myogenic constriction.

1.2.2 Integrins and Their Associated Kinases The other plausible sensors for the myogenic phenomenon are integrins, the heterodimeric receptors of ECM. They are considered as stress sensing and transducing elements in many cell types. Of the 24 known integrins, 16 have been reported in various aspects in vascular biology. Among them, α1 β1 , α2 β1 , α3 β1 , α4 β1 , α5 β1 , α6 β1 , α7 β1 , α8 β1 , α9 β1 , αv β1 , αv β3 , αv β5 , and α6 β4 are expressed in VSMCs (Glukhova et al., 1994; Moiseeva, 2001). They connect the ECM to the internal cytoskeleton (CSK) via the short cytoplasmic tail of the β-subunit and transduce both “outside-in” (modification of cellular events on integrin ligation) and “inside-out” signals (modification of the ECM in response to integrin ligation). An

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increase in perfusion pressure might provide the mechanical signal to activate the VSMC integrins via attachment between native ECM and integrin during passive dilation. Stretch of ECM may induce conformational changes in integrin resulting in the activation of focal adhesion associated secondary messenger pathways, such as focal adhesion kinase (FAK), Src family kinases, and integrin-linked kinase (ILK) (Lal et al., 2009). FAK is a cytosolic soluble tyrosine kinase, which plays a major role in integrinmediated signaling. It consists of an N-terminal functional domain that interacts with β-integrin and growth factor receptors, a catalytic domain for tyrosine kinase activity, and a focal-adhesion targeting domain that localizes FAK to focal adhesions and binding sites for associated signaling molecules (Schaller, 2001). Integrins induce auto-phosphorylation of FAK Tyr397 creating binding sites for cellularsarcoma (c-Src) family of tyrosine kinase. These FAK/c-Src complexes trigger other downstream signals including stimulating mitogen-activated protein kinases (MAPKs). Activation of MAPKs like extracellular signal-regulated kinase 1/2 (ERK 1/2) lead to cell proliferation and/or trigger other vascular remodeling signaling in response to mechanical stimulus. ERK1/2 has been shown to be activated by mechanical stretch in a time and strength-dependent manner in the VSMCs isolated from rat aorta (Hu et al., 1998). Moreover phosphorylation of ERK 1/2 is essential for integrin-mediated phenotypic change of cultured aortic smooth muscle cells (Roy et al., 2001). ILK is a serine/threonine protein kinase that binds to the cytoplasmic tail of the β integrin subunit. It plays a central role in translating the external mechanical stimulus in to intracellular biochemical signals. ILK has been shown to involve in Ca2+ -independent phosphorylation of smooth muscle myosin light chain, hence it may provide an alternative pathway through which integrin activation regulates vascular resistance independent of Ca2+ (Deng et al., 2001). It has been shown as an essential component of mechanical stretch sensor by controlling the contractility in the zebrafish heart (Bendig et al., 2006); and deletion of ILK in zebra fish using antisense morpholino oligonucleotides results in marked patterning abnormalities of the vasculature and is lethal (Friedrich et al., 2004). Furthermore, inhibiting ILK using siRNA was shown to inhibit vascular oxidative stress mediated by hypertension (Vecchione et al., 2009). In addition, integrins is known to mediate the activation of the serine/threonine protein kinase Akt through FAK-dependent and independent mechanisms (Chen et al., 1996; Velling et al., 2004), and members of Rho GTPases, including RhoA and Rac1, which interact with a wide-variety of downstream effectors to engage in specific vascular responses.

1.3 Integrins as Mechanotransducers in the Vascular Cells There is considerable evidence suggests that integrins can transduce mechanical force across plasma membrane and initiate intracellular signaling (Ingber, 1990; Sadoshima and Izumo, 1993; Wang et al., 1993; Vuori, 1998; Boudreau and

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Jones, 1999; Davis et al., 2001). Extracellular mechanical force can be transmitted across the plasma membrane via integrins to initiate intracellular signaling in non-muscle cells (Ingber, 1990; Wang et al., 1993; Martinez-Lemus et al., 2003; Katsumi et al., 2004). It is known that an intact ECM- integrin-CSK axis is essential for mechanosensing and mechanotransduction in the endothelial cells and fibroblasts. The tensegrity model proposed by Ingber suggests that cellular tension or “prestress” is balanced and stabilized by the forces within the intracellular actomyosin cytoskeleton network (Ingber, 1997, 2000, 2003a, b; Wang et al., 2001). Disturbances in external mechanical stress can restructure the microtubules in the cytoskeleton to offset the tension in adherent cells. This transduced force and restructuring of microtubules can then initiate secondary messengers and signaling cascades including an increase in the [Ca2+ ]i causing contraction. Accordingly, a change of perfusion pressure in an intact blood vessel may alter the stress in the attachment sites between native ECM, and VSMC integrins, triggering downstream signaling events for myogenic response.

1.3.1 Evidence in Endothelial Cells and Fibroblasts The participation of integrins in mechanotransduction in endothelial cells was demonstrated by directly applying force to integrins using magnetic twisting cytometry technique with integrin ligand-coated ferromagnetic beads (Wang and Ingber, 1994; Wang, 1998). Force applied to β1 integrin induced focal adhesion formation and supported a force-dependent stiffening response. The cytoskeletal stiffness increased in direct proportion to the applied stress and required intact microtubules, intermediate filaments, and microfilaments. Mechanical force transduced via different integrins increased cytoskeletal stiffness to different degrees; for example, β1 integrin had the maximum effect (Wang et al., 1993; Wang and Ingber, 1995). These observations indicate that integrins serve as the mechanoreceptors to transduce force across the plasma membrane and trigger stiffening of cytoskeleton by inducing structural rearrangements within the tensionally integrated cytoskeletal network in endothelial cells. Magnetic force transduced via integrins using RGDcoated beads also increased the expression of endothelin-1 in endothelial cells (Chen et al., 2001). This integrin dependent transcriptional response is dependent on intact CSK. Inhibition of myosin ATPase, myosin light chain kinase, and disruption of actin filament abolished the twist-induced endothelin-1 upregulation. Glogauer and colleagues showed an increase in the [Ca2+ ]i when fibroblasts were pulled using collagen coated ferromagnetic beads (Glogauer et al., 1995). This process was presumably mediated via α1 -, α2 -, and α3 -integrins of collagen receptors, and was dependent on stretch-activated cation channels, whose sensitivity could be modulated by filamentous actin. Moreover, integrin dependent stretch-induced ERK2 activation was seen in rat cardiac fibroblasts (MacKenna et al., 1998). These findings highlighted the importance of integrin-CSK axis in transducing mechanical force into different physiological processes, including cellular contraction, Ca2+ signaling, gene regulation, and vascular remodeling.

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1.3.2 Evidence in Vascular Smooth Muscle Cells Using a Fluorescence Resonance Energy Transfer (FRET)-based cytosolic Src reporter in live smooth muscle cells, RGD- and β1 - integrin antibody coated beads rapidly (< 0.3 s) activated Src when force was applied through magnetic twist cytometry (Na et al., 2008). While forces applied with beads coated with nonadhesion ligand or non-adhesion integrin antibody did not elicit a response. These observations confirmed that mechanical force transduced by β1 – integrin activates Src. Application of cyclic mechanical force with fibronectin-, vitronectin-, and β3 – integrin antibody coated beads induced activation of ERK 1/2; α2 - and β1 - integrin antibody coated beads did not significantly activate the kinase (Goldschmidt et al., 2001). Using atomic force microscopy (AFM), Meininger and colleagues detected micromyogenic events in VSMCs that counteract the pulling force when these cells were pulled with fibronectin coated bead fused to the AFM probe tip (Sun et al., 2008). These microevents were suggested to be equivalent to the myogenic response observed in VSMCs in intact arterioles. These micromyogenic events were abolished in the presence of function blocking antibodies to α5 β1 and αV β3 integrins (Sun et al., 2008). α5 β1 integrins are the primary receptors for fibronectin in VSMCs. Using traction force microscopy to monitor the changes in traction force of individual renal VSMC in response to pulling of fibronectincoated beads, it was found that the traction force remained elevated even after the pulling force was terminated (Balasubramanian et al., 2008a). However, there was no residual traction force when beads coated with the non-adhesion ligand lowdensity-lipoprotein (LDL) were used to apply this pulling force. The persistence of elevated traction force might be interpreted as the sustained contraction triggered by the integrin-mediated mechanotransduction. Integrin mediated mechanotransduction has also been studied in isolated arterioles. Exogenous RGD-containing peptide impaired myogenic response in cremaster muscle arterioles and rat afferent arterioles (Martinez-Lemus et al., 2005; Balasubramanian et al., 2007). Myogenic response was also significantly inhibited when cremaster arterioles (Martinez-Lemus et al., 2005) and renal afferent arterioles were treated with either anti-α5 , anti-β1 or anti-β3 integrin antibodies (Yip et al., unpublished data). The ability to exhibit vasoconstriction is not compromised as seen by the response to KCl and norepinephrine. Hence, blocking integrins only impaired the pressure-induced vasoconstrictor response in the arterioles. These observations strongly suggested that the interactions between ECM and integrins are essential for myogenic constriction.

1.4 Mechanisms of Coupling Integrin Activation to Myogenic Constriction 1.4.1 Calcium-Dependent Mechanisms Mechanotransduction through integrins may lead to myogenic constriction by Ca2+ dependant and independent mechanisms. The Ca2+ dependent mechanism may

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involve multiple Ca2+ influx and release pathways. Activation of integrins has been implicated in the modulation of L-type Ca2+ channels. Early studies in rat cremaster VSMCs showed that beads coated with fibronectin or α5 integrin antibody enhanced, whereas αv β3 ligands or β3 antibody inhibited L-type Ca2+ currents (Wu et al., 1998). Another study showed that the activation of α4 β1 integrins also triggered an increase in Ca2+ influx via L-type channels (Waitkus-Edwards et al., 2002). Leu-Asp-Val (LDV) peptide, a sequence of an alternatively spliced fibronectin variant, caused vasoconstriction and increase in [Ca2+ ]i in cremaster arterioles; and the response could be blocked by nifedipine and anti-α4 antibody. The α5 β1 and α4 β1 integrin induced L-type Ca2+ current potentiation is mediated through tyrosine phosphorylation involving Src proteins in native VSMCs (Wu et al., 2001; Waitkus-Edwards et al., 2002). It was confirmed in a heterologous expression system that α5 β1 integrin regulates L-type Ca2+ channels (Cav 1.2) by PKA and c-Src dependent phosphorylation of α1C C-terminal residues Ser1901 and Tyr2122 , respectively (Gui et al., 2006). Furthermore, activation of α5 β1 integrin using antibodies or fibronectin also potentiated BK channel activity in rat cremaster VSMCs through both Ca2+ and c-Src dependent mechanism (Wu et al., 2008), suggesting negative feedback regulation of Ca2+ entry through L-type Ca2+ channel. In cerebral arteries, TRPC6 and TRPM4 channels have been shown to be involved in myogenic response (Welsh et al., 2002; Earley et al., 2004; Spassova et al., 2006; Reading and Brayden, 2007). But the connection between these TRP channels and integrin has not been established. In renal VSMCs, integrin mediated mechanotransduction triggers a global increase in [Ca2+ ]i . Application of Arg-Gly-Asp (RGD)-containing peptide initiated local increase in [Ca2+ ]i which propagated as recurrent Ca2+ waves across renal VSMCs (Chan et al., 2001; Balasubramanian et al., 2008b). However, the RGD-induced Ca2+ mobilization in renal VSMCs was ryanodine-sensitive, but not inhibited by nifedipine or removal of extracellular Ca2+ . Moreover, mechanical force applied by pulling freshly isolated renal VSMCs with fibronectin-coated beads triggered ryanodine-sensitive local Ca2+ release (Ca2+ spark) which was followed by global Ca2+ increase (Balasubramanian et al., 2007). In intact arterioles, stepwise increase of pressure from 80 mm Hg to 120 mm Hg induced an increase in the frequency of Ca2+ sparks (Yip et al., 2007; Balasubramanian et al., 2008a). Preincubation with ryanodine or RGD-containing peptide inhibited pressure-induced myogenic constriction (Balasubramanian et al., 2007). These findings support the concept that ligation of integrin modulate myogenic constriction in renal arterioles by activating Ca2+ release from the ryanodine-gated Ca2+ stores. Similar to renal VSMCs, a study in VSMCs of rat pulmonary arteries showed that the GRGDSP peptide (Gly-Arg-Gly-Asp-Ser-Pro) triggers Ca2+ release from ryanodine-sensitive stores and lysosome-like organelles; and Ca2+ influx is not required for the integrin mediated Ca2+ response (Umesh et al., 2006). It is interesting that myogenic response and integrin activation trigger both global and local Ca2+ response. Global mobilization of [Ca2+ ]i activates Ca2+ /calmodulin dependent myosin light chain kinase to initiate actin-myosin interactions and smooth muscle contraction; whereas local Ca2+ events may modulate ion channels and Ca2+ effectors in local vicinity to regulate vascular reactivity. In cerebral arterioles, increase in intraluminal pressure elicits myogenic response and activates

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Ca2+ sparks (Nelson et al., 1995; Jaggar et al., 1998), which are local Ca2+ release events originate from clusters of ryanodine receptors. It is well established that Ca2+ sparks causes large increase in local [Ca2+ ] (10–100 μM) in subsarcolemmal microdomains and activate nearby Ca2+ activated K+ channels (KCa ), leading to membrane hyperpolarization, reduction of Ca2+ influx via L-type Ca2+ channels and vasodilatation. They operate as the negative feedback modulators of membrane potential in cerebral arterioles. In contrast, the integrin mediated Ca2+ sparks in renal arterioles contribute to the myogenic response because their inhibition with ryanodine reversed the active myogenic vasoconstriction to passive vasodilatation (Yip et al., 2007; Balasubramanian et al., 2008a). Integrin induced Ca2+ sparks in renal arterioles may support vasoconstriction by functioning as elementary Ca2+ events underlying the global Ca2+ transients (Cheng et al., 1993). Alternatively, Ca2+ sparks may activate Ca2+ activated Cl– channels (ClCa )(Gordienko et al., 1999) to initiate myogenic response via membrane depolarization and the subsequent Ca2+ entry through voltage gated Ca2+ channels. The latter is supported by the observations of spontaneous transient inward ClCa currents activated by Ca2+ sparks in airway (Kotlikoff and Wang, 1998; ZhuGe et al., 1998; Bao et al., 2008), portal vein (Mironneau et al., 1996), and corpus cavernosum smooth muscle cells (Williams and Sims, 2007). A significant contribution of Ca2+ sparks in vasoconstriction has also been suggested in rat pulmonary arterial smooth muscle cells (Remillard et al., 2002; Zhang et al., 2003). Nevertheless, the roles of local Ca2+ signaling in integrin mediated myogenic response require further investigations.

1.4.2 Ca2+ Independent Mechanisms By examining the temporal and steady-state relationships between lumen diameter and vascular smooth muscle [Ca2+ ]i in isolated arterioles of hamster cheek pouch exposed to step changes in perfusion pressure, D’Angelo et al. has suggested the presence of a Ca2+ -independent regulatory system in the mechanotransduction of myogenic response (D’Angelo et al., 1997). As aforementioned, integrins are associated with kinases which phosphorylate proteins pertinent to smooth muscle contraction. Integrin-linked kinase (ILK) is an integrin associated protein which binds to the cytoplasmic tail of β1 and β3 integrins (Dedhar, 2000; Wu and Dedhar, 2001). ILK is a serine/threonine kinase with a low basal kinase activity, which is stimulated by cell-ECM interactions and certain growth factors (Dedhar, 2000). Mechanical stress evoked by high blood pressure in carotid artery increased the expression and activity of ILK (Vecchione et al., 2009). ILK can function as an Ca2+ -independent myosin light chain kinase to phosphorylate 20-kDa light chain of myosin and to induce contraction of vascular smooth muscle (Deng et al., 2001; Wilson et al., 2005). ILK also inhibits myosin light chain phosphatase via phosphorylation, and thus increases the Ca2+ sensitivity of VSMCs (Muranyi et al., 2002). It has been shown that pressure induced myogenic constriction in mouse tail artery required the generation of reactive oxygen species (ROS) via NADPH oxidase pathway (Nowicki et al., 2001; Keller et al., 2006). ILK activation might attribute to the activity of NADPH oxidase (Vecchione et al., 2009).

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1.4.3 Integrins as Part of a Larger Mechanotransduction Complex In addition to direct linkage to ECM and CSK, integrins could interact with other mechanosensory elements on the surface of VSMCs to form a larger mechanosensory complex. It has been suggested that cellular responses and signaling cascades initiated by mechanosensitive channels may be regulated more effectively if such channels were organized around integrins, kinases and cytoskeletal complexes (Shakibaei and Mobasheri, 2003). One possible mechanosensory protein that might interact with integrin in renal VSMCs is epithelial Na+ channel (ENaC). Members of the degenerin/Epithelial Na+ channel (DEG/ENaC) family of proteins are known to function as mechanosensitive channels in C. elegans. ENaC message and protein expression in interlobar arteries of mouse kidney have been confirmed by RT-PCR, Western blotting and immunofluorescence (Drummond et al., 2004; Jernigan and Drummond, 2005). Inhibiting DEG/ENaC with amiloride and benzamil also impaired myogenic constriction in mouse interlobar arteries, suggesting that their presence is vital for myogenic response in renal circulation (Drummond et al., 2004; Jernigan and Drummond, 2005). Similarly amiloride and benzamil impaired myogenic constriction in juxtamedullary afferent arterioles of rat kidney (Guan et al., 2009). Afferent arterioles are the major resistant arterioles in renal circulation. However, RT-PCR studies did not detect mRNA for α-, β-, or γ- subunits of ENaC in rat afferent arterioles (Wang et al., 2008). It has been speculated

Fig. 1.1 Potential interactions among integrins, Ca2+ sparks, ion channels in myogenic response. cADP – cyclic adenosine diphosphate; ClCa – calcium activated chloride channel; DEG/ENaC – degenerin/epithelial Na+ channel; ECM – extracellular matrix; ERK – extracellular signalregulated kinase; FAK – focal adhesion kinase; KCa – calcium activated potassium channel; KV – voltage gated potassium channel; MAPKs – mitogen-activated protein kinases; NKCC – sodium potassium chloride cotransporter; NCX – sodium calcium exchanger; NSCC – non-selective stretch-sensitive cation channel; RGD – Arg-Gly-Asp motif; RyR – ryanodine receptor; SR – sarcoplasmic reticulum; Src – sarcoma family of tyrosine kinases; TRP – transient receptor potential superfamily of channels; VGCC – voltage gated calcium channel

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that ENaC subunits form the pore and interact with integrins, ECM and CSK to form a larger mechanosensitive complex (Shakibaei and Mobasheri, 2003; Jernigan and Drummond, 2005; O’Hagan et al., 2005; Drummond et al., 2008). There is no direct evidence for these mechanosensitive complexes in VSMCs yet. However, β1 integrin subunit has been shown to co-localize with ENaC in mouse limb-bud chondrocytes by co-immunoprecipitation and immunofluorescence (Shakibaei and Mobasheri, 2003; Fig. 1.1).

1.5 Integrin-Mediated Mechanotransduction in Hypertension and Vascular Remodeling 1.5.1 Roles of Integrins in Systemic Hypertension Aside from myogenic vasoconstriction, the elevated mechanical stress imposed on the integrin signaling complex may play important roles in vascular remodeling and hypertension. Mechanical stimuli are initially transmitted via ECM-integrin-CSK axis to regulate VSMC functions (Dajnowiec et al., 2007); and vascular cells then react reciprocally by remodeling their surrounding ECM (Dajnowiec and Langille, 2007). Many integrins have been implicated in differentiation, migration, hypertrophy and proliferation of VSMCs, as well as ECM synthesis and deposition. These processes are essential for the inward eutrophic remodeling in resistance vessels, and outward hypertrophic remodeling in conduit vessels in hypertension. It has been reported that the expression of αV β3 and α5 β1 integrins in mesenteric arteries were upregulated in adult spontaneous hypertensive rats (SHR)(Bezie et al., 1998; Intengan et al., 1999); α1 , α5 , α8 , β1 , and β3 integrin expressions were enhanced in the aorta and carotid arteries of angiotensin II-induced hypertensive rats and mice, respectively (Brassard et al., 2006; Louis et al., 2007); and α8 integrin immunoreactivity was increased in VSMC of renal vasculature in desoxycorticosterone acetate (DOCA) induced hypertensive rats (Hartner et al., 2002). αV β3 and α5 β1 integrins are known to play major role in myogenic response. Their upregulation in SHR may account for the enhanced myogenic tone in these animals (Falcone et al., 1993; Shibuya et al., 1998; Falcone and Meininger, 1999). The sustained vasoconstriction caused by vasoconstrictors or myogenic tone in resistance arterioles is believed to initiate eutrophic inward remodeling by stimulating autorelengthening and repositioning of VSMCs, cross-linkage formation with ECM, structural rearrangement of matrix materials, and entrenchment of vascular wall (Bakker et al., 2002; Martinez-Lemus et al., 2004; Dajnowiec and Langille, 2007). Evidence from the TGR(mRen2)27 hypertensive rat model suggests that αV β3 integrin is involved in vascular remodeling (Heerkens et al., 2006). In these animals, hypertension was associated with upregulation of αV subunit and prominent arteriolar inward eutrophic remodeling. Inhibition of αV β3 integrin with a specific peptide inhibitor abolished inward remodeling but enhanced vascular smooth muscle hypertrophy. Inward eutrophic remodeling in isolated rat skeletal muscle arterioles in vitro

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was also inhibited with an antibody directed to β3 integrins (Bakker et al., 2004). Since αV β3 integrin is associated with myogenic response (Martinez-Lemus et al., 2005), VSMC migration (Jones et al., 1996) and ECM remodeling (Bendeck et al., 2000; Sajid and Stouffer, 2002), these observations are consistent with the notion that ECM-αV β3 interactions can trigger eutrophic remodeling. In addition, the collagen receptor α1 β1 integrin was upregulated in conduit arteries of angiotensin induced hypertensive rats and mice (Brassard et al., 2006; Louis et al., 2007). Evidence from α1 integrin knockout mice suggested that α1 β1 integrin play a significant role in smooth muscle cell hypertrophy in conduit arteries of hypertensive mice through the FAK and p38 MAPK signaling pathways (Louis et al., 2007). Furthermore, the tissue-type transglutaminase, a cross-linking enzyme that associates with β1 and β3 integrins to form co-receptors for fibronectin (Akimov et al., 2000), also contributed significantly to enthothelin-1 induced inward eutrophic remodeling in small resistance arteries (Bakker et al., 2005), suggesting that associated protein partners of integrin are involved in the remodeling process.

1.5.2 Roles of Integrins in Pulmonary Hypertension Integrin expression has been reported in pulmonary vasculatures (Damjanovich et al., 1992; Schnapp et al., 1995; Buck et al., 1996; Gotwals et al., 1996; Jones et al., 1997b, 1999; Yao et al., 1997; Medhora, 2000; Singh et al., 2000). Our survey with RT-PCR showed the presence of α1 -, α2 -, α3 -, α4 -, α5 -, α7 -, α8 -, αv -, β1 -, β3 -, and β4 - integrin mRNA in rat pulmonary arteries, similar to those expressed in rat aorta (Umesh et al., 2006). Their relative expression were in the orders of α8 >αv >α5 >α7 >α1 and β1 >β3 >β4 (Umesh and Sham, unpublished); and α5 , αv , β1 , and β3 integrin proteins were also detected in rat pulmonary arteries and aorta. Compare to systemic circulation, the basal vascular resistance and tone of pulmonary circulation are low and myogenic response is not evident under normal conditions except in fetal circulation. However, many studies have reported major contributions of ECM and integrins in vascular remodeling associated with pulmonary hypertension, which is characterized with outward hypertrophic and hyperplasia remodeling in larger vessels and neomuscularization in small arterioles of all forms of pulmonary hypertension (Stenmark et al., 2006, 2009). Increased deposition of extracellular matrix components, particularly collagen, elastin, tenascin-C, and fibronectin have been reported in pulmonary vasculature of human and animal models of the disease (Botney et al., 1992; Jones et al., 1996, 1997a, 2001; Novotna and Herget, 1998; Ihida-Stansbury et al., 2006). It is attributed to the increase in the activity of serine elastase, and a change in the balance of matrix metalloproteinase (MMP) and tissue inhibitors of metalloproteinase (TIMP) activity (Rabinovitch, 1999; Cowan et al., 2000; Zaidi et al., 2002; Hassoun, 2005). The increase in serine elastase activity activates MMPs to degrade native type I collagen, causing clustering of β3 containing integrins and increase tenascinC transcription. Tenascin-C in turn interacts with αv β3 integrin in pulmonary arterial smooth muscle cells to amplify the proliferative response by promoting epidermal

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growth factor receptor clustering and phosphorylation (Jones et al., 1997b, 1999). The evidence that αv β3 integrin blockade with selective antagonists induced apoptosis of pulmonary arterial smooth muscle cells and reduced medial wall thickness in organ culture of pulmonary arteries of monocrotaline-treated rats (Merklinger et al., 2005) supports an important role for αv β3 integrin in hyperplasia and hypertrophic remodeling. The expression of integrins in pulmonary vasculature with respect to pulmonary hypertension has not been systemically examined. Our recent preliminary study found significant changes in the expression of multiple α and β integrins in pulmonary arteries of chronic hypoxia and monocrotaline-induced pulmonary hypertensive rats (Umesh and Sham, unpublished). These changes in integrin expression could be related to the phenotypic changes of VSMCs occur during vascular remodeling. It is noteworthy that mild myogenic tone was observed in pulmonary microvessels of pulmonary hypertensive animals (Broughton et al., 2008), but the involvement of integrins has not yet to be determined.

1.5.3 Roles of Integrins in Vascular Remodeling Caused by Mechanical Injury Integrins are also major contributors in vascular remodeling and neointimal formation in vascular injury, atherosclerosis, and restenosis. α1 β1 , α5 β1 , αV β3 , α7 and β3 integrin expression were increased (Gotwals et al., 1996; Stouffer et al., 1998; Pickering et al., 2000; Kappert et al., 2001; Sajid and Stouffer, 2002; Chao et al., 2004) and the expression of α8 and β1 integrins were reduced in vascular cells of neointima formed after vascular injury (Gotwals et al., 1996; Zargham and Thibault, 2005). Among these integrins, αV β3 integrin has attracted most attention. Increased expression of αV β3 integrin in the medial and neointimal layers have been observed following mechanical injury in various animal models, including those in mice, rats, rabbits, baboons, and pigs, and in human atherosclerotic graft and arteriopathic coronary arteries (Hoshiga et al., 1995; Sajid and Stouffer, 2002; Sadeghi et al., 2004; Zhang et al., 2005). Many potential ligands of αV β3 , including osteopontin, thrombospondin, prothrombin, fibronectin, vitronectin, MMP-2 were found at the sites of injury with increased abundance. Studies using αV β3 antagonists to inhibit neointima formation and vascular remodeling suggested a central role of αV β3 integrins in neointima formation through multiple mechanisms including reduction of VSMC proliferation and migration, MMP production, and increased smooth muscle cell apoptosis (Slepian et al., 1998; van der Zee et al., 1998; Bendeck et al., 2000; Dufourcq et al., 2002; Honda et al., 2005). Similar to αV β3 antagonists, β3 integrin blockade with peptide inhibitors or anti-β3 antibodies reduced neointimal thickening, VSMC migration and proliferation, MMP activity in injured arteries (Slepian et al., 1998; Stouffer et al., 1998; Bendeck and Nakada, 2001). The β3 integrin mediated VSMC migration could be related to activation of β3 -FAK signaling

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pathway by osteopontin (Yue et al., 1994; Wang et al., 1996; Han et al., 2007). In contrast to pharmacological inhibition of β3 -integrin, deletion of β3 -integrin gene in mice provides no protection to transluminal probe induced mechanical injury, but inhibited VSMC accumulation in neointima after milder injury induced by carotid ligation (Smyth et al., 2001; Choi et al., 2004). The discrepancy could be explained by species differences in the response to vascular injury, mechanistic differences between β3 -integrin deficiency and blockade with antagonists, and compensatory mechanisms in β3 -integrin knockout animals. Other integrins, such as α1 β1 , α5 β1 , α7 β1 and α8 β1 , may also contribute to neointima formation. For example, the expression of α8 β1 integrin is downregulated in the medial layer in the early stage and is upregulated in the intima later during inward constrictive remodeling after balloon angioplasty (Zargham and Thibault, 2005; Zargham et al., 2007). It has been shown that α8 integrin is required for maintaining the contractile, differentiated phenotype of VSMCs (Zargham and Thibault, 2006). Gene silencing of α8 integrin increased VSMC migration and expression of smooth muscle cell de-differentiation markers; and overexpression of α8 integrin attenuated smooth muscle cell migratory activity and restored contractile phenotypes (Zargham et al., 2007a, b). The alteration in α8 β1 integrin expression, hence, correlated with de-differentiation of VSMCs for migration during early neointima formation, and the contractile phenotype of VSMCs during late constrictive remodeling in restenosis.

1.6 Conclusion and Perspectives It is beyond any doubt that integrins are major mechanosensors or mechanotransducers in many different cell types because of their unique properties of transducing extracellular mechanical signals exerted by ECM into intracellular signals via CSK, kinases and other associated proteins. In this chapter, we have reviewed the major evidence pertinent to the pivotal role of integrins as the mechanosensors of myogenic response in resistance arteries. Depending on the types of integrin involved and the specific vascular beds, integrin dependent myogenic vasoconstriction can be mediated through Ca2+ dependent mechanisms of L-type Ca2+ channel potentiation, activation of global and local Ca2+ release from ryanodine sensitive Ca2+ stores; as well as Ca2+ independent mechanisms of FAK, c-Src, MAPK, and ILK activation and reactive oxygen species release. Furthermore, integrin-mediated mechanotransduction of VSMCs is not only involved in acute adaptation of vascular resistance, but it also actively participates in chronic adaptation such as hypertension and vascular remodeling. Since integrins are ubiquitous transmembrane proteins, their role and significance in mechanotransduction should be pursued beyond myogenic response and vascular remodeling. Acknowledgements This work is supported by the NIH grants R01-HL071835, R01-HL075135, and American Heart Association.

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Ingber DE (2000) Opposing views on tensegrity as a structural framework for understanding cell mechanics. J Appl Physiol 89(4):1663–1670 Ingber DE (2003a) Tensegrity I. Cell structure and hierarchical systems biology. J Cell Sci 116 (Pt 7):1157–1173 Ingber DE (2003b) Tensegrity II. How structural networks influence cellular information processing networks. J Cell Sci 116(Pt 8):1397–1408 Inoue R, Jensen LJ, Shi J, Morita H, Nishida M, Honda A, Ito Y (2006) Transient receptor potential channels in cardiovascular function and disease. Circ Res 99(2):119–131 Intengan HD, Schiffrin EL (2000) Structure and mechanical properties of resistance arteries in hypertension: role of adhesion molecules and extracellular matrix determinants. Hypertension 36(3):312–318 Intengan HD, Thibault G, Li JS, Schiffrin EL (1999) Resistance artery mechanics, structure, and extracellular components in spontaneously hypertensive rats : effects of angiotensin receptor antagonism and converting enzyme inhibition. Circulation 100(22):2267–2275 Jaggar JH, Stevenson AS, Nelson MT (1998) Voltage dependence of Ca2+ sparks in intact cerebral arteries. Am J Physiol 274(6 Pt 1):C1755–C1761 Jernigan NL, Drummond HA (2005) Vascular ENaC proteins are required for renal myogenic constriction. Am J Physiol Renal Physiol 289(4):F891–F901 Johnson PC (1980). The Handbook of Physiology. The Cardiovascular System. Vascular Smooth Muscle. Am. Physiol. Soc., Bethesda, MD Jones FS, Meech R, Edelman DB, Oakey RJ, Jones PL (2001) Prx1 controls vascular smooth muscle cell proliferation and tenascin-C expression and is upregulated with Prx2 in pulmonary vascular disease. Circ Res 89(2):131–138 Jones JI, Prevette T, Gockerman A, Clemmons DR (1996) Ligand occupancy of the alpha-V-beta3 integrin is necessary for smooth muscle cells to migrate in response to insulin-like growth factor. Proc Natl Acad Sci USA 93(6):2482–2487 Jones PL, Cowan KN, Rabinovitch M (1997a) Tenascin-C, proliferation and subendothelial fibronectin in progressive pulmonary vascular disease. Am J Pathol 150(4):1349–1360 Jones PL, Crack J, Rabinovitch M (1997b) Regulation of tenascin-C, a vascular smooth muscle cell survival factor that interacts with the alpha v beta 3 integrin to promote epidermal growth factor receptor phosphorylation and growth. J Cell Biol 139(1):279–293 Jones PL, Jones FS, Zhou B, Rabinovitch M (1999) Induction of vascular smooth muscle cell tenascin-C gene expression by denatured type I collagen is dependent upon a beta3 integrinmediated mitogen-activated protein kinase pathway and a 122-base pair promoter element. J Cell Sci 112 (Pt 4):435–445 Kappert K, Blaschke F, Meehan WP, Kawano H, Grill M, Fleck E, Hsueh WA, Law RE, Graf K (2001) Integrins alphavbeta3 and alphavbeta5 mediate VSMC migration and are elevated during neointima formation in the rat aorta. Basic Res Cardiol 96(1):42–49 Karlsen FM, Andersen CB, Leyssac PP, Holstein-Rathlou NH (1997) Dynamic autoregulation and renal injury in Dahl rats. Hypertension 30(4):975–983 Katsumi A, Orr AW, Tzima E, Schwartz MA (2004) Integrins in mechanotransduction. J Biol Chem 279(13):12001–12004 Keller M, Lidington D, Vogel L, Peter BF, Sohn HY, Pagano PJ, Pitson S, Spiegel S, Pohl U, Bolz SS (2006) Sphingosine kinase functionally links elevated transmural pressure and increased reactive oxygen species formation in resistance arteries. Faseb J 20(6):702–704 Kirber MT, Walsh JV Jr, Singer JJ (1988) Stretch-activated ion channels in smooth muscle: a mechanism for the initiation of stretch-induced contraction. Pflugers Arch 412(4): 339–345 Kotlikoff MI, Wang YX (1998) Calcium release and calcium-activated chloride channels in airway smooth muscle cells. Am J Respir Crit Care Med 158(5 Pt 3):S109–S114 Lal H, Verma SK, Foster DM, Golden HB, Reneau JC, Watson LE, Singh H, Dostal DE (2009) Integrins and proximal signaling mechanisms in cardiovascular disease. Front Biosci 14: 2307–2334

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Chapter 2

The Role of the Actin Cytoskeleton in Mechanosensation Tianzhi Luo and Douglas N. Robinson

Abstract Cells are capable of sensing mechanical stimuli and translating them into biochemical signals. This ability allows cells to adapt to their physical surroundings by remodeling their cytoskeleton, activating various signaling pathways, and changing their gene expression. These phenomena involve two essential processes – mechanosensing and mechanotransduction. In these processes, force or deformation needs to be transmitted from the outside environment to the proteins and organelles inside the cell. The actin cytoskeleton composed of actin filaments, myosin motors, and actin crosslinking proteins plays a critical role in force propagation and in response to deformations. Cellular adaptation to these deformations is often associated with feedback loops, and proteins in the actin cytoskeleton accumulate and function cooperatively in response to mechanical stimuli. Mutations in these proteins cause failure in cellular mechanosensing, which eventually leads to cellular errors associated with disease progression. Keywords Mechanosensing · Mechanotransduction · Actin cytoskeleton · Actin-crosslinking proteins · Myosin II

2.1 Introduction A mechanotransduction system requires at least one sensor and one transducer. The sensors, commonly located next to the outer layer of the cell membrane,

D.N. Robinson (B) Department of Cell Biology, Department of Pharmacology and Molecular Science, Johns Hopkins University School of Medicine, 725 N. Wolfe Street, Baltimore, MD 21205, USA; Department of Chemical and Biomolecular Engineering, Johns Hopkins University Whiting School of Engineering, 3400 N. Charles St, Baltimore, MD 21218, USA e-mail: [email protected]


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sense the mechanical stimuli, such as force, pressure and flow speed, and transmit the mechanical signals into the inside of cells. Ion channels, protein kinases, integrins, membrane glycocalyx, G proteins, intercellular junction proteins and other membrane-associated signal-transduction molecules are capable of sensing mechanical stimuli as shown in Fig. 2.1 (Ingber, 2006; Vogel and Sheetz, 2006; Wang et al., 2009). The transducers, mainly situated inside the cell membrane or inside the cytosol, convert the mechanical signals to chemical and biological signals. Physiological examples of mechanosensation include hearing (Fettiplace and Hackney, 2006), blood flow regulation in the circulatory system (Chien, 2007), and bone remodeling (Robling et al., 2006). Auditory sensing occurs in the inner ear in a region known as the cochlea that is covered with hair cells characterized by stereocilia. The mechanoelectrical transduction (MET) ion channels in one stereocilium are linked to a neighboring stereocilium through tip linkers. These linkages are then coupled internally to the core actin bundles through myosin I motors. When one stereocilium is flexed due to sound vibrations, this induces tension along the tip link, leading to channel opening and conversion of the sound into an electrical signal. Subsequent adaption and associated closing of the METs occurs partially through the unbinding of myosin I from the actin bundles. In the circulatory system, endothelial cells sense the shear and stretch forces due to blood flow and activate a number of mechanosensors, such as membrane proteins, integrins, G proteins and ion channels. The mechanosensing triggers a cascade of signaling pathways and consequently modulates gene expression, resulting in cytoskeleton remodeling and cell realignment. Bone adapts its structure to mechanical stimuli and repairs structural damage through remodeling. At bone surfaces, osteoclasts and osteoblasts control the bone resorption and formation, respectively. However, the osteocytes, which are embedded deep within the bone, appear to have the primary

Fig. 2.1 Schematic cartoon of mechanosensation and mechanotransduction in cells

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The Role of the Actin Cytoskeleton in Mechanosensation

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job of sensing where the bone needs to be remodeled and then relaying this information to the osteoblasts and osteoclasts located on the bone surface. One appealing model suggests that the osteocytes sense changes in fluid flow inside canaliculi and the associated viscous drag creates tensile forces along the central actin filament bundle in the osteocytes. The osteocytes then release second messengers such as prostaglandins and nitric oxide, which activate the osteoblasts and osteoclasts. At the molecular level, the bone mechanotransduction is also thought to involve ion channels, focal adhesions, and G protein-coupled receptors, but detailed molecular mechanisms are still unclear. The actin cytoskeleton composed of actin filaments, actin crosslinking proteins (ACLPs) and myosin II motors, is unambiguously involved in these mechanosensing processes. To understand how mechanical stimuli are propagated through actin filaments and how the actin cytoskeleton responds to these stimuli, it is essential to understand the mechanical behaviors of the individual players, the reconstituted networks in vitro and the whole network in vivo.

2.2 Microstructures and Deformations of the Actin Cytoskeleton Cellular mechanics originate from mechanical features of the cytoskeleton that traverse many length- and time-scales. On the molecular level, the mechanical properties of proteins are determined by their molecular structures, such as peptide sequences, folding states and assembly states. On the next complexity level, the mechanical properties of the actin network depend not only on its morphology/microstructures, such as mesh size, filament length, bundle diameter and homogeneity, but also on the binding strength between the actin filaments and different crosslinkers and motor activities. In the past few decades, tremendous effort has been invested in characterizing the mechanical behaviors of ACLPs and motor proteins using single molecule techniques, and many attempts have also been made to study the mechanical properties of reconstituted actin networks. However, the understanding of individual ACLPs is still far from complete due to the complexity of these proteins and the limitations of the techniques. Additionally, the mechanical strength of reconstituted actin networks are often several orders lower than that of the intact cells. The central core protein of the actin cytoskeleton is monomeric actin, a 5-nm diameter, 42 kDa globular protein (G-actin) found in all eukaryotic cells. Each monomer is organized into four subdomains, flanking an internal cleft that binds ATP and magnesium ions. The end of G-actin close to the base of the cleft is called the plus end and the opposite end is called the minus end. G-actin monomers undergo polymerization and form microfilaments (F-actin) upon the addition of salt. Since the plus end of one G-actin is connected to the minus end of the neighboring G-actin, microfilaments also have well-defined plus and minus ends. The inclusion of ATP assists in G-actin stability and dramatically reduces the critical concentration for assembly. The 8-nm wide actin filament can be considered to have a left-handed

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helical morphology with 13 actin monomers per pseudo-repeat and a pseudo-repeat length of 37 nm. Alternatively, the actin filament can be considered to have a righthanded helical structure with two strands slowly twisting around each other. Each actin monomer is rotated 166◦ rotation with respect to its two nearest neighbors across the strand (Holmes et al., 1990). Within the strand, subdomains 2 and 4 contact subdomains 1 and 3 in the next monomer in the strand, and each monomer reaches across to the other strand through a hydrophobic plug that links the two strands together. The persistence length Lp of pure F-actin is around 17 μm. The intrinsic bending stiffness, κb = Lp kB T, and the elastic Young’s modulus E are 7 × 10−26 N·m−2 and 2.6 × 109 N · m−2 , respectively (Gittes et al., 1993; Kojima et al., 1994). The effective stretching stiffness is 44 pN·nm−1 provided the crosssectional area of a fully filled filament is 25 nm−2 , and the torsional rigidity is 8 × 10−26 N·m−2 (Tsuda et al., 1996). Non-muscle myosin II, a member of the conventional myosin II superfamily, is composed of functional hexameric monomers, consisting of two heavy chains, two essential light chains (ELCs) and two regulatory light chains (RLCs), which combine to form a ∼500 kDa complex (shown in Fig. 2.2). The amino-terminal motor domain consists of the catalytic core, which is structurally conserved with the Rasfamily small GTPase and includes the switch I and switch II helices and a Walker p-loop-family nucleotide-binding pocket. This catalytic core is functionalized with an actin-binding interface and a converter domain. The converter domain connects to an 8-nm long α-helix, which is wrapped by an ELC and RLC, forming the lever arm. The lever arm links to the carboxyl-terminal coiled coil domain (Warrick et al., 1987). Upon binding to actin, the motor domain remains rigid, whereas the lever domain is rotated through a ∼70◦ angle as the products of ATP hydrolysis are released; this lever arm rotation leads to the translation of the actin filament relative to the coiled coil of the myosin. To generate force and to bear load, myosin II must assemble into bipolar thick filaments (BTFs). These assemblies can range from as few as eight (Acanthamoeba)

Fig. 2.2 Domain structure of myosin II. HMM and LMM represent heavy meromyosin and light meromyosin, respectively

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to as many as 400 hexamers (mammalian skeletal muscle). Most mammalian nonmuscle myosin IIs (nonmuscle heavy chain IIA, IIB and IIC) assemble into BTFs ranging from 10–30 monomers, and Dictyostelium discoideum myosin II is thought to assemble into BTFs with up to 70 monomers. For Dictyostelium, the assembly process is thought to occur through the formation of two monomers into a parallel dimer and two of these parallel dimers join together to create an anti-parallel tetramer. Once the anti-parallel tetramer is formed (the stable nucleus), the BTF grows through side-by-side (lateral) addition of dimers, resulting in no extra elongation of the BTF as more dimers are added (Fig. 2.3) (Mahajan and Pardee, 1996). In the thick filament, myosin II molecules are thought to stack their rod tails in parallel with a small overlap where the subunits are held together through electrostatic interactions (Hostetter et al., 2004). Therefore, unlike muscle myosin II thick filaments which grow in length as they are assembled, the Dictyostelium myosin monomer is 250 nm whereas the thick filaments are just 400 nm long, independent of the number of monomers assembled. The assembly process is regulated by myosin heavy chain kinases (MHCKs), which phosphorylate three critical threonines in the tail region (Liang et al., 1999). This phosphorylation prevents the myosin monomer from assembling into BTFs, which is necessary to maintain a free pool (80–90% of total myosin II) of monomeric myosin in the Dictyostelium cell. ACLPs can organize actin filaments into bundles and branched networks, depending on their molecular structures (as shown in Fig. 2.4), kinetic properties, and concentration (Revenu et al., 2004). Bundle forming ACLPs include fascin, forked, villin, fimbrin, espin, scruin, plastin, cortexillin and dynacortin, whereas examples of meshwork forming ACLPs are filamin, Arp2/3, gelsolin and ERM (ezrin, radixin, and moesin) proteins. Some of these proteins have additional properties. For example, Arp2/3 also nucleates actin assembly and ERMs link actin filaments to the membrane. However, this classification of bundlers versus meshwork formers is an oversimplification as some ACLPs, such as α-actinin and dynacortin, can form bundles or meshworks depending on concentration and actin:crosslinker ratios. Some of these ACLPs, such as villin, espin, forked, fimbrin

Fig. 2.3 Illustration of the assembly process of non-muscle myosin II

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Fig. 2.4 Schematic graph of a few actin crosslinking proteins:(a) human filamin A; (b) αactinin; (c) cortexillin-I. In all cases, “N” and “C” represent the N-terminus and C-terminus, respectively

and fascin, are monomeric with at least two actin-binding motifs, allowing them to crosslink as monomers. By contrast, other ACLPs, such as filamin A, α-actinin, dynacortin, and cortexillin are dimeric, and filamin A, cortexillin, and most likely dynacortin form parallel dimers while α-actinin forms anti-parallel dimers. Actinbinding domains are localized at the amino-terminus of filamin A and α-actinin. Due to the differences in their structures, Y-shaped filamin A dimers can only crosslink actin filaments into nearly orthogonal networks while α-actinin forms either networks or loose bundles. Some of the most common filamentous actin-binding domains (ABDs) are the calponin homology (CH) domain, the Wiskott-Aldrich syndrome homology region 2 (WH2 ) domain, the gelsolin homology (GH) domain, and the formin homology (FH) domain (Sjöblom et al., 2008). Different ABDs generally associate with different surfaces on the actin monomer and with different apparent affinities. These network structures may experience four types of deformations (Vogel, 2006; Ferrer et al., 2008): (1) deformation in actin filaments, (2) deformation of the binding between actin filaments and myosin-II, (3) deformation of the binding between actin filaments and ACLPs and (4) intramolecular deformation of ACLPs. In other words, intermolecular and intramolecular deformations exist at the same time within the same network. One additional feature of how these networks respond to deformation is the relationship of the time-scale of the deformation to the time-scale of the turnover/remodeling of the network. For example, profilin binds monomeric actin, inducing it to exchange its nucleotide and shuttling the actin monomer to sites where elongation has been stimulated. By contrast, cofilin binds cooperatively to sever the actin filaments, which has a complicated effect on the actin network. Severing provides free actin plus ends that may grow whereas the free minus ends can disassemble. These proteins influence the time-scales over which the actin polymers can grow and disassemble, modulating the time-scales of remodeling of the actin network. These features of network remodeling may provide a phenomenological viscous character to the network, allowing it to flow and remodel over the long time-scales of many biological processes.

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2.2.1 Intermolecular and Intramolecular Deformations Intermolecular interactions, including hydrogen bonds, electrostatic interactions, van der Waals interactions and hydrophobic interactions, are non-covalent in nature. Based on the transition state theory, there are two popular bond models to describe the single-bond behaviors of intermolecular interactions: the catch-bond model and the slip-bond model. A simplified definition of the catch-bond model is the lifetime of the bond increases as the stretching force increases. In contrast, a slip-bond shows the opposite behavior, i.e., the bond lifetime decreases as the force increases. A schematic diagram of the energy landscapes of the two different models are shown in Fig. 2.5. In the spirit of Bell (Bell, 1978), both models can be described by the unified formula (Evans and Ritchie, 1997; Evans, 2001): 0 exp koff = koff

f x , kB T

(2.1)

0 = ν exp −G k T is the unbinding rate in the absence of force, ν where koff B is the vibration frequency, G is the energy difference between the transition state and the bound state, kB is Boltzmann’s constant, T is temperature, f is the external force that pulls the two molecules apart, and x is the bond length change along

D

Energy slip bond pathway

catch bond pathway

bound state

unbound state

unbound state

Force Direction

Fig. 2.5 A schematic diagram of the force dependence of protein-protein interaction strength: (a) catch-bond model; (b) slip-bond model; (c) three typical bond behaviors; (d) two pathways between bound state and unbound state. Red and blue lines are energy landscapes in the presence and absence of external load, respectively

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the force direction. When the force pulls the system close to the transition state, i.e., f x > 0, Eq. (2.1) describes the slip-bond model; however, when f x < 0, it refers to the catch-bond model. Equation (2.1) can only be applied to a single pathway (either catch-bond or slip-bond). However, sometimes there are multiple pathways between the bound state and unbound state, and catch-slip and slip-catch transitions usually occur (as shown in Fig. 2.5). In that case, the unbinding rate is a superposition of the rates of two different pathways: koff =

ks0 exp

f xs kB T

+ kc0 exp

f xc kB T

,

(2.2)

where the subscript s and c represent slip and catch, respectively. To characterize intermolecular interactions, two kinds of experiments are often conducted: the measurement of the bond life-times at a constant force and the measurement of rupture forces at a constant loading speed. Under constant force conditions, the surviving probability P (t) of a bond in the bound state satisfies (Thomas, 2008) dP (t) = −koff (f ) P (t) , dt

(2.3)

where koff (f ) is described by Eq. (2.1). Apparently, P (t) decreases exponentially over time and the slope of P (t) in log scale is −koff (f ). For the slip-bond model, the surviving probability shows a larger negative slope when force increases; whereas for the catch-bond model, it shows the opposite trend. Under constant loading rate conditions, the rupture force is x kB T kB T ln 0 ln rf , + f = x x koff kB T ∗

(2.4)

where rf is the loading rate and f (t) = rf t (Ackbarow et al., 2007). Equation (2.4) predicts a linear relationship between the rupture force and logarithm of the constant rate. If the intermolecular interaction involves multiple bonds, the f ∗ ∼ ln loading rf will show different slopes associated with different bond energies. Intramolecular interactions include covalent bonds, hydrogen bonds, electrostatic interactions, van der Waals interactions, and hydrophobic interactions. Mechanical deformation can cause twisting and stretching of a single bond or the whole protein, general conformational changes and the unfolding of domains. Since intramolecular interactions are much more complicated than intermolecular interactions, there is no simple mathematic model to describe how mechanical force affects the intramolecular interactions even though protein unfolding can still be probed experimentally using similar techniques. However, for a linear polyprotein, such as an actin filament and titin, the force-extension relationship can be predicted using a worm-like-chain (WLC) model that was originally developed to describe the mechanical behaviors of double stranded DNA molecules (Bustamante et al., 1994; Marko and Siggia, 1995):

2


kB T f = Lp

x 1 − , 2 + Lc 4 4 1 − x Lc 1

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(2.5)

where Lp is the persistence length, Lc is the contour length, and x is the extension. Another popular model of the force-extension of proteins is the free-joint-chain (FJC) model (Bueche, 1962): fl kB T − , x = Lc coth kB T fl

(2.6)

where l is the Kuhn monomer size. In the WLC model, the force is written as a function of the chain extension while in FJC model the extension is written as a function of the force. However mathematically, both models give similar force-extension curves.

2.2.1.1 Force Generation Associated with Actin Polymerization Actin polymerization, like microtubule polymerization, is force generating. During actin polymerization, actin monomers bound with a nucleotide assemble into helical filaments. The nucleotide binding cleft sits between subdomain 2 and subdomain 4, which faces the minus or pointed end of the actin filament. The opposite end is the fast growing so-called barbed- or plus-end of the filament. The plus and minus ends have different affinities for ADP-actin or ATP-actin monomers with the plus end having the highest overall affinity for ATP-actin monomer. After some time delay after the ATP-actin monomer incorporates into the polymer, the ATP is hydrolyzed to ADP•Pi . Subsequently, the Pi is released, leaving behind an ADP-actin monomer within the actin filament. The effect of these kinetics is that a density gradient of ATP-, ADP•Pi -, then ADP-actin monomers extends from the plus- to minus-ends of the actin polymer. In comparison to ATP-monomers, ADP-monomers have different conformations and are less stable in the filament form. Therefore, at concentrations between the critical concentrations of the two ends, net polymerization at the plus end and net depolymerization at the minus end lead to treadmilling: the net flow of actin monomers from plus end to the minus end. During polymer assembly, the filament can generate a force, measured at 1 pN, which is close to the theoretical estimate (for a given concentration of G-actin) according to f =

k+ cA kB T , ln δ k−

(2.7)

where δ is the length increment of one monomer addition, k+ (k− ) is the on(off) rate of polymerization at the plus end, and cA is actin concentration (Kovar and Pollard, 2004; Footer et al., 2007). Therefore, filament assembly by itself is force generating, and compressive forces can suppress filament growth. Furthermore, the

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theoretical estimate implies that simply by increasing the free actin concentration, greater forces may be generated by filament assembly. However, this process is limited by the flexural rigidity of the filament, which sets a force limit beyond which the filament buckles. However, actin-associated proteins may increase the flexural rigidity, increasing the range of forces that might be generated by filament assembly.

2.2.1.2 Force-Dependent Behaviors of Actin-Myosin Binding Myosin II is one type of an actin-based motor that converts chemical energy into mechanical work by amplifying a small conformational change associated with ATP hydrolysis in its motor domain and translating it into the relative movement of the myosin II and the actin filament. The actin-activated myosin II ATPase cycle is shown in Fig. 2.6 (Spudich, 2001). Initially, ATP binds to the nucleotide-binding pocket in the myosin head (motor domain), which results in the unbinding of the motor from an actin filament. Upon ATP hydrolysis, the myosin rotates its lever arm, moving the head into the pre-stroke state. In this state, the motor can weakly sample the actin filament in search of its binding site. Upon binding, the motor locks on tightly and releases the Pi as the head begins to swing the lever arm through its working stroke (a ∼70◦ rotation). Upon completing the full working stroke, the ADP-bound motor remains locked onto the actin filament. Subsequently, the motor releases the ADP, forming the so-called rigor state. ATP can then rebind, starting the cycle over again, which will continue until ATP is depleted. The time the motor spends tightly bound to the actin filament is the strongly bound state time. The ratio of the strongly bound state time to the entire ATPase cycle time gives rise to a duty ratio, which specifies the fraction of time each motor domain spends tightly bound to actin and correspondingly, the fraction of motor heads tightly bound to actin at any time. Because the motor generates force as it translates the actin filament, the

Fig. 2.6 Myosin II ATPase cycle

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motor performs work as it undergoes its conformational change. For most myosin family members (with myosin VI having some unique and exquisitely complicated twists on this theme (Phichith et al., 2009)), the head region is highly rigid while the lever arm is considered to be elastic. Furthermore, the ADP-release step itself is not force-sensitive for most myosin isoforms. Rather the conformational changes that precede the formation of the ADP-bound post-stroke configuration are forcesensitive. Therefore, as the motor swings the elastic lever arm through its power stroke, resistive tension can lead to deformation (strain) of the lever arm, inhibiting the lever arm swing and locking the motor in the load-bearing, transition state. Moreover, the step-size of the myosin is related to the length of its lever arm, and this relationship has been shown to be linearly proportional to the lever arm length for Dictyostelium myosin II. A further hypothesis is that the maximum force depends on the length of the lever arm (Uyeda et al., 1996). Lower force is required to stall a motor with a long lever arm whereas the same motor with a short lever arm powers through greater loads before being stalled by load, implying that the strain on the lever arm prevents the full conformational change needed to allow for the motor to acquire the conformation where it can release the ADP. This load-dependency is consistent with a catch-slip-like behavior, which can be interpreted using Eq. (2.4).

2.2.1.3 Force-Dependent Binding Between Actin Crosslinkers and Actin Filaments In order to crosslink or bundle two actin filaments, an ACLP must have a minimum of two actin-binding domains associated with each functional unit through which the ACLP contacts the actin filaments. This can occur by either dimerizing a monomer that contains one ABD or by having two or more ABDs within a monomer. Furthermore, the strength of the actin interactions and the conformation of the actin network (meshwork or bundle) varies for different ABDs and the different conformations of ABDs within the ACLP. For example, the ABDs may be closely linked through short spacers (e.g. fimbrin), resulting in tightly packed actin bundles or there can be elongated spacers (e.g. α-actinin and spectrin), which can form relatively loose networks (Bañuelos et al., 1998). The most common ABD module is composed of two tandem CH domains; this module is found in ACLPs such as α-actinin, spectrin, dystrophin, fimbrin, filamin, plectin and cortexillin. For comparison, the dissociation constants of Acanthamoeba α-actinin, chicken smooth muscle α-actinin and Dictyostelium α-actinin are 4.7, 0.6 and 3 μM, respectively even though the first two have very similar structures (Wachsstock et al., 1993). In single molecule measurements, filamin A and rabbit muscle α-actinin have actinunbinding energies of 4.3 kB T and 3.6 kB T, respectively, while displaying increasing rupture forces with increasing loading rates (Ferrer et al., 2008). The loading rate dependence of the rupture forces of these crosslinkers is indicative of catch-bond behavior.

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2.2.1.4 Force-Dependent Intramolecular Deformation of ACLPs During the deformation of the actin cytoskeleton, ACLPs undergo intramolecular deformations, which involve domain unfolding and shearing and stretching between two actin-binding domains. The unfolding process can be assessed experimentally by single molecule stretching, and the resulting saw tooth-like force-extension curve usually agrees well with the WLC model. Molecular dynamics simulations can also be used to computationally pull on proteins from various directions at relatively high pulling speeds to reveal detailed unfolding schemes. The most common structures of interest typically include bundles of α-helices or β-sheets (Rohs et al., 1999; Ackbarow et al., 2007; Buehler and Keten, 2008). Two studied examples are filamin and α-actinin, both of which form anti-parallel homodimers. However, filamin is constructed from multiple β-sheet-like immunoglobulin (Ig) domains, the number of which differs between various family members. By contrast, α-actinin includes multiple spectrin repeats, each of which consists of a bundle of three α-helices. Dictyostelium filamin (DdFLN) consists of an ABD at the amino-terminal end followed by a rod domain, containing six Ig domains. Single molecule stretching of DdFLN revealed that the sequence of the fourth Ig domain is unique because it has a lower unfolding force (Schwaiger et al., 2004). Additionally, it was found that the fourth Ig domain has an intermediate unfolded state in the low force regime. Based on the WLC model, the persistence length is 0.5 and 0.9 nm for the high force regime and the low force regime, respectively. The corresponding unfolding force ranges from ∼50–250 pN and the periodicity of extension also ranges from ∼14–17 nm. Similarly, the human endothelial filamin A has one ABP and 24 Ig repeats plus two flexible hinges. However, despite the diversity in the structures between the two filamin family members, the force extension curve reveals a similar persistence length of 0.33 nm and an unfolding force, ranging from 50 to 200 pN (Furuike et al., 2001). The unfolding of spectrin repeats requires an unfolding force of ∼30–50 pN, a persistence length of 0.8 nm and an extension period of 31 nm (Rief et al., 1999). In the Ig domains, β-strands are arranged almost in antiparallel fashion with small twist angles mainly through hydrogen bond interactions. During mechanical unfolding, two anti-parallel strands slide in opposite directions while breaking the hydrogen bonds between them (Lu and Schulten, 2000; Keten and Buehler, 2008). Many studies have shown that the unfolding of β-sheet proteins depends highly on the pulling directions (Brockwell et al., 2003; Nome et al., 2007; Dietz et al., 2006; Bertz et al., 2009). In each spectrin repeat, three antiparallel α-helices linked by loops are folded into a left-handed coiled coil (Pascual et al., 1997; Djinović-Carugo et al., 1999). Despite the elasticity of coiled-coil structures, the unfolding of these repeat domains mainly relies on the stretching of the linkage between different helices (Altmann et al., 2002). The linkage dependent unfolding has also been observed in other proteins (Carrion-Vazquez et al., 2003). In summary, the full force-extension relationship of individual proteins can be obtained by linear superposition of the force-extension of each domains and linkages while considering the corresponding folding and unfolding probability at certain forces (Li et al., 2002).

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2.2.2 Mechanical Properties of an Actin Network The mechanical properties of an in vitro assembled actin network depend on the average length of actin filaments, the mesh size of the actin network, the concentration of myosin II, the assembly states of myosin II thick filaments, the concentration of ACLPs, the binding strength between ACLPs and actin filaments, the mechanical properties of each ACLP and the heterogeneity of the actin network (Gardel et al., 2004a; Wagner et al., 2006; Bausch and Kroy, 2006; Ferrer et al., 2008). The addition of myosin II can alter the fundamental character of the actin network. In response to mechanical stimuli, the actomyosin system undergoes continuous remodeling of its microstructure. The remodeling of the actin-myosin II contractile system includes assembly/disassembly of actin filaments and myosin II thick filaments and bundling/unbundling of actin filaments by ACLPs. During remodeling, the whole network is more or less out of mechanical equilibrium, which leads to transient behaviors within the network (Mizuno et al., 2007; Wilhelm, 2008). 2.2.2.1 Mechanical Properties of Pure Actin Gels Polymers can be divided into three groups based on two length scales: the persistence length Lp and the contour length Lc . A filament is considered rigid if Lp >> Lc or flexible if Lp 20, respectively, and transient behaviors exist in the range of 2 < λ < 20. For a fixed contour length (actin dynamics reaches steady state), elasticity is dominated by the stretch modulus under high crosslinker density or small lc conditions (affine); the bending modulus dominates under low crosslinker density or large lc conditions (nonaffine). The apparent strainstiffening during deformation is basically a nonaffine to affine transition (Onck et al., 2005; Gardel et al., 2004b). The nonaffine-affine transition theory also successfully

Fig. 2.9 Mechanical properties of actin networks crosslinked by α-actinin and fascin. The synergistic effect on the elasticity of α-actinin and fascin is shown in a. The elasticity displays [fascin]/[α-actinin] dependence in b. The exponent of the elasticity as a function of frequency is shown in c. The total concentration of ACLPs is 0.96 μM in b and c and the actin concentration is 24 μM for all panels. Reproduced from Tseng et al. (2005) with permission of Elsevier

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interprets the scalings between G (or K ) and cA , ς and the pre-stress observed in these experimental systems. The corresponding microstructural transition was observed in scruin-bundled actin network by confocal microscopy (Liu et al., 2007).

2.2.2.3 Effects of Myosin II on the Mechanical Properties of the Actin Network In the absence of ACLPs, myosin II motors pull on actin filaments along their axial directions as shown in Fig. 2.10b. Over time, actin filaments aggregate, resulting in a heterogeneous distribution of actin filaments (super-precipitation). In the presence of saturating ATP, active myosin II does not change the shear modulus of an actin filament solution, whereas in the absence of ATP, inactive myosin II crosslinked Factin, leading to a >10-fold enhancement of the shear modulus (Humphrey et al., 2002). Additionally, the decreasing of G G and the relaxation time associated with active myosin II indicates that myosin II increases the fluidity of the actin network. This apparent fluidization likely arises from cycles of pulling and releasing, which generate local fluctuations inside the actin network.

Fig. 2.10 Microstructures and mechanical properties of actin meshwork with muscle myosin II and human filamin A. Electron micrographs of pure actin, actin+myosin ([myosin]/[actin] = 0.02 and 5 mM ATP) and filamin A +myosin+actin ([filamin]/[actin] = 0.005, [myosin]/[actin] = 0.02 and 5 mM ATP) are shown in a, b and c, respectively. The corresponding fluorescent images are found in the inserts. The illustration of an active stiffening mechanism is shown in d. Stiffening behaviors of active networks (with myosin to actin ratio of 0.02 (blue squares), 0.005 (green squares), and 0.001 (red squares)) and a passive network at fixed [filamin]/[actin] of 0.005 (white triangles) are shown in e. The filamin concentration dependence of rupture force is shown in f. Critical strain decreases as a function of the internal stress generated by myosin II. Scale bars are 10 μm in all panels. Reproduced from Koenderink et al. (2009) with permission by Proc Natl Acad Sci USA

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In the presence of ACLPs, internal tensile stresses associated with the myosin II motor stiffen the actin network, leading to stress-stiffening such that G (ω) ∼ G (ω) ∼ ω1/2 (Mizuno et al., 2007). In assembled networks, myosin II and filamin A work together to enhance the network stiffness as shown in Fig. 2.10 (Koenderink et al., 2009). Increasing the myosin concentration leads to higher differential modulus and increasing the filamin A concentration makes the network able to sustain higher stresses. The critical strain displays a power-law decay of the internal stress that is dependent on the myosin II concentration. Because ACLP binding to actin is dynamic, myosin may break the binding between ACLPs and actin when the myosin concentrations become high enough, resulting in the power-law decay of the critical strain with increasing myosin concentration.

2.2.3 In Vivo Measurements of Cell Mechanics In comparison to reconstituted actin cytoskeletons, the shear moduli of intact cells are several orders higher (Hoffman et al., 2006; Girard et al., 2004; Reichl et al., 2008). One reason for this difference is that there are tens of ACLPs with concentrations on the order of 1 μM in intact cells compared to just one or two ACLPs as in the in vitro experiments. Another reason is that myosin II motors generate contractile forces that not only stiffen the actin network but also enhance the binding between some ACLPs and actin filaments that further increases the stiffness of the actin cytoskeleton (Reichl et al., 2008). In Dictyostelium, myosin II null cells have decreased cortical tension and elastic moduli compared to wild type cells. Similarly, mutant cells depleted with various ACLPs also have softer cortices as shown in Fig. 2.11 (Girard et al., 2004; Reichl et al., 2008). In the myosin II null background, a complex relationship between the ACLPs and myosin II is observed. Some ACLPs (for example, dynacortin) have large effects upon their depletion from a wild type background but smaller effects when depleted from a myoII mutant background. By contrast, the depletion of fimbrin has an even more complicated effect being both time-scale sensitive and myosin II dependent: fimbrin contributes only to the viscoelastic moduli on fast time-scales in a wild type background, but also impacts the cortical tension of myoII null cells. More generally, similar to the reconstituted actin cytoskeletons, the cytoskeleton of intact cells also display G (ω) ∼ G (ω) ∼ ω3/4 in the high frequency regime, indicating that the mechanical properties of cells are dominated by the entropic vibrations of actin filaments at these frequencies. However, in the low frequency range, cells exhibit elastic moduli, G (ω) ∼ G (ω) ∼ ωβ , where 0 < β < 0.3 (Hoffman et al., 2006; Deng et al., 2006; Girard et al., 2004; Reichl et al., 2008). The difference in the low frequency regimes between intact cells and artificial actin cytoskeletons has been primarily attributed to the forces generated by motors that push the system out of thermal dynamic equilibrium (Lau et al., 2003), which is consistent with the in vitro observations (Humphrey et al., 2002; Mizuno et al., 2007). Indeed, myosin II null cells have a fundamentally different character in the low frequency range that is more consistent with a passive network (Girard et al., 2006).

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Fig. 2.11 Mechanical properties of Dictyostelium cells. a and c show the frequency dependence of complex moduli of different cell-lines. b and d show the effective cortical tension for these cells. “wt” and “myoll” represent wild-type and myosin II-null cells, respectively. “dynhp” and “fimhp” refer to dynacortin-hairpin and fimbrin-hairpin, respectively. Dynacortin and fimbrin are ACLPs, and hairpin constructs are used to silence gene expression through RNA interference. Reproduced from Reichl et al. (2008) with permission of Elsevier

Many of the studies of living cells draw upon laser-based tracking of single or multiple particles embedded in the living network. Because the particles may fluctuate due to thermal or active forces that act on the particles, the fluctuation-dissipation theorem (FDT) is used to extract viscoelastic parameters. However, the FDT should only be applied to systems at equilibrium, not out of equilibrium. Therefore, by measuring the mean square displacement (MSD) of the particles as a function of correlation time, it was found that the FDT cannot describe the particle behaviors in the low frequency regime (Lau et al., 2003; Bursac et al., 2005; Girard et al., 2006; Mizuno et al., 2007; Wilhelm, 2008; Reichl et al., 2008). Furthermore, in some contexts, the FDT is violated because the apparent diffusive behavior has a much larger magnitude than expected, considering the viscous damping for the particle size. This suggests that local active processes can essentially facilitate the stirring of the cytoplasm (Brangwynne et al., 2009). It was further demonstrated that the mechanochemistry of myosin II motors in combination with ACLPs regulates the

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MSDs in the low frequency regime (Girard et al., 2006). ACLP mutant cells also display significant effects on the complex modulus over a wide frequency range (Girard et al., 2004). The coupled effect of myosin and ACLPs was also investigated (Girard et al., 2006; Reichl et al., 2008). The frequency-dependent mechanical behaviors of cells imply that microscopic processes, such as unbinding events and conformational changes of ACLPs and motors with a broad distribution of characteristic times play important roles in regulating cell mechanics. Based on these types of experimental observations, three major cell mechanics models have been proposed: tensegrity (Ingber, 2003), soft glassy rheology (Fabry et al., 2001; Trepat et al., 2007; Zhou et al., 2009) and the sol-gel hypothesis (Janmey et al., 1990). In the tensegrity model, the actin network, including myosin II and ACLPs, is considered to be primary sources of pre-stress (Wang et al., 2001), and cell stiffness is proportional to the pre-stress that cells experience. Tensegrity can help explain the stress-stiffening, but it does not predict the power-law behaviors of cells. The soft glassy rheology (SGR) model considers the cells glassy materials that are microstructurally disordered and thermodynamically close to a glass transition. The deformations in glassy materials are microscopically inelastic, localized and time-dependent. The SGR model successfully accounts for the power-law behavior in the low frequency regime but it does not capture the stress-stiffening. The sol-gel model assumes that the cell is a gel of filamentary polymers embedded within a fluid cytosol, which predicts a weaker frequency dependency in the low frequency regime than is observed experimentally. Thus, all models capture some aspects of cell mechanics, but a single model that accounts for all of cellular behaviors has yet to emerge.

2.3 Functions of the Actin Cytoskeleton in Mechanosensation Mechanosensing, the sensing of mechanical force, is crucial for a range of processes that extend over a wide range of length- and time-scales and from the molecular to organismal levels. At the cellular level, stretch receptors in the plasma membrane and many components of the cytoskeletal network are obvious targets of external forces. However, internally generated forces are also felt by the same machinery, allowing intrinsic regulation and cross-talk to occur through the heterogeneous cytoskeletal network. Here, we will focus on two mechanisms for mechanosensing in the actin network: the crosslinked actin network complete with myosin II motors, which governs the cell shape changes particularly during cytokinesis (Effler et al., 2006; Ren et al., 2009) and the actin-associated proteins found in focal adhesions (Vogel and Sheetz, 2006).

2.3.1 Mechanosensing Through Myosin II and Actin Crosslinking Proteins Dividing cells have a mechanosensory system that they use to monitor their shape as they cleave into two daughter cells. The system could be activated in a controlled

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Fig. 2.12 Mechansensing system in Dictyostelium. a and b show the accumulation of fluorescently labeled cortexillin I and myosin II, respectively. Cortexillin I and myosin II are green. The microtubules are labeled red with RFP-tubulin. The lever arm length of different myosin II mutants is shown in c. The percentage of cells displaying mechanosensing increases with elevated pressure as shown in d. Reproduced from Ren et al. (2009) with permission of Elsevier

fashion, using micropipette aspiration (Effler et al., 2006). Here, myosin II and the actin crosslinker cortexillin I accumulate cooperatively in highly deformed regions in dividing wild type cells as shown in Fig. 2.12. Both myosin-II and cortexillin-I are necessary for this function since cell-lines devoid of either protein are unable to respond to cellular deformations (Ren et al., 2009). Furthermore, only fully wild type myosin-II and wild type cortexillin are able to fully restore mechanosensing while many of the functions of each protein are expendable for cytokinesis contractility (Ren et al., 2009). Except for cortexillin-I, other ACLPs that play a major role in cytokinesis and the microtubules may be dispensed for mechanosensing in dividing Dictyostelium cells (Effler et al., 2006). In sum, these observations indicate that the cooperative interaction between myosin II and cortexillin I provide the core of the mechanosensor. Myosin-II mechanochemistry appears to be the essential active component of the mechanosensory module. Regulatory light chain phosphorylation leads to the activation of Dictyostelium myosin-II. Interestingly, this phosphorylation step is essential for mechanosensing (Ren et al., 2009) but is not required for cytokinesis (Beach and Egelhoff, 2009). The lack of a requirement for light chain phosphorylation in cytokinesis is likely due to the fact that RLC phosphorylation only leads to a 3-5fold activation of the actin-activated ATPase activity in Dictyostelium myosin II, and that the myosin II mechanochemistry is not rate limiting for cytokinesis over at least a 30-fold range (Zhang and Robinson, 2005; Chen et al., 1995). By contrast, many

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other myosin II isoforms are activated ∼30-fold by RLC phosphorylation. However, with respect to mechanosensing, this observation suggested that the motor domain of myosin-II was the critical active component of cellular-scale mechanosensing. To test this, the maximum force (Fmax ) of the myosin-II was shifted by altering the lever arm length (Fmax ∝ lever arm length−1 ), which shifted the applied pressuredependency of the mechanosensory response (Uyeda et al., 1996; Ren et al., 2009). Because myosin-II must undergo a full lever arm swing before it can release its ADP, this result strongly indicates that the local accumulation of myosin-II is regulated by the kinetics of its binding/unbinding to actin filaments. It is also likely that the extent of accumulation during mechanosensing is proportional to the magnitude of the applied force as shown in Fig. 2.12 (Ren et al., 2009), which is consistent with catch-bond behaviors of myosin II (Guo and Guilford, 2006). The regulated assembly and disassembly of myosin II into bipolar thick filaments (BTFs) is also required for the mechanosensory system. Without assembling into BTFs, myosin II cannot generate contractile force and therefore experience tension. Because myosin II accumulates during the mechanosensory response, unassembled monomeric myosin II must be able to diffuse to the site, requiring these myosins to be disassembled. Therefore, the full assembly/disassembly dynamic is required for this process. It should be noted that myosin-II BTF assembly in Dictyostelium is independent of myosin-II light chain phosphorylation but is fully dependent on the heavy chain phosphorylation. In contrast, mammalian myosin II assembly is regulated by both heavy chain and regulatory light chain phosphorylation (Beach and Egelhoff, 2009 and references therein). Regulatory light chain phosphorylation results in a conformational change from a thick filament assembly incompetent state to an assembly competent state (the so-called 10S – 6S transition) (Craig et al., 1983). Because Dictyostelium myosin-II does not undergo this transition, it does not require the RhoA-ROCK kinases pathway for regulation of contractility. Consistently, the Dictyostelium genome is devoid of ROCK kinase. Dividing cells have distinctive mechanical properties (the cells soften from anaphase through cytokinesis completion) and the global/polar actin crosslinkers (dynacortin, fimbrin, and enlazin) become more cytoplasmic as compared to interphase cells (Robinson and Spudich, 2000; Reichl et al., 2008). Similarly, wild type cells show a much stronger mechanosensory response during cell division than during interphase. However, the mechanosensitive localization occurs very strongly in interphase RacE null cells (Ren et al., 2009). Unlike RhoA that regulates myosin-II functions, RacE, a Rac-family small GTPase, is known to be upstream of global ACLPs, such as dynacortin, enlazin and fimbrin (Robinson and Spudich, 2000; Zhang and Robinson, 2005). Experimental data shows that the cortical stiffness of RacE null cells is much lower (70% lower) than that of wild type cells, indicating RacE helps to maintain the mechanical integrity of the actin cortex. It is not known yet whether RacE directly inhibits the mechanosensory pathway or if the mechanosensory response only occurs within a certain range of cortical stiffness. The molecular mechanisms for the cooperative accumulation of myosin II and cortexillin I remain to be fully defined. However, there are four processes that may contribute to their accumulation. The first is that forces stabilize the bipolar

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thick filaments in highly deformed region by increasing the binding life time of myosin-actin. This tight binding then promotes additional myosin accumulation through cooperative interactions between motor domains of assembled (in BTF form) and unassembled myosin monomers. The second mechanism is that the binding of cortexillin and the binding of myosin to actin filaments facilitate each other, i.e. cortexillin and myosin II may bind actin cooperatively. A third possible mechanism is that the curvature sensitivity of phosphatidylinositol 4,5-diphosphate (PIP2 ) lipid molecules might lead to PIP2 accumulation in the pipette, resulting in cortexillin accumulation since cortexillin binds PIP2 (Stock et al., 1999). Consequently, the cortexillin accumulation increases the local stiffness of the actin network and enhances the force propagation, which can potentially lead to myosin II accumulation through the first mechanism. Finally, the fourth possible mechanism is that BTF assembly regulatory enzymes, including MHCK and myosin heavy chain phosphatase (which is less well characterized), may be force sensitive. The reduced phosphorylation (by inhibiting MHCK or activating the myosin heavy chain phosphatase) of myosin heavy chains with increasing force could promote local accumulation of the BTFs. However, force-sensitive activation or inhibition of these enzymes may not be essential; rather the enzymes may only be required to maintain the available free pool of myosin monomers, which is essential for the mechanosensory response. These four mechanisms are not mutually exclusive. In the next sections, we will show how force-dependent promotion of BTF assembly and cooperativity between myosin II and cortexillin might lead to the inter-dependent, mechanical stress-induced accumulation of these proteins. 2.3.1.1 How Force Might Modulate Myosin II Bipolar Thick Filament Assembly To consider where force might act in myosin BTF assembly, a sensitivity analysis of the BTF assembly pathway was assessed. The reactions of myosin II assembly and the corresponding reaction rates are listed in Table 2.1. M0 , M, D and T represent phosphorylated (assembly incompetent) monomer, unphosphorylated (assembly competent) monomer, dimer and tetramer, respectively. BTF3 , BTFn and Table 2.1 Kinetics of myosin II assembly based on the dimer addition model. Reproduced from Ren et al. (2009) with permission of Elsevier

M0

k1

k−1

M+M D+D T +D

k1 = 0.0008 s−1 k−1 = 0.1 s−1

M, k2

k−2

T,

k3 = 0.0395 μM−1 s−1 k−3 = 0.0045 s−1

BTF3 ,

k4 = 1.25 μM−1 s−1 k−4 = 0.025 s−1

k3

k−3 k4

k−4

k2 = 0.37 μM−1 s−1 k−2 = 0.01 s−1

D,

BTFn + D

k5

k−5

BTFn+1 ,

k5 = 10 μM−1 s−1 k−5 = 0.2 s−1

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BTFn+1 are the bipolar filaments having 6, 2n and 2(n+1) monomers, respectively. The formation of the anti-parallel tetramer T is the nucleation step, and the subsequent steps are the growth phase of BTF assembly where dimers D are added to the BTF. The reaction rates are derived from a combination of in vitro kinetic studies of BTF assembly as well as in vivo fluorescence recovery after photobleaching experiments (Mahajan and Pardee, 1996; Moores and Spudich, 1998; Reichl et al., 2008; Ren et al., 2009). From this analysis, the most likely process that may be affected is the ratio of k1 to k−1 . Figure 2.13 shows the kinetics of assembly after this ratio is shifted ten-fold, which could mimic inhibition of MHCK, activation of myosin phosphatase, or a lowering of the energy barrier required for incorporating M0 into

Fig. 2.13 Accumulation of myosin II in response to force in Dictyostelium cells during anaphase: (a) spatial distribution of myosin II intensity in pipette region normalized by the intensity at the opposite pole of the cell; (b) averaged transient curve of myosin II accumulation where intensity is normalized by the intensity in the cytoplasm; (c) simulation result of the kinetics of myosin II accumulation when k–1 in Table 2.1 is decreased by 10 fold to mimic the force effect; (d) the BTFs distribution before (without force) and 60 s after 10-fold change in k–1 . Reproduced from Ren et al. (2009) with permission from Elsevier

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a pre-existing BTF. The force dependence of these steps generally can have the form similar to Eq. (2.1). Overall, this analysis suggests that the largest impact on the assembly mechanism might be in the transition from assembly-incompetent to assembly-competent states. Although force might act on MHCK or a myosin heavy chain phosphatase, this scenario requires additional enzymes and steps. A very appealing mechanism is one where motor domains found in pre-existing mini-BTFs are stabilized in the mechanical transition state by force, which then leads to the local accumulation of myosin monomers, putting them in close proximity to the BTF where they can be directly inserted. The motors found in the HMM form of myosin II (dimeric but unassembled myosin monomers; see Fig. 2.2) binds actin in a highly cooperative fashion but only during the transition state of the actomyosin-ADP+Pi complex (Tokuraku et al., 2009) or if the actin structure has been altered such as by being assembled with Ca2+ cations (Ca2+ •ATP-actin – as opposed to Mg2+ •ATP-actin found in cells) (Orlova and Egelman, 1997). This mechanism has the appeal that no additional enzymes are required, and that all of the necessary parts are included directly in the myosin motor and thick filament assembly region. In this model, the MHCK and myosin heavy chain phosphatase are still required to maintain the pool of available myosin monomers M0 and to ensure that the system relaxes back once the mechanical signal subsides. However, more experiments are required to see if this mechanism accounts for mechanosensitive BTF assembly.

2.3.1.2 Cooperativity Between Myosin II and Cortexillin Since myosin II-binding to actin can be highly cooperative depending on the actin conformation and the transition state of the motor-actin complex (as discussed in the previous section), it is very tempting to consider that this cooperativity may be extended to interactions between myosin and actin-associated proteins. For muscle myosin II, the motors bind cooperatively to actin filaments, but only in the presence of tropomyosin-troponin (Geeves and Halsall, 1987; Hill et al., 1980; Chen et al., 2001). On the other hand, the binding of actin-binding proteins (ABPs), such as scruin (Owen and DeRosier, 1994) and formins, induces noticeable structural changes in F-actin. Since F-actin is the common binding substrate of myosin II and ACLPs, it is possible that the binding of myosin and ACLPs to actin are cooperative, i.e. the structural changes in F-actin due to one kind of binding facilitate another kind of binding (Williamson, 2008). Mathematically, the cooperativity between myosin II and cortexillin can be written as M dCC dCM M C C = g (CC ) − kon = h (CM ) − kon . (2.11) and CM − koff CC − koff dt dt where C is the concentration. The superscripts/subscripts M and C represent myosin and cortexillin, respectively. g and h are functions characterizing the cooperativity between the two proteins. Here, only positive cooperativity is considered and

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both g and h always have non-negative values. Furthermore, because of the forceM is assumed to be a function of force, i.e. dependence of myosin-actin binding, koff M (f ) = k0 exp koff off

f x kB T

.

2.3.2 Mechanosensation Through Focal Adhesion Complexes Mechanosensitive behaviors of focal adhesions (FAs) are important in many cellular processes such has cell growth, differentiation and motility. FAs are mechanical linkages between the cytoskeleton and the extracellular matrix (ECM). FAs are large multi-molecular complexes that can extend several-micrometers and consist of a large number of different proteins, including integrins and ABPs such as talin, vinculin, paxillin, and tensin (Zamir and Geiger, 2001; Geiger et al., 2009). In FAs, integrins form heterodimers consisting of α and β subunits non-covalently bound and each consisting of an extracellular domain, a single-pass transmembrane helix, and a short cytoplasmic tail (Puklin-Faucher and Sheetz, 2009). The tail of β-integrin binds to the talin head domain. Talin may then anchor directly to actin or indirectly through vinculin. FAs can be stationary or mobile while displaying a continuous exchange of components with the cytoplasmic pool. FAs grow with increasing local force (Tan et al., 2003) and tend to orient in the direction of applied force (Riveline et al., 2001), which has been attributed to stretching forces, which may enhance the binding affinity of integrins to the ECM (Katsumi et al., 2005). FA formation is initiated with the activation of integrins. Inactive integrins adopt a bent shape whereas the active forms have an extended shape (Hynes, 2002 and references therein). Integrins may be activated either by their head binding to the ECM (so-called outside-in signaling) or by the tail binding to talin (so-called insideout signaling). The activation of the head and the resulting binding to the ECM are thought to occur through long-range conformational changes that propagate through the integrin extracellular domain. Activation of integrin α5 β1 in cells can be switched on mechanically, and the corresponding strength of FAs increases with the rigidity of the extracellular matrix, indicating that the integrin-ECM interaction fits a catch-bond model (Friedland et al., 2009). Single molecule measurements have also shown that the catch-bond behavior of integrin α5 β1 may be attributed to the mechanical activation of the headpiece, but not integrin extension, over a force range of 4–30 pN (Kong et al., 2009). The next mechanosensitive protein in the FA is talin, a large protein consisting of an N-terminal head region and a long rod region. Near talin’s amino-terminus is a FERM (band 4.1, ezrin, radixin, moesin) domain through which talin binds to integrin, focal adhesion kinases (FAKs) and other receptors. Talin-binding to the integrin β tail disrupts an intracellular salt bridge between the α- and β-integrin subunits, increasing the integrin affinity for ECM (Tadokoro et al., 2003). Additionally, talin has eleven vinculin binding sites (VBSs) in its rod region. Single molecule measurements discovered that only one VBS is active in the absence of force and

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two more VBSs appear when the talin rod is stretched by relatively low (12 pN) forces (del Rio et al., 2009). In the absence of force, these two force-sensitive VBSs are thought to remain buried in adjacent amphipathic helices through hydrophobic interactions. Talin then links to vinculin, a 116 kDa actin-binding protein, which links the core FA proteins to the actin cytoskeleton. The vinculin head domain consists of seven α-helices arranged as two four-helical bundles (eight α-helices), and its tail domain has five α-helices that form an anti-parallel bundle. The strong interaction between the head and the tail masks the binding sites for other proteins such as talin, F-actin, α-actinin and paxillin and keeps vinculin in its inactive states. Upon talin-binding to α-actinin, the tail domain of vinculin is displaced away from the head domain, which activates vinculin (Izard et al., 2004) and enables its binding to F-actin and other molecules. The mechanical stretching of FAs triggers many downstream signaling pathways by activating the SH2 domain-containing phosphatase SHP-2 and non-receptor protein tyrosine kinases, such as Src and FAK (Tamada et al., 2004; Giannone and Sheetz, 2006). These enzymes regulate the assembly/disassembly of FAs by controlling the actin stress fiber (actin bundles with myosin II thick filaments) formation (Vicente-Manzanares et al., 2009). Activity of some of these kinases, such as FAK kinase, requires actin and myosin II-dependent tension (Tilghman and Parsons, 2008). Furthermore, stretching of p120Cas in cells or using an in vitro reconstitution system exposes more Src kinase binding sites and leads to its local activation (Sawada et al., 2006). This stretch-induced activation can be very fast as Src at remote sites may be activated within 0.3 s, demonstrating just how fast signals can propagate through the elastic cytoskeleton (Na et al., 2008).

2.3.3 The Actin Cytoskeleton Works as a Force-Transmission Highway Both chemical and mechanical signals can be transmitted over long distances. Propagation of chemical signals occurs mainly through the diffusion of molecules in the cytosol and its speed is limited by the chemical reaction (such as phosphorylation) rates, unbinding/binding rates and diffusion rates. Diffusion is usually the limiting step since the diffusion coefficient of molecules in cells is in the range of 0.01–100 μm2 s−1 , depending on the molecular size and shape and the viscosity of cytoplasm on the length-scale of the diffusing particle (Howard, 2001). For example, it can take one molecule 1–100 s to travel 10-μm using diffusion alone. On the other hand, mechanical signals may be transmitted through the deformation of cytoskeleton along actin filaments, microtubules and intermediate filaments. The speed of transmission depends on the elastic modulus of the cytoskeleton where signals may propagate over a 10-μm distance on sub-second time-scales, indicating that proteins in cells can sense mechanical stimuli much more rapidly than chemical signals (Forgacs, 1995). This propagation of mechanical deformations likely

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depends on the pre-stress as well as the elastic modulus of the cytoskeleton, and the magnitude of deformation decays exponentially in space with a characteristic length that is comparable to or larger than the size of the cell (Wang and Suo, 2005). One consequence of signal propagation through the integrated elastic actin cytoskeleton is that signals can be transmitted over long distances and broad areas and to a range of organelles. For example, actin filaments are connected to the nuclear envelope through a complex of SUN and nesprin proteins (Wang et al., 2009), to the plasma membrane through proteins such as ezrin, radixin, and moesin (ERM proteins) (Sato et al., 1992), to stretch-activated channels (SACs) by myosin I motor proteins (Fettiplace and Hackney, 2006), and to mitochondria by mitochondrial ABPs (Boldogh et al., 1998). Forces transmitted by the actin cytoskeleton to the nucleus alter gene expression, which may in turn regulate actin remodeling. The SACs can also be activated or deactivated by cytoskeleton stretching, resulting in ion flux, regulating stress fiber formation and orientation and myosin II bipolar thick filament assembly. Mechanical stimuli propagated to membranebound or associated proteins through the actin-membrane connections can lead to changes in activity of membrane-bound signaling molecules and other ion channels.

2.4 Remodeling of the Actin Cytoskeleton During Mechanosensation The actin cytoskeleton is composed of highly dynamic structures. Besides mechanosensing and transmitting mechanical signals, the cytoskeleton can rearrange its structures in response to the mechanical stimuli – this is referred to as remodeling. Actin remodeling is determined by ABPs that control linear elongation, shortening and organization of actin filaments in response to signaling cascades (Stossel et al., 2006). However, superimposed over these mechanosensitive actin-binding proteins are signaling molecules, such as kinases, Rho-family GTPases (e.g. RhoA, Cdc42 and Rac), phosphoinositides and WASP-family proteins, which are also spatially and temporally coordinated biochemically and, indirectly, mechanically. Most Rho proteins switch between active (GTP-bound) and inactive (GDPbound) conformations. The activities of Rho proteins are regulated by Rho guanine nucleotide exchange factors (Rho-GEFs) and Rho GTPase-activating proteins (Rho-GAPs). The Rho-GEF promotes the exchange of GDP for GTP while the Rho-GAPs enhance GTP hydrolysis. Rho guanine-nucleotide dissociation inhibitors (GDIs) also bind to prenylated GDP-bound Rho proteins and allow their translocation between membrane and the cytosol (Buchsbaum, 2007). RhoA and its effector Rho-kinase elevate myosin II light chain phosphorylation and thereby promote myosin II activation. Cdc42 activates WASP, which subsequently mediates the branched actin-network formation by activating Arp2/3 (Pollard, 2007). PIP2 can induce G-actin dissociation from actin-monomer-binding proteins and

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uncapping of the actin filament barbed ends, and can enhance the linkages between the actin cytoskeleton and the plasma membrane by activating ERM proteins (Nebl et al., 2000). PIP2 can also activate vinculin, promoting FA assembly (Sechi and Wehland, 2000), and WASP, increasing actin polymerization (Pollard, 2007).

2.4.1 How Mechanically Activated Kinases Regulate the Actin Cytoskeleton Mechanically induced FAs trigger the activation and recruitment of many downstream kinases (Brakebusch and Fässler, 2003), and these kinases affect actin remodeling by regulating small GTPases and ABPs. Focal adhesion kinase binds to integrin, talin and paxillin, and this binding enhances FAK activities, which promotes stress fiber formation by increasing the recruitment of talin and paxillin. Integrin-linked kinase (ILK) binds the tails of β integrin subunits, paxillin and phospholipids, which induces the phosphorylation of PKB/AKT protein kinases that are upstream of actin polymerization. ILK also forms a complex with other proteins to recruit F-actin to FAs. The FAK-Src complex stimulates Rac1 activity, recruits the GEF for Cdc42 and Rac1, and mediates the suppression of Rho-GTP by regulating Rho-GEFs and Rho-GAPs (Huveneers and Danen, 2009 and references therein).

2.4.2 Crosstalk Between Microtubules and Actin Cytoskeleton Increasing evidence shows crosstalk between microtubules and actin cytoskeleton. For example, centrosome separation and positioning during mitosis depend on the integrity of the actin cytoskeleton and F-actin cortical flow (Rosenblatt et al., 2004). By contrast, actin nucleation near the plasma membrane is coordinated by microtubules and microtubule-associated proteins (Martin et al., 2005; Siegrist and Doe, 2007; Rosales-Nieves et al., 2006). Similar to the concept of the tensegrity model, cortical motors may pull on astral microtubules and conversely, astral microtubule polymerization exerts a pushing force against the actin cortex, promoting centrosome separation and positioning. The activation of Src was observed at cortical sites where microtubules appear to deform the cortex (Na et al., 2008). Microtubules also affect the spatial distribution of active small GTPases, thereby regulating the organization of the actin cortex (Siegrist and Doe, 2007 and references therein), and many unproven mechanisms have been proposed for this crosstalk. However, the list of structural linkages between microtubules and F-actin continues to grow. Among the first identified linkages was the splice-variant of the mitotic kinesin-like protein (MKLP1) called CHO1. CHO1 has a kinesin-family motor domain, which can move on microtubules, and an additional microtubule-binding domain and an F-actin binding domain in its tail (Robinson and Spudich, 2004). Indeed, MKLP1 proteins help organize the central spindle microtubules, and CHO1 can integrate this

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system with the cortical actin network. In Drosophila melanogaster, cappuchino (an FH2 containing protein) and spire (a WH2 containing protein) can crosslink microtubules and actin filaments (Rosales-Nieves et al., 2006). In Schizosaccharomyces pombe, tea4p and tea1p also localize to the plus ends of microtubules, and a complex of tea1p, tea4p and the formin for3p is necessary for the establishment of cell polarity and actin nucleation at new cell ends (Martin et al., 2005). Thus, several linkages between the actin and microtubule networks promote their integration and may facilitate force-propagation, and therefore signal-propagation, through these systems.

2.5 Experimental Techniques for Measuring Mechanosensation – In Vitro and In Vivo Methods In addition to traditional micropipette aspiration (MPA), the past few decades have witnessed the development of various new techniques using the combinations of nanomanipulation, microfabrication, magnetic techniques, and optical techniques (Bao and Suresh, 2003; Addae-Mensah and Wikswo, 2008). These methods include atomic force microscopy (AFM), magnetic tweezers, optical tweezers, magnetic twisting cytometry (MTC), particle-tracking microrheology (PTM), microfluidic devices, stretching devices, traction force microscopy (TFM) and MEMS-based devices. Classic MPA still offers a number of advantages in that it is relatively easy to implement and can be readily adapted to a broad array of cell-types, particularly those which are not highly adherent. MPA can be used to measure effective tension and elastic and viscous properties of the cell. Perhaps more significantly, MPA is very useful for imposing deformations to cells so that the cell’s response may be monitored using an array of fluorescence methods. However, the major limitation of MPA is that it is difficult to measure properties occurring on fast sub-second timescales or to measure frequency-dependent features. For these sorts of measurements, many of the other methods, for example AFM, MTC and PTM, are more suitable (Girard et al., 2004; Hoffman et al., 2006). PTM can be categorized into singlebead, two-bead and multiple-bead modes (Wirtz, 2009). Depending on the driving force, PTM has two working modes: passive and active. To measure the mechanical properties of single molecules, AFM is commonly used for the high force ranges (>10 pN) whereas optical tweezers and magnetic tweezers are commonly used for relatively low forces (Neuman and Nagy, 2008; Finer et al., 1994). AFM and optical tweezers also allow three-dimensional manipulation of molecules. AFM is often used to study protein folding/unfolding and protein-protein interactions while tweezers are usually used to study biological motors, including cytoskeletal motors and DNA and RNA polymerases. Additionally, many of these methods have been combined with fluorescence microscopy techniques, such as total internal reflection fluorescence (TIRF) imaging and fluorescence resonance energy transfer (FRET) (Sarkar et al., 2004; Moffitt et al., 2008; del Rio et al., 2009). Microfluidic devices and flow chambers are used to study the cell responses to shear flow (del Álamo et al., 2008; Wang and Levchenko, 2009). Stretching devices

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are usually coupled with fluorescence imaging methods to quantify the effects of stretch on actin cytoskeleton rearrangement, activation of kinases, gene expression and cell differentiation (Sawada et al., 2006; Kurpinski et al., 2006). TFM was invented to investigate the traction force that cells apply to the substrates (Pelham and Wang, 1997). Initially, TFM methods utilized fluorescent beads embedded in polymeric substrates so that traction forces exerted by adherent cells on the substrate could be calculated from the displacements of the beads. Recently, micropatterned substrates by soft photolithography have been used to control the cell adhesion areas and cell shapes (Balaban et al., 2001; Tan et al., 2003; Théry and Bornens, 2006). Therefore, a broad range of mechanical measurements and manipulations are now possible across a broad array of length- and time-scales and from the molecule to cellular levels.

2.6 Conclusion and Perspectives Mechanical inputs must have been among the first signals that cells received and had to respond to, and the ability of cells to sense and react to these inputs likely evolved at a very early time point. Thus, it is not surprising that there are numerous overlaps between the mechanotransduction and “traditional” chemical signal transduction pathways. Because of the overlap between these pathways, mechanical-chemical coupling and feedback loops are a natural consequence of this system integration. Because the actin cytoskeleton is structurally integrated with nearly every aspect of the cell, mechanical inputs can be transmitted quickly throughout the cell. Furthermore, individual proteins may be involved in multiple pathways and contributing multiple functions. Therefore, to fully understand the roles of individual proteins and the cooperativity among them in the actin cytoskeleton, in vitro experiments involving single molecule measurements, reconstituted actin networks, and computational simulations of protein folding/unfolding and protein-protein interactions have to be combined with quantitative in vivo observations. Challenges for understanding mechanosensation through the actin cytoskeleton include revealing how proteins function cooperatively over short nanometer-length-scales and fast sub-second time-scales. Direct observations of force propagation in cells and eventually between cells within tissues will be essential. Novel designs of mechanical strain sensors using fluorescence readouts such as FRET pairs and inventive imaging setups will be needed to fulfill these demands. Applying a repertoire of these approaches to a genetically tractable organism, such as Dictyostelium cells as they perform physiologically and medically important processes like cell division, will continue to provide unique insights into cellular mechanosensing and mechanotransduction. Acknowledgements We are grateful to the insightful discussions and comments on the manuscript from Alexandra Surcel and Sheil Kee. We acknowledge the support of the National Institute of Health (Grant #GM066817) and the American Cancer Society (Grant #RSG CCG114122).

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Glossary List Affine Non-affine Stress Strain ABD ABP ACLP BTF ELC FJC HMM RLC WLC D E fb G∗ G G K Lc Lp Le ξ ς

Describes an actin network, which is co-linear during stretching and shearing Describes an actin network in which co-linearity is absent during stretching and shearing The force applied on a unit area The ratio between the length-change associated with deformation and the original length (no deformation) Actin-binding domain Actin-binding protein Actin-crosslinking protein Bipolar thick filament Essential light chain Free-joint-chain Heavy meromyosin Regulatory light chain Worm-like-chain Bundle size Young’s modulus: the proportionality between stress and the resulting strain Bending modulus: the proportionality between bending momentum and the resulting curvature Complex modulus Shear modulus – the real part of G∗ : the proportionality between shear stress and the shear strain Loss modulus – the imaginary part of G∗ Differential shear modulus: the differential proportionality between shear stress and the shear strain Contour length: the integrated length along the polymer chain Persistence length: the length over which correlations in the direction of the tangent are lost Distance between entanglements Mesh size Crosslinking density: the ratio between the cross-linker concentration and actin concentration

References Ackbarow T, Chen X, Keten S, Buehler MJ (2007) Hierarchies, multiple energy barriers, and robustness govern the fracture mechanics of α-helical and β-sheet protein domains. Proc Natl Acad Sci USA 104:16410–16415

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Chapter 3

Effect of Cytoskeleton on the Mechanosensitivity of Genes in Osteoblasts Qiang Fu, Yiping Zhang, Yajuan Xu, Yourui Li, Ling Guo, and Minfeng Shao

Abstract Mechanosensitivity is the ability of tissues and cells to detect and make response to mechanical stimuli. Osteoblast is a kind of important mechanosensitive cell in bone tissue, while cytoskeleton plays an important role in the mechanotransduction in osteoblasts. This article reviewed the roles of cytoskeleton on the mechanotransduction of genes in osteoblasts, summarized that cytoskeleton integrity is essential for the expression of bone formation–related genes in osteoblasts, and concluded that cytoskeleton reorganization inhibition can enhance the mechanosensitivity of some genes in osteoblasts. Further researches on the specific mechanisms of cytoskeleton will shed new light on the transmission mechanisms of mechanical stress in osteoblasts, and will lay the foundation for the further investigations on the biomechanical behaviors of bone cells and even bone tissues. Keywords Cytoskeleton · Osteoblast · Mechanotransduction · Mechanosensitivity

3.1 Introduction Cytoskeleton is the protein fiber network system in eukaryotic cells. The general concept of cytoskeleton includes nucleus cytoskeleton (intranuclear cytoskeleton, nuclear lamina, and chromosome cytoskeleton in mitotic state), cytoplasm cytoskeleton (microfilament, microtubule, intermediate fibre, and microtrabecular lattice), cell membrane cytoskeleton and extracellular matrix. The narrow sense of cytoskeleton only refers to the cytoplasm cytoskeleton. Cytoplasm cytoskeleton, which is a fibrous structure system mainly distributed in the eukaryotic cells, consists of three basic filaments: microtubule, microfilament and intermediate fibre. Q. Fu (B) Department of Prosthodontics, Guanghua School & Hospital of Stomatology, Sun Yat-Sen University, Guangzhou 510055, Guangdong, China e-mail: [email protected]; [email protected]


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These three filaments are polymerized from different protein subunits (skelemin) in different specific ways (Qian et al., 2009). The cytoskeleton we reported in this review is mainly about the cytoplasm cytoskeleton.

3.2 Characteristics and Roles of Cytoskeleton 3.2.1 Characteristics of Cytoskeleton One of the greatest characteristics of cytoskeleton is its dynamics variance which occurs in a short time by accommodating to the structure and function of cells. Dynamics variance is based on the continual polymerization of skelemin, which prolongs the filaments; or on the continual depolymerization of skelemin, which decurtates the filaments, and even clears it away. So, the cytoskeleton in cells is in a condition of continual dynamic reorganization. The other characteristic of cytoskeleton is its connection with cell nucleus, cell membrane and some other organelles, and such the connection is reversible due to the dynamics variance of cytoskeleton itself. In this connected network, the cytoskeleton system is the main fiber network, around which some other cellular structures and cytoplasmic matrix of biomacromolecules are cohered and embedded. The dynamic variance of cytoskeleton also endows the cytoplasmic matrix with the same characteristic.

3.2.2 Roles of Cytoskeleton The cytoskeleton plays a crucial role not only in maintaining the cellular morphology, bearing loads and keeping the intracellular structures in order, but also in lots of life activities, for examples: guiding the chromosome segregation during cell division, transporting kinds of vesicles and organelles along it in a directional transport. Recently, the increasing evidences indicate that several cellular activities, including activation of several signal transduction pathways and transcriptional activity of many genes, are based on the reorganization of the cytoskeleton. The cytoskeleton has the function of transferring and dispersing stress, as well as responding to the mechanical signals (Higuchi et al., 2009). 3.2.2.1 Tensegrity Model The cytoskeleton, as the core of the ECM-Integrin-CSK-Nucleus network system, transmits extracellular signals into cells in two different ways: One is that mechanical stress induces the redistribution of the tension in cytoskeleton, and then the rearrangement of cytoskeleton and morphological change of cells. The role of the cytoskeleton in this way is mainly the physical conduction. The other is that the cytoskeleton, entirety or partly as a translator of mechanical-chemical signals,

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makes come true the transformation from mechanical signals to biological signals and chemical signals. The transmission and transduction of mechanical signals can be completed by the cooperation of these two ways (Charras and Horton, 2002). The function of cytoskeleton, directly as the conductive track to transmit mechanical signals into cells, can be explained by the tensegrity model. In this model, actin microfilaments are the major part to bear the tension, meanwhile, microtubules can be viewed as the load-bearing element for compression, and intermediate fibres, as the potent integrated components, connect the microtubules with the inotropic microfilaments and fix them to the membrane surface and the nucleus. If the conducting system is based on the tensegrity, it must be instant for the mechanical signals to be transmitted into cells, and faster than any signal conducting system based on diffusion, which enhances the conducting efficiency of the mechanical signals (Ingber, 2003). Mechanical stimulus can be transmitted to cytoskeleton through integrin-microfilament pathway and cadherins-microtubule pathway, which can change the distribution and polymerization of cytoskeleton, increase the permeability of ion channels coupled with cytoskeleton and the activity of some surface receptors, and alter the function of the proteins and functional enzymes in many signal conducting systems coupled with cytoskeleton, then produces the second messengers and transmits the signals to nucleus, and finally promotes the metabolic activities of the proliferation and differentiation of cells and the synthesis and secretion of proteins (Sawada and Sheetz, 2002). The extracellular signals can be transmitted to cells through this continuum structure, vice versa. Such is the major way for microfilaments to participate in signal transduction (Chou et al., 2001).

3.2.2.2 Response of Cytoskeleton to Mechanical Forces The mechanical signals transmitted to cytoskeleton could cause the rearrangement of microfilaments and microtubules, as well as the change of orientation of cells. The reorganization of cytoskeleton is good for the extracellular signals to be transmitted into cells, and then the signals spread along the cytoskeleton, which induces the gene transcription and the change in cell cycle and cell shape. The response of the cells to mechanical signals is mainly based on the F-actin cytoskeleton. Studies have indicated that the cytoskeleton integrity plays a crucial role in the expression of c-fos and cox-2 genes in osteoblasts induced by fluid shear stress (Chen and Fu, 2008; Wu and Fu, 2009).

3.2.2.3 Cytoskeleton and Genes’ Transcription and Expression The cytoskeleton not only participates in the mechanical signal-chemical signals transduction, but also affects the genes expression in nucleus. Chen et al. (2000) have confirmed that the increasing of the expression and synthesis of c-fos and cox-2 genes induced by fluid shear force needs the recombination of actin into stress fibers. The structural change of cytoskeleton could directly activate NF-KB, which greatly enhances the binding activity of NF-KB with DNA, and then affects the expression

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of nuclear genes. Now this function is thought to be determined and performed mainly by the state of microtubules. Concerning the mechanism of how the mechanical agents affect the gene expression through cytoskeleton, Ingber (2003) reported that the distortion of cytoskeleton could expose some genes around the nucleus, which made them much easier to be identified and activated. Stein et al. (1999) thought that the rearrangement of cytoskeleton induced by mechanical stress may stretch the promoter through karyoskeleton, change its spatial conformation, and then regulate its transcription activity, which may be one of the pathways for microgravity to affect the genes expression. Lammerding et al. (2004) have observed that gene mutations of nuclear lamina would change the mechanical properties of nuclear lamina and the gene expression induced by tension force. When some intranuclear genes are activated by the mechanical signals, the immediate early genes will respond rapidly, and the mRNA localization of these immediate early genes is related to the cytoskeleton. The c-fos mRNAs of most interphase cells are located in cytoskeleton, but when the microfilaments are depolymerized by Cytochalasin D, most mRNAs will be transferred into the endochylema. The actin microfilaments and microtubules in cytoskeleton also participate in the RNA transfer; the former in short distance transportation, while the latter in long distance. And both of them have their own corresponding molecular motors. While the intermediate fibres are tied up with nuclear matrix, their roles in intranuclear signal transduction and genetic transcription should not be ignored (Guzman et al., 2006).

3.3 Effect of Cytoskeleton on the Genes Expression in Osteoblasts Bone formation-related genes, which could promote the proliferation and activation of osteoblasts, include c-fos, early growth response gene-1 (egr-1), osteopontin (OPN), cyclooxygenase-2 (COX-2), extracellular signal-regulated kinase (ERK), and so on. When these genes are activated, some bone formation-related proteins are expressed, and then the proliferation and differentiation of osteoblasts and new bone formation are promoted (Kapur et al., 2004; Lee et al., 2008).

3.3.1 C-fos Gene C-fos gene, as an important member of the immediate-early gene (IEG) family, can respond to external stimuli in a few minutes (Lau and Nathans, 1987). C-Fos protein, as the product of c-fos gene, can combine with c-Jun through leucine zipper to form the activator protein-1 (AP-1) heterodimer. AP-1, as an important transcription factor, which has the DNA-binding activity, can combine with the AP-1 binding sites of the DNA sequences of downstream target genes, and then activate the transcription

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activity of these genes. AP-1 plays a significant role both in the proliferation and differentiation of osteoblasts and the promotion of bone fracture healing (Cowles et al., 2000; Zayzafoon et al., 2005). The DNA sequences of the important bone formation-related genes that are known so far, such as OPN (Nomura and TakanoYamamoto, 2000), COX-2 (Ogasawara et al., 2001) and so on, have AP-1 binding sites. That means, the expression of these genes would be regulated by c-fos gene. Studies have found that tensile stress and compressive stress could activate c-fos gene expression of osteoblasts, but when the cytoskeleton is destructed by Cytochalasin D, the cytoskeleton would depolymerize, which could interrupt the c-fos gene expression of osteoblasts induced by tensile stress and compressive stress (Nomura and Takano-Yamamoto, 2000; Ogasawara et al., 2001). Our previous work has showed that Cytochalasin D used to destruct the cytoskeleton could also interrupt the induction of c-fos gene in osteoblasts under fluid shear stress (Chen and Fu, 2008).

3.3.2 Egr-1 Egr-1, also a member of the immediate-early gene family, can be activated by a kind of stimulus such as mechanical stress and then express the corresponding protein quickly. Egr-1, which is fairly conservative in evolution, generally exists in the eukaryocytes from yeast to human. Egr-1 protein, with 533 amino acids, is a karyo-phosphorylated protein of 80 kD and a transcription factor. Egr-1 has three functional areas in structure (Thiel and Cibelli, 2002): the N-terminal end as the transcriptional activation domain, the structural domain with three duplicate zinc fingers as the central zone, which has the characteristics of combination with DNA, and the area between the N-terminal end and the zinc fingers as the transcription inhibition zone. McMahon et al. (1990) indicated that egr-1 protein regulates the transcription of the target gene by combining with the specific sequence of the gene, and then promotes the cell proliferation. High level expression of egr-1 could be detected in places including periosteum during the bone formation (Chen and Fu, 2008). Moalli et al. (2000) showed that mechanical stress could induce the gene expressions of egr-1 and c-fos, meanwhile, could enhance the cytoactive of osteoblasts, which comes to the conclusion that egr-1 and c-fos play an important role in enhancing the cytoactive of osteoblasts. Ogata (1997) have reported that fluid shear stress could induce the expression of egr-1, but Cytochalasin D could interrupt this induction (the expression of egr-1 was reduced 40%).

3.3.3 OPN OPN, a non-collagenous bone matrix protein, is one of the phenotypes of osteoblasts. In different species, the sizes and forms of OPN are not exactly the same, but they all have the same specific conserved sequences: signal peptide

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sequences, seven to ten consecutive Asp sequences, RGD sequences and tyrosine kinase-αphosphorylation recognition sequences. The RGD (arginine-glycineaspartic acid) sequence, a special structure of OPN, could combine with integrin, and then facilitate the bone cells to adhere to the extracellular mineralized matrix. So, it mediates lots of significant physiological functions. Many studies showed that mechanical stress could induce the expression of OPN in osteoblasts (Toma et al., 1997). Morinobu et al. (2003) have established an osteogenesis model induced by stress to observe the role of OPN in bone formation with stress. They found that the osteogenesis under stress was obvious in wild-type mouse, and the expression of OPN protein was significant in new bone formation area, but not in OPN knock-out mice. Fujihara et al. (2006) found that bone remodeling induced by mechanical stress was significantly inhibited in OPN knock-out mice, which suggested that OPN also plays an important role in bone remodeling. But other studies showed that OPN is also an important regulatory factor in bone loss (disuse atrophy) (Ishijima et al., 2006). So, OPN may play important roles in both bone formation and bone resorption. After studying the OPN signaling pathway induced by mechanical stress, both Toma et al. (1997) and Carvalho et al. (2002) have found that the cytoskeleton destruction with Cytochalasin D could interrupt the induction of OPN under mechanical stress.

3.3.4 ERK ERK, as a member of the mitogen-activated protein kinases (MAPKs) family, with a wide catalytic activity, could phosphorylate a series of plasmosins, membrane proteins, intranuclear transcription factors and transcription regulons. ERK is also considered as the upstream molecule that cells make response to external signals, and as the junction of many kinds of important signal conduction pathways. Present studies suggested that ERK can be quickly activated by mechanical stress (Liu et al., 2006; Li et al., 2007; Wadhwa et al., 2002; Kapur et al., 2003). Many bone formation-related genes as we know, such as c-fos, COX-2, OPN and egr-1, are regulated by ERK, because the genes expression could be interrupted when the activation of ERK is inhibited (Kapur et al., 2004, 2003). Meanwhile, ERK plays an important role in the proliferation and differentiation of osteoblasts promoted by mechanical forces (Lee et al., 2008; Kapur et al., 2003). Liu et al. (2006) indicated that tensile stress and compressive stress can activate ERK, but when the cytoskeleton was destructed by Cytochalasin D, the activation would be interrupted.

3.3.5 COX-2 Cyclooxygenase, also named prostaglandin synthetase, could oxidize the arachidonic acid into prostaglandin E2 (PGE2). PGE2 could increase the contents of

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calcified tissue in bone, accelerate fracture healing, enhance the proliferation and differentiation of osteoblasts (Norrdin and Shih, 1988), and also mobilize the precursor cells of osteoblasts to differentiate into osteoblasts (Suponitzky and Weinreb, 1998). Cyclooxygenase includes two isoenzyme types: Conservative COX (COX-1) and induction COX (COX-2). COX-1 is widely distributed in various tissues and is maintained at a stable level under physiological conditions to regulate the physiological function of organisms, while COX-2, as an inducible enzyme, which can be induced by mechanical stress, is a rate-limiting enzyme for PGE2 synthesis under mechanical stress. Studies showed that COX-2 plays a significant role in the PGE2 expression induced by fluid shear stress (Bakker et al., 2003). Fluid shear stress could induce the expression of COX-2 and the release of PGE2 in osteoblasts, and both of them are necessary for mechanical stress to stimulate bone formation (Bakker et al., 2001; Klein-Nulend et al., 1997; Reich et al., 1997; Smalt et al., 1997; Chow and Chambers, 1994). The selective inhibition of COX-2 gene expression could interrupt the bone formation in vivo induced by mechanical stress (Chow and Chambers, 1994; Forwood, 1996). FU et al. had indicated that cytoskeleton integrity plays an important role in the expression of cox-2 gene in osteoblasts induced by fluid shear stress (Wu and Fu, 2009). To sum up, cytoskeleton integrity is necessary for the expressions of bone formation-related genes (such as c-fos, egr-1, OPN, ERK, COX-2, and so on) induced by fluid shear stress.

3.4 Cytoskeleton Reorganization Inhibition Enhances the Mechanosensitivity of Some Genes in Osteoblasts Mechanosensitivity refers to the ability that tissues or cells detect and make response to mechanical stimuli. Many studies showed that the response of bone cells to external stress could be changed (Turner, 1999; Schriefer et al., 2005; Jaasma et al., 2007). Cytoskeleton plays an important role in the biotransformation of mechanical signals transmitted to bone tissues. The previous work of Fu et al. has proved that LIMK2 gene plays a crucial role in cytoskeleton reorganization of osteoblasts induced by fluid shear stress (Fu et al., 2008; Fig. 3.1). Presently, Fu et al. have studied the effects of cytoskeleton reorganization inhibition on c-fos and cox-2 genes in osteoblasts induced by fluid shear stress. They found that cytoskeleton reorganization inhibition with RNAi (RNA interference) could enhance the expression of c-fos and cox-2 genes induced by fluid shear stress (Chen and Fu, 2008; Wu and Fu, 2009). Thus, Fu et al. concluded that on condition of maintaining the cytoskeleton integrity, cytoskeleton reorganization inhibition can promote the expression of c-fos and cox-2 genes in osteoblasts induced by fluid shear stress. In other words, cytoskeleton reorganization inhibition can enhance the mechanosensitivity of c-fos and cox-2 genes in osteoblasts.

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Fig. 3.1 Cytoskeleton immunofluorescence staining of 1–4 groups of osteoblasts (×200). F-actin was stained using FITC-phalloidine (green fluorescence), and the nucleus was stained using DAPI (blue fluorescence) (Fu Q et al., 2008). The cells were taken photographs in the same condition. As shown in the pictures, compared with the negative control (Group1), when LIMK2 gene was interfered with siRNA, cytoskeleton reorganization of osteoblasts was significantly inhibited in loading groups, and the F-actin became disordered and was in filamentary arrangement (Group 2)

3.5 Conclusion and Perspectives About the mechanisms that mechanical signals can be transformed into biological signals and chemical signals, and then induce the expression of some bone formation-related genes when osteoblasts are loaded by mechanical stress, it is not clear until now. For the c-fos and cox-2 genes, Fu et al. have indicated that cytoskeleton reorganization inhibition could enhance their mechanosensitivity. But for the other genes of osteoblasts, similar investigations should be conducted in the future. Further researches on the specific mechanisms of cytoskeleton will shed new light on the transmission mechanisms of mechanical stress in osteoblasts, and will lay the foundation for the further investigations on the biomechanical behaviors of bone cells and even bone tissues. Acknowledgments This work was supported by the Science and Technology Projects of Guangdong Province (No. 2008B030301121 and 2009B050700027), China.

References Bakker AD, Klein-Nulend J, Burger EH (2003) Mechanotransduction in bone cells proceeds via activation of COX-2, but not COX-1. Biochem Biophys Res Commun 305:677–683 Bakker AD, Soejima K, Klein-Nulend J, Burger EH (2001) The production of nitric oxide and prostaglandin E(2) by primary bone cells is shear stress dependent. J Biomech 34:671–677

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Carvalho RS, Bumann A, Schaffer JL, Gerstenfeld LC. (2002) Predominant integrin ligands expressed by osteoblasts show preferential regulation in response to both cell adhesion and mechanical perturbation. J Cell Biochem 84:497–508 Charras GT, Horton MA (2002) Single cell mechanotransduction and its modulation analyzed by atomic force microscope indentation. Biophys J 82:2970–2981 Chen NX, Ryder KD, Pavalko FM, Turner CH, Burr DB, Qiu J, Duncan RL (2000) Ca(2+) regulates fluid shear-induced cytoskeletal reorganization and gene expression in osteoblasts. Am J Physiol Cell Physiol 278:C989–C997 Chen R, Fu Q (advisor) (2008) Effects of cytoskeleton reorganization inhibition on the expression of c-fos in osteoblasts induced by fluid shear stress. Master’s Thesis, Sun Yat-Sen University, Guangzhou Chou YH, Helfand BT, Goldman RD (2001) New horizons in cytoskeletal dynamics: transport of intermediate filaments along microtubule tracks. Curr Opin Cell Biol 13:106–109 Chow JW, Chambers TJ (1994) Indomethacin has distinct early and late actions on bone formation induced by mechanical stimulation. Am J Physiol 267:E287–E292 Cowles EA, Brailey LL, Gronowicz GA (2000) Integrin-mediated signaling regulates AP-1 transcription factors and proliferation in osteoblasts. J Biomed Mater Res 52:725–737 Forwood MR (1996) Inducible cyclo-oxygenase (COX-2) mediates the induction of bone formation by mechanical loading in vivo. J Bone Miner Res 11:1688–1693 Fu Q, Wu C, Shen Y, Zheng S, Chen R (2008) Effect of LIMK2 RNAi on reorganization of the actin cytoskeleton in osteoblasts induced by fluid shear stress. J Biomech 41:3225–3228 Fujihara S, Yokozeki M, Oba Y, Higashibata Y, Nomura S, Moriyama K (2006) Function and regulation of osteopontin in response to mechanical stress. J Bone Miner Res 21:956–964 Guzman C, Jeney S, Kreplak L, Kasas S, Kulik AJ, Aebi U, Forro L (2006) Exploring the mechanical properties of single vimentin intermediate filaments by atomic force microscopy. J Mol Biol 360:623–630 Higuchi C, Nakamura N, Yoshikawa H, Itoh K (2009) Transient dynamic actin cytoskeletal change stimulates the osteoblastic differentiation. J Bone Miner Metab 27:158–167 Ingber DE (2003) Tensegrity II. How structural networks influence cellular information processing networks. J Cell Sci 116:1397–1408 Ishijima M, Ezura Y, Tsuji K, Rittling SR, Kurosawa H, Denhardt DT, Emi M, Nifuji A, Noda M (2006) Osteopontin is associated with nuclear factor kappaB gene expression during tail-suspension-induced bone loss. Exp Cell Res 312:3075–3083 Jaasma MJ, Jackson WM, Tang RY, Keaveny TM (2007) Adaptation of cellular mechanical behavior to mechanical loading for osteoblastic cells. J Biomech 40(9):1938–1945 Kapur S, Baylink DJ, Lau KH (2003) Fluid flow shear stress stimulates human osteoblast proliferation and differentiation through multiple interacting and competing signal transduction pathways. Bone 32:241–251 Kapur S, Chen ST, Baylink DJ, Lau KH (2004) Extracellular signal-regulated kinase-1 and -2 are both essential for the shear stress-induced human osteoblast proliferation. Bone 35:525–534 Klein-Nulend J, Burger EH, Semeins CM, Raisz LG, Pilbeam CC (1997) Pulsating fluid flow stimulates prostaglandin release and inducible prostaglandin G/H synthase mRNA expression in primary mouse bone cells. J Bone Miner Res 12:45–51 Lammerding J, Kamm RD, Lee RT (2004). Mechanotransduction in cardiac myocytes. Ann NY Acad Sci 1015:53–70 Lau LF, Nathans D (1987) Expression of a set of growth-related immediate early genes in BALB/c 3T3 cells: coordinate regulation with c-fos or c-myc. Proc Natl Acad Sci USA 84: 1182–1186 Lee DY, Yeh CR, Chang SF, Lee PL, Chien S, Cheng CK, Chiu JJ (2008) Integrin-mediated expression of bone formation-related genes in osteoblast-like cells in response to fluid shear stress: roles of extracellular matrix, Shc, and mitogen-activated protein kinase. J Bone Miner Res 23:1140–1149 Li J, Chen G, Zheng L, Luo S, Zhao Z (2007) Osteoblast cytoskeletal modulation in response to compressive stress at physiological levels. Mol Cell Biochem 304:45–52

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Liu J, Liu T, Zheng Y, Zhao Z, Liu Y, Cheng H, Luo S, Chen Y (2006) Early responses of osteoblast-like cells to different mechanical signals through various signaling pathways. Biochem Biophys Res Commun 348:1167–1173 McMahon AP, Champion JE, McMahon JA, Sukhatme VP (1990) Developmental expression of the putative transcription factor Egr-1 suggests that Egr-1 and c-fos are coregulated in some tissues. Development 108:281–287 Moalli MR, Caldwell NJ, Patil PV, Goldstein SA (2000). An in vivo model for investigations of mechanical signal transduction in trabecular bone. J Bone Miner Res 15:1346–1353 Morinobu M, Ishijima M, Rittling SR, Tsuji K, Yamamoto H, Nifuji A, Denhardt DT, Noda M (2003) Osteopontin expression in osteoblasts and osteocytes during bone formation under mechanical stress in the calvarial suture in vivo. J Bone Miner Res 18:1706–1715 Nomura S, Takano-Yamamoto T (2000) Molecular events caused by mechanical stress in bone. Matrix Biol 19:91–96 Norrdin RW, Shih MS (1988) Systemic effects of prostaglandin E2 on vertebral trabecular remodeling in beagles used in a healing study. Calcif Tissue Int 42:363–368 Ogasawara A, Arakawa T, Kaneda T, Takuma T, Sato T, Kaneko H, Kumegawa M, Hakeda Y (2001) Fluid shear stress-induced cyclooxygenase-2 expression is mediated by C/EBP beta, cAMP-response element-binding protein, and AP-1 in osteoblastic MC3T3-E1 cells. J Biol Chem 276:7048–7054 Ogata T (1997) Fluid flow induces enhancement of the Egr-1 mRNA level in osteoblast-like cells: involvement of tyrosine kinase and serum. J Cell Physiol 170:27–34 Qian A, di S, Gao X, Zhang W, Tian Z, Li J, Hu L, Yang P, Yin D, Shang P (2009) cDNA microarray reveals the alterations of cytoskeleton-related genes in osteoblast under high magneto-gravitational environment. Acta Biochim Biophys Sin (Shanghai) 41:561–577 Reich KM, McAllister TN, Gudi S, Frangos JA (1997) Activation of G proteins mediates flowinduced prostaglandin E2 production in osteoblasts. Endocrinology 138:1014–1018 Sawada Y, Sheetz MP (2002) Force transduction by Triton cytoskeletons. J Cell Biol 156:609–615 Schriefer JL, Warden SJ, Saxon LK, Robling AG, Turner CH (2005) Cellular accommodation and the response of bone to mechanical loading. J Biomech 38:1838–1845 Smalt R, Mitchell FT, Howard RL, Chambers TJ (1997) Mechanotransduction in bone cells: induction of nitric oxide and prostaglandin synthesis by fluid shear stress, but not by mechanical strain. Adv Exp Med Biol 433:311–314 Stein GS, van Wijnen AJ, Stein JL, Lian JB, Pockwinse SH, McNeil S (1999) Implications for interrelationships between nuclear architecture and control of gene expression under microgravity conditions. FASEB J 13 Suppl:S157–S166 Suponitzky I, Weinreb M (1998) Differential effects of systemic prostaglandin E2 on bone mass in rat long bones and calvariae. J Endocrinol 156:51–57 Thiel G, Cibelli G (2002) Regulation of life and death by the zinc finger transcription factor Egr-1. J Cell Physiol 193:287–292 Toma CD, Ashkar S, Gray ML, Schaffer JL, Gerstenfeld LC (1997) Signal transduction of mechanical stimuli is dependent on microfilament integrity: identification of osteopontin as a mechanically induced gene in osteoblasts. J Bone Miner Res 12:1626–1636 Turner CH (1999). Toward a mathematical description of bone biology: the principle of cellular accommodation. Calcif Tissue Int 65:466–471 Wadhwa S, Godwin SL, Peterson DR, Epstein MA, Raisz LG, Pilbeam CC (2002) Fluid flow induction of cyclo-oxygenase 2 gene expression in osteoblasts is dependent on an extracellular signal-regulated kinase signaling pathway. J Bone Miner Res 17:266–274 Wu CJ, Fu Q (advisor) (2009). Effect of cytoskeleton reorganization inhibition on the mechanosensitivity of COX-2 in osteoblasts induced by fluid shear stress. Master’s Thesis, Sun Yat-Sen University, Guangzhou Zayzafoon M, Fulzele K, McDonald JM (2005) Calmodulin and calmodulin-dependent kinase IIalpha regulate osteoblast differentiation by controlling c-fos expression. J Biol Chem 280:7049–7050

Chapter 4

Involvement of the Cytoskeletal Elements in Articular Cartilage Mechanotransduction Emma J. Blain

Abstract The cytoskeleton of all cells is a three-dimensional network comprising actin microfilaments, tubulin microtubules and intermediate filaments. The cytoskeletal networks are highly organised in structure enabling them to fulfil their biological functions. They are involved in many diverse cellular processes including alteration of cell shape, migration, cell division, movement of organelles, endocytosis, secretion and extracellular matrix assembly. This review will primarily focus on the organisation and function of the three major cytoskeletal networks in articular cartilage chondrocytes. Articular cartilage is a major load-bearing tissue of the synovial joint; it is well known that the cytoskeleton acts as a physical interface between the chondrocytes and the extracellular matrix in “sensing” mechanical stimuli. The effect of physiological and abnormal mechanical load on cytoskeletal element expression and organisation will also be reviewed. Keywords Cytoskeleton · Actin microfilaments · Tubulin microtubules · Vimentin intermediate filaments · Articular cartilage · Chondrocyte · Mechanotransduction

4.1 Introduction 4.1.1 Structure and Function of the Three Major Cytoskeletal Elements The cytoskeleton is a three-dimensional network which provides a physical interface between the cell and the extracellular matrix in “sensing” a mechanical stimulus. Converting a mechanical stimulus into a biochemical signal to initiate downstream biosynthetic responses is primarily mediated by the three major E.J. Blain (B) Connective Tissue Biology Laboratories, Biomedical Sciences Building, School of Biosciences, Cardiff University, Museum Avenue, Cardiff, CF10 3AX, UK e-mail: [email protected]


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cytoskeletal elements comprising actin microfilaments, intermediate filaments and tubulin microtubules (Benjamin et al., 1994). The cytoskeletal proteins have distinct roles in many diverse cellular processes as discussed below.

4.1.1.1 Actin Actin, a 43-kDa globular protein, is reported to be the most abundant protein in eukaryotic cells (Disanza et al., 2005) and comprises three different monomers (α, β and γ). Under physiological conditions, the actin monomers (globular/G-actin) assemble into long, highly organised filaments (filamentous/F-actin) (Fig. 4.1), and it is the formation of these F-actin filaments which are critical to the activities of many fundamental cellular events including cell migration (Heath and Holifield, 1991) and adhesion (Turner and Burridge, 1991), movement of organelles (Simon and Pon, 1996), secretion (Sontag et al., 1988; Koukouritaki et al., 1996), alteration of cell shape (Sims et al., 1992) and extracellular matrix assembly (Hayes et al., 1999). Actin assembly/disassembly is coupled with continuous ATP hydrolysis; a cleft containing the nucleotide ATP (or ADP) and the cations Mg2+ and Ca2+ exists within the G-actin moiety to meet the energy demands of actin polymerisation. This ATP-dependent process of actin filament dis/assembly is referred to as “treadmilling”, and it is the mechanism controlling cell locomotion, morphogenetic movements and intracellular transport (Pollard and Borisy, 2003). Actin filaments are polarised structures comprising a fast-growing (plus or barbed) end and a slow-growing (minus or pointed) end (Ono, 2007). Under physiological conditions, Mg2+ -ATP-bound G-actin is incorporated into growing filaments at the barbed end (Cooper and Schafer, 2000; Ono, 2007). ATP is hydrolysed during F-actin polymerisation and a phosphate moiety is released. The resulting ADP–actin filaments are disassembled through the loss of ADP–G-actin from the pointed end (Cooper and Schafer, 2000; Ono, 2007). The released ADP–G-actin undergoes nucleotide exchange to produce ATP–G-actin enabling continuation of F-actin polymerisation at the barbed end.

Monomer Subunits

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Fig. 4.1 Schematic representation of the assembly of the F-actin microfilaments

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4.1.1.2 Actin-Binding Proteins Actin treadmilling and its organisation into functional cytoplasmic networks is regulated by a variety of actin-binding proteins (Pollard and Borisy, 2003) which function by: (1) promoting nucleation of actin e.g. Arp2/3 complex (Goley and Welch, 2006), (2) inducing actin filament depolymerisation e.g. the actin-depolymerising factor family including ADF and cofilin (Ono, 2007), (3) associating with G-actin moieties e.g. profilin and thymosin β4 (Yarmola and Bubb, 2004), (4) capping the F-actin filament end e.g. gelsolin (Cooper and Schafer, 2000). The actin-binding proteins are therefore an essential prerequisite for regulating actin treadmilling, and this is fulfilled by a series of tightly controlled cell signalling pathways which co-ordinate the activities of these proteins (Disanza et al., 2005; Pullikuth and Catling, 2007).

4.1.2 Intermediate Filaments The intermediate filaments (10–12 nm: intermediate in size between the microfilaments and microtubules) are highly organised fibrous protein structures which connect the plasma with the nuclear membrane. Although intermediate filament function(s) has not been completely characterised to date, they, in addition to the other cytoskeletal elements, have been implicated in intracellular mRNA transport (Glotzer and Ephrussi, 1996; Lopez de Heredia and Jansen, 2004) and distribution of organelles and proteins (Herrmann and Aebi, 2000). The intermediate filaments are involved in the localisation of organelles including the Golgi apparatus, endosomes, lysosomes and nuclei in the cell cytoplasm (Styers et al., 2005). However, compelling evidence exists to demonstrate their involvement in signal transduction (Traub, 1995), and in particular, they fulfil a critical role in mechanotransduction i.e. the propagation of a mechanical stimulus into a downstream biochemical response(s) (Lazarides, 1980; Wang et al., 1993). The role of intermediate filaments in mechanotransduction has been verified in several human genetic diseases (Omary et al., 2004) and in knockout studies (Eckes et al., 1998; Broers et al., 2004) where cells lacking their usual complement of intermediate filaments have been demonstrated to be mechanically fragile. These studies infer that the intermediate filaments confer mechanical integrity to the cells and tissues. Five classes of intermediate filaments exist including: class I – cytokeratins (epithelia), class II – desmin (muscle cells), class III – vimentin (mesenchymal cells), class IV – glial fibrillar acidic protein (glial cells) and neurofilaments (neurons), and the class V – nuclear lamins which are present in the nuclei of all cell types. Unlike the other two major cytoskeletal elements, intermediate filament expression can be both developmentally regulated and cell-type specific i.e.

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only vimentin class III intermediate filaments are observed in chondrocytes, and may reflect their diversity in biological function. Intermediate filaments all contain common highly-conserved structural features including a central α-helical rod domain flanked by non-α-helical N-(head) and C-terminal (tail) domains. The highly-conserved central α-helical rod primarily consists of seven-residue repeats which configures a coiled-coil structure (Parry et al., 2007). Intermediate filaments assemble through the association of two monomeric subunits aligned in parallel and in register to form a homodimer. Two homodimers associate to form anti-parallel, half-staggered tetramers, and eight of these tetramers associate laterally to form a supercoiled sheet which has the characteristic appearance of a rope-like filamentous structure (Fuchs and Weber, 1994; Sokolova et al., 2006) (Fig. 4.2). However, it is the degree of heterogeneity in the non-α-helical head and tail domains (size and sequence can differ) which provides the array of intermediate filament classes indicated above. The intracellular organisation of intermediate filament networks i.e. the dynamics of assembly and disassembly is under the control of specific protein kinases and phosphatases (Izawa and Inagaki, 2006). Phosphorylation of the intermediate filament protein occurs on specific serine and threonine residues, resulting in filament disassembly; most of these phosphorylation sites are located in the N-terminal non-α-helical head domain (Beuttenmuller et al., 1994; Izawa and Inagaki, 2006) (Fig. 4.3). As a consequence of phosphorylation, filament disassembly generates a pool of soluble subunits in preparation for the next phase of polymerisation. Conversely, dephosphorylation of the intermediate filament instigates filament assembly, and the architecture undergoes a rearrangement (Herrmann and Aebi, 2000). The direct involvement of intermediate filaments in signal transduction has been demonstrated in many cell types, as many of the regulatory proteins appear to be dependent upon their interaction with the filaments. Several studies have demonstrated that protein kinases phosphorylate intermediate filaments at specific sites in vitro (reviewed in (Izawa and Inagaki, 2006; Minin and Moldaver, 2008)). These phosphorylation sites are located mainly on the N-terminal protein domains, which are imperative for intermediate filament assembly. In vivo, vimentin is one of the predominant phosphoproteins in the cytoplasm, and in vitro provides a good substrate for second messenger protein kinases (Fig. 4.3). The dynamic status of vimentin de/phosphorylation is critical in numerous signal transduction pathways

8–12nm Monomer

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Rope-like filament (8 tetramers)

Fig. 4.2 Schematic representation of the assembly of the vimentin intermediate filaments

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Phosphorylated serine residues in vimentin head domain 6

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Rho-kinase

+

CaM kinase II

+

cdc2 kinase

55

64

65

71

72

86

+ + +

+

+

Plk1 Aurora-B

82

+

+ +

+

+

+

+

+

+

+

Fig. 4.3 Schematic representation of the phosphorylation sites in the vimentin head domain. Serine residues are phosphorylated by PKC (protein kinase C), PKA (protein kinase A), PAK (p21-activated kinase), Rho-kinase, CaM kinase II (Ca2+ /calmodulin-dependent protein kinase II, cdc2 kinase, Plk1 (polo-like kinase 1) and Aurora-B (adapted from Izawa and Inagaki, 2006; Minin and Moldaver, 2008)

including the MAP kinases ERK 1/2 (Perlson et al., 2006); Blain, unpublished observations), various 14-3-3 signalling protein isoforms and apoptotic factors (Kim and Coulombe, 2007). 4.1.2.1 Nuclear Lamins The ubiquitous nuclear lamins are the principal components of the nuclear lamina; their primary role is to provide a structural framework to maintain interphase nuclear shape, but the nuclear lamins are also essential for many aspects of normal nuclear function including DNA replication, chromatin organisation and gene expression (Dechat et al., 2008). The nuclear lamina, which resides beneath the inner nuclear envelope, comprises four major lamin proteins: A, B1, B2 and C (Burke and Stewart, 2006), and as with the class III filaments, the nuclear lamins are also developmentally regulated; all cells express at least one lamin B, however the A-type lamins are absent from early embryonic development, embryonic stem cells and certain stem cell populations in adults (Rober et al., 1989; Cohen et al., 2008).

4.1.3 Tubulin Microtubules Tubulin microtubules are highly conserved α/β dimeric proteins, each monomer being approximately 450 amino acids in length with a molecular mass of 55 kDa

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(Valiron et al., 2001), that self-assemble to form the polymeric microtubular structure. Many functions have been attributed to the microtubules including cell motility, mitosis, cell morphogenesis, organisation and transport of organelles. The principal function of microtubules occurs during mitosis when they rearrange to form the “mitotic spindle” to orientate the plane of cell cleavage (Mitchison et al., 1986). The microtubules also operate as an intracellular supramolecular motor to segregate the chromosomes to the cell poles during anaphase (Mitchison et al., 1986). The ability of the microtubules to act as a supramolecular motor allows them to also act as a highway for “bi-directional” transport: protein trafficking and secretion results from the delineation of microtubule tracks along which the cargo are shuttled during endo- and exocytosis (Vale, 1987; Thyberg and Moskalewski, 1999). Microtubules organise the cell interior and are also involved in assembling intracellular organelles e.g. endoplasmic reticulum and the Golgi apparatus (Rogalski and Singer, 1984). Microtubules are also involved in sensing and responding to mechanical stimuli as they are one of the principal constituents of the axonemal structures found in cilia (Gibbons, 1981; Poole et al., 2001) and flagella (Woolley, 2000). Tubulin assembly/disassembly is coupled with GTP hydrolysis; α-tubulin binds a GTP molecule in a non-exchangeable manner whereas GTP-β-tubulin binding is exchangeable. GTP hydrolysis occurs upon the addition of a tubulin heterodimer to the microtubule end, with the β-tubulin subunit of the heterodimer exposed on the faster growing end (plus end) and the α-tubulin subunit exposed on the slow growing end (minus end) (Valiron et al., 2001). Microtubules organise into linear protofilaments that assemble to form 24 nm-wide cylindrical structures (Fig. 4.4). The numbers of protofilaments which constitute a microtubule is generally considered to be 13 in vivo (Wade and Hyman, 1997), although the number may vary depending on assembly conditions. As a result of the unique organisation of the α- and β-tubulin heterodimers, the microtubule protofilament is polarised resulting in differing kinetic and structural properties. Microtubule polarity is critical for the correct implementation of the microtubule motor proteins (comprehensively reviewed in (Mallik and Gross, 2004; Wu et al., 2006)). Recognition of microtubule

Subunit α-tubulin

GTP

GDP

β-tubulin

24nm

Protofilament

8nm

Fig. 4.4 Schematic representation of the assembly of the tubulin microtubules

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polarity by the motor proteins enables the specific motor employed to determine the direction of transport along the microtubule tracks (Mallik and Gross, 2004). Dynein shuttles cargo toward the microtubule minus-end (Hirokawa, 1998), sustaining microtubule-dependent trafficking of newly synthesised proteins from the endoplasmic reticulum to the cis side of the golgi stacks, and providing vesicular transport for endosomal clearance (Thyberg and Moskalewski, 1999). In contrast, the motor protein kinesin moves toward the plus-end of the microtubules bringing cargo toward the cell periphery (Hirokawa, 1998), e.g. by trafficking membrane and secretory proteins (following post-translational modifications) from the trans side of the golgi stacks. Kinesin is also involved in propagating the return transport of endoplasmic reticulum components from the cis side of the golgi stacks (Thyberg and Moskalewski, 1999).

4.2 Articular Cartilage Structure and Function The major load-bearing tissue of the synovial joint is articular cartilage which provides a smooth and resilient surface for joint locomotion. It is unique in being avascular, aneural and alymphatic and contains a sparse population of cells – the “chondrocyte” interspersed in a dense extracellular matrix. The ability of articular cartilage to withstand highly repetitive stresses, brought about by movement, depends on the structure and composition of this extracellular matrix.

4.2.1 Tissue Composition The unique mechanical properties of articular cartilage are dependent on the organisation (Fig. 4.5a) and composition of the tissue (Fig. 4.5b). Morphologically, there are four distinct zones including the superficial, transitional, deep, and calcified zone which merges into the subchondral bone. Articular cartilage (wet weight) comprises approximately 65–80% water, with 80% being in the superficial zone and 65% in the deep zone; the highly hydrated nature of the tissue allows for load-dependent deformation, as well as providing a medium for lubrication to maintain a low-friction bearing surface zone (Buckwalter and Mankin, 1998). Collagens comprise 10–20% of the wet weight of articular cartilage with type II forming the principal component (90–95%); parallel arrangement of the collagen fibrils in the surface zone is responsible for providing the greatest tensile and shear strength (Duance et al., 1998). The other principal constituent of articular cartilage is proteoglycan (10–20%); two major classes of proteoglycans are found in articular cartilage: large aggregating monomers e.g. aggrecan and the small leucine-rich proteoglycans e.g. decorin and biglycan (Hardingham et al., 1992). Aggrecan is most abundant in the transitional and deep zone, where due to its viscoelastic nature (hydrated gel) provides cartilage with its function of compressibility under load. Articular cartilage also contains a multitude of other extracellular matrix macromolecules which contribute to its unique structural/mechanical integrity, but this lies outside the scope of this review.

84

E.J. Blain hyaluronate link protein

A.

B. aggrecan decorin SZ

COMP

HA receptor glycosaminoglycan

TZ

biglycan DZ

type IX collagen chondrocyte

SB

fibromodulin

type II collagen core type XI collagen

Fig. 4.5 Organisation and composition of articular cartilage. (a) Haematoxylin and Eosin staining of articular cartilage from the carpal metacarpal joint of a 7-day-old bovine calf. (b) Schematic cartoon illustrating the interactions between the major components of articular cartilage (adapted from Silver and Glasgold, 1995). Collectively the composition and organisation of these matrix macromolecules provides the tissue with its unique biomechanical properties to withstand compressive load

Chondrocytes envelop themselves in “basket-like networks of fine fibrils of elaborate structure” – termed a chondron (Muir, 1995). As individual units, chondrons appear to be compression-resistant, fluid-filled elements that dampen mechanical, osmotic and physicochemical changes induced by mechanical loading (Muir, 1995). The elaborate structure of articular cartilage allows for many complex interactions between the chondrocyte and its surrounding environment, and it acts as an ideal intermediary for sensing and responding to alterations in mechanical load.

4.2.2 Tissue Function Articular cartilage is a specialised connective tissue that functions in dissipating mechanical loads across the joint. All components of the joint i.e. articular cartilage, subchondral bone, meniscus, synovium, muscles, ligaments and tendons participate in load transmission but it is the articular cartilage itself, which provides a resilient and compliant articulating surface to the bones in diarthrodial joints. In simple terms, the articular cartilage prevents potentially damaging local stress concentrations from being experienced, whilst providing a low friction-bearing surface to enable free movement of the joint for locomotion (Jeffrey et al., 1997). During normal walking, articular cartilage is subjected to a complex state of dynamic

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loading which is primarily applied perpendicular to the surface of the articular cartilage (Armstrong et al., 1984). Dynamic, compressive loading of articular cartilage induces cell deformation, changes in hydrostatic or fluid pressure and deformation of the charged extracellular matrix, resulting in alterations in osmolality and pH (Urban, 1994; Hall et al., 1996a), as depicted (Fig. 4.6). Chondrocytes can detect these “physico-chemical” signals, and respond by altering their metabolism via the activation of specific intracellular signalling pathways (Palmoski et al., 1980; Palmoski and Brandt, 1984; Kiviranta et al., 1988; Sah et al., 1989).

synovial membrane

pericellular matrix

territorial matrix

Mechanical Load

synovial fluid

soluble factors fluid flow hydrostatic pressure deformation

fluid expression

growth factor release

compression of matrix [cations] osmolality α5β1 integrin

F-actin FAK

IL-4

Ca2+ NO

iNOS

vimentin ERK

tubulin ECM macromolecules

transcriptional responses macromolecule synthesis

ECM assembly/turnover

Stretch-activated channels

MMPs Aggrecanases TIMPs

Fig. 4.6 A schematic representation of how mechanical deformation, acting on articular cartilage, can alter both the intra- and extracellular environment of the cells. Exposure of cells to mechanical stimulation disseminates further effects across the extracellular matrix including opening of stretch-activated channels, focal adhesion kinase signalling via the integrins, and rearrangement of the cytoskeletal elements to induce changes in gene and protein expression (adapted from Urban, 1993)

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Under compressive load, cartilage chondrocytes are subject to deformation which induces an alteration in cell volume and/or surface area (Guilak et al., 1995; Knight et al., 1998), and in native tissue this deformation is associated with an increase in extracellular osmolality (Guilak et al., 1995; Buschmann et al., 1996). Compressioninduced adaptations in chondrocyte volume and/or surface area activate various membrane ion transporters (Urban et al., 1993; Hall et al., 1996b), stretch sensitive ion channels and induce alterations in membrane potential (Guilak et al., 1994; Wright et al., 1996). Compressive load also results in nuclear deformation, concomitant with a reduction in nucleus volume (Guilak, 1995; Buschmann et al., 1996). Load-induced deformation of the intracellular organelles e.g. the nucleus is thought to effect alterations in matrix synthesis (Buschmann et al., 1996; Janmey, 1998). The mechanism by which compressive load is mediated through the cell to the nucleus to effect a metabolic response is thought to involve the cytoskeleton (Ben-Ze’ev, 1991; Wang et al., 1993; Ingber et al., 1994; Maniotis et al., 1997; Janmey, 1998).

4.3 Cytoskeletal Element Composition in Articular Chondrocytes The chondrocyte cytoskeleton comprises actin microfilaments, vimentin intermediate filaments and tubulin microtubules (Fig. 4.7), in addition to the ubiquitous nuclear lamins (Benjamin et al., 1994).

4.3.1 Organisation of the Cytoskeletal Elements in Articular Chondrocytes The architecture of the three major cytoskeletal networks in chondrocytes have been characterised in articular cartilage explants (Durrant et al., 1999; Langelier et al., 2000), in primary chondrocytes cultured in agarose (Idowu et al., 2000; Knight et al., 2001; Trickey et al., 2004; Sasazaki et al., 2008) and in high-density monolayer cultures (Blain et al., 2006). Key morphological features are described (Table 4.1A). F-actin is predominantly cortical in localisation and distributed at the periphery of the chondrocyte (Idowu et al., 2000; Langelier et al., 2000; Knight et al., 2001; Trickey et al., 2004; Blain et al., 2006; Sasazaki et al., 2008). In contrast, the vimentin intermediate filaments extend throughout the cytoplasm and form a highly organised architecture in chondrocytes, traversing from the plasma to the nuclear membrane (Idowu et al., 2000; Langelier et al., 2000; Holloway et al., 2004; Trickey et al., 2004; Blain et al., 2006; Sasazaki et al., 2008). Tubulin microtubules are uniformly distributed throughout the chondrocyte cytoplasm and exhibit a loose mesh-like network (Jortikka et al., 2000; Langelier et al., 2000; Trickey et al., 2004; Blain et al., 2006). Zonal differences are also evident in the organisation of the cytoskeletal networks in articular cartilage chondrocytes (Langelier et al., 2000). A comparison of cytoskeletal element distribution through the depth of

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Table 4.1 Organisation of the three major cytoskeletal elements: actin, vimentin and tubulin in [A] normal and [B] osteoarthritic articular chondrocytes. All three structures are highly organised in normal chondrocytes and their localisation in the cytoplasm reflects their biological functions within the cell. However, in chondrocytes from osteoarthritic cartilage there is a general loss of actin, vimentin and tubulin organisation, and a reduction in protein amounts. The loss of vimentin organisation in osteoarthritic cartilage may be attributed to an increase in vimentin degradation products which are derived from N-terminal domain cleavage i.e. that which is responsible for filament assembly Cytoskeletal element

Organisation – normal chondrocytes

F-Actin

• cortical distribution • predominantly localised to cell periphery

Vimentin

• highly organised structural network • traverses cytoplasm connecting nuclear membrane with cell periphery • loose, basket-like meshwork • uniformly distributed throughout cell cytoplasm

Tubulin

Reference Langelier et al. (2000), Idowu et al. (2000), Knight et al. (2001), Trickey et al. (2004), Blain et al. (2006), Fioravanti et al. (2003), and Sasazaki et al. (2008) Langelier et al. (2000), Idowu et al. (2000), Holloway et al. (2004), Trickey et al. (2004), Blain et al. (2006), Fioravanti et al. (2003), and Sasazaki et al. (2008) Jortikka et al. (2000), Langelier et al. (2000), Trickey et al. (2004), Blain et al. (2006), and Fioravanti et al. (2003)

Table 4.1B Cytoskeletal element F-Actin Vimentin

Tubulin

Organisation – osteoarthritic chondrocytes

Reference

• less well-defined and localised diffusely in the cytoplasm or limited to cell periphery [Human] • vimentin-rich multiple, elongated processes (correlating with greatest histological and macroscopic signs of OA) [Human] • loss of expression – 37.1% reduction. Predominantly localised to the superficial zone; ranged from simple disorganisation to total disruption of architecture [Rat partial menisectomy model] • diffuse cytoplasmic distribution [Human] • vimentin degradation products – derived from cleavage of N-terminal domain (responsible for filament assembly) [Human] • often absent [Human] • loss of expression – 20.1% reduction [Rat partial menisectomy model]

Fioravanti et al. (2003) Holloway et al. (2004)

Capin-Gutierrez et al. (2004)

Lambrecht et al. (2008) Lambrecht et al. (2008)

Fioravanti et al. (2003) Capin-Gutierrez et al. (2004)

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Fig. 4.7 The cytoskeleton of primary bovine chondrocytes depicting F-actin microfilaments, vimentin intermediate filaments and tubulin microtubules cultured in high density monolayer for 7 days. Cells were visualised using FITC-conjugated antibodies in conjunction with scanning confocal microscopy F-actin microfilaments

Vimentin intermediate filaments

Tubulin microtubules

skeletally mature bovine articular cartilage demonstrated that F-actin was uniformly distributed; in contrast, both vimentin and tubulin were predominantly detected in cells of the superficial zone (Langelier et al., 2000). Interestingly, there was an increasing gradient in both vimentin and tubulin network organisation in cartilage situated in proximity to weight-bearing regions (Langelier et al., 2000). The exhibition of specific structural characteristics in chondrocyte cytoskeletal network organisation reflects their distinct functions within the cell.

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4.3.2 Cytoskeletal Element Functions in Articular Chondrocytes In addition to the generic functions already outlined above (Section 4.1), the cytoskeletal elements provide further, more specific functions in articular chondrocytes.

4.3.2.1 Actin Microfilaments The ability of a chondrocyte to withstand compressive loads is dependent on the organisation of the F-actin cytoskeleton (Guilak, 1995). A chondrocyte’s response to mechanical stimulation is thought to occur as a consequence of an alteration in cell shape and volume which activates intracellular signalling cascades. Deformation of the chondrocyte nucleus is involved in relaying intracellular signals via the F-actin network. Disruption of the F-actin cytoskeleton with cytochalasin D affected the relationship between matrix deformation and alterations in nucleus shape and height (Guilak, 1995). In the presence of cytochalasin D, chondrocyte stiffness decreased by 90% and viscosity by 80% (Trickey et al., 2004), demonstrating the importance of the actin cytoskeleton in maintaining the mechanical integrity of the cell (refer to Section 4.4 for more detail). Actin microfilaments are also crucial in maintaining cell – matrix interactions. Studies have demonstrated that cytoskeletal actin disruption within articular cartilage uncouples the chondrocytes from the matrix, altering tissue synthesis and structure by inhibiting type II collagen and sulphated glycosaminoglycan synthesis (Takigawa et al., 1984; Brown and Benya, 1988; Newman and Watt, 1988; Loty et al., 1995). Cytochalasin D induced disruption of F-actin reduced chondrocyte pericellular matrix assembly and the retention of proteoglycans within the cartilage tissue (Nofal and Knudson, 2002). This loss of function was attributed to a reduction in the anchorage of CD44 in the chondrocyte membrane which reduced the capacity of CD44 to bind to its extracellular ligand.

4.3.2.2 Vimentin Intermediate Filaments Vimentin intermediate filament organisation has been characterised in articular cartilage tissue (Durrant et al., 1999; Langelier et al., 2000; Holloway et al., 2004), in isolated chondrocytes (Idowu et al., 2000; Langelier et al., 2000; Holloway et al., 2004; Trickey et al., 2004; Sasazaki et al., 2008) and in high-density monolayer cultures (Blain et al., 2006), but their function(s) are still being unravelled. The vimentin knock-out mouse displays no obvious phenotype, being able to develop and reproduce normally; the skeleton is apparently normal (Colucci-Guyon et al., 1994), although the morphology of the articular cartilage was not fully assessed. Interestingly, vimentin –/– fibroblasts cultured in vitro displayed reduced mechanical stability when exposed to external stimuli (Eckes et al., 1998). Vimentin –/– mesenteric resistance arteries decreased their response to blood flow (Henrion et al., 1997), and in vimentin –/– carotid arteries flow-induced remodelling was reduced

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(Schiffers et al., 2000), demonstrating a requirement of vimentin in the mechanotransduction of shear stress. These studies suggest that the absence of a vimentin filament network does not impair basic cellular functions, but that cells are mechanically compromised; therefore Eckes et al. proposed that cellular functions that are dependent upon mechanical stability become impaired. This argues well with the observation of a more prominent vimentin expression in weight-bearing areas of articular cartilage in situ (Eggli et al., 1988). More recently, the contribution of the vimentin filaments in withstanding mechanical load has been investigated in chondrocytes (Lahiji et al., 2004; Trickey et al., 2004). In conjunction with actin microfilaments, the vimentin cytoskeleton confers some mechanical integrity to the cell, as a decrease in stiffness and viscosity was observed in chondrocytes treated with 40 mM acrylamide to disrupt the vimentin filaments (Trickey et al., 2004). We have demonstrated a novel function of the vimentin network in contributing to the maintenance of the chondrocyte phenotype (Blain et al., 2006). Chondrocyte matrix synthesis is regulated by the state of vimentin network assembly/disassembly (using 5 mM acrylamide to disrupt vimentin architecture), as evidenced by a significant reduction in de novo collagen and sGAG biosynthesis at both the mRNA and protein level (Blain et al., 2006). Preliminary studies indicate that this response may be due to sustained phosphorylation of ERK1/2 (unpublished observations), which has previously been reported to inhibit the chondrocyte phenotype (Seghatoleslami et al., 2003). 4.3.2.3 Tubulin Microtubules The microtubule network does not directly confer any mechanical integrity to the chondrocyte, as disruption of tubulin with colchicine did not alter the stiffness or viscosity of the cell (Trickey et al., 2004). However, a sophisticated tubulin network, in the form of acetylated α-tubulin, has been observed in the primary cilia of chondrocytes (Poole et al., 2001). The primary cilium is a highly conserved, single cytoplasmic organelle which projects into the pericellular matrix and monitors the mechanical environment of the cartilage tissue (McGlashan et al., 2008). The primary cilium interacts with pericellular matrix components including collagen types II and VI (Poole et al., 1997; McGlashan et al., 2006). Ultrastructural studies have also identified a direct connection between extracellular collagen fibres and the proteins which decorate the ciliary microtubules, suggesting a matrix-cilium-Golgi continuum in chondrocytes to detect mechanical fluctuations in the surrounding matrix (Poole et al., 2001). However, as in all cell types, the microtubules have a critical function in protein trafficking and secretion (Thyberg and Moskalewski, 1999). The integrity of the tubulin network, manipulated using disrupting agents i.e. colchicine or nocodazole, is also essential for the synthesis and secretion of both collagens and proteoglycans in chondrocytes (Jansen and Bornstein, 1974; Lohmander et al., 1976, 1979; Madsen et al., 1979; Takigawa et al., 1984). Colchicine dissociates the Golgi apparatus of chondrocytes (Moskalewski et al., 1975), therefore its negative impact on

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matrix synthesis may reflect alterations to the normal functioning of the Golgi apparatus, or its disruption of normal routes of intracellular transport.

4.4 Biomechanics and the Chondrocyte Cytoskeleton 4.4.1 Contribution of the Cytoskeletal Elements to the Mechanical Properties of the Chondrocyte Cytoskeletal element organisation is a highly dynamic process, as demonstrated in isolated chondrocytes whereby F-actin, vimentin and tubulin assembly increased markedly over a culture period of days (Lee et al., 2000a; Blain et al., 2006). The biomechanical integrity of the chondrocyte is dependent on the organisation of these major cytoskeletal elements, as each of the networks serve an important function in the cell’s ability to resist and recover from mechanical forces. Ofek et al. hypothesised that each cytoskeletal component would differentially contribute to the compressive properties and behaviour of single chondrocytes, which would have specific ramifications for mechanotransduction pathways (Ofek et al., 2009). Chemical disruption of individual cytoskeletal elements prior to the application of a range of compressive strains demonstrated that F-actin was the greatest contributor to chondrocyte stiffness, vimentin and tubulin contributed significantly to volumetric changes/compressibility and that tubulin was essential for cell recovery post-deformation (Ofek et al., 2009). These specific functions relate to the distribution of the cytoskeletal elements within the chondrocyte (Fig. 4.7). F-actin is localised to the periphery of the cell where it provides mechanical reinforcement for the cytoplasm (Guilak et al., 1995; Trickey et al., 2004; Knight et al., 2006). Conversely, tubulin forms a loose meshwork throughout the cell cytoplasm, increasing the ability of the cell to revert to its original morphology post-compression. Vimentin, comprising a coiled-coil structure with flexible regions at its head and tail (Fig. 4.2) acts primarily to resist tensile forces and counteract compression-induced volumetric changes (Ofek et al., 2009). Chondrocyte stiffness can play a critical role in the cell’s interpretation and ability to function under an altered mechanical environment; the functional role of the cytoskeleton in resisting compression is particularly important as cell volume alterations can influence matrix synthesis and tissue homeostasis, potentially predisposing the tissue to pathological conditions e.g. osteoarthritis (OA).

4.4.2 Mechanical Load Influences Cytoskeletal Element Organisation The application of a physiological mechanical load is imperative for maintaining articular cartilage integrity (Fig. 4.6); extracellular matrix synthesis, i.e. sGAG and collagen production, is regulated by mechanical load, and sustained synthesis is

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in itself required to maintain the ability of the cartilage to withstand the forces that are associated with tissue deformation (Urban, 1994). However, abnormal nonphysiological loads including disuse (Palmoski et al., 1980; Palmoski and Brandt, 1984) and high, repetitive loads (Vasan, 1983; Radin et al., 1984) inhibit extracellular matrix synthesis, and is detrimental to the properties of the tissue in vivo. Altering the biomechanical properties of the chondrocyte influences cell-matrix interactions (Guilak and Mow, 2000), supporting the role of the cytoskeleton in mechanical signal transduction. Collectively, the three major cytoskeletal networks play a fundamental role in maintaining the phenotype of chondrocytes and acting as a physical interface between the chondrocyte and the extracellular matrix in “sensing” mechanical stimuli (Table 4.2).

4.4.2.1 Actin Microfilaments The F-actin cytoskeleton is intimately involved in the conversion of a mechanical stimulus into a biochemical response during articular chondrocyte mechanotransduction (Wang et al., 1993; Banes et al., 1995; Guilak, 1995; Grodzinsky et al.,

Table 4.2 The effect of mechanical load on the expression and organisation of cytoskeletal elements in articular cartilage chondrocytes Cytoskeletal element

Effect of load on chondrocyte cytoskeleton

Reference

Loss of actin stress fibres with increasing hydrostatic pressure (15–30 MPa); reversible after 2 h Compressive strain (10–15%) induced punctuate reorganisation of F-actin replacing uniform cortical distribution in unloaded cells Actin reorganisation in response to osmotic stress – loss of cortical F-actin distribution due to severing and detachment from cell periphery

Parkkinen et al. (1995)

Increased thymosin β4 mRNA: 0.5 MPa, 1 Hz, 10 min reorganisation of thymosin β4 and F-actin architecturesa Increased cofilin and destrin mRNA: 15%, 1 Hz, 10 min temporal disassembly of cortical F-actin

Blain et al. (2002) and Blain et al.(2003)

Vimentin

Increased content in weight-bearing regionsa Increased filament organisation: 0–4 MPa, 1 ha Reduction in vimentin mRNA after continuous or cyclic (0.5 Hz) stretch (5 MPa) Increased vimentin mRNA in response to cyclic strain (24 h)

Eggli et al. (1988) Durrant et al. (1999) Karjalainen et al. (2003) Lahiji et al. (2004)

Tubulin

Loss of organisation: (24 MPa, 3 h)

Fioravanti et al. (2005)

F-Actin

Actinbinding proteins

a Cartilage

explants

Knight et al. (2006)

Erickson et al. (2003) and Chao et al. (2006)

Campbell et al. (2007)

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2000). The application of hydrostatic pressure to isolated articular chondrocytes cultured in monolayer (Parkkinen et al., 1995) and alginate (Fioravanti et al., 2005), or dynamic compression of cells embedded in agarose (Knight et al., 2006) reorganises the F-actin cytoskeleton. A polygonal arrangement of F-actin was observed throughout the cytoplasm of non-loaded cells however, this organisation was lost under increasing levels of hydrostatic pressure (Parkkinen et al., 1995; Fioravanti et al., 2005). Also, the number of cells containing an intact F-actin network decreased in cells subjected to a cyclic pressure of 15 MPa, with a near total disappearance of the F-actin networks, concomitant with retraction of the cells, after exposure to 30 MPa (Parkkinen et al., 1995). Interestingly, a recovery period of 2 h postload was sufficient to reverse the effect of hydrostatic pressure on the F-actin network. In contrast, the application of compressive (10 or 15%, 1 Hz) or static compression (15%) on chondrocytes resulted in F-actin remodelling, with a more punctate organisation replacing the uniform cortical distribution observed in nonloaded cells (Knight et al., 2006). However, a recovery period of 1 h was sufficient to reorganise the actin filaments to that observed prior to loading. An increase in punctate actin cytosolic features was also observed in cyclically compressed chondrocytes embedded in agarose (0–10% strain, 0.5 Hz, 20 min) (Haudenschild et al., 2008a, b). The actin cytoskeleton is also sensitive to mechanical distortions as a consequence of osmotic loading (Erickson et al., 2003; Chao et al., 2006). Prolonged mechanical compression or release of compression can alter the osmotic environment of the chondrocyte; manipulation of the osmotic pressure to 250 mOsm in vitro resulted in F-actin rearrangement from a predominantly cortical distribution to the severing and subsequent detachment from the chondrocyte periphery (Erickson et al., 2003; Chao et al., 2006). Conversely, exposure to hypo-osmotic stress led to a progressive disorganisation of F-actin followed by a gradual reorganisation (Erickson et al., 2003; Chao et al., 2006). Clearly, the F-actin cytoskeleton is highly dynamic in nature, responding to mechanical stimulation by reversibly disassembling and reorganising the filaments to maintain the structural integrity of the chondrocyte. Furthermore, mechanically induced actin remodelling may provide a feedback mechanism through which mechanical stimuli can modulate chondrocyte mechanosensitivity (Knight et al., 2006). The mechano-responsiveness of the F-actin cytoskeleton is likely mediated by the integrins, which form a link between the extracellular matrix and the cell. Integrins are heteromeric transmembrane glycoproteins, consisting of a combination of α- and β-subunits, of which the α5β1 integrin is a major mechanoreceptor in articular chondrocytes (Wright et al., 1997). The integrin receptor has an extracellular domain that acts as a ligand binding site, and a cytoplasmic tail which interacts with intracellular molecules, including the actin cytoskeleton, enabling the integrin receptor to transduce mechanical signals into intracellular biochemical responses (Hynes, 1992). Integrin ligand binding, in response to mechanical stimulation (3700 μstrain, 0.33 Hz, 1–5 min), influences F-actin assembly concomitant with phosphorylation of actin accessory proteins e.g. FAK and paxillin (Lee et al., 2002) and activation of downstream intracellular signalling cascades e.g. IL-4 signalling

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(Millward-Sadler et al., 2000). Included in the downstream signalling cascades is the Rho GTPase pathway which was activated in cyclically-compressed (0–10% strain, 0.5 Hz, 20 min) human articular chondrocytes embedded in agarose (Haudenschild et al., 2008a, b). Rho activation was followed by Rho kinase (ROCK)-dependent rearrangements of the actin cytoskeleton and by ROCK-dependent changes in gene expression (Haudenschild et al., 2008a). The Rho GTPases promote Factin rearrangement and enhance the formation of focal adhesions, creating a link between the cytoskeleton and the matrix. A major effector pathway downstream of Rho is the activation of ROCK, which phosphorylates and activates Lim kinase, which in turn phosphorylates and inhibits the actin-depolymerising protein cofilin. Chondrocyte deformation is accompanied by hydrostatic pressure gradients, fluid flow and streaming potentials generated by the displacement of water and ions from the matrix (Urban et al., 1993). A local change in ion concentration is sufficient to activate ion channels; stretch-activated Ca2+ -activated K+ channels were shown to be active in compressed chondrocytes (Hall, 1999; Roberts et al., 2001). The mechanical induction of intracellular Ca2+ release by the integrins is required for the association of the actin accessory proteins (FAK, paxillin) with the F-actin cytoskeleton. This effect may result from a requirement of Ca2+ ions for the activity of actin-binding proteins and other actin components in load-induced actin reorganisation. Actin-Binding Proteins Actin remodelling in cartilage chondrocytes subjected to a mechanical stimulus has been shown to involve the differential expression of the actin-binding proteins (Blain et al., 2002, 2003; Knight et al., 2006; Campbell et al., 2007). We have previously shown that thymosin β4, an actin-binding protein, is regulated by mechanical load in articular chondrocytes (Blain et al., 2002, 2003). Thymosin β4 mRNA levels were significantly elevated (20-fold) in articular cartilage explants subjected to a dynamic low-physiological load (0.5 MPa, 1 Hz, 10 min); the response was transient as the thymosin β4 mRNA signal returned to basal levels after 60 min of load (Blain et al., 2003). In unloaded explants chondrocytes, thymosin β4 protein was cytoplasmically diffuse, but after 60 min of load, the protein was organised into punctate loci throughout the cytoplasm; the distribution of thymosin β4 coincided with a more abundant immunolabelling of F-actin in the loaded explant chondrocytes (Blain et al, unpublished data). Thymosin β4, an actin sequestering protein, actively disassembles the actin filaments by binding in a 1:1 complex with Gactin, thereby preventing its availability for polymerisation (Weber et al., 1992). Other actin-disassembling proteins are also up-regulated by cyclic compression in chondrocytes. mRNA levels of cofilin and destrin were significantly increased after 10 min of mechanical stimulation (15% strain, 1 Hz) in chondrocytes embedded in agarose, concomitant with a temporal disassembly of cortical F-actin (Campbell

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et al., 2007). Increased cofilin expression concomitant with F-actin disassembly may be mediated by the regulation of the Rho kinase pathway (Haudenschild et al., 2008a). 4.4.2.2 Vimentin Intermediate Filaments Cartilage chondrocyte vimentin networks are also sensitive to mechanical perturbations in the surrounding environment. Physiological loading conditions induce vimentin filament assembly (Eggli et al., 1988; Durrant et al., 1999; Lahiji et al., 2004), whereas the application of abnormal, non-physiological loads promote filament disassembly (Karjalainen et al., 2003; Henson and Vincent, 2008). In vivo, increased vimentin content was observed in the more prominent weight-bearing regions of young rabbit articular cartilage (Eggli et al., 1988). In vitro, reorganisation of the vimentin network was observed in cartilage explant chondrocytes subjected to increasing levels of physiological load (0–4 MPa, 1 h); with minimal load, the explant chondrocytes behaved as free-swelling cultures as characterised by vimentin network disassembly (Durrant et al., 1999). Interestingly, the vimentin network in human osteoarthritic cartilage chondrocytes also reassembled when subjected to cyclic strain (Lahiji et al., 2004). In contrast, when a non-physiological single impact load (0.16 J) was applied to cartilage explants, vimentin disassembly was observed in the “injured” chondrocytes as evidenced by peri-nuclear collapse (Henson and Vincent, 2008). Vimentin mRNA levels were also significantly decreased in human chondrosarcoma cells subjected to a non-physiological continuous or cyclic (0.5 Hz) strain (5 MPa) (Karjalainen et al., 2003). 4.4.2.3 Tubulin Microtubules Very few studies have been conducted on the effect of mechanical load on the chondrocyte tubulin cytoskeleton (Jortikka et al., 2000; Trickey et al., 2004; Fioravanti et al., 2005). Tubulin network organisation was altered in chondrocytes exposed to a continuous hydrostatic pressure (24 MPa, 3 h) (Fioravanti et al., 2005), but appeared unaffected in response to a slightly lower hydrostatic pressure (> 15 MPa, 20 h) (Jortikka et al., 2000). Chemical disassembly of the chondrocyte tubulin cytoskeleton (using colchicine) had no discernable effect on the mechanical properties of the cell, suggesting that the F-actin and vimentin architectures provide the viscoelastic properties of the chondrocyte (Trickey et al., 2004). However, α-tubulin, the principal cytoskeletal component of the mechano-sensing cilium is responsive to strain. In a very recent study, it was demonstrated that the incidence and length of cilia i.e. acetylated α-tubulin increased in chondrocytes seeded in agarose subjected to cyclic compressive strain (0–15%, 1 Hz) (McGlashan et al., 2010). It was previously determined that approximately 50% of chondrocyte cilia are decorated with connexin 43 (Knight et al., 2009); further to this, the expression of a range of purine receptors (P2X and P2Y receptor subtypes) were also identified through which

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ATP activates downstream signalling events, suggesting a mechanism by which mechanical load activates ATP release as part of a purinergic mechanotransduction pathway involving the cilium (Knight et al., 2009).

4.5 Cytoskeletal Elements in Cartilage Chondrocyte Pathology Abnormal mechanical load, be it highly repetitive, injurious or the absence of, can result in degeneration of the articular cartilage; continued dissolution of the tissue can ultimately lead to the development of osteoarthritis (OA). OA is a multifactorial disorder, but one of the primary risk factors involved in its progression is inappropriate mechanical loading of the synovial joint (Cooper et al., 1994). OA is characterised by a loss of cartilage tissue homeostasis where catabolic events exceed synthesis culminating in a loss of matrix. OA is associated with altered chondrocyte gene expression (Reginato and Olsen, 2002) and metabolism i.e. reduced proteoglycan and type II collagen synthesis, compounded by increased secretion of pro-inflammatory cytokines and proteases (Goldring, 2000).

4.5.1 Cytoskeletal Element Organisation in Osteoarthritic Cartilage Chondrocytes Cell morphology, via cytoskeletal element organisation, influences cartilage chondrocyte metabolism (Ingber et al., 1994). Chondrocytes are normally devoid of actin microfilament bundles with F-actin localising to the cell periphery (refer to Section 4.3 for detail); the predominantly cortical distribution of F-actin ensures that the cell maintains a rounded morphology for preservation of the chondrocyte phenotype (von der Mark et al., 1977; Benya and Shaffer, 1982). Morphological differences between the cytoskeletal element(s) organisation in normal and OA chondrocytes has been hypothesised to influence the cells’ biochemical responses to external stimuli e.g. mechanical load, pro-inflammatory cytokines. In recent years an association, either directly or indirectly, between the presence of disorganised cytoskeletal networks in chondrocytes and cartilage pathology has been alluded to (Fioravanti et al., 2003; Capin-Gutierrez et al., 2004; Holloway et al., 2004; Lambrecht et al., 2008). Key morphological features of cytoskeletal element organisation in osteoarthritic cartilage chondrocytes are described (Table 4.1B). Characterisation of the cytoskeleton of human osteoarthritic cartilage chondrocytes, using transmission electron microscopy, demonstrated that the intensities and distribution patterns of the F-actin, vimentin and tubulin elements contrasted with observations in normal cells (Kouri et al., 1998). This early study was one of the first to hypothesise that a modified cytoskeleton might contribute to the altered phenotype in OA (Kouri et al., 1998). It was subsequently demonstrated that F-actin was apical in normal human chondrocytes, but was cytoplasmically diffuse or limited to the cell periphery in human OA chondrocytes (Fioravanti et al., 2003). Although tubulin was uniformly localised at the periphery of normal chondrocytes, it was

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often absent in OA cells (Fioravanti et al., 2003). Fioravanti et al. postulated that the loss of cytoskeletal element assembly in OA chondrocytes could compromise, not only the metabolic activities of the cells, but also their biomechanical function (Fioravanti et al., 2003). Disparities in cytoskeletal element expression and/or organisation are not exclusive to human pathology but have also been reported in animal models of OA. In a rat partial menisectomy model of OA, there was a reduction in the number of OA chondrocytes immuno-positive for F-actin (4.7%) and tubulin (20.1%) expression (Capin-Gutierrez et al., 2004). Concomitant with this was the observed reduction in vimentin expression (37.1%). Vimentin filaments which predominantly localised to the superficial zone, ranged from a simple disorganisation to a total disruption of cytoplasmic architecture; this contrasted with the uniform distribution of vimentin filaments as perinuclear bundles in normal rat chondrocytes (Capin-Gutierrez et al., 2004). The vimentin cytoskeleton presented as a complex fibrous structure extending throughout the cytoplasm in both normal and OA human chondrocytes (Holloway et al., 2004). Interestingly, multiple elongated processes containing elaborate vimentin filament networks, extending up to 30 μm into the surrounding matrix, were also observed in the OA cartilage which corresponded to sites with the greatest histological and macroscopic signs of OA (Holloway et al., 2004). Thus, the aberrant arrangement of the cytoskeletal networks in OA cells will impact on the mechanosensitivity of the cell, and the downstream signalling cascades activated as a consequence.

4.5.2 Mechanotransduction in Osteoarthritic Cartilage: Effect(s) on the Cytoskeleton Chondrocyte stiffness i.e. the mechanical properties of the cell, is an important indicator for how a cell will respond to mechanical perturbations in the surrounding matrix environment (refer to Section 4.1). The observed aberration in cytoskeletal element organisation in OA chondrocytes impacts on the mechanical properties of the cell. Significant increases in elastic and viscous properties were observed in OA chondrocytes from end-stage pathology (Trickey et al., 2000). Interestingly, a recent study reported that the mechanical properties of cartilage chondrocytes changed profoundly with ageing, and not OA pathology (Steklov et al., 2009). Viscoelastic properties increased in aged cartilage (< 55 years of age), but there were no apparent differences between the aged and age-matched OA cartilage. Clearly, the biomechanical properties of the cartilage are dependent on cytoskeletal element organisation irrespective of whether age and/or pathology are/is the confounding influence. An alteration to the biomechanical properties of the cell i.e. as a consequence of cytoskeletal element aberrations, undoubtedly affects mechanotransduction responses. Cyclical pressure (1–5 MPa, 0.25 Hz, 3 h) did not affect F-actin or tubulin distribution in OA chondrocytes, although the elements were subject to reorganisation in normal chondrocytes (Fioravanti et al., 2005). It was suggested

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that the cytoskeletal structures, which are not assembled appropriately in OA cartilage (Fioravanti et al., 2003), were unable to respond to similar loading conditions, possibly becoming “desensitised”. This could compromise the metabolic activities of the chondrocytes and hence the biomechanical integrity of the cartilage leading to pathology (Fioravanti et al., 2005). However, it was demonstrated in OA tissue that the incidence and length of cilia, detected using an anti-acetylated tubulin antibody, increased at the eroding articulating surface, resulting in an overall increased proportion of ciliated cells with OA severity (McGlashan et al., 2008). This study (McGlashan et al., 2008) would argue against the hypothesis that the tubulin cytoskeleton becomes desensitised to mechanical stimulation in pathological tissue (Fioravanti et al., 2003). The mechanisms of cartilage chondrocyte mechanotransduction have also been demonstrated to differ between normal and OA cells. As previously discussed (Section 4.2.1), a mechanical stimulus is perceived by the chondrocyte and α5 β1 integrin-mediated signalling is activated (Salter et al., 2001). This involves the opening of stretch-activated channels, an alteration in membrane potential and the rapid phosphorylation of focal adhesion proteins e.g. FAK125 , paxillin and β-catenin concomitant with F-actin treadmilling and the activation of downstream signalling cascades e.g. PKC (Lee et al., 2000b). However, although the initial response of OA chondrocytes to mechanical stimulation involved activation of α5 β1 integrin, the mechanotransduction route was independent of the F-actin cytoskeleton (Millward-Sadler et al., 2000), which may imply an adaptation by the cell to circumvent an aberrant F-actin network. Compressive stimulation activated an integrin-dependent interleukin-4 (IL-4) autocrine/paracrine loop in normal chondrocytes, whereas mechanotransduction was mediated via the activation of an IL-1β autocrine/paracrine loop in OA cells (Salter et al., 2002). Interestingly, IL-1 has been shown to increase F-actin amounts in chondrocytes (Pritchard and Guilak, 2006) suggesting that a finely balanced interplay of soluble factors e.g. cytokines may exist to regulate cytoskeletal element dynamics and hence organisation. Recently, a whole genome array demonstrated that IL-1β inhibited the mRNA expression of both tubulin and vimentin, as well as the LIM protein FHL2 which is associated with the actin cytoskeleton (Joos et al., 2008). Therefore in an inflamed or pathological environment i.e. in an OA synovial joint, the organisation and expression levels of these cytoskeletal proteins may be compromised which, in turn, will adversely influence the biomechanical properties preventing normal modes of cartilage chondrocyte mechano-signalling.

4.6 Conclusions and Perspectives The cytoskeleton acts as a physical interface between the chondrocyte and the extracellular matrix in “sensing” mechanical stimuli, and the cytoskeletal elements are themselves subject to load-induced reorganisation. Although several mechanisms of mechanotransduction involving the three major cytoskeletal elements have been

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elucidated in articular cartilage chondrocytes there is undoubtedly much still to discover. The reported loss of cytoskeletal element expression and/or organisation in OA chondrocytes has profound effects on cell morphology, negatively impacting on both its biological and biomechanical functions within the cell. To determine whether loss of cytoskeletal element architecture is a cause or effect of abnormal mechanical load, we are currently assessing whether there is a “threshold” above which load can disassemble the cytoskeleton. However, many more questions remain to be answered to elucidate the signalling pathways that are activated downstream of load-induced cytoskeletal element reorganisation, and how non-physiological loads may adversely impact on mechanotransduction pathways leading to matrix dissolution and pathology. Acknowledgements I wish to acknowledge the Arthritis Research Campaign (Grant No. 18221) for funding.

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Part II

Molecular Mechanisms of Mechanotransduction and Ion Channels Modulation

Chapter 5

The Role of Nitric Oxide in the Regulation of Mechanically Gated Channels in the Heart Victor Kazanski, Andre Kamkin, Ekaterina Makarenko, Natalia Lysenko, Natalia Lapina, and Irina Kiseleva

Abstract The article presents the effects of NO on myocardial functions including its pronounced influence on myocardium contraction and heart rhythm. Attention is given to cell signaling of nitric oxide in the heart. It is demonstrated that in general the final effect of NO depends on the cellular source of NO, amount of NO release, the prevailing redox balance and antioxidant status, stimuli such as coronary flow rate and heart rate, the target tissue, interaction with neurohumoral and other stimuli, activity level of the immune system and activation of cGMP-dependent and independent intracellular cascades. A number of experiments conducted on whole hearts lets us suppose that NO and NO-synthases as NO origins, directly regulate the conductivity of mechanically gated channels (MGCs). This study discusses experimental data obtained from isolated ventricular myocytes of mouse, rat and guinea pig by means of patch-clamp in the whole-cell configuration about the role of NO in the regulation of MGCs. Presented data demonstrate that NO donors lead to MGCs activation and appearance of MG-like currents in unstretched ventricular myocytes, while in stretched cells with activated MGCs NO donors lead to inactivation and inhibition of the conductivity of these channels. The NO scavenger PTIO causes inactivation of all MGCs. In unstretched cells the conductance through MGCs is blocked, which is present in control before deformation. PTIO causes complete inhibition of stretch induced MG-current during presence of cellular stretch. Application of non selective inhibitors of NO-synthases L-NAME or L-NMMA resulted in a complete blockade of MGCs. The presented data are instituted on cells of transgenic mice. In ventricular myocytes of wild-type mice, NOS1–/– and NOS2–/– stretching of cells results in an activation of typical MG-currents. On the contrary, in cells from NOS3–/– mice stretch does not activate MG-currents. The results suggest that NO plays an important role in the activation and inactivation of MGCs in cardiomyocytes and demonstrate that NOS3 dominates as NO origin. A. Kamkin (B) Department of Fundamental and Applied Physiology, Russian State Medical University, Ostrivitjanova 1, Moscow 117997, Russia e-mail: [email protected]


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Keywords Heart · Nitric oxide · Myocardium contraction · Heart rhythm · Isolated ventricular myocytes · Mechanically gated channels · PTIO · L-NAME · L-NMMA · NO donors · NOS–/–mice

5.1 Introduction Nitric oxide (NO) is a universal biological regulator contained in practically all human body tissues. NO was first identified in 1987 as an endothelium-derived biological messenger causing dilatation of blood vessels (Ignarro et al., 1987). Since then the amount of works devoted to NO has been growing fast. The “Science” Journal proclaimed it in 1992 as the molecule of the year. The 1998 Nobel Prize in physiology and medicine went to F. Murad, R. Furchgott and L. Ignarro, for discovering the functional role of NO in the cardiovascular system. That was the first boost to the gargantuan amount of publications devoted to research of the functional qualities of this simple chemical compound. NO possesses an obvious functional advantage over most biological regulators, having a small molecular weight and no charge, enabling it to quickly diffuse and freely penetrate through tight cellular layers and the intracellular space (Hughes, 2008). Going through the plasma membrane, NO acts not only as an intercellular transmitter but also as a part of intracellular effectoral systems like other known second messengers. NO has an unpaired electron, possesses high chemical activity and easily reacts with numerous cell structures and chemical components (Hughes, 2008). For these reasons, NO insures exclusively diverse biological effects. This gas is highly toxic in big concentrations, but in low or moderate concentrations it has a wide range of regulatory effects (reviewed in Butler et al., 1995; Marletta et al., 1994; Nathan and Xie, 1994; Schmidt et al. 1994; Stamler et al., 1994; Liaudet et al., 2000; Bruckdorfer, 2005). In human and animal organisms NO is produced as a result of enzymatic oxidation of N-terminal L-arginine (Moncada et al., 1991; Mayer and Hemmens, 1997; Hughes, 2008):

At the first stage, the hydroxylation of L-arginine occurs forming NG -hydroxyL-arginine, which oxidizes to NG -hydroxyarginine at the next stage. When the latter splits, NO gets free and intermediate carbodiimide forms that hydrolyzes to citrulline. At each of the two main stages of catalysis, one NADPH and one oxygen molecules are used. The peroxidate of heme is the intermediate product of the catalytic process (Marletta et al., 1994; Korth et al., 1994; Mayer and Hemmens, 1997; Hughes, 2008).

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The NO synthesis is carried out by a family of ferments – NO-synthases (NOS) that are presented in 3 different isoforms, named according to the place of their discovery: (1) nNOS, subtype NOS1 (neuronal or brain), (2) iNOS, subtype NOS2 (inducible or macrophagal), (3) eNOS, subtype NOS3 (endothelial) (Moncada and Higgs, 1993; Feron et al., 1996; Mayer and Hemmens, 1997; Xu et al., 1999). NOS differ in their localization in the cell, activity regulation and substrate inhibitory profile (Nathan and Xie, 1994; Moncada and Higgs, 1993). According to their induction character and activity NOS are divided into 2 types: more active – calcium-independent inducible NO-synthase (NOS2); and less active – calciumcalmodulin-dependent constitutive NO-synthases (NOS1 and NOS3) (Moncada and Higgs, 1993; Mayer and Hemmens, 1997). In the heart the key role in NO level regulation plays the constitutive NOsynthases. Thus, in mammals NOS1 is discovered in cardiomyocytes (Xu et al., 1999; Papapetropoulos et al., 1999; Damy et al., 2003; Ziolo et al., 2008), as well as in the preganglionic and postganglionic fibers innervating sinoatrial and atrioventricular nodes, subepicardial and endocardial neuronal cells. NOS3 is mainly expressed in cardiac vessels and endocardial endotheliocytes, less – in cardiomyocytes and sinoatrial and atrioventricular node cells (Papapetropoulos et al., 1999; Shah and MacCarthy, 2000; Ziolo et al., 2008). As opposed to NOS1 that is found in the perimembrane area of cardiomyocite sarcoplasmic reticulum, NOS3 is located in caveoles (Feron et al., 1996; Xu et al., 1999; Williams et al., 2006; Ziolo et al., 2008), which play a significant role in limiting NO diffusion in heart cells with high concentration of myoglobin (that binds NO with high affinity) and, what is especially important in pathological processes, of superoxide-anions (that react with NO and restrict its biological activity) (Casadei and Sears, 2003). Also, inducible NOS2 was discovered in the heart which is expressed in infiltrating inflammatory cells, microvessels, endocardial endotheliocytes, vascular smooth muscle cells, fibroblasts and cardiomyocytes, often simultaneously with inflammatory cytokines (Papapetropoulos et al., 1999; Shah and MacCarthy, 2000; Casadei and Sears, 2003). The wide presence of NO-synthases in different types of cardiac cells proves the necessity of nitric oxide for heart functioning. During the recent 20 years, the role of NO as an important agent in myocardial function regulation has become clear. It includes such effects like contractile activity, energy balance, substrate metabolism, cellular growth and survival in physiological and pathophysiological conditions (Shah and MacCarthy, 2000; Casadei and Sears, 2003). In this review, rather than discussing the known effects of nitric oxide, we will discuss in detail its effects on mechanically gated channels in heart.

5.2 Nitric Oxide and Cardiac Function Since the first report on NO effects on myocardial contractility in 1991 (Smith et al., 1991) a lot of data, often contradicting, have been accumulated on NO effects on heart functions. In general it can be said that the final effect of this regulator depends

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on the following: (1) the cellular source of NO, (2) amount of NO release, (3) prevailing redox balance and antioxidant status, (4) stimuli such as coronary flow rate and heart rate, (5) the target tissue, (6) interaction with neurohumoral and other stimuli, (7) activity level of immune system or disease, and (8) activation of intracellular cGMP-dependent and independent subcellular cascades (Shah and MacCarthy, 2000). NO takes part in regulating cardiac functions already at the embryogenesis stage. NO affects the growth and development of cardiac cells in fetal and postnatal lives (Strijdom et al., 2009). Moreover, several authors have considered NO as the key signaling molecule in early embryonic development of the heart, at least in rodents (Malan et al., 2004). NO is cyclically secreted in the working heart with fast growing concentration during early diastolic filling, which can be connected to the autoregulation during the cardiac cycle (Pinsky et al. 1997). Endogenous NO causes different effects in the heart, one of which is early relaxation of the left ventricle, that was shown on isolated heart and isolated cardiomyocytes (Shah et al., 1994; Casadei and Sears, 2003). It was discovered that specific inhibition of NOS1 or destruction of NOS1 genes results in increased left ventricular contractility in vitro and in vivo (Ashley et al., 2002; Casadei and Sears, 2003; Dawson et al., 2005). Clinical studies have also shown that under NO there were changes in the onset time of left ventricle relaxation, that manifests itself in an earlier onset of relaxation and decreased peak and end-diastolic pressure of the left ventricle (Paulus et al., 1994). Stimulation of NOS3 activity in healthy patients is also associated with changes in left ventricular end-diastolic pressure–volume relationship. This suggests a sharp decrease of left ventricular stiffness (Paulus et al., 1994). NO also has a pronounced effect on myocardial contractility and heart rate, and these NO effects have biphasic and concentration-dependent character. Low concentrations of NO-donors (0.1–10 μM) increase myocardial contractility and heart rate, while high concentrations (over 100 μM) produce negative inotropic and chronotropic effects (Koja at al., 1996, 1997; Vila-Petroff et al., 1999; Brady et al., 1993; Shah and MacCarthy, 2000; Casadei and Sears, 2003). It should be noted that the positive inotropic effect is close to maximum, within the range of NO concentrations corresponding to those in the normal heart with intact endothelium (e.g., 1–3 μM, according to Pinsky et al., 1997) and therefore reflects the physiological role of NO in healthy hearts (Casadei and Sears, 2003). Supporting this hypothesis is the fact that coronary infusion of certain concentrations of NO-synthase inhibitors into isolated rat hearts and healthy persons causes a small but reliable decrease of left ventricle contractility (Kojda et al., 1999; Cotton et al., 2001). NO plays an important role in regulating myocardium reaction to mechanical stimuli, in particular to stretch. Myocardium response to stretch includes significant and fast increases of contractile force (Frank-Starling response), followed by a slower and less pronounced increase of contractility (Anrep effect). NO can be involved in the realization of both effects. Endogenous NO strengthens the Frank-Starling response probably by increasing diastolic distensibility (Shah and MacCarthy, 2000; Casadei and Sears, 2003; Zhang et al., 2009). It was shown that

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the increases of NO production in the endothelium of coronary vessels or of intracardial infusion of NO-donors causes fast left ventricle relaxation and reduction of the end-diastolic resistance of left ventricle, that can affect the realization of the FrankStarling response by increasing end-diastolic volume of left ventricle and length of myocardial myofibriles (Paulus et al., 1995). Vice versa, the inhibition of NO synthesis blocks the increase of cardiac output in response to increased left ventricle loading in isolated hearts (Prendergast et al., 1997), which attests that stretchmediated stimulation of constitutive NO production in myocardium plays a role in regulation of cardiac output. An alternative autocrine mechanism for stretch induced increase of contractive activity is the Anrep effect. NO release in myocardium corresponds to increased levels of intracellular Sa2+ , which results in positive inotropic effect in response to stretch. This effect includes NOS3 activation as it is completely blocked by NO-synthase inhibition or NOS3-gene destruction (Petroff et al., 2001). Endogenous NO affects the expression of force-frequency relationship in cardiomyocytes. Direction and expression of these effects depend on the experimental conditions. Thus, positive force-frequency relation in myocardium is suppressed by endogenous NO: this relation significantly rises after administering NOS1inhibitors in rat cardiomyocytes. The opposite effect is observed in vivo in NOS1–/–mice. This testifies towards the ability of NOS1-produced NO to increase the positive force-frequency relation (reviewed in Shah and MacCarthy, 2000; Casadei and Sears, 2003). NO is involved in the regulation of the response to adrenergic and cholinergic stimulation in the heart. Thus, NO-donors modulate β-adrenergic inotropic effects, decreasing it in small doses and increasing it in high doses (Kelly et al., 1996; Shah and MacCarthy, 2000; Massion and Balligand, 2003). It was also shown that increased myocardial expression of NOS3 in pathophysiological conditions results in inhibiting β-adrenergic reactivity (Casadei and Sears, 2003). Moderate myocytespecific NOS3 overexpression results in inhibiting α-adrenergic responses, but these data are not confirmed in animals with high level of cardiomyocyte-specific NOS3 overexpression (Casadei and Sears, 2003). NO also mediates the effects of cholinergic stimulation of the heart: the first direct confirmation of the fact was given in the work presented by Smith T.W. et al. (1995), similar data were received later using NOS-inhibitors and NOS–/–mice (Shah and MacCarthy, 2000). One more component of NO-mediated regulation of cardiac activity is its effect on energy and metabolism qualities of myocardium. Paracrine NO release from the endothelium of microvessels causes reversible inhibition of cardiac muscle oxygen consumption (Shen et al., 1994; Trochu et al., 2000). Thus, stimulation of NO release, by e.g., bradykinin, lowers mitochondrial respiration, and administration of NO-synthase inhibitors results in increased total oxygen consumption in the heart irrespective of changes of haemodynamics and contractile activity (Shen et al., 1994, 1995). Decreased oxygen consumption in the heart is connected with increased substrate utilization of free fatty acids into glucose. Besides, there are data that NO inhibits glycolysis (Zhang and Snyder, 1992). One of the most important nitric oxide functions is modulation of cell survival and death: it can both assist cell death and prevent it. The double role of NO in cell

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death depends on its concentration inside the cell and interaction with other biological molecules, such as oxygen and superoxide (Tsang et al., 2004; Calabrese et al., 2009). In low concentrations NO produces cardioprotective, anti-apoptotic, anti-hypertrophic effects protecting heart cells from damage caused by pathological influences, like, for example, ischemia. In high concentrations NO is harmful when it is presented in excessive amounts: pro-apoptotic, pro-necrotic effect on cardiomyocytes (Casadei and Sears, 2003; Strijdom et al., 2009; Calabrese et al., 2009). NO is involved in the regulation of cardiac activity in pathophysiological conditions. Thus, expression and activity of NO-synthases increases in rat and mouse myocardium under experimental infarction, as well as in humans with heart diseases (Wildhirt et al., 1995; Casadei and Sears, 2003). NOS1 and NOS3 produced NO influences contractility, energy balance and gene expression in myocardium (e.g., with cardiomyopathies, infarctions), that can be an adaptive effect protecting the injured heart from harmful influences of excessive catecholamine stimulation, as well as from free radicals (Michel and Smith, 1993; Paulus, 2001). NOS2 participates in developing immune-mediated heart diseases, e.g., infarction. NOS2 produced NO, on the contrary, causes apoptosis and is involved in inflammatory processes under functional pathology of heart (Michel and Smith, 1993; Wildhirt et al., 1995).

5.3 The Role of Nitric Oxide in the Regulation of Mechanically Gated Channels in Isolated Ventricular Cardiomyocytes from Guinea Pig, Rat and Mouse Mechanical stretch is an important physiological and pathological stimulus in the heart. It is well known that stretch increases myocardial contractility, but the mechanisms underlying this effect are not yet clear (Zhang et al., 2009). Thus, the mechanical events can modify electric processes in the cardiomyocite membrane by direct effects on mechanically gated channels (MGCs) (Kamkin et al., 2000). It was shown in a number of works that mechanical stimuli (e.g., circumferencial or longitudinal stretch) cause increased release of endocardial NO from vascular endothelium and left ventricle cardiomyocites (Pinsky et al., 1997, Petroff et al., 2001). Besides, in response to stretch in cardiac cells NO-synthase activity significantly increases, in particular NOS3 (reviewed in Shah and MacCarthy, 2000; Seddon et al., 2007). In Part 2 of this article we have described in brief the participation of NO in heart response to stretch. One of the possible mechanisms of realizing cardiomyocites reaction to mechanical stress is the stretch stimulated NO effect on MGCs. Still, the NO input into regulating the activity of mechanically gated currents (MG-currents) has been hardly studied up till now due to serious methodical difficulties of such experiments, although studies in this field have been conducted for a long time simultaneously by two groups. One group included G. Isenberg, V. Dyachenko and U. Rueckschloss and the second group included A. Kamkin, V. Kazanski and I. Kiseleva. The authors

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used similar methods of stretching cells and registering MG-currents. However Dyachenko et al. (2009b) preincubated isolated ventricular mouse myocytes with the PTIO, SNAP and L-NMMA (thus, not getting MG-currents control registration from each cell under recording) after which they tried to stretch the cells and obtain MG-currents. Preincubation of the cells means that the authors loaded drugs by storing the cells in drug containing KB-solution (see later) for the quoted period of time. In the works presented by Kazanski et al. 2010a, b (See also this chapter) ventricular myocites of young guinea-pigs (3 months old), young Wistar rats (3 months old) and mice were first perfused in physiological salt solution (PSS), and after registering the control MG-currents the perfusion with PSS containing the tested drugs was started. With that, at each stage any and every drug was first tested without deforming the cells, and secondly on the background of the previous deformation of the cells. In both cases we tried to wash out the drug. The perfusion chamber had at volume of 0.5 ml that was changed within 15–20 s. Ventricular myocytes were dispersed using the standard collagenase dissociation technique. Briefly, the isolated hearts were perfused retrogradely (36◦ C). After an initial 5-min period with Ca2+ -free PSS, a 15-min period followed where Ca2+ -free PSS was supplemented with 0.1% collagenase type II (Worthington, Lakewood, N.J., USA) and 20 μM CaCl2 . Finally, the ventricles were chopped and gently triturated to release the cells into “Kraftbrühe” (KB) medium (Isenberg and Klockner, 1982) containing (mmol/L) 30 KCl, 0.5 KH2 PO4 , 50 glutamic acid, 20 taurine, 10 glucose, 3 MgSO4 , 0.5 H4 EGTA, adjusted to pH 7.2 with KOH. The cell suspension was filtered, resuspended and kept in KB medium at room temperature for at least 2 h. Ventricular myocytes were perfused with (37◦ C) PSS containing (in mM) 150 NaCl, 5.4 KCl, 1.8 CaCl2 , 1.2 MgCl2 , 20 glucose, 5 HEPES/NaOH, pH 7.4 (Kout ). Patch pipette solution (Kin ) was composed of (in mM) 140 KCl, 5.5 MgCl2 , 5 Na2 ATP, 0.05 EGTA, 10 HEPES/KOH (pH 7.2). Patch pipettes had tip resistances between 1.8 and 2.2 M (Kamkin et al., 2000, 2003). The myocyte adhered with its bottom to the cover-slip. The patch pipette pressed the cell to the glass bottom. After whole-cell access of the patch pipette, a firepolished glass stylus was attached to the membrane (Kamkin et al., 2000, 2003). When the stylus was freshly polished and the surface membrane was clean, attachment succeeded in approximately 70% of attempts. The stylus was then lifted 2 μm to prevent “scratching” of the lower cell surface on the cover-slip during stretch. A motorized micromanipulator (MP 285, Sutter, Novato, Calif., USA, accuracy 0.2 μm) increased the pipette – stylus distance stepwise by up to 12 μm, with the pipette being the fixed point (Kamkin et al., 2000, 2003). Still, in most cases we used a standard stretch equal to 10 μm, as sufficiently “soft” for the cell. Stretch and release of stretch could be repeated 3–5 times with the same cell, on average. Currents were recorded with an RK-300 amplifier (Biologic, Echirolle, France), digitized (PowerCED, Cambridge Instruments, Cambridge, UK) and stored in a computer. Cells were voltage-clamped to a holding potential of –45 mV in order to inactivate voltage-dependent sodium currents. Changes in voltage-dependent

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membrane currents were studied with ramp-like repolarizations. To exclude a contribution of voltage-dependent calcium currents the membrane was first clamped to +50 mV for 50 ms and than repolarized from +50 to –100 mV at a rate of –100 mV/s. The resulting currents were plotted versus the potential as I-V-curves. Since the amplitude of the currents depends on cell length and diameter, cells of similar geometry were selected. The effect of a different size of the stretched membrane was minimized by adjusting the glass tools to the same 40 μm glass stylus – patch pipette distance prior to application of stretch. Since mechanical stretching of the cell was restricted to a small unknown area between the stylus and the patch pipette, we did not divide the mechanically gated currents (MG-currents) by whole membrane capacitance. Currents in response to trains of short 5 mV pulses (applied at −45 mV) were evaluated in terms of membrane capacitance (time integral) and the access resistance (time constant divided by time integral (Gillis, 2000)). Results are presented as mean ± S.D., n is the number of cells studied. Significant differences were detected by analysis of variance (ANOVA) with the Bonferroni test as post-hoc test. This data demonstrates that the result of measurements in the three animal groups do not differ significantly. There were no differences between experimental series conducted on ventricular myocytes of different animals but with the same drug. Before stretch, the I-V curve was N-shaped (Fig. 5.1a: IC ). The intercept of the I-V curve with the zero-current axis (Dyachenko et al., 2008) is the zero-current potential (V0 ) which is equivalent to the diastolic membrane potential under current clamp (V0 = −85 mV: mean±S.D. V0 = −85 ± 5 mV, n = 11). The data demonstrated that under ramp-like repolarizations of ventricular myocytes (in Kin /Kout solution) the hump of outward current at −60 mV was attributed to the inwardly

Fig. 5.1 Voltage dependence of membrane currents in a ventricular myocyte from young guineapig in control and under cell stretch. a – I-V curves recorded with repolarizing ramp commands before (IC ) and during 10-μm stretch (IS ). The main changes in the I-V curve during cell stretch include reduction of V0 (intersection of the curve with zero-current-axis), induction of a negative current at EK (equilibrium potential of potassium). The arrows show the direction of the I-V curve shift. Compared to the control, 10-μm stretch significantly reduces V0 , activates Ins and deactivates IK1 . b – Stretch-induced difference current (IS–C : the difference of IS minus IC ). Modified from Kazanski et al., 2010a with permission

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rectifying K+ -current IK1 . At potentials positive to −40 mV, curve fit required the superimposition of outwardly rectifying current components. As we have shown, the 10 μm stretch (1 min) shifted the I-V curve downwardly (Fig. 5.1a: IS ), reduced the hump of IK1 and depolarized V0 to −42 mV (−41±5 mV, n = 11). At the potassium equilibrium potential the potassium current IK1 is zero. Therefore, the stretch-induced negative current I(EK ) should be interpreted as stretch-induced non-selective cation current Ins (Dyachenko et al., 2008), otherwise called current through stretch activated channels − ISAC (Hu and Sachs, 1997; Kamkin et al., 2000), or current through mechanically gated channels − IMGC . (Zhang Y and Hamill OP, 2000; Hamill and Martinac, 2001; White, 2006; Kamkin and Kiseleva, 2008). I(EK ), the current at EK = −89 mV (61 mV log[5.4/155] = −89 mV), yielded a first estimate of the stretch-activated current Ins . In absence of mechanical deformation, Ins (−89 mV) was in the order of −0,03 nA (Dyachenko et al., 2009a). The stretch-induced difference current Is−c (Fig. 5.1b) reversed polarity at −15 mV (−11 ± 2 mV, n = 11). Fit of the net and difference current according to Ins = Gns (V − Ens ) and Ins = Gns (V − Ens ) with a voltage-independent Gns yielded a straight line. At potentials negative to −20 mV mechanosensitivity is composed of both Gns and GK1 . Gns has been reported to be voltage-independent and to operate with a reversal potential Ens =−10 mV (measurements with blocked K+ -currents (Kamkin et al., 2000, 2003). The deviation of the difference current from the straight line was attributed to the stretch-deactivation of GK1 because it was abolished by substituting extracellular K+ by Cs+ ions (Kamkin et al., 2000, 2003). At potentials positive to −20 mV, mechanosensitive currents are carried by K+ ions through both (Dyachenko et al., 2009a) TRPC6 – outwardly rectifying channels (Hofmann et al., 1999; Spassova et al., 2006; Onohara et al., 2006) and TREK (1 and 2) − K2P 2.1 and K2P 10.1 – leak channels (Honoré et al., 2006; Li et al., 2006; Patel and Honoré, 2005) which are mechanosensitive. Dyachenko et al. (2009a) proposed, that together with stretch-activated Gns (Kamkin et al., 2000, 2003), stretch-induced deactivation of Kir2.3 and activation of TRPC6 can destabilize the diastolic membrane, eventually leading to pacemakerlike depolarizations and extra systoles (Kamkin et al., 2000, 2003; Lozinsky and Kamkin, 2010; Zeng et al., 2000). After block of TRPC6 channels, stretch-induced depolarizations remained small suggesting that stretch-activation of TRPC6 (and not deactivation of Kir2.3 channels) is the key event Dyachenko et al. (2009a). Thus, due to mechanosensitive currents carried by K+ ions, the difference current Is−c can only conditionally be called the MG-curent. Due to the simultaneous activation of Gns , deactivation of Kir2.3, activation of TRPC6 and mechanosensitivity of TREK Dyachenko et al. (2008) have introduced the term “stretch modulation of ion currents” (SMIC: ISM ). But later these authors used the term “MG-current” (Dyachenko et al., 2009b). Not to complicate the terminology here and further on, in Figures and legends, we stick to the standard definition of a difference current as Is−c , and in presenting the data we use the term “MGcurrent”, as MGC make the main contribution to control I-V curve modulation under cell stretch (Dyachenko et al., 2009a). In our experiments the changes in

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MG-current increased with the amplitude of stretch and were reversible, i.e. they have disappeared upon relaxation from stretch. These data coincide with the data obtained under ramp-like repolarizations of ventricular myocyte Dyachenko et al. (2008). Since during cellular stretch (for example 10 μm) both Ins (Ins(Vo) = −0.50 nA: Fig. 5.1) and mechanosensitive currents carried by K+ ions contribute to the stretchinduced difference current Is−c , we calculated the value of stretch-induced currents at the level of the holding potential, referring to it as IMGC(−45 mV) (IMGC(−45 mV) = −40 nA: mean±S.D. IMGC(−45 mV) = −0.41±0.02. Fig. 5.1). It seemed to represent both MG-current and MG-like current.

5.3.1 NO Scavenger PTIO The NO scavenger PTIO or 2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-11-oxy-3-oxide was used for testing the involvement of NO in the regulation of MGcurrent activity. Dyachenko et al. (2009b) preincubated mouse ventricular myocytes for the period of 30 min in a solution of PTIO (100 μmol/L). Then they recorded the I-V relationship, altered by PTIO preincubation, and tried to stimulate MG-currents by application of 10 μm cellular stretch. Figure 5.2 shows the I-V curve registered after the PTIO preincubation (IPTIO ) and the I-V curve of the same cell registered after it was stretched by 10 μm (IS,PTIO ). The absence of any changes caused by stretch signifies the block of MG-currents on the background of PTIO. According to the authors’ data the I-V curve crossed the voltage axis (zero current potential V0 ) at −89 mV. It is a large enough resting potential for an isolated mouse cell. We presupposed that incubation of cells with PTIO can result in closing a number of MGCs that for certain reasons are activated in control, i.e. without artificial stretch of the cell.

Fig. 5.2 Preincubation of mouse ventricular myocytes with the nitric oxide scavenger PTIO (100 μmol/L, 30 min preincubation) blocks the induction of MG-currents by 10-μm stretch. (From Dyachenko et al., 2009b, with permission from Elsevier)

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First of all, we studied the reaction of the cell to PTIO. After whole-cell access of the patch pipette, a glass stylus was attached to the membrane but no stretch of the cell was conducted. After registering the control I-V curve we started perfusing the cell with PTIO solution (500 μmol/L). Such concentration of PTIO was used, for example, for studying the regulation of cardiac calcium current by NO (Gallo et al., 2001). Figure 5.3 shows the control I-V curves (IC ) as compared to I-V curves under PTIO (IPTIO ) without stretching the ventricular myocyte from guinea-pig after 1 min (Fig. 5.3a) and 3 min (Fig. 5.3b) of perfusion, and besides that the cell’s reaction to washing out PTIO (W IPTIO ) for the period of 5 min (Fig. 5.3c). As the crossing point of the I-V curve characterizes the diastolic membrane potential, it is obvious that under the PTIO effect V0 increased by approximately 10 mV from −84 mV (−84±5 mV; n = 10) to −94 mV (−94±2 mV; n = 10). PTIO shifted the I-V curve downwardly, reduced the hump of outward current and shifted the hump from

Fig. 5.3 Voltage dependence of membrane currents in ventricular myocytes from guinea-pig under the perfusion of unstretched cell by PTIO solution. a – I-V curve measured in control (IC ) and after 1 min PTIO perfusion (IPTIO ). b – After 3 min PTIO perfusion (IPTIO ) as compared to the control (IC ). c – After 5 min of PTIO wash out (W IPTIO ) as compared to control (IC ). Note: In panels (a) and (v) the arrows show the direction of the I-V curve shift under PTIO perfusion of the cell after 1 and 3 min after the beginning of the perfusion accordingly. Modified from Kazanski et al., 2010b with permission

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−60 mV to −80 mV (Fig. 3a,b: IPTIO ). The value of IPTIO−C is equal to −0.13 nA (IMGC(−45 mV) = −0.15 ± 0.02 nA, n = 10) for 3 min PTIO perfusion. As it is known that NO increases cardiac IK1 (Gómez et al., 2009), it is understandable that PTIO, reacting stoichiometrically with NO, decreases inwardly rectifying K+ currents − IK1 . It is shown that after 3 min of PTIO perfusion the 5 min long wash out did not result in any shifts of the curve towards the control, i.e. the unstretched cell did not show even any partial release from PTIO effect during the registered time. Similar data were registered from ventricular myocytes of mouse and rat. In the following series of experiments after registering the control I-V curve we first stretched the cell and registered the I-V curve. Then we perfused it with the PTIO solution (500 μmol/L) and registered the I-V curves during the perfusion. Figure 5.4 shows I-V curves registered from ventricular myocytes of

Fig. 5.4 Voltage dependence of membrane currents in ventricular myocyte from young guineapig under PTIO perfusion on the background of cell stretch. a – I-V curve measured before stretch (IC ) and during 10 μm stretch (IS ). The arrows show the direction of the I-V curve shift. b –, c – I-V curve modifications under PTIO perfusion with continued stretch for 2 and 8 min accordingly (IS,PTIO ). The arrows show the direction of I-V curve shift at the given cell stretch under PTIO perfusion. For better perception the panel D combines the control I-V curve (IC ) and the I-V curve under 8 μm cell stretch after 8 min of PTIO perfusion (IS,PTIO ). It is obvious that during the PTIO perfusion the activity of MGCs is inhibited. Modified from Kazanski et al., 2010b with permission and the author’s data

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young guinea-pig in control, under cell stretch and under further PTIO perfusion. The voltage dependency of IC and its modulation by stretch IS is shown in Fig. 5.4a. Before stretch (IC ), the I-V curve was N-shaped and crossed the voltage axis at −85 mV (−86 ± 5 mV, n = 10). The 10-μm stretch shifted the I-V curve downwardly (Fig. 5.4a: IS ), reduced the hump of inwardly rectifying K+ -current IK1 and depolarized V0 to −42 mV (−39 ± 4 mV, n = 10). Close to −10 mV, the I-V curves recorded before and during stretch crossed each other, and at positive potentials IS was increased. The stretch-induced difference current (IS−C ) – the difference of IS minus IC (Fig. 5.5a corresponding to Fig. 5.4a) reversed polarity at −15 mV (−12±3 mV, n=10) and equaled −0.41 nA (IMGC(−45 mV) = −0.40 ± 0.02 nA, n = 10). At potentials negative to −20 mV mechanosensitivity is composed of both Gns and GK1 . The deviation of the difference current from the straight line was attributed to the stretch-deactivation of GK1 . Thus, stretch increases the MG-currents and depolarizes the cell (Fig. 5.4a). Figure 5.4b shows the change of IS on the background of 10 μm cell stretch 2 min after beginning the PTIO perfusion (IS,PTIO ), and Fig. 5.4c demonstrates IS,PTIO under the same stretch but after 8 min of PTIO perfusion. Disregarding the continuous cell stretch PTIO perfusion returns the I-V curve to the original state, at which the curve got back to N-shaped and crossed the voltage axis at −84 mV (−86±4 mV, n=10). The stretch-induced difference current − current during stretch minus current during stretch with PTIO IS−S,PTIO (Fig. 5.5b corresponding to Fig. 5.4c) reversed polarity at −10 mV and showed deviation from the straight line and equals (+)0.33 nA IMGC(−45 mV) = (+)0.35 ± 0.03 nA, Erev =

Fig. 5.5 Stretch-induced difference currents from ventricular myocyte of young guinea-pig. a – Different current activated by 10 μm of stretch (the difference of IS minus IC ), corresponding to Fig. 5.4a (reversal potential Erev = −15 mV, IMGC(−45 mV) = −0.41 nA. b – Different current under PTIO perfusion on the background of myocyte stretch: the difference of the current during stretch (IS ) by 10 μm minus same stretch with PTIO (IS–S,PTIO ). Panel (b) corresponds to Fig. 5.4c (reversal potential Erev = −5 mV, IMGC(−45 mV) = (+)0.33 nA). Note: two curves for stretch of 10 μm (a) and stretch of 10 μm plus PTIO (b) show a nearly linear voltage dependence. Modified from Kazanski et al., 2010b with permission

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−2 ± 2 mV, n = 10)1 . Thus, PTIO perfusion on the background of cell stretch resulted in inhibiting MG-currents, and in partial restoration of IK1 . The I-V curve registered under those conditions is practically the same as the I-V curve under control (Fig. 5.4d). Only the hump of outward current at −60 mV was slightly decreased and smoothed. Similar data were obtained from ventricular myocites of mouse and rat. It is of interest, that under preceding cell stretch the PTIO perfusion did not result in the shift of the hump of outward current and V0 towards negative potentials. We considered a possibility of PTIO wash out at least at the first stages of the effect development. Figure 5.6 shows the experimental results obtained from mouse ventricular myocytes. The 10-μm stretch shifted the I-V curve downwardly, reduced the hump of IK1 , depolarized V0 and caused the MG-current (Fig. 5.6a: IS ). PTIO perfusion under up to 10 μm cell stretch resulted in progressed inhibition of MGcurrents (Fig. 5.6b: IS,PTIO during 2 min and Fig. 5.6c: IS,PTIO during 3 min). To check the possibility of PTIO wash out, it was started after 3 min, i.e. when the PTIO effect came into force. Interestingly, the wash out immediately prevented further inhibition of MG-currents but even after 5 min of wash out the curve remained unchanged (W IS,PTIO − Fig. 5.6d as compared to IS,PTIO – Fig. 5.6c). Thus, the treatment of PTIO for 3 min blocked a part of MSCs irreversibly, which manifested itself in a decrease of MG currents, while the other part continued to function under continuous stretch.

5.3.2 NO Donors 5.3.2.1 NO-Donor SNAP The NO-donor SNAP or S-Nitroso-N-acetylpenicillamine was used for testing the involvement of NO in regulating MG-current activities. First of all, we demonstrate the reaction of an undeformed cell to SNAP. After registering the control I-V curve we started perfusing the cell with SNAP (100 μmol/L). Figure 5.7a demonstrates the control I-V curve (IC ) registered in mouse ventricular myocyte and the I-V curve received 1 min after the SNAP perfusion started (ISNAP ) without any cell stretch. Figure 5.7b shows, as compared to the control curve, the I-V curve received 2 min after SNAP perfusion started (ISNAP ). Before perfusion of SNAP (IC ), the I-V curve was N-shaped and crossed the voltage axis (zero current potential V0 ) at −74 mV. It is obvious that SNAP perfusion shifts V0 from −74 mV (−85 ± 4 mV, n = 8)

1 Here

we must make the following note. The differential current that occurs under cell stretch as a result of I increase (e.g., the difference of IS minus IC ) or the differential current occurring as compared to the control under the drugs effect (the difference of IDrug minus IC ) was marked with a minus sign. To make it easier for the reader and to avoid confusion with the differential current under cell relaxation after stretch or the differential current resulting from I return registered in a stretched cell to the control under the drugs effect, the latter we mark conventionally with a plus sign in brackets (+).

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Fig. 5.6 Voltage dependence of MG currents in ventricular myocyte from mouse under PTIO perfusion of the cell and its wash out. a – I-V curve measured before stretch (IC ) and during 10 μm stretch (IS ). b-, c- modified I-V curve under PTIO perfusion with continuous stretch for 2 and 3 min accordingly (IS,PTIO ). In both panels IS characterizes MG-currents during 10 μm stretch. d – 5 min of PTIO wash out under stretch (W IS,PTIO ). Note that the I-V curve W IS,PTIO did not change as compared to I-V curve IS,PTIO in panel C. The author’s data

to −50 mV (52 ± 3 mV, n = 8) in 1 min and to −30 mV (−28 ± 4, n = 8) in 2 min. Besides, you can see a reduction of the slope of ISNAP at potentials negative to V0 which could be attributed to stretch-deactivation of GK1 (Dyachenko et al., 2009a). The appearing MG-like current equals to −0.38 nA and corresponds to the stretch induced current appearing at 10 μm cell stretch. The SNAP-induced difference current (ISNAP−C : during perfusion of SNAP minus control) plotted in the I-V relations in Fig. 5.7c, d (for 1 and 2 min, respectively) reversed polarity at −2 mV, and at 2 mV (near zero potential: Erev = 1 + 1 mV). The deviation of the difference current from a straight line was attributed to the stretch-deactivation of GK1 . The value of ISNAP−C is equal to −0.38 nA (IMGC(−45 mV) = −0.39 ± 0.02 nA, n = 8) for 2 min SNAP perfusion vs. IMGC(−45 mV) = −0.40 ± 0.02 nA, n = 11 during 10 μm cell stretch in control (P=NS). Thus, SNAP perfusion resulted in progressed activation of MG-currents.

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Fig. 5.7 Voltage dependence of membrane currents in ventricular cardiomyocytes from mouse under SNAP perfusion of unstretched cells. a – I-V curve measured in control (IC ) and after 1 min SNAP perfusion (ISNAP ). b – After 2 min SNAP perfusion (ISNAP ) as compared to the control (IC ). c – Difference current activated by SNAP ISNAP-C (the difference of ISNAP minus IC ), corresponding to Panel A 1 min after the beginning of perfusion (reversal potential Erev = –2 mV, IMGC(−45 mV) = −0.13 nA). d – ISNAP-C , corresponding to Panel B 2 min after the beginning of perfusion (reversal potential Erev = 2 mV, IMGC(−45 mV) = −0.38 nA). Note 1: In panels (a) and (b) the arrows show the direction of the I-V curve shift under SNAP perfusion of the cell. Note 2: After 2 min the different MG-like current (Panel d) corresponds to 10 μm cell stretch. Modified from Kazanski et al., 2010a with permission and the author’s data

After 2 min of SNAP perfusion the wash out for 2 min results in an I-V curve shift towards the control, i.e. the unstretched cell shows removal of SNAP effect during registration time (Fig. 5.8a). After 5 min of wash out the control I-V curve and the I-V curve received after SNAP wash out coincide completely, and the latter crosses the voltage axis at −72 mV (−82 ± 5 mV, n = 8) (Fig. 5.8b). The value

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Fig. 5.8 Voltage dependence of membrane currents in ventricular cardiomyocytes from mouse after SNAP wash out. a – After 2 min of SNAP wash out (W ISNAP ) as compared to the control (IC ) and to the perfusion of SNAP during 2 min (ISNAP ). b – After 5 min SNAP wash out (W ISNAP ) as compared to the control (IC ). The author’s data

of W ISNAP is equal to (+)0.39 nA (IMGC(−45 mV) = (+)0.40 ± 0.03 nA, n = 8) for 5 min SNAP perfusion (not shown). It is interesting that earlier Dyachenko et al. (2009b), who preincubated cells with SNAP (200 μmol/L), has shown that superfusion of cardiomyocytes with SNAP did not significantly activate Gns or deactivate GK1 . We demonstrate that MG-like currents (Gns ) in cells isolated from all the animals change quite significantly, as well as GK1 deactivation is observed. SNAP perfusion for approximately 2–3 min caused the same changes as 10 μm stretch. Since application of the NO-donor SNAP in the absence of mechanical stimulation failed to induce MG-like currents we concluded that availability of NO is crucial for MSC activation, and an increase in NO alone is sufficient for its activation. Further on we conducted a series of experiments in which we first stretched cells by 10 μm, and then perfused them in this stretched state with SNAP solution (100 μmol/L). Figure 5.9 shows the results of experiments carried out on ventricular myocytes from mouse. The voltage dependency of IC and its modulation by stretch (IS ) is shown in the I-V curves in Fig. 5.9a. Before stretch (IC ), the I-V curve was N-shaped and crossed the voltage axis at V0 = −70 mV (−85 ± 4 mV, m = 8). The 10-μm stretch shifted the net currents to more negative values (IS ), and V0 changed to −35 mV (−40 ± 5 mV, n = 8). Close to 10 mV, the I-V curves recorded before and during stretch crossed each other. The stretch-induced difference current (IS−C ) is plotted in the I-V relations in Fig. 5.9c and equals −0.46 nA (IMGC(−45 mV) = −0.42 ± 0.02 nA, Erev = −8 ± 2 mV, n = 8). The deviation of the difference current from a straight line was attributed to the stretch-deactivation of GK1 , because at

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Fig. 5.9 Voltage dependence of membrane currents in ventricular myocyte from mouse under SNAP perfusion on the background of the cell stretch. a – Current-voltage relation (I-V curve) measured before stretch (IC ) and during 10 μm stretch (IS ). The arrows show the direction of the I-V curve shift at the given cell stretch. b – modified I-V curves on the background of SNAP perfusion at continuous stretch during 2 min (IS,SNAP )and 3 min accordingly (IS,SNAP ). The arrows show the direction of the I-V curve shift at the given cell stretch under SNAP perfusion. It is obvious that during SNAP perfusion the activity of MGCs is inhibited. c – Difference current activated by 10 μm of stretch (IS–C ), corresponding to panel A (reversal potential Erev = −10 mV, IMGC(−45 mV) = −0.46 nA). d – Difference current (IS-S,SNAP ): the difference of the IS minus same stretch with SNAP IS,SNAP , corresponding to Fig. 9B (reversal potential Erev = −3 mV, IMGC(−53 mV) = (+)0.22 nA). Modified from Kazanski et al., 2010a with permission and the author’s data

potentials negative to −20 mV mechanosensitivity is composed of both Gns and GK1 . Thus, stretch increases MG-currents and depolarizes the cell. Figure 5.9b shows the change of IS on the background of 10 μm stretch at the beginning of SNAP perfusion (2 min) and then after 3 min. Despite the continuing cell stretch the SNAP perfusion returned the I-V curve in 2 min to the state close to the original, and the curve again turned N-shaped and crossed the voltage axis at −68 mV, in 3 min V0 = −78 mV (−87 ± 4 mV, n = 8). The stretch-induced difference current

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(IS−S,SNAP ) with SNAP 3 min after perfusion of SNAP is plotted in the I-V relations in Fig. 5.9d and equals (+)0.53 nA (IMGC(−45 mV) = (+)0.48 ± 0.05 nA, n = 8 vs control, P < 0.001; Erev = −3 ± 1 mV). Thus, SNAP perfusion under 10 μm cell stretch resulted in inhibited MG-currents, probably due to the blocking of MGCs. The I-V curve received in these conditions practically coincided after 2 min with the I-V curve registered in the control. But the following SNAP perfusion already at 3 min shifts the membrane potential value to −78 mV, which happens, in our opinion, as a result of inhibiting MGCs that have background activity. Similar data were obtained on ventricular cardiomyocytes from guinea-pig and rat. 5.3.2.2 NO-Donor DEA-NO The NO-donor DEA-NO or 2-(N,N-Diethylamino)-diazenolate-2-oxide.diethylammonium salt is a nitric oxide donor, useful for reliable generation of nitric oxide (NO) in vitro or in vivo. We perfused isolated ventricular cardiomyocytes with DEA-NO (250 μmol/L) solution without deforming the cell. After registering the control I-V curve we started perfusing the cell with DEA-NO. Figure 5.10a shows the control I-V curve (IC ) registered in a ventricular myocyte from rat and the I-V curve received 2 min after the beginning of DEA-NO perfusion (IDEA-NO ) without stretching the cell. Before perfusion with DEA-NO control I-V curve was N-shaped and crossed the voltage axis at −80 mV (−86 ± 5 mV, n = 9). It is obvious that DEA-NO induced MG-like currents, and already after 2 min the I-V curve crossed the voltage axis at −48 mV (−45 ± 3 mV, n = 9). The originating MG-like current corresponds to the stretch-induced current originating to 10 μm cell stretch. The difference current activated by DEA-NO (IDEA-NO−C ) is plotted in the I-V relations in Fig. 5.10c and equals −0.46 nA (IMGC(−45 mV) = −0.43 ± 0.03 nA, Erev = −3 ± 2 mV, n = 9) for 2 min DEA-NO perfusion. It is shown that after 2 min perfusion with DEA-NO the wash out for 2 min results in I-V curve shifts towards the control, i.e. the unstretched cell showing the loss of DEA-NO effect during registration time (Fig. 5.10b). After 2 min of wash out the control I-V curve (Fig. 5.10a) and the I-V curve registered after DEA-NO wash out (Fig. 5.10b) coincided completely, at which the latter crossed the voltage axis at −79 mV (−86±4 mV, n = 9). The difference of the currents activated by DEA-NO without stretch minus currents under wash out (IDEA-NO−W ) is plotted in the I-V relations in Fig. 5.10d. IDEA– NO–W = (+)0.47 nA (IMGC(−45 mV) = (+)0.45 ± 0.03 nA, Erev = −3 ± 2 mV, n = 9) Our data, accumulated on cells from different animals have demonstrated a pronounced DEA-NO effect as an activator of MG-like currents. Since application of the NO-donor DEA-NO in the absence of mechanical stimulation failed to induce MG-like currents we have concluded that, although availability of NO was crucial for MSC, an increase in NO alone was insufficient. Further on, we conducted a series of experiments in which we first stretched cells by 10 μm and then perfused them with DEA-NO (250 μmol/L). Figure 5.11 shows the results of the experiments conducted on ventricular myocyte from mouse. The voltage dependency of IC and its modulation by stretch (IS ) is shown in the I-V curves in Fig. 5.11a. Before stretch (IC ), the I-V curve was N-shaped and

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Fig. 5.10 Voltage dependence of membrane currents in ventricular cardiomyocytes from rat under DEA-NO perfusion of unstretched cell. a – I-V curve measured in control (IC ) and after 2 min DEA-NO perfusion (IDEA-NO ). b – after 1 and 2 min of DEA-NO wash out (W IDEA-NO ) as compared to IDEA-NO . Note: In panels (a) and (b) the arrows show the direction of the I-V curve shift under DEA-NO perfusion and under cell wash out. c – Difference current activated by DEA-NO (IDEA-NO–C : the difference of IDEA-NO minus IC ), corresponding to Panel a 2 min after the beginning of perfusion (reversal potential Erev = −5 mV, IMGC(−45 mV) = −0.46 nA). At 2 min IDEA-NO–C corresponds to 10 μm cell stretch. d – IDEO-NO–C activated by DEA-NO without stretch, 1 min after the beginning of wash out (reversal potential Erev = −4 mV, IMGC(−45 mV) = (+)0.22 nA. Modified from Kazanski et al., 2010a with permission and the author’s data

crossed the voltage axis at V0 = −69 mV (−83 ± 4 mV, n = 9). The 10-μm stretch shifted the net currents to more negative values (IS ), and V0 changed to −35 mV (−38±3 mV, n = 9). Close to −20 mV, the I-V curves recorded before and during stretch crossed each other. The difference of currents during stretch minus before stretch (IS−C ) is plotted in the I-V relations in Fig. 5.11c and equals −0.31 nA (IMGC(−45 mV) = −0.38 ± 0.04 nA, Erev = −15 ± 3 mV, n = 9). Thus, stretch increases MG-currents and depolarizes the cell. Figure 5.11b shows the change of IS on the background of 10 μm stretch under DEA-NO perfusion of the cell (3 min).

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Fig. 5.11 Voltage dependence of membrane currents in ventricular myocyte from mouse under DEA-NO perfusion of the cell on the background of its stretch. a – I-V curve measured before stretch (IC ) and during 10 μm stretch (IS ). The arrows show the direction of the I-V curve shift at the given cell stretch. b – modified I-V curves on the background of DEA-NO perfusion under continuous stretch during 3 min accordingly (IS,DEA-NO ). The arrows show the direction toward which the I-V curve shifts in response to DEA-NO application to prestretched cell. It is obvious that in the process of DEA-NO perfusion the activity of MGCs is inhibited. c – Difference current (IS–C ) activated by 10 μm of stretch corresponding to Fig. 5.11a (reversal potential Erev = −20 mV, IMGC(−45 mV) = −0.31 nA). d – Difference current (IS–S,DEA-NO ): the difference of IS minus same stretch with DEA-NO IS,DEA-NO , corresponding to Fig. 5.11b (reversal potential Erev = −20 mV, IMGC(−45 mV) = (+)0.34 nA). Modified from Kazanski et al., 2010a with permission and the author’s data

Despite the continuing cell stretch, the DEA-NO perfusion returns the I-V curve to the state close to the original, at which the curve turns again N-shaped and crosses the voltage axis at −75 mV (−87±3 mV, n = 9). The difference of IS minus during stretch with DEA-NO (IS,DEA-NO ) 3 min after perfusion is plotted in the I-V relations in Fig. 5.11d and equals (+)0.34 nA (IMGC(−45 mV) = (+)0.40 ± 0.03 nA, n = 9 vs.

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control, P = NS; Erev = −14 ± 4 mV). Thus, DEA-NO perfusion on the background of 10 μm cell stretch resulted in inhibition of MG-currents, probably due to blocking of MGCs. The I-V curve registered under these conditions shifted the value of membrane potential after 3 min to −78 mV, which occurs, in our opinion, as a result of inhibiting MSC that have background activity. Similar data were received on ventricular cardiomyocytes from guinea-pig and rat.

5.3.3 Nitric Oxide Synthases Inhibitors – NOS Inhibits We used a non-selective inhibitor of nitric oxide synthase - L-NAME (hydrochloride) or L-NG -Nitroarginine methyl ester (hydrochloride). Ventricular myocytes were preincubated in L-NAME solution (20 μmol/L) for 2 h after which they were put into the experiment. Figure 5.12 shows the results of the experiments conducted on ventricular myocyte from guinea-pig. The 10 μm stretch of the preincubated cell does not cause any changes in the I-V curve, therefore MGCs are completely blocked. An increase of L-NAME concentration to 100 μmol/L with the same incubation time does not cause any changes, there is no reaction of the cell to stretch. But a concentration decrease to 2 μmol/L with the same incubation time preserves completely the cell’s reaction to stretch and relaxation returns the I-V curve to the original state. Comparing Fig. 5.12 to Fig. 5.3 or Fig. 5.4 shows on guinea pig cell samples that the cell preincubation in L-NAME solution shifted the I-V curve downwardly and reduced the hump of outward current. The use of nitric oxide synthase inhibitor – L-NMMA or Nω -Methyl-L-arginine acetate (200 μmol/L, 2 h preincubation) showed results similar to the ones reported above. In L-NMMA-pretreated cells MG currents were absent (Dyachenko et al., 2009b). Thus, NO-synthase inhibitor L-NMMA completely blocked MGCs.

Fig. 5.12 Voltage dependence of stretch-induced membrane currents, K+ currents not suppressed. Ventricular myocyte from guinea pig. Current-voltage relation (I-V curve) measured before stretch (IC ) and during 10 μm stretch (IS ) after preincubation of cells in L-NAME solution (20 μmol/L, 2 h). Please, note that the cell preincubation in L-NAME solution changes the shape of the control I-V curve. See Fig. 5.3 compared to Fig. 5.4. Modified from Kazanski et al., 2010b with permission

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5.3.4 Cardiomyocytes Derived from NOS–/–Mice Research of physiological effects of NO is complicated by the fact that it is a fast inactivating molecule. Interpretation of the data is also complicated because in a cell there are other compounds very similar to NO in their chemical qualities, e.g., free radicals that have analogical biological effects practically impossible to divide pharmacologically. The solution is in using transgenetic animals, e.g. NOS–/– mice. MG currents were analyzed in cardiomyocytes derived from NOS–/– mice (Dyachenko et al., 2009b). In cardiomyocytes from NOS1–/– mice, 10-μm stretch activated Gns similar as in cells from their wild-type littermates (10μm Gns = 7.3 ± 2.1 nS, n = 5 in NOS–/– vs. 10μm Gns = 6.2 ± 2.5 nS, n = 8 in wild-type littermates). In contrast, in cardiomyocytes from NOS3–/– mice, 10-μm stretch did not significantly induce MG currents (0μm Gns = 0.2±0.6 nS vs. 10μm Gns = 2.0±3.4 nS and 0μm GK1 = 24 ± 10 nS vs. 10μm GK1 = 23 ± 11 nS, n = 18; Fig. 5.12). This result clearly points to NOS3 as the dominant source of NO involved in MGCs and is in agreement with studies showing that NOS3 is activated by stretch (Petroff et al., 2001; Kuebler et al., 2003; Fig. 5.13)

5.3.5 Possible Explanations of NO Involvement into Regulation of MG-Currents The data presented in Part 4 about the involvement of NO into regulating the activity of MG-currents come down to several basic issues. In ventricular myocytes from guinea pig, mouse and rat in all experimental conditions the NO scavenger PTIO produces complete inhibition of stretch-activated MG currents and deactivation of K1-currents. The extent of MG current block depends on incubation duration. The

Fig. 5.13 NOS3-derived NO in MGCs. Cardiomyocyte isolated from a NOS3–/– mouse. 10-μm stretch of the myocyte does not induce MSC (black: before stretch; red: during 10-μm stretch) (From Dyachenko et al. (2009b) with permission of Elsevier)

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wash-out of PTIO does not reverse the blockade of MG current. Wash-out stops the development of the blockade. We believe that application of PTIO scavengers NO, thus blocking MG currents. However during the beginning of application of PTIO, its removal prevents the development of the MG current blockade. PTIO blockade of MG current cannot be reversed by its washout. The NO-donors SNAP and DEO-NO in undeformed cell result in activation of MG-like currents. Wash out removes these currents. On the contrary, SNAP and DEO-NO inhibit MG currents in a previously stretched cell. This is an interesting phenomenon that correlates with described bimodal effects of NO. It is known that the effect of NO is bimodal, with a positive inotropic effect at low amounts of NO exposure but a negative one at higher amounts (Massion et al., 2003). Admittedly, defining what low or high amounts really mean is difficult, both in terms of actual quantity of bioactive NO delivered (e.g., with different exogenous NO donors) as well as the correspondence with amounts endogenously produced in vivo. Similarly, Kojda et al. (1996) observed that low concentrations of the NO donors SNAP and DEA/NO caused a moderate positively inotropic effect in adult rat ventricular myocytes. Higher concentrations of either SNAP or DEA/NO suppressed myocyte contractile function. Mohan et al. (1996) have reported a similar biphasic inotropic response to NO donors in isolated feline papillary muscle strips that appeared to be dependent on a GMP-mediated signaling pathway. Since the direction of NO donors mediated effects can be dose dependent we suggest that NO donors open MGCs in ventricular cardiomyocytes, which underlie MG-like currents. We also register MG currents when stretching the cell, but with that we increase the concentration of endogeneous NO. We speculate that in both first and second cases we have that low concentration of exogenous or endogenous NO, that activates MGCs. In case the cell, when stretched, shows higher endogenous NO concentration, then an additional increase of exogenous concentration using NO donors results in increased summary concentration, which probably is the reason for inhibiting MGCs. As we have no data on MGCs ion channel structure in ventricular myocytes, our speculation is based on analogues with voltage gated L-type calcium channels which also show dose dependent activatory and inhibitory effects of NO donors on ICa−L . Thus, e.g., Mery et al. (1993) originally reported that SIN-1 (NO donor – 3-morpholino-sydnonimine), the active metabolite of molsidomine, has a biphasic effect on ICa−L in enzymatically dissociated frog ventricular myocytes. The data of these authors suggest that the activatory and inhibitory effects of NO donors on ICa−L result from an inhibition of the cGMP-inhibited cAMP-phosphodiesterase and an activation of the cGMP-stimulated cAMP-phosphodiesterase, respectively, both linked to the activation of guanylyl cyclase, possibly a membrane form of the enzyme. The use of NOS inhibitors L-NAME and LNMMA results in MG currents that do not appear as a response to cell stretch. In NOS1–/– the reaction to stretch is present, whereas in NOS3–/– there is no reaction to stretch. This allows to consider NOS3 as the dominant source of NO involved in MGCs and also define that NOS3 is activated by stretch.

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5.4 Cell Signaling of Nitric Oxide in the Heart and Possible Role in Regulation of MG-Currents NO has a wide range of physiological effects, but the main intracellular target for NO is considered soluble guanylyl cyclase that causes an increase of the concentration of cGMP (Fig. 5.14), which can modulate the activity of cGMP-dependent cyclic nucleotide phosphodiesterases and, respectively cAMP level, as well as cGMP-dependent protein kinase G, that causes phosphorylation of a number of proteins (Moncada et al., 1991; Murad, 1998; Lane and Gross, 1999; Shah and MacCarthy, 2000; Bryan, 2009). The cGMP-mediated NO effect also includes influence on such effectors as cyclooxygenases, phosphatases, phospholipase S (Murad, 1998; Lane and Gross, 1999; Bryan, 2009). The effect of the cGMP-dependent pathway on the function of the myocardium consists of modulating Sa2+ influx from the sarcolemma, lowering myofilaments sensitivity to Sa2+ , changing the functional activity of sarcoplasmic reticulum, changing the action potential, changing cell volume, and decreasing oxygen consumption (Ji et al., 1999; Shah and MacCarthy, 2000). cGMP-dependent NO effects in the heart are: (1) early onset of relaxation mediated by troponin I phosphorylation by protein kinase G, resulting in lower myofilament sensitivity to Sa2+ ; (2) positive inotropic effect of NO at low concentrations by affecting cGMP-inhibited cyclic nucleotide phosphodiesterase resulting in increased levels of intracellular cAMP, but cGMP participation can not be excluded, (3) negative inotropic effect of higher NO doses, realized probably by activating protein kinase G, (4) modulation of β-adrenergic and cholinergic responses, and (5) protective effect on cardiomyocites under myocardial hypoxia at the expense cGMP-mediated opening

Fig. 5.14 NO-cGMP-mediated signal transduction in myocardium (Reproduced from Shah and MacCarthy, 2000 with permission from Elsevier.)

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of KATP -channels (Ji et al., 1999; Casadei and Sears, 2003; Shah and MacCarthy, 2000). The cGMP-independent mechanisms for realizing NO effects include its reactions with amino, thiol (SH), diazo and tyrosyl groups in proteins, as well as haem-, iron- or sulphur centers of proteins (Mateo and De Artiñano, 2000; Landar and Darley-Usmar, 2003; Hughes, 2008; Bryan et al., 2009). The most important fact is the direct interaction of NO with chemically active thiol protein groups causing their posttranslational modifications, which results in significant functional changes. S-nitrosylation of reactive thiol groups can affect the activity of ion channels, transporters and Sa2+ -binding proteins participating in the regulation of Sa2+ cycling in myocytes. Possible targets are ryanodine receptor Ca2+ release channels (RYR) of sarcoplasmic reticulum, SERCA 2a, phospholambam, Sa2+ -ATP-ase of the sarcolemma (Shah and MacCarthy, 2000; Casadei and Sears, 2003; Lim et al., 2008; Petroff et al., 2001; Jaffrey et al., 2001). A NO mediated increase of Sa2+ concentration in cardial myocytes is the main mechanism of NO modulation of myocardial excitation–contraction coupling and realization of cardiac cell response to mechanical impacts (Shah and MacCarthy, 2000; Seddon et al., 2007; Lim et al., 2008; Fig. 5.15). One more important mechanism for realizing NO effects is its interaction with active peroxide radicals. NO, especially under pathophysiological conditions, can interact with high reactivity superoxide, which in norm is tied by superoxide dismutase. As a result a high-activity compound is formed, such as peroxynitrite

Fig. 5.15 Regulation of cardiomyocyte functions by nitric oxide in normal heart (Reproduced from Seddon et al., 2007 with permission from Elsevier)

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(ONOO− ) that can modulate ion channel functions, oxidizing, like NO itself, the regulatory thiol groups of channel proteins (Mateo and De Artiñan, 2000; Landar and Darley-Usmar, 2003; Hughes, 2008). Also shown was the effect of peroxynitrite on the physiological activity of a number of enzymes, such as phospholipases of different types (Yuen et al., 2000; Guidarelli et al., 2000). Protonation of peroxynitrite, especially under low rN, observed in heart pathologies, results in formation of peroxynitrous acid. As a result, highly reactive hydroxyl-like species are formed that can cause toxic effects related to nitration and oxidation of not only functional but also structural proteins (reviewed in Shah and MacCarthy, 2000).

5.5 Conclusion and Perspectives There is a wide discussion going on about, how mechanical energy is transferred to MGC: through the lipid bilayer of the membrane or through the cytoskeleton and which of these mechanisms prevail. Several groups reported that MGCs are activated by the stretch of the lipid bilayer, while other papers are focused in the role of the cytoskeleton in MGCs activation. Some authors brought up the role of extracellular matrix, although the forces distribution in the extracellular matrix remain unknown. In addition to those issues we would like to add another one – the possibility of MGCs modulation by intracellular second messengers and/or pharmacological compounds. We have already mentioned that in our experiments stretch or several pharmacological compounds, besides changing current through cation-nonselective MGCs, also change mechanosensitive currents carried by K+ ions through both TRPC6 and TREK which are mechanosensitive, together with inwardly rectifying K+ -current IK1 (possibly through Kir2.3). Especially hard to explain is the appearance of Ins in absence of cellular stretch. Regarding nitric oxide, it is possible that S-nitrosylation of reactive thiol groups can affect the activity of MGCs. In general, different studies report that besides direct activation of MGCs by mechanical forces (via cytoskeleton or bilayer) fast indirect modulation of MGCs by pharmacological compounds is also possible. Moreover some of those compounds are capable of activating MGCs in the absence of actual cellular stretch, while others inactivate MGCs despite continuous presence of cellular stretch. If this is true for compounds other then nitric oxide, this provides an excellent opportunity for development of new drugs for treatment of mechano-induced arrhythmias as well. Acknowledgments This work was supported by grants from RFBR (09-04-01277a), DFG (Tr 02-A3) and a travel grant from the Humboldt-University (Berlin, Germany). VK, AK and IK thank Prof. G. Isenberg and Prof. P. Persson for providing the opportunity to perform some of experiments and general support of this work.

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Chapter 6

Role of Signaling Pathways in the Myocardial Response to Biomechanical Stress and in Mechanotransduction in the Heart Danny Guo, Zamaneh Kassiri, and Gavin Y. Oudit

Abstract The heart is a mechanosensitive organ that adapts its morphology to changing hemodynamic conditions via a process named mechanotransduction, which is the primary means of detecting mechanical stress in the extracellular environment. In the heart, mechanical signals are propagated into the intracellular space primarily via integrin-linked complexes, and are subsequently transmitted from cell to cell via paracrine signaling. The biochemical signals derived from mechanical stimuli activate both acute phosphorylation of signaling cascades, such as in the PI3K, FAK, and ILK pathways, and long-term morphological modifications via intracellular cytoskeletal reorganization and extracellular matrix remodelling. Mechanotransduction plays a fundamental role in cardiac (and vascular) function and involves interaction between extracellular matrix and intracellular cytoskeletal proteins via cell adhesion complexes, which are modulated by PI3Ks. Loss of PI3K signaling enhances the susceptibility to biomechanical stress while the loss of its negative regulator, PTEN, is associated with a wide variety of adaptive mechanisms necessary to resist the progression of maladaptive ventricular remodelling and heart failure. In this chapter, we discuss several of the key players involved in mechanotransduction in the heart. Keywords Mechanotransduction · ECM · Integrin · ILK · FAK · Dystrophin · PI3K · PTEN · Cadherin · Remodelling

G.Y. Oudit (B) Division of Cardiology, Department of Medicine, Mazankowski Alberta Heart Institute, University of Alberta, Edmonton, AB T6G 2S2, Canada e-mail: [email protected]


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6.1 Introduction Biomechanical signaling involves complex interactions between intracellular and extracellular components. While whole organisms respond to “external” stimuli, organs, tissues, and cells respond to mechanical signals from the extracellular space. Common cellular responses to such signals include alterations in morphology, intracellular structure, and extracellular structure. This extracellular transmission of mechanical force followed by the subsequent intracellular conversion into biochemical signals is termed mechanotransduction (Fig. 6.1). This process describes the dynamic process of a mutually dependent relationship between cells and their surrounding extracellular matrix (ECM). Responding to mechanical stress allows cells to acclimate and adapt to changing environments. However, the benefits from this intricate process are reversed if the carefully balanced equilibrium is tilted, often leading to abnormal signaling and pathophysiological remodelling of cells and tissues. By detecting and responding to hemodynamic changes in the environment via cell-ECM interactions, the heart develops compensatory responses utilizing intracellular signaling cascades to maintain adequate function (Frey et al., 2004; Heineke and Molkentin, 2006; Ruwhof and van der Laarse, 2000). Even at rest, the heart is constantly exposed to much biomechanical and biochemical signals, causing it to continuously adapt various aspects of its morphology such as the infrastructure of its cytoskeleton as well as the composition of cell-ECM adhesion complexes. This chapter will discuss several sub-cellular processes and signaling pathways involved in mechanotransduction and the interactions between cardiomyocytes and fibroblasts with the ECM.

Fig. 6.1 A conceptual framework of the key players involved in the detection of and the translation of mechanical signals in the heart

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6.2 Cell-ECM Adhesion The ECM is like a framework of springs maintaining cells in a state of physical equilibrium, which, if deformed, mechanically stimulates the cells (Sussman et al., 2002). The structural component of the ECM is a complex lattice of macromolecular proteins such as fibronectin and collagen; these proteins facilitate even distribution of exogenous mechanical signals to the recipient cells (Berrier and Yamada, 2007). Cellular attachment to the ECM via adhesion complexes such as integrin and dystrophin-glycoprotein complexes warrants effective transmission of extracellular mechanical signals into intracellular domains, stimulating changes in cellular processes such as the cell cycle (Berrier and Yamada, 2007; Sussman et al., 2002). Depending on the type and the mode of stimulation, downstream signals may vary from mediating acute responses, such as the phosphorylation and activation of signaling cascades and the generation of second messengers, to long-term cellular modifications, such as changes in gene expression, initiation of compensatory pathways, and/or modification of intracellular and extracellular structural compositions. For instance, cyclic and static stretch, elongation and compression, and unilateral and bilateral strain all generate unique changes in transcriptional profile and total gene expression in cardiac fibroblasts (Lee et al., 1999; Simpson et al., 1999). Similarly, in vitro studies of cardiomyocytes cultured on deformable membranes suggest that different magnitudes and directional orientations of stretch and compression yield distinct intracellular effects (Kumar et al., 2002; Ruwhof et al., 2000). Mechanical stimulation of one cell can spread to its neighbouring cells via paracrine release of chemical messengers such as angiotensin 2 (Ang II), endothelin-1 (ET-1), and transforming growth factor β (TGF- β) (van Wamel et al., 2001) (discussed in Section 6.3.2). In order for cardiac cells to maintain the intricate cell-ECM interactions necessary for mechanotransduction, sensitive adhesion complexes are required. Though more than 50 proteins have been reported to be associated with these complexes (Zamir and Geiger, 2001), cell-ECM adhesions in cardiomyocytes can be categorized into two major groups: integrin-linked and dystrophin-glycoprotein complexes (Fig. 6.2).

6.2.1 Cell-ECM Adhesion: Integrin-Linked Complexes An increased pressure and/or volume hemodynamic load on the heart typically results in the development of pathological ventricular hypertrophy and systolic and diastolic dysfunction. One of the key players in mediating this response is the integrin-linked complex, which contains a heterodimeric pair of integrins (α and β) that spans across the cytoplasmic membrane, binding to structural proteins in the ECM, such as collagen, laminin, and fibronectin (Fig. 6.2) (Berrier and Yamada, 2007; Carver et al., 1994; Hynes, 1992; Schwartz et al., 1995). Integrin-linked complexes also bind to cytoskeletal actin and sarcomeric components primarily

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Fig. 6.2 A schematic of integrin based complexes and their associated signaling pathways. Integrin complexes linked to the Z-disc are called costameres. The ones directly linked to actin are called focal adhesions

through the β-subunit’s intracellular domain (Calderwood et al., 2000; Schwartz et al., 1995). This bridging of intracellular and extracellular structural polymers allows integrins to propagate bidirectional biomechanical communication between cytoskeletal components and the ECM (Hynes, 2002). Mice with cardiomyocyte specific ablation of β1-integrin have reduced cardiac function, intolerance to increased hemodynamic workload, and gradual development of heart failure with age (Shai et al., 2002) while mice with overexpression of β1-integrin develop an augmented hypertrophic response to mechanical stress (Ross et al., 1998). In concordance, studies involving transgenic and adenoviral models of β1-integrin both show that integrin’s downstream signaling cascades directly promote growth, proliferation, and survival (Ieda et al., 2009; Ross et al., 1998) implicating that β1-integrin is critical to and is directly responsible for inducing cardiac hypertrophy. Integrin-linked complexes are divided into costameres and focal adhesions in cardiomyocytes (Fig. 6.2), both of which form in response to stretch (Sharp et al., 1997). Structurally, these two categories only differ in the proteins clustered at their intracellular domains and their intracellular anchoring points. While costameres connect to sarcomeres via Z-disc proteins, focal adhesion complexes associate

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with cytoskeletal actin filaments (Ervasti, 2003; Laser et al., 2000; Samarel, 2005; Sharp et al., 1997). To some degree, both types of integrin-linked complexes share overlapping signaling cascades with maintaining some key differences in signaling. Integrins moderate cellular behaviour partially through physically bridging the ECM to intracellular components. Nevertheless, the formation of mutli-protein complexes at their intracellular domains is necessary for mechanotransduction (Berrier and Yamada, 2007; Hynes, 2002). Because integrins do not exhibit inherent enzymatic activity, they must recruit signaling proteins to their cytoplasmic tails to convert mechanical stimuli into biochemical signals (Berrier and Yamada, 2007). In general, the proteins recruited to integrin-linked complexes are categorized into 3 major groups (Berrier and Yamada, 2007). The first group is integrin regulating proteins such as talin, which bind to integrins and regulate their signaling and activity (Anastasi et al., 2009; Calderwood, 2004; Chen et al., 1995; Ulmer et al., 2003). The second group is adaptor proteins, which act as scaffolds and link integrins to structural and enzymatic proteins (Berrier and Yamada, 2007). An example is paxillin, a β-integrin binding protein that recruits enzymes to integrin-linked complexes (Schlaepfer et al., 1999; Tachibana et al., 1995). Another example is vinculin, which in response to mechanical stimulation relocates to costameres and connect integrinlinked complexes to cytoskeletal actin (Sharp et al., 1997; Wood et al., 1994). The third group is enzymatic proteins such as focal adhesion kinase (FAK), integrinlinked kinase (ILK) and phosphoinositide-3-kinase (PI3K) and which can initiate a broad spectrum of chemical signaling cascades (Berrier and Yamada, 2007; Ross et al., 1998). 6.2.1.1 Cell-ECM Adhesion: Focal Adhesion Kinase Focal adhesion kinase (FAK), a non-receptor tyrosine kinase (Peng et al., 2008), can be associated with focal adhesion complexes but can also localize at costameres depending on the stimulus (Fig. 6.2) (Sharp et al., 1997; Torsoni et al., 2003). In response to stretch and compression, FAK is recruited to integrin-linked complexes via paxillin (Domingos et al., 2002; Tachibana et al., 1995), strategically allowing FAK to receive mechanical signals directly; moreover, multiple FAKs can cluster at these complexes in a number proportional to the strength of stimulation, sustaining and enhancing downstream signals accordingly (Katz et al., 2002; Torsoni et al., 2003). However, it is important to note that FAK translocates differentially in response to mechanical and chemical stimuli. In stretched cardiomyocytes in vitro, FAK associates with Src adaptor proteins at costameres and autophosphorylates at Tyr-397 (Laser et al., 2000; Torsoni et al., 2003). Src subsequently phosphorylates FAK at Tyr-576 and Tyr-577, activating its kinase activity. The disruption of the FAK-Src complex via Src inhibitors prevents FAK from binding to costameres leading to the loss of FAK mediated signaling (Domingos et al., 2002; Torsoni et al., 2003). One of the many roles of the FAK-Src complex in cardiomyocytes is to mediate stretched induced up-regulation of gap junctions (Yamada et al., 2005). Stretching cardiomyocytes causes the release of VEGF in a FAK dependent manner, increasingly ERK phosphorylation, an effect which is prevented via neutralization

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of VEGF suggesting that VEGF increases ERK phosphorylation via paracrine signaling (Seko et al., 1999). The FAK mediated release of VEGF also up-regulates the gap junction protein connexin-43 in neighbouring cells via paracrine signaling (Li et al., 1997; Yamada et al., 2005). Increased expression of connexin-43 improves overall electrical signaling in the heart (Yamada et al., 2005). FAK-Src can also up-regulate the expression of cell-cell adhesion proteins, such as N-cadherin, desmoplakin, and plakoglobin (Yamada et al., 2005). Because stretch induced localization of FAK to integrin-linked complexes increases with the magnitude and the duration of stretch (Katz et al., 2002; Simpson et al., 1999), up-regulation of the aforementioned cell-cell adhesion proteins in cardiomyocytes is likely proportional to increases in work load. Cardiomyocyte adhesion to fibronectin stimulates a higher level of N-cadherin and connexin-43 expression than adhesion to collagen suggesting that matrix composition may alter focal adhesion binding and signaling (Shanker et al., 2005). Together, these demonstrate that mechanical stimulation of FAK is involved in the maintenance of both electrical and mechanical connectivity in the heart. Contrary to mechanical stimulation, chemical stimulation in cardiomyocytes recruits FAKs to focal adhesions instead (Fig. 6.2) (Torsoni et al., 2003). For instance, application of Ang II stimulates FAK mediated hypertrophy (Salazar and Rozengurt, 2001). In concordance, application of AT1 receptor antagonists impairs Ang II stimulation of FAK signaling without inhibiting cyclic stretch stimulation of FAK signaling (Torsoni et al., 2003). This bimodal activation of FAK is one of many examples that demonstrate the specificity of mechanotransduction and its associated signaling cascades. While costameric and focal adhesion FAKs are, on the most part, distinct in their respective signaling cascades, it should be noted that there are circumstances of overlap. For instance, studies have shown that FAK activates phosphoinositide-3-kinase (PI3K) and its subsequent effectors by binding to PI3K’s p85 adaptor subunit regardless of FAK’s location (Chen and Guan, 1994; Thamilselvan et al., 2007). Similarly, when activated, both costameric and focal adhesion FAKs can bind to the mitogen activated protein kinases, ERK, JNK, and p38, via Grb2 and Sos (Schlaepfer et al., 1999; Torsoni et al., 2003). Focal-adhesion kinase importance to cardiac development is illustrated by the heart-specific FAK-deficient embryonic mice (Table 6.1). These mutant mice develop right ventricular eccentric hypertrophy due to ineffective FAK mediated stimulation of physiological hypertrophic programs, ultimately resulting in decompensation and maladaptive right ventricular remodelling (Peng et al., 2008). On the other hand, after birth, these mice gradually develop left ventricular eccentric hypertrophy overtime due to age (Peng et al., 2008). Similar pathologies are observed acutely when these mice are treated with Ang II or pressure overload (Peng et al., 2008). This shift from right to left ventricular eccentric hypertrophy is likely due to differences in ventricular dependence and workload between embryonic and postnatal mice (Peng et al., 2008). This model implies that FAK signaling is necessary for myocardial tolerance of mechanical stress, possibly through eliciting hypertrophy via the regulation of various transcription factors such as NF-κB (Crosara-Alberto et al., 2009; Gupta et al., 2002).

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Table 6.1 Mutations in various genes involved in mechanotransduction and their effects on signalling Gene mutation

Phenotype

Reference

β1-integrin

Whole body mutant: Embryonic lethality. Cardiac specific mutant: Reduced cardiac function. Intolerance to stress. Overexpression leads to augmented hypertrophic response. Whole body mutant: Embryonic lethality. Cardiac specific mutant: Right ventricular eccentric hypertrophy during embryonic stage and left ventricular eccentric hypertrophy postnatal. Whole body mutant: Embryonic lethality. Cardiac specific mutant: Attenuation of hypertrophic response. Weakened FAK/PI3K signaling. Whole Body mutant: Embryonic lethality. Cardiac Specific mutant: Reduce AKT signaling. Whole body knock out: Increased ERK1/2 stimulation. Decreased AKT activation.

Ross et al. (1998) Shai et al. (2002)

FAK

ILK

N-Cadherin

Dystrophin

Peng et al. (2008)

White et al. (2006) Lu et al. (2006)

Tran et al. (2002)

Khairallah et al. (2007) Kumar et al. (2004)

6.2.1.2 Cell-ECM Adhesion: Integrin Linked Kinase Integrin linked kinase (ILK) plays a key role in mediating hypertrophic programs at both costameres and focal adhesion complexes (Fig. 6.2) (Li et al., 1999; Sakai et al., 2003; White et al., 2006). When integrins in cardiomyocytes bind to the ECM, ILKs are recruited to costameres via paxillin and are subsequently activated; this allows ILKs to interact with the cytoplasmic domain of β1-integrin and phosphorylate several downstream targets such as Rac1, AKT, and GSK-3β (Hannigan et al., 2005; Li et al., 1999; Lu et al., 2006; Qian et al., 2005; Ross et al., 1998), which have all been associated with mechanically stimulated cardiomyocyte hypertrophy (DeBosch et al., 2006; Matsuda et al., 2008; Satoh et al., 2006; Sugden, 2003). In addition to its enzymatic roles, ILKs also act as scaffolding proteins by forming multi-protein complexes at focal adhesions (Hannigan et al., 2005, 2007). In cardiomyocytes, ILK is recruited to focal adhesions via binding to PINCH1 (BockMarquette et al., 2004; Chen et al., 2005; Li et al., 1999), which if inhibited, leads to reduced phosphorylation of AKT (Delcommenne et al., 1998; White et al., 2006) and GSK-3β (Delcommenne et al., 1998) implying that the formation of the ILKPINCH1 complex may be necessary for ILK signaling. On the other hand, one study suggested that PINCH1 is only necessary during embryonic development and is dispensable in postnatal cardiomyocytes (Liang et al., 2005). Integrin-linked kinase interaction with two regulatory proteins, α-parvin and β-parvin, can further illustrate its role as a scaffolding protein (Chen et al., 2005;

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Tu et al., 2001). The mutually exclusive binding of α-parvin and β-parvin respectively increases and reduces ILK’s prohypertrophic activity (Hannigan et al., 2005). Interestingly, formation of either α-parvin-ILK or β-parvin-ILK complexes have been suggested to increase cell-ECM adhesion complexes (Hannigan et al., 2005; Mongroo et al., 2004; Tu et al., 2001; Yamaji et al., 2001) demonstrating ILK’s importance as a scaffold in mediating the formation of integrin-linked complexes. This also puts forth the idea that the enzymatic and scaffolding functions of ILK may be independent of each other (Hannigan et al., 2005). Structural analysis of ILK reveals a highly conserved Pleckstrin homology domain, which activates ILK by binding to phosphoinositide-3,4,5-phosphate (PIP3 ) (Delcommenne et al., 1998), the primary product of PI3K activity (Vanhaesebroeck et al., 1997) positing that activation of PI3K can subsequently activate ILK. In addition, inactivation of PTEN, which is a lipid phosphatase that dephosphorylates PIP3 into PIP2 (Oudit and Penninger, 2009; Oudit et al., 2004; Sun et al., 1999a), results in constitutive activation of ILK (Edwards et al., 2005). Previous studies have also shown that both enhancing PI3K activity and impairing PTEN activity can enhance intracellular PIP3 levels, leading to cardiac hypertrophy (Crackower et al., 2002; Shioi et al., 2000; Sun et al., 1999a). Together, these suggest that PI3K activity can directly modulate ILK activity. Finally, ILK also directly phosphorylates AKT and GSK-3β (Troussard et al., 1999), two well studied downstream effectors of FAK/PI3K signaling, supporting the notion that ILK may contribute to the myocardial compensatory response to mechanical stress. Since PI3K is involved in mediating acute responses to both chemical stimuli via RTK and GPCR receptors (Oudit and Penninger, 2009; Oudit et al., 2004) and mechanical stimuli via FAK (Chen and Guan, 1994), it is possible that ILK may also be activated by chemical agonist stimulation. For instance, the application of Ang II induces concentric hypertrophy in wildtype mice (Sadoshima and Izumo, 1993; Sadoshima et al., 1993), an effect that is mitigated in ILK kinase dead mutants (Lu et al., 2006). This partial attenuation of hypertrophy in the mutant strain suggests that ILK amplifies, but is not necessary for, PI3K signaling. On the other hand, one study suggested that ILK activation may be entirely PI3K dependent, as pharmacological inhibitors of PI3K fully inhibited mechanically induced ILK activity (Khwaja et al., 1997). However, an in vivo study stated that during maladaptive cardiac hypertrophy, increased ILK association with Rac1 was observed without phosphorylation of the PI3K signaling cascade (Lu et al., 2006). The same study also demonstrated that ILK kinase dead mutants are unable to develop ventricular hypertrophy in response to mechanical stress (Lu et al., 2006). Similarly, another study postulated that ILKs can directly respond to RTK stimulation irrespective of PI3K, further implicating that ILK and PI3K pathways may be independent (Knoll et al., 2007). Though the results from these studies appear to conflict, their discrepancies could be attributed to differences in the heart’s condition; while ILK activity may be PI3K independent during pathological states, ILK is likely PI3K dependent during physiological conditions. As one might expect, ILK is necessary for survival as mice with cardiomyocytespecific ablation of ILK develop heart failure and sudden death within 6 weeks

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of age and develop spontaneous dilated cardiomyopathy (DCM) without the need for a secondary stressor such as pressure overload or Ang II (Table 6.1) (White et al., 2006). Similarly, a single mutation in the human ILK gene (A262V) has been correlated with contractile dysfunction, ventricular dysmorphism, and DCM (Knoll et al., 2007). As mice with cardiomyocyte specific ablation of PINCH1 do not deviate from the wildtype phenotype (Liang et al., 2005), it is likely that ILK has other physiological targets of interaction in cardiomyocytes allowing it to sustain its signaling even in the absence of PINCH1. Interestingly, mice lacking ILK have impaired FAK signaling indicating that ILK either contributes to FAK activation or mediates a part of FAK’s downstream signaling (White et al., 2006). One possible mechanism is that FAK indirectly stimulates ILK activity via PI3K/PIP3 , which is not impossible due to their close spatial proximity at integrin-linked complexes (Chen and Guan, 1994; Thamilselvan et al., 2007). This is further supported by FAK and ILK’s overlapping downstream signaling (Antos et al., 2002; Bueno and Molkentin, 2002; Delcommenne et al., 1998; Shioi et al., 2002; Troussard et al., 1999).

6.2.2 Cell-ECM Adhesion: Dystrophin-Glycoprotein Complex The dystrophin-glycoprotein complex (DGC) is another major class of actin-linked adhesion complex (Hanft et al., 2006; Rybakova et al., 2000). DGCs are only briefly covered in this report but its role in disease and cardiac physiology and disease have previously been discussed elsewhere (Danialou et al., 2001; Ervasti, 2007; Lapidos et al., 2004; Rybakova et al., 2000). DGCs were traditionally regarded as being only involved in mechanical linkage, but recent studies illustrate that the loss of DGCs results in altered protein expression and cellular contractile dysfunction (Danialou et al., 2001; Hanft et al., 2006; Lapidos et al., 2004). Specifically, mutants lacking dystrophins develop various diseases such as Duchenne muscular dystrophy, Becker muscular dystrophy, and X-linked DCM, all of which are accompanied with the development of cardiomyopathy (Lapidos et al., 2004). One well studied DGC mutant model is the mdx mouse, which has a point mutation in the dystrophin gene (Quinlan et al., 2004). These mice show no signs of cardiomyopathy at 10–12 weeks of age, but inevitably develop DCM and myocardial contractile dysfunction at 40 weeks of age (Table 6.1) (Quinlan et al., 2004; Quinlivan et al., 1996). It should be noted that though these mice do not have cardiac dysfunction at 10–12 weeks, they are less tolerant of hemodynamic stress and are more susceptible to work load induced myocardial injury and adverse ventricular remodelling (Danialou et al., 2001). Though DGCs primarily modulate cell-ECM adhesion, its role in coordinating intracellular chemical signaling can also be illustrated in mdx mice, which have increased ERK1/2 activation both at rest and in response to mechanical stress as compared to wildtypes (Kumar et al., 2004). On the other hand, a metabolic study demonstrated that mdx hearts have a shift in basal metabolic substrate dependence from fatty acids to carbohydrates (Khairallah et al., 2007) suggesting

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that a weakened energy supply may be responsible for its intolerance to stress. However, since the overall cellular AMP to ATP ratios in the mdx cardiomyocytes are not different from wildtypes (Khairallah et al., 2007), it is possible that mdx cardiomyocytes either utilize a less efficient pattern of ATP allocation to sustain normal contractile function or require more ATP to sustain physiological conditions resulting in energy deficiency. It is interesting to note that mdx hearts have decreased activation of AKT in vivo (Khairallah et al., 2007) indicating that DGCs, like integrins, may associate with the PI3K signaling cascade. Consistent with the mdx phenotype, AKT has been shown to be necessary for physiological growth and cellular survival in the heart (DeBosch et al., 2006). Moreover, since DGCs are located closely to integrin-linked complexes within the cell (Rybakova et al., 2000), they could very likely interact with ILKs or FAKs, posing a potential cause for the decreased activation of AKT observed in mdx mice.

6.3 PI3K and PTEN 6.3.1 PI3K PI3K, as previously described, is a highly conserved lipid kinase and an important mediator of acute cellular responses to mechanical and chemical stimuli (Figs. 6.2 and 6.3) (Shioi et al., 2000; Oudit and Penninger, 2009; Oudit et al., 2004).The PI3K signaling cascade is both directly and indirectly involved in mechanically stimulated pathways. The direct mode of PI3K activation is at integrin complexes via interactions with enzymes such as FAK and ILK (refer to Sections 6.2.1.1 and 6.2.1.2.). On the other hand, PI3K is indirectly activated during mechanical stimulation via autocrine and paracrine signaling (refer to Section 6.4.2). The primary PI3K isoforms expressed in the heart are PI3Kα and PI3Kβ, which bind to receptor tyrosine kinases (RTKs) and are activated by growth factors, and PI3Kγ, which binds to GPCRs and are activated by, among others, Ang II stimulation (Oudit and Penninger, 2009). Though PI3Kβ has traditionally been associated with RTK signaling, later studies have given controversy to this belief revealing that PI3Kβ can be stimulated by binding to G-proteins (Kurosu et al., 1997; Maier et al., 1999; Yart et al., 2002). The generation of PI3Kα knockout mice was unsuccessful as PI3Kα is necessary for embryonic development (Shioi et al., 2000). Thus, studies made use of PI3KαDN instead, which overexpress a dominant but inactive form of PI3Kα in their cardiomyocytes after birth (Shioi et al., 2000). PI3Kα-DN hearts are smaller than wildtypes but do not display any signs of ventricular dysfunction under resting conditions. Furthermore, unlike wildtypes, PI3Kα-DN hearts do not hypertrophy when exercised, suggesting that PI3Kα is necessary for cellular growth and mechanosensitivity to physiological stress (McMullen et al., 2003). Moreover, PI3Kα may also protect the heart against pathological stress as PI3Kα-DN hearts rapidly develop

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Fig. 6.3 The PI3K signaling cascade. PI3K signals through phosphatidylinositol-3,4,5triphosphate (PIP3 )/Akt pathway and stimulates growth, proliferation, and survival. RTK = receptor tyrosine kinase, GPCR = G-protein coupled receptor

DCM in response to pressure overload (McMullen et al., 2007). PI3Kα’s cardioprotective properties are also observed in mutants that overexpress PI3Kα, which are protected from PKCβ induced pathological hypertrophy (Bowman et al., 1997; Rigor et al., 2009). PI3Kα’s effects on the heart are perhaps due to changes atrial natriuretic peptide (ANP) expression, which increases in PI3Kα-DN mutants exposed to pressure overload but not exercise (McMullen et al., 2003). On the other hand, in exercised but not pressure overloaded wildtype mice, brain natriuretic peptide (BNP) expression increases (McMullen et al., 2003), which can attenuate ERK activation and pathological hypertrophy (Takahashi et al., 2003). Together these observations denote that PI3Kα regulates physiological and pathological hypertrophy. Similarly to PI3Kα-DN, PI3Kγ-KO mutant hearts are also more susceptible to pathological stress as they rapidly develop DCM and ventricular lesions within 1 week after aortic banding (Patrucco et al., 2004). However, the cardiomyopathies observed in these two mutant strains likely arise from distinct cellular pathologies due to isoform specific functional differences. PI3Kγ-KO, but not PI3Kα-DN, mutants develop an increase in intracellular levels of cAMP due to reduced PDE activity(Crackower et al., 2002; Patrucco et al., 2004). Increases in cAMP have been shown to up-regulate expression of matrix metalloproteinase (Melnikova et al., 2006), proteolytic enzymes of that target the ECM and cell-cell adhesion complexes (Covington et al., 2006; Dwivedi et al., 2009), illustrating a potential

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mode of maladaptive ECM remodelling and a likely cause of heart failure in pressure overloaded PI3Kγ-KO hearts. Despite their redundant kinase activity, PI3K isoforms demonstrate unique effects on cardiac hypertrophy. For instance, while PI3Kα activity was shown to mitigate maladaptive hypertrophy (McMullen et al., 2007), PI3Kγ activity may exacerbate it. Studies of PI3Kγ-KO and PI3Kα-DN cardiomyocytes demonstrated that PI3Kγ and PI3Kα respectively activate and inhibit ERK (McMullen et al., 2007; Patrucco et al., 2004), whose activation has been associated with pathological hypertrophy (Ainscough et al., 2009; Lorenz et al., 2009).

6.3.2 PTEN PTEN is a lipid phosphatase which dephosphorylates PIP3 into PIP2 (Stambolic et al., 1998), and hence is a negative regulator of PI3K activity (Fig. 6.3) (Oudit and Penninger, 2009). PIP3 is the primary catalytic product of PI3K which activates AKT, stimulating physiological growth (Vanhaesebroeck et al., 1997). AKT signaling also protects the heart from pathological hypertrophy (Rigor et al., 2009). The PTEN null and caPI3Kα mutants, which have excessive PI3K activities, are resistant against pressure overload induced heart failure (McMullen et al., 2007; Oudit et al., 2008). Though PTEN-KOs are resistant to pressure overload, their maladaptive response to GPCR agonists worsens (Oudit et al., 2008) suggesting that loss of PTEN is specifically protected against biomechanical stress. Indeed, PTEN is a key regulator of cell-matrix adhesion and migration (Gu et al., 1999, 1998; Larsen et al., 2003; Oudit et al., 2004) and PTEN regulate several distinct pathways involved in remodeling of cell adhesion complexes and cytoskeletal organization (Gu et al., 1999, 1998; Larsen et al., 2003; Oudit et al., 2004). For example, PTEN directly dephosphorylates Shc and FAK which in turn modulates cell adhesion complexes and the intracellular actin cytoskeleton (Gu et al., 1999, 1998; Larsen et al., 2003), independent of AKT/PKB activity.

6.4 Intercellular Propagation of Mechanical Signals 6.4.1 Cell-Cell Adhesion Intercellular transmission of mechanical signals is propagated in two distinct mechanisms: by directly stretching and straining neighbouring cells, and via paracrine signaling. The direct transmission of force between adjacent cardiomyocytes is propagated by cell-cell adhesion complexes called adherens junctions, which are composed of several linkage proteins, with the traditional components being cadherins (with N-cadherin being the primary isoform in the heart) and catenins (α and β) (Fig. 6.4a). Though it is often assumed that like many striated muscles, cardiomyocytes only experience mechanical stress along its longitudinal axis

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Fig. 6.4 A schematic of adherens junctions (a) and gelsolin activity (b). The intracellular domain of the transmembrane protein N-cadherin is anchored to actin filaments by β- and α-catenin while gelsolin cleaves actins and prevents repolymerization by capping the actin filament

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via intercalated discs, it seems unlikely that cardiomyocytes do not experience lateral strain from the surrounding ECM or from neighbouring cells, or exert lateral force during systole (Schoenberg, 1980). In fact, a study elegantly demonstrated that roughly 80% of N-cadherin junctions between adjacent cardiomyocytes are located in the peripheral (lateral) membrane (Pedrotty et al., 2008). Furthermore, these lateral junctions allow cardiomyocytes to adhere to other cell types as intercalated discs are only formed between cardiomyocytes. Aside from propagating mechanical force, adherens junctions have also been postulated to interact with PI3K to mediate AKT signaling (Tran et al., 2002). Increased and decreased adherens junctions were shown to respectively improve and weaken basal AKT phosphorylation (Tran et al., 2002). However, it is possible that this loss of AKT stimulation could be indirectly due to weakened mechanosensitivity as there is still little evidence of direct interactions between cadherins and the PI3K pathway.

6.4.2 Autocrine and Paracrine Signaling While the conversion of mechanical signals into chemical signals occurs in the intracellular compartment, the spreading of this chemical signal from cell to cell occurs through autocrine and paracrine signaling; hence, these processes play key roles in stretch induced cardiac hypertrophy (Cingolani et al., 2001; van Wamel et al., 2001). For instance, Ang II, ET-1, and TGF-β have all been implicated in mediating cardiac hypertrophy in hemodynamic overload models (Arai et al., 1995; Ishiye et al., 1995; Ito et al., 1994; Ruzicka et al., 1995; Takahashi et al., 1994). In response pressure overload, the heart increases expression of both endogenous ET-1 and ET-1 receptors appropriating ventricular hypertrophy (Arai et al., 1995), which can be attenuated by applying ET-1 receptor blockers (Ito et al., 1994). Similarly, in response to volume overload, the heart develops ventricular hypertrophy as a result of increased Ang II levels (Ruzicka et al., 1995), but can be reversed via pharmacological inhibition of Ang II receptors (Ishiye et al., 1995). TGF-β is primarily expressed by non-cardiomyocytes during basal condition, but in response to pressure overload, cardiomyocytes surprisingly become the primarily secretor of TGF-β (Takahashi et al., 1994). TGF-β paracrine signaling also stimulates excess collagen deposition by fibroblasts leading to fibrosis and stiffness of the myocardium (Eghbali et al., 1991). Cardiac hypertrophy following mechanically induced paracrine signaling has also been demonstrated in vitro. Cultured cardiomyocytes exposed to stretch release ET-1 and TGF-β, which in turn causes an increased expression of the hypertrophy marker ANP (van Wamel et al., 2001). Whether cardiomyocytes express ET-1 in the absence of stretch appears to be under debate (Suzuki et al., 1993; van Wamel et al., 2001), but in either case, the level of ET-1 excretion in the absence of stretch is likely insufficient for stimulating the expression of hypertrophic markers (van Wamel et al., 2001). Similarly, studies investigating whether Ang II is expressed by acutely stretched cardiomyocytes have yielded mixed results (Sadoshima et al.,

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1993; van Wamel et al., 2001; Yamazaki et al., 1999). Nevertheless, multiple studies have confirmed that endogenous secretion of Ang II mediates, but is not necessary for inducing, hypertrophy in stretched cardiomyocytes (Sadoshima and Izumo, 1993; Sadoshima et al., 1993; van Wamel et al., 2001; Yamazaki et al., 1999). Furthermore, it should be noted that Ang II mediated hypertrophy occurs partially via increased production and excretion of ET-1 (Cingolani et al., 2001). The release of Ang II by stretched cardiac fibroblasts can also induce hypertrophy in unstretched cardiomyocytes (van Wamel et al., 2001, 2000). Similarly, stretched cardiac fibroblasts also produce TGF-β, altering gene expression in neighbouring cardiomyocytes (van Wamel et al., 2001). Recently, a co-culture study demonstrated that paracrine signaling from cardiac fibroblasts can induce cardiomyocyte proliferation (Ieda et al., 2009). Together, these in vitro and in vivo studies describe the prominent potential of intercellular dependence and regulation between cardiomyocytes and cardiac fibroblasts in mechanosensitivity. An array of other factors, such as ANP, is also excreted to mediate cellular changes in response to mechanical stress (described in Sections 6.3.1 and 6.3.2) (Ruwhof et al., 2000; van Wamel et al., 2001, 2000). Hence, given the current knowledge of autocrine and paracrine effectors in mechanically stimulated pathways in the heart, it is clear that diffusible chemical factors are crucial in mediating mechanotransduction.

6.5 Myocardial Remodelling 6.5.1 Extracellular Remodelling The loss of cell-ECM linkage proteins in the heart almost always leads to cardiac dysfunction due to the inability to tolerate stressful conditions. Similar results are observed from the loss of signaling proteins such as FAK and ILK. As previously discussed, the inability to detect and to respond to mechanical stress can incapacitate the heart’s ability to initiate proper compensatory mechanisms ultimately leading to heart failure. Increased mechanical work load and hemodynamic stress is often closely correlated with concomitant intracellular and extracellular remodelling. In the heart, like in many other organs and tissues, extracellular remodelling is primarily regulated by MMPs (Kassiri and Khokha, 2005; Spinale, 2007). MMPs are a family of potent proteolytic enzymes that digest, among others, collagen, laminin and fibronectin and are inhibited by interactions with tissue inhibitors of metalloproteinases (TIMPs) (Kassiri and Khokha, 2005). Indeed, loss of TIMP3 leads to an exacerbation of pressure-overload induced adverse myocardial remodelling and heart failure (Kassiri et al., 2005). Several mechanically induced signaling pathways, specifically PI3K pathways, stimulate MMP expression and activation (Grote et al., 2003; Hess et al., 2003; Ispanovic and Haas, 2006; Zahradka et al., 2004). For instance, stretch induces NAD(P)H mediated formation of reactive oxygen species (ROS), increasing expression and activation of pro-MMP2 and the release of active MMP2 (Grote et al.,

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2003). As this suggests, chronic exposure to stressful hemodynamic conditions may lead to over-expression and over-activation of MMPs resulting in excess ECM degradation. In turn, this could reduce the efficiency and effectiveness of cell-ECM adhesions making the heart unable to respond to subsequent increases in biomechanical stress and increasing the susceptibility to the development of a dilated cardiomyopathy and heart failure. Aside from their influence on the ECM, several MMP isoforms also cleave the extracellular portion of cadherins (Ito et al., 1999; Uglow et al., 2003), proteins that make up cell-cell adhesion complexes such as adherens junctions and desmosomes (Stokes, 2007; Zuppinger et al., 2000). In addition to the reduction of cadherin-based intercellular junctions, this process also releases β-catenin, an intracellular component of cadherin complexes which translocates to the nucleus to stimulate, among others, cell proliferation and survival (Table 6.1) (Dwivedi et al., 2009; Uglow et al., 2003). A recent study illustrated that β-catenin shedding is necessary for postinfarction adverse left ventricular remodelling (Zelarayan et al., 2008) implying that excess or perhaps even normal level of mechanical stress post-infarction may exacerbate ventricular damage through differential gene expression and imbalanced matrix remodelling.

6.5.2 Intracellular Remodelling 6.5.2.1 Actin Intracellular remodelling of the actin cytoskeleton plays an important role in mechanotransduction because many cell-cell and cell-ECM adhesion complexes rely on coupling to cytoskeletal actin filaments to gain anchorage and to maintain location. Cytoskeletal actin regulation is mediated via a combination of polymerization, depolymerisation, and modifications of actin crosslinking proteins (Pollard, 2007; Vicente-Manzanares et al., 2005), maintaining actin structures in a dynamic equilibrium. Cytoskeletal actin can also relay extracellular force to the nucleus, altering its morphology (Maniotis et al., 1997). This immediate transmission of mechanical stimulus to the nucleus has been demonstrated to be capable of directly inflicting changes in gene expression (Guilak et al., 2000), though this hypothesis still remains to be elucidated. More details of the mechanical properties of ECM-nuclear connectivity are described elsewhere (Dahl et al., 2008). Disruption of actin polymerization prevents cell adhesion (Sanger and Holtzer, 1972), whereas cellular adhesion to matrix stimulates actin reorganization (DeMali et al., 2003); albeit, the resulting actin infrastructure depends on the cell’s external environment (DeMali et al., 2003; Hannigan et al., 2005). For instance, fibroblasts in 2D cultures spread out and cover a large surface area while fibroblasts in 3D cultures have enhanced migration and adopt a star like shape (DeMali et al., 2003) suggesting that environmental differences detected via cell-ECM adhesion help direct migration and modulate morphology. Interestingly, the formation of focal adhesions and the association of FAK at focal adhesions are much more prominent in 3D cultures than 2D cultures (Berrier and Yamada, 2007; DeMali et al., 2003)

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suggesting restricted FAK signaling in 2D cultures. In addition to adhesion and nuclear changes, stretch also stimulates changes in actin structure (Simpson et al., 1999). For instance, pressure overload has been observed to induce changes in actin isoform expression prior to ventricular decompensation (Berni et al., 2009). Also, stretched cardiomyocytes induces reinforcement and thickening of actin filaments and increase actin turnover in vitro via proteins such as zyxin and gelsolin (Liepina et al., 2003; Simpson et al., 1999; Sun et al., 1999b; Yoshigi et al., 2005; Yu et al., 1992) as discussed below. 6.5.2.2 Zyxin Zyxins are subcellular proteins found at the cellular substratum attached to cell adhesion complexes, such as focal adhesions, as well as to cytoskeletal actin (Beckerle, 1998; Bershadsky et al., 2003; Hoffman et al., 2003). Zyxin’s molecular structure reveals its role as both a scaffold for focal adhesion complexes as well as a docking site for the ENA/VASP family of proteins (Hoffman et al., 2003; Krause et al., 2003), which are abundantly found in intercalated discs of cardiomyocytes (Brancaccio et al., 2006). Under resting conditions, zyxins are found bound to vinculin and α-actinin at focal adhesions close to Z-discs (Crawford and Beckerle, 1991; Yoshigi et al., 2005). Interestingly, unlike other integrin complex proteins such as FAK, ILK, vinculin, or paxillin, (Li et al., 1999; Sawada and Sheetz, 2002; Sharp et al., 1997; Torsoni et al., 2003), zyxins move away from integrin-linked complexes toward actin filaments in response to stretch, causing realignment of actin and thickening of actin filaments according to the direction of stretch (Yoshigi et al., 2005). Interestingly, zyxin can localize between the nucleus and the cytoplasmic membrane and can thus potentially mediate gene regulation (Nix and Beckerle, 1997; Nix et al., 2001). For instance, exposure of cardiomyocytes to atrial natriuretic peptide accumulates zyxin at the nucleus inducing anti-apoptotic effects (Kato et al., 2005). Zyxin is not necessary for survival as zyxin knockouts are both viable and fertile (Hoffman et al., 2003), yet the lack of zyxin compromises the tolerance to mechanical stimuli (Yoshigi et al., 2005). When stretched, zyxin mutant cells have impaired thickening, but not reorganization of actin filaments, which can be rescued via the addition of exogenous zyxin (Yoshigi et al., 2005). Reorganization of actin filaments was likely not impaired due to compensation from other actin remodelling proteins such as gelsolin. 6.5.2.3 Gelsolin Gelsolin is a calcium dependent monomeric protein that regulates actin reorganization by severing and capping actin filaments (Fig. 6.4b) (Liepina et al., 2003; Sun et al., 1999b; Yu et al., 1992). Increases in intracellular calcium concentrations stimulates gelsolin to sever actin, following by its binding to the truncated actin filament acting as a cap to prevent subsequent regeneration and repolymerization (Sun et al., 1999b). The removal of the gelsolin cap requires the binding of PIP2 (Liepina et al., 2003; Yu et al., 1992). Cell adhesion causes gelsolin to co-localize with members

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of the integrin complex such as FAK, PI3K, c-Src, paxillin, vinculin, and talin suggesting that mechanical stimulation recruits gelsolin to integrin-linked complexes (Chellaiah et al., 2001). Gelsolin is up-regulated in DCM, ventricular hypertrophy and heart failure (Yang et al., 2000). Moreover, gelsolin null mice have better contractile function post infarction compared to wildtypes (Li et al., 2009) suggesting that gelsolin activity may be necessary in mediating ventricular damage and various forms of cardiomyopathy (Nishio and Matsumori, 2009). However, gelsolin activity has been argued to be beneficial to the survival of cardiac cells as it can stimulate anti-apoptotic signaling (Azuma et al., 2000; Koya et al., 2000). Considering gelsolin’s potency in cardiac cells as well as other cell types and the central role of actin in both cellcell and cell-ECM adhesion complexes, Gelsolin’s role in cardiac remodelling and mechanotransduction poses an important target for future investigation.

6.6 Conclusion and Perspectives Mechanosensitive signaling pathways in cardiac cells orchestrate the dynamic process of intracellular and extracellular cardiac remodelling, stimulating an abundance of physiological processes such as hypertrophy. However, when exposed to pathological stress, such as pressure- and/or volume-overload, the same signaling pathways act adversely causing cardiomyopathy and heart disease. In conclusion, mechanical signaling in the heart is a fundamental adaptation to biomechanical stress and suggests that a shift of focus toward repairing damage in cell adhesion complexes and restoring normal mechanotransduction may have therapeutic benefits. Ultimately, the manipulation of these various signaling cascades may indeed provide novel therapeutic avenues for the prevention and/or treatment of heart failure. Acknowledgements GYO is a Clinician-Investigator Scholar of the Alberta Heritage Foundation for Medical Research and ZK is a New Investigator of the Heart and Stroke Foundation of Canada. We acknowledge the financial support from the Canadian Institute for Health Research (GYO grant 86602, ZK grant 84279), the heart and Stroke Foundation of Canada (GYO) and the Alberta Heritage Foundation for Medical Research (GYO).

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Chapter 7

Atomistic Molecular Simulation of Gating Modifier Venom Peptides – Two Binding Modes and Effects of Lipid Structure Kazuhisa Nishizawa

Abstract GsMTx4, a gating-modifier peptide obtained from tarantula venom has been a valuable tool for investigating the gating mechanisms of mechanosensitive channels. GsMTx4 is thought to act at the channel/lipid interface by modifying the structure of the surrounding lipid molecules. However, the atomistic details of these actions are poorly understood. Here, the studies of GsMTx4 and related peptide toxins that inhibit the voltage activation of various ion channels are reviewed, with emphasis on the results of molecular dynamic (MD) simulation analyses. Free energy profile analyses suggest that these toxins exhibit two modes of binding to lipid membrane, namely, the shallow mode and the deep mode. These toxins favor the deep mode, especially in membranes rich in saturated lipid acyl chains, which make the headgroup layer tight. It is hypothesized that in the case of HaTx the deep mode is the action mode, while for GsMTx4 the two modes can explain the concentration-dependent (biphasic) effect of GsMTx4 that has recently been reported. The possibility that such toxins seek out specific types of lipid molecules is discussed. Simulation results support the view that the channel/GsMTx4 (or HaTx)/lipids make a tertiary complex crucial to the effectiveness of the toxin and therefore binding of the toxin to channels occurs only in the presence of lipid molecules with appropriate structures. Keywords GsMTx-4 MscS · MscK · Stretch-activated channels · Mechanosensitive channels · Inhibitory cysteine knot peptides · Spider venom · Molecular dynamics simulation · HaTx

K. Nishizawa (B) Department of Laboratory Medicine, Teikyo University School of Medical Technology, Kaga, Itabashi, Tokyo 173-8605, Japan e-mail: [email protected]


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7.1 Introduction Since the identification and characterization of the tarantula toxin GsMTx4 by Sachs and coworkers (Chen et al., 1996; Suchyna et al., 2000), this cationic hydrophobic polypeptide has been utilized as a specific blocker of stretch-activated cation channels (SACs). As such, GsMTx4 has proven to be a valuable tool in studies analyzing the involvement of SACs in various phenomena. The seminal papers by Chen et al. (1996) and Suchyna et al. (2000) have reported a role for SACs in hypotonic cell swelling-induced Ca2+ increase with the use of the venom from which GsMTx4 was later isolated. More recently, several additional targets of GsMTx4, including TRPC6 (Spassova et al., 2006) and MscK and MscS of E. coli have been reported (Hurst et al., 2009). The number of studies showing pharmacological potentials of GsMTx4 is also increasing (for review, Bowman et al., 2007). In addition to such medical interests, GsMTx4 has drawn much interest from biophysicists after the elucidation of a wide variety of target channels of different phyla of organisms. GsMTx4 inhibits cationic SACs in a variety of vertebrate cell types such as chick heart, rat astrocytes and skeletal muscle and human smooth muscle. How does it modify the gating of such different channels with different kinetic properties and presumably different structures? It is likely that, after GsMTx4 adsorbs to lipid bilayer membranes, it modifies their mechanical properties, thereby modifying the energy from bilayer tension required for gating SACs. It is also likely that the effects of GsMTx4 on membrane proteins are dependent on its partition into the interface between channels and lipid molecules. However, structural and dynamical features enabling these effects on various channels and motor molecules are not well understood. This review primarily focuses on recent studies using MD simulation analyses and experimental results of GsMTx4 and related gating modifier peptides including HaTx, a gating modifier of Kv channels. Based on these findings, we propose a model of membrane behaviors of these toxins and the mechanisms by which such toxins affect the gating of channels. A limited amount of information on the potential medical applications of GsMTx4 can be covered here, thus interested readers are referred to more comprehensive reviews on GsMTx4 and ICK peptides. Bowman et al. (2007) reviewed topics of history, properties, mechanisms and pharmacology of GsMTx4. HaTx and other gating modifier toxins are detailed in Swartz (2007). Recent reviews on ICK peptide gating modifiers include Sollod et al. (2005) and Escoubas and Rash (2004). Computational methods for ion channels, including homology modeling and MD simulations, have been reviewed by Tai et al. (2008). Computer modeling and simulation studies of bacterial mechanosensitive channels have been reviewed by Corry and Martinac (2008a, b). Recent progress on MD simulations of membrane proteins is covered by Lindahl and Sansom (2008).

7.2 Discovery and Functions of GsMTx4 GsMTx4 is a 34 amino acid residue peptide isolated from the venom of a spider, the Grammostola spatulata tarantula. The inhibitory activity of the crude venom on

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SACs was originally discovered in a study to evaluate the importance of L-type Ca2+ channels and SACs in the hypotonic cell swelling-induced Ca2+ increase (HICI) in GH3 cells (Chen et al., 1996). In their study, the finding that the extracellular Ca2+ is the source of the Ca2+ influx of HICI led the authors to determine the route for the Ca2+ influx; both L-type Ca2+ channels and SACs were likely involved in the HICI. Importantly, the venom was found to block the HICI without blocking L-type Ca2+ channels, suggesting the sensory role for SACs in the events of HICI. Sachs and coworkers continued their study and isolated the molecule responsible for the inhibition and named it GsMTx4 (Grammostola mechanotoxin number 4) (Suchyna et al., 2000). They further showed that GsMTx4 inhibits SACs in astrocytes and heart cells (Suchyna et al., 2000). The group also determined the structure of GsMTx4 in solution using NMR (Oswald et al., 2002). Important insights into the mechanism for gating modification by GsMTx4 were provided by an experiment using the enantiomer. GsMTx4 and its enantiomer, enGsMTx4, exhibit similar efficacy in inhibiting stretch-activated channels of rat astrocyte, suggesting that membrane distortion, rather than specific molecular recognition, plays an important role (Suchyna et al., 2004). Despite that GsMTx4 is unlikely related to gramicidin A, GsMTx4 causes a 10–25 fold increase in the channel appearance rate of gramicidin A, leading the authors to conclude that GsMTx4 exerts its effect on the gramicidin A channel-formers by altering the lipid packing at the bilayer/solution/channel interface. Clearly, the action of GsMTx4 is not dependent on the traditional lock-and-key model of ligand protein interactions postulated for a number of inhibitors of enzymes (Suchyna et al., 2004). This finding also drew the interest of researchers with respect to how GsMTx4 can modify the mechanical properties of bilayer membranes. While the potential pharmacological value of GsMTx4 cannot be fully covered here, notable findings include GsMTx4-dependent inhibition of atrial fibrillation (Bode et al., 2001) and Yeung et al. (2005) have shown that GsMTx4 inhibits Ca2+ elevation in muscle fibers from the mdx mouse, a mouse model of Duchenne Muscular Dystrophy. This suggests that GsMTx4 may be effective in protecting dystrophic muscle cells in which SACs activation, leading to Ca2+ influx, is easily triggered due to muscle cells defective in the dystroglycan complex (Yeung et al., 2005). As reviewed in the next section, GsMTx4 is a member of a peptide family named ICK (inhibitory cysteine knot) peptides (Narasimhan et al., 1994). Different ICK peptides modify the gating property of a variety of channels (Swartz, 2007). Among ICK peptides, GsMTx4 is unique in its high specificity to SACs. Recently, several genes belonging to the TRP (transient response potential) channel family have been cloned and a consensus is emerging that TRP channels form an important group of SACs (Yin and Kuebler, 2009). A recent example of the specificity of GsMTx4 for SACs was also obtained from the TRC family channels. GsMTx4 inhibits TRPC1 (Maroto et al., 2005, Bowman et al., 2007), although the presence of endogenous TRPC1-like SACs confounded the effect of GsMTX4 on TRPC1 (Bowman et al., 2007). On the other hand, inhibitory effects of GsMTx4 on TRPC6 channels were reported in a more reliable system (Spassova et al., 2006). In their study, HEK293 cells were transfected with the TRPC6 cDNA under the control of an inducible

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promoter. TRPC6 was shown to be indeed a direct sensor of the mechanically and osmotically-induced membrane stretch. These stretch responses are blocked by GsMTx4. TRPC6 channel is also activated by diacylglycerol (DAG), which in turn is induced by the receptor-induced phospholipase C activation. Moreover, they showed that GsMTx4 blocks DAG-induced TRPC6 activation, suggesting that both chemical and mechanical lipid sensing by the channel have a common molecular basis (Spassova et al., 2006). Intriguingly GsMTx4 also has an effect on prokaryotic mechanosensitive channels. Using E.coli spheroplasts, Martinac and coworkers reported that GsMTx4 exhibits a biphasic effect on the gating of MscS and MscK (Hurst et al., 2009). That is, at low concentrations of 2–4 μM the pressure sensitivity of the gating of MscS and MscK was decreased and therefore gating was hampered, whereas at concentrations of 12–20 μM the pressure sensitivity was increased and gating was facilitated (Hurst et al., 2009). This finding suggests that there are at least two different kinds of interaction between GsMTx4 and lipid membrane or interaction among GsMTx4, lipid membrane and channels as considered below. While it is reasonable to argue that GsMTx4 is specific to the stretch-activated channels, an even wider spectrum of membrane proteins may turn out to be targets of GsMTx4 in the future. GsMTx4 has recently been shown to have effect on the voltage sensing of a Kv channel; Elinder and coworkers found that 5 μM GsMTx4 causes a positive shift (+5.7 mV) in the G(V) curve of the Shaker K channel) (Börjesson and Elinder, 2008), meaning that depolarization to the higher voltage (by +5.7 mV) is necessary to induce the similar level of conductance . Strikingly, Fang and Iwasa (2006) found that GsMTx4 has an inhibitory effect on the outer hair cell (OHC) motor; the presence of 5 μM GsMTx4 causes a positive shift (−39.8 to −18.4 mV) in the bell-shaped curve of the membrane capacitance. Let us briefly review the OHC motor. It is well known that the primary function of the organ of Corti of cochlea is to change sound-induced vibrations into electrical signals. OHCs, however, not only detect and convert sound in this way but also they convert the electrical signals into changes in cell length. This response is considered to be essential to mechanical tuning unique to mammalian ears. The mechanical response is accompanied by a current similar to the gating current observed for voltage-activated ion channels, although in the OHC case the current is manifested as nonlinear membrane capacitance. This transient capacitive current reflecting changes in membrane voltages is dependent on motor proteins such as the membrane protein prestin (Zheng et al., 2000). Therefore, while GsMTx4 is relatively specific toward SACs, the range of target proteins may be wider. Overall findings thus far suggest that the structural requirement of the target molecule is not stringent. Rather, GsMTx4 appears to be located at the protein/lipid boundary for many membrane proteins and a large part of its action is likely to be related to the change in structural and/or mechanistic properties of lipid molecules in the vicinity of the channel. Lipid partition has been investigated from the point of view that it enhances the binding or the local concentration of toxins. Posokhov et al. (2007) have shown that GsMTx4 bonds strongly to both anionic and zwitterionic membranes. By contrast,

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SGTx1, another member of the ICK peptide family (see below) acts only with anionic lipids. However, this study used POPC (palmitooleoyl-phosphatidylcholine) and POPC-POPG (palmitooleoyl-phosphatidylglycerol) membranes that did not contain DPPC (dipalmitoyl-phosphatidylcholine), sphingomyelin or other lipid mixtures that have potential to create domains rich in saturated lipid tails. As considered below, our MD simulation study showed that the use of DPPC has a dramatic increase of the binding energy (∼ G = −100 kJ/mol) compared with POPC, indicating the necessity to take the lipid tail structure into account.

7.3 Gating Modifier Toxins Related to GsMTx4 Besides GsMTx4, a variety of peptide toxins isolated from the venom of spiders, scorpions, sea anemone, cone snails and snakes have drawn the attention of researchers because of their ability to inhibit voltage-gated ion channels (Catterall et al., 2007; Swartz, 2007). In a recent thorough review, Swartz (2007) classified such toxins into two groups. The first group consists of toxins, such as charybdotoxin and agitoxin, which inhibit Kv channels by binding to the pore forming regions (MacKinnon and Miller, 1989). The second group of toxins, often called gaitingmodifier toxins, influences the gating mechanism by altering the relative stability of the closed, open or inactivated states. For example, β-scorpion toxin is a wellstudied example of the second group and specifically enhances activation of sodium channels by holding the S4 segment in domain II in its outward position. (Cestèle et al., 1998; Catterall et al., 2007). For β-scorpion toxins, homology modeling and extensive docking analyses have been done by Cestèle et al. (2006). Such toxins have been utilized to study the structure and function of voltage sensing machinery of ion channels. ICK peptides constitute a well-known subclass of gating-modifier toxins. The name “ICK” comes from the fact that these peptide toxins contain a common structural motif known as “inhibitory cysteine knot (ICK)” (Narasimhan et al., 1994; Takahashi et al., 2000; Sollod et al., 2005; Bosmans et al., 2006). Table 7.1 shows examples of ICK peptides and their targets. (Table 7.1 is not an exhaustive list of ICK peptides but rather focuses on the cases where the time for the development of the effects or the affinity to the channel in the membrane can be inferred at least approximately.) GsMTx4 is unique in terms of its specificity to SACs rather than voltage-gated ion channels (Table 7.1). Figure 7.1d shows an alignment of amino acid sequences of several ICK peptides. ICK peptides have of six cysteine residues forming three disulphide bonds (Fig. 7.1d), creating the cysteine knot motif. All these peptides share a common structural feature, a “hydrophobic patch (or protrusion)” surrounded by a unique “charge belt” (Fig. 7.1a, b) (Oswald et al., 2002; Takahashi et al., 2000). The hydrophobic patch contains either a F or W residue (shown in green in Fig. 7.1d). The overall feature of the structure is likely similar to that of GsMTx4, yet the sequence similarities are generally low. Amino acid identity of GsMTx4 is 47% with GxTX-1E, 35% with VSTx1 and 21% with HaTx (Fig. 7.1d;, Herrington et al., 2006; Swartz, 2007)

α1A-Cav , Xenopus

α1A-Cav , Xenopus

Kv2.1, Xenopus

Kv2.1, Xenopus

KvAP, POPE/POPG¶ Kv4.2, COS cell

Kv2.1, COS cell

Kv2.2, COS cell

ω–GrTx-SIA

ω–AgaIVA

ω–AgaIVA

SGTx

SGTx

VSTx1

ScTx1

ScTx1

ScTx1

Kv2.1/F274A, Xenopus Kv2.1 in Xenopus

HaTx

IC50 = 1.2 nM (0 mV) IC50 = 12.7 nM (0 mV) IC50 = 21.4 nM (0 mV)

Kd = 2.7 μM (4 sites) Kd ∼2 μM (4 sites) IC50 = 30 nM

Kd = 160 nM (1:1)

Kd = 19 μM

Lee et al. (2004) Jung et al. (2005) and Lee and MacKinnon (2004) Escoubas et al. (2002) Escoubas et al. (2002) Escoubas et al. (2002)

τ = ∼3000 s at 20 nM steadyc at 93 s, 30 nM steadyc at 101 s,100 nM steadyc at 113 s,100 nM

Li-Smerin and Swartz (1998) Winterfield and Swartz (2000) Winterfield and Swartz (2000) Wang et al. (2004)

Swartz and MacKinnon (1995) Swartz and MacKinnon (1997) Swartz and MacKinnon (1997) Li-Smerin and Swartz (1998) Phillips et al. (2005b)

Reference

τ = ∼20 s at 500 nM

τ = ∼10 s at 2.5 μM

τ = ∼50 s at 1 μM

τ = ∼200 s at 125 nM

τ = ∼5 s at 25 μM

τ = 5.1 s at 2 μM

τ = ∼20 s at 10 μM

α1A-Cav , Xenopus

HaTx

Kd = 40 μM (4 sites) Kd = 2.8 μM

τ = ∼4 s at 5 μM

Kv2.1, Xenopus

HaTx

τ = ∼500 s at 50 nM

Kd = 42 nM (1:1)

Kv2.1, Xenopus

HaTx

τ = 114 s at 200 nM

Kd = 42 nM (1:1)

Kv2.1, Xenopus

HaTx

Time for onsetb

Kd

Targeta

Toxin

Table 7.1 Target and time for onset of ICK peptide toxins

172 K. Nishizawa

Kv2.1, CHO cell

Ito1, rat heart myocyte Kv4.2, Xenopus

Kv4.3, Xenopus

Nav1.2/β1, Xenopus SACs, astrocytes capacitance, outer hair cell

GxTX-1E

HpTx2

PaTx1

CcoTx2

Diochot et al. (1999) Bosmans et al. (2006)

τ = ∼20–50 s at 50 nM τ =∼594 ms at 5 μM τ = ∼17.6 s at 5 μM

Kd = 630 nM Kd =3.1 μM

Suchyna et al. (2000) Fang and Iwasa (2006)

Sanguinetti et al. (1997)

Sanguinetti et al. (1997)

Herrington et al. (2006)

steadyc at 90 s, at 50 nM

τ = ∼50 s at 43 nM

Reference

IC50 = 1.5 nM (80 mV) IC50 = 15.8 nM (−10 mV) IC50 = 67 nM (−5 mV) IC50 = 28 nM (0 mV) IC50 = 8 nM

Time for onsetb

Kd

channel or current shown along with the expression system used for the experiment. ¶ Artificial bilayer system. b Time constant, τ , based on exponential curve fitting is shown along with the toxin concentration in the experiment. Some data are rough estimations from the reference figure. c The time at which the steady state was reached

a Target

Note that this list is not exhaustive, but focuses on the toxins/targets for which the time course of effect can be inferred from the literature, at least approximately. For quick reference, readers are referred to the SwissProt file (http://au.expasy.org/sprot/) for each toxin using the accession number shown in the Fig. 7.1 legend

GsMTx4 GsMTx4

HpTx3

Targeta

Toxin

Table 7.1 (continued)

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Fig. 7.1 (a–c) Structure of GsMTx4 and HaTx. The bottom and side view of GsMTx4 (a) and HaTx1 (b) and the initial orientation used for simulations (c). The hydrophobic patch is oriented around the center of the normal view and at the bottom of the side view. (a–c) are similar to Figure 1 of Nishizawa and Nishizawa (2007), in which the following coloring scheme is used; Hydrophobic residues (Ala, Cys, Ile, Leu, Met, Phe, Pro, Trp, Tyr and Val) are green; basic (Arg and Lys) are blue and acidic (Asp and Glu) residues are red. (d) Amino acid sequence alignment of ICK peptides. SwissProt accession numbers are as follows: GsMTx4 (Grammostola mechanotoxin number 4, Q7YT39), GxTx-1E (Guangxitoxin-1E, P84835), ω-Aga4A (P30288), CcoTx2 (P84508), HaTx1 (P56852), SGTx1 (P56855), VSTX1 (voltage sensor toxin 1, P60980), ProTx2 (Beta-theraphotoxin-Tp2a, P83476), HmTx1 (heteroscordatoxins, P60992), ScTx1 (stromatoxin 1, P60991), HpTx3 (heteropodatoxin3, P58427), PaTx1 (phrixotoxin1, P61230), ω-GrTx-SIA (Omega-grammotoxin-SIA, P60590)

Hanatoxin (HaTx) is a 35 amino acid residue gating-modifier of Kv channels (Swartz and MacKinnon, 1995, 1997). HaTx was isolated from the venom of the Chilean tarantula (Grammostola spatulata), the same venom from which GsMTx4 was found. HaTx (a mixture of HaTx1 and HaTx2, which are identical except for residue 13 which is Ser in HaTx1 and Ala in HaTx2) can bind to VSDs of closed Kv2.1 channels and dramatically shift the channel’s voltage dependence towards more positive voltages (Lee et al. 2003). It exhibits very slow action (τ = 114 s) despite its high affinity to the voltage sensor (Kd = 42 nM) (Swartz and MacKinnon, 1995). Besides GsMTx4 and HaTx, the ICK peptide gating modifiers include: SGTx1 from Scodra griseipes, which resembles HaTx and inhibits Kv2.1 (Marvin et al., 1999; Wang et al., 2004), VSTx1 isolated from Grammostola spatulata by screening inhibitory activity against KvAP (Ruta et al., 2003; Lee and MacKinnon, 2004), ProTx2 (Beta-theraphotoxin-Tp2a) from Thrixopelma pruriens which inhibits various Nav s (voltage-gated Na channels) and Cav s (Middleton et al., 2002), HmTx1 (heteroscordatoxins) from Heteroscodra maculate and ScTx1 (stromatoxin 1) from Stromatopelma calceata which are Kv2 and Kv4 inhibitors (Escoubas et al., 2002),

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and GxTx1E (guangxitoxin) from Plesiophrictus guangxiensis, which is a high affinity inhibitor of Kv2.1 (Herrington et al., 2006). Other ICK peptides include; ω−GrTx-SIA, which is another gating modifier toxin from Grammostola spatulata and inhibits voltage gating of Cav2.1 (Li-Smerin and Swartz, 1998); ω−Aga-IVA from Agelenopsis aperta, which is an inhibitor of P-type Ca2+ channels (Winterfield et al., 2000; Mintz et al., 1992); CcoTx2 (β-theraphotoxin-Cm1b) from Ceratogyrus marshalli (Straighthorned baboon tarantula), which is an inhibitor of several subtypes of Nav s (Bosmans et al., 2006); HpTx2 and HpTx3 (heteropodatoxin-2 and -3) from Heteropoda venatoria (Brown huntsman spider), which are inhibitors of different sets of Kv4 channels (Sanguinetti et al., 1997); PaTx1 (phrixotoxin-1) from Paraphysa scrofa (Chilean copper tarantula), an inhibitor of Kv4.2 and Kv4.3 (Diochot et al., 1999). For some ICK peptide gating modifiers, the time required for the development of the effect is known at least approximately (Table 7.1). In general, gating modifiers have a high affinity for voltage-activated ion channels, the dissociation constant (Kd ) or the half maximal inhibitory concentration (IC50 ) being in the 10–100 nM range. Such is the case with HaTx (Kd = 42 nM), VsTX1 (IC50 = 30 nM), GxTX1E (IC50 = 1.5 nM), HpTx2 (IC50 =15.8 nM) and PaTx1 (phrixotoxin1) (IC50 =28 nM). GsMTx4 is unique in its relatively low degree of affinity with Kd >500 nM; in the case of the OHC motor (Fang and Iwasa, 2006), Kd is ∼3 μM. It is somewhat puzzling that, even in the case of the OHC motor, the time for action, τ =∼17.6 s at 5 μM, is approximately in the same range as for HaTx. The affinity of VsTx1 to KvAP in the ‘absence’ of membranes is quite low (100–500 μM) (Lee and MacKinnon, 2004). Nonetheless, VSTx1 inhibits KvAP in membranes at a low concentration in solution (∼10 nM). This discrepancy led the authors to suggest that VSTx1 first binds to the membrane and then diffuses laterally until it finds its binding site on the channel (Lee and MacKinnon, 2004). For HaTx, the binding site in the Kv2.1 channel has been determined to be several residues in the S3b and S4 helices of the voltage-sensing domain, and that at least a large part of the interaction is mediated by a hydrophobic interaction (Li-Smerin and Swartz, 2000). (Of note, S4 is the helix which mainly carries the gating charges and constitutes a main part of votlage-sensing domain. (Börjesson and Elinder, 2008; Swartz, 2008) Swartz and coworkers have also shown using SGTx1 that the binding site is made up of a small number of residues and the most critical two residues, based on their alanine-scanning analysis, are F6 and W30 of SGTx1, implying the importance of hydrophobic interactions. (SGTx1, being similar to HaTx1 and easily expressed, was used for amino acid alteration analysis.) More recently, HaTx is thought to influence the interaction between the voltage sensing domain and lipids, suggesting the formation of a tertiary complex made up of the toxin, voltage sensor and lipids (Milescu et al., 2007). These findings point to a view that, while the toxin/channel interaction is mediated by only a small number of mainly hydrophobic residues (as in the case of SGTx1), the membrane/toxin interaction in the vicinity of the channel may cause deformation facilitating the toxin/channel interaction. Thus, the strength of the interaction between the ICK peptide toxin and channel maybe greatly altered by and even dependent on the interaction between the toxin and membrane.

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7.4 Unanswered Questions About HaTx, VsTx and GsMTx4 Because of the variety of functions of ICK peptides, researchers studying different ICK peptides tend toward quite different views about this family of toxins. For example, while GsMTx4 has low affinity and the target specificity also appears weak (Table 7.1), several toxins including HaTx and VSTx1 have higher affinity and specificity toward the target. Notwithstanding such differences, the following five qualities appear to be relevant to many of these toxins: (1) Slow action. They have small apparent association and dissociation rate constants, 103 –104 times slower than the pore blocking toxins (Swartz and MacKinnon, 1995; Ruta et al., 2003). In the case of HaTx, despite of its high affinity Kd = 42 nM, action is on the order of minutes. The action tends to become slow especially when the concentration is low (Table 7.1). One example is the slow equilibration rate of VSTx1 in inhibition of KvAP (∼60 min for 20 nM VSTx1) reconstituted in a planer membrane (POPE (palmitooleoyl-phosphatidylethanolamine): POPG=3:1) (Lee and MacKinnon, 2004). Intriguingly, in a recent experiment with the Xenopus oocyte expression system, after the removal of HaTx from the aqueous solution, the Kv2.1 channel activity recovered very slowly, requiring ∼1,000 s (Milescu et al., 2007), suggesting the presence of an energy barrier between the aqueous phase and the action site. (2) Small binding site and high affinity. It is puzzling why the apparent binding affinity of HaTx (and grammotoxin and VSTx1) to the voltage sensor is so high, considering the binding site on the channel consists of only a few amino acids on S3b and S4. However, from the VSTx1 study by MacKinnon and coworkers, a consensus is emerging that, not only the toxin partition into the lipid membrane enriches the toxin in the membrane, the tertiary complex toxin/channel/lipid is created and this strengthens the binding of toxin to channel proteins (Schmidt and MacKinnon, 2008). (3) Not activation, but inhibition. Unlike β-scorpion toxin that stabilizes activated states of Nav channels (Cestèle et al., 1998), HaTx and VSTx1 (and most other ICK peptide toxins) inhibit channel activation. It is unknown how they do so. (4) Effects of lipid mechanical states. Lipid mechanical state or structure influences the inhibition efficacy of these peptides. A recent study by Schmidt and MacKinnon (2008) employing a Kv1.2 channel with a voltage sensor paddle from Kv2.1 showed that VsTx1 activity depends on the mechanical state of the membrane. While VSTx1 has no effect on channel analyzed in whole cell recording of oocytes, VSTx1 causes a slowing of voltage activation similar to the effect observed on the channel embedded in planar bilayers. This suggests that voltage sensor toxins exert their effect by perturbing the interaction between the channel and the membrane lipid. The importance of lipid head groups is also shown by Milescu et al. (2007) in the finding that sphingomyelinase D, which removes choline from sphingomyelin, dramatically reduces the inhibitory effect of HaTx1.

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(5) Concentration-dependent effect. Martinac and coworkers showed that GsMTx4 action is biphasic; at low concentration they enhance but at high concentrations they inhibit the SACs (Hurst et al., 2009). With respect to (1) The membrane diffusion coefficient for a molecule like ICK peptides is probably ∼10−8 cm2 s−1 (Lee and Mackinnon, 2004). The authors also speculate that the collision frequency between VSTX1 and the KvAP channel is likely to be orders of magnitude faster than the observed kinetics of inhibition in their experimental system. Also with respect to (1), note that the time for action depends on the concentration of toxin; the lower concentration leads to slower action (e.g., HaTx, ω–AgaIVA and SGTx in Table 7.1). Although it is speculative, one possibility is that the toxin concentration influences the location of toxin, which in turn influences the rate of its access to the site of action. From (5) (concentrationdependent effects of GsMTx4) and the consideration given below, it is possible that HaTx at high concentration may be located at a distinctly different position from the position preferred at low concentration. On the other hand, even with the same concentration, the time for action differs significantly among the systems examined. For example, while GsMTx4 at 5 μM inhibits SACs on outside-out patch of astrocyte within ∼0.6 s, the effect of the same concentration of GsMTx4 on the OHC motor takes ∼17.6 s and the Kd is also high. It is currently difficult to address this issue because both lipid composition and peptide-protein interaction are influence the dynamics. With respect to (5), a recent study by Martinac and colleague (Hurst et al., 2009) using E. coli spheroplast is of particular interest. They measured two channel activities MscS (small conductance) and MscK (small conductance, K+ dependent), both activities being mediated by mechanosensitive channels but with distinct gating characteristics where the membrane tension required for activation is different. For both MscS and MscK, when GsMTx4 is applied to the outer leaflet of the bilayer at concentrations of 2–4 μM, channel activity is inhibited, while at higher concentrations of 12–20 μM, channel activity is potentiated. Moreover they reported that the initial inhibiting effect of 2–4 μM GsMTx4 becomes less pronounced with increasing time of exposure. They repeated the pressure application three times, each lasting ∼30 min including lag-times of 20 min. Their free-energy parameter analyses showed that the initially inhibiting 4 μM GsMTx4 had only a minimal degree of effect during the third pressure cycle (Hurst et al., 2009). As the authors discuss, these puzzling findings can be accounted for by considering the two binding modes, which we proposed based on our MD simulation analyses (see below).

7.5 The Deep Mode Hypothesis – an Insight from MD Simulations When we performed numerous simulations of a HaTx/DPPC membrane system with various positions and orientations of HaTx in the membrane, HaTx exhibited a tendency to move inward such that some of the charged residues interact with the inner

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leaflet of the membrane (Nishizawa and Nishizawa, 2006). This result prompted us to apply the free energy analysis to HaTx and GsMTx4 in lipid membranes. In this and the next sections, technical issues for such free energy analyses are considered.

7.5.1 Free Energy Analysis The free energy profile of a molecule along the membrane normal represents the density corresponding to the probability of localization of the molecule at different depths in the lipid bilayer membrane. For most atomistic peptide/membrane simulations, the timescale covered is within the range of ∼100 ns which is too short to sample the entire conformation/orientation of the peptide and lipids to the degree necessary for free energy analysis. Fortunately, a free energy profile can be obtained with use of several efficient methods. Current MD simulation packages such as NAMD (Phillips et al., 2005a) and Gromacs (Lindahl et al., 2001) provide a module for such free energy profile analyses. For small molecules or ions, the umbrella sampling may be the most popular approach (Roux, 1995). With this method, the position of the center of the harmonic potential that restrains the molecule (or atom) is successively varied among the simulations such that the successive distribution of the molecule overlaps. The free energy profile can be recovered from the resulting overlapping distributions using the weighted-histogram analysis method (Kumar et al., 1992). Recently, adaptive biasing force simulations have also been utilized, in which, instead of the harmonic potential, a potential that counteracts the free energy profile is used (Hénin and Chipot, 2004). However, while these methods are useful for very small molecules, the amount of computation necessary for peptide/membrane systems is not clear. Another approach is the one based on the steered molecular dynamics (Izrailev et al., 1998). This procedure allows one to move a set of particles in a specified direction. From the work done to move the particle, the free energy can be analyzed. The steered dynamics is effective in systems with strong binding such as avidin-biotin binding (Izrailev et al., 1998). However, the binding energy of most peptide/membrane interactions is so weak that the force measured is no stronger than the range of the frictional and elastic force. The lipids are so viscous that very ´ slow pulling (e.g., ∼1 Å/ns) is necessary to make the frictional and elastic forces negligible (Nishizawa, unpublished result). This is inconvenient because it is often difficult to establish whether the pulling rate is sufficiently slow. Yet another approach is the mean force measurement. We chose this method for the toxin/membrane free energy analyses (Nishizawa and Nishizawa, 2006, 2007). As described in Marrink and Berendsen (1994), the average force exerted on a particle constrained at a point can be measured directly. The constraining procedure is repeated at different positions and is briefly represented in Fig. 7.2a–d. The z-position (i.e., the vertical position with respect to the membrane plain) of the center of mass (COM) of the two lysine residues (K8/K28) in GsMTx4 is constrained and the average force acting on them measured (the circle of Fig. 7.2c). The advantage of this approach is that there is no complication from the frictional force.

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However, unlike the mean force on a water molecule in a membrane (Marrink and Berendsen, 1994), the mean force acting on a peptide in a membrane depends on the strong electrostatic interactions between the peptide and the lipid head groups and on the energy associated with lipid packing. Therefore, how to avoid the membrane deformation represents a major problem, as discussed in the next section. Another weakness of this method is that the requirement of a long equilibration time for each position necessitates the coarse discretization, which may lead to inaccuracy in the free energy analyses.

7.5.2 Technical Consideration of Free Energy Analysis of GsMTx4/Membrane System For both the steered dynamics analyses and the mean force measurement analyses, it is crucial to avoid the concerted movement of the proximal part of the membrane, which makes the meaning of ‘the position within the membrane’ obscure (Nishizawa and Nishizawa, 2006, 2007). Related to this, elasticity of the lipid molecules and the membrane may cause a bias in the mean force measurement. For example, if the mean force is measured immediately after the peptide is pulled to the position of interest, the force in the opposite direction becomes dominant due to the elasticity of the membrane lipid. Pre-equilibration required for this may become too long. To reduce such complications, introduction of some restraints on the lipid head group could be a reasonable compromise. In our analyses, we chose to restrain the z-coordinate of “C13” of DPPC (C13 forms an O-ester with the sn-2 acyl chain) in order to avoid the gross deformation of the membrane when the constraining position was varied (Nishizawa and Nishizawa, 2006, 2007). In practice, the ‘template frame’ was chosen from a trajectory exhibiting a mild but not intense membrane thinning (Fig. 7.2b). The mean force measurement in the presence of the restraint on C13 atoms was carried out with the use of the template frame and GsMTx4. In practice we chose the membrane for which the thickness of the proximal part of membrane was 2.4 nm based on the average distance of the C13 atoms between the leaflets (i.e., l2 =1.2 nm of Fig. 7.2b). Our preliminary mean force measurement showed that, with the presence of GsMTx4 in the shallow mode, the membrane thinning (from ∼2.8 to 2.4 nm) per se requires energetic cost about ∼15 kJ/mol (Nishizawa and Nishizawa, 2007). This value is much smaller than the energy difference between the shallow and deep binding modes. Even with the use of such a method, further care may become necessary to avoid a systematic bias arising from the preparation of the initial structures for the mean force measurement. One possibility, which we have not explored as of yet, is that charged residues may be neutralized by protonating Glu and Asp (or deprotonating Lys and Arg) sidechains so as to smoothen the peptide movement during the settingup procedure. Another possibility is to avoid the pulling procedure. The peptide may be placed and restrained, then the lipid molecules moved horizontally so that they fit to the peptide. These techniques may improve the free energy analyses in the future.

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In addition, the difference in the density of lipid molecules between the two leaflets may cause inaccurate measurement of the mean force. In our first paper in ICK peptides (Nishizawa and Nishizawa, 2006), a set of DPPC bilayers, namely 58/64, (number of DPPC of the upper/lower leaflets) was used. We also tested 61/64 but this alteration did not have a significant effect on the result. Although we have not tested this extensively, embedding the GsMTx4 in one of the leaflets a 64/64 DPPC membrane may lead to too high a density of lipid on the side containing the GsMTx4, which may in turn artificially promote the deep positioning of the toxin. To minimize such a density effect, we also tried a 6 nm × 12 nm wide membrane and embedded two GsMTx4 molecules on the two sides of the membrane to approximate the C2 symmetry. In this way the lipid density of the two leaflets become the same. This consideration also has a bearing on the issue of the local lateral pressure of the membrane and the effect of peptide toxins on the local pressure profile. Lindahl and Edholm (2000) developed an important method to calculate the pressure profile of membranes, which shows the lateral pressure at different depths of the lipid membrane. Gullingsrud and Schulten (2004) applied this method to several membranes with and without the applied tension. In the future, the effect of GsMTx4 on the pressure profile may well be investigated. In such studies, the number of lipid molecules in two leaflets may have to be carefully varied. The flip-flop of lipid molecules (i.e., the translocation of lipid molecules from one leaflet to the other leaflet of the bilayer) is a slow process (∼minutes) even with the help of fllipase (Zachowski and Devaux, 1990) and therefore, the uneven distribution of the lateral pressure among the two leaflets may be of relevance also in the real experimental system. After the binding of GsMTx4 to the membrane, the lateral pressure profile of the membrane may slowly change during the experiment as such flip-flop events occur.

Fig. 7.2 (continued) Our measurement showed that, with the presence of GsMTx4 in the shallow mode, the membrane thinning (from l1 (1.4 nm) to l2 (1.2 nm)) requires ∼15 kJ/mol of energy for DPPC and ∼14 kJ/mol for POPC (Supplementary File 6 of Nishizawa and Nishizawa, 2007). This value is much smaller than the energy difference between the shallow and deep binding modes (Fig. 7.2). (c) The mean force measurement. The z-position of the center of mass (COM) of the sidechain atoms of K8 and K28 was restrained and the mean force experienced by the COM was measured. (d) Deep mode binding. (e) Free energy profile of GsMTx4 and HaTx in a DPPC membrane. For GsMTx4, K8/K28 COM was used as the reaction coordinate (Nishizawa and Nishizawa, 2006). For HaTx, R24/K26 COM was used as the reaction coordinate. (f) Free energy profile same as (e) but with the POPC membrane (Nishizawa and Nishizawa, 2007). Cartoons below (e) and (f) represent our hypothesis on the factors influencing the stability of the deep mode relative to the shallow mode and our hypothesis on the transition from the shallow to the deep mode. Movements of GsMTx4 toward saturated acyl chain-rich domains and/or movements of such lipids toward GsMTx4 may promote the transition toward the deep binding mode. The transition may also be facilitated by binding of GsMTx4 to the membrane at a high density. (g) Two mode-based model for the GsMTx4 biphasic effect on E.coli MscK and MscS (Hurst et al., 2009). Note that in this illustration the Msc channel is represented by a simple tetramer composed of four transmembrane cylinders. (h) A deep mode-based model for the inhibition of the voltage sensor movement by HaTx

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Fig. 7.2 Free energy analyses and two-binding-mode model of GsMTx4 effects. (a–d) Steps assumed for energy analyses of GsMTx4 binding and penetration into the membrane. A simple illustration of the GsMTx4 structure is used; the shaded area represents the “charge band”, whereas the Lys and Arg residues are shown by long sidechains (zigzag lines). (a) GsMTx4 binds to the membrane from bulk water, forming the shallow mode. l1 is the distance from the bilayer center based on the average positions of phosphorus of DPPC for representative simulations. In our study, l1 was ∼1.4 nm. (b) A mildly deformed membrane was subjected to the membrane fixation, which was useful for the mean force measurement (see text). To make the interpretation easy, a membrane with a mild degree of thinning (l2 = 1.2 nm) was used. Of note, the approximate free energy associated with the membrane thinning can also be measured by the mean force measurement.

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7.6 Two Binding Modes of GsMTx4/Lipid Bilayer Membrane Results show that there are at least two binding modes (the shallow and deep mode) in the HaTx (and GsMTx4)-lipid bilayer membrane interaction (Nishizawa and Nishizawa, 2006, 2007) (Fig. 7.2e, f). This result is different from the MD simulation analyses of VSTx1 by other researchers (Bemporad et al., 2006; Wee et al., 2008), in which only the shallow mode was reported. However, their simulations appear to have been started with a limited range of position of the toxin relative to the membrane and used limited types of lipids, giving rise to a condition favorable to the shallow mode. In the following, we discuss from the viewpoint that the many interesting behaviors the toxins exhibit can be better explained by considering the two bonding modes rather than only the shallow mode binding. Phillips et al. (2005b) has shown that the fluorescence of W30 of HaTx was quenched by C9, C10, which are located near the middle of the hydrocarbon tail of the outer leaflet lipid of bilayer made of POPG:POPE=1:1. The quenching effect of C6, C7 was weaker than C9, C10 and even weaker than C11, C12, i.e., carbon atoms deeper into the hydrocarbon core. This finding does not support the shallow mode as the only binding mode; our simulations that started from different positions showed that the data was more consistent with the deep mode than with the shallow mode (Nishizawa and Nishizawa, 2006). In our view, the simple interfacial positioning (i.e., shallow mode) does not explain the very slow action of the toxin at least for HaTx. At least in our simulations, there is no substantial energy barrier between the extracellular space and the shallow mode (Nishizawa and Nishizawa, 2007). Although orientation can be varied exhaustively, our analyses on HaTx and Kv2.1 VSD interaction in the DPPC membrane suggest that there is no substantial barrier between the extracellular space and the shallow mode. Moreover, the shallow binding mode alone does not provide a good explanation for the stabilization by HaTx of the Kv2.1 channel in the resting state. Current understanding of the VSD structure in the resting state places S4 at a position lower than the HaTx binding site can access. Therefore, the shallow binding does not appear to significantly hinder the S4 upward movement. Also puzzling is the concentration-dependent effect of GsMTx4 (see above). To our knowledge, MD simulation analyses of ICK peptide carried out thus far have not directly addressed the structural features of the “tertiary complex” of toxin/channel/lipids. Nonetheless, based on the currently available findings, we hypothesize that the following model may be relevant to many of the ICK peptide toxins. (1) First, the toxins bind to the membrane at the interface (the shallow mode) (Fig. 7.2a, e). Due to the high energy barrier between the two modes, a spontaneous transition to the deep mode is a rare event, initially. (2) Several factors act to promote the transition to the deep mode (Fig. 7.2e). As the surface density of the toxin increases, which may happen with a high concentration of toxins in the solutions, pressure imbalance between membrane leaflets may become so high that it facilitates the transition of some toxins to the deep mode. Lipid structure and mechanical state may also be a factor. In our analyses, DPPC, which contains saturated acyl chains only, favored the deep mode over the shallow mode (Fig. 7.2e), whereas

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POPC membrane favored the two modes equally (Fig. 7.2f). (3) It is possible that, after several minutes, the toxin moves into the domains rich in the saturated acyl chains and/or the toxin ‘attract’ such lipids in its vicinity (Fig. 7.2e). In this way, deep mode binding may acquire stability. (4) Then, the toxin may interact with the binding site on the channel, if such a site is accessible (Fig. 7.2 g, h). In the case of HaTx, primarily the effect of (3) drives the toxin downward and this force may inhibit the S4 upward movement (Fig. 7.2 h). The concentration-dependent (biphasic) effect of GsMTx4 can be explained by our two mode-based model (Hurst et al., 2009). As we hypothesize in Fig. 7.2 g, GsMTx4 in the shallow mode may inhibit the opening of SACs. In the deep mode, further thinning of the membrane induced by the GsMTx4 molecules may facilitate opening the SACs. As our free energy analyses have shown, an energy barrier between the two binding modes explains the slow equilibration and recovery (after removal) for GsMTx4, HaTx and VSTx1 (Nishizawa and Nishizawa, 2007). Moreover, given the recent finding showing the low position of S4 in the resting state (Börjesson and Elinder, 2008) and the finding that VSTx1 interact at S4 of the KvAP, the deep binding mode appears to be more suitable to stabilize the resting state than the shallow mode. Finally, extensive chimera analyses showed that, for the Kv2.1 channel containing KvAP S4, in the presence of VSTx1, strong depolarization (∼100 mV) allows channel opening to some extent and this opening is accompanied by the dissociation of VSTx1 from the S4 of the chimera channel (Alabi et al., 2007). This finding is also consistent with the view that the deep mode is the action mode of VSTx1; considering that the partial channel opening is accompanied by upward movement of S4, it is difficult to explain, based only on the shallow mode, why VSTx1 exhibits the higher affinity in the resting (down) state than in the activated state. Shiau et al. (2002) carried out modeling by the docking method and suggested a binding model for HaTx1 and the Kv2.1 channel, although their analyses were based on a lipid-free system (for review, Huang et al., 2007). Intriguingly, our preliminary analyses show that the model by Shiau et al. (2002) is more consistent with the deep binding mode (our manuscript in preparation). Further simulation analyses are underway. In general, for ICK peptides other than GsMTx4, more experimental and modeling studies are necessary before a model based on the two modes of interaction can be proposed.

7.7 Conclusion and Perspectives The above considerations show that there is an array of gating-modifier peptides for voltage-dependent channels, but gating-modifiers specific to SACs are very few. Therefore, GsMTx4 is a valuable tool to study the structural and dynamical details of SACs. There is some evidence supporting the view that ICK peptides change the mechanical properties of lipid membranes in the proximity of channels (e.g., Schmidt and MacKinnon, 2008; Suchyna et al., 2004). To characterize the structure of the GsMTx4/lipid membrane/channel complex, MD simulation will continue to be useful. Although the number of studies of ICK peptides utilizing MD simulation

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is currently limited, importance of MD simulation as a powerful tool to integrate physiological and structural data is increasing. On the other hand, it is always true that the value of MD simulation depends on the structural and molecular physiological information, given the weaknesses and limitations of current MD simulations. Here, general limitations of MD analyses are considered first. Despite a wide array of technical advances, there are a number of limitations in simulation that impede faithful representation of the experiments. A major drawback is the limited simulation time that can be covered by MD simulations. The limitation in timescale is severe for analyses of protein-lipid interactions. Large-scale motions of lipids, internal dynamics of peptides in a viscous lipid environment, or peptide-peptide association in a lipid environment are beyond the timescale of tens of nanoseconds that can be achieved in many of simulation studies (Mátyus et al., 2007). For the purpose of the free energy analyses, combinations of the aforementioned techniques may help perform more meaningful simulations, but we cannot be free of the essential problem. For example, combination of the membrane restraints and the temporary neutralization of charged residues may be worth consideration. Recent developments on coarse-grained (CG) models of lipids and peptides are also promising for long timescale simulations involving channel peptides embedded in a membrane (e.g., Marrink et al., 2004; Bond and Sansom, 2007). Since CG models have a limited degree of accuracy in the representation of interactions, for example, between charged residues of the peptide and the lipid headgroups, it is always desirable that the results from CG analyses are evaluated with atomistic simulation systems. Another technical limitation is the accuracy of parameters. Given that ion channel activation/deactivation depends on many interactions between amino acid residues and interactions between amino acid and lipids, the accuracy of the parameters used to describe lipids, peptides and their interaction are critical. It should be kept in mind that these parameters are not perfect, but intensive research for improvements is on going (Kandt et al., 2007; Mátyus et al., 2007). Therefore, to draw useful conclusions from simulation results, one usually has to take into account the potential effects of the force field chosen and other simulation parameters used. The good news is that, while the validity of an individual simulation result is often difficult to show, a set of simulations for comparative analyses (variation in mutations, initial configurations, lipid compositions, etc.) often become more effective than one isolated simulation. Our analyses suggested that GsMTx4 and HaTx can interact with lipid membranes in at least two different modes. The two-mode hypothesis explains the biphasic effect by Hurst et al. (2009) well. In the future, many more analyses including mechanosensitive channels themselves are necessary. To this end, structural information of SACs in membranes is crucial. The crystal structure of MscL from Mycobacterium tuberculosis (Tb-MscL) in the closed state (Chang et al., 1998) has become a useful starting point for computational modeling and simulation studies. The crystal structure of E.coli MscS was solved by the Rees group at a resolution of 3.9 A (Bass et al., 2002). This structural information facilitated many modeling and simulation studies (for review, Corry and Martinac (2008a, b)). More recent computational efforts on MscL include Yefimov et al. (2008) and Jeon and Voth

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(2008), and those on MscS include Anishkin et al. (2008a) and Anishkin et al. (2008b). Recently, Yefimov et al. (2008) used the CG model of protein and lipid (DOPE). In the study, starting from the closed crystal structure, considerable opening of the channel was observed close to the rupture tension of the bilayer. This demonstrates the potential usefulness of the CG approach, but it should be noted that the CG representation introduces artificial smoothness into the channel gating. In fact, in the atomistic simulations by Jeon and Voth (2008), the membrane tension and/or the structural asymmetry (i.e., due to the difference in the number of lipid molecules between two leaflets) of the membrane was rather ineffective in inducing MscL structural changes within 20 ns. Recent determination of the structure of a truncated MscL from Staphylococcus aureus is illuminating because it likely corresponds to the ‘pre-expanded’ state based on the tilted transmembrane helices and overall flattening of the channel (Liu et al., 2009). This flattening is accompanied by an increase in the cross sectional area of the MscL channel. Flattening accompanying channel opening has also been pointed out in several simulation studies (e.g., Anishkin et al. 2008a, b). These studies should deepen our understanding of the gating mechanism by membrane stress, and also provide opportunities for simulation studies of the tertiary complex ICK peptide/channel/lipid membrane. In future studies, lipid composition should also be varied, since different compositions will likely alter the relative stability of the two binding modes. It is possible that the presence of ICK peptides influences the formation of microdomains containing acyl chains with a distinct degree of saturation. Besides acyl chains, lipid headgroup structure/composition may also be influenced by ICK peptides. Such considerations are interesting because the phospho-headgroups of membrane lipids, together with certain acidic channel residues, are believed to provide the necessary counter-charges for the positively charged residues of S4 during individual steps of the voltage-sensor movement (Xu et al., 2008; Ramu et al., 2006). Therefore, some ICK peptides may have direct effects on lipid headgroups, which then modify the gating property of channels. Besides such potential effects of ICK peptides on channel-lipid interaction, the effect of the peptides on the lateral pressure profile of the membrane could exerts indirect effects on the gating of mechanosensitive channels. The binding of GsMTx4 to the membrane may change the lateral pressure profile. For the pure bilayers of POPE or POPC, the layer of the lipid headgroups has a ‘tension trough’ that has a weak lateral pressure (Lindahl and Edholm, 2000; Gullingsrud and Schulten, 2004). The presence of GsMTx4 in the shallow mode may nullify the tension trough. The GsMTx4 transition to the deep mode may further alter the pressure profile, thereby changing the free energy required to open the gate. Clearly, many more analyses are necessary to discuss this issue quantitatively. To improve future simulation studies of ICK peptides, the importance of comparing simulations starting from various positions/orientations is again emphasized. In particular, when studying the deep binding mode, it is important to take into account that various combinations of two or three charged residues may interact with the inner leaflet of the membrane to form the deep mode. Therefore, reorientation may be necessary until the peptide finds the orientation/position with the lowest free

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energy. Although, for both experiments and simulations, the reorientation may be a time-consuming step because interaction between the charged residues and lipid headgroup has to be remodeled. Notwithstanding such difficulties, homology modeling and MD simulations provide useful measures to study many themes of mechanosensitive channels. In many channel studies, it is difficult to say that MD simulations can stand alone as a method, yet they further the understanding of how ion channels work in atomistic details. At the same time, ICK peptide analyses suggest that understanding the mechanics of mechanosensitive channels at the atomic level will help us design molecules that can modify gating dynamics in a channel-specific manner. On the other hand, it can be anticipated that many future MD simulation studies suffer from discrepancies between the simulation results and experimental findings. As is the case with large membrane proteins in general, careful consideration is necessary when applying computational methods to ion channels; for example, how close is the simulation system to reality, or how robust is the result? As with the experimental approaches, deliberate designing of a series of simulations should always be helpful.

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Part III

Mechanosensitivity and Mechanotransduction in Vascular Cells

Chapter 8

Cellular and Molecular Effects of Mechanical Stretch on Vascular Cells Kou-Gi Shyu

Abstract The vascular endothelium is a dynamic cellular interface between the vessel wall and the blood stream. It plays an important role by sensing the alterations in biological, chemical, and physical properties of blood flow to maintain homeostasis. Cells in the cardiovascular system are permanently subjected to mechanical forces due to pulsatile nature of blood flow and shear stress, created by the beating hearts. These haemodynamic forces play an important role in the regulation of vascular development, remodeling, wound healing, atherosclerotic lesion formation, and endothelial progenitor cell function. Mechanical stretch can modulate cell alignment and differentiation, migration, survival or apoptosis, vascular remodeling, and autocrine and paracrine functions in smooth muscle cells. Laminar shear stress exerts anti-apoptotic, anti-atherosclerotic, and anti-thrombotic effects on endothelial cells. However, low shear stress or high laminar shear stress exerts atherogenic effect on endothelial cells. Knowledge of the impact of mechanical stretch on the cardiovascular system is vital to the understanding of pathogenesis of cardiovascular diseases and is also crucial to provide new insights in the prevention and therapy of cardiovascular diseases. Keywords Mechanical stretch · Shear stress · Smooth muscle cell · Endothelial cell · Cardiac myocyte

8.1 Introduction Cells in the cardiovascular system are permanently subjected to mechanical forces due to pulsatile nature of blood flow and shear stress, created by the beating heart. These haemodynamic forces play an important role in the regulation of vascular development, remodeling, wound healing, and atherosclerotic lesion formation, and K.-G. Shyu (B) Division of Cardiology, Shin Kong Wu Ho-Su Memorial Hospital, Taipei, Taiwan; Graduate Institute of Clinical Medicine, College of Medicine, Taipei Medical University, Taipei, Taiwan e-mail: [email protected] A. Kamkin, I. Kiseleva (eds.), Mechanosensitivity and Mechanotransduction, Mechanosensitivity in Cells and Tissues 4, DOI 10.1007/978-90-481-9881-8_8, C Springer Science+Business Media B.V. 2011

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endothelial progenitor cell function. For the cardiovascular systems, endothelial cells (ECs) and smooth muscle cells (SMCs) are the major cells that face mechanical forces. Blood pressure is the major determinant of vessel stretch. Vascular smooth muscle cells (VSMCs) are a main cellular component of the blood vessel wall. They are subjected to a dynamic mechanical environment modulated by pulsatile pressure and oscillatory shear forces. VSMCs are primarily subjected to the cyclic stretch resulting from pulsatile changes in blood pressure. ECs are mainly subjected to shear stress by the flowing blood. Knowledge of the impact of mechanical stretch on the cardiovascular system is vital for the understanding of pathogenesis of cardiovascular diseases and can provide new insights in the development of therapeutic strategies of cardiovascular diseases. In the past years, several review articles have been published discussing the molecular mechanisms of mechanical stretch on ECs and VSMCs (Kakisis et al., 2004; Gunningham and Gotlieb, 2005; Haga et al., 2007; Lehoux, 2006; Lehoux et al., 2006; Li and Xu, 2007; Orr and Helmke, 2006; Riha et al., 2005; Shyu, 2009). In this book chapter, I summarize the recent findings about the cellular and molecular effect of mechanical stretch on vascular cells, including ECs and VSMCs. Because several types of devices have been applied to induce mechanical stretch in vitro (Brown, 2000; ReinhartKing et al., 2008), the cellular and molecular responses in each type of vascular cells may be different. Most of the effects of shear stress on the ECs are beneficial (including anti-apoptotic, anti-atherosclerotic, and anti-thrombotic effects) because atherosclerosis preferentially occurs in areas of disturbed flow or low shear stress, whereas regions with steady laminar flow and physiological shear stress are protective. However, the effect of mechanical stretch on SMCs may be beneficial or detrimental. Although various models of mechanical stimuli (static or dynamic) have been used in the past, the majority of research used the Flexercell Stress Unit (Flexercell Corp., USA). This model, a 2-dimensional cell culture, is controlled by a computer program and provides a physiological representation of the in situ environment of repetitive mechanical stimuli. However, this model is a poor representation of natural tissue environment of vascular cells, which is 3-dimensional, mechanically dynamic, and involves the interaction of multiple cell types (Nikolovski et al., 2003). Organ culture model for study the effect of mechanical stretch of vascular cells is the best in vitro representation of the vessel in its in vivo environment, where multiple cell types and the extracellular matrix participate in response to mechanical stimuli (Lehoux et al., 2000). Regardless of the 2- or 3-dimensional model used, in vitro studies do not allow easy distinction between stretch effects due to transmembrane force transfer and stretch effects due to a global change in cell morphology which causes generalized deformation of the plasma membrane and the cytoskeleton (Wang et al., 1993).

8.2 Effect of Mechanical Stress on Endothelial Cells The vascular endothelium is a dynamic cellular interface between the vessel wall and the blood stream. It plays an important role by sensing the alterations in biological, chemical, and physical properties of blood flow to maintain homeostasis.

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In addition to interacting with blood constituents and circulating cells, the vascular endothelium is exposed to a distinct mechanical environment consisting of haemodynamic shear stress and mechanical stretch from blood pressure. Disturbance of normal haemodynamic load can contribute to cardiovascular diseases including hypertension, intimal hyperplasia, vascular restenosis, and atherosclerosis (Cummins et al., 2007). Laminar shear stress, the frictional force created by the flowing blood, exerts a variety of cellular and molecular effects on endothelial structure and functions. The molecular mechanism of shear stress on ECs has been extensively reviewed (Cummins et al., 2007; Kakisis et al., 2004; Lehoux, 2006; Lehoux et al., 2006; Li et al., 2005) In addition to anti-apoptotic and antiatherosclerotic effects of laminar shear stress on ECs, laminar shear stress has a profound impact on endothelial metabolism and can alter gene expression leading to changes in the endothelial phenotype and vessel wall homeostasis. The genes regulated by shear stress can modulate several endothelial functions including vessel diameter, cell proliferation, migration and angiogenesis, cell-cell communication, coagulation and fibrinolysis, anti-inflammation, and immune modulation (Cummins et al., 2007; Kakisis et al., 2004; Lehoux, 2006; Li et al., 2005). In this book chapter, additional novel findings about the impact of shear stress on ECs are discussed.

8.2.1 Effect of Shear Stress on ECs Protein Alteration DNA mircoarrays have been used to analyze a large number of genes in ECs exposed to shear stress (Wasserman et al., 2002; Andersson et al., 2005). Proteomic analysis shows that a broad spectrum of proteins is altered by shear stress. Wang et al. found 142, 213, and 186 candidate proteins up- or down-regulated at least two-fold after 10 minutes, 3 hour, and 6 hour of shear stress, respectively (Wang et al., 2007). These proteins include transcriptional regulators, enzymes, protein kinases, G-protein-coupled receptors, cytokines, protein degradation related proteins, cytoskeletal and matrix proteins. These findings suggest that shear stress has profound effects on the molecular response and the physiological function of the vascular endothelium.

8.2.2 Vasculoprotective Effect of Shear Stress on ECs The vasculoprotective effects of shear stress on ECs have been documented more than one decade ago (Traub and Berk, 1998). Novel findings extending this aspect have been reported recently. Bone morphogenetic protein (BMP) is a TGF-β family member cytokine that exerts proinflammatory effect on the endothelium and plays a role in atherogenesis. BMP is upregulated at “athero-prone” regions in blood vessels and may contribute to vascular calcification and development of atherosclerotic plaques (Csiszar et al., 2006). Csiszar et al. have reported that laminar shear stress activates cAMP and cAMP-dependent protein kinase and downregulates the BMP-4 expression in coronary artery ECs (Csiszar et al., 2007). This finding supports the

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antiatherogenic and vasculoprotective effect of shear stress because BMP-4 elicits endothelial activation, dysfunction, hypertension, and vascular calcification. The transcription factor Kruppel-like factor-2 (KLF2) is an important mediator of the anti-inflammatory and anti-thrombotic properties of the endothelium (Dekker et al., 2005; Lin et al., 2005). Prolonged shear stress stabilizes the KLF2 mRNA and induces KLF2 protein expression, especially in the presence of pro-inflammatory cytokine TNF-α stimulation (van Thienen et al., 2006). The atheroprotective effect of prolonged shear stress is superior to statin, a lipid lowering agents, in the presence of TNF-α in endothelial cell culture model. This finding also supports the atheroprotective effect of prolonged shear stress on ECs. Vascular injury and atherogenesis can be induced by complement activation. The complement inhibitory protein CD59 can be upregulated by shear stress through KLF2 activation (Kinderlerer et al., 2008) in venous and aortic ECs, indicating a vascular protection by shear stress in complement-mediated injury. Laminar shear stress also activates nuclear factorerythroid 2-related factor 2 (Nrf2) which regulates redox levels by activation of numerous anti-oxidant genes including heme oxygenase-1 and ferritin (Ridger et al., 2008). Both KLF2 and nrf2 are candidate regulators of the anti-inflammatory effects of laminar shear stress. Elevated shear stress has been shown to enhance classical pathway complement activation on vascular ECs in vitro (Yin et al., 2007). Shear stress actually can enhance C4 deposition on ECs surface (Yin et al., 2008). Regulation of complement activation at the cell surface by complement regulatory proteins may modulate the inflammatory response and attendant pathological sequelae. Stearoly-CoA desaturase-l (SCD1) is a rate-limiting enzyme in the biosynthesis of monounsaturated fatty acids. SCD1 converts palmitate and stearate into palmitoleate and oleate by catalyzing the 9-cis desaturation of saturated fatty acids (Ntambi and Miyazaki, 2004). Palmitoleate and oleate are the predominant unsaturated fatty acids in membrane phospholipids. Qin et al. recently have reported that shear stress increases SCD1 expression in human vascular EC through a peroxisome proliferator-activated receptor-γ (PPARγ) mechanism (Qin et al., 2007). The metabolic effect of shear stress provides another evidence for its atheroprotective effect. The liver X receptors participates in cholesterol transport and lipid metabolism causing atheroprotective effect. Laminar shear stress increases liver X receptor function via a PAPARγ-sterol 27-hydroxylase dependent mechanism (Zhu et al., 2008), further supporting the atheroprotective role of laminar shear stress in ECs. Li et al. provide additional evidence for the atheroprotective effect of shear stress in ECs by demonstrating that shear stress decreases TNF-α-mediated vascular adhesion molecule-1 by inhibiting JNK through MEK5-BMK1 signaling pathways (Li et al., 2008). They found that flow inhibits TNF-α-mediated signaling events in ECs by a mechanism dependent on activation of MEK5-BMK1, but not MEK1-ERK1/2. Heme oxygenase-1 catalyzes the degradation of heme to liberate free iron, carbon monoxide and biliverdin in mammalian cells. Heme oxygenase-1 induction represents a cytoprotective defense mechanism against oxidative insults. Recently, Di Francesco et al. reported that laminar shear stress up-regulates heme oxygenase 1

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Fig. 8.1 Schematic summary of vasculoprotective effects of laminar shear stress on endothelial cells. A diagram summarizing the laminar shear stress-induced mechanosensing and intracellular signaling that lead to the modulation of gene expression and cellular function, resulting in vasculoprotection, as discussed in the text

by cyclooxygenase2-dependent prostacyclin induction in human umbilical vein ECs (Di Francesco et al., 2009). The up-regulating heme oxygenase-1 attenuates TNF-α generation. The cyclooxgenase2 and heme oxygenase-1 are the vasoprotective genes up-regulated by steady laminar shear stress, which characterized “atherosclerotic lesion-protected areas”. The mechanism and signal pathways for vasculoprotective effects of laminar shear stress on endothelial cells are summarized in Fig. 8.1.

8.2.3 Effect of Shear Stress on ECs Polarity and Morphology Mechanical forces regulate ECs polarity and directional migration. Mechanical stress also affects ECs morphology and function (Chien, 2007). This is important for vascular function, remodeling and wound repair (Chien et al., 2005). Shear stress can modulate microtubule-organizing centers polarity and microtubule stability in vitro and in vivo by activation of glycogen synthase 3β signaling pathway (McCue et al., 2006). The application of shear stress causes of EC elongation in the direction of flow. Shear stress induces reorientation of the microtubule organizing center to the leading edge of migrating cells in a cdc42-dependent manner (Simmers et al., 2007). Vascular ECs respond to laminar shear stress by aligning in the direction of flow. Goldfinger et al. have reported that localized α4 integrin phosphorylation leads localized Rac1 activation and subsequent stress fiber alignment and ECs elongation parallel to the flow direction in response to shear stress (Goldfinger et al., 2008). The shear-induced α4 integrin phosphorylation is dependent on cAMP-dependent protein kinase A. The α4 integrin and cAMP-dependent protein kinase A regulate

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vascular ECs adhesion, migration and survival, and angiogenesis. The shear-induced cAMP-dependent protein kinase A-dependent α4 integrin phosphorylation may be an important regulatory step in endothelial function during vascular development and remodeling (Goldfinger et al., 2008). The reorientation of ECs after cyclic strain has been related to activation of transient receptor potential vanilloid 4 (TRPV4) ion channels (Thodeti et al., 2009). TRPV4 is one of the stretch-activated channels. Cyclic stretch of capillary ECs activates mechanosensitive TRPV4 ion channels that stimulate PI3K-dependent activation and binding of additional β1 integrin receptors, which promotes cytoskeletal remodeling and cell reorientation. Simmers et al. have also demonstrated that shear stress-induced directed migratory polarity is modulated by exogenous growth factors and dependent on Par6 activity, a major downstream effector of Cdc-42-induced polarity, and shear stress direction (Simmers et al., 2007). Shear stress regulates ECs bulk migratory characteristics as well as morphology. Lee et al. reported adaptation of ECs shape in arteries under axial stretch using an organ culture model. They demonstrated that ECs were initially elongated by the axial stretch but eventually adapted to the axial stretch, regaining their normal shape (Lee et al., 2008). The capillary ECs reorientation after mechanical stress is important for angiogenesis. These data indicate that mechanical stress regulates ECs bulk migratory characteristics as well as morphology. These mechanical sensitive ion channels may represent new targets for therapeutic intervention in angiogenesis-dependent disease.

8.2.4 Anti-Inflammatory and Anti-Oxidant Effect of Shear Stress on ECs Endothelial inflammation is a major initiator of atherosclerosis. ECs exposed to disturbed flow experience oxidative stress, increased expression of markers of inflammation, and monocyte recruitment as early signs of atherosclerosis (Harrison et al., 2006). Anti-inflammatory and anti-oxidant defenses are critical for the protection of cellular macromolecules and progression of atherosclerosis. Laminar shear stress upregulates peroxiredoxins as important antioxidants in ECs (Mowbray et al., 2008). Laminar shear stress increases ERK5 and PPARγ transcriptional activity and decreases adhesion molecule expression in ECs. The laminar shear stress increases eNOS expression through ERK5 to inhibit the formation of ROS (Woo et al., 2008). Shear stress also activates AMP-activated protein kinase in ECs, which contributes to elevated eNOS activity and subsequent NO production (Zhang et al., 2006). AMP-activated protein kinase α is the primary kinase phosphorylating eNOS Ser633/635, which is functionally linked to NO bioavailability (Chen et al., 2009). Shear stress also activates phosphatidylinositol 3 (PI-3) kinase to increase eNOS expression. Sud et al. recently reported a key role for decrease in protein kinase Cδ (PKCδ) signaling in the up-regulation of eNOS in response to shear stress was also due to inhibition of phosphorylation of STAT3 (Sud et al., 2009). The decrease in PKCδ caused by shear stress was mediated by a decrease in

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STAT3 binding to the eNOS promoter. Laminar shear stress up-regulates antioxidant genes and activates the transcriptional factor, NF-E2-related factor-2, which is a major transcriptional factor for EC redox homeostasis (Dai et al., 2007). Laminar shear stress alters the functions of NF-κB by inhibiting its capacity to induce proinflammatory molecules such as VCAM-1, e-selectin, interleukin-8, and ICAM-1 and simultaneously enhancing the induction of NF-κB-dependent cytoprotective transcripts such as manganese superoxide dismutase, Bcl-2 and A1 (Patridge et al., 2007). High glucose and arachidonic acid synergistically decrease cell viability and increase glutathione oxidation and lipid peroxidation in human umbilical ECs, while laminar shear stress attenuates the oxidative stress induced by high glucose and arachidonic acid (Mun et al., 2008). The antioxidant effect of laminar shear stress could be attributed to increased biosynthesis of tetrahydrobiopterin and glutathione. These findings strongly support the crucial role of laminar shear stress as anti-inflammatory and anti-oxidative force. The mechanism and signal pathways for anti-inflammatory and anti-oxidative effects of laminar shear stress on endothelial cells are summarized in Fig. 8.2. High wall shear stress haemodynamics mediate adaptive outward remodeling and cerebral aneurysm development. High wall shear stress stimulates ECs proliferation and suppresses apoptosis (Metaxa et al., 2008). The stimulation of EC proliferation by high wall shear stress is dependent on NO signaling pathways. The mechanism by which ECs sense high wall shear stress is through stretch-activated calcium channels on the luminal surface of the endothelium (Brakemeier et al., 2002). High wall shear stress by stimulation of ECs proliferation plays an important role in maintaining continuity of the endothelium in regions where the vessel wall is subjected to high levels of haemodynamic stress.

Fig. 8.2 Schematic summary of anti-inflammatory and anti-oxidative effects of laminar shear stress on endothelial cells. A diagram summarizing the laminar shear stress-induced mechanosensing and intracellular signaling that lead to the modulation of gene expression and cellular function, resulting in anti-inflammatory and anti-oxidative stress, as discussed in the text

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8.2.5 Effect of Disturbed Flow on ECs While laminar shear stress plays an atheroprotective role on ECs, cyclic stretch or oscillatory shear stress induces different cellular responses (Fig. 8.3). Pulsatile flow is steady and laminar in the straight part of vessels, whereas disturbed flow is not steady with large oscillation near bifurcations and curvatures. Disturbed flow stimulates the pro-inflammatory transcription factor NF-κB through integrinand Rac-dependent production of ROS. Cyclic stretch is the repetitive mechanical deformation of the vascular cells as it rhythmically distends and relaxes with the cardiac cycle. These biomechanical forces promote atherosclerosis by increasing formation of reactive oxygen species in ECs and by upregulating pro-atherogenic cytokine expression. We have reported that cyclic stretch augments TNF-α production and matrix metalloproteinase expression in human umbilical vein ECs (Wang et al., 2003). CD40 is a co-stimulatory molecule playing an important role in controlling inflammatory responses, including atherosclerosis. Recently, Korff et al. have demonstrated that cyclic stretch increases the abundance of CD40 in ECs co-cultured with VSMCs through TGF-β1/activin-receptor-loke kinase-1 (ALK1) signaling, whereas EC CD40 abundance is down-regulated by exposure to cyclic stretch in ECs alone (Korff et al., 2007). Cyclic stretch also activates Akt, glycogen synthase kinase (GSK)-3 to enhance survival of ECs (Nishimura et al., 2006). Akt is important in preventing apoptosis but is not involved in EC proliferation. These findings indicate that haemodynamic forces present in atherosclerosis-resistant and -susceptible region of the vasculature induce different responses in the vessel wall.

Fig. 8.3 Schematic summary of atherogenic effects of disturbed flow on endothelial cells. A diagram summarizing the disturbed flow-induced mechanosensing and intracellular signaling that lead to atherogenesis, as discussed in the text

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Atherosclerotic lesions frequently develop in areas of the vasculature exposed to disturbed flow, whereas areas that experience pulsatile laminar flow are relatively protected from plaque formation. Actually, prolonged high laminar shear stress suppress endothelial tissue-type plasminogen activator expression through downregulation of JNK pathway (Ulfhammer et al., 2009) and may contribute to the enhanced risk of arterial thrombosis in hypertensive disease. Therefore, high laminar shear stress is atherogenic on endothelial cells. The role of JNK by mechanical stress in ECs is controversial. Li et al. reported that laminar flow inhibits JNK activation in ECs by inflammatory cytokines such as TNF-α (Li et al., 2008). Inhibition of JNK is mediated through MEK5 and ERK5/BMK1. Recently, Hahn et al. reported that JNK activation by laminar flow and long-term oscillatory flow is matrix-specific, with enhanced activation on fibronectin compared to basement protein or collagen (Hahn et al., 2009). The JNK activation is mediated by MKK4 and p21-activated kinase. The JNK activation is also seen in vivo at atheroprone regions of arteries coincident with fibronectin in the subendothelial matrix. P21activated kinase acts as a critical upstream mediator of matrix-specific NF-κB activation by disturbed flow (Orr et al., 2008). Low shear stress has been shown to induce interleukin-8 gene expression through ERK1/2, JNK1/2 and p38 MAP kinase, which triggers a cascade of events that lead to inflammatory events, which in turn contribute to the initialization of atherosclerosis (Cheng et al., 2008). More recently, Thomas et al. reported that human EC exposed to an atherosclerosis-prone flow pattern, as in vascular regions susceptible to the development of atherosclerosis, exhibited a significant increase in PDGF-DD expression (Thomas et al., 2009). PDGF-DD inhibits expression of multiple SMCs genes, including SM α-actin and SM myosin heavy chain, and up-regulated expression of the potent SMC differentiation repressor gene Kruppel-like factor-4 at the mRNA and protein levels. These findings establish a link between low-oscillatory shear stress blood flow, EC-mediated PDGF-DD expression, and SMCs phenotype modulation. Circulating leukocytes are recruited into the tissue mainly in small vessels such as capillaries and venules. Glutamate-leucine-arginine (ELR) tripeptide motif plays an important role in leukocyte trafficking into the tissues. ELR expression is higher in microvascular endothelium than in aortic ECs (Shaik et al., 2009). Low intensity shear stress at 4 dynes/cm2 activated endothelial ELR chemokine production via cell surface heparin sulfates, β3 -integrins, focal adhesion kinase, MAPK p38β, mitogenand stress-associated protein kinase-1 and NF-κB. The preferential activation of endothelial chemokine expression by low shear is consistent with the concentration of endothelial-leukocyte interaction in capillaries and post-capillary venules. These chemokines may provide a mechanism for the high neutrophil concentrations seen in the capillaries in the normal state. Low shear stress increases the expression of adhesion molecules including VACM-1 and ICAM-1 which engage integrins expressed by activated leukocytes thus reducing the rolling speed of inflammatory cells over the endothelium (Ridger et al., 2008). The mechanism and signal pathways for atherogenic effects of low shear stress on endothelial cells are summarized in Fig. 8.4.

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Fig. 8.4 Schematic summary of atherogenic effects of low shear stress on endothelial cells. A diagram summarizing the low shear stress-induced mechanosensing and intracellular signaling that lead to the modulation of gene expression and cellular function, resulting in atherogenesis, as discussed in the text

8.2.6 Effect of Shear Stress on Endothelial Progenitor Cells Endothelial progenitor cells (EPCs), mobilized from bone marrow, contribute postnatal neovascularization. EPCs are exposed to shear stress generated by flowing blood and tissue fluid flow during the process of EPC incorporation into tissues and neovascularization (Obi et al., 2009). EPCs appear to be responsible to shear stress and the vasculogenic activities of EPCs may be modulated by shear stress. The effect of shear stress on progenitor cell fate has been reviewed by Stolberg and McCloskey (Stolberg and McCloskey, 2009). Shear stress can accelerate the proliferation, differentiation, and capillary-like tube formation of EPCs (Yamamoto et al., 2003). The endothelial differentiation of EPCs by shear stress has been demonstrated by activation of Akt (Ye et al., 2008) and by increasing ephrinB2 expression and Sp1 activation (Obi et al., 2009). Shear stress increases the gene expression of arterial endothelial markers in EPCs, such as ephrin B2, Notch1/3, Hey1/2 and ALK1 but decreases the gene expression of venous endothelial markers such as EphB4 and NRP2 (Obi et al., 2009). Shear stress within physiological range can enhance tissue type plasminogen activator and prostaglandin I2 secretion (Yang et al., 2007) and decrease plasminogen activator inhibitor-1 (Yang et al., 2006) in human EPCs, which improves the antithrombogenic potential of human EPCs. Shear stress has also been demonstrated to increase Cu/Zn superoxide dismutase activity and NO production, as well as superoxide dismutase and eNOS mRNA expression (Tao

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et al., 2007). Shear stress also induces endothelial differentiation in a mouse mesenchymal progenitor cell line (Wang et al., 2005). Wang et al. reported that TGF-β1 was down-regulated in mesenchymal progenitor cell line by shear stress (Wang et al., 2008). The negative regulation of the TGF-β1 system may be involved in shear-induced endothelial cell differentiation in the mouse mesenchymal progenitor cell line.

8.3 Effect of Mechanical Stretch on VSMC Function Mechanical stretch can modulate several different cellular functions in VSMCs. These functions include, but are not limited to cell alignment and differentiation, migration, survival or apoptosis, vascular remodeling, and autocrine and paracrine functions. However, different kinds of VSMCs (venous or arterial type) and several species of animals (mouse, rat, rabbit, swine, and others) were used in different studies, resulting in sometimes controversial findings. Most of the studies used in vitro models. The cellular functions induced by in vitro mechanical stretch may not really represent the in vivo cellular function. Further and detailed studies are needed to elucidate the real effect and mechanisms of mechanical stress on VSMC functions.

8.3.1 Effect of Mechanical Stretch on VSMC Alignment and Differentiation Arterial SMCs are aligned primarily in the circumferential direction in the media of artery. The mechanical stretch from pulsatile blood flow is one of the key factors in regulating vascular remodeling (Taber, 1998). Mechanical environment in vivo modulates the distinct patterns of VSMC orientation in the arterial wall. The predominant mechanical force influencing VSMCs structural organization and signaling is cyclic stretch (Halka et al., 2008). There are at least three elements included in the cyclic stretch: magnitude, frequency, and duration. Cultured VSMCs in vitro can be induced to reorient to a uniform alignment almost perpendicular to stretch vector alignment by mechanical stretch (Lehoux et al., 2000; Standly et al., 2002). The response of cell reorientation depends on the stretching magnitude and frequency (Dartsch et al., 1986; Liu et al., 2008; Wang et al., 1995). The signaling pathways involved in the stretch-induced VSMCs alignment include p38 mitogen-activated protein (MAP) kinase (Liu et al., 2008), nitric oxide and reactive oxygen species (Chen et al., 2003; Standly et al., 2002). The mechanosensor and outside-in signal of integrin-β1 is also involved in the stretch-induced VSMCs alignment (Chen et al., 2003). An intact cyctoskeleton is important for the stretch-induced VSMCs alignment. Destroying the actin filament system by cytochalasin D inhibits the effect of stretch-induced alignment (Liu et al., 2008). H1-calponin, a family of actin-associated protein, is a specifically differentiated

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marker in SMCs. Cyclic strain can upregulate the expression of Rac and downregulate its negative regulator Rho-GD dissociation inhibitor alpha in a nonlinear frequency-dependent pattern, then cause the activation of p38 pathway followed by increasing expression of h1-calponon, which marked VSMC differentiation (Qu et al., 2008). Rho-GD dissociation inhibitor alpha, a member of Rho-GD dissociation inhibitor, can negatively regulate the activities of small G proteins of the Rho family by shutting off their GDP/GTP cycling and cytosol/membrane translocation. Not only VSMCs alignment is affected by stretch frequency, but also the phenotype of VSMCs. Cyclic strain increases smooth muscle and decreases nonmuscle myosin expression in VSMCs (Reusch et al., 1996). Mechanical stretch increases both smooth muscle α-actin protein expression and promoter activity (Tock et al., 2003). The induction of smooth muscle α-actin is mediated by activation of JNK and p38 MAP kinase pathways. Mechanical stretch could promote a frequency-dependent redifferentiation of synthetic VSMCs in vitro, mediated at least in part by the activation of p38 MAP kinase (Qu et al., 2007). Mechanical stretch modulates cell shape, cytoplasmic organization and intracellular processes leading to migration, proliferation, or contraction. Rho and intact actin filaments play an important role in mechanical stretch-induced extracellular signal-regulated kinase (ERK) activation and cell growth (Numaguchi et al., 1999). RhoA signaling plays a major role in the serum responsive factor (SRF)-dependent regulation of SMC differentiation marker gene expression (Mack et al., 2001). The primary genes encoding SMCs contractile proteins are regulated by the stretch-induced RhoA pathway and associated transcription factors, most importantly the SRF (Hellstrand and Albinsson, 2005). In vitro, RhoA enhances actin polymerization and stimulates the SRF homodimer binding to their CArG boxes (Mack et al., 2001). SRF binds to the serum response element region containing the 10-bp CArG-box sequence facilitating activation of this motif alone or as a macromolecule bound to myocardin, its specific co-activator (Halka et al., 2008). Myocardin increases the promoter activity of the CArG-dependent VSMCs contractile markers. Stretch of the vascular wall can stimulate increased actin polymerization, activating synthesis of smooth muscle-specific proteins via Rho-associated kinase and cofilin downstream of Rho (Albinsson et al., 2004). Rho/Rho kinase, p44/p42 MAP kinase and phosphatidylinositol-3 (PI-3) kinase pathways are all involved in the stretch-induced human saphenous vein SMC proliferation and inhibition of either of them prevents stretch-induced SMCs proliferation (Kozai et al., 2005). The effect of mechanical stretch on SMCs phenotype has been reviewed by Halka et al. (Halka et al., 2008).

8.3.2 Effect of Mechanical Stretch on VSMCs Migration VSMCs migration is important in the development of vascular diseases including atherosclerosis and post angioplasty restenosis. VSMCs migration is found more frequently in curved and bifurcating blood vessels, which are exposed to nonlaminar blood flow, than in straight arterial segments exposed to laminar blood flow

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(Liu, 1999). In a vein graft model of SMC study, vortex blood flow induces VSMCs migration and neointimal hyperplasia in control vein graft, whereas reduction of vortex blood flow in the vein graft strongly suppresses the migration and hyperplasia (Goldman et al., 2007). In this model, VSMCs migration is regulated through the mediation of ERK1/2 and myosin light chain kinase. In vitro, mechanical stretch of arterial VSMCs translocates PKCδ from membrane to cytoskeleton and increases the migration of VSMCs (Li et al., 2003). In our laboratory, we could also demonstrate that mechanical stretch increased migration of VSMCs (Shyu et al., 2005). The increased migration of VSMCs by mechanical stretch involves p38 MAP kinase and transforming growth factor-β1 (TGF-β1) (Li et al., 2003). Inhibition of p38 MAP kianse and TGF-β1 activity decreased the migration activity. Qi et al. used proteomic analysis to demonstrate that low shear stress-induced VSMC migration and apoptosis are mediated by a down-regulation of Rho-GDP dissociation inhibitor alpha (Qi et al., 2008). The effect of Rho-GDP dissociation inhibitor alpha on VSMCs migration is dependent on the PI3K/Akt signal transduction pathway.

8.3.3 Effect of Mechanical Stretch on Proliferation, Survival and Apoptosis of VSMCs The effect of mechanical stretch on survival and apoptosis of VSMCs has been extensively reviewed by Kakisis et al. (2004) and Haga et al. (2007). In this chapter, I would like to extend these studies by data published recently. In a mouse SMCs cultured model, Cheng et al. have reported that mechanical stretch prevents apoptosis of VSMCs in response to oxidized low-density lipoprotein (Cheng et al., 2007). The mechanism of increased survival of VSMCs induced by mechanical stretch includes αVβ3 integrin expression, stabilization of PINCH-1, a survival protein that is linked with integrin and the cytoskeleton (Xu et al., 2005), and remodeling of cytoskeleton. Small interfering RNA (siRNA) against integrin β3 as well as VSMC isolated from integrin β3 knockout mice abolishes the antiapoptotic effect of mechanical stretch. Down-regulation of PINCH-1 by siRNA enhanced the ability of oxidized low density lioporotein to cause apoptosis of VSMCs, while up-regulation of integrin β3 stabilizes PINCH-1 and protects VSMCs from apoptosis. Disruption of cytoskeleton also abolishes the antiapoptotic effect of stretch. In cultured rabbit VSMCs, mechanical stretch also stimulates SMCs growth and hypertrophy (Richard et al., 2007). The increased SMCs survival by mechanical stretch is induced by nuclear protein import and nuclear pore protein expression that is mediated via MAP kinase. In cultured bovine pulmonary artery VSMCs, mechanical stretch stimulates proliferation of VSMCs and RhoA is essential for stretch-induced VSMCs proliferation (Liu et al., 2007). Blocking Rho completely inhibits the proliferation of VSMCs induced by stretch. In cultured vein VSMC, mechanical stretch stimulates proliferation of venous SMCs (Cheng and Du, 2007). The proliferation of venous SMCs induced by mechanical stretch is mediated by activation of insulin-like growth factor-1 (IGF-1) and IGF-1 receptor. When IGF-1 receptor is knocked out, the mechanical

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stretch-induced increase in VSMCs proliferation is blocked. IGF-1 receptor level is increased in neointima in vein grafts and IGF-1 receptor deletion reduces neointima formation in vein grafts. Cysteine- and glycine-rich proteins (CRP) regulate SMCs proliferation and differentiation and are important for the maintenance of the contractile apparatus of SMC. Campos et al. reported that CRP3/muscle LIMdomain protein is expressed mainly in arteries and can be induced in veins during the arterialization process in vitro and in vivo (Campos et al., 2009). The induction of CRP3/muscle LIM-domain protein is dependent on increased stretch in SMCs, rather than increased shear stress in ECs. Pulsatile mechanical pressure has been shown to provoke proliferation of human aortic SMCs and mechanical pressure initiates an up-regulation of angiotensin-converting enzyme protein, activity and mRNA expression (Iizuka et al., 2008). The mechanism of up-regulating angiotensin-converting enzyme by pulsatile pressure is regulated by ERK-related signal transduction cascades. In a porcine VSMCs cultured model, Su et al. have demonstrated that mechanical stretch-induced VSMCs apoptosis is phenotype dependent (Su et al., 2006). Mechanical stretch induces apoptosis in differentiated VSMCs, but not in proliferating VSMCs. The stretch-induced apoptosis in VSMCs is associated with Bcl2-associated death factor expression. Vascular endothelial growth factor (VEGF) and overexpression of the antiapoptotic protein Bcl-2 decrease Bcl-2-associated death factor expression and apoptosis induced in response to stretch. Recently, we also have demonstrated that mechanical stretch induces apoptosis in VSMCs from rat thoracic aorta (Cheng et al., 2008). The mechanical stretch induced VSMCs apoptosis is load-dependent. In contrast to 10% stretch, stretch of 20% induces apoptosis. Cyclic stretch enhances GADD153 protein and mRNA expression in VSMCs. Stretch-induced GADD153 protein expression in VSMCs is mediated by JNK Cyclic stretch increases AP-1 binding activity. Cyclic stretch increases GADD153 promoter activity through AP-1 (Fig. 8.5). The mechanism of apoptosis induced by mechanical stretch in our study is mediated by GADD153, a component of endoplasmic reticulum stress-mediated apoptosis factor (Oyadomari and Mori, 2004). Caspase 3 is involved in the GADD153-induced apoptosis of VSMCs after mechanical stretch. An in vivo model of aorta-caval shunt also increases aortic GADD153 protein expression. These results indicate that GADD153 plays an important role in stretch-induced VSMCs apoptosis. Fitzgerald et al. also confirm that laminar shear stress stimulates SMCs apoptosis (Fitzgerald et al., 2008). SMCs respond to laminar shear stress with diminished Akt activity that in turn promotes the intrinsic apoptotic pathway. Pulsatile equibiaxial stretch, acting through NO and cGMP, can prevent the ability of thrombin to stimulate Rho signaling pathways that contribute to pathophysiological proliferative and inflammatory responses (Haga et al., 2008). The inhibition of thrombin-induced RhoA activation by stretch suggests that mechanical forces provide an additional level of regulation of SMCs proliferation and inflammatory signaling. Using cells from different species and modifications in intensity and duration of stretch may cause the differences observed in previous studies. Therefore, the controversial effect of mechanical stretch on survival and apoptosis of VSMCs

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Fig. 8.5 Schematic summary of regulation of mechanical stretch on GADD153 expression in VSMCs. The increased GADD153 expression by mechanical stretch induces apoptosis of VSMCs

Fig. 8.6 Schematic summary of various effects of mechanical stretch on VSMCs. A diagram summarizing mechanical stretch-induced mechanosensing and intracellular signaling that lead to the modulation of gene expression and cellular function, resulting in migration, differentiation, proliferation, and survival of VSMCs, as discussed in the text

under mechanical stretch needs further investigation. The mechanism and signal pathways for effects of mechanical stretch on migration, proliferation, and survival in VSMCs are summarized in Fig. 8.6.

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8.3.4 Effect of Mechanical Stretch on Vascular Remodeling Vascular inward remodeling results in decreased lumen size and increased vessel resistance. The signaling cascades that modulate vascular remodeling process in response to mechanical stretch include reactive oxygen species (ROS), nitric oxide (NO), nuclear factor κB (NF-κB), epidermal growth factor receptor (EGFR), MAP kinase and protein kinase C (PKC) (Kouri and Eickelberg, 2006). Mechanical force can be transduced via ROS-dependent autocrine and paracrine EGFR activation and may regulate VSMCs proliferation and synthetic activity through NF-κB pathway (Lemarie et al., 2006). TGF-α is a potential specific target for vascular remodeling induced by mechanical stretch. In vivo, increased hemodynamic forces in a model of hypertension by angiotensin II (AngII) infusion, activation of NF-κB and associated cell proliferation and wall thickening are reduced in TGF-α- mutant mice, compared with wild-type animals. Syndecan-1 and -4 belong to a family of tranasmembrane proteoglycans, acting as co-receptors for growth factor binding as well as cell-matrix and cell-cell interactions, and are induced in neointimal SMCs after balloon injury (Julien et al., 2007a, b). Both syndecan-1 and -4 expression and shedding are upregulated by mechanical stretch (Julien et al., 2007a, b), which may contribute to the vascular pathology induced by mechanical microenvironment in vivo. Recently, Albinsson and Hellstrand have reported that remodeling of SMCs to stretch requires a dynamic cytoskeleton (Albinsson and Hellstrand, 2007). The stabilization of actin filaments is essential for the growth and synthesis of contractile proteins in response to physiological levels of mechanical stretch. Mechanical stretch enhances VEGF and hypoxia-inducible factor-1α (HIF-1α) gene expression through transcriptional regulation in VSMCs (Chang et al., 2003; Shyu et al., 2001). The transient increase in VEGF and HIF-1α gene expression induced by mechanical stretch may be relevant to pathological complications in the cardiovascular system, including atherosclerosis, plaque stability and hypertension. The induction of VEGF and HIF-1α gene by mechanical stretch may play a role in vascular remodeling. Prolonged cyclic strain produced up-regulation of small proline-rich repeat protein (SPRRP3), a protein highly expressed in advanced atheromas of human arteries, RNA and protein in VSMCs (Pyle et al., 2008). SPRR3 regulation by cyclic strain required type I collagen, whereas VSMCs grown on poly-L-lysine or pronectin F failed to regulate SPRR3 with cyclic strain. The α1β1 collagen-binding integrin is required for mechanoregulation of SPRR3. The data presented by Pyle et al. indicated that SPRR3 may play a role in altered biomechanical compliance of SMCs within an atheromatous lesion because SPRR3 is exclusively enriched in VSMC within atheromas in response to mechanical stress. EC can regulate VSMCs proliferation. Heparin and EC soluble heparin sulfate proteoglycans (HSPG) are potent inhibitors of VSMCs proliferation and fibroblast growth factor-2 induced mitogenesis. Baker et al. reported that mechanical strain stimulated the production of perlecan and HSPGs by EC (Baker et al., 2008). HSPGs are a key component in an integrated feedback control loop regulating vascular remodeling through the modulation of paracrine endothelial inhibition of VSMCs growth. ERK and TGF-β signaling were required for pressure-induced

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up-regulation of endothelial HSPG using in vitro and ex vivo studies. Net arterial remodeling to haemodynamic forces is controlled by a dynamic interplay between growth stimulatory signals from VSMCs and growth inhibitory signals from ECs.

8.3.5 Autocrine and Paracrine Effect of Mechanical Stretch on VSMCs Mechanical stretch may induce secretion or synthesis of bioactive molecules from VSMCs. These secreted bioactive molecules can act on neighboring cells or the cells secreting them. The autocrine and paracrine effect of mechanical stretch has been demonstrated recently and these effects regulate individual intracellular signaling pathways, VSMCs growth and initiate the cellular and molecular effect of mechanical stretch on VSMCs. Platelet derived growth factor was initially found to play autocrine function in VSMCs after mechanical stretch (Wilson et al., 1993). Mechanical stretch using the portal vein SMCs induces endothelin-1 release and promotes synthesis of smooth muscle specific proteins by a mechanism requiring an intact cytoskeleton (Zeidan et al., 2003). Mechanical stretch also stimulates autocrine IGF-1 production from arterial and venous VSMCs (Cheng and Du, 2007; Standley et al., 1999). TGF-α has been shown to modulate the NF-κB activation and vascular remodeling under stress (Lemarie et al., 2006) and TNF-α has been reported to modulate the GADD153 expression and VSMCs apoptosis in VSMCs

Fig. 8.7 Schematic summary of different effects of mechanical stretch on VSMCs. A diagram summarizing mechanical stretch-induced mechanosensing and intracellular signaling that lead to the modulation of gene expression and cellular function, resulting in vascular remodeling, autocrine, and paracrine effects in VSMCs, as discussed in the text

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under mechanical stretch (Cheng et al., 2008). The intracellular signaling pathways of autocrine and paracrine effect of TGF-α under mechanical stress involve ROS and NFκB. The intracellular signaling pathways of stretch-induced GADD153 expression involve JNK and AP-1 pathway. AngII and TGF-β1 play autocrine and paracrine action on discoidin domain receptor 2 (DDR2) expressions in VSMC under mechanical stretch (Shyu et al., 2005). The stretch-induced DDR2 is mediated by p38 MAP kinase and Myc-Max pathway. DDR2 can regulate cell proliferation and extracellular matrix remodeling mediated by MMP activities. The intracellular signaling pathways of stretch-induced DDR2 involve p38 MAP kinase and Myc pathway. The mechanism and signaling pathways for effects of mechanical stretch on vascular remodeling, autocrine and paracrine effects in VSMCs are summarized in Fig. 8.7. In an ECs-SMCs co-culture system, SMCs secrete interleukin-1β and interleukin-6 after application of shear stress resulting in inhibition of Eselectin expression (Chiu et al., 2007). In this model, SMC induces endothelial E-selectin expression, while shear stress inhibits the SMC-induced E-selectin expression via the inhibition in SMCs activation of interlukin-1 receptor associated kinase/glycoprotein-130, JNK/p38 MAP kinase, and NF-Kβ (Haga et al., 2007; Lehoux, 2006).

8.4 Conclusions and Perspectives Mechanical stretch activates in ECs, VSMCs, and cardiac myocytes multiple intracellular signaling networks and regulates gene expressions and functional responses. Specific cell types may respond differently to mechanical forces. Different mechanisms of response may be observed by using different duration, load, and frequency of mechanical forces. Although the in vitro mechanical stretch model is assumed to mimic the in vivo haemodynaimc overload, the findings obtained from the in vitro mechanical models must be considered with caution, because the in vivo haemodynamic overload is more complex than the in vitro mechanical stretch model. The cellular and molecular effects of mechanical stretch on vascular cells may provide new insights in the pathogenesis of vascular diseases and therapeutic options. Understanding the molecular mechanisms regulating electrical remodeling under mechanical stretch supports the clinical application of angiotensin converting enzyme inhibitor and ARB in the cardiac protection and in the prevention of atrial fibrillation (Healey et al., 2005). Recently, we have used siRNA technology in the carotid artery balloon injury model to demonstrate the potential therapeutic utility of DDR2 siRNA for prevention of neointimal formation induced by balloon injury (Shyu et al., 2008). This finding supports previous studies indicating that DDR2 increases SMCs migration and proliferation in response to mechanical stretch (Shyu et al., 2005). Therefore, knowledge of the impact of mechanical stretch on the ECs and VSMCs is vital for the understanding of pathogenesis of cardiovascular diseases and crucial to provide new insights in the prevention and therapy of cardiovascular diseases.

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Chapter 9

Role of Proteoglycans in Vascular Mechanotransduction Aaron B. Baker

Abstract Vascular mechanotransduction is the process through which the arterial system adapts to changing hemodynamic and pathophysiological stimuli to maintain homeostasis. This adaptation is made possible by the major cellular components of the artery including vascular smooth muscle cells of the arterial wall and endothelial cells that line the luminal surface. This review focuses on the role of proteoglycans in vascular mechanobiological responses of the arterial system. Proteoglycans are proteins that are post-translationally modified with polysaccharide glycosaminoglycan chains. These molecules are intimately involved in controlling cellular organization, proliferation and migration. In this chapter, we discuss how these complex molecules allow cells to sense mechanical forces and alter arterial structure. Keywords Vascular smooth muscle cells · Endothelial cells · mechanotransduction · Proteoglycans · Heparan sulfate · Syndecans · Vascular remodeling · Shear stress · Mechanical stretch · Integrins · Cytoskeleton

9.1 Introduction The vascular system is under continual exposure to a complex mechanical environment due to variations in blood flow and pressure during the cardiac cycle. Biomechanical signals are potent regulators of the growth, structure and function of the cardiovascular tissues (Dzau and Gibbons, 1993). Blood flow through the arterial conduits leads to cyclical distension of the arterial wall and concomitant shear stress (Humphrey, 2008). Shear stresses are regulators of endothelial nitrous oxide production and, consequently, vasomotor tone (Smiesko et al., 1985). When the mechanics of the vascular system are perturbed by disease processes or changes A.B. Baker (B) Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, 77 Massachusetts Avenue, E25-442, Cambridge, MA 02139, USA e-mail: [email protected]


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in tissue metabolism the system must adapt to seek a new level of homeostasis. For example, hypertension is the most common clinical finding in the United States and is associated with increased risk of stroke, atherosclerosis and myocardial infarction (Whisnant, 1996; Rosendorff, 2007). While the fundamental mechanisms causing essential hypertension remain elusive, the consequences to the vascular system are profound and include arterial stiffening and vascular smooth muscle cell hypertrophy (van den Akker et al., 2009). Fundamental to the processes vascular adaptation are the mechanosensing mechanisms of the vascular cells themselves. Numerous in-vitro studies have been done examining vascular cell mechanotransduction in response to alterations in shear stress or mechanical stretch (Davies, 2009). Vascular cells respond to mechanical forces and are able to translate them into biochemical events including gene expression and protein secretion. Various mechanisms have been proposed to explain vascular mechanotransduction at the cellular and molecular levels. Vascular cells have a variety of cell surface receptors including the integrins that allow the artery detect and respond to mechanical forces (Hahn and Schwartz, 2009). In addition, studies have implicated signaling mechanisms such as receptor tyrosine kinases, mechanosensitive ion channels, and G proteins in cellular mechanotransduction (Li et al., 2005; Orr et al., 2006). Also implicated are the cytoskeleton and other structural components that can transmit and modulate cellular tension through integrins/focal adhesion complexes that link to the extracellular environment (Li et al., 2005; Orr et al., 2006). This chapter focuses on the role of proteoglycans in vascular mechanotransduction. Proteoglycans are complex biomolecules that consist of a core protein that is post-translationally modified to have one or more glycosaminoglycan chains. Glycosaminoglycans are linear polymers of repeating disaccharides that is modified by multiple enzymes to have a complex sulfation and acetylation pattern (Sasisekharan and Venkataraman, 2000). The addition of the glycosaminoglycan chains occurs in the transgolgi and proceeds in a processive manner to create a heterogeneous structure and with intricate microdomains. These complex molecules are known to play an important role in controlling vascular homeostasis and have an emerging role in mechanobiology. We first describe the various types of proteoglycans in the artery and then discuss how these molecules are involved in the sensing and adaptation of the vascular system to mechanical stimuli. Finally, future directions in examining the role of proteoglycans in vascular mechanobiology are discussed.

9.2 Proteoglycans of the Cardiovascular System 9.2.1 Glycosaminoglycans The most common types of glycosaminoglycans in the artery include heparan sulfate, chondroitin sulfate and hyaluronic acid (Fig. 9.1). Of these four, only hyaluronic acid is synthesized without being attached to a protein core. The other

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Fig. 9.1 The chemical structure of glycosaminoglycans found in the vascular system. Shown in include one potential structure for: (a) heparan sulfate, (b) chondroitin sulfate and (c) hyaluronan

glycosaminoglycans are found as post-translational modifications to core proteins. In the case of heparan sulfate, proteins are targeted for glycosylation in the transgolgi by having a particular amino acid sequence Ser-Gly (Ala)-X-Gly (Ala). This site accepts an initial tetrasaccaride synthesized by four enzymes (Esko and Zhang, 1996). Heparan sulfate synthesis is initiated by the heparan sulfate copolymerase that adds glucuronic acid and N-acetylglucosamine to produce the initial heparan sulfate structure (Rosenberg et al., 1997a). This initial chain is then heterogeneously modified by deacetlyation, epimerization and sulfation to create a intricate fine structure (Rosenberg et al., 1997b).

9.2.2 Heparan Sulfate Proteoglycans 9.2.2.1 Perlecan In the vascular system heparan sulfate is found on several core proteins including perlecan, glypicans, and syndecans. Perlecan is a large heparan sulfate proteoglycan found in the basement membrane (Fig. 9.2a). In humans it is the product of the HSPG2 gene with a molecular weight of 470 kDa and approximately 800 kDa after

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Fig. 9.2 The predominant proteoglycans of the vascular system: (a) perlecan, (b) glypican-1 (c) the syndecans (d) versican

post-translational glycosylation (Iozzo, 2005). It has a modular structure possessing a myriad of interactions with growth factors, extracellular matrix molecules and adhesion molecules. Its name is derived from its “pearls on string” appearance under

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rotary shadowing electron microscopy (Noonan et al., 1991). Perlecan has been shown to have a role in angiogenesis, atherosclerosis and vascular injury (Nugent et al., 2000; Iozzo and San Antonio, 2001). Heparin and heparan sulfate have also been shown to inhibit neointimal proliferation in animal models of vascular injury and disease (Clowes and Karnovsky, 1978; Guyton et al., 1980; Hoover et al., 1980; Lindner et al., 1992; Edelman et al., 1993; Volker et al., 1995). This inhibition is dependant on heparan sulfate proteoglycans but also requires a protein component (Ettenson et al., 2000). 9.2.2.2 Glypicans The glypicans are a family of cell surface heparan sulfate proteoglycans having a globular structure with membrane association due to a glycosylphosphatidylinositol (GPI) anchor (Fig. 9.2b) (Fransson, 2003). Predominantly heparan sulfate groups are attached to serine residues in consensus sequences located between the central domain and the C-terminal GPI-anchor. The GPI anchor can be cleaved phospholipase C or D and leads to shedding of glypican from the cell surface. The GPI anchor also localizes the protein to cholesterol and sphingolipid-rich lipid rafts within the cell membrane (Fransson, 2003). 9.2.2.3 Syndecans The syndecans are a family of transmembrane heparan sulfate proteoglycans found on the cell surface and shed in a soluble form (Bernfield et al., 1999). Each syndecan consists of an extracellular domain that contains glycosaminoglycan attachment sites, a single pass transmembrane domain, and a short cytoplasmic domain with multiple phosphorylation sites (Fig. 9.2c). The heparan sulfate and chondroitin sulfate glycosaminoglycan chains allow syndecans to interact with a large number of ligands including FGF-2, VEGF, PDGF and TGF-β (Tkachenko et al., 2005). The interaction of syndecans with FGF-2 is probably the most characterized of these interactions. Syndecans and the attached heparan sulfate proteoglycan are essential for effective binding and signaling of the FGF receptor (Nugent and Iozzo, 2000). On the cell surface syndecans stabilize the FGF-2/FGFR complex and are essential for downstream signaling (Nugent and Iozzo, 2000). When shed from the surface, syndecan-1 can inhibit FGF-2 induced cell proliferation (Mali et al., 1993). However, physiologic degradation of syndecan by heparanase may lead to heparan sulfate fragments that enhance FGF-2 signaling (Kato et al., 1998). Syndecan-4 has also been shown to interact with FGF-2 and promote FGF-2 signaling (Volk et al., 1999). Recent work has also found that syndecans can act independently of the FGF receptor to act as a transmembrane receptor of FGF (Chua et al., 2004). The transmembrane domain and two regions of the short cytoplasmic domain of the syndecans are highly conserved. Conserved region 1 and 2 (C1 and C2) are separated by a variable domain (V) that is specific to each syndecan type (Fig. 9.2c). The C1 region has been found to bind src (Kinnunen et al., 1998), ezrin (Granes et al., 2000), and tubulin (Brockstedt et al., 2002). The C2 region

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contains a binding site for PDZ-domain proteins linking the syndecans to CASK, syntenin and other molecules (Grootjans et al., 1997; Cohen et al., 1998; Hsueh et al., 1998; Ethell et al., 2000; Gao et al., 2000). This region has been shown to control trafficking of syndecan-1 in epithelial cells (Maday et al., 2008). The variable region has differential functions in each syndecan type. The variable region of syndecan-4, in particular, is involved in focal adhesion formation, protein kinase C activation and binding of α-actinin (Couchman, 2003). The cytoplasmic variable region of syndecan-1 is known to affect cellular adhesion, migration and spreading (Chakravarti et al., 2005). Transforming growth factor-β (TGF-β) has been shown to interact with the heparan sulfate chains on syndecans. Syndecan-2, in particular, has been shown to interact with TGF-β via a protein-protein interaction (Chen et al., 2004). The exact nature and role of this interaction is complex and still remains to be elucidated. The presence of syndecan-2 may serve to compete with betaglycan (TGF receptor type III) for the binding of synectin. Synectin stabilizes betaglycan on the cell surface and, consequently, syndecan-2 may serve to reduce signaling in the TGF pathway (Chen et al., 2004; Tkachenko et al., 2005). The syndecans have an intricate role in orchestrating development and are known to be involved in cell-cell and cell-matrix adhesion. Syndecan-1 has been shown to be important for cell adherence to type-I collagen (Sanderson et al., 1989). In addition, syndecan-1 stabilizes the interactions of vitronectin with αvβ3 integrin (Beauvais et al., 2004). During migration syndecan-1, syndecan-4 and calveolin are directed to the region of cell contraction (Baciu and Goetinck, 1995; Borset et al., 2000; Beardsley et al., 2005). Syndecan-4 has also been shown to be an essential component for the activation of focal adhesion kinase and is known to bind fibronectin with its heparan sulfate chains (Mukai et al., 2002; Wilcox-Adelman et al., 2002; Hsia et al., 2003). In vascular smooth muscle cells exposed to shear stress syndecan-4 has been shown to dissociate from focal adhesions (Li and Chaikof, 2002). Several studies have revealed differential regulation of syndecans by various growth factors and cytokines. Fibroblast growth factor-2 (FGF-2) has been shown to increase syndecan-4 expression in vascular smooth muscle cells (Cizmeci-Smith et al., 1997). Arterial injury and myocardial infarction have also been shown to increase syndecan-4 expression (Geary et al., 1995; Li et al., 1997; Li and Chaikof, 2002). Stimulation with tumor necrosis factor-α (TNF-α) increases syndecan-2 expression and decreases syndecan-1 in endothelial cells (Halden et al., 2004). Transforming growth factor β2 increases syndecan-4 and decreases syndecan-1 in epithelial cells (Dobra et al., 2003).

9.2.3 Chondroitin Sulfate/Dermatin Sulfate Proteoglycans 9.2.3.1 Versican Versican is the major extracellular chondroitin sulfate bearing proteoglycan in arteries (Wight and Merrilees, 2004). This molecule has multiple sites for chondroitin

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sulfate attachment and reaches a molecular weight exceeding 1000 kDa (Fig. 9.2d). Versican has been linked to controlling the viscoelastic behavior in many tissues (Ludwig, 2007). It also has a pro-migratory affect in vascular smooth muscle cellsand various tumor cell types. Versican has multiple molecular interactions with other extracellular matrix components including hyaluronan, CD44, tenascin R and the fibulins (Wu et al., 2005). 9.2.3.2 Biglycan Biglycan is the predominant chondroitin sulfate-bearing proteoglycan found on the endothelial cell surface. It has been found to be associated with lipid deposition and atherosclerosis. Biglycan binds to TGF-β, collagen I/IV, fibronectin and other extracellular matrix molecules. It is thought to have pro-inflammatory activity and has a role in activating TNF-α and MIP-2 in macrophages. Biglycan can be positively regulated by TGF-β in response to abnormal mechanical stresses. In addition, nitric oxide (NO) can down regulate biglycan and can thereby link bilgycan expression to shear stress-induced NO levels (O’Brien et al., 1998; Williams, 2001). 9.2.3.3 Decorin Decorin is another arterial proteoglycan that is similar to biglycan in that it is a member of the small leucine-rich proteoglycan (SLRP) gene family. Decorin, howver, has different biological activity in comparison to biglycan. It can bind to IGF-R and IGF-I and regulate this pathway in endothelial cells. In addition, decorin induces expression of p21WAF1, a cyclin-dependent kinase inhibitor, that can arrest the cell cycle (Williams, 2001).

9.2.4 Hyaluronan Hyaluronan is a glycosaminoglycan with several distinct properties including its lack of attachment to a protein core, broad range of molecular weights (5–20000 kDa) and lack of sulfated groups (Pure and Assoian, 2009). It is a non-branching polymer of repeating disaccharides of D-glucuronic acid and N-acetylglucosamine. Hyaluronan is strongly expressed in the extracellular matrices of cells that are proliferating or migrating. In particular, it is highly expressed during healing, inflammation and development (Fraser et al., 1997). One of the major receptors for hyaluronan is CD44, a transmembrane glycoprotein. Through its binding to CD44 hyaluronan can be linked to the actin cytoskeleton, rho family GTPases and the ERM (ezrin/radixin/moesin) proteins (Wight and Merrilees, 2004; Cain et al., 2005).

9.2.5 Sialic Acid In addition to the glycosaminoglycans/proteoglycans mentioned above there are a number of sialic acid modified surface proteins found on the vascular cell surface

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(von Gunten and Bochner, 2008). While sialic acid is a term that encompasses either the N- or O-substituted derivatives of neuraminic acid, N-acetylneuraminic acid is the predominant form found in mammalian cells. Sialic acid modification is found in gangliosides and glycoproteins located at the cell membrane. Gangliosides are glyocolipids anchored in the cell membrane that have been found to mediate cellcell recognition, inflammation and growth factor signaling. Sialic acid groups serve as ligands for sialic acid-binding, immunoglobulin-like lectins (Siglecs). These lectins bind to specific forms of sialic acids mediating a number of immunological processes (von Gunten and Bochner, 2008).

9.2.6 Arterial Distribution of Glycoproteins 9.2.6.1 Glycocalyx The glycocalyx is a structure found on the luminal surface of the endothelium consisting of the glycoproteins and glycosaminoglycans (Fig. 9.3). In-vivo, the glyocalyx varies from 100 to 500 nm in thickness depending on location within the vascular tree and the local flow environment (Weinbaum et al., 2007). Notably, this structure is almost completely lost in in-vitro endothelial cell lines (Potter and

Fig. 9.3 Arterial distribution of proteoglycan in the artery. Boxes show magnified regions of the artery including the composition for the basement membrane underlying the endothelial layer and the glycocalyx that extends into the lumen of the artery

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Damiano, 2008; Chappell et al., 2009). The glycocalyx interacts directly with blood flow and is thus a prime candidate for the mechanotransduction of shear forces in the endothelial cells. In-vivo, this relatively thick structure is sufficient to reduce the fluid shear forces to nearly zero at the endothelial surface (Weinbaum et al., 2003). 9.2.6.2 Basement Membrane The basement membrane of the endothelial layer consists of multiple components (Fig. 9.3). Most notably there is a high concentration of perlecan that serves to inhibit vascular smooth muscle proliferation and migration as well as inhibit the proliferation of the endothelial cells themselves (Ettenson et al., 2000; Nugent et al., 2000; Baker et al., 2008; von Gunten and Bochner, 2008). In addition there are many protein constituents that include laminins, nidogens, agrins and nonfibrillar collagens (Iozzo et al., 2009). 9.2.6.3 Arterial Wall The arterial wall contains versican, biglycan, decorin and heparan sulfate proteoglycans (Fig. 9.3). Chondroitin sulfate proteoglycans are distributed throughout the wall in areas not occupied by fibrous components. Versican and biglycan both bind to lipoproteins and are markedly enhanced in atherosclerosis and neointimal hyperplasia following vascular injury (O’Brien et al., 1998; Williams, 2001; Wight and Merrilees, 2004).

9.3 Proteoglycans in Sensing Mechanical Forces 9.3.1 Proteoglycans in the Control of Flow-Induced Vasodilation Increases in blood flow cause arteries to dilate in an endothelium dependent manner (Kamiya et al., 1984; Pohl et al., 1991). This process is predominantly mediated by the release of nitric oxide (NO). There is a rapid response to the rate of change of the shear stress that is both G-protein and Ca2+ -dependent. The later phase of the response to shear depends on absolute shear level and is independent of G-protein and Ca2+ signaling pathways (Kuchan and Frangos, 1994; Kuchan et al., 1994). Several studies have supported the role of elements of the glycocalyx in control of the shear induced vasodilator response of arteries and are discussed below. Hecker et al. examined the vasoconstriction of rabbit femoral arteries in ex-vivo organ culture and found that incubation with neuraminidase, an enzyme that digests sialic acid moieties, reduced shear stress-induced NO release (Hecker et al., 1993). In contrast, neuraminidase digestion did not have an effect on acetylcholine-induced NO release. They also examined the release of the eicosanoid prostacyclin 2 (PGI2) in response to shear stress. This molecule is an important inhibitor of the platelet aggregation in blood clot formation and also has vasodilatory properties. Luminal

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neuraminidase digestion did not attenuate the shear stress dependent release of PGI2 (Hecker et al., 1993). In a parallel study, Pohl et al. examined diameter changes in response to flow and pressure changes in the mesenteric arteries of the rabbit (Pohl et al., 1991). They found that in-situ perfused mesenteric arteries had a three stage response to changes in perfusion flow and pressure. The first phase was a passive distention due to increased transmural pressure. The second phase is the myogenic response to the arterial stretch (Johnson, 1986). The myogenic response is the propensity of blood vessels to respond to transmural pressure elevation with constriction and to pressure reduction with dilation. This behavior is inherent to smooth muscle and is most pronounced in arterioles but can be demonstrated occasionally in arteries, veins, and lymphatics (Johnson, 1986). The late phase of the response to changes in perfusion is a flow-dependent dilation that overcomes the myogenic constriction leading to increased vessel diameter. Neuraminidase digestion completely inhibited the late phase of flow-induced vasodilation and enhanced the myogenic constriction of the artery (Pohl et al., 1991). More recently, Florian et al. studied the production of NO in cultured endothelial cells under shear stress (Florian et al., 2003). The cells were treated with an enzyme to digest heparan sulfate groups (heparinase) and then exposed to various shear stress regimes. This study demonstrated that heparan sulfate digestion leads to about a three-fold decrease in the NO production after exposure to steady shear stress as well as a four-fold decrease in the NO production following treatment with oscillatory shear stress. Similar studies found that pre-digestion with heparinase did not inhibit bradykinin-induced increases in NO production. Additional studies on endothelial cells in culture from this group also demonstrated that digestion with neuraminidase and hyaluronidase both attenuated shear induced NO release but digestion with chondroitinase did not. In addition, shear induced increases in PGI2 were not affected by removal of heparan sulfate, sialic acid, chondroitin sulfate or hyaluronan (Pahakis et al., 2007). In a recent study, VanTeefelen et al. hypothesized that the binding of heparin to the glycocalyx would alter the mechanotransduction of shear stress in the endothelium (Van Teeffelen et al., 2007). They found that treatment with clinical levels of heparin led to a reduction in the duration of vasodilation in response to occlusioninduced reactive hyperemia in mice. This again adds to the evidence that heparan sulfate is involved in NO-mediated vasodilation in response to changes in fluid flow. While there are multiple studies supporting a role for glycocalyx proteins in the arterial mechanotransduction of shear forces, the mechanisms remain to be uncovered. The majority of studies have used digestive enzymes whose activity is not necessarily restricted to the luminal surface or to endothelial cells versus vascular smooth muscle cells. Thus, it remains unclear if the model of fluid shear acting on the glycocalyx is correct in the case of heparan sulfate proteoglycans or if these effects are due to alterations in heparan sulfate mediated extracellular matrix interaction. Another notable complexity is that heparan sulfate can mediate the vasodilatory effects of released factors such as FGF and can consequently have indirect effects on these processes (Cuevas et al., 1991). One recent study examined the role of

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the glycocalyx and the reactive oxygen species in the shear-induced NO release (Kumagai et al., 2009). In this work, it was found that the superoxide dismutase mimic superoxide scavenger (Tempol) restored the vasodilatory response and NO increase in heparinase or neuraminidase digested arteries. However, Tempol did not inhibit the vasodilation/NO response in hyaluronidase digested arteries. Endothelial superoxide dismutase (SOD) catalyses the dismutation of superoxide into oxygen and hydrogen peroxide and thus has anti-oxidative properties. Heparan sulfate binds to endothelial SOD and can localize in the extracellular matrix. It has an important role in maintaining endothelial NO as it diffuses to the vascular smooth muscle cells in the arterial wall. Thus, the SOD binding properties of heparan sulfate may represent a potential mechanism of involvement of heparan sulfate in flow-induced vasodilatory responses.

9.3.2 Glycocalyx in Shear Stress Induced Vascular Smooth Muscle Cell Contraction While shear-stress induced responses are most often thought of as being associated with endothelial cells, vascular smooth muscle cells can also be exposed to fluid flow shear stress after endothelial denudation and through transmural interstitial flow (Cohen et al., 1995). Although vascular stretch has been proposed as the major mechanism in determining myogenic response, other studies suggest that this response can occur in the presence of increased pressure without vascular stretch. Ainslie et al. examined the contraction of vascular smooth muscle cells in culture after expose to a step increase in shear stress (Ainslie et al., 2005). Digestion of the chondroitin sulfate or heparan sulfate chains inhibited contraction after exposure to shear stress, suggesting a potential role of chondroitin sulfate/heparan sulfate in this behavior.

9.3.3 Role of HSPGs in Controlling Mechanically-Induced Changes in Cell Migration, Proliferation and Adhesion Moon et al. found that disruption of HSPGs by heparinase decreased endothelial adhesion and strength of adhesion (Moon et al., 2005). Further, heparinase digestion decreased actin stress fiber formation, focal adhesions and enhanced cellular motility. Under flow conditions cells with digested HSPGs lost the directional migration induced by fluid flow. In addition, loss of heparan sulfate lead to a reduction in focal adhesion formation and recruitment of focal adhesion kinase (FAK) aligned with fluid flow. Fluid flow is also known to reduce the proliferation of endothelial cells and induce alignment in the direction of flow. Yao et al. showed that both of these phenomenons did not occur after heparinase III digestion of the cultured endothelial cells (Yao et al., 2007). In addition, they found that fluid flow induced a cellular redistribution of HSPGs to the cell-cell interface of the endothelial layer.

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9.4 Mathematical Models of Glycocalyx-Based Mechanotransduction Weinberg and colleagues have proposed a general model of fluid glycocalyx interaction and force transmission (Weinbaum et al., 2003). Their model assumed that the glycocalyx has a quasiperiodic structure based on experimental observations (Squire et al., 2001). These studies revealed the ultrastructural details of the glycocalyx as a network with a characteristic spacing of 20 nm with periodic centers spaced at 10–12 nm. In their model the glycocalyx elements are linked to the cortical layer of actin that links the dense peripheral actin band. Their model predicts that the glycocalyx is fairly rigid and reduced the fluid shear stress at the surface of the cell to almost zero. As fluid flows over the surface of the glycocalyx the resultant torque is applied through first ~25 nm of the glycocalyx. This torque is transmitted to the cortical actin and applied to the dense peripheral actin band causing disruption of this actin network. Their theory was consistent with the observations of fluid stress causing disruption of this actin network (Weinbaum et al., 2003).

9.5 Long Term Adaptations of the Vascular System to Alterations in Mechanical Stresses While the majority of studies have examined the role of proteoglycans in acute cellular and arterial vasodilatory responses, the role of these molecules in the long term changes that occur in vascular remodeling has been less well studied. Tissue level vascular remodeling is highly dependent on the mechanical environment. An increase in shear stress in arteries and arterioles causes the artery to remodel to expand the luminal diameter. Conversely, reduction in flow leads to decreased arterial diameters in the long term. These observations lead Murray to suggest that the artery adapts to normalize the wall shear stress to a preferred, homeostatic level (Murray, 1926). “Murray’s Law” has been generally validated for most arteries (Kamiya et al., 1984). Cyclic circumferential arterial stress arises due to blood pressure during the cardiac cycle. Similar to the control of luminal diameter to normalize arterial shear stress, the artery adjusts the number of elastic laminae and arterial thickness to obtain a preferred wall stress due to arterial pressure. The role of proteoglycans in these long term adaptations to mechanical stimuli are relatively unknown. The normal arterial structure of the vessel results from the dynamic communication between the endothelium, vascular smooth muscle cells, tissue resident/recruited inflammatory cells and adventitial cells surrounding the artery. In the context of controlling arterial structure, endothelial cells produce several inhibitory factors that limit the growth of vascular smooth muscle cells within the artery (Ettenson et al., 2000; Nugent et al., 2000). Among these, the heparan sulfate proteoglycans are potent inhibitors of vascular smooth muscle proliferation and migration. Both heparin and endothelial cell heparan sulfate proteoglycans are potent inhibitors of vascular smooth muscle cell proliferation and FGF-2 induced

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mitogenesis (Clowes and Karnowsky, 1977; Ettenson et al., 2000; Nugent et al., 2000). Heparan sulfate-induced regulation is growth state dependent, with subconfluent cultures of endothelial cells stimulating vascular smooth muscle cell growth and postconfluent cultures inhibiting vascular smooth cell growth (Ettenson et al., 2000). Consistent with these findings, both, perlecan and endothelial-derived heparan sulfate proteoglycans have been shown to be essential in inhibiting the neointimal response to vascular injury (Nugent et al., 2000). A recent study from our group aimed to examine how mechanical forces alter the dynamic interplay of regulatory signals between endothelial and vascular smooth muscle cells (Baker et al., 2008). In this work, we applied mechanical stretch to cultured endothlelial cells and examined their production of growth-inhibitor heparan sulfate proteoglycans. Mechanical stretch stimulated increase endothelial inhibition of vascular smooth cell proliferation. Heparan sulfate proteoglycans and the core protein perlecan were both in creased by exposure to mechanical stretch. Knock down studies demonstrate that perlecan was required to increase vascular smooth muscle cell inhibition. Experiments using small molecule inhibitors and neutralizing antibodies demonstrated that a complex autocrine signaling cascade controlled the mechanical-load induced production of the growth inhibitory heparan sulfate proteoglycan perlecan (see Fig. 9.4). This pathway involved autocrine TGF-β, ERK and p38 MAPK signaling to induce perlecan expression in endothelial cells and lead to increased vascular smooth muscle growth inhibition. Using ex-vivo arteries exposed to high pressure under physiological flow conditions, these pathways were found to regulate the production of heparan sulfate proteoglycans in intact arterial segments.

Fig. 9.4 Mechanical feedback control mechanisms of arterial structure in response to mechanical stretch. The molecular signaling pathway shown refers to the pathways involved in controlling the mechanical load-induced endothelial production of growth inhibitory heparan sulfate proteoglycans. These factors are increased with mechanical load and partially inhibit the pro-growth signals in vascular smooth muscle cells

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The finding that endothelial cells produce more inhibitory factors to vascular smooth muscle cells with mechanical load is in contrast with the behavior of vascular smooth muscle cells under similar mechanical strain conditions. Mechanical stretch induces vascular proliferation (Hishikawa et al., 1997; Morita et al., 2004), release of FGF (Cheng et al., 1997) production of PDGF (Ma et al., 1999), and production of cell surface associated heparan sulfate proteoglycans (Li and Chaikof, 2002) (Fig. 9.4). Thus, under mechanical stimulation the endothelium provides negative feedback control inhibiting the progrowth signals produced by vascular smooth muscle cells under mechanical strain. Consequently, in the absence of an intact endothelium due to disease of injury mechanical stretch would induce vascular smooth muscle cells to produce growth stimulatory factors that are unchecked by load-enhanced endothelial cell paracrine growth inhibition. With endothelial repair, the inhibitory responsiveness to mechanical strain is restored as well as additional means of responding to hemodynamic forces can be applied including regulation of vascular tone and inhibition of vascular smooth muscle cell proliferation.

9.6 Conclusion and Perspectives There remain many unanswered questions in terms of the role of proteoglycans in mechanotransduction. While there is ample evidence that these molecules participate in sensing and responding to mechanical forces, there is little data that directly addresses the role of individual core proteins in mechanotransduction. Several studies have demonstrated that the expression of syndecan-1 and syndecan-4 are increased in mechanically stimulated cells. The syndecans are good candidates for mechanotransduction pathways as they have multiple interactions with integrins, focal adhesion, cytoskeletal elements and growth factor signaling (Morgan et al., 2007). While the last decade has yielded many new insights into the basic biology of proteoglycans, future studies will hopefully be able to shed additional light on the roles these molecules play in mechanotransduction and vascular remodeling.

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Borset M, Hjertner O, Yaccoby S, Epstein J, Sanderson RD (2000) Syndecan-1 is targeted to the uropods of polarized myeloma cells where it promotes adhesion and sequesters heparin-binding proteins. Blood 96:2528–2536 Brockstedt U, Dobra K, Nurminen M, Hjerpe A (2002) Immunoreactivity to cell surface syndecans in cytoplasm and nucleus: tubulin-dependent rearrangements. Exp Cell Res 274:235–245 Cain SA, Baldock C, Gallagher J, Morgan A, Bax DV, Weiss AS et al. (2005) Fibrillin-1 interactions with heparin. Implications for microfibril and elastic fiber assembly. J Biol Chem 280:30526–30537 Chakravarti R, Sapountzi V, Adams JC (2005) Functional role of syndecan-1 cytoplasmic V region in lamellipodial spreading, actin bundling, and cell migration. Mol Biol Cell 16:3678–3691 Chappell D, Jacob M, Paul O, Rehm M, Welsch U, Stoeckelhuber M et al. (2009) The glycocalyx of the human umbilical vein endothelial cell: an impressive structure ex vivo but not in culture. Circ Res 104:1313–1317 Chen L, Klass C, Woods A (2004) Syndecan-2 regulates transforming growth factor-beta signaling. J Biol Chem 279:15715–15718 Cheng GC, Briggs WH, Gerson DS, Libby P, Grodzinsky AJ, Gray ML et al. (1997) Mechanical strain tightly controls fibroblast growth factor-2 release from cultured human vascular smooth muscle cells. Circ Res 80:28–36 Chua CC, Rahimi N, Forsten-Williams K, Nugent MA (2004) Heparan sulfate proteoglycans function as receptors for fibroblast growth factor-2 activation of extracellular signal-regulated kinases 1 and 2. Circ Res 94:316–323 Cizmeci-Smith G, Langan E, Youkey J, Showalter LJ, Carey DJ (1997) Syndecan-4 is a primaryresponse gene induced by basic fibroblast growth factor and arterial injury in vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 17:172–180 Clowes AW, Karnovsky MJ (1978) Suppression by heparin of injury-induced myointimal thickening. J Surg Res 24:161–168 Clowes AW, Karnowsky MJ (1977) Suppression by heparin of smooth muscle cell proliferation in injured arteries. Nature 265:625–626 Cohen AR, Woods DF, Marfatia SM, Walther Z, Chishti AH, Anderson JM (1998) Human CASK/LIN-2 binds syndecan-2 and protein 4.1 and localizes to the basolateral membrane of epithelial cells. J Cell Biol 142:129–138 Cohen MI, Wang DM, Tarbell JM (1995) Measurement of oscillatory flow pressure gradient in an elastic artery model. Biorheology 32:459–471 Couchman JR (2003) Syndecans: proteoglycan regulators of cell-surface microdomains? Nat Rev Mol Cell Biol 4:926–937 Cuevas P, Carceller F, Ortega S, Zazo M, Nieto I, Gimenez-Gallego G (1991) Hypotensive activity of fibroblast growth factor. Science 254:1208–1210 Davies PF (2009) Hemodynamic shear stress and the endothelium in cardiovascular pathophysiology. Nat Clin Pract Cardiovasc Med 6:16–26 Dobra K, Nurminen M, Hjerpe A (2003) Growth factors regulate the expression profile of their syndecan co-receptors and the differentiation of mesothelioma cells. Anticancer Res 23: 2435–2444 Dzau VJ, Gibbons GH (1993) Vascular remodeling: mechanisms and implications. J Cardiovasc Pharmacol 21(Suppl 1):S1–5 Edelman ER, Nugent MA, Karnovsky MJ (1993) Perivascular and intravenous administration of basic fibroblast growth factor: vascular and solid organ deposition. Proc Natl Acad Sci U S A 90:1513–1517 Esko JD, Zhang L (1996) Influence of core protein sequence on glycosaminoglycan assembly. Curr Opin Struct Biol 6:663–670 Ethell IM, Hagihara K, Miura Y, Irie F, Yamaguchi Y (2000) Synbindin, A novel syndecan-2binding protein in neuronal dendritic spines. J Cell Biol 151:53–68 Ettenson DS, Koo EW, Januzzi JL, Edelman ER (2000) Endothelial heparan sulfate is necessary but not sufficient for control of vascular smooth muscle cell growth. J Cell Physiol 184:93–100

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Part IV

Mechanotransduction in the Lung

Chapter 10

Control of TRPV4 and Its Effect on the Lung James C. Parker and Mary I. Townsley

Abstract The transient receptor potential vanilloid 4 (TRPV4) non-selective cation channel has emerged as a critical channel for initiating the increased vascular permeability induced by high airway or vascular pressures in the lung. TRPV4 gating is regulated by multiple factors: mechanical stress, heat, epoxyeicosatrienoic acids (EETs) – the arachidonic acid metabolites of P450 epoxygenases, and phorbol esters. Increased pulmonary venous pressure and ventilation with high peak inflation pressures increase endothelial calcium influx, nitric oxide production, and vascular permeability in a TRPV4 dependent fashion in intact lungs. The permeability response to excess mechanical stress is attenuated by inhibition of cytosolic phospholipase A2 or P450 epoxygenases, and permeability increases in response to infusion of EETs. Various molecular mechanisms have been implicated for regulating TRPV4 gating, including channel translocation, direct ligand binding and phosphorylation. However, the mechanisms for EET dependent regulation of TRPV4 or amplification of TRPV4 by phosphorylation in intact lungs subjected to mechanical stress have not been clarified. Keywords Transient receptor potential vanilloid · Ventilator induced lung injury · Calcium · Pulmonary hypertension · Pulmonary edema · Epoxyeicosatrienoic acids

10.1 Mechanical Stress in Lung and TRPV4 The lung is continuously subjected to mechanical movement and cellular strain during breathing. As the lung inflates, extra-alveolar vessels, airways and alveolar sacs elongate and/or distend. Similarly, with an increase in alveolar volume, lateral forces in the alveolar septal wall lead to expansion of its surface area and elongation J.C. Parker (B) Department of Physiology and Center for Lung Biology, MSB 3074, University of South Alabama, Mobile, AL 36688, USA e-mail: [email protected]


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and flattening of capillaries. The pulmonary circulation is also in constant motion due to circumferential and shear stresses which accompany pulsatile blood flow. While lung function is maintained over a physiological range of mechanical stress, excessive stress is associated with acute lung injury. The transient receptor potential vanilloid 4 (TRPV4) channel is an important sensor for detecting mechanical stress induced by hypotonicity (cell swelling), shear, and stretch (Nilius et al., 2004; Cohen, 2005; Nilius and Voets, 2005; Liedtke, 2006; Townsley et al., 2006). TRPV4, a member of the TRPV family of cation channels, is expressed in lung epithelium, endothelium, fibroblasts and macrophages (Alvarez et al., 2006). Among other members of this TRPV family, TRPV4 has the greatest responsiveness to mechanical perturbation and is likely involved in cell volume regulation and mechanical sensing in all lung cells. Single channel conductance for TRPV4 is approximately 100 pS for outward currents and 60 pS for inward currents; inward currents can be blocked by ruthenium red. The channel permeability for Ca2+ is ∼6- to 10-fold higher than that for sodium (Everaerts et al., 2010; Nilius et al., 2004). Yin et al. (2008) reported inward calcium currents in whole cell patch clamp recordings in pulmonary microvascular endothelial cells after treatment with the TRPV4 agonist, 4α-phorbol-12,13-didecanoate (4αPDD) which was inhibited with 8Br-cGMP. TRPV4 channels are gated by multiple stimuli and likely integrate a number of external stimuli for cell responses. Activation occurs during hypotonic stress and at temperatures above 27◦ C. Vreins et al. (2006) demonstrated that cell swelling-induced activation of TRPV4 required phospholipase A2 -mediated release of arachidonic acid and subsequent metabolism of arachidonic acid by P450 epoxygenases to epoxyeicosatrienoic acids (EETs). Further, the P450 epoxygenase products 5,6- and 8,9-EET appear to activate TRPV4. Watanabe et al. (2003a) reported Ca2+ transients via heterologously expressed TRPV4 in HEK293 cells in response to arachidonic acid, which were completely abolished by blockade of P450 epoxygenase, indicating that EETs were involved in TRPV4 activation. Both 5,6and 8,9-EET evoked Ca2+ responses via the heterologously expressed channel and the endogenous TRPV4 in endothelium, with a smaller response to other EETs. Blockade of the cyclooxygenase and soluble epoxide hydrolases which degrade EETs also enhanced the swelling- and arachidonic acid-induced Ca2+ transients in mouse aortic endothelial cells (Vriens et al., 2005). Similar to hypotonicity-induced activation of TRPV4, shear stress activates heterologously expressed TRPV4 as well as endogneous TRPV4 in systemic endothelium in a phospholipase A2 -and EET-dependent manner (Hartmannsgruber et al., 2007; Loot et al., 2008; Wegierski et al., 2009). The activation of TRPV4 elicited by by 4α-phorbol-12,13-didecanoic acid (4αPDD) and heat does not appear to require EETs (Vriens et al., 2004).

10.1.1 Stretch Activated Cation Channels and Lung Injury Calcium entry through stretch activated cation channels and arachidonic acid metabolites have been previously implicated in acute lung injury. Parker et al.

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(1998a) reported a 3.7-fold increase in the filtration coefficient (Kf ), a sensitive measure of lung endothelial permeability, in isolated perfused rat lungs ventilated with 30 cmH2 O peak inflation pressure (PIP). This mechanical stretch-induced injury was completely prevented after treatment with gadolinium, a blocker of stretch activated cation channels. Subsequent studies confirmed that the increases in Kf and lung edema induced by mechanical ventilation were prevented by gadolinium (Parker and Yoshikawa, 2002). Although these studies implicated a mechanically gated Ca2+ permeable channel, the molecular nature of this stretch activated channel was unknown at the time. Previous studies had also implicated several signaling pathways now acknowledged as important for regulation of TRPV4 channel gating. Townsley et al. (1990) observed a synergistic impact of arachidonic acid and high vascular pressure on increases in Kf in isolated, perfused dog lungs. In both intact mice and isolated mouse lungs, the permeability lesion and edema resulting from high PIP ventilation were attenuated after blockade of cytosolic phospholipase A2 (Miyahara et al., 2008; Yoshikawa et al., 2005). Alvarez et al. (2004) reported an increase in Kf in isolated rat lungs of approximately 2.5-fold after infusion of 5,6- and 14,15-EET, whereas 8,9- and 11,12-EET had no significant impact. The permeability responses to 5,6- and 14,15-EET were attenuated by low extracellular Ca2+ but not by phospholipase A2 or P450 epoxygenase inhibitors. Figure 10.1 summarizes the impact of extracellular Ca2+ on Kf measurements in isolated rat lungs treated with the TRPV4

Fig. 10.1 Filtration coefficients (Kf ) in isolated rat lungs showing the effect of treatment with 4αphorbol-12,13-didecanoate (4αPDD), epoxyeicosatrienoic acids (EETs), and thapsigargin (TG). During baseline (BL) Kf and the initial Kf measurement after addition of channel agonists, a low Ca2+ perfusate was used. Subsequently, Ca2+ addback in the same lungs brought extracellular Ca2+ to a physiologic concentration (Alvarez et al., 2006)

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agonists 4αPDD, 5,6-EET or 14,15-EET, compared to that in lungs treated with thapsigargin, which activates store-operated Ca2+ channels (Alvarez et al., 2006). Kuebler et al. (2002) demonstrated that increases in venous pressure as small as 5 cmH2 O evoked Ca2+ entry in venular endothelium in isolated perfused lungs. These Ca2+ increases were abolished by gadolinium and Ca2+ -free perfusate. In other studies this group found that elevated left atrial pressure in isolated lungs induced increases in endothelial Ca2+ accompanied by P-selectin expression (Kuebler et al., 1999). However, Wu and colleagues recently documented a dependence of P-selectin expression on the voltage-gated T-type Ca2+ channel, rather than TRPV4 (Wu et al., 2009). Lung stretch induced by either high vascular pressure or by ventilation with high airway pressure induced production of endothelial nitric oxide (Kuebler et al., 2003), which could be blocked by inhibitors of nitric oxide synthase (NOS) or phosphatidylinositol-3-0H kinase. Kuebler et al. (2003) demonstrated a cumulative, linear, time dependent NO production measured by DAF-FM fluorescence in venular capillary endothelium in situ in isolated rat lungs induced by an increase in venous pressure of only 10 cmH2 O. Finally, Ito et al. (2010) reported that stretch of cultured pulmonary microvascular endothelial cells induced increases in intracellular Ca2+ that were sensitive to the TRPV antagonist, ruthenium red.

10.1.2 TRPV4 is Critical for Pressure Induced Lung Injury The TRPV4 channel has now been identified as a stretch-activated cation channel responsible for lung injury produced by both high vascular and airway pressures. Hamanaka et al. (2007) observed that high airway pressure increased Kf in isolated lungs of TRPV4+/+ mice (solid bars) but not in lungs of TRPV4-/- mice (shaded bars). Heat (40◦ C) amplified the mechanical injury only in TRPV4+/+ mice. The endothelial Ca2+ transients observed in TRPV4+/+ lungs during distention by high airway pressure were absent in TRPV4−/− mice and in TRPV4+/+ mice treated with ruthenium red, a TRPV channel antagonist. Lung injury was also blocked by inhibition of anandamide hydrolysis or cytochrome P450 epoxygenases. Figure 10.2 indicates the effects of high PIP ventilation, increased temperature, and inhibition of these pathways on Kf in isolated mouse lungs from both phenotypes. Jian et al. (2008) also measured a 4.5-fold increase in Kf in isolated lungs from TRPV4+/+ mice after venous pressure elevations of 30 cmH2 O, which was attenuated by approximately 80% in lungs from TRPV4−/− mice and ruthenium red-treated lungs from TRPV4+/+ mice. Significant attenuation of Kf was also achieved by phospholipase A2 and cytochrome P450 epoxygenase inhibitors. Endothelial Ca2+ transients and alveolar edema volumes were attenuated after pressure stress in TRPV4−/− lungs and by inhibition of Ca2+ entry, TRPV channels, and P450 epoxygenases. Figure 10.3 shows the increase in intracellular Ca2+ after an increase in pulmonary venous pressure in isolated mouse lungs from wild type mice untreated, treated with ruthenium red or treated with the P450 epoxygenase inhibitor (PPOH), and lungs from TRPV4 knockout mice either untreated or treated with gadolinium.

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Fig. 10.2 Impact of high airway pressure ventilation on filtration coefficients in lungs isolated from TRPV4+/+ and TRPV4−/− mice; (a) at 35◦ C, (b) at 40◦ C, and (c) after treatment of TRPV4+/+ lungs with inhibitors of TRPV, P450 epoxygenase and arachidonic acid (Hamanaka et al., 2007). PIP, peak inflation pressure

In situ fluorescence studies of lung endothelium by Yin et al. (2008) also indicated an increase in intracellular Ca2+ after a venous pressure increase of only 15 cmH2 O. Calcium entry was enhanced by blockade of soluble guanylate cyclase (sGC) or nitric oxide synthase (NOS), but decreased by the nitric oxide generator, S-nitrosoglutathione (GSNO), a sGC activator drug, or the cGMP analog, 8BrcGMP. Kf measurements in the isolated rat lungs mirrored the changes in endothelial Ca2+ . Using a nitric oxide (NO) sensitive dye, endothelial production of NO also

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Fig. 10.3 Relative Ca2+ fluorescence in lungs isolated from TRPV4+/+ (WT) and TRPV4−/− (KO) mice exposed to increased venous pressure (Pv) with and without treatment with P450 epoxygenase and TRPV inhibitors (Jian et al., 2008)

increased as venous pressure was increased, but was attenuated by the sGC activator or cGMP analogue. The vascular pressure-induced increase in Kf was attenuated by gadolinium or the NO donor GSNO, but was further increased by blockade of nitric oxide synthase (NOS). The increase in endothelial Ca2+ elicited by 4αPDD was also blocked by both ruthenium red and the cGMP analogue. Sildenafil, an inhibitor of PDE5, also attenuated the pressure-induced increases in lung Kf , protein leak and edema formation. These data indicate a TRPV4-mediated Ca2+ increase in lung endothelium after venous pressure elevation which increased vascular permeability, but which simultaneously activates a negative feedback control of the Ca2+ and permeability increases mediated by cGMP.

10.1.3 Phosphorylation and Mechanical Injury Activation of Src family kinases or myosin light chain kinase (MLCK) during mechanical distention of the lung appear to mediate the increased vascular permeability. Yin et al. (2008) found that inhibition of myosin light chain kinase (MLCK) protected against the permeability response to increased vascular pressure in isolated rat lung. Inhibition of MLCK or tyrosine kinases also attenuated airway pressure induced injury, whereas inhibition of tyrosine phosphatase augmented injury (Miyahara et al., 2007; Parker et al., 1998b). The mechanism by which MLCK amplifies TRPV4-mediated Ca2+ entry and lung injury is unknown, but may relate to the dependence of TRPV4 channel activity on actin cytoskeleton attachments and tension within the cytoskeleton (Becker et al., 2009; Ramadass et al., 2007). Alternatively, direct phosphorylation of TRPV4 may play a role. For example, the Src family kinases have been shown to amplify TRPV4 activity through phosphorylation of tyrosine 253 (Xu et al., 2003). Involvement of other kinases such as Akt/PKB may play a protective role. In isolated mouse lung, inhibition of Akt amplified the high airway pressure vascular permeability lesion (Miyahara et al., 2007), suggesting a protective pathway that likely acts through NO feedback.

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10.1.4 Segmental Vascular Effects of TRPV4 Figure 10.4 shows segmental and total Kf data in isolated rat lungs ventilated at high peak inflation pressures (PIP) in untreated lungs (solid bars) and lungs treated (shaded bars) with gadolinium chloride (Parker and Yoshikawa, 2002). The largest impact of ventilator-induced lung injury was on the lung microvascular compartment. Notably, segmental Kf values in high-pressure ventilated, gadolinium-treated lungs were not significantly different from baseline measurements. Although TRPV4 is expressed in endothelial cells in both alveolar and extra-alveolar vessels, activation of TRPV4 has the greatest effect on permeability in alveolar capillaries. Using electron microscopy, Alvarez et al. (2006) observed preferential separation of endothelial and epithelial layers as well as breaks in these layers in the alveolar septal wall in rat or mouse lung after TRPV4 activation. In contrast, activation of store-operated Ca2+ channels with thapsigargin preferentially disrupted inter-endothelial junctions of extra-alveolar vessels. Activation of storeoperated channels in cultured alveolar capillary endothelial cells results in only a small Ca2+ response with no concomitant permeability response (Cioffi et al., 2009). In contrast, pulmonary extra-alveolar vessels and pulmonary artery endothelial cells in culture which express TRPC1, 3, 4, and 6 display robust Ca2+ currents during store depletion and significant permeability increases (Townsley et al., 2006; Chetham et al., 1999). The impact of such heterogenous responses to activation of TRPV4 vs storeoperated channels on lung function has been investigated in a series of recent studies. In lungs treated with doses of either 4αPDD or thapsigargin sufficient to increase Kf by threefold, markedly different distributions of edema formation resulted. Alvarez et al. (2006) reported that only activation of TRPV4 was associated with alveolar flooding, as measured by the alveolar fluid volume fraction. Subsequently, Lowe et al. (2007) confirmed the impact of TRPV4 activation on

Fig. 10.4 Segmental and total filtration coefficients in isolated rat lungs after ventilation with high peak inflation pressures (PIP), showing attenuation of the injury by pretreatment with gadolinium (Parker and Yoshikawa, 2002, Reproduced with permission from the American Physiological Society)

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Fig. 10.5 Photomicrographs of toluidine blue-stained sections of rat lung showing preferential distribution of lung edema in perivascular cuffs induced by thapsigargin (a) and in alveoli after activation of TRPV4 with 14,15-EET (b). Pa, pulmonary artery; AF, alveolar fluid

alveolar flooding and further noted that TRPV4-mediated lung injury was associated with little perivascular cuff formation. In contrast, thapsigargin- treated lungs had cuff formation without alveolar flooding. Although the lungs in both groups gained approximately the same amount of edema, the thapsigargin-treated lungs had a threefold greater decrease in dynamic lung compliance compared to 4αPDDtreated lungs. Perivascular cuff formation presumably facilitated a collapse of the small airways to reduce compliance. Figure 10.5 contrasts the cuff formation in isolated rat lungs treated with thapsigargin (A) with the alveolar flooding resulting from TRPV4 activation via 14,15-EET (B), even when the increase in Kf was similar in these lungs. In addition to specificity for segmental vascular permeability, the TRPV4 channel also exhibits functional specificity within the alveolar septal endothelium. Wu et al. (2009) observed that TRPV4 activation increased endothelial cytosolic Ca2+ and Kf in isolated mouse lungs, without causing P-selectin expression. The same increase in intracellular Ca2+ elicited by a high potassium perfusate and activation of T-type Ca2+ channels resulted in P-selectin expression without any increase in permeability.

10.2 TRPV4 Structure and Potential Regulatory Sites As previously discussed, growing evidence supports a role for EETs as the signaling link between mechanical stress, evoked by hypotonicity, stretch or shear, and gating of the TRPV4 channel. All of the EET regioisomers have been shown to activate TRPV4 in vitro. Pinpointing the endogenous EET regioisomer involved is difficult, since each epoxygenase metabolizes arachidonic acid to produce a distinct mixture of EETs. Based on in vitro studies with microsomal fractions from

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rat and human lung, 14,15- and 11,12-EET proved to be the predominant products (Zeldin et al., 1996), though all were present in some amount. Thus, availability of EETs during mechanotransduction in lung cells will depend upon the epoxygenase expressed in the lung cell exposed to mechanical stress, as well as expression of the key enzyme responsible for EET degradation – soluble epoxide hydrolase (Spector, 2009). Interestingly, epoxygenase expression is itself up-regulated by mechanical stress (Fisslthaler et al., 2001). While in vitro studies have identified a number of regulatory domains in TRPV4, the molecular mechanisms underlying mechanotransduction-initiated and EET-dependent activation of TRPV4 have not been resolved. The TRPV4 channel consists of 871 amino acids with 6 transmembrane spanning segments and cytoplasmic C and N termini. The pore region lies between transmembrane segments 5 and 6, as shown in Fig. 10.6. EETs could potentially impact TRPV4 gating via a number of mechanisms.

10.2.1 Oligomerization and Membrane Localization Crystallography has resolved six ankyrin repeats in TRPV4’s N-terminus (reviewed in (Everaerts et al., 2010), which function in protein recognition. Splice variants of human TRPV4 lacking exons 5 and/or 7 which code for sequences within this region failed to produce Ca2+ transients in response to hypotonicity, 4αPDD, or arachidonic acid (Arniges et al., 2006). Arniges et al. reported that these variants also failed to localize to the plasma membrane due to their inability to oligomerize in the endoplasmic reticulum (Arniges et al., 2006). More recently, Becker and colleagues determined that the most distal C-terminal sequence (amino acids 828–871) of TRPV4 was also required for oligomerization and membrane trafficking (Becker et al., 2008).

Fig. 10.6 Diagram of TRPV4 structure showing potential sites for regulation of channel trafficking or gating. See text for more detail

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In addition to proper assembly, surface expression of TRPV4 is also modulated by ubiquitination, binding of PACSIN3 or microfilament-associated protein 7, and glycosylation (Wegierski et al., 2006; Cuajungco et al., 2006; Suzuki et al., 2003; Xu et al., 2006). PACSIN3 specifically interacts with a proline-rich domain upstream of the ankyrin repeats (Cuajungco et al., 2006). While these in vitro observations lend insight into assembly and trafficking of TRPV4, there are no reports which specifically evaluate whether active shuttling of TRPV4 plays a role in the response to mechanical stress in the intact lung, that is, whether mechanical stress evokes insertion of channels into the plasma membrane. Regulated trafficking is recognized for some TRP channels (reviewed in (Niemeyer, 2005; Cayouette and Boulay, 2007). Loot and colleagues (Loot et al., 2008) observed a shift in TRPV4 localization from a perinuclear distribution to the plasma membrane in cultured endothelial cells exposed to shear. This active trafficking required P450 epoxygenase activity. These observations need to be confirmed in lung cells exposed to mechanical stress in vivo.

10.2.2 Ligand Binding Pocket Activation of TRPV4 by 4αPDD is independent of the pathways utilized by cell swelling. The ligand-binding pocket between transmembrane domains 3 and 4 (S3 and S4) binds synthetic ligands such as the phorbol ester 4αPDD, while the binding domain for GSK1016790A has not been defined (Everaerts et al., 2010; Watanabe et al., 2002; Willette et al., 2008). Mutation of critical amino acids in these two transmembrane domains, specifically Y556, L584, and W586, markedly diminished the Ca2+ response to 4αPDD (Vriens et al., 2007). However, this binding pocket is not a likely site for EET interaction with TRPV4, since binding efficacy appears to require a dual ring structure (Everaerts et al., 2010) not present in EETs. Further, the Ca2+ response to 5,6-EET remained intact when these S3/S4 mutants were heterologously expressed. Vriens and colleagues consider the diminution of the Ca2+ response to hypotonicity, arachidonic acid and 5,6-EET resulting from mutation of more distal residues in S4 (Y591 and R594) to be due to altered channel gating rather than altered ligand binding. Phorbol esters other than 4αPDD, such as phorbol myristate acetate (PMA), also activate TRPV4 (Vriens et al., 2007), though in part activation is secondary to PKC-mediated phosphorylation. Although 4αPDD activates TRPV4 directly, heat appears to act through a soluble mediator.

10.2.3 Direct Binding of EETs Watanabe et al. found that 5,6-EET activated TRPV4 in inside-out membrane patches prepared from HEK293 cells or endothelial cells expressing heterologous or endogenous TRPV4, respectively (Watanabe et al., 2003a). This evidence has let to the conclusion that EETs activate TRPV4 in a membrane-delimited fashion and

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thus by direct binding to the channel. However, based on evidence derived from studies of EET-dependent regulation of other ion channels, the mechanism of action could be more complex. In coronary artery smooth muscle cells, EETs activate the large conductance Ca2+ -activated potassium channel (BKCa) in inside-out patches via a mechanism which requires GTP and Gαs (Li and Campbell, 1997). Further, EETs appear to elicit responses via binding to G protein-coupled receptors (Wong et al., 1997, 2000; Yang et al., 2008), suggesting the possibility for outside-in signaling. EETs activate Gαs , resulting in synthesis of cAMP and activation of PKA (Yang et al., 2008; Carroll et al., 2006; Node et al., 2001; Spector and Norris, 2007). While PKA-mediated phosphorylation has been implicated in TRPV4 activation with hypotonicity (see below), a role for G protein-coupled receptors, Gαs activation or cAMP synthesis has not been evaluated in lung’s response to mechanical stress.

10.2.4 Phosphorylation Domain mapping of the TRPV4 sequence has highlighted a number of potential phosphorylation sites (Nilius et al., 2004; Everaerts et al., 2010; Fan et al., 2009) for PKC, PKA and Src family tyrosine kinases. Kuebler and colleagues have recently reported that NO- and cGMP-dependent negative feedback regulation of TRPV4 gating in lung endothelium exposed to hydrostatic stress (Yin et al., 2008). However, this feedback control may not be direct, as there does not appear to be a PKG phosphorylation site in TRPV4. Most relevant to mechanical stress-induced EET-dependent regulation of TRPV4 are PKA and the Src kinases. EET-induced activation of Gαs leads to recruitment of cAMP-PKA signaling (Yang et al., 2008; Carroll et al., 2006; Node et al., 2001; Spector and Norris, 2007; Imig et al., 2008). While EETs are known to activate a number of tyrosine kinase cascades via Src kinases, MAPK or PI3 kinase (Spector and Norris, 2007), a role for tyrosine phosphorylation of TRPV4 in response to mechanical stress remains controversial. Xu et al. surveyed a number of Src kinase family members, investigating a role for tyrosine kinase-mediated phosphorylation of TRPV4 in the response to hypotonicity (Xu et al., 2003). Using a mutagenesis approach, these investigators mapped the Src kinase-mediated phosphorylation to Y253, within the first ankyrin repeat domain. While others found no role for tyrosine phosphorylation in hypotonicity-induced TRPV4 activation (Vriens et al., 2004), a recent study utilizing a mass spectrometric analysis has implicated Y110 in regulation of TRPV4 function (Wegierski et al., 2009). While translation of these observations to the in vivo setting is limited, the available evidence does support a role for tyrosine phosphorylation in the response to mechanical stress. Parker and colleagues have observed tyrosine phosphorylation in lung following high pressure mechanical ventilation and have implicated TRPV4, PI3K, Src kinases and Akt in the resultant ventilator-induced lung injury (Parker et al., 1998b; Hamanaka et al., 2007; Miyahara et al., 2007). Similarly, Src tyrosine kinases and TRPV4 have been implicated in mechanical hyperalgesia (Alessandri-Haber et al., 2008).

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10.2.5 Other Regulatory Domains TRPV4 contains several domains potentially regulated by protein-protein interactions. In addition to the contribution of PACSIN3 binding to channel trafficking, PACSIN3 binding to the N-terminal proline-rich domain in TRPV4 also inhibits the basal activity of the channel and attenuates TRPV4 activation by hypotonicity and heat (D hoedt et al., 2008). Further, deletion mutants lacking the N-terminal proline-rich domain were unresponsive to hypotonicity and to 4αPDD despite proper localization of TRPV4 to the plasma membrane (Garcia-Elias et al., 2008). This in vitro evidence highlights this region as critical for overall channel regulation. Two additional protein-binding sites exist in the C-terminus: a calmodulin-binding domain and a PDZ-like domain. Deletion of the PDZ-like domain does not alter the response to hypotonicity or 4αPDD (Garcia-Elias et al., 2008), but could participate in interaction of TRPV4 with other PDZ domain scaffolding proteins (Everaerts et al., 2010). The calmodulin-binding domain appears to participate in Ca2+ -dependent regulation of TRPV4 (Garcia-Elias et al., 2008; Strotmann et al., 2003). Interestingly, IP3 potentiates the Ca2+ response to 5,6-EET via IP3 receptor binding to this calmodulin domain (Fernandes et al., 2008). Activation of TRPV4 is also tightly controlled by both intracellular and extracellular Ca2+ levels. Strotman et al. (2003) demonstrated potentiation of TRPV4 Ca2+ entry during the initial increase in intracellular Ca2+ from baseline followed by a vigorous feedback inhibition at higher Ca2+ levels. Feedback inhibition of the Ca2+ entry was also greatly facilitated by increased extracellular Ca2+ concentration (Watanabe et al., 2003b).

10.2.6 Homo- or Heteromultimeric Channels Current evidence for heteromultimeric assembly of TRPV subunits into functional channels, based upon heterologous expression of FRET-TRPV reporter constructs, is conflicting. Hellwig et al. concluded that most TRPV channel subunits including TRPV4 preferentially assemble as homotetramers (Hellwig et al., 2005). In contrast, Cheng and colleagues found that TRPV proteins can form heteromultimeric channels with other proteins in this TRP subfamily (Cheng et al., 2007). TRPV4 has also been reported to form heteromultimers with TRPP2 (Köttgen et al., 2008). Since channel subunit stoichiometry can impact channel conductance and regulation, more definitive information regarding assembly of TRPV4 alone or with other potential TRP proteins in cultured lung cells and in the intact lung is needed. If indeed TRPV4 in lung is expressed as a component of a heterotetrameric channel, then EET-dependent regulation of the channel with mechanical stress becomes potentially more complex.

10.3 Conclusion and Perspectives TRPV4 is ubiquitously present in lung cells and appears to be the major channel for transduction of mechanical signals in these cells. Calcium influx occurs

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with relatively modest increases in pulmonary vascular or airway pressures, and the channel is essential for initiating the increased vascular permeability induced by high vascular and airway pressures. Mechanical gating is mediated by epoxyeicosatrienoic acids (EETs), whereas heat and phorbol ester activation occur through different mechanisms. TRPV4 is a major mediator of injury to pulmonary microvascular endothelium, which lacks the store operated TRPC channels of macrovascular endothelium. Several potential regulatory sites on the TRPV4 molecule have been identified for phosphorylation by Src family kinases, PKA, and PKC. Altered membrane trafficking, oligomerization and binding to scaffold proteins have been proposed as other potential regulatory mechanisms, but details of the mechanism whereby EETs or phosphorylation control channel opening during mechanical stress remain to be elucidated. Acknowledgements Research support by NIH HL066299 and HL092992

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Chapter 11

The Role of Protein-protein Interactions in Mechanotransduction: Implications in Ventilator Induced Lung Injury Matthew Rubacha and Mingyao Liu

Abstract Critically ill patients often require mechanical ventilation to support their breathing. This is especially true for patients with Acute Respiratory Distress Syndrome, where therapeutic intervention still remains largely ineffective. A clinical study supported by the National Institutes of Health has indicated that low tidal volume ventilation is beneficial, which has established mechanical ventilation an important contributor to lung injury in these patients. Mechanical ventilation can lead to increased production of cytokines and chemokines related to inflammation and tissue damage. Further understanding of mechanotransduction may reveal targeting strategies for therapeutic intervention. It is known that cells can sense mechanical forces across the plasma membrane through a variety of mechanisms. In addition, intracellular force sensors have been proposed to play an important role in conversion of physical forces into biochemical signals through protein-protein interactions. In this chapter we reviewed this novel mechanism for mechanosensation and mechanotransduction, and proposed to inhibit Src protein tyrosine kinase activation as a potential therapy for ventilator induced lung injury. Keywords AFAP · p130Cas · Src · Acute lung injury · ARDS

11.1 Introduction Like in many other solid organs, physical forces play an important role in the lung physiology, and are involved in the regulation of fetal lung development (Liu and Post, 2000), airway branching and alveolarization, and the regulation of pulmonary physiology (Liu et al., 1999). The lung is unique from other organs in that it is M. Liu (B) Faculty of Medicine, University of Toronto, 101 College Street Toronto Medical Discovery Tower, Room 2-814, Toronto, ON, M5G 1L7 Canada e-mail: [email protected]


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constantly expanding and contracting to facilitate the flow of air into and out of a complex network of airways. The physical force associated with normal breathing does not lead to aberrant side effects. Abnormal physical forces however, are known to contribute to the pathogenesis of lung diseases, such as asthma, pulmonary hypertension, chronic obstructive pulmonary disease, etc. Mechanical ventilation as a therapeutic modality can be used to provide life support during surgical operation, or to patients with respiratory distress. This externally applied physical force which ventilators use to force air into the lungs can lead to tissue damage and is associated with several side effects known as ventilator-induced lung injury (VILI) (Girard and Bernard, 2007). Before clinicians can address VILI, we need to recognize and understand more about the mechanisms and pathways of mechanotrasduction initiated both by normal breathing and by mechanical ventilation. Acute lung injury (ALI), and its most severe form, acute respiratory distress syndrome (ARDS), is defined by The American-European Consensus Conference on ARDS as severe gas exchange dysfunction and radiographic abnormalities after a predisposing injury in the absence of heart failure (Bernard et al., 1994). The age-adjusted incidence of ALI is around 86.2 per 100,000 person-years. There are approximately 190,600 patients diagnosed with ALI each year in the United States which is associated with 74,500 deaths and 3.6 million hospital days, representing a significant cost to the healthcare system (Rubenfeld et al., 2005). It has been known for some time that physical force associated with high tidal volume ventilation contributes to the progression of ALI/ARDS (Liu, 2007). Since the first recommendations from the ARDS network were published in 2000 (ARDSNetwork, 2000), the importance of controlled mechanical ventilation conditions has been realized. Physicians are required to perform a balancing act between using mechanical ventilation to support the lives of critically ill patients and limiting the injurious side effects that this treatment includes. The acute inflammatory response associated with ARDS is thought to be one of the major mechanisms associated with lung injury (Suter, 2006). Injurious mechanical ventilation leading to VILI is a contributor to this inflammatory reaction. There has been growing interest in the role that mechanotransduction plays in VILI in an effort to better understand how physical forces are transmitted into biochemical signals in the lung and how these signals can be controlled to reduce or eliminate the deleterious effects of mechanical ventilation while keeping the benefit (Liu, 2007; Oeckler and Hubmayr, 2007). There are a variety of mechanisms that cells can use to detect mechanical force including stretch mediated ion channels (Hamill and Martinac, 2001), extracellular matrix (ECM)-cytoskeleton perturbations (Ingber, 1991), and disruptions in cellcell adhesions (Ko et al., 2001). This chapter will focus on the role that proteinprotein interactions play in mechanosensation and mechanotransduction. There is evidence to show that multiple pathways can be activated by mechanical ventilation and researchers have begun looking at blocking the signals in the lung initiated by the mechanical force associated with injurious mechanical ventilation. We will discuss the possible therapeutic application of blocking Src protein tyrosine kinase (PTK) pathway to reduce ALI/ARDS associated with VILI.

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11.2 Basic Lung Mechanics vs. Mechanical Ventilation The lung is considered to be a gravitationally deformed elastic solid organ which shape matches that of its surroundings, namely, the chest wall and the heart. The shape conforming nature of the lungs leads to a non-uniform strain during inspiration. Since the weight of the lung itself is negligible when determining this strain, the movements of diaphragm and chest-wall are actually the main influences on regional ventilation (Vlahakis et al., 1999). It has been shown that the lung can expand in volume up to 40% from the functional reserve capacity to the total lung capacity. Because of the heterogeneity that exists at the micro-scale that cannot be accounted for by the force of gravity, this predicted expansion of lung tissue is likely an overestimate of the actual forces experienced at the cellular level. Airways and blood vessels are known to be more resistant to mechanical forces than the lung parenchyma itself. Based on observations in fixed tissue, airway micromechanics are strongly influenced by the role that elastin and collagen fibers play in bearing much of the mechanical strain experienced during normal ventilation (Vlahakis and Hubmayr, 2005). At the alveolar level, the shape of the airway is maintained by surface tension, and is thought to fold and unfold during the constriction and expansion cycle of normal respiration (Oldmixon and Hoppin, 1991). Using electron microscopy observations of the length of the basement membrane surrounding alveoli, it was estimated that the area of the basement membrane increased approximately 35% during inspiration, representing a cell stretch of around 15% from the functional reserve (Tschumperlin and Margulies, 1999). This prediction is limited by the fact that fixing tissue affects the hydration and surface tension of the specimen, which could impair accurate observations of the real lung architecture. In the future, it is hoped that living, unfixed samples can be studied to improve our understanding of the forces that the airway and alveoli experiences while breathing (Bachofen et al., 2002). Often the only life saving therapy for patients with ALI/ARDS, mechanical ventilation can lead to the production of inflammatory mediators. Understanding the basic concepts of lung mechanics helps to understand why mechanical ventilation may induce tissue injury. Vlahakis and Hubmayr have summarized the two leading theories as to why the forces acting on injured lungs leads to VILI while ventilation of normal lungs (i.e. during surgery) typically does not lead any significant side effects (Vlahakis and Hubmayr, 2005). First, as the injury to the lung progresses, alveoli become less able to facilitate the exchange of gas across the alveolar-blood barrier. This results in a smaller number of functional aerated alveoli being preferentially recruited, which end up bearing most of the tidal volume during an inspiratory maneuver. As a result, there is a regional over-distension in portions of the lung where airway epithelial cells experience a higher than normal mechanical force ultimately leading to further injury (Maunder et al., 1986). Secondly, because of local decreases in the ability of certain areas of the lung to expand during inspiration caused by edema, infection, or restrictive forces, there is heterogeneity in lung compliance. As the expansive force of a tidal volume inspiration leads to a non-uniform

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expansion of the lung there is shear stress between neighboring regions of different compliance which is not seen in normal lungs (Mead et al., 1970).

11.3 Ventilator Induced Inflammatory Response A biotrauma hypothesis has been proposed to account for the relationship between mechanical ventilation and the associated inflammatory response. The injurious mechanical ventilation may lead to the release of cytokines and pro-inflammatory mediators, resulting in excessive immune system activation (Tremblay et al., 1997). These mediators can cause lung damage themselves, or in the case of ventilated patients, become a contributing factor to their ongoing lung injury by further recruiting inflammatory cells into the lung (Girard and Bernard, 2007). In an isolated perfused rat lung model, injurious ventilation is associated with the production of TNF-α, IL-1β, and IL-6 among other inflammatory cytokines found in BAL fluid (Tremblay et al., 1997). In vitro, it has been shown that the stretch of A549 cells, a human alveolar epithelial cancer cell line, can cause the secretion of IL-8, an inflammatory cytokine, into the culture medium (Vlahakis et al., 1999). The release of both IL-8 and TGF-β from A549 cells in response to mechanical stretch is dependent upon the amount of physical force applied to the cells as measured by the percentage of elongation from rest (Yamamoto et al., 2002). The amount of force required to elicit cytokine response from A549 cells appears to be non-physiological, much more than cells are predicted to experience in vivo during normal breathing. BEAS-2B cells, a human normal bronchiole airway epithelial cell line, can also respond to mechanical stretch by secreting IL-8 (Oudin and Pugin, 2002). This cell line has been used to uncover the mechanism by which mechanical stretch force leads to the production of cytokines and chemokines. With 20% elongation, a significant increase in IL-8 release over unstimulated cells was observed. On top of this, paxillin, a component of the focal adhesion complex, was translocated from the plasma membrane to the peri-nuclear area. Under the same stretch conditions, application of Y-27632, a Rho-activated kinase inhibitor, completely abolished the increase in IL-8 secretion. Based on these observations, the authors concluded that integrin mediated signalling is critical for mechanotransduction in airway epithelial cells (Thomas et al., 2006). It is also believed that mechanical ventilation can inhibit lung repair after injury. Waters et al. have shown that mechanical stretch, simulating injurious ventilation can inhibit airway epithelial cell migration (Savla and Waters, 1998), an important process for repair and regeneration of airway epithelium after injury. It has been shown that cyclic mechanical stretch impairs airway cell migration via a pathway that involves FAK (focal adhesion kinase), JIP3 (JNK-interacting protein 3) and JNK (Jun-N-terminal kinase) (Desai et al., 2009). Mechanical stretch also decreased migration of primary cultured rat alveolar epithelial cells through inhibition of Rac1 related signalling (Desai et al., 2008). These studies and others have demonstrated activation of signal transduction pathways related to inflammatory responses. How physical forces associated with

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mechanical ventilation are sensed by cells in the lung and converted to signalling event is the focus of this chapter. Several basic mechanisms for mechanosensation and mechanotransduction have been discussed in this book. We will focus on one of the novel mechanisms, protein-protein interaction in mechanosensation and mechanotransduction.

11.4 Protein-Protein Interactions Signal transduction between cells, within cells, and from the extracellular to intracellular environment is critical for sustaining life. Proteins are responsible for mediating these signals through their intrinsic dynamic properties. Recently, there has been a large influx of information regarding protein interaction networks based on genetic and bio-physical studies (Yu et al., 2008). Adaptor proteins contain multiple protein-protein interaction domains which function to link binding partners together, creating a signaling complex. Through various combinations of protein interaction domains, adaptor proteins can link specific proteins together in response to cellular signals providing specificity and proper subcellular localization of signaling partners. In this manner, adaptor proteins are important for regulating the specificity of downstream cell signaling and regulating proteins which are important for individual signaling cascades in a spatial and temporal manner (Flynn, 2001). Studies on several different families of signal adaptors suggest that these proteins are not only used to temporally and spatially couple successive signaling proteins but are highly dynamic interactors capable of directing and regulating downstream signals. Typically, adaptors contain functional domains which can selectively recognize activated cellular surface receptors or intracellular signaling molecules and use one or more of their other domains to couple the receptor with their downstream effector(s) (Feller, 2001). A simple example of this phenomenon is the SH2/SH3 adaptor Grb2. Grb2 contains an SH2 (Src homology 2) domain which recognizes tyrosine phosphorylated YXN motifs on activated receptor tyrosine kinases (RTK) (Fig. 11.2a). Two SH3 (Src homology 3) domains are then able to recruit effector proteins, such as the Ras guanine nucleotide exchange factor Sos or docking proteins such as Gab1/2. Recruited proteins become phosphorylated and interact with other SH2 domains on downstream effectors such as PI3K or Shp2. In this manner, Grb2 is capable of facilitating the downstream signal cascade that links an activated RTK to Ras-MAPK or PI3K signaling pathways, ultimately having an effect on cellular function such as cell proliferation, migration, and differentiation (Rozakis-Adcock et al., 1993). In depth structural analysis of Grb2 demonstrated that the interaction domains of this adaptor function almost independently of each other (Maignan et al., 1995). This observation implies a very simple “linker” or organizer function for adaptor proteins as one binding motif can interact with upstream signaling proteins while another binds to downstream effectors, propagating a signaling cascade. As more adaptor proteins are studied, the complexity of their function is becoming

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more evident and the “linker”/organizer model for adaptor proteins appear to be the exception rather than the rule (Burns K, 1998). Adaptor proteins themselves can be dynamically regulated, adding an extra layer of complexity to the signaling cascade (Pawson, 2007). It has been demonstrated that the function of adaptor proteins can even be regulated by intracellular signals such as DNA damage (Stucki et al., 2005). The association of signaling proteins with adaptors can act cooperatively to enhance signaling or even antagonistically to gate a signal cascade (Hoshi et al., 2005). The complexity of signal adaptor proteins doesn’t end there. A single adaptor protein can be associated with different proteins depending on the signaling pathway functioning at any given time. In this way they serve to link different signaling proteins to different downstream effectors depending on the cell type or even subcellular compartment being investigated. A classic example of this is the intracellular protein trafficking of signaling proteins. This important cellular function is dependent on the diversity and specificity of adaptors involved in endocytosis and sorting in the endosomes (Traub, 2003). The various capabilities of adaptors demonstrate that the interaction subunits of these proteins are capable of much more than the simple “beads on a string” model with which they were originally labeled (Pawson, 2007). Knowing that protein-protein interaction can mediate and coordinate signal transduction pathways in the cell, we would like to know whether protein-protein interaction can be the first step to activate signal transduction, especially to convert physical forces to activation of protein tyrosine kinase for signal transduction. Several mechanosensory complexes have been defined and demonstrated that adaptor proteins can act as the actual mechanosensor or play an integral role in linking the mechanosensor to the downstream signaling pathways.

11.5 Unfolding of p130Cas as a Mechanosensor Previously it was thought that the force sensing function of cells occurs at the cytoplasm membrane via mechanisms such as calcium movement and ion channel gating (Hamill and Martinac, 2001), activation of cell surface receptors by their ligands (Correa-Meyer et al., 2002; Tschumperlin et al., 2004), ECM-integrin interactions on basolateral side of the cells by mechanical stretch (Chen et al., 1999; Ingber, 1991), or shear stress on adhesion molecules on the apical membrane of the cells (Tzima et al., 2005). Through the aforementioned studies it was unclear as to whether cells can sense mechanical force without intact cellular membranes. It is difficult to separate the role of ion channel and membrane protein function from that of the cytoskeleton cells use to bear much of the tension and physical force. One interesting way to solve this problem is to remove the non-cytoskeletal components of the cell through Triton X-100 treatment of cells cultured on a collagen coated matrix, leaving behind the Triton insoluble cytoskeleton. This process removes transmembrane ion channels as well as minimizes the influence of ion flux on mechanotransduction. In this manner, any measurable changes in protein status (i.e. protein phosphorylation) resulting from mechanical stretch can only result from interactions between proteins associated with the cytoskeleton. Sawada and Sheetz

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used this method to remove the cellular membrane and soluble proteins from the cytoplasm, and stretched the remaining cytoskeleton attached to the extracellular matrix. Detergent soluble proteins were then replaced for a short period of time, during which some were found to undergo tyrosine phosphorylation (Sawada and Sheetz, 2002). This interesting study suggests that in the absence of membrane related proteins and local changes in ion concentration mechanical force still can be sensed by cells. It has been reported that small GTPases can be involved in the mechanical force induced signal cascade (Sawada et al., 2001). Biaxial stretch was found to inhibit Ras activity, decrease lamellipodia formation, and activate Rap1. C3G (Rap1 guanine nucleotide exchange factor) is known to contain multiple SH3 domain binding motifs that are able to associate with the Crk family of adaptor proteins (Knudsen et al., 1994; Tanaka et al., 1994). It is believed that the association of the C3G-Crk complex with other proteins containing a phosphotyrosine residue allows for the release of cis-acting negative regulatory domain of C3G, facilitating its activation (Ichiba et al., 1999). Interestingly, Crk, C3G, and Rap1 have all been shown to be associated with Src PTK in the regulation of cell adhesion (Li et al., 2002), prompting researchers to suspect that this pathway may be associated with mechanotransduction. Using the model of cytoskeletal stretch, Tamada et al. showed that the Crk-C3GRap1 signaling pathway was activated. In response to stretch, Crk was recruited to ECM-cell contacts where it is believed to sense force applied to the cytoskeleton (via ECM stretch) and facilitates the activation of this signaling pathway. In addition, they showed that Src PTK was associated with the cytoskeleton after stretch, confirming the hypothesis that mechanical force can recruit Src for local activation by protein-protein interactions (Tamada et al., 2004). These studies showed that protein-protein interactions are able to facilitate mechanotransduction in the absence of transmembrane ion channels or cytoplasmic ion flux; however the question of “what is the actual mechanosensor?” remained unanswered. It has been reported that p130Cas, when tyrosine phosphorylated on its multiple internal YXXP repeats, can bind to the N-terminal SH2 domain of Crk. Interestingly, under normal conditions, the YXXP repeats of p130Cas are hidden through its folded tertiary structure and can be exposed only under certain conditions. p130Cas contains an N-terminal SH3 domain that can bind FAK as well as a C-terminal binding site for Src PTK. These two binding domains help to link p130Cas to focal adhesions where it is believed to play a role in mechanotransduction. When cells are mechanically stretched, p130Cas underwent extensive tyrosine phosphorylation at its multiple internal YXXP repeats (Sawada et al., 2006). Phosphorylation of p130Cas allows for Crk binding, C3G activation, and increased GTP-loading of Rap1. It was also shown that this effect was not the result of an increase of Src PTK activity, implying the possibility that under mechanical stretch, p130Cas is unfolded exposing internal tyrosine phosphorylation targets for Src that are not apparent under non-stretch conditions (Sawada et al., 2006). In this way, p130Cas can be considered a dynamic mechanosensor that can detect a cell stretch force, and facilitate the beginning of a signal cascade through the aforementioned protein-protein interactions (Pawson, 2007) (Fig. 11.1).

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Fig. 11.1 Mechanical stretch-induced unfolding of p130Cas as a mechanosensor. p130Cas is represented as an accordion-like structure with an N-terminal SH3 domain and a C-terminal Src binding region (top). The SH3 domain binds to FAK while the Src binding motif binds to a Src PTK. These interactions localize p130Cas to focal adhesion sites. Under stretch conditions, the accordion-like structure extends, and reveals multiple internal YXXP domains to Src PTK for phosphorylation (middle). The phosphorylation of YXXP motifs allows for the binding of the CrkC3G complex. This interaction leads to the activation of C3G and subsequent activation of the small GTPase Rap1. Reproduced from Pawson (2007) with permission from Elsevier

11.6 AFAP as an Activator of Src PTK In the above p130Cas model of mechanotransduction, the adaptor protein was stretched and unfolded by mechanical force to expose its internal tyrosine phosphorylation sites to Src PTK. However, it has been seen that the activities of Src PTK are increased by mechanical forces in multiple cell types (Franchini et al., 2000; Han et al., 2004; Naruse et al., 1998; Sai et al., 1999). This raises the interesting question: can adaptor proteins function as activators of protein kinases in order to convert physical force to biochemical reaction for signal transduction? Liu et al. found that actin filament associated protein (AFAP) could be a good candidate for protein-protein interaction related mechanotransduction in fetal rat lung cells (Liu et al., 1996). They used a model of cell culture where fetal rat lung cells were grown in a gelfoam matrix, an artificial 3-dimensional structure which provides a scaffold to support cells. Interestingly, under this condition, mixed fetal lung cells develop into alveolar like structures that mimic the architecture of the lung parenchyma (Liu et al., 1992). Cells were subjected to intermittent stretch that is believed to closely mimic the physiological forces experienced by alveolar epithelial cells during fetal breathing movements. It was shown that the stretch force could enhance cell proliferation (Liu et al., 1992, 1995b, 1993) through activation of c-Src (Liu et al., 1996), which can subsequently induce tyrosine phosphorylation of phospholipase C, generation of secondary messengers IP3 and DAG, increase in intracellular free calcium, and activation of protein kinase C (Liu et al.,

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1995a). Activation of c-Src also resulted in increased tyrosine phosphorylation of multiple proteins related to cytoskeleton, including FAK, p130Cas, cortactin, and paxillin among others, however, only the binding of c-Src to AFAP was increased by mechanical stretch, as determined by co-immunoprecipitation (Liu et al., 1996). AFAP is distributed along stress fibers and is closely associated with F-actin (Flynn et al., 1993). This puts AFAP in an excellent position to respond to small perturbations affecting the cytoskeleton. Indeed AFAP is unique in that its position changes with stress fibers in response to mechanical stretch. This may increase interaction of AFAP with other cytoplasmic proteins, including c-Src. Because AFAP contains high affinity binding sites for c-Src, it is possible that the interaction between AFAP and c-Src may lead to binding of these two proteins, and subsequently alter the confirmation of c-Src, resulting in its activation. This could be one of the initial steps of mechanotransduction in a well defined signal pathway (Liu and Post, 2000; Liu et al., 1999). In order to fully understand this hypothesis, more must be known about c-Src itself (Fig. 11.2b). When inactive, c-Src exists in a closed confirmation bound by two intra-protein interactions. Firstly, in the closed state the c-terminal tail is tyrosine phosphorylated and folds in on itself to interact with the SH2 domain. Secondly, the SH3 domain binds to the linker region between the SH2 and the kinase domain (Fig.11.3). c-Src can be activated either by dephosphorylation of tyrosine in its C-terminal tail, or by high affinity binding of its SH2 and/or SH3 domain(s) with another protein, which can be considered a Src activator. When active, c-Src unfolds to expose Y416 , located in the kinase domain. Once phosphorylated at Y416 , c-Src

Fig. 11.2 Proteins can be considered as molecules built with functional modules. (a). The multiple modular domain structure of Grb2 is typical for that of an adaptor protein. (b). Molecular structure of c-Src protein tyrosine kinase. (c). Molecular structure of actin filament associated protein (AFAP)

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Fig. 11.3 Mechanical stretch-induced Src activation via competitive binding with AFAP. In the inactive state, phosphorylation at Y527 in the C-terminus of c-Src binds to its own SH2 domain and the linker region between the SH2 and kinase domains binds to its own SH3 region. This intraprotein folding locks c-Src in a closed confirmation (left). When cells are stretched, deformation of F-actin stress fiber presents AFAP to c-Src. The SH3 and SH2 domain binding motifs of AFAP bind c-Src with high affinity, disrupting the intra-protein inhibitory interactions of c-Src (middle). While stabilized in the open confirmation by interactions with AFAP, c-Src can be phosphorylated at the Y416 residue, locking it in the open and active confirmation (right). Reproduced from J Biol Chem 2004;279:54793–54801. Permission granted from the Journal of Biological Chemistry

goes on to be a potent tyrosine kinase, involved in many different intracellular signaling processes (Xu et al., 1999). AFAP is a complex signal adaptor protein in that it contains a putative SH2 domain binding motif and a putative SH3 domain binding motif in its N-terminus. This is followed by a WW binding motif, then two PH domains which flank multiple internal Ser/Thr phosphorylation sites which in turn are flanked by two SH2 domain binding motifs finally followed by a C-terminal leucine zipper/actin binding domain (Baisden et al., 2001b) (Fig. 11.2c). With multiple protein interaction domains, it is easy to hypothesize that AFAP could be involved in many different biological functions which are linked to the actin cytoskeleton. Through the analysis of the modular domains of AFAP, it can be seen that the actin binding domain in the C-terminus allows it to associate with actin filaments, and the special structure of N-terminus of AFAP could bind the SH2 and/or SH3 domains of Src with high affinity. Mechanical stretch-induced deformation of cytoskeleton may present AFAP (along the stress fiber) to c-Src (in the cytoplasm). This interaction has the potential to lead to the translocation of Src to the actin cytoskeleton and activation of c-Src. Subsequent downstream tyrosine phosphorylation of other substrate proteins completes the conversion of the physical force to a biochemical signal (Han et al., 2004).

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AFAP was first discovered through research directed at discovering the mechanism by which constitutively activated v-Src could induce morphological transformations in cells. AFAP was found to form a stable complex with v-Src through its SH2 or SH3 domain binding motifs. The observation that AFAP was normally associated with stress fibers and with rosette-like structures in v-Src transformed chicken embryonic fibroblast cells indicated that AFAP may be involved in Src activity-dependent alteration of actin filaments (Baisden et al., 2001a; Kanner et al., 1991). The avian form of AFAP was first cloned by Flynn et al. who reported the sequence (Flynn et al., 1993). The mammalian form was first cloned in rats (Lodyga et al., 2002) and later in humans (Han et al., 2004). The mammalian sequences have a >80% homology with the avian form (Han et al., 2004; Lodyga et al., 2002). Importantly, the putative Src binding and activation motifs and actin binding domain described above are well conserved from avian to mammalian forms (Han et al., 2004). It has been shown that AFAP can interact with c-Src through both its SH2 and SH3 domain binding motifs. Mutation of five putative tyrosine phosphorylation sites or a single amino acid mutation from proline to alanine in the SH3 domain biding motif significantly reduced the ability of AFAP to bind to c-Src and to lead to its activation. Han et al. further over-expressed these mutant forms of AFAP as dominant negative inhibitors and demonstrated these mutants blocked stretch-induced c-Src activation in multiple cell types (Han et al., 2004). Based on these results, Han et al. have proposed that deformation of the cytoskeleton increases the competitive binding between AFAP and c-Src SH2 and SH3 domains. The interaction between AFAP and c-Src leads to a conformational change in Src, facilitating its activation. Activation of Src allows for the initiation of a downstream signaling cascade (Han et al., 2005) (Fig. 11.3).

11.7 Src PKT Activation by Multiple Physical Forces In multiple models of mechanotransduction, Src activation has been found to be one of the early upstream signals (Han et al., 2005). Whether using models of cell stretch in fibroblasts (Sai et al., 1999) or endothelial cells (Naruse et al., 1998), pressure overload in vivo (Franchini et al., 2000), or sheer stress (Okuda et al., 1999), Src tyrosine kinase activation has been detected by the applied physical force. Src PTK activation is associated with mechanotransduction in lung tissue (Liu et al., unpublished observation). Parker et al. found that when they inhibited protein tyrosine phosphotase activity, there was an increase in susceptibility to VILI resulting from a high peak airway inflation pressure in mechanically ventilated rat lungs, whereas inhibition of protein tyrosine kinase reduced ventilation-induced lung injury (Parker et al., 1998). One of the particularly damaging features of VILI is the breakdown of epithelial and endothelial barriers. Barrier disruption leads to an increase in vascular permeability allowing proteins and fluid to leak from circulation into the alveolar space

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ultimately leading to edema formation and the interruption of gas exchange across the air-blood barrier. It has been shown that after only 20 min of injurious ventilation, vascular permeability is significantly increased in isolated, perfused mouse lungs, much sooner than expected if this was the result of cytokine influence alone. These results implied a rapid mechanism for mechanotransduction leading to lung injury independent of cytokine response (Miyahara et al., 2007). The force associated with mechanical ventilation is not limited to the airways of the lung. This force is also transmitted to the vasculature, where a different type of mechanical force is experienced. During normal breathing the airway and alveolar pressure remains within an accepted range, however during mechanical ventilation, the airway and alveolar pressure increase during inspiration to facilitate air flow (von Bethmann et al., 1998). As the airway and alveolar pressure increases from mechanical ventilation, the lung vasculature is stretched causing vessel narrowing and an increase in pulmonary circulation pressure. This increase in pressure may increase shear stress, a force parallel to the plane of reference, which is known inducer of mechanotransduction in endothelial cells (Uhlig, 2002). Observations from studies directed at determining the role of β-catenin in the development of increases in vascular permeability have implicated Src PTKs as upstream modifiers of this process. Normally found at adherens junctions, β-catenin can be tyrosine phosphorylated by Src, a process leading to the re-distribution of β-catenin to the cytoplasm where it is degraded, resulting in an increase in vascular permeability, possibly thorough the destabilization of the adherens junction (Mehta and Malik, 2006). The PI3K-Akt signaling pathway has also been shown to act on β-catenin. Activation of this signaling pathway can lead to the negative regulation of GSK3β, which when active, can phosphorylate β-catenin on serine residues, tagging it for degradation. PI3K signaling, however, can also activate Src. Depending on the strength of these signaling pathways, PI3K signaling could be both a positive and negative regulator of vascular permeability (Miyahara et al., 2007). PI3K is a diverse kinase that is activated in response to many different stimuli (Uhlig and Uhlig, 2004). Mechanical ventilation of isolated, perfused mouse lungs leads to the activation of PI3K. Inhibition of PI3K during mechanical ventilation leads to a decrease in vascular permeability and a similar effect is seen when Src is inhibited directly. Interestingly, inhibition of Akt, a downstream kinase of PI3K, lead to the augmentation of VILI induced vascular permeability. Further study revealed that mechanical ventilation induced activation of PI3K signaling may lead to a two pronged signaling cascade. One prong activates Src leading to the removal of β-catenin from adherens junctions and an increase in vascular permeability. The other prong activates the Akt-GSK3β signaling pathway leading to the preservation of β-catenin at the adherens junction and the maintenance of endothelial barrier function (Miyahara et al., 2007). It appears that during high pressure or injurious mechanical ventilation, activation of Src is stronger than Akt signaling leading to VILI. A recent paper by Tzima et al. provides a very comprehensive overview of endothelial mechanotransduction (Tzima et al., 2005). They focused on investigating how integrins became activated in response to shear stress. Through a series of

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inhibitor experiments, they were able to show that the Src-PI3K-integrin pathway was responsible for the intracellular changes associated with shear force. It is known that shear stress induces a conformational change in integrins and cell alignment with the direction of flow, as well as the activation of NF-κB (Tzima et al., 2002; Tzima et al., 2001). The addition of a PI3K inhibitor (LY294002 or wortmannin) is able to block integrin activation. PI3K itself has been found to become rapidly activated in response to mechanical force (as measured through phosphorylation of the p85 regulatory subunit of PI3K, the increase of PtdIns(3,4,5)P3 and kinase activity (Tzima et al., 2005). Src is also known to become active within seconds of the onset of shear stress (Okuda et al., 1999; Pampori et al., 1999), even faster than PI3K. Src inhibition with PP2 or SU6656 prevented p85 phosphorylation as well as integrin activation (Tzima et al., 2005). To determine what exactly is the mechanosensor in shear stress-induced mechanotransduction, endothelial cells lacking VE-cadherin or PECAM-1 (VE-cadherin−/− and PECAM-1−/− respectively) were tested for their ability to respond to shear stress. Both types of cells were unable to undergo integrin activation in response to shear stress. When VE-cadherin and PECAM-1 were ectopically reexpressed in these cells, normal functionality was restored. Further studies demonstrated that PECAM-1 is upstream of VE-cadherin and is required for Src activation, and VE-cadherin acts as an adaptor protein to link Src to PI3K allowing for the Src mediated activation of the PI3K signal cascade (Tzima et al., 2005).

11.8 Blocking Src PTK as a Potential Therapy for VILI By investigating the role of protein-protein interactions in mechanotransduction, we hope to find some novel strategies to treat or prevent VILI. This is an important goal to strive for because with the limitations of technology available to us today, mechanical ventilation is unavoidable for patients with ALI/ARDS, and sometimes a reduction in tidal volume to reduce mechanical strain while maintaining acceptable blood gas parameters is unattainable (Villar et al., 2004). Activation of Src PTK has been implicated in acute inflammatory responses and tissue injuries in multiple organs, including the lung (Okutani et al., 2006). Therefore, although mechanotransduction in the lung may be activated by multiple mechanisms, the mechanical forces induced Src PTK activation may represent a common pathway that leads to acute lung injury. It has been shown that Src PTK activity can be involved in many of the processes that we associate with lung injury including cytokine and chemokine production, immune cell recruitment, and increased vascular permeability (Okutani et al., 2006). Indeed Src PTK inhibition has been shown to be protective in animal models of stroke (Paul et al., 2001), myocardial infarction (Weis et al., 2004), and ALI (Khadaroo et al., 2004; Severgnini et al., 2005). In multiple models of mechanotransduction, Src activity was found to be one of the upstream events in a defined signal cascade that responds to a mechanical force (Han et al., 2005; Pawson,

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2007; Sawada et al., 2006). As mentioned above, Src is believed to be directly activated by AFAP in response to mechanical stretch (Han et al., 2004) and is thought to phosphorylate p130Cas on its YXXP repeats when it is unfolded in response to mechanical force (Pawson, 2007; Sawada et al., 2006), it is also one of the first signaling proteins activated in shear stress induced mechanostransduction in endothelial cells (Tzima et al., 2005). Therefore, Src PTKs are an attractive target for the treatment of VILI. The role of protein-protein interactions in mechanotransduction is likely not limited to adaptor proteins. Interactions between proteins and other molecules, such as membrane lipids, calcium and other ions, RNA and DNA molecules may also be affected by physical forces. At every level of the signal cascade, there is a complex and largely unknown mechanism for the interaction of signaling molecules. These interactions may occur locally in a particular sub-cellular compartment to determine a specific cellular function; they may also be affected further by other factors cells are exposed, such as hyperoxia, hypoxia, inflammatory mediators and environmental conditions, etc. To further explore the role of protein-protein interaction and interaction between proteins and other molecules will improve our understanding of mechanosensitivity and mechanotransduction under physiological and pathophysiological conditions and reveal new targets for clinical therapies, such as VILI in ALI/ARDS.

11.9 Conclusions and Perspective VILI is a complex problem that will likely require a multifaceted approach to solve including work from basic scientists, clinicians, and even engineers. Interestingly there are research groups currently investigating ways to remove or limit the use of mechanical ventilation in the treatment of patients with ALI/ARDS and replacing it with a modified extracorporeal membranous oxygenator (Fischer et al., 2006). If successful, this technology will eliminate the need to apply external force to injured lungs, as all the blood gas exchange could occur outside of the body. Unfortunately, early trials of such treatment have indicated several critical limitations on this technology making it unsuitable for some of the most critically ill patients. Based on the current standard of this treatment, it does not appear that the technology to completely remove mechanical ventilation from the treatment of critically ill patients will be available in the near future. With this in mind, it is as important as ever continue to strive to understand how mechanical ventilation leads to lung injury through various mechanotransduction mechanisms and look for novel strategies to manipulate these pathways to benefit patients. Based on the information in this chapter, we propose that protein-protein interactions play an important role in intracellular mechanotransduction. We believe that the mechanisms presented are physiologically relevant and are part of an ongoing process that contributes to lung injury in mechanically ventilated patients. It should be noted that missing from this review are several other types of

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mechanotransduction mechanisms which may be involved in the development of VILI such as ion flux or lateral intercellular space compression (Tschumperlin et al., 2004) that were omitted to maintain a focus on what we believe to be a relatively novel and understudied mechanism. The idea that an adaptor protein can respond to physical force and initiate a signal cascade is an exciting one. It allows for the identification of the initial step in the mechanotransduction pathway that is interesting not only from a bioengineering standpoint, but will hopefully reveal therapeutic targets for future treatment of VILI. We have emphasized that Src activation appears to be a common theme in the mechanotransduction pathways that have been described and propose that intervention directed at dampening Src activity could be therapeutically beneficial. Although this is an exciting thought, the body of literature to draw upon in the field of protein-protein interactions in VILI limited, meaning that there is still much work to be done before we are able to fully understand this process lead alone make the transition from the bench to the bedside.

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Part V

Mechanosensing and Mechanotransduction in Bone and Joint Tissues

Chapter 12

Cellular Mechanisms of Mechanotransduction in Bone Suzanne R.L. Young and Fredrick M. Pavalko

Abstract Bone is a dynamic tissue that adjusts its structure over time to adapt to changes in mechanical load. This adaptive ability is critical to skeletal development and maintenance of optimal skeletal health throughout life. Imbalances in the ability of bone to keep pace with demands placed on it by mechanical loading results in bone that is fragile and susceptible to fracture. The cells within bone, i.e. osteocytes and osteoblasts, are responsible for detecting and responding to mechanical loading. Understanding how bone cells sense and respond to mechanical signals (the process known as mechanotransduction) has been the focus of considerable research. Despite excellent progress over the past two decades, the precise mechanisms by which bone cells perceive and mediate proper responses to mechanical loading are not fully understood. When bone is loaded, movement of interstitial fluid is generated within the small spaces that surround bone cells. This movement generates fluid shear stress (FSS) that stimulates bone cells, resulting in enhanced anabolic activity. Most research on cellular mechanisms of mechanotransduction is now focused on understanding the mechanisms through which bone cells sense FSS and generate a proper biochemical response. Mechanisms of cellular mechanotransduction likely involve the adhesive junctions, integrins, mechanically-sensitive ion channels, purinergic receptors, gap junctions and primary cilia. The integrin family of cell adhesion molecules may, as their name implies, “integrate” mechanical signals across the cell surface. It is also likely that ion channels, particularly calcium changes, play a critical role in mediating mechanical responses to FSS. This review will focus on recent progress in understanding how bone cells detect and respond to mechanical stimuli using novel multi-protein signaling complexes called “mechanosomes”. Keywords Fluid shear stress · Integrins · Focal adhesion · Primary cilia · Mechanosome F.M. Pavalko (B) Department of Cellular and Integrative Physiology, Indiana University School of Medicine, 635 Barnhill Drive, MS 346A, Indianapolis, IN 46202, USA e-mail: [email protected] A. Kamkin, I. Kiseleva (eds.), Mechanosensitivity and Mechanotransduction, Mechanosensitivity in Cells and Tissues 4, DOI 10.1007/978-90-481-9881-8_12, C Springer Science+Business Media B.V. 2011

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S.R.L. Yong and F.M. Pavalko

12.1 Introduction Cellular mechanotransduction is the process by which cells sense and respond to physical cues from their surrounding environment. This mechanical sensitivity is evident from the simplest isolated cells on two-dimensional substrates to cells integrated within the most complex tissues. Understanding the biological processes that mediate cellular mechanotransduction has been a growing area of interest during the past 20 years and is particularly important in understanding how bone responds to mechanical load (Allori et al., 2008; Chen et al., 2010; Huang et al., 2004; Papachristou et al., 2009; Papachroni et al., 2009; Robling and Turner, 2009a). Bone is an extremely dynamic tissue that normally undergoes constant turnover, a process that defines bone remodeling (Turner and Pavalko, 1998). The mechanical responsiveness of the cells that reside in bone, including osteocytes, the most abundant cells, and osteoblasts, the cells that secrete new osteoid, has been a primary focus in efforts to understand how bone turnover is regulated in response to the mechanical demands placed on the skeleton (Bonewald, 2006; Bonewald and Johnson, 2008; Turner et al., 1994, 2009). This article will discuss recent progress in the identification and characterization of molecules at the bone cell membrane that are responsible for detecting mechanical signals. We then discuss progress in identifying intracellular signaling pathways that are responsible for propagating those signals inside the cell and regulate bone cell function.

12.2 Detection of Mechanical Stimuli 12.2.1 Focal Adhesions and the Mechanosome Hypothesis Focal adhesions are sites of attachment between the cell and the extracellular matrix and have been proposed as cell mechanosensors. Focal adhesions appear to be integral for the detection and propagation of mechanical signals across the plasma membrane and into the cytoplasm and nucleus. Direct and indirect connections between the extracellular matrix (ECM), integrin cell adhesion molecules, and focal adhesion-associated signal transduction molecules play an important role in mechanotransduction in many types of cells, including bone cells. Each of these three key elements (matrix, integrins and intracellular signaling molecules) converges at these sites of structural attachment between cells and the ECM. Although there is strong evidence that focal adhesions are involved in mechanical signaling, the molecular mechanisms through which mechanical signaling via focal adhesions occurs is not completely understood. Focal adhesions are characterized by the presence of integrin cell adhesion molecules that link the ECM outside the cell to the cytoskeleton and signaling molecules inside the cell (Burridge and ChrzanowskaWodnicka, 1996). Focal adhesions serve as important sites of structural attachment between cells and the ECM (Burridge and Chrzanowska-Wodnicka, 1996), however, focal adhesions also function as important organizing centers for signal

12

Cellular Mechanisms of Mechanotransduction in Bone

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transduction activity (Alahari et al., 2002; Burridge and Chrzanowska-Wodnicka, 1996). Importantly, signaling proteins associated with focal adhesions are also found in the nucleus (Carvalho et al., 2003; Cattaruzza et al., 2004; Ogawa et al., 2003; Yi et al., 2003). This has led us to propose the concept of the mechanosome (Pavalko et al., 2003) in which proteins that can associate with focal adhesions respond to mechanical stimuli by participating in signaling events leading to alteration of transcriptional activity in the nucleus. The mechanosome hypothesis states that the release of molecules from adhesion complexes following mechanical loading alters transcriptional activity regulating genes that control osteoblast function. The mechanosome should be considered as a multi-protein complex comprised of adhesion-associated and DNA-binding proteins that, upon translocation to the nucleus in response to mechanical loading, transfers mechanical information from adhesion complexes to target genes. The protein complexes we describe as mechanosomes are not specific to osteoblasts, or even bone, but may be particularly well-suited to mediate mechanotransduction in this tissue. Additional reports showing that proteins associated with focal adhesions can also be present in the nucleus include focal adhesion kinase (FAK) (Lobo and Zachary, 2000; Stewart et al., 2002), NMP-4/CIZ (Feister et al., 2000; Nakamoto et al., 2000), p130cas and zyxin (Nix et al., 2001). Evidence that integrins are activated by fluid shear stress (FSS) is compelling and comes both from direct demonstration of conformational changes in integrin structure (Tzima et al., 2001) and from inhibition studies using integrin-specific inhibitory antibodies and Arg-Gly-Asp (RGD) peptides (Ponik and Pavalko, 2004). Using antibodies that recognize conformational changes in integrin structure associated with integrin-ligand affinity changes, Tzima et al. (2001) showed increased immunostaining of endothelial cells subjected to FSS indicating a modulation of αvβ3 integrin affinity by shear. Similarly, Jalali et al. (2001) using antibodies that specifically recognize the high affinity binding state of β1 and β3 containing integrin heterodimers have shown that FSS also causes conformational changes of these integrins in endothelial cells. In complimentary studies in endothelial cells, others have inhibited integrin function using blocking antibodies and RGD peptides to inhibit FSS induced activation of intracellular signaling pathways and cell motility (Urbich et al., 2002). These include blockade of shear induced signaling by extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK) and the IκB complex (Labrador et al., 2003). Similarly, Liu et al. (2002) used integrin function blocking strategies to inhibit shear induced activation of key transcription factors that regulate sterol and lipid homeostasis called sterol regulatory element-binding proteins (SREBPs). Shear induced secretion of basic fibroblast growth factor (bFGF) was also blocked by anti-integrin antibodies (Gloe et al., 2002) as were the anti-apoptotic effects of fluid shear (Urbich et al., 2000). Data from our laboratory demonstrates that inhibition of integrin-ECM interactions using RGD peptides inhibits both shear induced up-regulation of cyclooxygenase-2 (COX-2) protein and shear induced release of PGE2 from cells (Ponik and Pavalko, 2004). Previously we found that osteoblasts rapidly (

Mechanosensitivity and Mechanotransduction (Mechanosensitivity in Cells and Tissues)

Mechanosensitivity of the Heart (Mechanosensitivity in Cells and Tissues)

Mechanosensitivity of the Nervous System (Mechanosensitivity in Cells and Tissues)

Oral Cells and Tissues

ANIMAL CELLS AND TISSUES

Cells, tissues, and skin

Plant Cells And Tissues (The Green World)

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Mast Cells and Basophils

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Cardiac Mechanotransduction (Medical Intelligence Unit)

Adhesive Interactions in Normal and Transformed Cells

Adipose Tissues

Proteases in Gastrointestinal Tissues (Proteases in Biology and Disease)

Endothelial Cells in Health and Disease

Stem Cells in Development and Disease

Taurine 4 - Taurine and Excitable Tissues

Mast Cells Methods and Protocols

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Immobilization Of Enzymes And Cells

Solar Cells and Their Applications

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Molecular Evolution and Genetic Defects of Teeth - Cells, Tissues, Organs Vol 186 No 1

Macrophages and Dendritic Cells Methods and Protocols

Osmotic and Ionic Regulation: Cells and Animals

Mechanosensitivity and Mechanotransduction (Mechanosensitivity in Cells and Tissues)

Mechanosensitivity of the Heart (Mechanosensitivity in Cells and Tissues)

Mechanosensitivity of the Nervous System (Mechanosensitivity in Cells and Tissues)

Oral Cells and Tissues

ANIMAL CELLS AND TISSUES

Cells, tissues, and skin

Plant Cells And Tissues (The Green World)

Stem Cells and Cancer

Immobilized Enzymes and Cells

Esau's Plant Anatomy: Meristems, Cells, and Tissues of the Plant Body: Their Structure, Function, and Development

Immobilized Enzymes and Cells

Mast Cells and Basophils

Stem Cells and Cancer

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Cardiac Mechanotransduction (Medical Intelligence Unit)

Adhesive Interactions in Normal and Transformed Cells

Adipose Tissues

Proteases in Gastrointestinal Tissues (Proteases in Biology and Disease)

Endothelial Cells in Health and Disease

Stem Cells in Development and Disease

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Mast Cells Methods and Protocols

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