Mechanical Forces and the Endothelium
The Endothelial Cell Research Series A series of significant reviews of basic and clinical research related to the endothelium. Edited by Gabor M.Rubanyi, Berlex Biosciences, Richmond, California.
Volume One Endothelium-Derived Hyperpolarizing Factor edited by Paul M.Vanhoutte Volume Two Endothelial Modulation of Cardiac Function edited by Malcolm J.Lewis and Ajay M.Shah Volume Three Estrogen and the Vessel Wall edited by Gabor M.Rubanyi and Raymond Kauffman Volume Four Modern Visualisation of Endothelium edited by J.M.Polak Volume Five Pathophysiology and Clinical Applications of Nitric Oxide edited by Gabor M.Rubanyi Volume Six Mechanical Forces and the Endothelium edited by Peter I.Lelkes Volumes in Preparation Morphogenesis of Endothelium W.Risau Vascular Endothelium in Human Physiology and Pathophysiology P.Vallance and D.Webb
This book is part of a series. The publisher will accept continuation orders which may be cancelled at any time and which provide for automatic billing and shipping of each title in the series upon publication. Please write for details.
Mechanical Forces and the Endothelium
Edited by
Peter I.Lelkes University of Wisconsin Medical School Sinai Samaritan Medical Center Milwaukee USA
harwood academic publishers Australia • Canada • China • France • Germany • India • Japan Luxembourg • Malaysia • The Netherlands • Russia • Singapore Switzerland
This edition published in the Taylor & Francis e-Library, 2004. Copyright © 1999 OPA (Overseas Publishers Association) N.V. Published by license under the Harwood Academic Publishers imprint, part of The Gordon and Breach Publishing Group. All rights reserved. No part of this book may be reproduced or utilized in any form or by any means, electronic or mechanical, including photocopying and recording, or by any information storage or retrieval system, without permission in writing from the publisher. Amsteldijk 166 1st Floor 1079 LH Amsterdam The Netherlands
British Library Cataloguing in Publication Data Mechanical forces and the endothelium.—(Endothelial cell research series; v. 6) 1. Endothelium—Mechanical properties I. Lelkes, Peter I. 611'.0187 ISBN 0-203-30384-9 Master e-book ISBN
ISBN 0-203-34317-4 (Adobe eReader Format) ISBN 90-5702-447-0 (Print Edition) ISSN 1384-1270
CONTENTS
Foreword Michael A.Gimbrone, Jr.
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Preface
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Contributors
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1 The Hemodynamic Environment of Endothelium In Vivo and its Simulation In Vitro Mark M.Samet and Peter I.Lelkes 1 2 Chloride Channels in Endothelium: The Role of Mechano-stimulation and Changes in Cell Volume Bernd Nilius, Jan Eggermont, Thomas Voets and Guy Droogmans
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3 Tyrosine Phosphorylation of Platelet Endothelial Cell Adhesion Molecule-1 (PECAM-1) and Mechanosignal Transduction Keigi Fujiwara, Michitaka Masuda, Masaki Osawa, Noboru Harada and Rosangela Bruno Lopes
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4 Protein Phosphorylation in Shear Stress Activated Endothelial Cells John Y-J.Shyy, Yi-Shuan Li, Song Li, Shila Jalali, Michael Kim, Shunichi Usami and Shu Chien
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5 In Vitro Simulation of Shear Stress and Mitogen-activated Protein Kinase Responses to Shear Stress in Endothelial Cells Oren Traub, Chen Yan and Bradford C.Berk
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6 Flow-induced Endothelial Gene Regulation Joji Ando, Risa Korenaga and Akira Kamiya
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7 Endothelial Gene Regulation by Fluid Shear Forces Nitzan Resnick, Efrat Wolfovitz and Shahar Zilberstein
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8 Shear Stress Mediated Gene Regulation Susan M.McCormick and Larry V.McIntire
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9 Flow-induced Endothelial Cell Activation and Gene Regulation by Mechanical Forces 189 Eugene A.Sprague, Antonio J.Cayatte, Robert M.Nerem and Sumathy Mohan 10 Hemodynamics and Endothelial Phenotype: New Insights into the Modulation of Vascular Gene Expression by Fluid Mechanical Stimuli James N.Topper and Michael A.Gimbrone, Jr.
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11 Differential Regulation of Endothelial Cell Surface Molecules by Diverse Hemodynamic Forces Peter I.Lelkes
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12 Endothelium and Cyclic Strain Ira Mills and Bauer E.Sumpio
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13 Effects of Hydrostatic Pressure on Endothelial Cells Eric A.Schwartz, Mary E.Gerritsen and Rena Bizios
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14 The Role of Hemodynamic and Mechanical Factors in Vascular Growth and Remodeling Olga Hudlicka, Margaret D.Brown and Stuart Egginton
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Index
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FOREWORD
This timely volume highlights an area of intensive investigation at the interface of the fields of vascular biology and biomedical engineering—the effects of mechanical forces generated by blood flow on the vascular endothelium. A central premise of modern vascular biology is that the endothelium, the gossamer-like lining of the cardiovascular system, is a dynamically mutable interface that is locally responsive to various biochemical stimuli delivered by the circulating blood or generated by neighboring cells and tissues. Early studies of the mechanisms underlying this plasticity of endothelial phenotype identified certain proinflammatory substances, such as cytokines and bacterial products, as important humoral stimuli, acting via receptors and intracellular signalling pathways, to ‘activate’ the endothelial cell. The biological consequences of this activation process (which often reflected genetic regulation) were found to be multiple and diverse, including enhanced macromolecular permeability, altered adhesivity for circulating blood cells, and changes in the balance of various endothelial products important in hemostasis and thrombosis, growth regulation and vascular reactivity. In addition to this form of biochemical stimulation, the endothelial lining is constantly subjected to various mechanical forces—shear stresses, cyclic strains, hydrostatic pressures—generated by pulsatile blood flow. There is increasing awareness that these biomechanical stimuli can also directly influence endothelial structure and function, both acutely and chronically, thus constituting a novel paradigm of endothelial activation. Although systematic experimentation in this fascinating area spans less than two decades, impressive strides have been made in elucidating the cellular and molecular mechanisms involved in biomechanical activation of the endothelium. Initially, this investigative process resembled the parable of the blind men and the elephant, with different groups of scientists attempting to dissect this complex phenomenon, each from their respective viewpoints and each using the specialized tools of their discipline. Cell biologists noted flow-induced cell shape changes and probed the underlying cytoskeletal events with electron microscopy and immunocytochemistry. Cell physiologists applied biophysical probes to measure ion fluxes and intracellular biochemical changes, while pharmacologists reached for inhibitors to dissect the relevant pathways involved. Circulatory physiologists catalogued changes in a remarkable array of secreted biological factors whose activities have important implications for cardiovascular function. Cardiovascular pathologists called attention to the remarkable correlation of certain vascular disease processes such as atherosclerosis with specific arterial geometries, and fluid mechanical engineers attempted to model the relevant hemodynamic parameters involved. Taken together vii
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these various initiatives have provided an impressive descriptive fund of knowledge about the overall response of the endothelium to hemodynamic forces, and the potential implications of this type of stimulation for cardiovascular physiology and pathophysiology. We now have a clear picture of the endothelial cell being impacted upon by various components of its biomechanical environment, and, as a consequence, undergoing a dramatic modulation in its functional phenotype. As detailed in several chapters of this volume, this discovery process has recently taken a quantum leap forward with the application of the tools of modern molecular biology. The level of experimental analysis has moved rapidly from secreted protein product, to cytoplasmic mRNA content, and then on to genetic regulatory events within the nuclear compartment. The response of individual ‘candidate genes’, important to specific aspects of endothelial biology and pathobiology, have been analyzed in detail. Dissection of their respective promoter regions has led to the characterization of ‘shear-stress-response elements’ that can interact with specific transcription factors in the initiation of gene expression. In parallel, various signal transduction pathways that appear to link externally applied mechanical forces to these nuclear events have been identified, and, in certain instances, their functional importance has been established by selective inhibition and/or overexpression of critical components. Recently, state-of-the-art techniques for comprehensive analysis of patterns of gene expression have begun to be applied to the problem. This strategy, which attempts to take a ‘snapshot’ of all the genes that are being expressed by an endothelial cell in response to a defined set of stimuli, has yielded a number of interesting insights. It appears that a given endothelial cell can respond to a particular biochemical stimulus by up-and/or down-regulating a set of genes, and that the functional relationship of the products of these structurally dissimilar genes may have important implications in a given pathophysiological context. For example, steady laminar shear stress stimulation of cultured human endothelial cells, analyzed by high-throughput differential display, results in the sustained upregulation of a subset of genes (including cyclooxygenase-2, the endothelial isoform of nitric oxide synthase, manganese-dependent superoxide dismutase) whose products and activities (e.g., prostacyclin, nitric oxide, oxidant stress resistance) can exert a ‘vaso-protective’ effect. This observation may help to explain the resistance of certain arterial geometries that are typically associated with uniform laminar flow to atherosclerotic lesion development. This strategy has also led to the identification of certain novel human genes whose expression appears to be dependent upon continuous biomechanical stimulation, thus suggesting that hemodynamic forces can function as ‘differentiation stimuli’ in vivo. This phenomenon of ‘flow-dependent genes’ has important implications for normal vascular development, physiological vascular remodeling, and pathophysiological responses to injury. Finally, the observation that different types of biomechanical stimulation (e.g., laminar versus turbulent flow, high and low hydrostatic pressures, various regimens of cyclic stretching) can elicit different patterns of endothelial gene regulation implies complexity at the level of sensing, transducing and coupling to downstream effector mechanisms. Dissection of the molecular mechanisms of this fascinating phenomenon, at the level of primary biomechanical sensors and their associated signalling pathways, is currently an area of active investigation.
Foreword
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Notwithstanding the remarkable progress that has been made, as documented in the contributed chapters of this volume, certain caveats need to be recognized. First, the majority of the in vitro studies to date have made use of simplified model systems, designed to isolate a single component of the complex biomechanical environment of the endothelial cell in vivo. Furthermore, this experimental mechanical stimulation usually represents an abrupt transition from the typically static (non-dynamic) cell culture situation, rather than a quasi-steady state condition. Second, these in vitro models take the endothelial cell out of its usual biological context, thus depriving it of input from neighboring cells (e.g., smooth muscle, pericytes) and extracellular matrix components. While this reductionist approach can be useful, ultimately there is a need to test the results obtained in a more integrated context. Practically, such validation can be sought either via the development of more complex in vitro models (mechanical and/or cellular), or through the demonstration of similar flow-dependent phenomena in an in vivo setting. Third, it will be important to assess the relative contributions of biomechanical stimuli, in conjunction with various humoral stimuli, in the maintenance and modulation of endothelial phenotype in a given biological context. The identification of selective markers of biomechanical activation of the endothelium, that can be detected in both in vitro models and various in vivo settings, should greatly facilitate this effort. Further development and application of high-throughput technologies for comprehensive phenotypic profiling, at either the mRNA or protein level, could conceivably allow this evaluation to be performed on individual endothelial cells in various defined flow environments in vitro or in different vascular geometries in vivo. A more complete appreciation of how biomechanical stimuli act to modify endothelial phenotype clearly would augment our understanding of vascular development and differentiation, and potentially the pathogenesis of vascular diseases such as hypertension and atherosclerosis. Beyond blood vessels per se, such knowledge may also provide insight into the role of mechanical forces in the integrative physiology and pathophysiology of highly vascularized organs, such as lung, heart, kidney and brain, in which the interaction between vessels and parenchyma are so important. Ultimately, it will also be of interest to systematically compare and contrast the molecular regulatory mechanisms that govern the responses of the vascular endothelial cell to the mechanical forces encountered in its in vivo environment with those of other biomechanically responsive cells in their respective tissue settings (e.g., osteoblasts and osteoclasts in bone, myocardiocytes in heart, airway epithelial cells in lung, etc.). How the endothelial cell, in the complex hemodynamic environment of the cardiovascular system, adaptively responds to mechanical stimulation remains a fascinating question—one that should continue to offer challenges to vascular biologists and biomedical engineers for several years to come. Michael A.Gimbrone, Jr.
PREFACE
For the better part of the past 30 years, experiments with isolated endothelial cells in conventional, ‘static’ tissue culture led to the discovery of the multifaceted properties of these fascinating cells, and defused the notion that the endothelium is merely ‘porous cellophane’ acting as a passive barrier between blood and tissue. The development of ‘dynamic’ cell culture chambers permitted the study of endothelial cells exposed to mechanical forces, i.e. under conditions which more realistically mimic the hemodynamic environment of the endothelium in vivo. The aim of this book is to summarize our current state of knowledge of Mechanical Forces and the Endothelium and to point at new areas to be explored in the near future. Obviously, a monograph like this one serves as a progress report and can only render a static, ‘still-life’ like impression of the fast-paced advances in this dynamic field. The complex subject matter is literally very much in flux. As this book goes to press, new discoveries are being published in rapid sequence, adding more sophistication to many of the issues discussed in the 14 chapters of this book. To set the stage, Michael Gimbrone introduces the subject by providing a historical perspective and looking into the future of using genomics-based studies for the discovery of novel genes affected by hemodynamic forces. Clearly, the long-term goal of our studies is to gain a detailed understanding of the molecular genetics of how endothelial cells perceive and transduce the stimuli derived from their hemodynamic environment, in particular of pathophysiological alterations therein. We are optimistic that such detailed knowledge will eventually lead to the identification of novel therapeutic modalities in treating and/or preventing prevalent cardiovascular diseases which, according to our recent understanding, are primarily affected by hemodynamic aberrations, such as locally disturbed flow patterns and/or hypertension. Samet and Lelkes describe in the first chapter the hemodynamic environment of the endothelium in vivo in health and disease and introduce some of the model systems developed for studying the effects of the distinct hemodynamic/mechanical forces, i.e. fluid shear stress, cyclic strain and elevated pressure, on cultured endothelial cells in vitro. During the past decade, analysis of hemodynamic forces has focused largely on the effects of laminar flow-induced shear stress. This, the simplest of all flow fields represents, however, the pattern of blood flow in only a subset of blood vessels under physiological conditions. While the literature abounds with reports on endothelial cell responses to laminar flow, significantly fewer publications are devoted to studying the effects of pulsatile, non-laminar, non-steady, or perturbed flow as encountered under pathophysiological conditions. Similarly, the role of other hemodynamic forces, xi
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caused by the pulsatile nature of blood flow, e.g. of cyclic strain, or of hydrostatic pressure is less well understood. In keeping with the preponderance of data derived from studying cultured endothelial cells in a simplified flow field, more than half of the chapters in this book describe, at the cellular and molecular level, the regulation of endothelial cell gene expression by laminar fluid shear stress. The overall picture emerging from these studies suggests that laminar flow-induced shear stress acts as a potent, distinct (and unique?) extracellular stimulant which ‘activates’ endothelial cells in a manner reminiscent of the receptor-mediated activation of cells following exposure to specific mitogens. While a flow-specific sensor/receptor remains, as yet, elusive, exposure to fluid shear stress leads to a cascade of intracellular signaling, which involves activation of ion channels (Nilius et al.), tyrosine phosphorylation of trans-membranous sensors (Fujiwara et al.), activation of well-known phosphorylation-based second messenger pathways (Shyy et al., Traub et al.), as well as of distinct ‘shear response elements’ (SSREs) in the promoter regions of shear-sensitive genes, which contain consensus binding sites for previously described nuclear transcription factors (Ando et al., Resnick et al.). This cascade of flow-induced signalization and nuclear translocation of diverse transcription factors leads to a distinct pattern of up- or down-regulation of numerous, shear sensitive genes and gene products, such as vasomodulatory compounds and cell adhesion molecules (McCormick and McIntire, Sprague et al.). It is well established that exposure of endothelial cells to cytokines and growth factors, which trigger some of the same signaling pathways as fluid shear stress (and also other mechanical forces), results in cellular ‘activation’, e.g. induction of proliferation and migration, upregulation of adhesion molecules, and expression of a prothrombotic phenotype. Remarkably, however, laminar flow-induced shear stress appears to have vasoprotective effects resulting in cellular ‘passivation’ and expression of a quiescent endothelial phenotype, reminiscent of the intact, healthy endothelium in vivo. As such, the study of endothelial cells exposed to laminar shear stress continues to provide a valuable paradigm for understanding ‘normal’ vascular physiology. In addition this system might serve as a suitable model for exploring the complex integration of unique extracellular stimuli which, although utilizing seemingly similar signaling pathways, result in diverse cellular responses. In extending the studies of the cellular effects of flow-induced shear stress to include more realistic representations of non-laminar and turbulent flow, more recent data suggest that gene-induction in endothelial cells is exquisitely sensitive to specific flow patterns (Topper and Gimbrone). Indeed, the finding that particular genes, which are down-regulated by laminar flow, are upregulated by perturbed flow (and also by other mechanical forces) stresses the importance of aberrant flow patterns in the etiology of cardiovascular pathophysiology (Lelkes). The ‘activation’ of endothelial cells by cyclic mechanical strain bears certain similarities to that by fluid-shear stress. Amongst the notable differences, however, is the fact that, to date, no unique ‘cyclic strain response element’ similar to a SSRE has been identified (Mills and Sumpio). Similarly, exposure of endothelial cells to (steady) hydrostatic pressure leads to their activation, as assessed for example by the release of growth factors (Schwartz et al.). Presumably, in the real world in vivo, pathophysiological modulations of all three components of vascular hemodynamic
Preface
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theory play a concerted role in the establishment and progression of vascular diseases. To approach this problem experimentally, novel, more sophisticated model systems will be required, which combine the individual actions of all three mechanical components of flowing blood. These systems will help to further substantiate the hemodynamic theory of the origin of numerous vascular diseases which postulates that the cumulative effect of complex mechanical forces plays a pivotal role in the initiation and progression of focal vasculopathies. The impressive progress achieved in vitro ultimately has to be corroborated and validated in vivo. The final chapter by Hudlicka et al. provides a fascinating glimpse into the important role of hemodynamic forces on organ-specific vascular growth/ angiogenesis and vascular remodeling. Clearly, more studies are needed to elucidate in greater molecular detail the pivotal role of hemodynamic forces in the establishment and maintenance of the vasculature during embryogenesis and under pathophysiological conditions. This book could not have been completed without the individual chapters by expert authors. I am grateful to all of them for accepting the challenge, interrupting their busy schedules and generously contributing their time and expertise. I’d like to acknowledge the assistance of Harwood Academic Publisher’s staff for patiently guiding me through the various phases of this project. I would also like to thank Ms Mary Simon for excellent secretarial support. This book is dedicated to my family: my children, Tamar, Efrat, Yphtach and Nadav, and above all to my wife, Iris, who for many years have endured and supported my ‘crazy’ schedules with patience, good humor and lots of love.
CONTRIBUTORS
Joji Ando Department of Biomedical Engineering Graduate School of Medicine University of Tokyo Kongo 7–3–1 Bunkyo-ku Tokyo 113 Japan
Antonio J.Cayatte Vascular Biology Department of Medicine Boston University 80 Concord Street Boston, MA USA
Bradford C.Berk Centre for Cardiovascular Research Box 679 University of Rochester Rochester, NY 14642 USA
Shu Chien Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA
Rena Bizios Department of Biomedical Engineering Rensselaer Polytechnic Institute Troy, NY 12180–3590 USA
Guy Droogmans Department of Physiology Campus Gasthuisberg KU Leuven B-3000 Leuven Belgium
Margaret D.Brown School of Sport and Exercise Sciences University of Birmingham Edgbaston Birmingham B15 2TT UK
Jan Eggermont Department of Physiology Campus Gasthuisberg KU Leuven B-3000 Leuven Belgium
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Stuart Egginton Department of Physiology University of Birmingham Edgbaston Birmingham B15 2TT UK
Shila Jalali Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA
Keigi Fujiwara Department of Structural Analysis National Cardiovascular Center Research Institute Suita, Osaka 565–8565 Japan
Akira Kamiya Department of Biomedical Engineering Graduate School of Medicine University of Tokyo Kongo 7–3–1, Bunkyo-ku Tokyo 113 Japan
Mary E.Gerritsen Genetech Inc. Building 10 10NA Way South San Francisco CA 94080 USA Michael A.Gimbrone, Jr. Vascular Research Division Department of Pathology Brigham and Women’s Hospital Harvard Medical School 221 Longwood Avenue, LMRC-401 Boston, MA 02115–5817 USA Noboru Harada Department of Structural Analysis National Cardiovascular Center Research Institute Suita, Osaka 565–8565 Japan Olga Hudlicka Department of Physiology University of Birmingham Edgbaston Birmingham B15 2TT UK
Michael Kim Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA Risa Korenaga Department of Biomedical Engineering Graduate School of Medicine University of Tokyo Kongo 7–3–1, Bunkyo-ku Tokyo 113 Japan Peter I.Lelkes Laboratory of Cell Biology Department of Medicine University of Wisconsin Medical School Milwaukee Clinical Campus Sinai Samaritan Medical Center P.O. Box 342 Milwaukee, WI 53201–0342 USA
Contributors
Yi-Shuan Li Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA Song Li Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA Rosangela Bruno Lopes Department of Structural Analysis National Cardiovascular Center Research Institute Suita, Osaka 565–8565 Japan Michitaka Masuda Department of Structural Analysis National Cardiovascular Center Research Institute Suita, Osaka 565–8565 Japan Susan M.McCormick Cox Laboratory for Biomedical Engineering Rice University P.O. Box 1892 Houston, TX 77251–1892 USA Larry V.McIntire Cox Laboratory for Biomedical Engineering Rice University P.O. Box 1892 Houston, TX 77251–1892 USA Ira Mills Department of Surgery
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Yale University School of Medicine 333 Cedar Street P.O. Box 208062 New Haven, CT 06510 USA Sumathy Mohan Department of Radiology University of Texas Health Science Center at San Antonio 7703 Floyd Curl Drive San Antonio, TX 78284–7800 USA Robert M.Nerem Institute for Bioengineering and Bioscience Georgia Institute of Technology Atlanta, GA 30332–0363 USA Bernd Nilius Department of Physiology Campus Gasthuisberg KU Leuven B-3000 Leuven Belgium Masaki Osawa Department of Structural Analysis National Cardiovascular Center Research Institute Suita, Osaka 565–8565 Japan Nitzan Resnick Department of Morphological Sciences Bruce Rappaport Medical Research Institute The Rappaport Faculty of Medicine-Technion P.O. Box 9697 Bat-Galim, Haifa Israel 31096
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Contributors
Mark M.Samet Laboratory of Cell Biology Department of Medicine University of Wisconsin Medical School Milwaukee Clinical Campus Sinai Samaritan Medical Center P.O. Box 342 Milwaukee, WI 53201–0342 USA Eric A.Schwartz Department of Biomedical Engineering Rensselaer Polytechnic Institute Troy, NY 12180–3590 USA John Y-J.Shyy Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA Eugene A.Sprague Department of Radiology University of Texas Health Science Center at San Antonio 7703 Floyd Curl Drive San Antonio, TX 78284–7800 USA Bauer E.Sumpio Department of Surgery Yale University School of Medicine 333 Cedar Street P.O. Box 208062 New Haven, CT 06510 USA James N.Topper Cardiovascular Division Department of Medicine Stanford University School of Medicine Falk Cardiovascular Research Center 300 Pasteur Drive Stanford, CA 94305–5406 USA
Oren Traub Department of Medicine University of Washington Seattle, WA 98195–7710 USA Shunichi Usami Department of Bioengineering and Institute for Biomedical Engineering University of California, San Diego La Jolla, CA 92093–0412 USA Thomas Voets Department of Physiology Campus Gasthuisberg KU Leuven B-3000 Leuven Belgium Efrat Wolfovitz Department of Morphological Sciences Bruce Rappaport Medical Research Institute The Rappaport Faculty of MedicineTechnion P.O. Box 9697 Bat-Galim, Haifa Israel 31096 Chen Yan Department of Pathology University of Washington Seattle, WA 98195–7710 USA Shahar Zilberstein Department of Morphological Sciences Bruce Rappaport Medical Research Institute The Rappaport Faculty of MedicineTechnion P.O. Box 9697 Bat-Galim, Haifa Israel 31096
1 The Hemodynamic Environment of Endothelium In Vivo and its Simulation In Vitro Mark M.Samet* and Peter I.Lelkes Laboratory of Cell Biology, Department of Medicine, University of Wisconsin Medical School, Sinai Samaritan Medical Center, P.O. Box 342, Milwaukee, WI 53201–0342, USA, Tel.: (414) 219–7753, Fax: (414) 219–7874. *Corresponding author: E-mail address:
[email protected].
INTRODUCTION The cardiovascular system transports oxygen and nutrients to all tissues and removes carbon dioxide and waste products of metabolism by continuously circulating its working fluid, blood, through an elaborate hydraulic network of large and small vessels. Blood flow in humans is neither turbulent nor completely laminar. In the arteries the flow is pulsatile and intermittently accompanied by small disturbances, but in the vicinity of curvatures (bends) and branches, where secondary flows are generated, the patterns are unsteady and complex. By contrast, blood flow in veins is generally quasi-steady. Flowing blood continuously exerts mechanical forces on the vascular wall and the vessels adapt to this hemodynamic environment accordingly. At physiological levels, blood-imposed forces play an important role in maintaining normal biology of vascular wall cells. By contrast, in aberrant situations the hemodynamic challenges result in abnormal cellular responses that may lead to various vasculopathies. The endothelial cells lining the entire vasculature are in direct contact with the flowing blood and, thereby, constantly exposed to its mechanically imposed forces, such as traction and pressure. Our understanding of the blood flow phenomena is fairly good. With proper approximations and basic physical principles we can quantitatively delineate the hemodynamic environment that affects the endothelium. However, as of yet, our state of knowledge of endothelial response to the bloodimposed mechanical challenges is incomplete and there are important questions awaiting to be answered. In addressing the unknowns, the cell-expressed biochemical clues are sought via in vitro experiments that are aimed at mimicking the hemodynamic environment of the cells. In the laboratory setup, this environment includes three types of mechanical effectors: flow-induced shear stress, elevated pressure and stretching/straining of the underlying substrate. 1
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The purpose of this chapter is to outline the fluid mechanical aspects associated with blood flow and the resulting forces acting on the vascular wall, in vivo. With this information in mind, we then examine some of the available experimental systems in vitro as they attempt to mimic select aspects of the hemodynamic conditions perceived by the endothelium.
BASIC FLUID MECHANICS CONCEPTS Hemodynamics is concerned with the physical principles that govern the flow of blood within the cardiovascular system. These principles are adapted from the general laws of fluid dynamics, with some modifications that are dictated by the complex properties of the vasculature and blood. Blood vessels are essentially circular pipes whose impermeable wall is composed of cells and cell products; blood is an incompressible liquid. When a liquid flows through a pipe its motion is resisted by the continuous action of viscous forces. The magnitude of these forces is maximal at the wall and it diminishes as the distance from the boundary, y, increases. Because of the viscous resistance, liquid particles are at rest at the wall (no-slip condition) and their velocity, u, increases towards the center of the pipe, as the resistance to their motion decreases. The radial difference in velocities shears the flow, with the rate of shear (velocity gradient) being maximal at the wall and diminishing as the distance from the boundary increases. Because of the velocity gradient, du/dy, work must be done to overcome the viscous resistance between adjacent layers of liquid particles that slide over each other. A measure of this work is represented by the tangential force per unit area , or shear stress. In the case of a liquid obeying Newton’s law of viscosity (Newtonian liquid), the shear stress is linearly related to the rate of shear: (1)
where µ, the dynamic viscosity, is a property of the liquid itself. By contrast, in a nonNewtonian liquid the strict requirement of linearity, as expressed by equation 1, does not hold. The distinction between Newtonian or non-Newtonian traits is particularly useful when describing blood behavior in different flow conditions. Flows can be classified in many ways such as, uniform, non-uniform; steady, unsteady; laminar or turbulent. When a body of liquid moves in a wall bounded space, the conditions within the liquid can vary from one point to another and, at any given point, from one instant to the next. If, at a given moment, the velocity of liquid particles is the same in magnitude and direction at every spacial location, the flow can be described as uniform. Otherwise, the flow is non-uniform. The latter category has relevance to hemodynamics because, in vivo, the size and shape of the crosssection of the blood stream as it passes through the vasculature is not constant. Also of relevance to hemodynamics is the temporal behavior of flow. If the velocity and pressure of the stream change from one spacial location to another but do not vary with time, the flow can be described as time independent or steady. By
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contradistinction, if, at a given point, flow conditions do change with time, the flow is termed unsteady. In vivo, the flow of blood in the cardiovascular system is time dependent: On the time scale of years, aging and diseases alter the pace of the heart and also introduce structural changes in the vasculature that manifest in continuous, albeit slow, alterations in blood circulation. On the time scale of seconds and minutes, the temporal dependency is more apparent because the flow in the arteries ceaselessly cycles between the systole and diastole of the heart. Additionally, due to posture changes and movement of the body there are always slight variations in pressure and velocity of the blood in the veins, hence the flow is only quasi-steady. Thus, the concept of steady flow in its strict sense does not apply to blood flow in the cardiovascular system but, as an idealization, it may provide a foundation for a simplified analysis of the circulation. The motion of a viscous liquid through a wall bounded space (like in a circular pipe), can be smooth or, if the conditions warrant, disorderly. If all liquid particles move deterministically in distinct and traceable trajectories that are tangent to the direction of flow, the flow can be described as laminar. Alternatively, laminar flow can be characterized by layers or stratums of liquid that smoothly slide one over the other. When laminar flow becomes destabilized and the motion of liquid particles transits from orderly to vortical and chaotic, the flow can be described as turbulent. Most naturally occurring flows are not laminar, primarily because their inertia force is much higher than the viscous resistance. A similar consideration also applies to blood flow in the cardiovascular system. The relative importance of liquid inertia and liquid viscous resistance in a given vessel is defined by a dimensionless parameter called Reynolds number (Re). In a pipe flow, the Reynolds number is given by: (2)
where U is the average velocity of the flow, D is the diameter of the pipe, and and µ are liquid density and dynamic viscosity, respectively (Tritton 1988). Since the average velocity in a circular pipe is given by: (3)
where Q is the volumetric rate of flow, the Re number can be conveniently expressed as: (4)
with v=µ/ denoting the kinematic viscosity of the liquid. Thus, in pipe flow, the value of Re increases proportionally to increase in flow rate, and vice versa. As implied above, Re scales the transition between laminar and turbulent flows; for Re>2000, the flow streams disorderly and becomes turbulent. On the other hand, when Re«2000, the flow is definitely laminar. From the information available for human circulation,
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the estimated Reynolds number in straight segments of veins and arteries is well below 2000 (Guyton, 1981), but it may reach values of 5000–12000 in the root of the aorta and the pulmonary artery during the rapid phase of blood ejection by the ventricles (Milnor, 1980b). By contrast, blood flow in the micro-circulation is characterized by very low Re numbers, typically of the order of 10-2, or less (Skalak, Ozkaya and Skalak, 1989), (Goldsmith and Karino, 1988). Consequently, blood flow in the cardiovascular system covers a wide domain of Re values in which the viscous forces prevail on one end and the inertia forces dominate on the other end. In many locations in the cardiovascular system, blood vessels bend or curve significantly. A typical example of such a configuration is the aortic arch, which makes almost a 180° turn as it connects between the ascending and descending segments of the aorta. When liquid flows in a curved pipe it is forced to accelerate centripetally as well as flow downstream. As a result, a secondary flow is established in radial planes just as the outward-driven liquid returns back along the pipe wall. The secondary flow consists of two counter-rotating vortices which are “superimposed” on the axial stream, causing the liquid particles to coil in a spiral path. This spiraling phenomenon occurs in both laminar and turbulent flows and becomes even more intricate when the flow is unsteady due to pulsatility or vibrations of the bounding wall. The characteristics of the secondary flow depend on the relative importance of centrifugal forces and liquid viscous resistance in a given vessel. This ratio is defined by a dimensionless parameter called Dean number (Dn) (Pedley, 1980): (5)
where d is the diameter of the curvature. For Dn<10, the secondary flow is negligible and the profile of the main stream is almost identical to the shape noted in straight pipe flow. By contrast, for 96
500 has much relevance to pathological hemodynamics because, in vivo, the resulting highly skewed velocity profiles and/or transient, local flow separations have been implicated in the initiation and exacerbation of certain vascular diseases, for example, atherosclerosis (Skalak, Ozkaya and Skalak, 1989). Blood flow in arteries is unsteady and its time-varying characteristics are governed by the pulsatile output of the heart. For practical purposes the pulsatile flow can be regarded as a stream of fluid, Q(t), consisting of a fast propagating periodic pulse, which oscillates between maximum systolic and minimum diastolic values, and a more slowly moving bulk of the steady flow, : (6)
The Hemodynamic Environment of Endothelium
In a more rigorous approach, proposed by Womersley (1957), the periodic term subjected to harmonic analysis using a Fourier series in the form:
5
is
(7)
In this series, each term is a sinusoidal wave of amplitude qi, phase i, and angular frequency i. The first harmonic, (i=1), represents the fundamental pulsation frequency of the flow and, for arterial flow, it corresponds to the rate of heart beats, . The relative importance of the pulsation forces and viscous resistance in a given vessel is defined by a dimensionless parameter called Womersley number (Wo), where: (8)
When Wo«1, viscous forces dominate. The velocity profile assumes a parabolic shape as in the case of steady pipe flow, and the centerline velocity varies periodically in the same phase as the driving pressure gradient (Uchida, 1956), (Womersley, 1957). By contrast, for Wo>10, pulsating forces dominate and the velocity profile assumes a flat shape while lagging by 90° behind the driving pressure gradient (Uchida, 1956; Womersley, 1957). Interestingly, in most large arteries in human vasculature the estimated Wo values range from 1 to about 10; only in the aorta Wo>10.
AN OVERVIEW OF THE CARDIOVASCULAR SYSTEM Conceptually, the cardiovascular system is a closed, pressurized hydraulic circuit consisting of two networks, systemic and pulmonary, that are connected in series to each other. As a self-contained unit, this hydraulic circuit includes four functional elements, viz. a working fluid, a pump, vessels and control, that operate continuously and in concert with one another. The working fluid in the cardiovascular system is blood. Blood is a viscous suspension of plasma and cells. Plasma occupies about 55–58% of blood total volume and contains close to 90% (v/v) water, 7% (w/v) organic solutes (proteins), and 1% (w/v) electrolytes (Conley, 1980). Plasma includes a large variety of proteins, among which the three most prominent types are albumin, globulins and fibrinogen. The presence of these macromolecules in the plasma makes it a slightly heavier liquid than water. Thus, plasma solution of healthy individuals has an average specific gravity of 1.026 at 37°C. This value remains virtually constant within the limited variations in body temperature, such as during fever, but it may vary to a considerable degree following excessive blood loss or in chronic renal failure. The viscosity of plasma is generally a function of temperature and local rate of shear. However, at the magnitudes of flow and the rates of shear normally found in the vascular circuitry, the plasma at 37°C may be considered a Newtonian fluid with viscosity values ranging from 0.012 to 0.016 Poise.
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The cellular composition of blood includes erythrocytes (RBCs), white cells and platelets. About 99% of the corpuscles are erythrocytes and, on the average, they occupy 40–45% of blood total volume. The other cell types take up less than 1% of blood total volume and due to their small number and size they are not important to the fluid mechanics of blood. Nonetheless, these corpuscles play a central role in blood clotting and in combating inflammatory processes. The RBCs, on the other hand, are dynamically important. They are highly flexible biconcave discoids with diameter size of 8 µm and thickness of about 2 µm. Also, as formed elements, the RBCs are slightly heavier than plasma and have a specific gravity of 1.093 (Conley, 1980). Because these corpuscles are so small, and the difference in densities between plasma and erythrocytes is less than 6.5%, the cells can track very closely the motion of blood and maintain it as a homogeneous continuum. Indeed, the continuum hypothesis works very well in flows through large and medium-sized blood vessels but is clearly invalid in capillaries where the diameter of the small vessels approaches the size of the corpuscles. Since the topic of blood flow through small vessels is beyond the scope of this work, flowing blood will be regarded from hereon as a homogeneous continua with specific gravity of 1.06 (for reviews on microcirculation and capillary flows see Zweifach, Ozkaya and Secomb, 1986; Skalak, 1986; Secomb, 1995 and Intaglietta, 1996). Whole blood is in essence a non-Newtonian fluid and its apparent viscosity depends on the volume of packed red blood cells in the specimen (hematocrit) and the rate of shear. The dependence on hematocrit is highly nonlinear and can be represented by an upward concaving curve. Typically, in an healthy individual with total volume of blood of 5.5 liters and hematocrit of about 42% (38% in women), the viscosity of whole blood is 4-times higher than that of water. However, in a hematologic disorder such as polycythemia, the hematocrit increases to more than 53% and the viscosity of blood becomes 6-times higher than that of water (Erslev and Gabuzda, 1979). The dependence of whole blood viscosity on the rate of shear is by far more complex. Constant-shear viscometer studies reported by Whitmore (1968) indicate that the apparent viscosity becomes independent of shear rates for values above 100 sec-1. By contrast, Merrill and Pelletier (1967) report that below the shear rate of 20 sec-1 blood behaves like a shear thinning fluid in which a certain yield stress must be exceeded in order to make the blood flow. Since the estimated mean wall shear rates in human vasculature range from about 200–300 sec-1 (in large arteries and veins) up to >1000 sec-1 (in vessels of 0.5–1.0 mm in diameter), (Chien, 1972; Goldsmith and Karino, 1988), the blood flowing in large vessels (>0.5 mm) can be regarded as a Newtonian fluid. The applicability of this assumption, though, remains questionable near the center of blood vessels or in areas of flow separation/recirculation, where the average rate of shear is certainly less than 100 sec-1. Conceivably, blood does not remain Newtonian in arteries throughout the entire pulsation cycle, but Kunz and Coulter (1967) suggest that this bi-periodic, non-Newtonian effect can be considered negligible. Nonetheless, for all practical purposes, blood can be regarded as an homogeneous Newtonian continua of apparent viscosity 0.04 Poise at 37°C (Ku, 1997) and kinematic viscosity of 4 mm2/sec. The heart is responsible for pumping blood through the entire cardiovascular system. The power required from the heart to pump the blood is estimated at 33
The Hemodynamic Environment of Endothelium
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calories/day (Zamir, 1977). Considering a typical cardiac efficiency of 20–25% (Guyton, 1981), this power consumption amounts to about 6% of the total metabolic rate for an average man at rest (Zamir, 1977). Of course, during exercise the heart works much harder and the consumption of energy is much higher, but, typically, it does not exceed the level of 17% of total metabolic rate. All in all, given a normal regimen of rest and activity, the heart beats about 5– 10 billion times over a period of 70 years. These self-didactic estimates are indicative of the unique structure and mode of operation of the heart that make it, energy-wise, a highly cost-effective flow-propelling machine. The heart is composed of four chambers, two atria and two ventricles. The two atria are relatively elastic, and act as reservoirs for the blood draining from the systemic and pulmonary veins. They also weakly propel blood into the ventricles. The ventricles are more muscular and provide the main force that ejects the blood into the outflow vessels. Although both ventricles eject almost the same amounts of blood into their respective circulations, they differ significantly in shape and their mode of operation. The left ventricular shape is that of a prolate spheroid, which, during ejection, contracts along the minor axes while maintaining the major axis constant (Pedley, 1980). The right ventricle, on the other hand, is wrapped around one side of the left ventricle and resembles a semi-lunar cavity. During ejection, the outer wall moves downwards along with the open orifice (Pedley, 1980), permitting the blood to exit at a pressure that is about 25% of its systemic counterpart (Mountcastle, 1980). The heart operates in a cyclic fashion. Each cardiac cycle includes a period of filling (diastole) and a period of ejection (systole). The filling period consists of three phases. During the first phase, blood accumulated in the atrium flows rapidly into the ventricle upon opening of the mitral (tricuspid, on the right side of the heart) valve. The entering stream reinforces vortical motion within the chamber which encompasses the entire lumen. During the middle third of diastole, the blood entering the atrium from the veins continues directly into the ventricle and contributes to the development of secondary flow. These two phases of diastole account for about 70% of the filling volume. The final third phase occurs when atrial pumping propels the remaining 30% of the filling volume into the lower chamber (Guyton, 1981). It is likely that the weak pumping helps energize the already existing vortex and, thereby, assists in smooth closing of the mitral (tricuspid) valve at the end of diastole. This description of ventricular flow is consistent with the cineradiographic studies of Taylor and Wade (1973) which portray the flow patterns in the left and right ventricles as stable, rather than laminar, with little evidence of gross mixing or occurrence of large scale disturbances. Furthermore, their data also show no indication of flow reversal at the time of valve closure. Consequently, these authors attribute the stability of the ventricular flow to the curvature and compliance of the cardiac wall. The period of systole begins about 0.04–0.05 seconds before the actual discharge of ventricular blood takes place (Mountcastle, 1980). During this time, which starts immediately after closure of the mitral (tricuspid) valve and ends just before the aortic (pulmonic) valve opens, the pressure within the ventricle rises slightly above that in the aorta (or pulmonary artery). The build up of pressure in left ventricle
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results from increase in tension in the ventricular muscle along with some shortening in the apex-to-base length (Guyton, 1981; Pedley, 1980). However, this decrease in length is compensated by a slight expansion in the ventricular cross-section, so that the lumenal volume of the ventricle remains unchanged (isometric) (Pedley, 1980). As the isometric pressurization ends, the motion of blood within the ventricle, including that of the vortex, becomes relatively small, but the change in ventricular shape promotes flow along the wall, thus protecting the endocardial endothelium from being subjected to complete stagnation. When the left and right ventricular pressures exceed 80 mm Hg and 8 mm Hg, respectively, the aortic and pulmonic valves open and blood is discharged into the respective arteries at a high Re number and a flat velocity profile (Pedley, 1980). The phase of ejection continues over the first three-fourth of ventricular systole with the valves completely open and aligned with the wall, to minimize obstruction to the flow. By the end of systole, as the muscles begin to relax, the ventricular pressure falls rapidly and the aortic and pulmonic valves close. This permits blood to fill the ventricles from about 60 ml at the bottom of systole to a peak volume of about 130 ml at the bottom of diastole. Normally the heart ejects about 70 ml with each beat at a rate of 72 beats/min. However, the output volume can increase by as much as 1.5 fold, and the heart rate can increase by as much as three fold during exercise; the maximal pace, though, is limited by age. The blood vessels forward blood under high pressure from the heart to all tissues and transport it back under much lower pressure from the tissues to the heart. Their unique structure and mechanical properties play an important role in maintaining normal hemodynamics in health, but these properties are also regarded as key contributors to formation of pathophysiological conditions associated with localized deviations from normal blood flow. By and large, arteries and veins of medium and larger caliber have the same wall structure, although the wall of veins is much thinner (Pedley, 1980; Monos, Berczi and Nadasy, 1995). Indeed, the thickness-to-diameter ratio in veins is of the order of 0.01, with vena cava bottoming at 0.006 and jugular vein topping at 0.015 (Pedley, 1980), whereas in systemic arteries this ratio is 9 fold (aorta) to 16 fold (femoral artery) higher (Ku and Zhu, 1993). Only in pulmonary circulation, the thickness-to-diameter ratio in arteries is comparable to that in the veins (Pedley, 1980). Despite the small thickness-to-diameter ratio, venous wall is very strong and it is capable of withstanding high pressure loads. For example, recent studies have determined the breaking pressure for saphenous vein as 56 lb/in2 or 2873 mm Hg (Monos, Berczi and Nadasy, 1995), which is about 24 times higher than the peak systolic pressure at normal heart output. Blood vessel walls are composed of three layers. The innermost layer, tunica interna (intima), consists of a monolayer of endothelial cells adhered to a self-assembled extracellular matrix, and an elastic lamina that separates the intima from the central layer (Griendling and Alexander, 1994), (Pedley, 1980; Dobrin, 1978). The central layer, media, is the thickest among the three layers, and consists of elastic laminae, bundles of collagen fibers, elastic fibrils and orderly arranged mantles of smooth muscle cells (Dobrin, 1978; Pedley, 1980; Mountcastle, 1980; Bergel, 1972). The outermost layer, adventitia, includes elastic and collagen fibers covered or intermingled
The Hemodynamic Environment of Endothelium
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with fibroblasts, smooth muscle cells, and nutrient vessels and nerves (Griendling and Alexander, 1994). Clearly, the specific makeup of the layers and their relative thickness depends on the type of blood vessel considered (vein or artery) and its anatomic location. In veins the adventitia is, generally, much thicker than the media and contains a more elaborated meshwork of collageneous material than elastin (typically at a ratio of 3:1) (Pedley, 1980). However, in veins experiencing high hydrostatic pressure, the media is as thick as the adventitia and rich in smooth muscle cells. Most of these vessels are also equipped with one way valves to prevent backflow of blood. By contrast to veins, the media in most arteries is the thickest layer of the wall and, via its composition, it also dominates the elastic behavior of the vessel. Accordingly, the medium and large arteries can be divided into two groups: conducting arteries which are elastic, and distributing arteries which are muscular. The conducting arteries, such as aorta, common carotid, and brachiocephalic artery serve as a pressure reservoir of the systemic network and their media contains more elastic fibers than collagen and smooth muscle cells to withstand high pressure and provide recoil; the smooth muscle cells within the layer help regulate vessel diameter. Distributing arteries, on the other hand, regulate blood inflow into limbs and organs, and their media contains more smooth muscle cells and collagen than elastic fibers. Consequently, the ratio of elastin-to-collagen in these vessels (including brachial, radial, splenic, and femoral arteries) is close to 0.5, as compared to 1.5 in the intrathoracic aorta (Pedley, 1980). Small arteries play a pivotal role in maintaining a nearly constant blood flow in the face of temporal changes in pressure. The media of these vessels contains almost exclusively smooth muscle cells that are arranged helically in several concentric laminae (Dobrin, 1978). The adventitia of the small arteries, albeit as thick as the media, is poor in elastin and collagen (Pedley, 1980) and, in most sites, it does not contribute much to the mechanics of the vessels. Thus, the diameter of the small arteries is almost exclusively determined by the state of contraction/relaxation of the smooth muscle cells. Further, the tonus of these cells is regulated in a paracrine fashion by the endothelium (Haller, 1997; Schiffrin, 1996; Griendling and Alexander, 1994), in accordance with the changing hemodynamic demands. At any given time the vasculature in a healthy individual circulates about 5–5.5 liters of blood. This volume is distributed between the pulmonary (11%) and systemic (82%) networks, and the heart and its related vessels (7%). Within the systemic circulation, the aorta and the arterial tree hold about 16% of blood volume, the capillaries contain about 4%, with the veins holding more than 60% of the volume (Johnson, 1992). These figures can vary significantly, particularly venous capacitance, which can alter, over a short period of time, by several tens of percent with relatively small changes in the diameter of central veins (Mountcastle, 1980). Indeed, venous capacitance is a potent modulator of cardiac output and, indirectly, also an important effector of the flow rate and fluid mechanics of blood in the arterial tree. The fluid mechanics of blood flow in the vasculature is complex (Pedley, 1980; Lucas, 1984; Pedley, 1995; Talbot and Berger, 1974) and involves pulsatility at high pressure in arteries (Nerem, 1981; Pedley, 1980; Ku, 1997; Pedley, 1995; Clark and Schultz, 1973), quasi-steadiness at low pressure in veins (Lemaire, 1978; Pedley, 1980; Moreno et al., 1970; Anliker, Wells and Ogden, 1969; Wexler et al., 1968) and various flow patterns in the vicinity of curvatures and bifurcations in both types
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Mark M.Samet * and Peter I.Lelkes
of blood vessels (Nerem, 1981; Caro, Parker and Doorly, 1995; Nakamura, Sugiyama and Haruna, 1993; Naruse and Tanishita, 1996; Pedley, 1995; Karino et al., 1987; Moore et al., 1992). Turbulence is rare in veins and has only been conjectured from non-invasive visualizations and temporal records of flow in diseased vessels. As the Re number in large veins ranges from about 150 to 2400 (Nerem, 1981), it is likely that this occurrence manifests disorders in blood flow due to lumen reduction or localized wall bound obstruction, as noted in, for example, varicosities, phlebothrombosis and thrombophlebitis, rather than break down of natural instabilities within the venous circulation. Venous flow, however, is not strictly laminar. In small veins the orthograde flow is accompanied by transient disturbances (most notably in the limbs) which become more numerous as the daughter vessels empty into larger, parent vessels. Eventually, the returning blood exhibits low-amplitude oscillatory characteristics (Nippa, Alexander and Folse, 1971), which amplify and become most prominent in the central vessels (Lemaire, 1978). These periodic disturbances proceed into the vena cava and influence its instantaneous velocity profiles, most notably in the abdominal section and at the entrance to the right atrium. In addition to the disturbances convected by the orthograde flow, venous circulation is also modulated by pressure waves that originate from venous pumping and the massaging action of skeletal muscles (Guyton, 1981). The transmission characteristics of these waves in vena cava have been studied extensively in the past (Anliker, Wells and Ogden, 1969; Anliker, Yates and Ogden, 1971; Pedley, 1980). Finally, blood flow in vena cava is also influenced by right atrial pressure pulses (Brawley et al., 1966) and by progressive changes in the elastic properties of the wall due to aging (Munari, 1967). Blood flow in arteries has been studied extensively over the years and the subject matter is now detailed in numerous publications aimed at biological and medical audience, applied mathematicians, physicists and bioengineers (Liepsch, 1990; O’Rourke, 1982; Alexander, Schlant and Fuster, 1998; Harcus and Adamson, 1975; Nichols and O’Rourke, 1998; Pedley, 1980; Nerem, 1981; Ku, 1997). Despite this wealth of information, the question as to whether the flow in arteries is normally disturbed or fully turbulent remains still unanswered. Generally, the velocity waveforms in the aorta lack high-frequency components from the start of systole until the end of the acceleration phase (Sbarbati-Del Guerra et al., 1996). This feature and the fact that the phase-locked averages of velocity signals resemble a “text-book” waveform (Nichols and O’Rourke, 1998) are indicative of the unperturbed nature of the flow during this period of the cardiac cycle. However, past the peak of systole and during the entire deceleration phase, the velocity waveforms contain high-frequency fluctuations associated with persistent disturbances in the flow (Nerem and Seed, 1972). The spectral analysis of these velocity fluctuations, which displays a wide and continuous range of frequencies from about 25 Hz up to 500 Hz, and the randomness and non-stationarity of the signal, lead various investigators (Nerem and Seed, 1972; Pedley, 1980) to the conclusion that blood flow during this period of the cardiac cycle is indeed turbulent. Also measurements in horse’s aorta (Nerem et al., 1974) and in a conscious human (Seed and Thomas, 1972), lend support to this notion. On the other hand, the occurrence of fully developed turbulence in arterial flow is challenged by the fact that neither the instantaneous velocity field nor the frequency spectrum of the
The Hemodynamic Environment of Endothelium
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fluctuations are the same as those in a steady, fully developed pipe flow (Laufer, 1954; Townsend, 1976). Hence, if the flow is not fully turbulent, it would appear to be three-dimensional and highly disturbed or in the transitional stages to turbulence. The latter characterizations also accord with the findings from dye visualization and MRI studies of the flow in models of the human abdominal aorta (Moore et al., 1994a; Moore et al., 1992). Whether highly disturbed, transitional, or turbulent, flow pulsatility plays an important role in defining the amplitude and direction of the hemodynamic forces exerted on the vessel wall.
HEMODYNAMIC FORCES ON VESSEL WALL Flowing blood subjects the wall of the vasculature to a force system consisting of two mutually orthogonal components: viscous friction and distension. The frictional force per unit area acts tangentially to the wall and is known as wall shear stress. The distending force per unit area (also known as normal stress) acts perpendicularly to the wall and is essentially equal to fluid pressure. The effects of these two hemodynamic forces are manifested both macroscopically as well as microscopically. Macroscopically, the shear stress and pressure stretch and distend the wall producing longitudinal, (albeit minor), and circumferential (or hoop) strains (Dobrin, 1978). The latter strains are mostly carried by the media and to a smaller extent by the subendothelial layer, whereas the former are predominantly constrained by the adventitia and vascular side branches (Dobrin, 1978; Kenner, 1972). On the microscopic level, the hemodynamic forces act directly on the individual endothelial cells shearing and pressing their luminal surface and, concomitantly, stretching their basement membrane. The endothelial cells, in turn, have the capability of sensing these hemodynamic challenges and adapting to their input, accordingly. Wall shear stress can be obtained directly from equation 1 by calculating the velocity gradient from experimentally derived velocity profiles. For example, using a MRI scanner, Moore et al. (1994b) made detailed velocity measurements in a model of human abdominal aorta from which they show that the wall shear stress in the suprarenal region averages at 1.32 dyn/cm2, but its temporal levels in a pulsatile flow can be as high as 8.38 dyn/cm2 and as low as -4.08 dyn/cm2, when <0 for 38±10% of the time. The same authors also report a mean shear stress value of 4.3 dyn/cm2 in the infrarenal region (except for the posterior wall where =-5 dyn/ cm2) and temporal levels of ranging from 10.4 dyn/cm2 down to -12.2 dyn/cm2 proximally to the iliac bifurcation. Unfortunately, direct measurements of velocity profiles in vessels smaller than the aorta are technically very difficult and, more often than not, the results obtained suffer from a considerable uncertainty associated with the spatial resolution of MRI or Doppler scanners, or misalignment of the hot-film sensor relative to the incoming velocity vector (Girard, Helmlinger and Nerem, 1993). Thus, most estimates of wall shear stress values in the cardiovascular system rely on theoretical considerations. The classical description of blood flow through straight segments in the vascular network is founded in the Hagen-Poiseuille flow model. The vessels are considered
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Mark M.Samet * and Peter I.Lelkes
rigid pipes and the liquid is assumed Newtonian. For steady, laminar and fully developed flow the velocity profile u(r) is a paraboloid (Tritton, 1988), given by: (9)
where r is the radial coordinate, and r=0 corresponds to the center of the pipe of diameter D. The rate of shear at the wall (using notation of equation 1) is: (10)
and the wall shear stress (from equation 1) is given by: (11)
where Q is the volumetric flow rate in the pipe. The magnitude of the wall shear stress is linearly proportional to the rate of blood flow but it is inversely proportional to the third power of vessel diameter. Thus, any increase in hemodynamic demand translates into a linear elevation in , whereas a narrowing in vessel lumen by, for example, pharmacological or pathophysiological events may lead to a significant increase in the shearing stress imposed on the endothelium, and vice versa. Tabulated below are exemplary values of the wall shear stress in select human blood vessels.
* ** *** **** ***** ******
(Mountcastle, 1980) (Nerem, 1981) (Wells, Archie and Kleinstreuer, 1996) (Zamir, 1977) (Moneta and Porter, 1995) (Pedley, 1980)
The trend noted from the table in the magnitude of shear stresses can be schematically described as bi-phasic. The lowest values of on the arterial and venous sides of the vascular network are obtained in vessels that are closest to the heart, and the levels increase consistently as the distance from the heart increases. This is consistent
The Hemodynamic Environment of Endothelium
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with the notion that most of the resistance to blood circulation comes from peripheral vessels of lumen diameter <0.4 mm (Mulvany, 1993; Schiffrin, 1996; Milnor, 1980a). Another point noted from the table is that the theoretically derived shear stress values are well below the critical magnitude of 400 dyn/cm2 that has been shown to cause an irreversible damage to the endothelial lining, even if applied only for a short time. For more details on this aspect, see Fry (1968). The Hagen-Poiseuille flow model fails to accurately describe the flow of blood in regions dominated by phenomena such as entrance effects, vessel curvature and flow unsteadiness. In most cases the flow is too complex for a theoretical analysis, and evaluation of the local wall shear stress requires lengthy numerical formats, which are only tractable by high speed computers. However, some simplified approaches are worth mentioning. In an entry flow, the flow field immediately downstream from the inlet consists of an inviscid core in the central portion of the pipe and a very thin boundary layer on the wall. The velocity profile in the core is flat and, as the flow proceeds downstream, the boundary layer thickens reducing the size of the core and accelerating the fluid at the center, so that the mass is conserved (Tritton, 1988). At a distance L from the inlet, termed entrance length, the boundary layer flow overtakes the entire cross-section and the flow becomes fully developed. The entrance length depends linearly on Re and is commonly expressed as (Nerem, 1981; Tritton, 1988): (12)
In the aorta (see the table above), for example, this distance translates to 182cm. Beyond the distance L, the flow conforms with the Hagen-Poiseuille model and the wall shear stress is given by equation 11. Yet, at axial distances shorter than L, 0<X
where (14)
and the shear stress at the wall is given by: (15)
which resembles the derivation provided by Schlichting (1987) for a boundary layer flow over a flat plate. Equations 13–15 give accurate results (±1–5%) for 0<=0.05
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Mark M.Samet * and Peter I.Lelkes
but may be stretched to about =0.08, if the accuracy sought is relaxed to ±15%. Using aorta as an example (see the table above), we obtain at 5cm distally to the inlet =0.06 and an inviscid-core velocity of Uc=40.91 cm/sec, which is consistent with the data published in Tritton (1988). The corresponding wall shear stress at this location is (X)wall=7.2 dyn/cm2, about 1.6 fold higher than the value obtained from equation 11. In curved vessels and bends the resistance to fluid passage is higher than in HagenPoiseuille flow due to the presence of secondary flow and its influence on the primary stream. The secondary flow forms under both laminar and turbulent conditions, but the effect of curvature is more pronounced at low and moderate Re numbers when laminar flow prevails. Most importantly, the wall shear stress in the curving vessel is no longer uniform and its magnitude varies along wall circumference. Thus, rather than determining the local shear stress value , it is common in engineering applications to represent the resistance to flow by the friction factor , where: (16)
and (-dP/dX) is the streamwise pressure gradient. In Hagen-Poiseuille flow the friction factor is: (17)
and the streamwise pressure gradient is linked to the local wall shear stress via: (18)
In curved vessels, the friction factor lc depends on the Dean number, Dn (equation 5). From the variety of formulae proposed over the years, the following appear to be most relevant to blood circulation: a) Dean’s formula for Dn<96 and D/d<0.0005 (Berger, Talbot and Yao, 1983)
(19) b) Ito’s formula for Dn>170 (Schlichting, 1987) (20)
c) Prandtl’s formula for 225
The Hemodynamic Environment of Endothelium
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all applying to laminar flow. At the onset of turhulence, the friction factor becomes independent of Dn and can be given by (Schlichting, 1987): (22) Flow unsteadiness associated with pulsatility has a significant impact on the local wall shear stress only when Wo»1; as mentioned above, at low Wo numbers the velocity profile assumes a parabolic shape (Uchida, 1956) and twall at any = constant can be estimated by equation 11. At large Wo numbers, however, the profile is no longer parabolic. For Wo»1, (according to Kenner (1972), Wo must be >5, but Dewey (1979) is more restrictive and requires Wo>10), the flow profile in the vicinity of the wall can be represented by: (23)
and the shear stress at the wall is given by: (24)
Using aorta as an example (see the table above) and a heart beat rate of 1 Hz, we obtain Wo=15.67 and ()wall=12.41 dyn/cm2. This result is about 2.8 fold higher than the value obtained from equation 11 and appears to be consistent with the experimental findings of Moore et al. (1994b). From the fluid dynamic point of view, the total blood pressure at any location in the vasculature is made up of four elements: ambient pressure, hydrostatic pressure, dynamic pressure, and potential energy pressure or pump head (Guyton, 1981). The ambient pressure, Po, is customarily referenced to the pressure in the right atrium and considered equal to the atmospheric pressure surrounding the body. However, for all practical purposes the value of Po is expressed in relative terms, namely, Po=0 mm Hg (Guyton, 1981). The hydrostatic pressure, Pg, results from the weight of blood and is given by: (25)
where g is gravity acceleration, h is the vertical distance from the tricuspid valve where Pg is conventionally taken as 0 mm Hg (Guyton, 1981). It follows that h<0 when measured downward, resulting in Pg>0. The dynamic pressure, Pd, results from the motion of blood and is given by: (26)
where u is the velocity of fluid at a given axial, X, and radial, r, locale. In arteries, u also depends on the time, t. However, in Hagen-Poiseuille flow model, u is only a function of r and the magnitude of Pd depends on the radial position in the vasculature.
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The potential energy pressure is introduced into the cardiovascular system by the heart. Because the heart continuously pumps blood in a pulsatile fashion into the aorta and the arterial tree, the potential energy pressure can be decomposed into a time-mean value, Ps, and a temporally varying pulse pressure, Pp, in which the amplitude equals, by definition, the difference between the systolic and diastolic values (Guyton, 1981). In the aorta, the time-mean pressure is approximately 100 mm Hg, it falls down to 95 mm Hg in the large arteries (where lumen diameter is ~0.3–0.5 cm), and decreases rapidly to 30 mm Hg at the entrance to the capillaries (Mountcastle, 1980; Guyton, 1981). At the beginning of the venous system the time-mean pressure is only 10–12 mm Hg, and decreases slowly along the network of veins, reaching the level of 0 mm Hg at the entrance to the right atrium (Mountcastle, 1980; Guyton, 1981). Like the magnitude of Ps, also the amplitude of Pp depends on the axial location along the arterial tree, (Pedley, 1980; Ku, 1997). Using the above definitions, the expression for total pressure in the blood, PT, can be written in the form: (27)
In this format, the second term on the right hand side of equation 27 represents the pressure in the blood due to the motion of the fluid, whereas the first term represents the actual pressure exerted by blood on the luminal surface of the vasculature. This term, commonly referred to as lateral pressure, is important because it determines the stretching and distension of the blood vessels. On the microscopic level, the lateral pressure determines the normal stress on the endothelial cells as well as the straining of their basement membrane. The lateral pressure exerted on the luminal surface of a blood vessel is balanced by the extravascular pressure, customarily approximated as Po, and by a stress system within the vascular wall which prevents it from rupturing. At any location in the three-dimensional space of the wall, this stress system is made up of nine, mutually perpendicular components: six shearing and three tensile (compressive) stresses. However, under normal conditions the radial, compressive stress is relatively small and for all practical purposes it can be neglected. Also the shearing stresses are negligible because the flow of blood within the vasculature occurs with almost no twisting of the vessels (Dobrin, 1978). Thus, in essence, the distension and stretching of the vascular wall is dominated by a two tensile stresses, axial and circumferential. Under static conditions and a given body posture the axial stress may be computed from: (28)
where (29)
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is the static transmural pressure and Do and Di represent the outer and inner diameters of the vessel, respectively. Comparably, the circumferential stress is computed from the law of Laplace for a finite size wall thickness, Do—Di: (30)
and the corresponding axial and circumferential wall strains are given by: (31)
(32)
where L and D stand for changes in vessel size from its unstressed length, L, and diameter, D, respectively. In vivo, the functional relationship between stress and strain is rather complex. Yet, it can be simplified greatly if the vessels are considered elastic and composed of an incompressible and isotropic material. Following Dobrin (1978), the axial and circumferential strains may be given by a set of two coupled equations: (33)
where E is the elastic modulus (equal in all directions), is Poisson’s ratio, and and represent changes in stress and strain, respectively, due to change in static loading (of transmural pressure) on vessel luminal surface. The latter may occur, for example, due to change in body posture or temporal movement of the limbs. Equation 33 can be simplified even further by assuming that the change in static loading does not alter the axial strain in tethered vessels, in vivo or in situ (Dobrin, 1978). With X=0, , assumes the form (Ku and Zhu, 1993): (34)
Interestingly, despite it simplistic portrayal of the mechanical properties of vascular wall, equation 34 gives accurate results at physiological transmural pressure loadings of 75–135 mm Hg. However, at much higher loadings the results become greatly overestimated. This is also true for pressures below 75 mm Hg, where the calculated results underestimate the actual values by about 20% (Dobrin, 1978). Thus, the usefulness of equation 34 appears to be limited to non-hypertensive, high-pressure loadings; in most cases, though, more adequate results may be obtained by employing the coupled system of equation 33. Analysis of vascular wall response to a time-dependent loading is, by far, more complicated. The passage of pressure pulse constantly changes the stresses within
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the vessel and brings into play the viscoelastic properties of the wall. These properties have been shown to strongly depend on the frequency of oscillations, as well as the anatomic origin of the artery considered (Bergel, 1972; Kenner, 1972; Dobrin, 1978; Pedley, 1980). Furthermore, considerable dependence has also been reported on age and hypertension and, presumably, on smooth muscle cell activation. In addressing the viscoelastic properties of the wall, a linear approximation is commonly employed in which the dynamic characteristics in the axial direction are decoupled from their circumferential counterparts. Each case is then separately represented by an analogue model, which consists of a parallel arrangement of a spring and a dashpot (Kenner, 1972). Accordingly, for a given discrete pulsation frequency, , the ratio between viscous damping and elastic moduli is given by (Kenner, 1972): (35)
where and ED are, respectively, the viscous damping and the dynamic elasticity coefficients of vascular tissue, and is the phase angle between transmural pressure and vessel diameter. In large arteries the change in vessel diameter lags behind the pressure pulse by 5° <10° (Dobrin, 1978) implying that the viscous damping is about 8–25% of ED. Yet, as mentioned above, the magnitude of ED depends on the artery examined and on the frequency of pulsations. Indeed, for frequencies below 2 Hz, the magnitude of ED decreases in a frequency dependent fashion and approaches its static value as approaches 0. However, above 2 Hz the magnitude of ED is virtually frequency independent and exceeds the static value by a factor of 1.1–1.7 (Bergel, 1972; Dobrin, 1978; Pedley, 1980). This information, and similar, can be used to analytically formulate the dynamic straining of arteries in response to a timedependent loading (Kenner, 1972; Pedley, 1980). During cardiac cycle the arteries exhibit variations of about 1% in their length which is indicative of constraints imposed on these vessels by the perivascular connective tissue, by tethering and by presence of arterial branches. By contrast, the variations in vessel circumference is quite pronounced. Typically, in healthy human subjects the expansion in vessel diameter during systole is about 9–12% for the aorta, 6–10% for the pulmonary arteries, and 1–2% for the carotid and about 2–15% for femoral arteries. By contrast to these changes, the external diameter of these vessels does not alter by more than 8–10%, reflecting geometric differences between the luminal and outer surfaces and a possible restraining influence of the connective tissue.
IN VITRO SYSTEMS At any given locale, the mechanically-imposed forces of the flowing blood shear and press the lumenal surface of the endothelial cells and, concomitantly, stretch their basement membrane. The cells, in turn, sense the hemodynamic stimuli and respond to their challenge by secreting numerous bio/vaso active molecules, including prostacyclin and nitric oxide, that regulate endothelial biology in an autocrine and
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paracrine fashion. The coupling between stimulus and response and, in a broader sense, the classic link between hemodynamic forces and endothelial cell biology have been the subject of numerous studies. Most of these investigations have been conducted in vitro using apparatuses that were designed to expose cultured endothelial cells to a well defined hemodynamic environment of either flow-induced shear stress, or elevated pressure, or (cyclic) strain, viz. stretching/flexing of the underlying substrate. Three types of perfusion systems are commonly used to study the effects of shear stress on endothelial cells: small-caliber tubes, cone-and-plate viscometers and parallel plate flow chambers. Small-caliber tubes of constant internal diameter are the simplest, commercially available perfusion devices. The tubes can be readily cut into the desired length, sterilized in an autoclave, and utilized for cell culturing and superfusion (Eskin et al., 1984). Also, the setup and connection of this device to the mock-loop circulation are rather simple, and Poiseuille flow conditions can be obtained over a large portion of the vessel. For circular tubes, the Re number and the wall shear stress can be calculated directly from the flow rate using equations 4 and 11, respectively. Generally, small-caliber tubes successfully mimic straight segments of blood vessels devoid of branches or bifurcations. In the past, square glass tubes of 1 mm in diameter were employed to study changes in cell morphology in response to sustained exposure to steady laminar flow and wall shear stress of 34 dyn/cm2 (Eskin et al., 1984). In other studies (Kesler et al., 1986; Anderson et al., 1987; Sentissi et al., 1986), tubular devices, made of Dacron, polytetrafluoroethylene (PTFE), or expanded-PTFE have been used to evaluate retention of cells under steady flow conditions as part of a program aimed at developing endothelial cell-lined vascular prostheses. Most tubular devices are fabricated from materials that are poor cell culturing substrates: their luminal surfaces must be precoated with an adhesive protein such as fibronectin or collagen type I prior to cell seeding, to ensure proper cell attachment and spreading. Due to the small bore of the tubes and the axisymmetric shape of the luminal surface, attention must be paid to achieving uniform coverage of the surface with adhesive proteins as well as homogeneous distribution of the cells during inoculation. These tasks can be successfully accomplished by employing axial rotation of the treated vessel on a device similar to the designs currently used for seeding vascular grafts (Mazzucotelli et al., 1993). Cone-and-plate viscometers, also known as Couette flow devices, have been used extensively since 1981 (Dewey, Jr. et al., 1981) to impose a predetermined shear stress on cultured endothelial cells, and study its effect on cell phenotypic and functional traits. Figure 1.1 illustrates a typical layout of this apparatus. The system consists of a stationary flat plate, which is the bottom of a cylindrical container, and a rotating, inverted cone. The cone is fabricated from a non-cytotoxic material such as stainless steel (Dewey, Jr. et al., 1981), transparent polycarbonate (Franke et al., 1984), plexiglass (Malek et al., 1995), or transparent polymethylmethacrylate (Schnittler et al., 1993) and attached to a gear-train that is driven by a variable-speed motor. The cylindrical container is custom designed to either house a large diameter Petri dish, or snugly accommodate up to 12 circular coverslips lined with cells. Some designs also include inlet/outlet ports on the sidewall for replacement of culture medium during lengthy experiments (Dewey, Jr. et al., 1981). The less flexible designs permit only short-term experiments, or require complete disassembly of the system
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Figure 1.1. Schematics of a cone-and-plate apparatus.
for medium exchange. The container and its content are mounted on a X-Y-Z positioning stage, carefully centered with cone axis, and raised until the apex of the cone just contacts the plate. To ensure laminar flow, the included angle between cone surface and the flat plate is limited to 0.5° 3° (Dewey, Jr. et al., 1981; Franke et al., 1984; Schnittler et al., 1993; Slack and Turitto, 1994); turbulence, on the other hand, is induced at =5° (Davies et al., 1986). The shear stress imposed on the cells is produced by rotating the fluid contained between the cone and the plate at an angular speed O. At low fluid speeds and 3°, the velocity vector, and, therefore, the shear stress at the flat wall are unidirectional and their orientation is tangential to the rotating fluid. Also, the magnitude of wall shear stress is independent of the radial distance from the center, and it can be calculated from (Dewey, Jr. et al., 1981): (36)
When excessive rotation is imparted on the fluid, or if 5°, the flow in the gap becomes turbulent. The fluid mechanics of secondary flow and turbulence in a coneand-plate device have been elegantly elucidated by Spdougos, Bussolari and Dewey (1984). A limited account of cellular response to turbulent shear stress was presented by Davies et al. (1986). Finally, the use of a cone-and-plate apparatus in pulsatile flow and other time-varying conditions was addressed by Malek et al. (1995). It is important to note, though, that, in neither case, laminar, turbulent or pulsatile, is the flow in the apparatus mimicking a physiologic situation in the vasculature. The apparatus also fails to model a particular segment in the cardiovascular system, it merely provides a well defined biologic model for studying, in vitro, cellular response to flow induced forces. The parallel plate flow chamber can be considered as a rigid-wall mimic of a straight segment of blood vessel, devoid of branches or bifurcations, that is
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expressed in a rectangular coordinate system rather than in its “natural”, cylindrical form. In accordance with this transformation, the axisymmetric velocity profile is replaced by a one-dimensional flow whose transverse velocity is uniform across the entire width of the chamber. These features can be readily addressed in a simple and inexpensive design. Indeed, a number of designs have been successfully used over the years in a wide variety of investigations (Viggers, Wechezak and Sauvage, 1986; Sakariassen et al., 1983; Levesque and Nerem, 1985; Ruel et al., 1995; Helmlinger et al., 1991; Nollert, Eskin and McIntire, 1990; Frangos et al., 1985). Figure 1.2 details a parallel plate apparatus currently used in our laboratory. The design includes three elements: the main body of the apparatus, spacers for adjustment of channel height, and cell culturing chambers. The main body of the apparatus is fabricated from two rectangular pieces of polycarbonate. It contains two streamlined settling chambers, one on the upstream end of the test section and the other on the downstream end. The upstream settling chamber is designed to minimize small-scale disturbances and provide uniform, twodimensional flow into the specimen test area. The downstream settling chamber is designed to prevent mock-loop disturbances from affecting the flowfield at the end of the test section. The test section is a long rectangular channel whose height, h, is much smaller than its width, b, thus satisfying the requirements for a parallel-plate flow-geometry, h can be varied in a stepwise manner by using interchangeable spacers of different thickness, e.g., 250, 500 or 1000 µm, or the combination thereof. All spacers are cut from commercially available polyethylene terephthalate glycol sheets, and their inner contour is carefully fabricated to precisely match the streamlined walls of the settling chambers. Cells are cultured to confluence under static conditions in commercially available Permanox® slide chambers (Nunc Inc.,
Figure 1.2. Design details of a parallel-plate flow chamber. The Permanox® slide chambers for cell culturing are commercially available from Nunc Inc.
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Naperville, IL). At confluence, the upper structure is detached from the cell covered slide and the latter is snugly fit into a recess within the upper wall of the main body. The final assembly of this device and its connection to the mock-loop circulation are rather simple, and the flow can be made steady or pulsatile. In a fully developed laminar flow, which is typically obtained for L/h=0.04 Re (Schlichting, 1987), the velocity profile is parabolic and the shear stress at the wall is given by: (37)
where Q is the flow rate of the perfusate. However, in turbulent flow, the axial velocity profile becomes more flattened and equation 37 must be replaced by a more suitable formula (for details see Tennekes and Lumley (1972), pp. 149–156). A common problem encountered in this apparatus is the presence of three-dimensional flow near the side walls which, in order to avoid biased data, limits the effective lateral size of the cell-covered test area. As elaborated elsewhere (Tran-Son-Tay, 1993), this difficulty can be minimized in most designs by using b/h50. Also, to ensure one-dimensional flow over the entire specimen area it is recommended that the value of b is chosen so that it exceeds the width of the cell culturing slide by at least 20%. The description of the perfusion systems will not be complete without inclusion of a brief account about a new class of apparatuses that have been developed to study cell structure and function in a complex flow environment. Figure 1.3 depicts a flow apparatus that is currently used in our laboratory. The perfusion chamber is
Figure 1.3. Design details of a flow-through, cylindrical-cavity apparatus. The flow in the cavity is characterized by four distinct patters: central jet, flow impingement, flow separation and recirculating eddies.
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designed as a flow-through cylindrical cavity, with the inlet and outlet ports centered on the axis of planar symmetry, half way between the top and bottom walls. The main body and the removable walls are fabricated from polycarbonate and the sealing O-rings (Great Lakes Rubber, Glendale, WI) are made of Viton, that was found stable under cell culture conditions and non-cytotoxic (Lelkes et al., 1992). Cells are grown to confluence under static conditions on 13 mm in diameter Thermanox™ coverslips (Nunc Inc., Naperville, IL). At confluence, the coverslips are snugly fit into prefabricated recesses in the removable walls, the apparatus is assembled and connected to a mock-loop circulation. The system is perfused by steady or pulsatile flow at mean flow rates ranging from 11 mL/min (laminar flow) to 500 mL/min (turbulent flow). At these flow rates the large scale motion of the fluid in the circular cavity is well organized and its structure reflects the planar symmetry of the chamber. There are four distinct flow patterns in the cavity: central jet, flow impingement, flow separation and recirculating eddies. These flow patterns and their distinct impact on cell morphology (at the respective sites of the coverslips) were studied extensively in the past and are documented elsewhere (Christensen et al., 1992; Samet et al., 1993; Samet and Lelkes, 1994a; Samet and Lelkes, 1994b). It is noteworthy, though, that this type of flow apparatus does not mimic a physiologic situation in the cardiovascular system, nor does it model a particular segment of the vasculature. From the bioengineering standpoint, this design merely provides a biologic model for studying, in vitro, cellular response to distinct flow patterns that are also commonly encountered in the circulation. A number of systems have been developed over the years to study the effects of elevated pressure on cultured cells. The simplest apparatus consists of a tissue culture flask sealed with a rubber cap (Hishikawa et al., 1992a; Hishikawa et al., 1992b; Hishikawa et al., 1995). The flask is lined with cells and maintained in a standard, 5% CO2 incubator. At the beginning of the study the cap is tightened, and compressed helium gas is pumped in to elevate the pressure to a predetermined level. Albeit simple, this system is only suitable for short-term experiments, because it is not equipped with an on-line supply of proper gas mixture to counter-balance the inevitable changes in medium pH, which result from the metabolic activity of the cells. Another simple apparatus, described by Acevedo et al. (1993), utilizes a tall cylindrical container filled with culture medium to a predetermined height. Cells are grown on pretreated glass coverslips, and the cultures are subjected to the weight of the column of medium in the container. The resulting hydrostatic pressure is evaluated from equation 25. Since the pressurized cultures and controls are maintained in a standard incubator, where the level of CO2 remains constant, the experiments can be conducted for a long period of time. Yet, despite this apparent advantage, the system may not be suitable for studying the secretory activity of pressurized cells due to the significant volumetric difference in media (and, thereby, dilutions) between controls and their pressurized counterparts. In the last few years more advanced apparatuses have been devised that could accommodate one or more standard tissue culture plates and sustain elevated pressures of more than 200 mm Hg (Kato et al., 1994; Sumpio et al., 1994; Samet, Wankowski and Lelkes, 1994; Vergne et al., 1996). Some of these designs were also equipped with a sampling port or a glove-box module to allow withdrawal of
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culture medium for on-line analysis (Kato et al., 1994; Sumpio et al., 1994). In others, access to the cells was made possible only after decompression of the system (Vergne et al., 1996; Mattana and Singhal, 1995). Details of the apparatus currently used in our laboratory (Samet, Wankowski and Lelkes, 1994; Kanda et al., 1996; Silverman, Samet and Lelkes, 1996a-d; Silverman et al., 1998) are illustrated in Figure 1.4. The pressurized cell culture system consists of three elements: 1) regulated CO2 (5%)-air (95%) supply, 2) pressurized humidifier with a backflow trap and an access port for a monitoring unit, and 3) pressurized cell culture chambers. The entire system is completely independent of the atmospheric conditions because the constant, elevated pressure is provided via the CO2-air mixture which is required for cell culturing purposes. The CO2-air supply is connected to the humidifier via a series of filters (DFU grade, Balston Inc.) and reinforced Tygon® tubing. For practical purposes this design utilizes three pressurized cell culture chambers that are connected in parallel to the humidifier. A detailed view of the pressurized cell culture chamber is given in Figure 1.5. The pressurized cell culture chamber is equipped with three inlet/ outlet ports. One port connects the unit to the humidifier. The second port is used as a sampling port and it also serves as a pressure relief sink at the end of the experiment. The third port serves as an input for pulsatile pressure. The chamber is designed to house up to six 35 mm Petri dishes or a tissue culture plate with 6, 24 or 96 wells. To allow easy and sterile access to the cavity, the top wall is removable and it seals the chamber via an arrangement of a Viton gasket and quick-release mechanisms. Finally, pulsatility is generated by the rotary mechanism of a pulsatile, dual phase Harvard respirator and this input is superimposed on the basal static pressure in the chamber through one of the inlet ports.
Figure 1.4. Schematics of a pressurized cell culture system.
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Figure 1.5. A close-up image of one of the pressurized cell culture chambers housing a 96-well tissue culture plate.
The interest in using in vitro models to study properties and activities of vascular cells subjected to stretching/straining of their underlying substrate dates back to the pioneering studies of Leung, Glagov and Mathews (1976). Since then various schemes of stretch-generating devices have been developed, but only three types were found biologically and physiologically sound and, therefore, are actively utilized at present. For comprehensive accounts on this subject see Mills, Cohen and Sumpio (1993), and Tran-Son-Tay (1993). The simplest designs of stretch-generating apparatus rely on the principle of uniaxial loading (Leung, Glagov and Mathews, 1976; Terracio, Miller and Borg, 1988; Dartsch and Betz, 1989; Ives, Eskin and McIntire, 1986). Cells are cultured on thin, rectangular membranes made of a distensible biomaterial, such as elastin (Leung, Glagov and Mathews, 1976), silicone (Terracio, Miller and Borg, 1988; Dartsch and Betz, 1989), or polyetherurethane (Ives, Eskin and McIntire, 1986). One membrane is clamped on all sides and serves as static control. The other membrane is clamped on three sides to a fixed frame, whereas the fourth side is firmly attached to a sliding rod. The sliding rod is connected to a motor-driven camshaft that cyclically moves it back and forth, thereby stretching and relaxing the membrane. Both membranes are housed in separate containers filled with medium, and the entire system is kept in a standard incubator. Uniaxial loading devices are easy to operate and, with a proper electro-mechanical coupling, they also permit accurate control of periodicity and degree of stretching. However, axial stretching of the membrane also causes it to shrink in the lateral direction by an amount determined by the Poisson ratio of the biomaterial used. Thus, the actual effect in these devices on cultured cells is not purely tensional, and the response obtained/detected may be biased by the compression of the underlying substrate.
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Another simple method of stretching cell-lined substrates involves circumferential loading of circular tubes by pulsatile flow (Moore et al., 1994a). The cells are grown on the luminal surface of thin, compliant tubes made of a distensible biomaterial. At confluence the tubes are connected to mock-loop circulation equipped with two pumps. One pump is used to circulate growth medium in the system at a predetermined flow rate, whereas the other provides pressure pulses (at an adjustable frequency) which are superimposed onto the steady flow. Hence, like in arteries in vivo, the passage of these pulse waves distends the wall of the tubes and imparts circumferential stretch on cell substrate. Concomitant with stretching, the flowing medium also exerts shearing stresses on tube walls, resulting in cellular response that addresses the combined stimulatory effect of both hemodynamic challenges. Interestingly, the versatility of this design also permits generation of shear stress without the accompanying effect of stretching (by using rigid-wall tubes) or, alternatively, generation of pure stretching by idling the primary pump. The third type of stretch-generating apparatuses relies on the principle of biaxial loading (Baskin, Howard and Macarak, 1993; Winston et al., 1989; Banes et al., 1990). The most popular device in this category is a commercially available strain unit, termed, Flexercell™ (Flexcell International Corp., McKeesport, PA). The apparatus, shown in Figure 1.6, includes a computer controlled pressure-regulation module and a manifold that can accommodate up to eight flexible-bottomed tissue culture plates. Each plate contains six wells. The bottom wall of each well consists of a flat silicone membrane that is firmly glued to a narrow flange that extends
Figure 1.6. A close-up image of a commercially available strain unit, termed Flexercell™ (by permission from Flexcell International Corp., McKeesport, PA).
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radially from the inner wall. When negative (or positive) pressure is applied from below, the membrane stretches downward (or upward) and deforms into a concave (or convex) shell. Upon relaxation of the pressure, the membrane returns to its flat shape. The Flexercell™ apparatus is operated in a cyclic fashion utilizing a computer to time the application/relaxation of the stretch-activating pressure and for continuously monitoring its level. The timing signals are passed from the computer to a pressure-regulation module, which connects the input from a vacuum pump to the manifold. This module is equipped with solenoid valves and regulators that permit precise adjustment of the duration, magnitude and frequency of the stretch-activating pressure applied to the individual wells. Further details on this apparatus can be found in the papers of Banes et al. (1990) and Mills, Cohen and Sumpio (1993), and manufacturer’s literature.
CONCLUDING REMARKS The present chapter focuses on the hemodynamic environment of the endothelium from two perspectives, in vivo and in vitro. The in vivo viewpoint echoes the current state of knowledge of the cardiovascular system, and also manifests our understanding of the blood flow phenomena in the vasculature. The in vitro perspective, on the other hand, reflects the advancements made thus far in engineering biological models that facilitate studying and understanding of cellular responses to well defined mechanical stimuli. At present, the difference between these two environments is substantial, in part because, historically, centuries of study and experience separate between the two fields and, in part, due to the fact that no engineer or biologist can do as good a job as mother nature. All the same, most current biological models are simplistic and do not mimic specific physiologic situations in, or represent a particular segment of the vasculature. In addition, from a bioengineering standpoint, the apparatuses utilized fall short in a) the materials they use as a substitute for the vascular wall, b) the liquid milieu they contain instead of blood, and c) the limited duration of the study they permit as compared to the life-long span of natural occurrences. Finally, a suitable control for the dynamic experiments is yet to be defined. Improvements in the present in vitro (biological) models require the coordinated attention from a multi-disciplinary team of scientists, and furthering the research and development of novel technologies. To extend the duration of studies, cell culturing and co-culturing techniques will have to be revisited, with particular attention paid to minimizing cell senescence. Also, advanced assays and measuring techniques will be needed to facilitate on-line acquisition and analysis of data. Bettering the current mimics of circulation calls for creation of blood analogues that will replace the “standard” culture medium in the apparatuses. It also requires development of cytocompatible, compliant biomaterials that will serve as more realistic substitutes of the vascular wall. Finally, novel computational schemes will have to be devised to more accurately characterize the mechanical forces in complex geometries. Clearly, the list of “to-be-done” is long and ambitious, and the tasks specified challenging, but not impossible.
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ACKNOWLEDGMENTS The present work was supported in part by the Milwaukee Heart™ Research Foundation and grants from the American Heart Association (AHA), National Center and Wisconsin Affiliate. The authors are grateful to Mr. N.P.Samet for skillfully drafting the various designs.
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of human vascular endothelial stress fibers by fluid shear stress. Nature, 307, 648–649. Fry, D.L. (1968) Acute vascular endothelial changes associated with increased blood velocity gradients. Circ. Res., 22, 165–197. Girard, P.R., Helmlinger, G. and Nerem, R.M. (1993) Shear stress effects on the morphology and cytomatrix of cultured vascular endothelial cells. In Physical Forces and the Mammalian Cell, edited by J.A.Frangos, San Diego: Academic Press, Inc. pp. 193–222. Goldsmith, H.L. and Karino, T. (1988) Flow-induced interactions of blood cells with vessel wall. In Endothelial Cells, edited by U.S.Ryan, Boca Raton, FL: CRC Press, Inc. pp. 139–171. Goldstein, S. (1965) Modern Developments in Fluid Dynamics. New York: Dover Publications, Inc. Griendling, K.K. and Alexander, R.W. (1994) Cellular biology of blood vessels. In The Heart Arteries and Veins, 8th edn., edited by R.C.Schlant and R.W.Alexander, New York: McGraw-Hill, Inc. pp. 31–45. Guyton, A.C. (1981) Textbook of Medical Physiology, 6th edn. Philadelphia: W.B.Saunders Comp. Haller, H. (1997) Endothelial function: General considerations. Drugs, 53, 1–10. Harcus, A.W. and Adamson, L. Symposium on Arteries and Veins, Royal College of Physicians of London, 1973 (1975) Arteries and Veins [Proceedings], Edinburgh; New York: Churchil Livingston. pp. 1–326. Helmlinger, G., Geiger, R.V., Schreck, S. and Nerem, R.M. (1991) Effects of pulsatile flow on cultured vascular endothelial cell morphology. J. Biomech. Eng., 113, 123–131. Hishikawa, K., Nakaki, T., Marumo, T., Suzuki, H., Kato, R. and Saruta, T. (1995) Pressure enhances endothelin-1 release from cultured human endothelial cells. Hypertension, 25, 449–452. Hishikawa, K., Nakaki, T., Suzuki, H., Saruta, T. and Kato, R. (1992a) Transmural pressure inhibits nitric oxide release from human endothelial cells. Eur. J. Pharmacol., 215, 329–331. Hishikawa, K., Nakaki, T., Suzuki, H., Saruta, T. and Kato, R. (1992b) New method of investigating functional roles of pressure-sensitive mechanoreceptor in human endothelial cells. J. Cardiovasc. Pharmacol ., 20, S66–S67. Intaglietta, M. (1997) Whitaker lecture 1996: microcirculation, biomedical engineering and artificial blood. Annals of Biomedical Engineerings, 25, 593–603. Ives, C.L., Eskin, S.G. and McIntire, L.V. (1986) Mechanical effects on endothelial cell morphology: in vitro assessment. In Vitro Cell. Dev. Biol., 22, 500–507. Johnson, L.R. (1992) Hemodynamics. In Essential Medical Physiology, New York: Raven Press, pp. 151– 164. Kanda, K., Silverman, M.D., Samet, M.M., Hayman, G.T. and Lelkes, P.I. (1996) Elevated static pressure does not alter VCAM-1 and ICAM-1 expression in cultured human vascular cells. J. Vasc. Res., 33, S45 (Abstract). Karino, T., Goldsmith, H.L., Motomiya, M., Mabuchi, S. and Sohara, Y. (1987) Flow patterns in vessels of simple and complex geometries. Ann. N.Y. Acad. Sci., 516, 422–441. Kato, S., Sasaguri, Y., Azagami, S., Nakano, R., Hamada, T., Arima, N. et al. (1994) Ambient pressure stimulates immortalized human aortic endothelial cells to increase DNA synthesis and matrix metalloproteinase 1 (tissue collagenase) production. Virchows Archiv., 425, 385–390. Kenner, T. (1972) Flow and pressure in the arteries. In Biomechanics: Its Foundations and Objectives, edited by Y.C.Fung, N.Perrone and M.Anliker, Englewood Cliffs: Prentice-Hall, Inc. pp. 381–434. Kesler, K.A., Herring, M.B., Arnold, M.P., Glover, J.L., Park, H.-M., Helmus, M.N. et al. (1986) Enhanced strength of endothelial attachment on polyester elastomer and polytetrafluoroethylene graft surfaces with fibronectin substrate. J. Vasc. Surg., 3, 58–64. Ku, D.N. (1997) Blood flow in arteries. Annu. Rev. Fluid Mech., 29, 399–434. Ku, D.N. and Zhu, C. (1993) The mechanical environment of the artery. In Hemodynamic Forces and Vascular Cell Biology, edited by B.E.Sumpio, Austin: R.G.Landes Company, pp. 1–23. Kunz, A.L. and Coulter, N.A. (1967) Non-Newtonian behavior of blood in oscillatory flow. Biophys. J., 7, 25–26. Laufer, J. (1954) The structure of turbulence in fully developed pipe flow. In NACA Report, 1174, pp. 1–18. Lelkes, P.I., Samet, M.M., Christensen, C.W. and Amrani, D.L. (1992) Factitious angiogenesis: endothelialization of artificial cardiovascular prostheses. In Angiogenesis in Health and Disease, edited by M.E.Maragoudakis, P.Gullino, and P.I.Lelkes, New York: Plenum Press, pp. 339–351. Lemaire, R. (1978) Blood flow in the venous system. Phlebologie, 31, 101–111. Leung, D.Y.M., Glagov, S. and Mathews, M.B. (1976) Cyclic stretching stimulates synthesis of matrix components by arterial smooth muscle cells in vitro. Science, 191, 475–477. Levesque, M.J. and Nerem, R.M. (1985) The elongation and orientation of cultured endothelial cells in response to shear stress. J. Biomech. Eng., 107, 341–347. Liepsch, D.W. (1990) International Symposium on Biofluid Mechanics, Palm Springs, Calif., April 27–29,
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2 Chloride Channels in Endothelium: The Role of Mechano-stimulation and Changes in Cell Volume Bernd Nilius*, Jan Eggermont, Thomas Voets and Guy Droogmans Department of Physiology, KU Leuven, B-3000 Leuven, Belgium. *Corresponding author: B.Nilius, Department of Physiology, Campus Gasthuisberg, KU Leuven, B-3000 Leuven, Belgium, Tel.: 32–16–345 937, Fax: 32–16–345 991, E-mail address: [email protected].
This review describes one aspect of mechano-sensing of endothelial cell, namely the activation of ion channels, especially the activation of a volume-regulated anion channel, VRAC. Mechanical forces such as shear forces resulting from blood flow and mechanical strain (biaxial tensile stress) induce also changes in cell shape and possible folding and unfolding of the plasma membrane which in turn activate a variety of mechano- and volume sensitive ion channels, and help to adjust the vessel diameter to the hemodynamic needs. VRAC channels will be introduced as outwardly rectifying anion permeable channels. The gating mechanisms are still unknown. A probable role of tyrosine kinases and small G-proteins (RhoA) will be discussed. It will be shown how VRAC channels are involved in electrogenesis of EC and thus the regulation of various electro-chemical driving f0orces for a variety of transports through the plasma-membrane (e.g. Ca2+ influx), in regulation of the EC volume and the intracellular pH, and vectorial transport. The contribution of VRAC to create mechano-sensitive pathways for permeation of amino acids is documented. The possible involvement of VRAC in the regulation of proliferation of EC will be discussed in detail. The molecular biology of these channels will be critically evaluated. KEYWORDS: Endothelium, mechano-sensors, volume sensors, ion channels, volume regulated anion channels, cell proliferation.
INTRODUCTION One of the most exciting features of vascular endothelial cells (EC) is their ability to respond to mechanical stimuli. ECs are constantly exposed to mechanical forces such as shear forces resulting from blood flow and mechanical strain (biaxial tensile stress) which also induce changes in cell shape and possible folding and unfolding of the plasma membrane (Davies, 1995; Malek and Izumo, 1994). These forces activate a variety of biological responses, such as gating of mechano- and volume sensitive ion channels, and help to adjust the vessel diameter to the hemodynamic needs (Davies, 1995; Davies and Barbee, 1994; Davies and Tripathi, 1993; Nilius, 1991; Resnick and Gimbrone, 1995). Our current knowledge of the mechanisms that regulate such ion channels in endothelium is very limited. This article focuses on mechano- and volume sensitive ion channels in endothelium and in particular on Cl- channels. 33
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MECHANO-SENSITIVE CHANNELS IN ENDOTHELIUM: AN OVERVIEW Several ion channels in endothelial cells have been reported to be mechano-sensitive. These channels have previously been described in detail (Nilius, Viana and Droogmans, 1997e).
Channels Activated by Tensile Stress Negative pressure applied via the patch pipette to ECs from pig aorta activates a channel which is permeable for monovalent cations (40–56 pS) and Ca2+ (19 pS), and has a permeability ratio PCa: PNa of around 6 (Lansman, Hallam and Rink, 1987). This channel senses tensile stress. Stretch-activated non-selective cation channels with a conductance of 20–30 pS for monovalent cations and 10–20 pS for Ca2+ and Ba2+ have also been described in endocardial endothelium and microvascular ECs (Hoyer et al., 1994; Popp et al., 1992). Activation of these channels induces an increase in the intracellular Ca2+ concentration, [Ca2+]i. This increase is sufficient to activate big conductance, Ca2+ activated K+ channels, BKCa, and to hyperpolarize the membrane thereby establishing a positive feedback on Ca2+ -entry by increasing the driving force (Nilius, 1991). Direct activation of Ca2+ -permeable ion channels by positive pressure (not stretch) has been suggested by several groups (Hoyer et al., 1996; Marchenko and Sage, 1996; Marchenko and Sage, 1997). Another group of ion channels which also sense tensile stress comprise Cl- channnels. These channels are regulated by mechanical forces which are connected with changes in cell volume or alterations in cell shape and surface structures of endothelium (Nilius et al., 1997a; Nilius et al., 1996a). Probably, tensile forces may induce small changes in cell surface thereby folding or unfolding membrane invaginations or caveolae. Such a process seems to be responsible for gating of volume-regulated Cl- channels (VRAC) possibly via an involvement of the small G-protein Rho (Okada, 1997). This current will be described in detail later on. Cell swelling mediated tensile stress also activates K+ channels which, if co-activated with Cl- channels, may be involved in volume regulation (De Smet et al., 1994; Perry and Oneill, 1993).
Channels Activated by Shear Stress Changes in shear stress due to an alternating frequency of pulsatile flow activate small conductance potassium channels (SKCa) as well as big conductance K+ channels (BKCa). A variation of the viscosity does also affect BKCa channels (Davies, 1995; Hutcheson and Griffith, 1994; Jacobs et al., 1995). Opening of these K+ channels induces hyperpolarization in native and in cultured ECs, which may modulate the membrane potential in electrically coupled cells (Daut, Standen and Nelson, 1994; Davies, 1995; Davies and Barbee, 1994; Davies and Tripathi, 1993; Nakache and Gaub, 1988). An inwardly rectifying, 30 pS K+ channel has been described as a possible mechano-sensor of shear stress (Jacobs et al., 1995; Olesen, Clapham and Davies, 1988). Its single
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channel conductance, its degree of rectification and its Ca2+ –sensitivity in excised patches suggest that this channel may belong to the class of medium conductance K+ channels or BKCa channels, but not to the SKCa channels. It also shares some properties with the ROMK-like channels, and the previously described Ca2+-and GTP-modulated inwardly rectifying K+-channels (Davies, 1995; Ohno et al., 1993; Olesen et al., 1988; Vaca, Schilling and Kunze, 1992). Shear stress also activates cation channels, which might be somewhat more permeable for Ca2+ than Na+ (Schwarz, Droogmans and Nilius, 1992b). It appears to be insensitive to activators of protein kinase C and is reversibly blocked by La3+ and non-steroid anti-inflammatory drugs such as mefenamic acid (Schwarz et al., 1992a; Schwarz et al., 1992b). This channel may be involved in shear stress activated Ca2+ influx and shear stress mediated Ca2+ transients (Schwarz et al., 1992a). Interestingly, shear stress activated Ca2+ influx is completely abolished after removal of sialic acid in the glycocalyx of ECs (Hecker et al., 1993).
Mechanically Activated Ca 2+ Entry Figure 2.1 shows a typical group of results. If an endothelial cell is mechanically stimulated in a Ca2+ free solution, it responds with activation of a Ca2+ transient (Figure 2.1 A). Because of the absence of extracellular Ca2+, it must be related to Ca2+ release. This release can be activated by all kinds of mechano-stimulation: shear stress, cell swelling, and cell stretch (Oike, Droogmans and Nilius, 1994a). If after mechanostimulation an agonist is applied, no further Ca2+ signal can be activated. Pre-treatment of the cells with thapsigargin also attenuates any further release of Ca2+. This indicates that a) the mechanical stimulus releases Ca2+ from IP3 sensitive stores and b) this stimulation apparently depletes these stores. We have proposed that the mechanicallyinduced release of Ca2+ is mediated via production of arachidonic acid (AA) because a) PLC is not involved b) heparin does not block this release c) PLA2 blockers inhibit this release d) AA application mimics the effects of mechano-stimulation even in the presence of heparin and PLA2 blockers. The PKA inhibitory peptide is inefficient, as well as cyclooxygenase and lipoxygenase inhibition (for details see Oike et al., 1994a). Since mechanical stimulation of EC seems to empty the Ins(1, 4, 5)P3 sensitive Ca2+ stores, it may also activate Ca2+-entry via ‘store-operated’ Ca2+ entry channels which open upon store depletion. Currents which are activated by this mechanism are referred to as Ca2+-release-activated Ca2+ currents (CRAC) or currents through store-operated channels (SOC) (Fasolato and Nilius, 1997; Hoth and Penner, 1993). Experimental evidence for activation of Ca2+-entry due to mechano-stimulation is depicted in Figure 2.1B. Mechano-stimulation induces a Ca2+-transient as already shown in Figure 2.1A but also activates a current which is independent of the changes in [Ca2+]i. This current is identical with the volume-regulated, Ca2+-independent anion (Cl–) current (VRAC) and has been described in detail below (Nilius et al., 1997a; Nilius et al., 1996b; Nilius, Viana and Droogmans, 1997d). After switching to a Ca2+ free solution and reapplication of Ca2+ (10 mM) a large Ca2+-signal appears which reflects Ca2+-entry. Without the mechanical stimulation, stepwise elevation of extracellular Ca2+ does not induce a Ca2+ transient (Oike et al., 1994a). No current can be detected during reapplication of Ca2+. Under the chosen buffering conditions for [Ca2+]i (0.1 mM
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Figure 2.1. Events in endothelial cells following mechano-stimulation. (A): Exposure of endothelial cells to hypotonic Ca2+-free bath solutions activates an inward current (upper trace) and induces a transient increase in [Ca2+]i (lower trace). The holding potential was –40 mV. Additional application of 100 µm histamine did not induce a further release of Ca2+ indicating that the stores are emptied (top). HTS, applied after depletion of intracellular Ca2+-stores with 2 µM thapsigargin, does not evoke a Ca2+-transient (below). The same Ca2+-signals can be evoked by direct stretch or shear stress. Cell swelling by HTS is induced by changing from a 290 mOsm/l to a 185 mOsm/l Kreb’s solution (Oike et al., 1994a) (B): Swelling of endothelial cells by HTS increases [Ca2+]i and activates a Cl- current (VRAC). The recovery of [Ca2+]i in isotonic solution is accelerated in Ca2+-free solution, but [Ca2+]i increases again after resubmission of extracellular Ca2+. Apparently, a Ca2+-entry pathway is activated by the exposure to HTS, but it is not accompanied by a significant change in transmembrane current (Oike et al., 1994a).
EGT A in the pipette solution), currents which are activated by store depletion, are in the range of 0.05 pA/pF at –40 mV and thus not detectable (Gericke et al., 1994; Oike et al., 1994b).
ACTIVATION OF CL - CHANNELS BY MECHANICAL STIMULI AND CHANGES IN CELL VOLUME Endothelial cells are constantly exposed to shear forces and also to biaxial mechanical stress induced by stretching or bending of the cell membrane. Both forces might be important for unfolding of the plasma membrane. Such an unfolding has been discussed as a possible signal to activate anion channels which are important for the regulation of cell volume after swelling (Okada, 1997). This unfolding mechanism could be of specific importance in endothelial cells because of the abundant presence of caveolae. Inflation of many cell types and even a short pulse of pressure positive pressure (5–10 s) is sufficient to elicit the whole sequence of events that leads to activation of such volume-sensitive anion channels (Doroshenko and Neher, 1992; Doroshenko, 1991;
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Lewis et al., 1993). These results suggest that activation of the Cl- conductance apparently occurs through membrane stretch. Similar if not identical anion channels are found in nearly every cell type. These anion channels are under physiological conditions mainly permeated by Cl-. They are referred to as volume-regulated anion channels (VRAC), because cell volume is the signal that most closely correlates with channel activation (Nilius et al., 1997a). In endothelial cells VRAC channels are not only activated by changes in cell volume, but also by changes in cell shape and by shear stress. The current through VRAC has already been introduced in Figure 2.1A and a more detailed description is given in Figure 2.2. A current which is mainly carried by Cl-, is activated by exposure to positive pressure via the patch pipette, by perfusion with an intracellular hypertonic solution, by a challenge by an extracellular hypotonic solution or by stimulation by shear stress (Figure 2.2A). The current can be repetitively activated by cell swelling. Panel A shows the time course of the current measured at -80 mV during a challenge with an extracellular 27% hypo-osmotic solution. As seen in the responses to the
Figure 2.2. Activation of VRAC in cultured pulmonary artery endothelial cells, CPAE. (A) VRAC can be activated by three different protocols: cell swelling by exposing the CPAE cell to a hypotonic extracellular solution or perfusion of the cell via the patch pipette with a hyperosmotic solution, inflation of the cell and application of a stream of solution along the cell surface (shear stress). (B) Time course of the current activated by challenging a cell with a 28% hypotonic mannitol Krebs’ solution measured at -80mV during repetitive voltage ramp protocols. Time course of the current at -80 mV. (C) Current traces during the step protocol under the indicated conditions (holding potential is 0 mV, 2 s steps ranging from -80 to +100 mV, spaced 20 mV, extracellular Mg2+ is 1 mM). Note the slow inactivation of the current at +100 mV. Arrows indicate zero current. Note the slowly inactivated currents present already under non-stimulated conditions. (D) Exposure of the cell to a 27% hypotonic solution to activated VRAC. (E) The cell is exposed to a 50 mM mannitol to induced cell shrinking. The outward current seen in panel C disappeared indicating its volume sensitivity.
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voltage steps, the current is already present in isotonic solution, which indicates that this current is partially activated in resting cells. Therefore, ICl, swell, the current through VRAC, is important for the electrogenesis of the resting potential in non-stimulated endothelial cells CPAE cells. Voltage steps (from -80 to +100 mV, increment 20 mV) evoked only small currents in resting cells (Figure 2.2B), but much larger currents during cell swelling (Figure 2.2C). These currents show outward rectification and are time-independent, except at potentials positive to +60 mV where they slowly inactivate. Cell shrinking again reduced the currents below their value in control conditions (Figure 2.2D), indicating the preactivation of VRAC under isotonic conditions. The I–V curves, obtained either from voltage ramps or from the step protocol, show outward rectification. The reversal potential is -19 mV and is close to the theoretical Cl- equilibrium potential, ECl, of -26 mV in the “hypotonic Krebs” solution (Figure 2.2E). We have shown previously that this current is mainly carried by Cl- but amino acids such as aspartate, which is a constituent of the pipette solution, also contribute to the current. This may account for the reversal potential more positive than ECl. Biophysics of VRAC In our previous experiments on mechano-sensitive responses of endothelium (Nilius, 1991; Nilius and Riemann, 1990; Oike et al., 1994a) we were constantly confronted with activation of VRAC under conditions of mechano-stimulation and we have published the first description of this current in EC (Nilius et al., 1994a). The mechanical stimulus for channel activation is under all these conditions probably related to unfolding of part of the cell membrane (Nilius et al., 1997a; Okada, 1997). The threshold for changes in cell volume is very small, a 3–5% change in cell volume already activates VRAC (Nilius et al., 1997a). Under hypotonic conditions, the current density of ICl, swell between +80 and +100 mV ranges for cultured endothelial cells from 100 to 150 pA/pF (Nilius et al., 1996b; Nilius et al., 1994a). For freshly isolated endothelial cells and cells in primary culture the density is less than 40 pA/pF (Nilius et al., 1994a; Nilius et al., 1994c). Permeation of VRAC is characterized by the following sequence: SCN - >I - >NO - 3>Br ->Cl - >HCO -3>F - > gluconate>glycine>taurine>aspartae, glutamate. Thus, amino acids and organic osmolytes also permeate through VRAC (Kirk, Ellory and Young, 1992; Nilius et al., 1997a; Strange, Emma and Jackson, 1996). The permeability ratios for amino acids correlate well with the degree of reduction in the cellular concentration of the different amino acids following hypo-osmotic swelling in endothelial cells (Manolopoulos et al., 1997b). The single channel conductance is approximately 40–50 pS at positive potentials and 10–20pS at negative potentials (reviewed in Nilius et al., 1997a; Nilius et al., 1996b; Strange et al., 1996; Strange and Jackson, 1995). Rectification is a property of the open channel. No reliable correlation could be found between the volume regulated anion currents and changes in membrane capacitance (Heinke et al., 1997). Activation of VRAC is Ca2+-independent (Szücs et al., 1996b). VRAC is further characterized by inactivation at positive potentials and a voltagedependent recovery from inactivation. Its kinetic properties are modulated by extracellular divalent cations, extracellular pH, the permeating anion and various
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channel blockers (Voets, Droogmans and Nilius, 1997a). Inactivation is accelerated by acidic extracellular pH and by an increase in the extracellular Mg2+ concentration, [Mg2+]e. Figure 2.3 shows this acceleration in bovine pulmonary artery endothelial cells. As compared with Figure 2.2C, which shows inactivation with 1.5 mM [Mg2+]e, an increase of [Mg2+]e from 7.5 to 60 mM drastically accelerated the decay of the current at potentials positive to +40 mV. It has been proposed that the influx of Clanions at positive membrane potentials drags the Mg2+ ions and/or protons into a blocking site within the channel pore, thus causing inactivation (Anderson, Jirsch and Fedida, 1995). Another possibility is the binding of Mg2+ ions and protons to negative charges on the channel protein, likely outside the electrical field, thereby altering the channel properties and accentuating inactivation (Nilius et al., 1997a; Nilius et al., 1996b; Voets et al., 1997a). Changes in inactivation observed for various cell types, might be explained by differences in Mg2+ or H+ affinities of the tentative binding site.
Possible Gating Mechanism of VRAC It is not known how mechano-stimulation or cell swelling affects gating of VRAC. The cytoskeleton is a likely candidate for transmitting volume changes and stretch forces to membrane channels (Ingber, 1997). In a variety of cells, it has been shown that depolymerization of the F-actin network by cytochalasine and various clostridial toxins interferes with activation of VRAC. Disrupting the microtubular network with cholchicine or colcemide or stabilising it with taxol had minimal effects on VRAC (for a detailled review see Nilius et al., 1997a). In endothelial cells, the F-actin
Figure 2.3. Modulation of inactivation of VRAC. As shown in Figure 2.2, VRAC inactivates slowly at low extracellular Mg2+ concentrations. Increasing extracellular Mg2+ substantially accelerated the decay of the current. VRAC is activated by challenge with a 27% hypotonic solution. A voltage step protocol is applied after maximal activation of the current (holding potential -100 mV, 2 sec steps from -80mV to 1+120 mV, sampling interval 2 ms).
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microfilamentous network and the microtubular system are most probably not important for activation of VRAC since cytochalasine nor taxol affected ICl, swell (Oike et al., 1994c). In contrast to the cytosolic cytoskeleton, there is experimental evidence pointing to the plasma membrane cytoskeleton being involved in mechano-signalling. In endothelial cells the annexin II-p11 complex might be involved in regulation of VRAC (Nilius et al., 1996c). Interestingly, annexins are involved in the formation of caveolae in EC. It is possible that annexins, as components of the subplasmalemmal cytoskeleton, transfer changes in cell volume or cell shape to transmembrane proteins. Interestingly, VRAC activation can be uncoupled from cell swelling at very low [Ca2+]i levels (50 nM to 100 nM; “permissive” or “threshold” concentration) (Szücs et al., 1996b). Interestingly, the annexin-P11 complex also dissociates from the plasma membrane below these Ca2+ concentrations (Kaetzel and Dedman, 1995). Integrins could act as transducers of mechano-sensitive and volume-induced effects by sensing the mechanical stretch on the membrane (Ingber, 1997). However, any connection between integrin function and VRAC has not yet been thoroughly evaluated. We have shown in macrovascular endothelial cells (bovine pulmonary artery) that the protein tyrosine kinase (PTK) inhibitors tyrphostin B46, tyrphostin A25 and genistein inhibited ICl, swell. Tyrphostin A1, a tyrphostin analogue with little effect on PTK activity, and daidzein, an inactive genistein analogue, were without effect on VRAC. The protein tyrosine phosphatase (PTP) inhibitors Na3VO4 and dephostatin potentiated ICl, swell when the current was preactivated by mild hypotonicity, but they could not activate ICl, swell under isotonic conditions. These observations indicate that one or more tyrosine phosphorylation steps are required for activation of VRAC by cell swelling. Interestingly, a swelling-activated tyrosine kinase activity has been reported in cardiomyocytes (Sadoshima et al., 1996). Intracellular perfusion with GTPS (100 µM) transiently activated a Cl- current with an identical biophysical and pharmacological profile as ICl, swell, the VRAC mediated current. This current was also inhibited by the tested PTK inhibitors and potentiated by the PTP inhibitors. Hypertonicity induced cell shrinking completely inhibited the GTPS-activated Cl- current. Intracellular perfusion with GDPßS (1 mM) caused a time-dependent inhibition of ICl, swell, which was more pronounced when the current was activated by mild hypotonicity. These results demonstrate that the activity of endothelial swelling-activated Cl- channels is dependent on tyrosine phosphorylation and suggest that a G-protein regulates the sensitivity to cell swelling (Voets et al., 1998). We therefore propose a model in which cell swelling activates a protein tyrosine kinase and that this might be an essential step in the activation cascade of VRAC. More general, we suppose that an important part of the mechanosensory machinery of macrovascular endothelial cells acts via a PTK/ PTP dependent mechanisms. In intestinal epithelial cells (Tilly et al., 1996) and in endothelial cells (own preliminary experiments) there is evidence that small GTP-binding proteins of the Rho family may modulate VRAC. Pretreatment of human intestine 407 cells with Clostridium botulinum C3 exoenzyme (an irreversible inactivator of p21Rho) reduced the swelling-induced efflux of iodide (Tilly et al., 1996). Conversely, the cytotoxic
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necrotizing factor CNF1 permanently activates RhoA by deaminating a glutamine at position 63 (Schmidt et al., 1997). Loading bovine pulmonary artery endothelial cells with CNF1 (Voets, Nilius, unpublished) can activate VRAC under isovolumic conditions. Consistent with these observations is the receptor-dependent potentiating effect of thrombin (an activator of Rho in endothelium) on ICl, swell (Manolopoulos et al., 1997a). At present it is not clear how Rho interacts with VRAC nor whether Rho can fully account for the GTPS effects on VRAC. VRAC can also be stimulated under isovolumic conditions by intracellular perfusion with a pipette solution of reduced ionic strength, i (Nilius et al., 1998). Reducing i at constant osmolarity and Cl- concentration activates an outwardly rectifying current that is mainly carried by Cl--ions and inactivates at positive potentials. The permeability ratio for various anions is PI>PBr>PCl>>Pgluc. Blockers of the swelling-activated Cl- current in CPAE cells also inhibit the current which is activated by a reduction in i. Hypertonic extracellular solutions rapidly and reversibly antagonised the i activated current, whereas increasing i precluded activation of ICl, swell by hypotonic shock. These experiments indicate that a reduction of i activates an anion current that is identical with ICl, swell. Similarly, the protein tyrosine kinase inhibitors tyrphostin B46 and genistein antagonise the i induced current indicating that the i activation requires tyrosine phosphorylation. Since the GTPS- and the i-activated currents are still sensitive to changes in cell volume and since they can be blocked by PTK inhibitors, it seems likely that both stimuli act upstream of the tyrosine phosphorylation step by inducing a shift in the volume-sensitivity of the PTK(s) (Figure 2.4). With respect to gating models it should be pointed out that activation of VRAC is most likely not achieved by an increase in the open probability of already accessible channels. There is indeed experimental evidence that VRAC activation occurs via an increase in the number of available
Figure 2.4. Possible activation cascade of VRAC. PTK: protein tyrosine kinase, PTP: protein tyrosine phosphatase, GGTP: G protein, possibly Rho, G: ionic strength. See text.
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channels which open with a very high open probability (˜ 1) (Jackson and Strange, 1995a; Jackson and Strange, 1995b).
Pharmacological Properties of VRAC VRAC has pharmacological properties which differ from other Cl- channels. The commonly used Cl- channel blockers, such as DIDS, SITS, N-phenylanthracillic acid (NPA), 9-AC (9-anthracene carboxylic acid) etc. have a low affinity for the volumeactivated Cl- current. Substances, such as NPPB, niflumic acid, 1, 9-dideoxyforskolin, verapamil, induce half-maximal block of VRAC in endothelium at concentrations of some 10 µM. Phenol-derivatives such as gossypol, a polyphenolic pigment found in cotton plants, are rather potent blockers of VRAC in endothelium (Szücs et al., 1996a). Furosemide inhibits volume-activated Cl- currents in epithelial and blood cells, but not in endothelium (Nilius, unpublished). Furthermore, antiallergic drugs from the chromone family also inhibit VRAC in endothelial cells (Heinke et al., 1995). Inhibitors of phospholipase A2 (PLA2) such as p-Bromophenacyl bromide (pBPB) and cyclosporin A, and arachidonic acid are efficient blockers of VRAC (Nilius et al., 1997a; Nilius et al., 1996b; Nilius, Sehrer and Droogmans, 1994b; Nilius et al., 1997d; Oike et al., 1994a). The block of VRAC by the anti-estrogen tamoxifen, a compound that has been widely used in the treatment of breast cancer, correlates nicely with its inhibitory effect on the proliferation of endothelial cells (Nilius et al., 1994b; Voets et al., 1995). Surprisingly, the anti-malaria compounds quinine and quinidine are also potent blockers of VRAC. Both drugs inhibit the current more efficiently at alkaline extracellular pH, indicating that they exert their action in the uncharged form (Voets, Droogmans and Nilius, 1996). Also, the anti-arrhythmic, anti-proliferative and antiischemic Ca2+-antagonist mibefradil efficiently inhibit VRAC (Nilius et al., 1997b). Table 2.1 gives an overview of the pharmacological modulation of VRAC in endothelium.
VRAC: Functional Significance
Electrogenesis Under iso-osmotic conditions VRAC is already partially activated in resting EC. This background Cl- conductance contributes to the resting potential. Activation of additional Cl- channels will shift the membrane potential towards the Cl- equilibrium potential. If a highly non-linear conductance, such as the inwardly rectifying K+ channel, is present, a bistable membrane potential may arise (Voets, Droogmans and Nilius, 1996; Nilius et al., 1997c). Small changes in Cl- conductance can under these circumstances induce drastic changes in membrane potential, shifting it from a value close to the K+ equilibrium potential to a value close to the Cl- equilibrium potential, or vice versa. Changes in membrane potential due to modulation of the volumesensitive Cl- conductance may thus significantly influence the electrochemical gradient
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Table 2.1a Blockers of VRAC in endothelial cells
Table 2.1b Modulators of VRAC (for details see text)
– indicates “no effect”, ? indicates “uncertain”
for a variety of transport systems. We have shown that administration of mibefradil induces hyperpolarization of macro vascular EC (Nilius et al., 1991c). Figure 2.5 shows an example. Application of 10 µM mibefradil induced a very fast
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Figure 2.5. Effects of mibefradil on the membrane potential of CPAE cells. (A). Membrane potential was measured in current clamp mode after breaking into the cell. Mibefradil (10 µM) induced a fast and reversible hyperpolarisation of the cells. Shown is a typical cell with a large Cl- conductance and a resting potential at -10 mV. (B). Distribution of the membrane potentials sampled at 2 Hz from the cell shown in panel A. The two peaks represent the membrane potential in the absence (control) and the presence of mibefradil. The mean values were taken from the Gaussian fits. Data obtained from the fits are: -16.4 mV, width 4.5 mV for control, 70.3 mV, width 5.6 mV for 10 µM mibefradil (bin width 2 mV).
hyperpolarization of the cell from –10 to –70 mV. This hyperpolarization is completely reversible. Mibefradil (Ro 40–597) is a novel calcium-antagonist that causes welltolerated antihypertensive and anti-ischemic effects and prevents is chemically induced ventricular fibrillation without negative inotropic effects (Bernink et al., 1996; Billman and Hamlin, 1996; Braun et al., 1996; Fang and Osterrieder, 1991). In addition to the Ca2+-antagonistic effects, it may facilitate the effects of endothelium-derived NO, affect eicosanoid production, have antiproliferative effects on smooth muscle cells after vascular injury and exert a vasodilating action via PKC-inhibition (Hermsmeyer and Miyagawa, 1996; Kung et al., 1995; Schmitt et al., 1995). The endothelial hyperpolarization shown in Figure 2.5 is due to the inhibition of the background VRAC (Nilius et al., 1997c). Hyperpolarization increases release of NO and other EDRF’s probably via an increased driving force for Ca2+ influx (Luckhoff and Busse, 1990; Nilius and Droogmans, 1995). It is therefore intriguing to speculate whether some of the above described beneficial cardiovascular effects of mibefradil are induced by their effect on the mechano-sensitive VRAC. Volume regulation One of the more important physiological features of the volume-regulated Cl- channel is that it provides volume-dependent mass transport of inorganic and organic osmolytes. VRAC is one of the main players in osmo- and volume regulation in many cell types
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(for some recent reviews see Hoffmann and Dunham, 1995; Lang et al., 1993; Nilius et al., 1997a; Nilius et al., 1996b; Sarkadi and Parker, 1991). It is not clear whether such a function is of relevance for EC under in vivo conditions. However, as discussed below this function might be important during the transition of EC through the cell cycle. In exocytotic cells, this channel seems to be important for volume regulation following degranulation (Dietrich and Lindau, 1994). This might also apply, although no experimental evidence has been given so far, for vascular endothelial cells. Intracellular pH regulation The exact contribution of VRAC to pH regulation is only anticipated. We have shown that and also lactate permeate through VRAC (PHCO/PCl–= 0.62 ±0.022, n=11, Plactate/ PCl=0.29±0.05, n=6; Nilius, unpublished). Obviously, VRAC provides a pathway for both metabolically important compounds. The high current density may change intracellular or compartmentalized Cl- concentrations and thereby affect other Cldependent transport mechanisms, such as the exchanger and the K+-Na+-2Cl transporter which are closely involved in the regulation of intracellular pH (Nilius et al., 1997a). Vectorial transport In many cell types VRAC is also involved in the directional transport of Cl- and, consequently, in salt and fluid secretion (McEwan et al., 1993; Strange, 1992; Strange et al., 1996). The vectorial transport of Cl- depends on its asymmetrical expression. Such a polarized distribution of VRAC in EC is not known. Exocytosis It has been shown that intracellular Cl- modulates exocytosis in several cell types. (Churcher and Gomperts, 1990; Dietrich and Lindau, 1994; Lindau and Gomperts, 1991; Rupnik and Zorec, 1992; Rupnik et al., 1994). It is again intriguing to speculate that VRAC could also be involved in exocytotic processes in endothelium. A pathway for amino acids Without doubt, VRAC provides a pathway for the transport of amino acids and non-ionic osmolytes (Manolopoulos et al., 1997b; Nilius et al, 1997a; Nilius et al., 1996a; Nilius et al., 1997d; Strange et al., 1996). The cytosol contains high concentrations of organic “osmolytes”, which include structurally dissimilar molecules such as amino acids (taurine, glycine), polyols (myo-inositol, sorbitol) and methylamines (betaine). Efflux of organic osmolytes (most often represented by taurine and myo-inositol) via VRAC has been observed in numerous cell types (Jackson and Strange, 1993; Kirk et al., 1992; Roy, 1995). We have shown that hyposmotic swelling of EC also activates the efflux of taurine, myo-inositol, and other organic solutes with the same pharmacological profile as VRAC. Also glycine, aspartate, and glutamate permeate the channel (Px:PCl=0.6:0.38:0.12:0.11 for glycine, taurine, aspartate, glutamate) and result in a reduction of its intracellular concentration (Manolopoulos et al., 1997b).
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Likely, VRAC is identical with the taurine pathway. As we have discussed already, the response of EC to changes in cell volume may be linked to responses to other mechanical stimuli such as shear stress, hydrostatic pressure, and stretch. Moreover, volume and shape changes take place during proliferation, contraction, migration, and other critical cellular processes. With its ability to mediate the efflux of anions, organic osmolytes and amino acids, VRAC may be a necessary component of these cellular processes. The controlled release of organic molecules from the cytosol may also serve cell homeostasis in ways unrelated to volume regulation. Taurine functions has been proposed to protect the cells from the disrupting effects of various stresses, such as hypoxia, reoxygenization, exposure to radicals (Huxtable, 1992). By regulating the release of molecules such as taurine, VRAC may be an indispensable component of the protective machinery developed by cells to deal with such stresses. Such a function would be particularly useful in EC, as these cells are often exposed to abnormal chemical and mechanical stimuli. Modulation of the driving force for Ca 2+-entry Activation of the swelling-induced conductance affects the membrane potential as already described. Therefore, the driving force for Ca2+ ions will be modulated by changes in activation of VRAC. We have already shown that block of VRAC substantially hyperpolarized the EC membrane. This may increase the driving force for the Ca2+ release-activated Ca2+ entry (CRAC) which is also activated during cell swelling (CRAC, see e.g. Hosoki and Iijima, 1994; Hosoki and Iijima, 1995; Oike et al., 1994a). Cell proliferation Inhibition of volume-activated Cl- currents suppresses cell proliferation in many cell types, e.g. myeloblastic leukemia cells, T-lymphocytes, glia cells, myogenic (BC3H1 or C2C12) or neuronal (PC12) cell lines (Schlichter et al., 1996; Schmitt et al, 1995; Schumacher et al., 1995; Ullrich, Gillespie and Sontheimer, 1996; Voets et al, 1997b). Swelling-induced Cl- currents are prominent in proliferating cell lines, but are largely attenuated if these cells switch to a differentiated state (Nilius and Riemann, 1990; Voets et al., 1997b). VRAC is also involved in growth of primary cell cultures of carcinoma in situ and in non-invasive cancer cells of the cervix, but not in cells from normal cervix tissue and has been associated with carcinogenesis (Chou, Shen and Wu, 1995). Proliferation of endothelial cells is arrested in the presence of structurally unrelated compounds that inhibit VRAC (Nilius and Riemann, 1990; Voets et al., 1995). Because endothelial cells induce vascularisation of neighbouring tissue in response to VEGF growth factors, this effect of Cl- channel blockers could be of therapeutic interest as they may provide a tool to inhibit angiogenesis and to impair neovascularization of tumour cells. Probably, VRAC activation is necessary as long as a cell is progressing through the cell cycle and is down regulated when it withdraws from the cycle. The underlying mechanism is unclear. It is also unclear whether such a mechanism may play a role in
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atherosclerosis or arteriolopathy which are somehow connected to EC proliferation (Waer, 1996). Metabolic functions Activation of VRAC may change the Cl- concentration in the cytosol or in a restricted space near the plasma membrane. This in turn could affect a recently described Cl-dependent GTP-utilising plasma membrane protein kinase (Treharne, Marshall and Mehta, 1994). Molecular Biology So far, the molecular identity of VRAC is a highly controversial issue. Over the past years several molecular candidates have been put forward: MDR1 P-glycoprotein (Gill et al., 1992; Valverde et al., 1992), pICln (Gschwentner et al., 1994), ClC3 (Coca-Prados et al., 1996; Duan, Hume and Nattel, 1997a; Duan et al., 1997b; Duan D, 1997), band 3 anion exchanger (Fiévet et al., 1995) and phospholemman (Moorman et al., 1995). However, there is as yet no conclusive evidence for any of these candidates and the identity of VRAC remains in our opinion still an unresolved issue. P-gp is a 170 kDa, glycosylated plasma membrane protein which in humans is encoded by the MDR1 gene. P-gp consists of two symmetric halves with each halve comprising a hydrophobic region (6 transmembrane domains) and a cytosolic ATP binding site, the so-called nucleotide binding fold. The domain organisation as well as the conserved structure of the ATP binding site place P-gp in the superfamily of ATP Binding Cassette (ABC) membrane transporters. P-gp is a transporter that utilises ATP hydrolysis to extrude hydrophobic compounds including many cytotoxic drugs used in cancer treatment out of the cell. MDR1 P-glycoprotein is expressed in endothelial cells, but the lack of correlation between P-gp expression and density of VRAC as well as the absence of a mutual interference between P-gp transport activity and VRAC argue against P-gp being VRAC (De Greef et al., 1995a; De Greef et al., 1995b; Viana et al., 1995). Furthermore, heterologous expression of P-gp in Xenopus oocytes did not induce swelling-activated chloride currents (Morin et al., 1995). Consequently, the hypothesis that P-gp is VRAC has been more or less abandoned, but it remains possible that it acts as regulator of VRAC (Okada, 1997). pICln, has also been proposed as VRAC or a regulator of VRAC (Krapivinsky et al., 1994a; Paulmichl et al., 1992b). pICln is a protein of 235 to 241 amino acids depending on the species with a predicted molecular mass of approximately 26 kDa and with a ubiquitous distribution pattern (Abe et al., 1993; Anguita et al., 1995; Buyse et al., 1996; Krapivinsky et al., 1994b; Paulmichl et al., 1992a). Secondary structure predictions fail to identify hydrophobic stretches that are sufficiently long to span a phospholipid bilayer as an a-helix. A striking and well conserved feature in the primary structure of pICln is its acidic nature. Because pICln is abundantly expressed in EC, we also addressed the relation between the pICln protein and VRAC. We cloned human pICln and studied its expression pattern at the mRNA and protein
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level (Buyse et al., 1996). pICln is mainly cytosolic located in EC and does not shift to the plasma membrane during cell swelling (Buyse et al., 1997). The elusive role of pICln as a candidate for VRAC is discussed in detail elsewhere (Nilius et al., 1997a). ClC-2 is a membrane protein (907 amino acids) that structurally belongs to the ClC (Chloride Channel) superfamily (Thiemann et al., 1992). Hydropathy analysis of ClC proteins predicts 13 putative transmembrane domains (D1 to D13), but domains D4 and D13 are no longer considered as membrane-spanning segments (Jentsch and Günther, 1997; Jentsch et al., 1995). D4 is now positioned as an extracellular loop and D13 resides in the cytosol. ClC-2 is a chloride channel activated by hyperpolarisation, extracellular acidification and hypotonicity (Jordt and Jentsch, 1997). However, the anion selectivity, the kinetics and the rectification pattern of ClC-2 are fundamentally different from those of VRAC which excludes ClC-2 as a molecular candidate for VRAC. Phospholemman was initially purified from cardiac sarcolemma as the major substrate for Protein Kinase A and Protein Kinase C (Palmer, Scott and Jones, 1991). It is a small intrinsic membrane protein (72 amino acids) with a single membranespanning domain, an extracellular N-terminus and an intracellular C-terminus. The cytosolic tail contains the phosphorylation sites for Protein Kinase A. The maxiconductance, the anion selectivity (Cl->Br-), the taurine over Cl- permeability and the cation-selective substate of the phospholemman channel do not correspond with the known features of VRAC. In addition, no relation to mechanical stimulation or cell swelling has been shown yet. Recently, ClC-3 has been shown to be a VRAC-like channel in cardiac cells (Duan et al., 1997b). ClC-3 is another membrane protein (760 amino acids) of the ClC superfamily. However, this channel is PKC modulated and not sensitive to tamoxifen which is clearly different from VRAC in EC.
CONCLUSION In this review we have discussed various endothelial ion channels that are activated by mechanical stimuli such as shear stress, biaxial stress and cell swelling. The different K+ channels activated by shear stress are only briefly discussed since they have already been well described. This article focused on a Ca2+ entry pathway which is activated by shear stress, membrane stretch and cell swelling probably via a mechano-sensitive phospholipase A2. The mechanism of Ca2+ entry seems to involve depletion of Ins(1, 4, 5)P3—sensitive intracellular Ca2+-stores (arachidonic sensitive release, mechanoactivation of leaks?) followed by activation of CRAC (Ca2+-release activated Ca2+ currents) channels. Another important player during mechanical EC activation is the volume-regulated anion channel, VRAC. In macrovasculair EC this channel is already active under resting conditions as a house-keeping Cl- channel. During mechanostimulation, changes in cell volume or shape VRAC is substantially enhanced and may carry not only anions but also osmolytes and amino acids. Activation of VRAC might be coupled to unfolding of the plasma membrane (caveolae?). The activation cascade of VRAC seems to involve one or more tyrosine phosphorylation steps and there is evidence for a modulation by GTP binding proteins (probably Rho) and by
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Figure 2.6. Scheme of the mechano-sensing mechanism for Ca2+ entry and VRAC. Mechanoinduced Ca2+-entry is probably via store depletion and subsequent activation of CRAC. A major response is the activation of VRAC in which protein tyrosine kinases (PTK), the small G-protein Rho and annexins might be involved. More hypothetically, unfolding of caveolae might be an intriguing mechanism for activation of VRAC. For details see text.
annexins. In addition to its role in mechano-sensitivity VRAC may also serve other cell functions or signaling cascades, but its overall functional impact still remains hypothetical. The molecular identity of endothelial VRAC is still not elucidated and it remains to be shown whether ClC-3, the recently identified cardiac VRAC, plays a role in endothelial cells. Figure 2.6 gives a scheme of the processes discussed in this article.
ACKNOWLEDGEMENT We thank Drs. T.Voets, V.Manolopoulos, G.Szücs, M.Kamouchi, D.Trouet and G.Buyse for many helpful discussions and R.Casteels for his continuous interest and support. The excellent technical support of J.Prenen, D.Hermans, A. Florizoone and M.Crabbé is greatly acknowledged. J.Eggermont is a Research Associate of the Fund for Scientific Research (FWO-Vlaanderen). The work was supported by grants form the Federal Belgian and Flemish Government (N.F.W.O. G.0237.95, IUAP Nr.3P4/ 23, C.O.F./96/22-A0659 and DWTC), and by the European Commission (concerted action BMH4-CT96–0602).
REFERENCES Abe, T., Takeuchi, K., Ishii, K. and Abe, K. (1993) Molecular cloning and expression of a rat cDNA encoding MDCK-type chloride channel. Biochimica et Biophysica Acta, 1173, 353–356. Andersen, J.W., Jirsch, J.D. and Fedida, D. (1995) Cation regulation of anion current activated by cell swelling in two types of human epithelial cancer cells. Journal of Physiology, 483, 549–57. Anguita, J., Chalfant, M.L., Civan, M.M. and Coca-Prados, M. (1995) Molecular cloning of the human volume-sensitive chloride conductance regulatory protein, pICln, from ocular ciliary epithelium. Biochemical and Biophysical Research Communications, 208, 89–95,
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Fiévet, B., Gabillat, N., Borgese, F. and Motais, R. (1995) Expression of band 3 anion exchanger induces chloride current and taurine transport: Structure-function analysis. EM BO Journal, 14, 5158–5169. Gericke, M., Oike, M., Droogmans, G. and Nilius, B. (1994) Inhibition of capacitative Ca2+ entry by a Clchannel blocker in human endothelial cells. European Journal of Pharmacology, 269, 381–4. Gill, D.R., Hyde, S.C., Higgins, C.F., Valverde, M.A., Mintenig, G.M. and Sepulveda, F.V. (1992) Separation of drug transport and chloride channel functions of the human multidrug resistance P-glycoprotein. Cell, 71, 23–32. Gschwentner, M., Nagl, U.O., Schmarda, A., Woll, E., Ritter, M., Waitz, W., Deetjen, P. and Paulmichl, M. (1994) Structure-Function Relation of a Cloned Epithelial Chloride Channel. Renal Physiology and Biochemistry, 17, 148–152. Hecker, M., Mulsch, A., Bassenge, E. and Busse, R. (1993) Vasoconstriction and Increased Flow—Two Principal Mechanisms of Shear Stress-Dependent Endothelial Autacoid Release. American Journal of Physiology, 265, H828–H833. Heinke, S., Raskin, G., Desmet, P., Droogmans, G., Vandriessche, W. and Nilius, B. (1997) Simultaneous Measurement of Membrane Capacitance and Whole Cell Currents During Cell Swelling In Macro vascular Endothelium. Cellular Physiology and Biochemistry, 7(1), 19–24. Heinke, S., Szucs, G., Norris, A., Droogmans, G. and Nilius, B. (1995) Inhibition of volume-activated chloride currents in endothelial cells by chromones. Br. J. Pharmacol., 115, 1393–1398. Hermsmeyer, K. and Miyagawa, K. (1996) Protein kinase C mechanism enhances vascular muscle relaxation by the Ca2+ antagonist, Ro 40–5967. J. Vasc Res., 33, 71–77. Hoffmann, E.K. and Dunham, P.B. (1995) Membrane mechanisms and intracellular signalling in cell volume regulation. International Review of Cytology, 161, 173–262. Hosoki, E. and Iijima, T. (1994) Chloride-Sensitive Ca2+ Entry by Histamine and ATP in Human Aortic Endothelial Cells. European Journal of Pharmacology, 266, 213–218. Hosoki, E. and Iijima, T. (1995) Modulation of cytosolic Ca2+ concentration by thapsigargin and cyclopiazonic acid in human aortic endothelial cells. European Journal of Pharmacology, 288, 131–137. Hoth, M. and Penner, R. (1993) Calcium Release-Activated Calcium Current in Rat Mast Cells. Journal of Physiology, 465, 359–386. Hoyer, J., Distler, A., Haase, W. and Gogelein, H. (1994) Ca2+ Influx Through Stretch-Activated Cation Channels Activates Maxi K+ Channels in Porcine Endocardial Endothelium. Proceedings of the National Academy of Sciences of the USA, 91, 2367–2371. Hoyer, J., Kohler, R., Haase, W. and Distler, A. (1996) Up-regulation of pressure-activated Ca(2+)-permeable cation channel in intact vascular endothelium of hypertensive rats. Proc. Natl. Acad. Sci. USA, 93, 11253–8. Hutcheson, I.R. and Griffith, T.M. (1994) Heterogeneous Populations of K+ Channels Mediate EDRF Release to Flow But Not Agonists in Rabbit Aorta. American Journal of Physiology, 266, H590– H596. Huxtable, R.J. (1992) Physiological functions of taurine. Physiological Reviews, 72, 101–154. Ingber, D.E. (1997) Tensegrity: the architectural basis of cellular mechanotransduction. Annual Review of Physiology, 59, 575–99. Jackson, P.S. and Strange, K. (1993) Volume-sensitive anion channels mediate swelling-activated inositol and taurine efflux. American Journal of Physiology, 265, C1489–500. Jackson, P.S. and Strange, K. (1995a) Characterization of the voltage-dependent properties of a volumesensitive anion conductance. Journal of General Physiology, 105, 661–676. Jackson, P.S. and Strange, K. (1995b) Single-channel properties of a volume-sensitive anion conductance. Current activation occurs by abrupt switching of closed channels to an open state. Journal of General Physiology, 105, 643–660. Jacobs, E.R., Cheliakine, C., Gebremedhin, D., Birks, E.K., Davies, P.F. and Harder, D.R. (1995) Shear activated channels in cell-attached patches of cultured bovine aortic endothelial cells. Pflügers Archiv European Journal of Physiology, 431, 129–131. Jentsch, T.J. and Günther, W. (1997) Chloride Channels: an Emerging Molecular Picture. BioEssays, 19, 117–126. Jentsch, T.J., Gunther, W., Pusch, M. and Schwappach, B. (1995) Properties of voltage-gated chloride channels of the ClC gene family. Journal of Physiology, 482, S19–S25. Jordt, S.-E. and Jentsch, T.J. (1997) Molecular dissection of gating in the ClC-2 chloride channel. EMBO Journal, 16, 101–11. Kaetzel, M.A. and Dedman, J.R. (1995) Annexins: novel Ca2+-dependent regulators of membrane function. News in Physiological Sciences, 10, 171–176. Kirk, K., Ellory, J.C. and Young, J.D. (1992) Transport of organic substrates via a volume-activated channel. Journal of Biological Chemistry, 267, 23475–23478. Krapivinsky, G.B., Ackerman, M.J., Gordon, E.A., Krapivinsky, L.D. and Clapham, D.E. (1994a) Molecular
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3 Tyrosine Phosphorylation of Platelet Endothelial Cell Adhesion Molecule-1 (PECAM-1) and Mechanosignal Transduction Keigi Fujiwara*, Michitaka Masuda, Masaki Osawa, Noboru Harada and Rosangela Bruno Lopes Department of Structural Analysis, National Cardiovascular Center Research Institute, Suita, Osaka 565–8565, Japan, Tel.:+81–6–6833–5012, ext. 2508, Fax: +81–6–6872–8092. *Corresponding author: E-mail address: [email protected]
INTRODUCTION Blood vessels were once considered to be an infrastructure of the body whose main function was to distribute blood throughout the body, and their biology was not a major concern for biologists, medical researchers, and physicians. When blood vessels were referred to as ‘living pipes’, this was understood to mean that they were made of alive cells and that new vessels could be made while existing ones could be remodeled and repaired. Recent progress in vascular biology, however, gave a new meaning to this term. We now know that blood vessels are metabolically highly active and perform multifaceted functions, such as synthesizing many types of physiologically active substances, receiving and transmitting chemical as well as mechanical signals, and controlling the passage of molecules and cells across the vessel wall. One of the major reasons for investigating the basic biology of an organ or a tissue is in relation to its diseases, and the recent interest in vascular biology is of no exception. Atherosclerosis is a cardiovascular disease that affects arteries. Pathologists have noted that atherosclerotic lesions do not develop anywhere in the artery but that there are certain regions in the artery where lesions are more likely to develop. Many factors are known to contribute to the development and progression of the disease. A number of them, such as high concentrations of lipids in the blood, hypertension, male hormones, the genetic make-up of an individual, stress, and smoking, are factors that act on the entire body, not just specific areas of the blood vessel. Thus, these factors are not responsible for the localized development of atherosclerotic lesions. Studies on fluid dynamics of blood flow inside blood vessels revealed that the high risk regions for atherosclerosis, such as the so-called “hip region” of blood vessel bifurcation points, were the areas of decreased fluid shear stress (Caro, Fitz-Gerald and Schrote, 1971; Zarins et al., 1983; Asakura and Karino, 1990). 55
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Early studies (Fry, 1968; Flaherty et al., 1972; Silkworth, McLean and Stehbens, 1975) have clearly demonstrated that blood flow is a determinant of endothelial cell morphology, indicating that endothelial cells have a mechanism for detecting fluid flow. It is now firmly established that fluid flow influences not only the structure but also the biosynthetic activity, gene expression, and physiology of endothelial cells (Davies, 1995). In spite of richly accumulated data on various effects of fluid flow on endothelial cells, however, the molecular mechanisms for flow sensing and the subsequent signal transduction by these cells are not yet elucidated. There are, undoubtedly, many reasons for this, but several years ago, we thought that one of the major reasons was the fact that no molecule had been identified which might be involved in sensing or early signal transduction of fluid flow. We, therefore, decided to look for molecules in endothelial cells that were biochemically modified within a few minutes of flow application. Here, we will review what we have found in our studies so far and present our view on mechanosensing by endothelial cells.
STRATEGIES AND EXPERIMENTAL APPROACHES Our goal was to identify polypeptides that were biochemically modified when fluid flow was applied to endothelial cells. The biochemical modification we chose to look for in proteins was tyrosine phosphorylation because many activated receptors and proteins in signal transduction pathways are tyrosine phosphorylated. In addition, it was technically simple to detect tyrosine phosphorylated proteins. Since our preliminary experiments indicated that a large number of proteins were tyrosine phosphorylated in endothelial cells exposed to fluid flow, it was necessary to limit our search to proteins that met certain requirements. Such requirements reflected our working model for flow sensing and mechanotransduction, which involved some molecule(s) in or associated with the plasma membrane being the sensing molecule(s) of fluid shear stress and some signal being transmitted from the cell surface to the cell interior by a chain of interactions of signal transduction molecules. The first requirement for the molecule was that it was a membrane protein. This condition was placed because we thought that the force from flow would affect the plasma membrane of endothelial cells and that flow sensing and early signal transduction molecules would be associated with the plasma membrane. The second requirement was that it was tyrosine phosphorylated shortly after flow stimulus was applied to cells. This requirement eliminates proteins that are tyrosine phosphorylated in later steps of signal transduction. A confluent culture of bovine arterial endothelial cells was exposed to steady laminar flow by using a cone-plate type viscometer (Harada, Masuda and Fujiwara, 1995). The sample cup of a viscometer was modified so that a 6cm tissue culture dish fits snugly in it. Rotation of a stainless steel cone-plate placed inside the dish creates steady laminar flow over the cell, and depending on the speed of rotation, the bottom surface of the culture dish is exposed to fluid shear stress ranging 0.4–30 dyn/cm2. Flow experiments lasting up to 30 minutes were easily conducted using this system. Cells were exposed to various levels of fluid shear stress for varying (0.5–10 minutes)
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length of time and then their extracts were analyzed for tyrosine phosphorylated proteins.
GP128 To identify membrane associated proteins that are tyrosine phosphorylated in flow stimulated endothelial cells, extracts of flow stimulated and non-stimulated endothelial cells were mixed with various lectin matrices, and bound polypeptides were probed with anti-phosphotyrosine in immunblotting analyses. Several types of lectins (such as Concanavala A, wheat germ agglutinin, phytohaemagglutinin, lentil lectin, and RCA120) were tested, but most of them failed to reveal polypeptides that were tyrosine phosphorylated in a flow stimulus dependent manner. RCA120 bound a limited number of polypeptides and among them was a protein with an apparent molecular mass of 128 kDa (GP128) that was tyrosine phosphorylated in the extract of cells exposed to flow (Harada, Masuda and Fujiwara, 1995). There was always a low level of tyrosine phosphorylation associated with GP128. However, when endothelial cells were exposed to fluid shear stress of more than 5 dyn/cm2, the level of phosphorylation of the polypeptide significantly increased. Although in some cases, we were able to detect the increase as early as 30 seconds of flow stimulation, it usually took 1–2 minutes before the increase was unambiguously detectable in all experiments. In separate experiments, we also identified several proteins including FAK (focal adhesion kinase) whose tyrosine phosphorylation increased when cells were stimulated by flow. However, their tyrosine phosphorylation occurred considerably later compared to GP128 phosphorylation. Our analyses indicate that tyrosine phosphorylation of GP128 is a fastest identifiable chemical change occurring in endothelial cells exposed to flow. This suggests that if GP128 tyrosine phosphorylation has some role in the molecular mechanism of flow signal transduction, it must be an event near the beginning of the mechanotransduction.
PECAM-1 Using RCA120 affinity resins, a gel filteration column, and SDS gel electrophoresis, we purified the 128 kDa glycoprotein, and its partial amino acid sequence was determined. Based on these limited amino acid sequence data, we performed RT-PCR and obtained a 3.4 kb cDNA which was then sequenced (Osawa et al., 1997). Both the amino acid and the cDNA sequence data showed a high degree of homology with both the human (Newman et al., 1990) and mouse (Xie and Muller, 1993) PECAM1 (platelet endothelial cell adhesion molecule-1). PECAM-1, which is also called CD31 and endoCAM, is a cell-cell adhesion molecule most abundantly expressed by endothelial cells. It is also expressed by platelets, monocytes, neutrophils, and a certain subset of T lymphocytes. In cultured solitary endothelial cells, PECAM-1 is diffusely distributed in the plasma membrane, but once a cell-cell contact is made, PECAM-1 is highly concentrated at the contact site. In
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confluent cultures, it is concentrated in the region where close plasma membrane apposition is formed between neighboring cells (Figure 3.1). In cultured endothelial cells, the band of PEC AM-1 localization is considerably wider than the cell-cell adhesion visualized by antibodies against ZO-1 (Figure 3.1). PECAM-1 appears to play some important role in forming and maintaining the contact inhibited state of endothelial cells in culture. Indeed, monolayer formation by cultured bovine endothelial cells was inhibited by antibodies against PECAM-1 (Albelda et al., 1990). In vivo, it is believed to have a role in transmigration of white blood cells (Muller et al., 1993; Vaporciyan et al., 1993). PECAM-1 was first cloned from human umbilical vein endothelial cells (Newman et al., 1990), and subsequently from mouse (Xie and Muller, 1993), and bovine (Osawa et al., 1997). At the amino acid sequence level, bovine PECAM-1
Figure 3.1. Immunofluorescence micrographs of confluent bovine arterial endothelial cells in culture stained with anti-PECAM-1 (A) or anti-ZO-1. Both antibodies stain cell-cell apposition areas, but their staining patterns are distinctly different, indicating that these two adhesion molecules form different structures for cell-cell association. Scale: Bar=10 µm.
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has 71% and 63% identity, and 80% and 77% similarity with human and mouse PECAM-1, respectively. PECAM-1 belongs to the immunoglobulin (Ig) superfamily and has 6 loops of the C2 type Ig-like domains, forming a large extracellular portion of the molecule (Figure 3.2). It has a single transmembrane domain and a short cytoplasmic piece. Within the extracellular domain of the molecule, there are many putative N-linked glycosylation sites. On an SDS gel, PECAM-1 migrates as a band of about 130 kDa, but the molecular mass calculated from its amino acid sequence is only about 80 kDa. This much higher apparent molecular weight is due presumably to glycosylation of the molecule. The cytoplasmic domain of PECAM1 is short and the bovine form consists of 118 amino acids. The primary structure of this domain is better preserved than that of the rest of the molecule among PECAM-1s from different species. The sequence homology is particularly high in the 40 or so of amino acids following the transmembrane domain (Osawa et al.,
Figure 3.2. A diagram showing the overall structure of PECAM-1. The 6 Ig loops formed by 6 disulfide bonds, the homophilic and putative heparan sulfate binding sites, and putative glycosylation sites are indicated.
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1997), suggesting functional importance of this part of the molecule, such as association with certain cytoskeletal components. PECAM-1 appears to have both homophilic and heterophilic binding activities. For homophilic binding, PECAM-1 molecules of neighboring cells interact via loops 1 and 2 (Figure 3.2) and establish cell-cell association (Fawcett et al., 1995; Sun et al., 1996). Some data also exist which indicate heterophilic interaction of PECAM-1 (Piali et al., 1995; Buckley et al., 1996). It is interesting that heterophilic binding is lost when 1/3 of the cytoplasmic domain is deleted from the C-terminus of the molecule (DeLisser et al., 1993; Sun et al., 1996; Famiglietti et al., 1997).
Tyrosine Phosphorylation of PECAM-1 Amino acid sequence data show that the cytoplasmic domain of PECAM-1 contains 12 serine, 4 threonine, and 5 (6 in the case of bovine) tyrosine residues. Of these phosphorylatable amino acids, only serine phosphorylation was reported earlier in thrombin stimulated platelets (Newman et al., 1992). The other residues were considered not to be phosphorylated, although the sequence data suggest that at least two tyrosine residues (tyr663 and tyr686) might be phosphorylated. We have recently demonstrated that tyr686 is phosphorylated in mechanically stimulated endothelial cells (Osawa et al., 1997). Also recently, PECAM-1 tyrosine phosphorylation was detected in aggregating platelets (Jackson et al., 1997) and in mast cells during aggregation of IgE receptors (Sagawa et al., 1997). In HEK293 cells transfected with human PECAM-1, both tyr663 and tyr686 were shown to be phosphorylated (Jackson, Kupcho and Newman, 1997). In endothelial cells, PECAM-1 is always tyrosine phosphorylated at a low level. When these cells were exposed to fluid shear stress of less than 4 dyn/cm2, this basal level of phosphorylation did not change. However, when the cells were stimulated by more than 5 dyn/cm 2 of fluid shear stress, the level of PECAM-1 tyrosine phosphorylation rapidly increased. Higher levels of shear stress appeared to give more pronounced increases. The kinetics of tyrosine phosphorylation in endothelial cells exposed to 16 dyn/cm2 is shown in (Figure 3.3). Immediately after flow stimulation, the level of tyrosine phosphorylation begins to increase, and within 5 minutes, it reaches a plateau. It has been shown that when cells are exposed to fluid flow, the apical cell surface is mechanically disturbed (Liu, Yen and Fung, 1994). Since fluid flow was able to increase tyrosine phosphorylation of PECAM-1 in endothelial cells, we thought that other means that disturbed the endothelial cell membrane might elicit the similar chemical modification in PECAM-1. Exposing cells to a high or low osmotic condition is a simple way to mechanically disturb the plasma membrane. When endothelial cells were treated with culture medium that was either diluted two times (hyposmotic) or supplemented with 0.3 M sucrose (hyperosmotic), we detected PECAM-1 tyrosine phosphorylation (Osawa et al., 1997). The time course of PECAM-1 phosphorylation was similar to that observed in cells exposed to flow (Figure 3.3). These experiments suggest that tyrosine phosphorylation of PECAM-1 is a common biochemical response in endothelial cells stimulated by mechanical stimuli. Because both flow and osmotic stimuli induce the
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Figure 3.3. The time course of PECAM-1 tyrosine phosphorylation. At time 0, confluent monolayers of bovine arterial endothelial cells were exposed to 16 dyn/cm2 of fluid shear stress (closed circle), to hypertonic medium (open circle), to hypotonic medium (triangle), or to normal medium (square) and PECAM-1 phosphotyrosine levels were measured by quantitative Western blot analyses. The level of tyrosine phosphorylation is expressed relative to the 0 time control value.
same response and because osmotic stimulation is easy to apply, this latter stimulation has been used to elucidate the signaling pathway involving PEC AM-1. The tyrosine phosphorylation of PECAM-1 is indeed a rapid response of endothelial cells induced by flow. However, there are other flow induced responses, such as a transient cytoplasmic Ca2+ increase (Ando, Komatsuda and Kamiya, 1988) and K+ channel activation (Olesen, Clapham and Davies, 1988), that also occur immediately after flow application. Thus, it is possible that the PECAM-1 phosphorylation is a downstream event of these responses. In order to test whether increased cytoplasmic concentrations of Ca2+ could induce PECAM-1 tyrosine phosphorylation, we treated endothelial cells with thrombin, ATP, histamine, or bradykinin, all of which are known to increase the cytoplasmic Ca2+ concentration, or with ionophores. None of these agents caused PECAM-1 tyrosine phosphorylation in endothelial cells (Table 3.1; Harada, Masuda and Fujiwara, 1995; Osawa et al., 1997). Thus, cytoplasmic Ca2+ mobilization is not involved in the PECAM-1 tyrosine phosphorylation. K+ channel inhibitors that are known to inhibit flow induced K-channel activation (Olesen, Clapham and Davies, 1988; Schilling, Mo and Eskin, 1992), such as Ba+ and TEA (tetraethyl ammonium ion), failed to block PECAM-1 tyrosine phosphorylation in osmotically stimulated endothelial cells (Table 3.1). Furthermore, gadolinium ion, a blocker of stretch-activated cation channels, failed to inhibit the PECAM-1 tyrosine phosphorylation. Activating protein kinase C (PKC) by PMA (phorbol myristate acetate) did not cause the PECAM-1 phosphorylation. These results indicate that tyrosine phosphorylation of PECAM-1 in mechanically stimulated endothelial cells is not a downstream event of Ca2+ mobilization, K+ channel activation, stretch activated cation channel activity, and PKC activation. It appears, therefore, that it is a unique response elicited in endothelial cells when their membrane is mechanically disturbed. Tyrosine
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Table 3.1 Effect of channel inhibitors and Ca2+ mobilization on PECAM-1 tyrosine phosphorylation
phosphorylation of PEC AM-1 may be a step in a novel mechanosensitive signaling pathway in endothelial cells.
PECAM-1 and Signal Transaction Our study described above suggests that PECAM-1 is a reasonable candidate for a signal transduction molecule and that its tyrosine phosphorylation is tied to its function as a signaling molecule. However, PECAM-1 does not appear to be an autophosphorylatable protein since there is no kinase domain in its cytoplasmic tail. In order to identify tyrosine kinase(s) for PECAM-1 and also other molecules associated with PECAM-1 in endothelial cells, coimmunoprecipitation was performed using antibodies against PECAM-1 (Osawa et al., 1997). In the immunoprecipitates, there was a tyrosine phosphorylated polypeptide with the same electrophoretic mobility as c-Src. In vitro, c-Src phosphorylated and bound to PECAM-1 and a GST fusion protein containing the cytoplasmic domain of PECAM-1. Thus, at least in vitro, c-Src can serve as a tyrosine kinase for PECAM-1, and in vivo, c-Src and PECAM-1 appear to be associated in endothelial cells as they can be coimmuno-precipitated. A GST fusion protein of PECAM-1 cytoplasmic domain that lack the exon 14, which contains tyr686, had no c-Src binding activity and was not phosphorylated. This experiment suggests that c-Src is a tyrosine kinase for tyr686 of PECAM-1. Since the exon 14 has the SH2binding YSEI motif containing the phosphorylatable tyr686 and since the exon 14less fusion protein failed to bind to c-Src, it is strongly suggested that c-Src binds to tyr686 of PECAM-1 via its SH2 domain. Our in vitro study has identified SH-PTP2 as a tyrosine phosphorylated PECAM1 binding protein (Masuda et al., 1997). The similar binding activity was found in platelets (Jackson et al., 1997). SH-PTP2 is a protein tyrosine phosphatase and is also known as Syp, SHP2, PTP2C and PTP1D. It is a homologue of Drosophila tyrosine phosphatase corkscrew (Freeman, Plutzky and Neel, 1992) which, when activated, transduces a positive signal to the MAP kinase pathway (Perkins, Larsen and Perrimon, 1992). In mammalian systems, SH-PTP2 is known to activate the MAP kinase (ERK)
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cascade (Noguchi et al., 1994). SH-PTP2 has two SH2 domains, and their binding to phosphotyrosine containing domains of another protein activates the phosphatase activity of the molecule (Ohnishi et al., 1996). We have proposed that the two SH2 domains of SH-PTP2 bind to the two tyrosine residues (tyr663 and tyr686) of PECAM1 when they are phosphorylated (Masuda et al., 1997). To support this view, we have shown that SH-PTP2 binds only to tyrosine phosphorylated PECAM-1 and not to unphosphorylated PECAM-1. Furthermore, we have found that the cellular distribution of SH-PTP2 changes in a mechanical stimulus-dependent way (Figure 3.4). SH-PTP2 is a cytoplasmic protein, and by immunofluorescence localization, it is generally distributed in the cytoplasm of unstimulated endothelial cells. In mechanically stimulated cells, however, staining appears at the cell-cell adhesion site where PECAM1 is localized. This indicates that when endothelial cells are mechanically stimulated, a portion of SH-PTP2 in the cell is translocated from the cytoplasm to the cell membrane. This localization shift of SH-PTP2 is a morphological manifestation of PECAM-1/SH-PTP2 association induced by mechanical stresses and supports the biochemical data. The nature of this PECAM-1/SH-PTP2 binding described above suggests that tyrosine phosphorylation of PECAM-1 is a positive signal for activating ERK in mechanically stimulated endothelial cells. Indeed, a transient activation of ERK which depends on tyrosine kinase activity has been reported when endothelial cells are exposed to flow (Ishida et al., 1996; Takahashi and Berk, 1996). We have also observed the similar ERK activation in endothelial cells treated with hyperosmotic medium. In this latter case, ERK activity peaked at 10 minutes and by 30 minutes, it returned to the basal level. We suggest that this ERK activation is a mechanism for increased transcription of certain genes in endothelial cells stimulated by fluid flow (Davies, 1995).
Figure 3.4. Immunofluorescence micrographs of confluent human umbilical cord vein endothelial cells stained with anti-SH-PTP2 before (A) or 4 minutes after (B) hyperosmotic stress. In unstressed cells, SH-PTP2 is in the cytoplasm, but after mechanical stress, accumulation at the cell-cell association site can be observed. Extended focus images reconstructed from the entire optical sections of cells obtained using a confocal microscope. Scale: Bar=10 µm.
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When PEC AM-1 expression in endothelial cells was down regulated by antisense S-oligonucleotides, the osmotically induced ERK activation was significantly reduced. In addition, the ERK activation was blocked by tyrosine phosphatase inhibitors, even though PECAM-1 tyrosine phosphorylation increased. PD 98059, a MEK inhibitor also blocked the ERK activation. Interestingly, however, ERK activation by VEGF was not inhibited by tyrosine phosphatase inhibitors. These observations suggest that the PECAM-1 expression is essential for mechanosensing and/or mechanosignal transduction in endothelial cells and that activation of SH-PTP2 by tyrosine phosphorylated PECAM-1 is an important step in the mehanosignal transduction to the MEK-ERK pathway. Although we have not yet investigated in detail, Src family kinases may also play important roles in mechanosignal transduction. c-Src is known to be activated in endothelial cells by mechanical stresses and can also phosphorylates tyr686 of PECAM-1. It binds to PECAM-1 at tyr686 via its SH2 domain (Osawa et al., 1997; Masuda et al., 1997). The Src family kinase is inactive when its N-terminal SH2 domain is bound to its own phosphotyrosine residue at the C-terminal tail. It is activated by breaking this intramolecular SH2/phosphotyrosine association. This is done either by dephosphorylating the C-terminal phosphotyrosine or by competitive binding of the SH2 to a higher affinity SH2-binding motif on another protein (Pawson, 1995). Dephosphorylation of the C-terminal phosphotyrosine can be achieved by the action of SH-PTP2 (Peng and Cartwright, 1995), and the phosphorylated tyr686 of PECAM-1 can serve as a specific competitive binding site for the SH2 domain of the Src family kinase (Masuda et al., 1997). It is quite possible that once PECAM-1 is tyrosine phosphorylated in endothelial cells, it may effectively recruit and activate Src family kinases. If Src family kinases are involved in tyrosine phosphorylation of PECAM-1 in mechanically stimulated endothelial cells, the Src/ PECAM-1/SH-PTP2 interaction may work as a self-accelerating system for Src family kinases. As mentioned above, PECAM-1 is not a self-phosphorylating membrane protein. Although Src family kinases can serve as tyrosine kinases for PECAM-1, they cannot interact with unphosphorylated PECAM-1. It is possible that they first bind to the subset of PECAM-1 molecules which are constitutively tyrosine phosphorylated and then catalyze tyrosine phosphorylation of unphosphorylated PECAM-1. However, it is also possible that there are other kinases that are specifically activated by mechanical stimuli. We are currently investigating whether or not such kinases are indeed present in endothelial cells.
CONCLUSIONS Our studies have indicated that there is a signal transduction pathway involving PECAM-1 tyrosine phosphorylation in endothelial cells that is specifically activated by mechanical stresses, such as fluid flow and osmotic changes. Tyrosine phosphorylated PECAM-1 appears to recruit SH-PTP2 and possibly also Src kinases to the plasma membrane of the cell-cell adhesion site. The formation of this molecular complex transmits a positive signal to ERK, which then activates transcription of
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Figure 3.5. Domain map of the cytoplasmic portion of PECAM-1 showing exons, the location of two phosphorylatable tyrosine residues (tyr663 and tyr686), the Src binding site, and the SH-PTP2 (SHP2) binding site. Possible ways for PECAM-1 to transmit signals are also indicated. Binding of SH-PTP2 to PECAM-1 activates SH-PTP2, sending a positive signal to ERK perhaps via Ras. PECAM-1 bound c-Src may also initiate a different signaling event. The actin cytoskeleton may bind to the N-terminal region of the cytoplasmic domain via one of the catenins.
certain genes (Figure 3.5). This molecular complex formation may also amplify signaling by Src family kinases. Although we did not discuss possible interaction of PECAM-1 with the cytoskeleton in detail in this chapter, some data are present that suggest PECAM-1 binding to the actin cytoskeleton. Like the cell-substrate attachment site, the cell-cell adhesion site is also a fixed point in the cell where externally applied mechanical forces can act on. PECAM-1 present in high concentrations at the cell-cell adhesion site not only is a good candidate for a mechanotransduction molecule, but also show characteristics expected for a signal transduction molecule. Our data summarized in this chapter make it tempting to suggest that PECAM-1 is involved in mechanosensing or mechanosignal transduction in endothelial cells as well as other cells expressing this molecule.
ACKNOWLEDGMENTS Various parts of the work presented here were supported by grants from the Ministry of Health and Welfare of Japan, Grants-in-Aid for Scientific Research from the Japanese Ministry of Education, Special Coordination Funds for Promoting Science and Technology from Science and Technology Agency of Japan, and the Program for Promotion of Fundamental Studies in Health Sciences of the Organization for Drug ADR Relief, R and R Promotion and Product Review of Japan.
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4 Protein Phosphorylation in Shear Stress Activated Endothelial Cells John Y-J. Shyy*, Yi-Shuan Li, Song Li, Shila Jalali, Michael Kim, Shunichi Usami and Shu Chien Department of Bioengineering and Institute for Biomedical Engineering, University of California, San Diego, La Jolla, CA 92093–0412, USA. *Corresponding author: Dr. John Y-J.Shyy, Department of Bioengineering and Institute for Biomedical Engineering, University of California, San Diego, La Jolla, CA 92093–0412, USA. Tel.: (619) 822–0785, Fax: (619) 534–3658, E-mail: [email protected].
Studies on cultured vascular endothelial cells (ECs) in flow channels have demonstrated that the application of shear stress leads to the phosphorylation of multiple cellular proteins. This mechanotransduction results from the activation of cellular protein tyrosine kinases (PTKs) and serine/threonine (Ser/Thr) kinases. Shear stress rapidly activates PTKs in the focal adhesion sites, including focal adhesion kinase (FAK) and Src-family PTKs (e.g., c-Src and Fyn). Shear stress activation of FAK is functionally linked to the Ras activation through the association of growth factor receptor binding protein-2 (Grb2) and Son of sevenless (Sos. Ras in turn activates cytoplasmic mitogen-activated protein kinases (MAPKs), including extracellular signal-regulated kinase (ERK) and c-Jun NH2-termmal kinase (JNK). The augmented Ser/ Thr kinase activities of ERK and JNK result in the increased phosphorylation of transcription factors c-Jun, Elk-1, and TCF/c-Fos and their binding to target cis-elements to cause the activation of appropriate genes in response to shear stress. Shear stress also causes the phosphorylation of other proteins, including Sp1, endothelial constitutive NO synthase (ecNOS), and platelet endothelial cell adhesion molecule-1 (PECAM-1). The interplay among various phosphorylation cascades catalyzed by multiple kinases in response to shear stress may play a significant role in vascular biology and pathobiology. KEYWORDS: Shear stress, endothelial cell, tyrosine kinase, MAPK, phosphorylation, mechanotransduction.
INTRODUCTION In the body, vascular endothelial cells (ECs) are constantly exposed to shear stress, which is the tangential component of hemodynamic forces acting on the vessel wall. While shear stress plays important roles in maintaining vascular homeostasis, it can also be pathophysiological factors in vascular disorders such as atherosclerosis. Flow channels with cultured ECs such as human umbilical vein endothelial cells (HUVECs) and bovine aortic endothelial cells (BAECs) have been used as in vitro models to study the endothelial responses to applied shear stress. Such studies have demonstrated that the application of shear stress leads to the phosphorylation of multiple cellular proteins in ECs, resulting in the activation of signaling pathways to modulate gene 69
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expression, cytoskeletal organization, vessel dilation/constriction, and cell proliferation/ apoptosis. These phosphorylation cascades have been shown to result from the activation of cellular pr otein tyrosine kinases (PTKs) and Ser/Thr kinases. PTKs can be generally divided into two major categories: receptor tyrosine kinases (RTKs) and nonreceptor PTKs. RTKs such as epidermal growth factor receptor (EGFR) and platelet-derived growth factor receptor (PDGFR) are PTKs with an extracellular ligand-binding domain and a cytoplasmic domain that functions as tyrosine kinases. Following the binding of the cognate growth factors (e.g., EGF and PDGF), these RTKs autophosphorylate the tyrosine residues at their cytoplasmic domains (see Ullrich and Schlessinger 1990 for review). In contrast, nonreceptor PTKs such as focal adhesion kinase (FAK) and c-Src represent cellular enzymes that have intrinsic kinase activity but do not have extracellular domain. Tyrosine phosphorylation of cellular proteins in response to extracellular stimuli has been investigated by immunoblotting the cell lysates with anti-phosphotyrosine mAb. In response to shear stress, many proteins in ECs, including those in focal adhesion sites and in the cytoplasm, are rapidly phosphorylated on tyrosines (Shyy et al., 1995a; Li et al., 1997). The level of shear stress-induced tyrosine phosphorylation of these proteins is significantly reduced by pretreating ECs with genistein, an inhibitor of PTKs, indicating the role of PTKs activation in the tyrosine phosphorylation (Jo et al., 1997; Li et al., 1997). The shear stress-activated Ser/Thr kinases include the members in the mitogen-activated protein kinases (MAPKs) family. The activation of multiple PTKs and Ser/Thr kinases by shear stress can exert profound influence on endothelial biology. This chapter provides a general review of the protein phosphorylation in ECs in response to shear stress and its functional relevance to vascular biology.
SHEAR STRESS ACTIVATION OF PTKs IN THE FOCAL ADHESION SITES Activation of FAK In response to shear stress, concomitant with the elongation of ECs and the alignment of stress fibers with flow direction, the focal adhesions on the abluminal side of ECs undergo dynamic, local reorientation at accelerated rates of association/dissociation without a noticeable change in the total attachment area (Davies et al., 1994). At the molecular level, such a dynamic rearrangement of focal adhesions may be related to the spatial and temporal responses of the associated proteins. Originally identified through its association with v-Src, FAK is present in focal adhesions and is tyrosinephosphorylated in response to cell adhesion and also to a number of growth factors (e.g., PDGF) and peptide hormones (e.g., angiotensin II, thrombin, bombesin, and endothelin) (see Parsons et al., 1994 for review). The increase in tyrosine phosphorylation of FAK in ECs in response to shear stress has been shown in confluent monolayers of BAECs subjected to a steady shear stress of 12 dyn/cm2 for different durations. Following the immunoprecipitation of FAK from cell lysates with polyclonal anti-FAK, the subsequent immunoblotting of the precipitated protein complex with
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an anti-phosphotyrosine PY20 mAb has shown that shear stress induces a rapid and transient tyrosine phosphorylation of FAK (Figure 4.1A). The tyrosine phosphorylation reaches a maximum level within 1 min and then decreases gradually. An increase in the kinase activity of FAK by shear stress has been demonstrated by autophosphorylation assay using the immunoprecipitated FAK and [–32P]ATP (Figure 4.1B). After a 5-min shearing treatment, the kinase activity of FAK increases by 3fold. The shear stress activation of FAK is dependent upon actin structure integrity, since it is attenuated by pre-treating BAECs with cytochalasin B (Figure 4.1B).
Activation of c-Src and Fyn At focal adhesions, integrins link the extracellular matrix proteins to the cytoskeletal proteins located on the cytoplasmic face of the cell membrane. Accordingly, FAK and other proteins, (e.g., FAK, paxillin, tensin, and Src-family PTKs) in the focal adhesions play an important role in the integrin-mediated signal transduction (Guan et al., 1992; Schaller et al., 1994a). The c-Src family members are also activated by shear stress. The performance of immunocomplex kinase assays (IP kinase assays) by using enolase and [–32P]ATP as the substrates shows that the kinase activity of c-Src is increased
Figure 4.1. Shear stress increases the tyrosine phosphorylation and the kinase activity of FAK. After serum-starvation for 15 hr, BAEC monolayers were either kept as static controls (represented by time 0) or subjected to a shear stress of 12 dyn/cm2. In (A), 500 µg of cell lysates from each sample following different durations of shearing was subjected to immunoprecipitation (IP) with a polyclonal anti-FAK antibody and immunoblotting (IB) with PY20 anti-phosphotyrosine mAb. The bound antibodies were detected by using the ECL system. In (B), BAEC monolayers were pre-treated with either 0.1 % DMSO or 1 µM cytochalasin B for 1.5 hr. The cells were then kept as static controls or subjected to a shear stress of 12 dyn/ cm2 for 5min. FAK was immunoprecipitated with a polyclonal anti-FAK antibody for kinase activity assays. Shear stress activation of FAK is indicated by the increased auto-phosphorylation of FAK in the DMSO samples, but not following cytochalasin B treatment.
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by shear stress as early as 1 min, peaks at 5–10 min, and decreases afterwards (Figure 4.2A). Densitometry shows that shear stress causes an increase in c-Src activity to 3.2-fold in 1 min, 4.1-fold at 5 min, and 7.0-fold at 10 min, but only 3.9 folds at 20 min. Similar IP kinase assays have shown that Fyn, a Src-family member, is also activated by shear stress. The normalized kinase activity of Fyn is increased to 3.2, 2.0, 1.8, and 1.7 folds in BAECs sheared for 5, 10, 30, and 60 min, respectively (Figure 4.2B). The rapid activation of focal adhesion-associated FAK and c-Src in these in vitro experiments demonstrates the importance of these non-receptor PTKs in the mechanotransduction in ECs in response to shear stress. Perfusion of isolated coronary arterioles ex vivo have shown that flow increases the vasodilatory responses and the binding of a fluorescein isothiocyanate-labeled phosphotyrosine antibody to the vessel (Muller et al., 1996). Genistein treatment reverses the flow-induced dilation as well as the increase in tyrosine phosphorylation. These experiments indicate that PTKs activation is a critical step in flow-induced vasodilation.
Figure 4.2. Shear stress increases the tyrosine kinase activity of c-Src and Fyn. BAEC monolayers were subjected to a shear stress of 12dyn/cm2 for periods of time as indicated or kept as a static control (represented by time 0). In (A), the kinase activities of c-Src were assessed by immunoprecipitation with anti-p60src mAb followed by kinase activity assays using enolase and [–32P]ATP as substrates. In (B), the kinase activities of Fyn were assessed using polyclonal anti-Fyn for immunoprecipitate kinase assays. Shear stress induction of c-Src and Fyn in ECs is demonstrated by the increased phosphorylation of enolase.
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SHEAR STRESS ACTIVATION OF ADAPTOR MOLECULES Induction of FAK-Grb2/Sos Formation The stimulation of monocytes with monocyte/macrophage colony stimulating factor (MCSF) and the adhesion of NIH3T3 fibroblasts to fibronectin have been shown to promote the interaction of growth factor receptor binding protein-2 (Grb2) with FAK (Kharbanda et al., 1995; Schlaepfer et al., 1994). This involves the FAK autophosphorylation on Tyr397, which leads to Src recruitment. The association of FAK with Src-family PTKs at focal adhesions further increases the phosphorylation of FAK at Tyr-925, creating a Grb2 binding site (Calalb et al., 1995; Cobb et al., 1994; Schaller et al., 1994b; Schlaepfer et al., 1994). Grb2, with a molecular weight of 24 kD, has a relatively simple structure with one Src-homology domain-2 (SH2) domain flanked by two SH3 domains (Matuoka et al., 1992). Grb2 associates with activated RTKs such as EGFR, PDGFR, and monocyte/ macrophage colony stimulating factor receptor (MCSFR) through interaction of its SH2 with the phosphotyrosines of these RTKs (Lowenstein et al., 1992; Suen et al., 1993; van der Geer and Hunter, 1993). The two SH3 domains, on the other hand, determine the localization of Grb2 to membrane ruffles (Bar-Sagi et al., 1993). When binding to activated RTKs, Grb2 also binds to Son of sevenless (Sos), a guanine nucleotide exchange factor. Sos activates Ras by converting the GDP-bound inactive state to the GTP-bound active state (Egan et al., 1993; Chardin et al., 1993). The shear stress-induced increase in FAK-Grb2 association has been shown by immunoblotting the FAK immunoprecipitates of BAEC cell lysates with a polyclonal anti-Grb2/Sem5. An increase in the amount of Grb2 co-immunoprecipitated with FAK occurs as early as 1 min after shearing (Li et al., 1997). The increased association of Grb2 with FAK lasts for at least 5 min, and then decreases to a level similar to that in the static controls. Immunoblotting of Sos immunoprecipitates of BAEC cell lysates with a polyclonal anti-Grb2/Sem5 reveals a constant level of Grb2/Sos association in both static and sheared cells, suggesting that shear stress increases the association of FAK with the binary complex of Grb2/Sos. Taken together, these results demonstrate that shear stress activation of FAK may functionally lead to the Ras activation through Grb2/Sos.
Involvement of Other SH2-containing Adaptor Proteins In addition to Grb2, the SH2-containing adaptor proteins also include p130cas, proto-oncogene product c-Crk, and She. p130cas has an SH3 domain and multiple SH2 binding motifs in the substrate domain, whereas Crk contains one SH2 and two SH3 domains and has been shown to interact with Sos (Matsuda and Kurata, 1996). p130cas was originally identified as a major tyrosine-phosphorylated protein in v-Crk- and v-Src-transformed cells. Subsequently, p130cas has been found to be tyrosine-phosphorylated in the integrin-mediated signal transduction and is regarded as one of the focal adhesion proteins. Shear stress causes the tyrosine phosphorylation of p130cas and its association with Crk (Takahashi et al., 1997). Pretreating ECs with cytochalasin D has little effect on the shear stress induction of p130cas tyrosine phosphorylation. Shearing fibroblasts isolated from
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Src-/-/Fyn-/- knockout mice reveals that the tyrosine phosphorylation of p130Cas is independent of the Src family tyrosine kinases. She is another adaptor protein which undergoes an increase in tyrosine phosphorylation when bound to RTKs (Daub et al., 1996; Ricci et al., 1995; van Biesen et al., 1995). She is associated with a subset of integrins, and this association is necessary and sufficient for the activation of ERK pathway in response to integrin ligation (Wary et al., 1996). Thus, p130cas and She are candidates for functioning as adaptor molecules in the shear stress-mediated signal transduction in ECs. The SH2-containing molecules can be divided into two main groups. The first group consists of the adaptor proteins, e.g., Grb2, p130cas, and She as discussed above. These molecules are composed of almost exclusively of SH2 and SH3 domains. The second group is comprised of proteins with enzymatic functions, e.g., GTPaseactivating protein of Ras (RasGAP), phosphatidylinositol 3-kinase (PI 3-kinase), phospholipase C- (PLC-), c-Src, and protein tyrosine phosphatases (SH-PTP1 and SH-PTP2) (see Montminy 1993 for review). It may well be that shear stress activates all these SH2-containing enzymes as well as the SH2-containing adaptor proteins. Multiple signaling events in ECs can be activated if the SH2-containing enzymes are involved in the shear stress induction mechanism. For example, the PI 3-kinase pathway can lead to the generation of intracellular diacylglycerol (DAG) and inositol 1, 4, 5trisphosphate (IP3); c-Src associates with Ras to activate the downstream ERKs and JNKs; and PLC- can activate the PKC pathway. Indeed, IP3 and PKC have been shown to be up-regulated by shear stress (Bhagyalakshmi et al., 1992; Prasad et al., 1993; Kuchan and Frangos, 1993). It remains to be investigated whether shear stress activates PI 3-kinase and PLC- which modulate the up-regulation of IP3 and PKC, respectively.
SHEAR STRESS ACTIVATION OF THE RAS-MAPK PATHWAYS Activation of ERKs, JNKs, and p38 MAPKs are a group of cytoplasmic kinases specific for Ser/Thr phosphorylation. MAPKs include three major subfamilies: extracellular signal regulated kinases (ERK 1/2), c-Jun NH2-terminal kinases/stress activated protein kinases (JNKs/ SAPKs), and p38 kinase. The activation of MAPKs is achieved in turn by cascades of protein kinases which are linked to the cell membrane-associated proteins such as Ras, c-Src, FAK, integrins, etc. ERK 1/2 were the first MAPKs identified (Ahn et al., 1991; Boulton et al., 1991). Upon growth factor stimulation, Ras becomes activated by replacing the bound GDP with GTP. Ras-GTP then activates ERKs by dual phosphorylation at the T-E-Y motif of ERKs through the cascade of MAP kinase kinase kinase and MAP kinase kinases (MEKs) (see Robinson and Cobb, 1997; Su and Karin, 1996). The phosphorylation/activation of ERKs in BAECs in response to shear stress has been demonstrated by immunoblotting with PY20 mAb (Shyy et al., 1995a) and by IP kinase assays using mylein basic protein (MBP) as the substrate (Tseng et al., 1995; Li et al., 1996; Takahashi and Berk, 1996). As shown in Figure 4.3A, the shear stress induction of ERK is transient, with its peak activity
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Figure 4.3. Shear stress increases the activity of ERK, JNK, and P38. BAEC monolayers were subjected to a shear stress of 12 dynes/cm2 for various lengths of time as indicated. In (A), ERK was immuno-precipitated and the kinase activity assay was performed in the presence of myelin basic protein (MBP) and [– 32P]ATP. In (B), the cell lysate was incubated with agarose-bound GST-c-Jun to precipitate JNK followed by the addition of [– 32P]ATP. In (C), p38 was immunoprecipitated followed by kinase activity assay using GST-ATF2 and [– 32P]ATP as substrates. Static controls are represented by time 0, and the phosphorylated GST-c-Jun, MBP, and GST-ATF2 are indicated by arrows. The induced ERK activity peaked at 10 min, whereas those for JNK and p38 were at 30 min.
occurring at 10 min after shearing. The ERKs can be activated by shear stress ranging from 1 to 17 dyn/cm2 (Tseng et al., 1995; Li et al., 1996; Jo et al., 1997). JNKs bind to the NH2-terminal of c-Jun and phosphorylate its ser-63 and ser-73 (Dérijard et al., 1994). JNKs, in turn, are activated by JNKKs (also known as MKKs) through dual phosphorylation on the T-P-Y motif (Lin et al., 1995; Hirai et al., 1997). MEKK1 has been identified as the upstream kinase that activates JNKK (Lin et al., 1995). MEKK1 is a Ser/Thr kinase that may cross talk to the ERK pathway by phsophorylating MEKs. Shear stress activation of JNKs has been demonstrated by the increased phosphorylation of GST-c-Jun fusion protein in IP kinase assays (Li et al., 1996; Jo et al., 1997). A shear stress of 0.5 dyn/cm2 is the threshold for JNK induction and similar degrees of induction are achieved with shear stresses up to 20 dyn/cm2 (Jo et al., 1997). Like that of ERKs, the activation of JNKs by shear stress has been shown to be transient, but with a time course somewhat longer than that of ERKs. At a shear stress of 12 dyn/cm2, JNKs activities in ECs reach a peak at 30 min and decreases afterward (Figure 4.3B).
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p38 kinase is the third major member of the MAPKs family. Dual phosphorylation of the T-G-Y motif of p38 leads to its activation. Overexpressing the active forms of MKK3 or MKK6 in CHO cells results in the activation of p38 (Raingeaud et al., 1996). Small GTPases Rac and Cdc42 and p21-activated kinase 1 (Pak 1) are also involved in the p38 signaling pathway (Zhang et al., 1995). Shear stress causes a transient activation of p38 in BAECs exposed to shear stress (Figure 4.3C). Using GST-ATF2 as the substrate in the IP kinase assay, the peak activity of p38 is found to occur at 30 min after shearing, which is similar to the temporal response of JNK (Figure 4.3A). However, the upstream signaling molecules activating p38 in response to shear stress is unknown. Whether other MAPK family members, such as ERK3, ERK5 and Fos-regulating kinase, are involved in the mechanotransduction in ECs remains to be investigated. Activation of ERKs, JNKs, and p38 pathways may lead to different functional consequences. ERK pathway mainly responds to growth stimuli, whereas JNK and p38 pathways are sensitive to the inflammatory cytokines and environmental stresses which may lead to apoptosis. Shear stress activates all three major MAPKs in ECs. The transient nature of the activation of ERKs, JNKs, and p38 caused by a sudden application of shear stress to cultured ECs in vitro may not reflect the responses of ECs in the blood vessels in vivo, where ECs are constantly exposed to the flow environment. In fact, the constant presence of laminar shear stress in the straight part of the arterial system may lead to a down-regulation of the MAPKs, which is mimicked by long term shearing in the in vitro experiments. In contrast, the branch points with unsteady and disturbed flow may be more prone to MAPKs activation (Chien et al., 1998).
Ras Regulation of ERKs and JNKs MAPKs can be activated by many upstream signaling events, which in turn can be augmented by multiple extracellular stimuli. What are the upstream signaling molecules that lead to the activation of ERKs and JNKs by shear stress? The shear-activated ERKs has been shown to be PKC-dependent but Ca2+-independent (Tseng et al., 1995). The requirement of PKC is demonstrated by the attenuation of the shear-activated ERKs by inhibiting PKC with staurosporine or down-regulating PKC with phorbol 12,13-dibutyrate. On the other hand, Ca2+ chelation has no inhibitory effect (Tseng et al., 1995). Ras is a common molecule regulating the growth factor activation of ERKs and stress activation of JNKs (Minden et al., 1994). Performance of guanine nucleotide binding assays on lysates of 32P-labeled BAECs shows a marked increase in the ratio of Ras · GTP/Ras · GDP after 1 min of shearing. Thereafter, the GTP-bound active form of Ras gradually returns to the GDP-bound form and becomes undetectable by 10 min, as in the static controls (Li et al., 1996). RasN17 is a dominant negative mutant of Ras in which Ser-17 in the wild-type has been replaced by Asn. Cotransfection of RasN17 with exogenous epitope-tagged Myc-ERK2 or HA-JNK1 into BAECs significantly reduces the shear stress activation of Myc-ERK2 or HA-JNK1 (Li et al., 1996). These results indicate that Ras regulates shear stress activation of ERKs and JNKs in ECs.
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JNKs, But Not ERKs, Regulate Shear Stress Activation of AP-1/TRE Transcription factors, which encompass a wide variety of DNA-binding motifs, have been implicated in the MAPK-mediated gene regulation (see Karin, 1995; Hill and Treisman, 1995 for review). For example, the activation of ERKs leads to the phosphorylation of the ternary complex factor (TCF)/Elk-1 (Gille et al., 1992), an important transcription factor involved in the regulation of c-fos gene expression (Treisman et al., 1992). On the other hand, JNKs cause the transcriptional activation of c-Jun (Dérijard et al., 1994), a major component of AP-1 transcriptional complex, which consists of either Jun:Jun homodimers or Jun:Fos heterodimers (see Karin et al., 1997 for review). The ERK and JNK pathways converge on serum responsive element (SRE) to mediate increased gene expression (Cavigelli et al., 1995; Gupta et al., 1995; Whitmarsh et al., 1995). The Ets motif of SRE is recognized by (TCF)/Elk1 (Marais et al., 1993). JNKs activates transcriptional activity of Elk-1 through the phosphorylation of Ser-383 and Ser-389 in the carboxyl-terminal (Cavigelli et al., 1995; Whitmarsh et al., 1995). JNK also phosphorylates the Thr-69 and Thr-71 in the transactivation domain of ATF2, causing enhanced transcriptional activity which is inhibited by the expression of a negative mutant of JNK (Gupta et al., 1995). Chimeric constructs of the luciferase reporter gene driven by TPA-responsive element (TRE) are shear stress inducible, indicating that shear stress activates the AP-1/TREmediated transcriptional activation (Shyy et al., 1995a). By using various negative mutants of MAPKs, it has been shown that JNKs, but not ERKs, are signaling molecules for TRE activation by shear stress (Li et al., 1996). Thus, co-transfection of expression plasmids encoding JNK(K-R), a kinase-deficient JNK1, in which the Lys-52 in the wild-type is replaced by an Arg, attenuates the shear-induced 4xTRE-Pl-Luc and MCP1Luc-540. 4xTRE-Pl-Luc is a chimeric construct in which the luciferase gene is driven by four copies of TRE fused to the rat prolactin minimal promoter, whereas MCP1Luc-540 is luciferase gene ligated to a 540-bp MCP-1 native promoter. In contrast, co-transfection of ERK(K71R) and ERK(K52R), negative mutants of ERK1 and ERK2 in which the respective Lys-71 and Lys-52 in the wild types has been replaced by Arg (Cobb et al., 1991), has little effect on the shear-induction of 4xTRE-Pl-Luc and MCP1-Luc-540.
RELATIONS OF INTEGRINS AND FOCAL ADHESIONS TO THE RASMAPKs PATHWAYS The FAK-Grb2/Sos Pathway is Upstream of the Ras-MAPKs Pathways In the integrin-mediated signaling, the FAK-Grb2/Sos pathway has been shown to modulate the ERKs activation through Ras (Schlaepfer et al., 1994). Dominant negative mutants of FAK and Sos have been used to test the hypothesis that the FAKGrb2/Sos pathway is critical for shear stress activation of the Ras-MAPKs pathways (Li et al., 1997). FAK(F397Y), which encodes a mutated HA-FAK in which Tyr-397 has been replaced by Phe, blocks the binding of both Src family and Grb2 to FAK (Schlaepfer and Hunter, 1997). Empty plasmid pCDNA3, HA-FAK(wild-type), or
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HA-FAK(F397Y) have been co-transfected with either the epitope-tagged MycERK2 or the epitope-tagged HA-JNK1 into BAEC to assess the functional roles of the Tyr-397 of FAK in shear stress activation of ERK and JNK. In cells co-transfected with the pCDNA3 empty vector or HA-FAK(wild-type), shear stress significantly increases the kinase activity of Myc-ERK2 and HA-JNK1. In contrast, FAK(F397Y) blocks the shear stress activation of Myc-ERK2 and reduces the HA-JNK1 activity by 50%. mSosl is a dominant negative mutant of Sos in which the guanine nucleotide exchange domain of the wild-type Sos cDNA has been deleted. Hence, the encoded protein can not activate Ras (Sakaue et al., 1995). mSosl causes a suppression of the shear stress-induced Myc-ERK2 and HA-JNK1 activities in BAECs. Thus, shear stress activation of the FAK-Grb2/Sos pathway in ECs is upstream of the Ras-ERKs and Ras-JNKs pathways.
c-Src is Upstream of the Ras-MAPKs Pathways In addition to FAK, c-Src at focal adhesions can be activated by shear stress (Takahashi and Berk, 1996; Jalali et al., 1998). Co-transfection of c-Src(K295R), a kinase-defective mutant of c-Src, with the epitope tagged Myc-ERK2 into BAECs attenuates the shear stress activation of Myc-ERK2. Co-transfection of c-Src(F527), a constitutively activated form of c-Src, with Myc-ERK2 increases the kinase activity of Myc-ERK2, and this effect is markedly reduced by RasN17. RasL61 (an activated form of Ras) also increases the Myc-ERK2 activity, but this effect is not decreased by the co-transfection of cSrc(K295R). These results indicate that c-Src is upstream of Ras in the shear stress activation of ERKs in ECs. Similar experiments have shown that c-Src is also upstream to Ras in the shear stress activation of JNKs in ECs (Jalali et al., 1998).
av ß3 Integrin is Upstream of the Ras-MAPKs Pathways The involvement of FAK and c-Src, which are PTKs in the focal adhesion sites, in the shear stress activation of ERKs and JNKs in ECs leads to the question whether the integrins linking the focal adhesion to ECM are engaged in the shear stress activation of MAPKs. Vitronectin receptor (i.e., avß3 integrin) is an integrin which is present in ECs. Confluent monolayers of BAECs were pre-incubated 2 hr with LM609 which is an anti-avß3 mAb that has been shown to inhibit endothelial spreading (Sriramarao et al., 1993) and to decrease angiogenesis in tumors (Brooks et al., 1995). Pre-incubating BAECs with LM609 mAb attenuates the shear stress activation of ERK2 and JNK1, as indicated by the decreased phosphorylation of MBP and GST-c-Jun (Li et al., 1997). These results suggest that avß3 integrin is involved in the mechanotransduction that mediates the shear stress activation of ERKs and JNKs. The regulation of MAPK pathways by FAK, c-Src, and avß3 integrin indicates the similarity between the cellular responses to integrin-mediated adhesion and those to shear stress. Thus, integrins are likely to serve as mechanosensors in ECs in response to shear stress (see Shyy and Chien, 1997 for review).
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SOME OTHER PHOSPHORYLATION EVENTS ACTIVATED BY SHEAR STRESS B Activation of NF The activity of the transcription factor NFB is tightly regulated by cytoines and other stimuli such as reactive oxygen species and ultraviolet radiation (see Thanos and Maniatis, 1995; Verma et al., 1995 for review). Several studies have demonstrated that shear stress also regulates the activity of NFB. Nuclear extracts isolated from sheared cells increase their binding activities to oligonucleotides containing the B sequence as demonstrated by electrophoresis moSbility shift assays (EMSA) (Lan et al., 1995). Furthermore, the NFB binding activity is significantly greater in ECs exposed to prolonged low shear than in ECs exposed to high shear (Mohan et al., 1997). The shear stress induction of the PDGF-B gene is due to the increased binding of NFB to the shear stress responsive element (SSRE) with a nucleotide sequence of GAGACC which constitutes a part of B sequence (Resnick et al., 1993; Khachigian et al., 1995). HIV(LTR) is another shear-inducible promoter that contains B sequence (Shyy et al., 1995b). In the quiescent state, NFB is sequestered in the cytoplasm due to binding of the inhibitory protein IBs, which prevent NFB from entering the nuclei. When cells are activated by the various stimuli, IBs are phosphorylated to result in their rapid degradation by proteasomes (DiDonato et al., 1996; Chen et al., 1996). When dissociated from IBs, NFB translocates into the nucleus where it binds to the B sequences in the promoter regions of target genes. A 900-kDa IB kinase complex (IKK) leading to the activation of NFB has been recently identified (DiDonato et al., 1997; Mercurio et al., 1997; Zandi et al., 1997). Serine kinases IKK and IKKß, which are two subunits of IKK, phosphorylate the Ser-32 and Ser-36 at the NH2-terminal of IB which leads to the ubiquitination and degradation of IB. It is possible that the shear stress activation of NFB in ECs is mediated by an induction of IKK.
Phosphorylation of Sp1 The tissue factor (TF) gene is another gene induced by shear stress in ECs. Shear stress has been shown to cause a transient increase of procoagulant activity in ECs, which is accompanied by a rapid and transient induction of the TF mRNA (Lin et al., 1997). Functional analysis of the 2.2-kb TF 5' promoter indicates that a GC-rich region containing three copies each of the EGR-1 and Sp1 sites is required for the induction. Mutation of the Sp1 sites, but not the EGR-1 sites, attenuates the response of the TF promoter to shear stress. EMSA shows no increase in binding of nuclear extracts from sheared cells to an Sp1 consensus site. Thus, other mechanism would be involved in the shear stress induction of Sp1 rather than the binding of the Sp1 transcription factor. An increase in Sp1 Immunoblotting of nuclear proteins prepared from BAECs with polyclonal anti-human Sp1 shows that a shear stress of 12 dynes/cm2 induces a transient increase in Sp1, as indicated by the increase in the phosphorylated/nonphosphorylated Sp1 ratio. This increase in phosphorylated Spl by shear stress is similar to that caused by okadaic acid (OKA) which is an inhibitor of Ser/Thr phosphatase
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(Vlach et al., 1995). As positive controls, BAECs were treated with OKA, and indeed the phosphorylated Sp1 was increased. Compared to the static controls, Sp1 phosphorylation in cells subjected to shear stress for 30min and 1 hr increase by 30±7% and 35±12%, respectively. The phosphorylated Sp1 in cells decreases thereafter to below the static controls (–25% following 12 hr of shearing) (Lin et al., 1997). Tyrosine Phosphorylation of EGFR in A431 Cells The binding of EGF to its cognate receptor (i.e., EGFR) induces the tyrosine phosphorylation of the cytoplasmic domains of EGFR, leading to the recruitment of the SH2-containing adaptor molecules such as Grb2 and She to the phosphorylated tyrosine (Batzer et al., 1994). Shearing A431 cells has similar effects as EGF stimulation in activating EGFR. Immunoprecipitation of EGFR followed by immunoblotting with PY20 reveals that shear stress rapidly induces the tyrosine phosphorylation of EGFR (Kim, Shyy and Chien, unpublished results). The increased phosphorylation can be detected as early as 30 sec after shearing. However, the tyrosine phosphorylation of EGFR is not detectable in ECs, which is likely due to the low level of expression of EGFR in EC lineage (Gospodarowicz et al., 1978). The mechanism by which shear stress activates EGFR is not clear, although this membrane-associated protein has been shown to aggregate in response to UV irradiation or osmotic stress (Rosette and Karin, 1996). Other Kinases Activated by Shear Stress In lymphocytes and fibroblasts, the p70/p85 S6 kinase (pp70S6K) phosphorylates the S6 polypeptide of the 40S ribosomal subunit to regulate the translation of mRNAs with pyrimidine-rich tracts and extensive secondary structures in their 5'-untranslated regions (see Chou and Blenis, 1995 for review). pp70S6K in HUVECs has been shown to be rapidly activated by shear stress. Pretreating cells with the pp70S6K inhibitors repamycin or wortmannin abolishes such an activation (Kraiss et al., 1997). These results suggest that shear stress may regulate cellular translation process by activating the pp70S6K pathway. Endothelial constitutive NO synthase (ecNOS) is a key enzyme in the regulation of the EC release of NO, which plays a critical role in vasodilation. The effects of shear stress on the phosphorylation of ecNOS have been investigated in BAECs metabolically labeled with [32P]orthophosphate. SDS-PAGE followed by autoradiographic analysis of ecNOS immunoprecipitated from cell lysate shows a drastic increase in ecNOS phosphorylation 1 min after a shear stress of 25 dyn/cm2 (Corson et al., 1996). Although ecNOS is not tyrosine-phosphorylated, the early phase of flow-dependent NO production can be blocked by genistein. Phosphorylation is one of several mechanisms that regulate ecNOS enzymatic activity. Whether shear stress activates other posttranslational modification of ecNOS, such as myristoylation and palmitoylation, remains to be determined. Shear stress also induces a rapid (as early as 30 sec) and sustained augmentation of tyrosine phosphorylation of a 128-kDa glycoprotein. Immunoprecipitation of cell
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lysates with a polyclonal anti-platelet endothelial cell adhesion molecule-1 (PECAM1) followed by immuoblotting with anti-PY mAb reveals that this glycoprotein is PECAM-1 (Osawa et al., 1997). c-Src possibly phosphorylates PECAM-1 since c-Src has been shown to phosphorylate and bind to a GST fusion protein containing the PECAM-1 cytoplasmic domain (Osawa et al., 1997). PECAM-1, also known as CD31 or endoCAM, is involved in cell-cell adhesion by a homophilic-binding mechanism and also by heterophilic binding to heparin sulfate proteoglycan or integrins (DeLisser et al., 1993; Piali et al., 1995). In ECs, PECAM-1 is concentrated at the cell junction and has been proposed to be important in maintaining the monolayer structure (see Newman, 1997 for review). The increased tyrosine phosphorylation PECAM-1 by shear stress indicates that PTKs in the cell junction may also be involved in the mechanotransduction.
CONCLUSION From the above discussion, shear stress activates a network of protein phosphorylation to modulate gene expression and other functional consequences in ECs. These events can be summarized as follows: Shear stress rapidly activates PTKs in the focal adhesion sites, i.e., FAK and Src-family PTKs (e.g., c-Src and Fyn). Through an integrin (e.g., vß3)-dependent process, these PTKs associate with the complex of Grb2/Sos to activate Ras through GDP/GTP exchange. Concurrently, Ras can be activated by RTKs signaling from the luminal side of membrane. The activated Ras augments the cytoplasmic MAPKs pathways through cascades of phosphorylation to activate the downstream ERKs, JNKs, and possibly IKKs. The enhanced Ser/Thr kinase activities of ERKs and JNKs result in the increased phosphorylation of transcription factors such as c-Jun, Elk-1, and TCF/c-Fos. On the other hand, phosphorylation of IB leads to its degradation followed by the translocation of NFB/Rel into the nucleus. Binding of these activated transcription factors to their target cis-elements causes the activation of appropriate genes in response to shear stress (see Figure 4.4 for summary). The signaling pathways and cis-elements involved in mechanical activation of genes have considerable similarities with those in chemical stimulation. However, most of the protein phosphorylation signaling events and the resultant gene expression induced by shear stress are transient (Chien et al., 1998). In shear stress-induced signaling, it seems that multiple receptors and pathways can be activated to form a highly coordinated mechanotransduction system. Many receptors have been postulated to be involved in sensing shear stress, e.g. membrane associated receptors, G-proteins, cytoskeleton structure, membrane-associated K+ channels, and integrins (see Davies 1995 for review). Shear stress acting on these putative shear stress receptors would lead to the activation of multiple PTKs and Ser/Thr kinases which initiate many cascades of protein phosphorylation reactions. The activated signal transduction pathways are not simply linear, but rather involve cross-talk, feedback, and bi-directional communication to form a signaling network. For example, integrin clustering causes the recruitment of many molecules in the RTKs signaling pathways such as c-Src, PI 3-kinase, and PLC- to the focal adhesion sites (Plopper et al., 1995).
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Figure 4.4. The phosphorylation-mediated signaling and gene expression in ECs in response to shear stress. RTKs and tyrosine kinases in the focal adhesion site of ECs, such as FAK and cSrc, are involved in the mechano-chemical transduction in a integrin-dependent manner. Through the SH2-containing adaptor Grb2, the small GTPase Ras is activated by Sos, a guanine nucleotide exchange factor which converts the inactive GDP-Ras to the activated GTP-Ras. As a result, ERK and JNK in the cytoplasm are activated to phosphorylate respectively p62TCF/c-Fos and cJun. Concomitantly, IB is phosphorylated by IKK to facilitate its degradation. In the nucleus, the action of the activated AP-1 on its target sequence, e.g., the TRE site in the promoter of the MCP-1 gene, causes an up-regulation of gene expression. Concurrently, the translocated NFB/ Rel binds to SSRE or B site to activate genes such as the PDGF-B gene.
PDGF stimulates tyrosine phosphorylation of FAK and paxillin and the association of FAK and PI 3-kinase (Rankin and Rozengurt, 1994; Chen and Guan, 1994). Activation of G protein-coupled receptors by ligands such as thrombin, bombesin, and lysophosphatidic acid, stimulate FAK phosphorylation (Sinnett-Smith et al., 1993; Seufferlein and Rozengurt, 1994). Furthermore, many of the activated pathways converge at Ras which can serve as a molecular switch to activate the downstream MAPKs. Thus, the activation of MAPKs is regulated as a result of fine tuning among different receptors and pathways. This notion is supported by the findings that the shear activation of ERKs is Gi2 dependent: but the shear activation of JNKs is Gß/ dependent (Jo et al., 1997), and that FAK and c-Src regulate the shear activation of both ERKs and JNKs (Li et al., 1997; Jalali et al., 1998). Each individual molecule in the various phosphorylation pathways has distinct structure-functional requisites, but their temporal responses to shear stress are highly orchestrated. For those molecules located close to the plasma membrane, e.g., FAK, c-Src, and PECAM-1, the activation by shear stress is within a time frame of 1–5min. The activation of cytoplasmic MAPKs requires approximately 30min to reach the peaks, whereas those in the nucleus, e.g., Sp1 phosphorylation, occur even later. Such temporal responses suggest the possibility that the earliest EC response to shear stress, and hence the mechanosensing machinery, is on the cell membrane. A highly coordinated mechanism of dephosphorylation may also be activated by shear stress to result in the transient nature of the protein phosphorylation. This could involve
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many Ser/Thr and tyrosine phosphatases to “switch off” the signaling relay. Indeed, it has been shown that laminar shear stress causes a sustained activation of phosphatases in ECs (Lin and Shyy, unpublished result). An intricate balance between the phosphorylation and dephosphorylation signaling pathways under physiological conditions is necessary to maintain the homeostasis of vascular functions. Pathophysiological hemodynamic conditions such as disturbed flow patterns at the branches of arterial tree may cause an impaired balance between the kinases and phosphatases, and lead to endothelial dysfunction.
ACKNOWLEDGMENT This work was supported in part by research grant HL19454, HL43026 (S.C.), and HL56707 (J.Y-J.Shyy) from the National Heart, Lung and Blood Institute of the National Institutes of Health.
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5 In Vitro Simulation of Shear Stress and Mitogen-activated Protein Kinase Responses to Shear Stress in Endothelial Cells Oren Traub, Chen Yan and Bradford C.Berk* Department of Pathology and Department of Medicine, University of Washington, Seattle, WA 98195–7710, USA and Center for Cardiovascular Research, University of Rochester, Rochester, NY 14642, USA. *Corresponding author: Bradford C.Berk, Center for Cardiovascular Research, Box 679, University of Rochester, Rochester, New York 14642, USA. E-mail: [email protected]
Mechanical forces are important modulators of cellular function in many tissues and are particularly important in the cardiovascular system where they may play a role in the pathogenesis of atherosclerosis and hypertension. As a result of its unique location, the endothelial cell has evolved important physiologic responses elicited by the mechanical force associated with fluid shear stress. While the effects of shear stress on endothelial cell function have been well studied, the mechanisms by which endothelial cells sense mechanical stimuli and convert them to biochemical signals are not well characterized. The class of mitogenactivated protein kinases (MAPKs) are excellent candidates to mediate mechanotransduction in endothelial cells. MAPKs respond to diverse stimuli, including physical stress, oxidative stress, and UV light, and have broad effects on cell physiology and gene expression. In this chapter, we detail the construction of two different apparatus and characterize their efficacy in simulating shear stress on cultured cells in vitro. We also discuss the responses and likely signal transduction mechanisms leading to shear stress-mediated activation of different members of the MAPK family. Finally, we demonstrate that the MAPK family member, ERK1/2 is an excellent biological marker for shear stress responsiveness and may aid in the study of mechanotransduction in endothelial cells. KEYWORDS: Shear stress, endothelium, mitogen-activated protein kinase, ERK1/2, parallel plate chamber, cone and plate viscometer.
INTRODUCTION Fluid shear stress is one of the most important hemodynamic forces recognized and transduced by endothelial cells, as it modulates vessel structure and function. Shear stress is also important in the pathogenesis of atherosclerosis because atherosclerotic plaques occur preferentially in areas which experience low shear stress and flow reversal. Changes in shear stress cause rapid secretion of vasoactive mediators, including nitric oxide, prostacyclin, and endothelin. Changes in shear stress also cause long term alterations in vessel structure and3 function by regulating gene and protein 89
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expression. For example, shear stress stimulates expression of platelet-derived growth factor (PDGF) A- and B-chains, tissue plasminogen activator, endothelial nitric oxide synthase and superoxide dismutase. An important question concerns the mechanisms by which shear stress transduces signals that modify endothelial cell function. Experimental evidence indicates that the cellular response to shear stress is similar to the response to classical growth factors (e.g. epidermal growth factor) which involves activation of a complex array of phosphorylation cascades. This concept has been strongly supported by study of the mitogen-activated protein kinase (MAPK) family which plays an integral role in growth factor-mediated signaling and in the endothelial cell response to fluid shear stress. In this chapter, we review mechanisms by which MAPKs are regulated by shear stress and discuss techniques whereby responsiveness to shear stress are easily studied.
IN VITRO APPLICATION OF SHEAR STRESS Since the signal transduction mechanisms by which the physical force of shear stress is transduced into biochemical signals have not been fully elucidated, the use of a system by which shear stress can be simulated on cells grown in culture is useful to characterize the biological consequence of alterations in blood flow. In this manner the parameters that define shear stress can be precisely controlled, while simultaneously eliminating other variables that are present in the in vivo or ex vivo system that respond to and influence changes in shear stress (e.g., reflexive smooth muscle contraction, flow-mediated vasodilation). Through the use of principles of physics, mathematics and biology, several different types of apparatus can be constructed to simulate shear stress over cultured cells and to evaluate the response to shear stress.
Extracellular Signal-Regulated Kinases as a Biological Marker for Shear Stress Responsiveness One consideration in determining the effect of shear stress on a biological system is the selection of an appropriate biological endpoint or marker that varies in response to shear stress. The family of MAPKs are excellent candidates to serve as a biological endpoint as they have been shown to be activated by various stimuli (Pelech and Sanghera, 1992), including physical stress (Yamazaki et al., 1995, Tseng et al., 1995, Brewster et al., 1993), and they have diverse effects on gene expression and cell physiology (Berk et al., 1995). As we will discuss below, the responses of the MAPK members, c-Jun NH2 terminal kinase (JNK), p38 kinase, and big MAP kinase (BMK-1), to shear stress are still being characterized, but activation of ERK1/2 in response to shear stress has been well documented and reproduced by several investigators (Tseng et al., 1995, Li et al., 1996, Traub et al., 1997, Pearce et al., 1996, Jo et al., 1997). Further, this ERK1/2 activation in response to shear stress is present in widely differing cell types from different developmental origins; in addition to the endothelial cell ERK1/2 response to shear stress, ERK1/2 activation in response to
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shear stress can also be observed in rat embryonic fibroblasts and Chinese hamster ovary cell lines (Figure 5.1), though the response in these cell lines is not as robust when compared to endothelial cells. There are several advantages to using ERK1/2 as a biological endpoint for studying shear stress signal transduction. Signaling pathways by which ERK1/2 is activated in response to classical growth factors, such as EGF, have been well elucidated (Pelech and Sanghera, 1992) (Figure 5.2). Thus, EGF is a useful positive control for stimulation of ERK1/2 activity through established signal transduction pathways. Another advantage is that determination of ERK1/2 activity is relatively simple, rapid, and inexpensive. We have previously reported that three different methods of determining ERK1/2 activity—the in vitro immune complex assay, the in-gel kinase assay, and the phosphorylation band shift assay—yield results that correlate very closely (Tseng et al., 1995). Within the past few years, an antibody that preferentially detects the dually phosphorylated form of ERK1/2 has been developed, making quantification of ERK1/ 2 phosphorylation even easier (ERK1/2 phosphorylation correlates well with ERK1/ 2 activity as shown in Figure 5.6). A final advantage is the ERK1/2 response to shear stress can be seen as early as 3 min and peaks at 10 min (Tseng et al., 1995); thus, relatively short stimulation by shear stress is sufficient to study signaling events which control the response to shear stress. In the discussions below we will present information regarding calculation of fluid shear stress in two different devices used to create fluid shear stress in vitro: the parallel plate chamber and the cone and plate viscometer. Next we will validate the mathematical derivation of shear stress in these devices by experimental determination of ERK1/2 activation. Differences in the two devices will be summarized. Finally studies of other MAPK kinase members will be presented.
Figure 5.1. Shear stress-mediated ERK1/2 in different cell lines. Different cell types grown in culture were exposed to shear stress (12 dynes/cm2) for 10 min. Lysates were separated by SDSPAGE and Western blotting with anti-phosphospecific-ERK1/2 antibody performed. Shear stress increased ERK1/2 phosphorylation in each cell type tested. Rat-1 and CHO cells were serum deprived for 1 day prior to shear stress stimulus in order to reduce baseline ERK1/2 activity.
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Figure 5.2. EGF-mediated ERK1/2 signaling pathway. The MAP and ERK kinase (MEK-1) is a dual specificity kinase that phosphorylates ERK 1/2 on T-E-Y. MEK-1 is itself regulated by a MAP kinase kinase kinase, one of which has been identified as Raf-1. Raf-1 is activated by translocation to the membrane and association with the small GTP-binding protein, ras. The GTPase activity of ras is regulated by a complex involving Grb2 and mSOS which are recruited and activated by a tyrosine kinase receptor.
Basic Physical Principles of Shear Stress As a physical principle, shear stress occurs when a tangential force is applied to one face of a body while the opposite face is held stationary by another force, such as friction or adhesion. If the object undergoing this force is a rectangular block, a shear stress results in a shape whose cross-section is a parallelogram. The shear stress in this simplistic example can be defined by the formula: (1)
where is the shear stress equal to the force, F, divided by the area of the face being sheared, A. When speaking of a more complex example of shear stress exerted by a liquid passing over an object, additional factors must be considered when calculating the magnitude of the shear stress, in particular the fluid dynamic state (Roark, 1965, Goldstein, 1996, Serway, 1996). When adjacent layers of a viscous fluid flow smoothly over each other in an ordered fashion, the stable streamline flow is called laminar flow, and the shear stress exerted by the fluid will be unidirectional and summative. However, at sufficiently high velocities or constrained geometry, the fluid flow will transition from laminar flow to
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a highly irregular and random motion of the fluid called turbulent flow. In this fluid dynamic state, the shear stress vectors are often opposing and the mean shear stress exerted over a body face can approach zero (in contrast to the high local shear gradients). The fluid velocity at which turbulence occurs depends on the geometry of the medium surrounding the fluid and the fluid viscosity. Experimentally, it is found that the onset of turbulence is determined by a dimensionless parameter called the Reynolds number (RN) given by: (2)
where is the fluid density, v is the velocity of the fluid, is the viscosity of the fluid, and d is the geometrical length associated with the fluid flow (for flow through a tube, d is equal to the diameter of the tube). Experimental observations have determined that the fluid dynamic state is laminar if the Reynolds number is below 2000 and is turbulent if the Reynolds number is above 3000 (Roark, 1965, Goldstein, 1996, Serway, 1996). Using these principles as a basis, two different types of apparatus have been constructed to simulate shear stress over cultured cells in vitro: the parallel plate chamber, and the cone and plate viscometer. Parallel Plate Chamber The parallel plate chamber (Figure 5.3a) is an apparatus consisting of a gravity-fed recirculating flow loop system. Gravity-fed medium between two reservoirs is placed in series with a parallel plate flow chamber and a flow meter (Figure 5.3b). During experiments, the upper reservoir and medium is warmed to 37°C and flow rate is measured through the use of a flow meter. A recirculating pump is connected to pump medium back to the upper reservoir. Fluid medium is a buffered physiological salt solution (containing in mM, NaCl 130, KC1 5, CaCl2 1.5, MgCl2 1.0, HEPES 20, pH 7.4). The cell chamber consists of a monolayer of cells grown on tissue culture plastic that is cut to precise dimensions (74 by 36 mm) from a tissue culture dish and a Plexiglas block sandwiching a mylar gasket of known thickness. These items define a cell chamber of known dimensions. Through a complex series of mathematical derivations discussed elsewhere (Rosenhead, 1963) and the use of readings from the flow meter, shear stress applied over cells in this apparatus can be directly determined by the formula: (3)
where =wall shear stress in dynes/cm2, µ=viscosity of the flow medium (poise), Q=flow rate (mL/sec), b=width of the cell chamber (cm), and h=height of the cell chamber (cm); in the present example, µ=0.006915; b=3.7 and h=0.025. In this case, the ratio of the flow channel width to height is large enough (>100) so that the Reynolds number is small and turbulent flow is minimized. This apparatus has a distinct advantage in applying defined magnitudes of shear stress over cultured cells, particularly because it provides a precise readout of flow rate through the system, thereby allowing the direct calculation of shear stress. However, this apparatus also has its limitations. Because of the relatively high volume
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Figure 5.3. (A) Parallel plate chamber for simulating shear stress. The cell chamber consists of a monolayer of cells grown on tissue culture plastic that is cut to precise dimensions (74 by 36 mm) from a tissue culture dish and a Plexiglas block sandwiching a mylar gasket of known thickness. Shear stress is calculated according to the following formula: t=(6µQ)/(h2b) where t=wall shear stress (dynes/cm2); µ=viscosity of the medium (poise), Q=flow rate (mL/sec); b=width of the cell chamber; h=height of the cell chamber. For the chamber and medium used in these experiments: µ=0.006915; b=3.7 cm; and h=0.025 cm. (B) Flow buffer is transferred to an upper reservoir by a roller pump. The height of this upper chamber determines the rate of flow of the buffer through the parallel plate chamber. The rate of flow is measured by a flow meter distal to the parallel plate chamber. The buffer returns to the lower reservoir and is recirculated. A heating pad regulates temperature of the buffer to 37°C. Fluorimetric data can be obtained during the experiment by optical capture above the parallel plate chamber. Approximate volume of the buffer utilized is 150 mL.
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of flow buffer needed to keep flow constant and free of air bubbles (minimum= 150 mL), large amounts of a pharmacological agent are needed in order to achieve sufficient concentration for biological effect. This large volume also makes measuring factors released from endothelial cells in response to shear stress, such as nitric oxide, difficult. Finally, because of its large size, this apparatus is relatively stationary. These constraints can be avoided through construction of an alternative apparatus, the cone and plate viscometer. Cone and Plate Viscometer The cone and plate viscometer consists of a cone that is inserted into a circular tissue culture dish containing cultured cells and rotated so that the medium within the dish circulates in a laminar fashion and at controlled velocities (Figure 5.4). This apparatus has the advantage of using lower volumes of fluid media (1–4 mL), thus allowing higher concentrations of drugs with minimal expense and facilitating measurement of factors released from endothelial cells in response to shear stress. Additionally, its size lends itself to portability. In deriving the mathematical formulae for applied shear stress using this device, it is useful to first examine the effects of a disk and plate apparatus (i.e. a flat surface inserted into the tissue culture dish) (Figure 5.5a). When the disk is rotated, a concentric fluid movement occurs in the culture medium in the dish. The endothelial cell monolayer cultured on the bottom of the dish is subjected to the
Figure 5.4. Cone and Plate Viscometer Apparatus. Polyacetal resin (Delrin) cones in two different sizes (for 60 mm and 100 mm Corning tissue culture dishes) were milled with precise angle measurements and attached to a stainless steel shaft which was in turn attached to a BC215GDAF model motor and 2GD10K gear head purchased from Oriental Motor Co. Torrence, CA. The motor was wired to a external controller (Model BLD15-AF) with adjustable potentiometer. A step down transformer (120 V to 100 V; Sanyo Model TSD-N0-GU) was added and the motor and gear head was placed on an adjustable platform so that the cone could be lowered onto the base containing the cell culture dish. Adjustable set screws were used to obtain reproducible heights from the cell culture dish. The apparatus (not including external controller) was placed in an adjustable incubator set at 37°C for experiments.
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Figure 5.5. (A) Geometry of the disk and plate apparatus defining radial [r], angular [q], and azimuthal coordinates [z]; (B) geometry of the cone and plate apparatus.
shear stress according to the velocity gradient near the cell surface. As the medium can be regarded as a Newtonian fluid, the induced flow can be expressed by the Navier-Stokes equations in cylindrical coordinates with coordinate directions, (r, , z) (See Figure 5.5a) as previously illustrated by Nomura et al. (1988). According to Lance et al. (1961), when the Reynolds number determined for this geometry is very low, the effects of the concentric force in the rotating fluid is negligible and therefore velocity for the components r and z can be regarded as negligible. The Navier-Stokes equation can be reduced to a partial differential equation, and when the angular velocity () of the disk rotation is constant, the wall shear stress exerted in the bottom of the dish is expressed as (4)
where µ is equal to the viscosity, is the angular velocity of the disk rotation, and d is the distance between the disk surface and the dish bottom. In this case, the sole variable is the radius; thus shear stress increases directly proportional to the increasing radial distance from the center of the disk. This variability in generated shear stress within a single dish is not desirable for our purposes as we wish to have a uniform shear stress applied to the cells. In order to rectify this, the rotating disk can be replaced with a rotating cone (Figure 5.5b) with a small, but constant angle (). Thus we now have two variables, the distance of the cone surface from the dish at each radius as well as the radial distance. However, by
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basic trigonometric principles, it is known that the ratio of these variables, given a constant angle on the cone, is constant, and thus: (5)
In other words, d always changes in proportion to r. Equation (4) can thus be altered so that shear stress on the bottom of the dish at each radius can now be described as: (6)
and shear stress is now theoretically uniform within the cell culture dish at each radius. However, with the change in geometry that occurs when replacing the disk with a cone, the Reynolds number which dictates fluid dynamic considerations also changes. The fluid dynamic patterns induced by a rotating cone have been previously characterized (Bussolari et al., 1982) and the modified Reynolds number defined by: (7)
where laminar flow is obtained when RNcone<1. Bussolari et al. (1982) goes on to describe local shear stress on the plate surface as: (8)
Through the use of the 1° angle cone, it is possible to minimize fluid turbulence at the levels of shear stress required for our experiments (0–12 dynes/cm2). Experimental Results Fluid shear stress leads to phosphorylation and activation of ERK1/2 in a force-dependent manner In order to demonstrate that ERK1/2 activation occurs in response to shear stress in a force-dependent manner, human umbilical vein endothelial cells (HUVEC) were exposed to shear stress of varying magnitudes for 10 min. Both activity and phosphorylation of ERK1/2 increased in a force-dependent manner (Figure 5.6a). An in-gel kinase assay was also performed on these samples to demonstrate that ERK1/2 phosphorylation correlates with ERK1/2 kinase activity (Figure 5.6b), indicating that this relatively rapid and inexpensive method of determining ERK1/2 phosphorylation correlates with ERK1/2 activity as well. When ERK1/2 phosphorylation force-response curves were compared for the parallel plate chamber and cone and plate viscometer, there was no significant difference between the results (Figure 5.6c). These findings indicate that the assumptions made for calculation of shear stress in the cone and plate viscometer are valid based on direct experimental comparison of results obtained with measured amounts of shear stress generated with the parallel plate chamber.
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Figure 5.6. (A) ERK1/2 phosphorylation and activation is increased by shear stress. HUVEC underwent shear stress of increasing magnitudes for 10 minutes. For measurement of ERK1/2 phosphorylation, lysates were separated by SDS-PAGE and Western blotting with antiphosphospecific-ERK1/2 antibody performed. For measurement of ERK1/2 activity, lysates were run on SDS-PAGE containing myelin basic protein. Protein was renatured and incubated with -32P-ATP for 1 hour. The gel was dried and autoradiography was performed to measure kinase activity. Shear stress increased ERK1/2 activity and phosphorylation in a force-dependent manner; (B) ERK1/2 phosphorylation correlated to ERK1/2 activity with r=0.95; (C) ERK1/2 phosphorylation force-response curves were not significantly different between the cone and plate viscometer and the parallel plate chamber when normalized to basal levels of ERK1/2 phosphorylation.
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Shear stress stimulates ERK1/2 in a time-dependent manner A time course of shear stress performed at 12 dynes/cm2 demonstrates that shear stress stimulates ERK1/2 in a time-dependent manner. Compared with static conditions, fluid shear stress at 12 dynes/cm2 in the parallel plate chamber activated ERK1/2 with a peak at 10 min and return to baseline by 60 min (Figure 5.7). Western blotting with an antibody for ERK1/2 that detects both the phosphorylated and unphosphorylated forms of the kinases showed that cellular ERK1/2 levels remained constant throughout the shear stress time-course. The time course for ERK1/2 phosphorylation using the cone and plate viscometer (Figure 5.7) was similar to results obtained with the parallel plate chamber for early time points (<30 min). However, levels of pERKl/2 were significantly higher at time points greater than 30 min when compared with results using the parallel plate chamber. One likely explanation for this persistent elevation is that vasoactive mediators released by the endothelial cell in response to shear stress become concentrated in the relatively low volume (1–4 mL) of media used in the cone and plate viscometer. Many vasoactive mediators are released by endothelial cells in response to shear stress (Davies, 1995) and could stimulate ERK1/2. These data indicate that the use of the cone and plate viscometer yields results similar to those obtained with the parallel plate chamber for early time points, but not for longer time points. Increased distance of the cone from the dish lowers ERK1/2 activation Theoretical considerations dictate that as the cone is moved away from the dish, the Reynolds number increases and turbulent flow will occur, resulting in a mean directional shear stress approaching zero. In order to determine whether distance of the cone from the dish (cone height) would affect shear stress generated, experiments were performed whereby the rotational velocity of the cone was held constant, but the cone was progressively moved away from the dish and ERK1/2 phosphorylation
Figure 5.7. Phosphorylation of ERK1/2 by fluid shear stress assayed by Western blotting: time dependence. Endothelial cells were washed free of culture medium with HBSS and maintained in static condition or exposed to 12 dynes/cm2 shear stress for varying times. Samples were harvested, protein lysates separated by SDS-PAGE and transferred to nitrocellulose for Western blotting. Time-course for ERK1/2 tyrosine phosphorylation by shear stress.
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subsequently measured. ERK1/2 phosphorylation was progressively lower as the cone was moved from 0 µm to 500 µm and then to 1000 µm height (Figure 5.8). At a height of 1000 µM, little difference in ERK1/2 phosphorylation was seen at any shear stress applied. These results indicate that reproducible cone height is critical to achieve similar levels of shear stress between different samples and demonstrate a critical variable in the cone and plate viscometer that is not present with the parallel plate chamber. Shear stress-mediated ERK1/2 phosphorylation is largely uniform at different radii in the cone and plate viscometer Since the Reynolds number is essentially an experimentally determined parameter (Serway, 1996, Bussolari et al., 1982), it is necessary to confirm that the response to shear is uniform at each radius and that turbulent flow does not result when testing a new apparatus. In order to determine whether the shear stress distribution is uniform at each radius, experiments were performed where cells were scraped from the dish just prior to experiments to yield defined areas (Figure 5.9a). The shear stress was then applied, and the cells were harvested for determination of ERK1/2 phosphorylation. Experiments determined that the ERK1/2 phosphorylation was not significantly different among the different areas tested except for the inner most area (Figure 5.9a). It is likely that the increased ERK1/ 2 phosphorylation in the inner most area results from actual physical deformation of the cell by the cone due to the small distances between the cone and the bottom of the tissue culture dish at the center of the cone. Further, it should be noted that this bias is relatively equal to that reported in the parallel plate chamber due to an “edge turbulence effect” (unpublished observations). These results confirm mathematical calculations indicating that shear stress is generally uniform, except for at the very center of the dish, for a 1° angle cone and a shear stress magnitude of 12 dynes/cm2.
Figure 5.8. Increasing cone distance from dish reduces ERK1/2 phosphorylation. Cone height was increased in sequential experiments and ERK1/2 phosphorylation measured by Western blotting. ERK1/2 phosphorylation was significantly decreased with increasing cone distance consistent with generation of turbulent flow.
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Figure 5.9. ERK1/2 phosphorylation is largely uniform at different radii, (a) Cells were scraped from the dish just prior to experiments to yield defined areas (b) After shear stress stimulus, cells were harvested and ERK1/2 phosphorylation determined by Western blotting. ERK1/2 phosphorylation was not significantly different among the different areas tested except for the inner most area.
Shear stress-mediated nitric oxide release from cultured cells Shear stress has been demonstrated to be a potent stimulus for nitric oxide release from endothelial cells (Rubanyi et al., 1986). Since nitric oxide has a relatively short half life, nitric oxide release is commonly measured by detection of nitric oxide metabolites. Previously, we have found it difficult to detect nitric oxide metabolites the parallel plate chamber apparatus, presumably due to the relative large volumes needed for this apparatus (>150 mL). However, medium taken from the cone and plate apparatus after shear stress stimulus showed high levels of nitric oxide metabolites, much greater than those obtained with the calcium ionophore, A23187 (Figure 5.10). These results suggest that the cone and plate viscometer is useful for detection of substances released from endothelial cells in response to shear stress, in contrast to the parallel plate chamber. Relative Utility of Parallel Plate Chamber Vs. Cone and Plate Viscometer Comparison of results obtained with the parallel plate chamber to the cone and plate viscometer (Table 5.1) show the two apparatus to be similar in terms of force-response and early time course (>30 min). There was a difference in results obtained between the two apparatus at later time points (>30 min). While the ERK1/2 phosphorylation
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Figure 5.10. Measurement of shear stress-mediated nitric oxide release. Medium taken from the cone and plate apparatus after shear stress stimulus showed high levels of nitric oxide metabolites that were significantly greater than those levels obtained with the calcium ionophore, A23187. Table 5.1 Comparison of the parallel plate chamber vs. the cone and plate viscometer in simulating shear stress
levels returned to baseline at approximately 30 minutes, the ERK1/ 2 phosphorylation levels obtained with the cone and plate viscometer remained elevated even at 60 min. The likely explanation for this disparity is that vasomediators released by endothelial cells in response to shear stress are more concentrated in the lower volume of the cone and plate viscometer (1–4 mL) compared to the parallel plate chamber (>150 mL). Thus, for longer time points, a medium replacement protocol (i.e. replace 1 mL of media with fresh media every 10 minutes) as reported by other investigators (Bussolari et al., 1982; Topper et al., 1997) may be helpful in determining whether released vasomediators may influence the results. It is this same property of the cone and plate viscometer which makes this apparatus very useful in measuring amounts of released vasomediators such as nitric oxide. We
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have previously been unable to detect significant levels of nitric oxide when using the parallel plate chamber. Through the use of the cone and plate viscometer, we have been able to detect the presence of nitric oxide metabolites upon exposure of the endothelial cells to shear stress that is much greater than those levels obtained with the calcium ionophore, A23187. This is consistent with reports that have shown that shear stress is a much more potent stimulus for nitric oxide release than A23187 (Berk et al., 1995). Variability in ERK1/2 phosphorylation occurs with the cone and plate viscometer in part from cells at the very center of the dish. ERK1/2 phosphorylation levels were significantly greater at the center of the dish compared to the remaining portions of the dish. The likely explanation for this is that the cells at the center are physically deformed by the cone itself. To diminish variability, cells from the center circle may be removed via scraping prior to shear stress stimulus. While the cone and plate viscometer does differ from the parallel plate chamber in some aspects, it also provides some advantages. These include reduced time and expense for each sample, less drug needed to achieve higher concentration, more protein harvested per sample, and less error due to leakage. Conversely, the cone and plate apparatus still provides greater difficulties in terms of fluid hemodynamics (though turbulent flow seems to be at a minimum at the levels of shear stress tested) and due to the fact that flow rate cannot be measured directly. Another major finding of these experiments is that the ERK1/2 phosphorylation and activity are excellent biological markers for shear stress responsiveness. ERK1/2 phosphorylation and activity varied with shear stress in a force- and time-dependent fashion and measurements of changes in ERK1/2 activation are relatively quick, simple and inexpensive. It should be stated, however, that the use of ERK1/2 activation as an endpoint is merely one of myriad possible biological markers for shear stress. Many different signal transduction elements are activated and vasomediators released in response to shear stress (Davies, 1995), each with signal transduction pathways that are likely unique from those that activate ERK1/2. In summary, ERK1/2 is an excellent biological marker for shear stress responsivity based on the following properties: (1) ERK1/2 responds to shear stress in a force- and time-dependent manner; (2) EGF-mediated ERK1/2 signal transduction pathways are already elucidated and serve as an excellent positive control; (3) shear stress-mediated ERK1/2 activation is present in several cell lines from different developmental origins; (4) measurement of ERK1/2 phosphorylation and/or activity is quick, simple, and relatively inexpensive and does not require radioactivity; (5) the ERK1/2 response to shear stress is rapid, so short periods of stimulation are sufficient for studies characterizing shear stress responsivity.
ACTIVATION OF MAPKs BY SHEAR STRESS MAPKs are serine/threonine protein kinases. Four subfamilies of MAPKs have been identified (Figure 5.11), including the ERK1/2, JNK, p38 kinase, and BMK1 (or ERK5) (Abe and Berk, 1998). ERK1/2 is activated by an upstream kinase (MEK1) via dual phosphorylation of a TEY motif. However, JNK and p38 kinase are activated by
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Figure 5.11. Putative signal transduction pathways for MAPKs. Depicted are schematic linear patterns showing protein kinases regulating each other in a cascade. It should be noticed that the signal transduction events in response to a certain stimulus may be varied in different cell types.
MEK4 and MEK3 (SEK1) via phosphorylation of TPY and TGY sequences respectively. BMK1 is a newly identified MAPK family member, which shares a TE Y activation sequence with ERK1/2 and is activated by MEK5 (Lee et al., 1995; Zhou et al., 1995). BMK1 has a long COOH-terminal domain which is absent in other MAPK family members, suggesting that its regulation and function may be unique.
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MAPKs play central roles in a variety of cell functions. The specificity of activation of MAPKs is reflected by specific stimuli and substrates for each MAPK member. For example, growth factors activate ERK1/2 strongly, but JNK and p38 weakly (Cano et al., 1994); hyperosmolar stress and inflammatory agents are strong stimuli for p38 and JNK (Han et al., 1994); and oxidative stress is a better stimulus for BMK1 than growth factors (Abe et al., 1996). Among the MAPK family members, ERK1/2, JNK, and BMK1 have been shown to respond to shear stress in endothelial cells. To our knowledge, no studies have been published regarding changes in p38 activity in response to shear stress. Transcriptional factors are important substrates for MAPKs. ERK1/2 can phosphorylate the ternary complex factor (TCF/Elk-1) (Marais et al., 1993; Gille et al., 1992) which is essential for transactivation (Treisman, 1994), while JNK phosphorylates c-Jun, increasing its transcriptional activating potential (Kyriakis et al., 1994). (D’Erijard et al., 1994). Further, ATF can be activated by both JNK (Gupta et al., 1995) and p38 (Raingeaud et al., 1995), and MEF2C is the first protein substrate recently identified for BMK1 (Kato et al., 1997).
Extracellular Signal-Regulated Protein Kinases (ERK1/2) ERK1/2 activation in cultured bovine aortic and human umbilical vein endothelial cells (BAEC and HUVEC) (Tseng et al., 1995) (Ishida et al., 1996) (Pearce et al., 1996) occurs within the physiological range at 1–2 dynes/cm2 and peak between 15– 30 dynes/cm2. The signal transduction events by which ERK1/2 is activated by flow shear stress have been characterized although the exact sequence of events has not been elucidated. The necessity for activation of a heterotrimeric G protein was first demonstrated in BAEC by Tseng et al. (1995) using the nonhydrolyzable GTP analog GDP-ß-S. Jo et al. (1997) further found that expression of mutant Gi2 and antisense i2 prevented shear stress-dependent activation of ERK1/2, suggesting that Gi2 is the G protein isoform mediating ERK1/2 activation by shear stress. The role of protein kinase C (PKC) in shear stress-induced ERK1/2 activation was also investigated (Tseng et al., 1995; Traub et al., 1997). Inhibiting PKC with staurosporine or down regulating PKC with phorbol 12, 13-dibutyrate (PDBu) completely blocked ERK1/2 activation by shear stress, while chelating Ca2+ with BAPTA-AM had no effect suggesting that a Ca2+-independent PKC isoform is required (Tseng et al., 1995; Traub et al., 1997). Using specific antisense PKC oligonucleotides, Traub et al. (1997) showed that PKC , but nor PKC- or PKC-, was specifically required for activation of ERK1/2 by shear stress in HUVEC. ERK1/2 activation by shear stress was found to be dependent on Ras by two groups (Jo et al., 1997) (Li et al., 1996). Shear stress activated Ras (Li et al., 1996) and the activation of ERK1/2 by shear stress was blocked in the cells over-expressing a dominant negative Ras (N17Ras) (Jo et al., 1997). In addition, an important role for tyrosine kinase(s) in shear stress-mediated ERK1/2 activation is suggested by the ability of herbimycin A (Takahashi and Berk, 1996) or genistein (Jo et al., 1997) to inhibit ERK1/2 activation completely. Src family kinase(s) may play an important role in the endothelial response to shear stress because many ERK1/2 activators, such as growth factor PDGF, UV, and shear stress, also activate Src
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(Takahashi and Berk, 1996, Courtneidge et al., 1991, Devary et al., 1992). In summary, Gi2, Ras, tyrosine kinase, PKC- are upstream regulators of shear stress-dependent activation of ERK1/2 (see Table 5.2). Activation of ERK1/2 by growth factors has been shown to be due to initial stimulation of Ras and subsequent activation of Raf-1. Raf-1, in turn, activates MEK1, the direct upstream activator of ERK1/2 (Pelech and Sanghera, 1992). It is very likely that a similar signaling pathway is involved in the ERK1/2 activation by shear stress. Although the mechanism by which PKC- activates ERK1/2 by shear stress is not clear, it seems likely that PKC- interacts directly with Raf-1 to activate ERK1/2 based on similar findings in NIH 3T3 and COS cells (Cai et al., 1997) and inhibitions of PKC- mediated ERK1/2 activation by the expression of mutant Raf-1 (Schaap et al., 1993).
c-Jun NH2 Terminal Kinase (JNK) The effects of shear stress on JNK activity are conflicting. Initial reports from two laboratories showed that JNK was activated in response to shear stress in BAEC even though the time for peak JNK activation by shear stress varied widely in these reports (Li et al., 1996, Jo et al., 1997). Shear stress activated JNK via a signaling pathway apparently different from ERK1/2. Expression of ßARK-ct to block Gß/ partially inhibited shear stress-dependent activation of JNK, suggesting that the ß and subunits of heterotrimeric G-proteins act as one of its upstream regulators. Expression of a dominant negative Ras completely prevented shear stress-dependent activation of JNK, showing its Ras dependence. In addition, because genistein prevented shear stress-dependent activation of JNK, a tyrosine kinase is likely involved. The identity of the tyrosine kinase(s) which mediates JNK activation by shear stress remains to be addressed. Pyk2 has been suggested to provide links between G-protein and JNK in response to stress signals such as UV light and osmotic shock (Tokiwa et al., 1996). Taken together, the shear stress-dependent activation of JNK appears to be regulated by mechanisms involving Gß/, Ras and tyrosine kinase(s).
Table 5.2 Regulation of various MAPK activities by shear stress and signaling molecules in endothelial cells
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In contrast, data from our lab showed that laminar flow (12 dynes/cm2, 10 min) did not activate JNK in HUVEC (Surapisitchat and Berk, 1998). Of potential physiological importance, shear stress prevented JNK activation mediated by a physiologic stimulus, TNF- (Surapisitchat and Berk, 1998). Possible explanations for the discrepancies in these studies include the cell types (BAEC vs. HUVEC) and culture condition (JNK is activated basally in serum-free medium). Big Mitogen-Activated Protein Kinase 1 (BMK1) Recent studies from our laboratory showed that fluid shear stress is among the strongest stimuli for BMK1 activation in both BAEC and HUVEC. However, BMK1 and its upstream kinase, MEK5, are not well activated by most growth factors and cytokines which activate ERK1/2 or JNK.(Abe et al., 1996) (English et al., 1995). Shear stress (12 dynes/cm2) activated BMK1 within 10 min with peak at about 60 min in a force-dependent manner (Yan et al., unpublished observation). The signal events which link shear stress to BMK1 are different from ERK1/2 (Yan et al., unpublished observation). For example, shear stress-stimulated BMK1 activation is dependent on Ca2+, and is particularly dependent on the Ca2+ release from internal stores, but not Ca2+ influx. BMK1 activation by shear stress was not blocked by PKC down-regulation with PDBu, suggesting that BMK1 activation is not dependent on any known PKC because none of the previously identified PKC are unresponsive to phorbol esters and still dependent on Ca2+. The requirement of tyrosine kinase(s) in BMK1 activation was also demonstrated by complete inhibition by herbimycin A. Src has been shown to be essential for BMK1 activation by H2O2 in fibroblasts because H2O2-induced BMK1 activation was blocked in fibroblasts derived from Src-knock out mice (Abe et al., 1997) However, Src is not required for BMK1 activation by shear stress because BMK1 activation was not blocked by over-expression of dominant negative Src using an adenovirus vector (Yan et al., unpublished observation). In summary, while some of the MAPKs responses to shear stress, the upstream molecules and exact sequences of the events involved in the mechano-transduction pathway for activation of MAPKs have been determined, they still remain to be fully defined. Furthermore, the signaling events by which MAPKs regulate gene expression should prove an exciting area for future research.
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6 Flow-induced Endothelial Gene Regulation Joji Ando, Risa Korenaga and Akira Kamiya Department of Biomedical Engineering, Graduate School of Medicine, University of Tokyo, Hongo 7–3–1, Bunkyo-ku, Tokyo 113, Japan Tel: +81–3–3912–2111 (ext. 3659), Fax: +81–3–5800–6928, E-mail: [email protected].
Shear stress generated by blood flow can modulate both the morphology and functions of vascular endothelial cells. In most cases, gene expression associated with endothelial function is also altered by shear stress. To date, the expression of nearly twenty endothelial genes has been shown to be up-and/or down-regulated by shear stress, and several related transcription factors and cis-acting shear-stress-response elements have been identified. We recently characterized a negative shear-stress-response element in the murine vascular adhesion molecule1 (VCAM-1) gene. Exposure of mouse venule endothelial cells to shear stress decreased VCAM1 protein cell surface expression and inhibited adhesion to lymphocytes. The decrease in protein expression was due to a decrease in VCAM-1 mRNA levels, which resulted from the suppression of VCAM-1 gene transcription induced by shear stress. A double AP-1 binding site in the VCAM-1 gene promoter was found to function as a cis-element for this negative transcriptional regulation. To determine the number of endothelial genes responsive to shear stress, differential display of endothelial mRNAs was performed. In cells exposed to a shear stress of 15 dynes/cm2 for 6 h, approximately 4% of the mRNA species increased more than two-fold or decreased to less than half the levels in static control cells. Thus, it seems that a large number of known or unknown shear-response genes are involved in blood flow-dependent phenomena including angiogenesis, vascular remodeling, and atherosclerosis. KEYWORDS: Shear stress, endothelial cell, vascular cell adhesion molecule-1, mRNA differential display, shear stress response element.
INTRODUCTION Endothelial cells (ECs) lining the inner surface of vessels have a variety of cellular functions and play an important role in the homeostasis of blood circulation and other body functions. For example, ECs regulate vessel tone by releasing a variety of smooth muscle cell relaxants and constrictors. They also express thrombomodulin and heparan sulfate on their cell surface to keep the vessel lumen anti-thrombotic. Furthermore, ECs actively interact with other cells through adhesion molecules and cell growth factors, and participate in tissue inflammatory and immune responses as well as vascular remodeling. It has long been considered that biochemical mediators such as hormones, cytokines and neurotransmitters control EC functions. In the last decade, however, it has become 111
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apparent that wall shear stress, a mechanical stress generated by blood flow, can also modulate EC functions. Increases in blood flow in vivo lead to enlargement of the vessel diameter, while decreases in blood flow reduce vessel diameter. This vessel response to blood flow is abolished when the endothelium is removed from the vessels, which indicates that ECs respond to blood flow changes. Actually, cultured ECs show alterations in function when exposed to shear stress in a flow-loading apparatus. For example, ECs increase the production of vasodilating substances such as prostacyclin, nitric oxide (NO), C-type natriuretic peptide (CNP) and adrenomedulin (AM), and decrease the production of vasocontricting factors such as endothelin (ET) and angiotensin converting enzyme (ACE) in response to shear stress. Shear stress affects transcription or mRNA stability which results in changes in mRNA levels. Study of the molecular mechanism for shear stress-mediated endothelial gene regulation is currently in progress. Some shear stress-associated transcription factors and shear stress-response elements have been identified in a couple of endothelial genes. In this review, the current knowledge of the action of shear stress on endothelial gene expression is discussed with a focus on our studies of the vascular cell adhesion molecule-1 (VCAM-1) gene, and of differential display in ECs exposed to shear stress. ENDOTHELIAL GENES THAT RESPOND TO FLOW In Vitro Data Endothelial genes that have been shown to respond to shear stress are listed in Table 6.1. The ET gene was first reported to be shear stress responsive by Yoshizumi et al. (1989). They exposed porcine ECs to shear stress (5 dynes/cm2) in a cone plate type flow-loading apparatus and observed a transient increase in ET mRNA levels reaching a peak at 2–4 h, after which it returned to control levels. In contrast, Sharefkin et al. (1991) demonstrated that when human umbilical vein ECs (HUVECs) were exposed to a shear stress of 25 dynes/cm2 for 24h, ET mRNA levels markedly decreased. Malek et al. (1992) also observed a sustained four- to Table 6.1 Shear stress-responsive genes in endothelial cells
up: mRNA levels are up-regulated by shear stress, down: mRNA levels are down-regulated by shear stress.
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five-fold decrease in ET mRNA in bovine aortic ECs (BAECs), which was evident within 1 h of the onset of shear stress (15 dynes/cm2) and completed by 2–4 h. Although the results vary, it may be that shear stress initially increases ET mRNA levels, but then decreases with time. Tissue plasminogen activator (tPA) has fibrinolytic activity, and Diamond et al. (1990) reported that tPA mRNA levels increased more than 10-fold after exposure of HUVECs to shear stress for 24 h (25 dynes/cm2). Hsieh et al. (1991) reported that shear stress (16 dynes/cm2) increased both platelet derived growth factor (PDGF)-A and -B mRNA levels in HUVECs, reaching a peak at 1.5–2 h and returning to control levels at 4h. The peak increase was a more than 10-fold increase for PDGF-A mRNA and a two- to three-fold increase for PDGF-B mRNA. Mitsumata et al. (1993) demonstrated in BAECs that although PDGF-A mRNA levels did not change, PDGF-B message began to increase within 3 h after the onset of shear stress (30 dynes/cm2), reaching a maximum at 6 h and remaining at high levels to 24h. Resnick et al. (1993) also reported an increase of PDGF-B mRNA induced by shear stress in BAECs. In contrast, Malek et al. (1993) demonstrated that shear stress (5 and 36 dynes/cm2) decreased the PDGF-B mRNA levels to around one-fourth the basal values in BAECs at 9 h. The exact reason for this discrepancy remains unclear. Both the basic fibroblast growth factor (bFGF) and heparin-binding epidermal growth factor-like growth factor (HB-EGF) genes are responsive to shear stress. Malek et al. (1993) showed an increase in bFGF mRNA levels increasing above control levels 4.8-fold at 6h and 2.9-fold at 9h after the onset of shear stress (35 dynes/cm2) in BAECs. Morita et al. (1993) observed that HB-EGF mRNA levels in HUVECs increased rapidly in response to shear stress (8 dynes/cm2), peaked at 3 h (4.5-fold increase) and returned to base line at 7 h. The NO synthase gene also responds to shear stress. Nishida et al. (1992) showed an significant increase in NOS mRNA levels induced by 24-h exposure of BAECs to shear stress (15 dynes/ cm2). Then their colleagues, Uematsu et al. (1995) reported that shear stress ranging from 1.2–15 dynes/cm2 increased ecNOS mRNA two- to three-fold the control level in a dose- and time-dependent manner in BAECs. Shear-induced upregulation of ecNOS mRNA expression was also observed in cultured human aortic ECs. The mRNA levels of protooncogenes such as c-jun, c-fos and c-myc also change in response to shear stress. Hsieh et al. (1993) observed a transient increase in c-fos mRNA levels, starting within 30 min after the onset of shear stress (16 dynes/cm2) and returning to the basal level within one hour. The mRNA levels of cjun and c-myc were also transiently increased by shear stress. Exposure of HUVECs to shear stress (16 dynes/ cm2) induced a two- to three-fold increase in monocyte chemotactic protein 1 (MCP-1) mRNA levels at 1.5 h, which returned to the control levels at 3–4 h (Shyy et al., 1994). Genes encoding endothelial adhesion molecules that mediate adhesion to leukocytes are also sensitive to shear stress. The first study of these molecules in the context of shear stress was done by Nagel et al. (1994), in which intercellular adhesion molecule1 (ICAM-1) mRNA in HUVECs was shown to increase in response to shear stress. ICAM-1 mRNA levels began to increase at 2 h after the onset of shear stress (10 dynes/cm2) and remained at increased levels at 24 h. Sampath et al. (1995) showed a
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transient increase in ICAM-1 mRNA levels induced by shear stress (25 dynes/cm2) in HUVECs, starting at 1–3 h and returning to the basal level at 6 h. Tsuboi et al. (1995) also observed that ICAM-1 mRNA levels increased and peaked at 8 h after the onset of exposure to shear stress (15 dynes/cm2). On the other hand, Ando et al. (1994) reported that VCAM-1 mRNA was decreased by shear stress (1.5 dynes/cm2) in murine lymph node venule ECs (MLVECs) in a time-dependent manner. Sampath et al. (1995) also observed the inhibition of VCAM-1 mRNA expression by shear stress (25 dynes/ cm2) in HUVECs. Nagel et al. (1994) and Sampath et al. (1995) reported that shear stress had no effect on E-selectin mRNA levels in HUVECs, but Ando et al. (1996) observed the induction of E-selectin mRNA by shear stress, the pattern of response of which was similar to that of ICAM-1 mRNA; E-selectin mRNA levels increased in a dose dependent manner (0–33 dynes/cm2) and reached a peak at 8 h after exposure to shear stress. The effect of shear stress on expression of the thorombomodulin (TM) gene, which plays a central role in anti-thrombotic activity of ECs, is controversial. Malek et al. (1994) reported that a shear stress of 4 dynes/cm2 did not change TM mRNA levels in BAECs, but that a shear stress of 36 dynes/cm2 resulted in a mild transient increase followed by a significant decrease in TM mRNA levels to 16% of its resting level by 9 h. In contrast, Takada et al. (1994) observed that the arterial levels of shear stress (15 dynes/cm2), but not the venous level of shear stress (1.5 dynes/cm2), increased TM mRNA levels in HUVECs, reaching a peak of a 3.5-fold increase over the control level at 8 h. The exact reason for this discrepancy is not clear but it may be due to differences in experimental conditions including the cell lines used. Ohno et al. (1995) showed that the transforming growth factor-beta (TGF-ß) gene was sensitive to shear stress. Exposure of BAECs to shear stress (20 dynes/cm2) induced a three- to five-fold increase in TGF-ß1 mRNA levels within 2h of exposure and elevated expression was sustained for over 12 h compared to static controls. The increase was in direct proportion to the intensity of the shear stress that ranged from 5–50 dynes/cm2. Expression of CNP mRNA was markedly increased by exposure to shear stress (24 dynes/cm2) at 3 h and this increase was maintained until 12 h (Okahara et al., 1995). Following this report, Chun et al. (1997) confirmed that the CNP gene was responsive to shear stress. A four-hour and a 24-hour exposure of HUVECs to shear stress (15 dynes/cm2) induced a six- and 30-fold increase in expression, respectively. Similar results were obtained in BAECs (4 h, two-fold increase; 24 h, three-fold increase) and in MLVECs (4 h, three-fold increase; 24 h, 10-fold increase). They also found that AM mRNA levels increased to 300% of the control levels in HUVECs at 24 h after the onset of shear stress (15 dynes/cm2). Superoxide dismutase (SOD) mRNA expression is also altered by shear stress. Inoue et al. (1996) observed an increase in Cu/Zn SOD mRNA reaching a peak at 24 h after the onset of shear stress (15 dynes/cm2) in human aortic ECs. Topper et al. (1996) observed an significant increase in Mn-SOD mRNA levels induced at 24 h after application of shear stress (10 dynes/cm2) in HUVECs, and demonstrated that cyclooxygenase (COX-2) mRNA levels were also increased by shear stress. ACE gene expression is down-regulated by shear stress. Rieder et al. (1997) observed that exposure of bovine pulmonary artery ECs to shear stress (20 dynes/cm2) decreased ACE mRNA levels to 18% of the control at 18 h. Lin et al. (1997) showed that
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exposure of HUVECs to shear stress (12 dynes/cm2) induced a transient response of tissue factor (TF) mRNA, where its levels began to increase at 1 h, reached a maximum at 2 h, and returned to the basal level by 6 h.
In Vivo Data The above mentioned data are from in vitro experiments using cultured cells and flow-loading devices, but recent studies have examined flow-induced endothelial gene expression in the animal. Kraiss et al. (1996) implanted bilateral aortoiliac prosthetic grafts and constructed femoral arteriovenous fistulas in baboons. Two months later they ligated one of the fistula, reducing shear stress. Four days after fistula ligation, they examined the changes in PDGF-A and -B mRNA expression using in situ hybridization and found that PDGF-B, but not PDGF-A, mRNA levels were significantly increased in low-flow grafts compared with high-flow grafts. Mondy et al. (1997) ligated branches of the right internal and external carotid arteries, reducing right common carotid artery blood flow while increasing flow in the left carotid. Compared with endothelium exposed to reduced blood flow significantly increased PDGF-A and -B mRNA expression, starting at 48 h and persisting until 72 h.
MOLECULAR MECHANISM FOR FLOW-INDUCED GENE REGULATION Using the VCAM-1 gene as a model, we propose a possible molecular mechanism for flow-induced gene regulation.
Fluid Flow Decreases Cell Surface VCAM-1 ECs express on their cell surface various adhesion molecules from the super immunogloblin, integrin, and selectin families, and can bind leukocytes via these adhesion molecules. Figure 6.1 shows microphotographs of cultured MLVECs immunostained with an antibody against VCAM-1. The lymph node venule is the base of lymphocyte homing phenomenon where a lot of lymphocytes adhere to the endothelium and migrate out into the extravascular space, eventually returning to the circulation via the lymphatic system. MLVECs, therefore, express abundant VCAM-1 which binds to very late activation antigen-4 on lymphocytes. When MLVECs were exposed to a shear stress of 1.5 dynes/ cm2 for 24 h in a parallel type of flow-loading chamber, cell surface VCAM-1 expression was markedly decreased (Figure 6.1). Flow cytometry confirmed that the VCAM-1 protein level was decreased by flow (Figure 6.2). Mean fluorescence decreased from 125.9 in static control cells to 82.3 in shear-stressed cells. In contrast, CD44 expression increased slightly in response to shear stress. The decrease in cell surface VCAM-1 was dependent on the magnitude and duration of shear stress. Prostacyclin and NO may affect the expression of adhesion molecules, and the release of these factors from ECs is enhanced by shear stress. However, in the presence of indomethacin or N -monomethyl- L-arginine,
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Figure 6.1. Fluorescence photomicrographs of MLVECs immunostained with antibody against VCAM-1. Static, static control cells; Flow, shear-stressed cells (1.5 dynes/cm2, 24h).
Figure 6.2. Flow cytometric analysis of flow-induced changes in the amount of cell surface VCAM-1 and CD44. Values in the column indicate mean fluorescence.
which inhibit production of prostacyclin or NO, respectively, VCAM-1 expression was decreased to an equal level by shear stress. The decrease in cell surface VCAM-1 induced by shear stress exerts an influence on the adhesiveness of MLVECs to lymphocytes. In a binding assay of MLVECs to murine lymphocytes, adhesion to lymphocytes was significantly inhibited in shearstressed MLVECs compared with static control ones.
Fluid Flow Lowers VCAM-1 mRNA Levels Shear stress lowered VCAM-1 mRNA levels in MLVECs. Figure 6.3 represents the time course of the changes in VCAM-1 mRNA levels induced by shear stress. Total RNA, which was extracted from static control and shear-stressed cells, was reverse
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Figure 6.3. Time course of the changes in VCAM-1 and CD44 mRNA levels induced by flow.
transcribed into cDNA and amplified by polymerase chain reaction (PCR). Shear stress decreased VCAM-1 mRNA levels in a time-dependent manner, whereas CD44 expression increased, showing a peak at 6 h after the onset of shear stress. GAPDH expression remained unchanged by shear stress. These results suggest that the decrease in cell surface VCAM-1 induced by shear stress is due to a decrease in VCAM-1 mRNA levels, and that the response to shear stress varies in individual genes.
Wall Shear Rate or Shear Stress Fluid flow has two modes of action. One is the flow-induced change in mass transport. Bio-active substances present in the fluid become increasingly available at the EC surface as flow rate or shear rate increases, leading to greater stimulation of ECs. The other mode of action is of shear stress as a mechanical stress which deforms and stimulates the ECs. To determine which action, shear rate or shear stress, is predominantly involved in the flow-induced decrease in VCAM-1 mRNA levels, flow-loading experiments using two perfusates with different viscosities were performed. This method allows us to apply different levels of shear stress to ECs at the same flow rate. MLVECs were exposed to flow of high (culture medium with 5% dextran) or low (culture medium) viscosity medium at various flow rates, and changes in VCAM-1 mRNA levels were examined. Figure 6.4A shows the relationship between mRNA levels and flow rate. The VCAM-1 mRNA levels decreased as flow rate increased, but the decreasing rate was larger in high viscosity medium, i.e., higher shear stress, compared with that seen in low viscosity medium.
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Figure 6.4. Shear stress-dependency of flow-induced decrease in VCAM-1 mRNA levels.
When the data were plotted against shear stress, they formed almost a single line as shown in Figure 6.4B. These findings suggest that the flow-induced decrease in VCAM1 mRNA levels is shear stress-rather than shear rate-dependent. Fluid Flow Affects Gene Transcription mRNA levels may be regulated either at the transcriptional level, or via mRNA turnover. The level at which shear stress regulates VCAM-1 gene transcription was first evaluated by the run-on assay. Transcription in MLVECs exposed to a shear stress of 3.5 dynes/cm2 for 24 h decreased to 68% of the static control levels. This was also confirmed by luciferase assay. A reporter gene consisting of the 5' flanking promoter region of chromosomal VCAM-1 gene (-3.7kb) cloned from MLVECs and luciferase vector was constructed and transfected into MLVECs. The cells were exposed to a shear stress of 3.5 dynes/cm2 for 24 h and their luciferase activities, which reflects the transcriptional activity of the VCAM-1 gene, were measured. As shown in Figure 6.5, VCAM-1 gene transcription was markedly suppressed by shear stress. Negative Flow-Response Element The fact that shear stress down-regulates VCAM-1 gene transcription suggests the presence of a cis-element in the promoter which is essential for shear responsiveness. Luciferase vectors with varying length of VCAM-1 promoter were transfected into MLVECs, and the cells were then exposed to shear stress. When the promoter region extended to -0.7 kb 5' upstream from the transcription start site, transeriptional activity decreased in response to shear stress, but when only –0.3 kb of promoter was present, there was no marked response to shear stress (Figure 6.5). These results suggest that there is a shear stress response element located between -0.7 kb and -0.3 kb of the VCAM-1 promoter. To isolate transcription factors that may bind this putative response element, twenty overlapping oligonucleotides of around 30 bases in length, based on the sequence from -0.7 kb to -0.3 kb, were synthesized and incubated with nuclear extracts obtained
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Figure 6.5. Deletion analysis of VCAM-1 promoter. -3.7 luc, -1.8 luc, -1.1 luc, -0.7 luc, and0.3 luc are reporter genes containing -3.7, -1.8, -1.1, -0.7, and -0.3 kb, respectively, 5' upstream from the transcription start site of the VCAM-1 promoter. * p<0.001.
from either static control or shear-stressed cells. Only three oligonucleotides formed distinct DNA-protein complexes in this gel shift assay (Figure 6.6). These oligonucleotides contains one or two tumor promoting agent response elements (TREs) with a sequence of TGACTCA. To ascertain whether the TRE actually functions as negative flow-response element, targeted mutations of the TRE sequence (GGACTTG) were made in the VCAM-1 promoter, which was then tested in the luciferase assay. The shear-induced suppression of VCAM-1 gene transcription was lost by mutation of one or both of the TREs. In a gel shift assay using oligonucleotides containing the mutated TRE and nuclear extracts obtained from shear-stressed cells, no distinct DNA-protein complexes were seen, indicating that no transcription factors bind to the mutated TREs. Taken together, it appears that a pair of TREs function as a single negative flow-response element in the flow-mediated down-regulation of VCAM-1 gene transcription in MLVECs (Korenaga et al., 1997). Transcription factor AP-1, which is either a homodimer of c-fos or a heterodimer of c-fos and c-jun, binds to the TRE, and AP-1 may be activated by shear stress in ECs. To examine whether AP-1 binds to TREs on the VCAM-1 promoter in shearstressed cells, a supershift assay using antibodies against c-jun or c-fos was performed. An antibody to c-jun partially inhibited the formation of the DNA-protein complex seen after the incubation of oligonucleotides containing TRE and nuclear extracts of shear-stressed cells. The c-fos antibody had no effect on mobility in this assay. Thus, c-jun and other transcription factors may be involved in the inhibition of VCAM-1 gene transcription by shear stress.
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Figure 6.6. Gel shift assay of DNA-protein complexes formed by oligonucleotides bearing double AP-1 consensus elements with nuclear extracts from shear-stressed of static cells. Lanes: 1, no nuclear extracts; 2–4, nuclear extracts from murine ECs exposed to shear stress of 3.5 dynes/cm2 for 24 h; 5–7, nuclear extracts from static murine ECs; 3 and 6, addition of the relevant unlabeled oligonucleotides as a competitor in 100-fold excess; 4 and 7, addition of unrelated DNA (EBNA-1); The arrow indicates the shifted band, protein-DNA complexes.
Positive Flow-Response Element Positive flow-response elements that are involved in the up-regulation of gene ex pression by shear stress have been identified in several endothelial genes. Resnick et al. (1993) showed that a GAGACC sequence locating in the PDGF-B gene promoter was essential for shear-induced up-regulation of PDGF gene expression, and named this the shear stress-response element (SSRE). Shyy et al. (1995) reported that proximal one of two TREs located in the MCP-1 gene promoter is a positive cis-element for shear stress. Lin et al. (1997) showed that the SP1 binding site (GCGGGGGCGGGG) in the TF gene is a positive shear stress-response element and Khachigian et al. (1996) indicated that an early growth response gene (Egr-1) binding site in the PDGF-A gene functions as a positive element. Although the sequence has not yet been determined, Malek et al. (1993) suggested that a shear inhibitory element exists between -2.9 and -2.5 kb 5' upstream region on the ET gene promoter. Ohno et al. (1995) reported the
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presence of a positive flow-response element between -453 and +115' upstream in the TGF-ß1 gene. These positive and negative flow-response elements reported to date are listed in Table 6.2. Results suggest that there may be a variety of shear stressresponse elements present in different genes. Transcription Factors Activated by Flow Several transcription factors that bind to flow-response elements have been identified (Table 6.2). Lan et al. (1994) showed that shear stress stimulates the DNA binding activity of nuclear factor kappa B (NF-B) and nuclear factor activator protein-1 (AP-1). When BAECs were exposed to a shear stress of 12 dynes/cm2, NF-B binding increased within 30 min, reaching and maintaining maximal levels at 1 h, while AP1 binding activity was biphasic, increasing four-fold within 20 min and returning to basal levels before steadily increasing by 2 h to a high level relative to basal values. Mohan et al. (1997) indicated that NF-B activation varied depending on the intensity of shear stress applied. Exposure of human aortic ECs to high shear force (16 dynes/ cm2) led to an early transient increase in NF-B expression, followed by return to basal levels in 2 h, whereas exposure to low shear stress (2 dynes/cm2) induced a significantly increased and prolonged NF-B activity. NF-B binds to the shear stressresponse element (GAGACC) of the PDGF-B gene (Khachigian et al., 1995), and AP1 binds to TRE which function as flow-response elements in the MCP-1 gene (Shyy et al., 1995). The transcription factors, SP1 and Egr-1, are also known to be activated by shear stress (Lin et al., 1997; Khachigian et al., 1997). Fluid Flow Affects mRNA Stability Recently, Kosaki et al. (1998) observed that shear stress stimulates ECs to produce the hematopoietic cytokine granulocyte/macrophage colony stimulating factor (GMCSF). GM-CSF mRNA levels were increased markedly by shear stress in a dosedependent manner. In this case shear stress had no influence on the transcriptional activity of the GM-CSF gene, but markedly lengthened mRNA half life. These results indicate that shear stress may regulate endothelial gene expressions at the post transcriptional level as well as at the transcriptional level. Table 6.2 Shear stress-response element of endothelial genes
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Flow Signal Transduction The fact that ECs alter their morphology and functions in response to shear stress strongly suggests the presence of a signal transduction pathway in ECs. However, this issue has not been fully clarified. A number of signal transduction factors are known to be activated by shear stress (Davies, 1996). For example, exposure of ECs to flow opens K+ channels leading to hyperpolarization of the cell membrane, activates Gproteins, increases inositol 1, 4, 5 triphosphate and intracellular Ca2+ concentrations, and increases a number of kinases such as protein kinase C and mitogen activated protein kinase. Recent work has suggested that changes in intracellular Ca2+ concentrations are closely related to activation of transcription factors. Although not associated with shear stress, a transient large increase in Ca2+ concentration occurring in mice B lymphocytes activates NF-B and c-jun N-terminal kinase, while a sustained small increase in Ca2+ concentration activates the nuclear factor of activated T cells (NFAT) (Dolmetsch, 1997). In a neuronal cell line, PC12, increases in cytoplasmic.Ca2+ concentration activate serum response factor (SRF), whereas increases in nuclear Ca2+ concentration activate cyclic AMP response element-binding protein (CREB), both of which lead to Ca2+-dependent induction of c-fos gene expression (Bading, 1997).
UNKNOWN ENDOTHELIAL GENES THAT RESPOND TO FLOW As shown in Table 6.1, to date around twenty endothelial genes have been identified as shear stress-responsive genes. This raises the question of the total number of genes that actually respond to shear stress. To answer this question, differential display of mRNA was carried out (Ando, 1996). Total RNA was extracted from HUVECs either incubated under static conditions or exposed to a shear stress of 15 dynes/cm2 for 6 h and reverse transcribed into DNA using the four oligo-dT primers (T12MG, T12MA, T12MT, and T12MC; M is a mixture of A, G and C) that anchor at the poly(A) tail of mRNA. The DNAs were amplified by PCR using four oligo-dT primers and twenty arbitrary primers anchoring at the 5' end of the mRNA molecule in the presence of DNA polymerase and -[32S]dATP. The amplified products were separated on a 6% DNA sequencing gel and the radioactivity of each band was measured. Increases in radioactivity of more than 200% and decreases to less than 50% were regarded as representing up-regulation and down-regulation, respectively, in response to shear stress. Laddering of the bands was seen in a total of 160 lanes. Only part of the differential display data is shown in Figure 6.7. Differential display of mRNAs was triplicated using different samples and only the reproducibly detected cDNA bands were analyzed. This method revealed a total of 1507 bands ranging from 100 to 500 bp in size. A total of thirty-three mRNAs were up-regulated by shear stress, while a total of twenty-seven were down-regulated (Table 6.3). Under the present experimental conditions, the ratio of shear-stress responsive mRNAs to total mRNAs detected was approximately 4% (60/1507). In general, mammalian cells express about 15,000 mRNAs. It is therefore possible that the expression of approximately 600 genes may be altered by 6-h exposure to shear stress (15 dynes/cm2).
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Figure 6.7. Differential display of endothelial mRNAs. S, mRNAs obtained from static control cells; F, mRNAs from flow-loaded cells (15 dynes/cm2, 6h).
cDNA products were cloned from sixteen of the shear stress-responsive bands and their nucleotide sequences were determined. They were all flanked by the sequences of the primer sets used and corresponded to the 3' end of the mRNAs, as expected. Six were known genes including laminin B-1 chain, H+-ATP synthase coupling 6, lysyl oxidase, myosin light chain kinase, ineterleukin-8 receptor and NADH dehydrogenase. Ten other cDNA fragments showed no significant homology to any known genes. Table 6.3 Number of endothelial mRNAs that are up- or down-regulated by shear stress
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CONCLUDING REMARKS Shear stress, a mechanical stimulus generated by blood flow, regulates the expression of a number of endothelial genes. The regulatory mechanism is as follows: shear stress is detected at the surface of ECs and the signal is transmitted into the cell interior via a second messenger system. Then transcriptional factors are activated and bind to shear stress-response elements regulating gene transcription positively or negatively, or in some cases altering mRNA stability. As shown in differential display of mRNA, shear stress affects the expression of a large number of genes, most of which remain unknown. The response of ECs to shear stress, therefore, can be considered to result from a net effect of a large number of known and unknown genes. Clarification of the details of the mechanisms of flow-induced gene regulation may provide new insights into the role of blood flow on blood flow-dependent phenomena such as angiogenesis, vascular remodeling and atherosclerosis. Furthermore, transcriptional factors activated by flow and flow-response elements may be used to develop new medicines and gene therapies for vascular diseases.
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changes in the expression of leukocyte adhesion receptors on human umbilical vein endothelial cells in vitro. Annals of Biomedical Engineering, 23, 247–256. Sharefkin, J.B., Diamond, S.L., Eskin, S.G., McIntire, L.V. and Dieffenbach, C.W. (1991) Fluid flow decreases preproendothelin mRNA levels and suppresses endothelin-1 peptide release in cultured human endothelial cells. Journal of Vascular Surgery, 14, 1–9. Shyy, J.Y-J., Hsieh, H.J., Usami, S. and Chien, S. (1994) Fluid shear stress induces a biphasic response of human monocyte chemotactic protein 1 gene expression in vascular endothelium. Proceedings of the National Academy of Sciences of the United States of America, 91, 4678–4682. Shyy, J.Y-J., Lin, M-C., Han, J., Lu, Y., Petrime, M. and Chien, S. (1995) The cis-acting phorbol ester “12O-tetradecanoylphorbol 13-acetate”-responsive element is involved in shear stress-induced monocyte chemotactic protein 1 gene expression. Proceedings of the National Academy of Sciences of the United States of America, 92, 8069–8073. Takada, Y., Shinkai, F., Kondo, S., Yamamoto, S., Tsuboi, H., Korenaga, R. and Ando, J. (1994) Fluid shear stress increases the expression of thrombomodulin by cultured human endothelial cells. Biochemical and Biophysical Research Communications, 205, 1345–1352. Topper, J.N., Cai, J., Falb, D., Michael, A. and Gimbrone, J. (1996) Identification of vascular endothelial genes differentially responsive to fluid mechanical stimuli: Cycloocygenase-1, manganese superoxide dismutase, and endothelial cell nitric oxide synthase are selectively up-regulated by steady laminar shear stress. Proceedings of the National Academy of Sciences of the United States of America, 93, 10417–10422. Tsuboi, H., Ando, J., Korenaga, R., Takada, Y. and Kamiya, A. (1995) Flow stimulates ICAM-1 expression time and shear stress dependently in cultured human endothelial cells . Biochemical and Biophysical Research Communications, 206, 988–996. Uematsu, M., Ohara, Y., Navas, J.P., Nishida, K., Nurphy, T.J., Alexander, R.W., Nerem, R.M. and Harrison, D.G. (1995) Regulation of endothelial cell nitric oxide synthase mRNA expression by shear stress. American Journal of Physiology, 269, C1371–C1378. Yoshizumi, M., Kurihara, H., Sugiyama, T., Takaku, F., Yanagisawa, M., Masaki, T. and Yazaki, Y.
(1989) Hemodynamic shear stress stimulates endothelin production by cultured endothelial cells. Biochemical and Biophysical Research Communications, 161, 859–864.
7 Endothelial Gene Regulation by Fluid Shear Forces Nitzan Resnick*, Efrat Wolfovitz and Shachar Zilberstein Department of Anatomy and Cell Biology, Bruce Rappaport Medical Research Institute, The Rappaport Faculty of Medicine—Technion P.O. Box 9697, Bat-Galim, Haifa, Israel 31096 Tel.: 972–4–8295201, Fax: 972–4–8520089, E-mail: [email protected].
The remodeling of Blood vessels accompanies physiological and pathological processes such as, angiogenesis and vasculogenesis, atherosclerosis, hypertension and restenosis. Vessel remodeling occurs in response to both biochemical and biomechanical stimuli, and has been shown to be dependent on the presence of an intact endothelial layer. By virtue of their anatomical position, endothelial cells are constantly exposed to hemodynamic forces generated by the flowing blood, forces that consists of fluid shear stress, cyclic strain and pressure. These forces affect endothelial cells structure and function, changes that are often mediated by the induction or shut-off of endothelial genes. Up to date few dozens endothelial genes have been found to be regulated by hemodynamic forces. Promoter analysis of some of the genes resulted in the definition of positive and negative cis-acting elements that are essential for their responsiveness to biomechanical forces. These shear stress response elements (SSREs) bind transcription factors, among them, NFB, NFATc2, Sp1, Egr1, Fos and Jun, that are activated themselves by hemodynamic forces. This review attempts to summarize the effects of hemodynamic forces, and more specifically fluid shear stress, on endothelial gene regulation in vitro and in vivo and point to several SSREs and transcription factors that are involved in this regulation. New technologies, as well as, new in vitro shear stress models facilitating the study of endothelial gene regulation by shear stress will be discussed. Finally, special attention will be given to results accumulating from recent studies on the regulation of endothelial genes by complex (pathological) shear stresses. Vascular remodeling is the sum of multiple events, some of which are regulated by biomechanical forces, and mediated by the regulation of vascular endothelial genes. It is thus hoped that unraveling the complexity of endothelial gene regulation by biomechanical forces contributes to our understanding of the physiological processes in the circulatory system and the pathogenesis of vascular diseases. KEYWORDS: Vascular wall, endothelial cells, fluid shear stress, gene regulation shear stress response element (SSRE), transcription factors.
HEMODYNAMIC FORCES IN THE ARTERIAL TREE The arterial tree which consists of large elastic and muscular vessels as well as arterioles
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and precapillaries vessels, is constantly exposed to hemodynamic forces varying widely in magnitude, frequency and direction. These forces consist of pressure acting perpendicular to the vessel wall, cyclic strain, and shear stress acting parallel to the wall, creating a frictional shear force on the surface of the endothelium. In large arteries the magnitude of shear stress is in the range of 10–40 dynes/cm 2, and it is over imposed with the pulsatile characteristic of the flow producing a range of shear stresses and shear stress gradients. In area of unique morphologies, such as curvatures and bifurcations, the steady laminar flow is disrupted to create regions of separated flow that include recirculation sites, which themselves may vary with the cardiac cycle. These secondary flows modify the profile of the original laminar flow therefore dictating the shear stress acting on the endothelium in these specific regions. Studies including in vitro modeling systems, as well as, in vivo measurements suggest that the values of shear stress in these regions vary from negative, to zero (in areas of flow separation) and up to positive values of 40 dynes/cm2. Under non-physiological conditions (hypertension) these values are even higher (1–8).
ENDOTHELIUM ADAPTATION TO BLOOD FLOW Large elastic arteries undergo dramatic adaptations in response to both acute and chronic changes in blood flow. This adaptation has been shown to depend upon the presence of an intact endothelial layer (9). The adaptation is mediated in part by the production of endothelial compounds that are vasodilators or vasoconstrictors, which act locally to modulate the vascular tone of the underlying smooth muscle cells (10–13). More chronic adaptation which occurs in pathological conditions or in experimental models also involves cellu lar remodeling that includes cell proliferation and programmed cell death (apoptosis) (14–16). The ability of the endothelium to sense hemodynamic forces was first documented more than 100 years ago by Virchow (17), who demonstrated the heterogenous morphology of the cells along the arterial tree. In a simple tubular section of the arteries, the cells are elongated and aligned in the direction of the flow. In regions of bifurcation, endothelial morphology varies widely from tightly packed cuboidal cells in areas of high shear stress, to more polygonal cells with no obvious orientation in the regions of disturbed non-laminar shear stress. These morphologies change abruptly over a very short distance of one or two cells. The dependency of vessel remodeling on the presence of an intact endothelium and the morphological fluctuations that reflect the local shear stresses, suggest that the vascular endothelium is a sensor of local fluid mechanical forces. The non-random distribution of early atherosclerosis lesions observed in naturally occurring disease in humans and in experimental animal models, points to local hemodynamic forces as crucial players in the development of this disease. Thus, hemodynamic forces are important stimuli effecting the biology of the endothelium, which plays a pivotal role in both physiological processes and pathological conditions (atherosclerosis, hypertension, thrombosis).
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ENDOTHELIAL RESPONSES TO SHEAR STRESS IN VITRO Evidence of the direct action of hemodynamic forces on endothelial structure and function come from in vitro studies in which cultured human and animal endothelial cells have been subjected to defined fluid mechanical forces (18–22). Results of these experiments mimic the in vivo response of the endothelium to different shear stresses. Endothelial cells exposed to unidirectional physiological laminar shear stress elongated and aligned in the direction of the flow, changes that were accompanied by the formation of stress fibers and redistribution of actin fibers and microtubules. Under disturbed laminar shear stress, these cells were polygonal with no directional orientation. Laminar shear stress led to cell cycle arrest, while low levels of either turbulent shear stress, for as little as 3 hours, or disturbed laminar shear stress triggered cell cycle entry without cell retraction or damage (23–26). Recent studies from several laboratories demonstrated the complex and wide broad effects of shear stress on endothelial cells. The response to shear stress begins almost instantaneously after the onset of flow, where acute changes in membrane structure, cytoskeleton organization and composition and phosphorylaton of focal adhesion proteins are observed (20, 27, 28). This response continues with changes in the activity and distribution of ion channels throughout the endothelial membrane (20, 29), alteration of intracellular calcium (30, 31), as well as activation and phosphorylation of many signaling molecules such as, G proteins, MAP kinase, Erk, Rho and others (32, 33), which will be broadly discussed throughout this book. Other immediate responses such as production of arachidonate metabolites and vasoactive mediators are also detected (10–13). Most of the immediate responses do not require protein synthesis and appear to involve regulation at the level of rate-limiting enzymes or substrate availability. The more delayed responses (minutes to hours after the onset of flow) include the up and down regulation of endothelial molecules many of which are key components in the physiological and pathological effector systems, located in the vascular endothelium that modulate, thrombosis and hemostasis, vascular tone, vascular growth, inflammatory and immune reactions. We and others have found that many of these molecules are regulated at the level of gene expression, thus providing a useful experimental paradigm for the investigation of gene regulation by biomechanical forces.
IN VITRO FLOW MODELS To explore the hypothesis that fluid shear stress acts directly on the endothelium to modulate its structure and function, several laboratories have designed and developed in vitro shear stress model systems. In the present chapter several of these systems will be described, and specific attention will be given to some new models recently developed. Two basic systems have been designed to study the effects of laminar shear stress on endothelial cells: First, the relatively simple parallel flow chamber, in which well developed laminar flows are generated by a pump device over a confluent endothelial
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monolayer grown on a transparent plastic or glass coverslip (21, 34, 35). In this device laminar shear stress is a linear function of the volume flow rate through the channels. The simplicity of this devise and the ability to couple it to an optical sieve and a microscope made it extremely useful in the study of endothelial cell—leukocyte interaction under flow. The hallmark of the parallel plate—the small amount of cells grown on the coverslip and the fact that each experiment consists of only one coverslip—brought to the development of a second model, the modified cone-plate viscometer (21, 23, 36). In this system shear stresses are produced in a layer of fluid contained between a stationary base plate and a rotating cone. By adjusting the angle of the cone, the speed it rotates, its distance from the plate and the viscosity of the medium a wide range of shear forces (1–50 dynes/cm2) and shear patterns (laminar, turbulent) can be generated. Once the cone is tipped in respect to the axis of the rotation, oscillatory flow can be also generated (37). The base of this apparatus can carry either small coverslips (up to 12), enabling the exposure of endothelial cells grown or treated under various conditions to the same flow field, or a single “maxi” plate exposing large number of cells to the same flow field. With the appreciation that complex shear stresses are involved in the development of cardiovascular diseases, more effort has been put in the design of new models mimicking these complex flow patterns, as well as combining shear stress with other hemodynamic forces, such as cyclic strain. The first model attempting to expose the cells to small defined areas of disturbed shear stress, was designed as a collaborative effort between the Vascular Research Division at the Brigham and Women’s Hospital and the Laboratory of Fluid Mechanics at MIT (38). This model which can be applied to both the parallel plate system or to the cone-plate apparatus, consists of a small bar added to the original coverslips where the endothelial monolayer is grown. The addition of the bar creates a barrier to the primary flow and forms complex secondary flows with large spatial variations in shear stress magnitude and pattern. Indeed, several studies using this model have demonstrated that cells in the region of flow separation behave differently then the cells on the very same coverslip located downstream from the bar, and exposed to simple physiological shear stress. Gene expression studies which will be described in details in this review also revealed pattern of expression unique to disturbed laminar shear stress (38–41). Three new models that are based on the original parallel plate, in which shear stress gradients are created were also described. In the arrow head model (42) which consists of two parallel plates in the shape of an arrow, endothelial cells are exposed to variations in both shear stress pattern and magnitude along the arrow and in the sides of the arrow head. Using this model it has been demonstrated that while the expression of tight junction molecules was elevated under laminar shear stress (and was dependent on shear stress magnitude), almost no expression of these molecules was observed in regions where the flow stagnates. In another model (43) laminar shear stress gradients are achieved by changes in the width of the parallel plate, which creates shear stress magnitudes ranging from 50 dynes/cm2 at the flow entrance to zero shear stress at the exit. Adhesion of platelets to endothelial cells using this model was reported to be shear stress magnitude dependent. The pulsatile nature of the flowing blood, shear stress gradients and complex shear stress patterns, were all taken into account in the third model based on the parallel plate which is described in details in chapter six in
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this book. Studies carried out in all four models, applying complex shear forces to endothelial cells, strongly agree with the in vivo observations that non-laminar and laminar shear stress have different effects on the cells. The main hallmark of all four models remains the small number of cells localized at the area of flow stagnation. Finally, the tubular shape of the vessel was taken into account in the following models (44, 45), in which the cells are grown within short silicon tubes coated with fibronectin, and subjected to laminar or pulsatile flow. The elastic properties of the tube allow to expose the cells to shear stress alone (the tube is casted with a rigid shell), cyclic strain alone (no flow) or the combination of both. Subjecting endothelial cells to the combination of cyclic strain and shear stress mimics better the complexity of hemodynamic forces to which the vascular endothelium is subjected in vivo. Shear stress gradients or non-laminar shear stress can be achieved in these models by ligation of the tube. ENDOTHELIAL GENE REGULATION BY LAMINAR SHEAR STRESS The development of in vitro shear stress models in which the cells are subjected to well defined shear stresses, together with the application of molecular biology techniques, lead in 1990 to the discovery that the regulation of several endothelial molecules by shear stress occurs at ‘the level of the transcript (46, 47). More than thirty endothelial genes have been found since than to be regulated by laminar shear stress in vitro, and they are grouped on Table 7.1 according to their cellular function. Among them are growth factors, cytokines and chemokines, vasoconstrictors and vasodilators, extracellular matrix molecules and their modifying enzymes, oxidizing enzymes, adhesion molecules, thrombolytic molecules, immediate early genes, transcription factors and signaling molecules (46–71). It is important to note that this is only a partial list of cloned genes which
Table 7.1 Endothelial genes transcriptionally regulated by laminar shear stress
Table 7.1 presents a list of genes that their cellular function is known and are regulated (up and down) by laminar shear stress, in both large vessels and lymph node endothelial cells. * N.Resnick, unpublished results.
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have a known function. Many additional genes (estimated as 600 novel sequences) were recently identified to be shear stress responsive, using differential display techniques (65, 68) and are waiting to be cloned. In the following chapter the patterns of the temporal expression of endothelial genes by laminar shear stress will be discussed with the emphasis on the source of the endothelium and the magnitude of shear stress. Most of the studies recently published, describing the regulation of endothelial genes by laminar shear stress, were carried out using endothelial cells from large vessels—namely aorta, arteries and veins. Interestingly, the pattern of regulation was not dependent upon the species (human, bovine, porcine) of the endothelium. In endothelial cells derived from large vessels three temporal patterns of regulation are found: The first group which is the largest one, includes genes that are induced by laminar shear stress. This group can be further divided into two subgroups—the immediate early genes which are rapidly induced (minutes) by shear stress and rapidly decline (minutes—hours) to their static levels, and genes that are rapidly induced by the onset of flow but slowly (hours—days) decline back to their static level of expression. The first subgroup includes c-fos, c-jun, c-myc, egr-1, as well as PDGF A and B chains (47, 61, 62). The second subgroup of genes includes—ICAM-1, tPA, TGF-B, ecNOS, COX2 (46, 48, 52, 68) and others (see Table 7.2). It is yet to be defined whether their prolonged induction is the result of message stabilization, or alternatively, prolonged upregulation of transcription. Within the second group are genes that respond to laminar shear stress in a biphasic pattern. These genes are upregulated acutely after their exposure to flow (1.5–4 fold), a phase that is followed by a dramatic drop in the transcript level (several folds) below static control. This group includes endothelin 1 (ET-1)(56), angiotensin converting enzyme (ACE) (59), thrombomodulin (70) and monocyte chemotactic protein1(MCP-1) (63). Both induction and suppression involve regulation at the transcription level, and are not the result of message stabilization/ Table 7.2 The pattern of endothelial gene regulation by uniform laminar shear stress
Table 7.2 summarizes the various patterns of endothelial gene regulation by laminar shear stress. The expression of all genes except for VCAM-1 was tested in large vessel endothelial cells.
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destabilization. One may speculate that the primary induction of these genes results from the transition of the cells from static conditions to shear stress, while the net effect of shear stress is the decrease in transcription. The molecular mechanisms underlying both the acute increase, and the chronic transcriptional decrease are not fully understood. To the last group of shear stress responsive genes belong two members, plasminogen activator 1 (PAI1) (55) and Endothelin Converting Enzyme (ECE) (57), which are suppressed transcriptionally by shear stress in a time dependent fashion. Several genes were demonstrated to be non-responsive to shear stress, many of which are “house keeping” genes. These include enzymes like GAPDH, or cytoskeleton elements like tubulin. There are also certain genes which are quiescent in unstimulated endothelial cells grown under static conditions, such as VCAM-1 and E-selectin. Because of their low transcript level under static conditions, it was hard to determine whether they are affected by shear stress. Two alternative approaches were taken to test the responsiveness of these genes to laminar shear stress: Endothelial cells from large vessels (HUVEC) were preconditioned for 24 hours under laminar shear stress. The flow was then halted and the cells were treated with cytokines. Under these conditions both E-selectin and ICAM-1 transcription was identical to their expression in HUVEC solely treated with cytokines. In contrast, the expression of VCAM-1 was much lower in cells that were preconditioned under shear stress prior to cytokine treatment (72). These results suggested that VCAM-1 expression is suppressed by shear stress, suppression which is prolonged enough to affect the transcription after flow cessation, and during the incubation with cytokines. Interestingly, if the cells are first treated with cytokines and then exposed to several hours of shear stress, all three molecules are transcriptionally induced, similarly to cells stimulated with cytokines alone (40). Taken together these results suggest that 1) E-selectin expression is not affected by shear stress and 2) VCAM-1 transcription is suppressed by laminar shear stress. Suppression of VCAM-1 by shear stress was further supported by the use of endothelial cells from lymph nodes (49, 73). Murine endothelial cells derived from lymph nodes exhibited significant expression of VCAM-1 under static conditions, without exposing the cells to cytokines or endotoxin. VCAM-1 expression in these cells, after their exposure to shear stress, was markedly reduced in a time dependent manner. The responsiveness of endothelial genes to shear stress can be also categorized according to the dependency on the magnitude of the laminar flow. As already noted by Malek et al. (74) the responsiveness of most genes is dependent on the magnitude of shear stress—higher magnitude of flow correlates with a more prominent change in the expression of the gene (whether they are induced or suppressed). Few exceptions are known to this phenomenon, ICAM-1 (48) PDGF-B and MCP-1 (74, 63) are regulated by shear rates varying from 2 to 46 dynes/cm2 to similar levels. A potential explanation to such a phenomenon may be the responsiveness of these genes not to the magnitude of force, but to changes in shear rates (static to flow, high flow to low flow etc.). This hypothesis is supported by the elevated expression of ICAM-1 in vivo in areas of shear stress gradients (75) as will be discussed later.
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TRANSCRIPTIONAL REGULATION BY SHEAR STRESS Run-on assays and treatment with actinomycin D verified that the regulation of most of the genes, described in the previous chapter and in Table 7.2, is trasncriptional. In order to further elucidate the mechanism of the transcriptional regulation our group has focused on PDGF-B as a gene model. A reporter gene construct consisting of 1.3 Kb fragment of the human PDGF-B promoter coupled to chloramphenicol acetyl transferase (CAT) reporter gene was transfected into endothelial cells. Interestingly, like the endogenous gene, the induction of the construct occurred minutes after the onset of flow, which further supported the fact that the regulation of PDGF-B is indeed trasncriptional. To localize element(s) within the promoter that are shear stress responsive 5'—nested deletion analysis was used. Upon deletion of a 50bp elements lying between positions –153 to –101, the responsiveness of the promoter construct to shear stress was ablated. Computer analysis revealed that this region encoded for one binding site for a known transcription factor Sp1. Electromobility shift assays (EM S A) enabled us to better localize the shear stress responsiveness to a 6bp element within this region. This element (GAGACC), termed shear stress response element (SSRE), is not a consensus binding site for known transcription factors, binds nuclear proteins from endothelial cells that were exposed to shear stress and is encoded in the promoters of many additional shear stress responsive genes (37). Addition of this SSRE (either in its sense or in its anti-sense orientation) to a viral SV40 promoter, in hybrid promoter vector system, rendered the non- shear stress responsive promoter into a responsive one (76). Screening for nuclear proteins that bind to this element revealed that heterodimers of p50 and p65 members of the NFB family are able to bind to the SSRE (77). This binding was demonstrated by EMSA and super-shift assays, as well as, by DNase footprinting analysis. To our surprise we have found that NFB is not the only factor able to bind to this SSRE. Nuclear factor of activated T cells (NFAT), which is mostly common in T cells and to a lesser extent in B cells, has been found by us to be activated by shear stress in endothelial cells. This factor that share some similarity in its DNA binding site to the Rel family, is able to bind to this SSRE as well, and to compete with NFB on the site (78). The cooperative binding of these factors to the SSRE under static and shear stress conditions is under current investigation. Interestingly, the induced binding of nuclear proteins to the SSRE is not limited to endothelial cells exposed to shear stress, but is also found in endothelial cells (but not smooth muscle cells) exposed to cyclic strain although in a different time kinetics (77). The nature of the proteins that bind to the SSRE in endothelial cells exposed to cyclic strain has not been revealed yet. In both PDGFB and tPA the presence of the SSRE did not mediate the responsiveness of these genes to cyclic strain, although it was crucial for their induction by shear stress (79). The heterogeneous response of endothelial genes to shear stress suggested that transcriptional regulation by shear stress is a complex process and involves various shear stress response elements. Indeed, shortly after the identification of the first PDGF-B/SSRE other SSREs were also defined. Although the promoter of MCP-1 contains the PDGF-B/SSRE, additional element—tPA response element (TRE) mediates the responsiveness to shear stress. Furthermore, although two TRE sites are
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localized in tandem in this promoter in a 38bp region (–53 to –90 upstream to the initiation of transcription), only the TRE site proximal to the transcriptional initiation mediates shear stress responsiveness. TRE is a binding site for the transcription factors c-fos and c-jun and mediates shear stress responsiveness of hybrid promoter constructs in both HeLa and endothelial cells (63). Recently two additional positive SSREs were defined in the promoters of PDGF-A and Tissue Factor genes (54, 62). PDGF-A is rapidly induced in endothelial cells following their exposure to shear stress, and is also regulated trasncriptionally. The promoter of PDGF-A is a G-C rich promoter, and does not contain neither PDGF-B/SSRE nor TRE. Using 5' nested deletion analysis a 50bp region proximal to the initiation of transcription was identified. This element contains a dual biding site for the transcription factors Egr1 and Sp1. Gel shift analysis with this binding site as a probe revealed the formation of several DNA/protein complexes under both static and shear stress conditions, with a unique complex formed under shear stress conditions. The addition of antibodies against either egr1 or sp1 in the binding reaction, as well as, competition assays with recombinant egr1 and sp1 proteins revealed that under static conditions the site is bound by sp1. Immediately after the onset of flow egr1 is rapidly (minutes) induced. The elevated levels of egr1 in the cell and its localization in the nucleus lead to the displacement of sp1 from the binding site, its replacement by egr1 and increased transcription of PDGF-A. Similar mechanism is responsible for the induction of PDGF-A by PMA. The fourth positive SSRE was recently defined in the Tissue Factor promoter and is a binding site for sp1. Recently, the first negative SSRE was also defined in the promoter of VCAM1(73). As mentioned in the previous chapter VCAM-1 is expressed under static conditions in murine lymph node endothelial cells, and is down regulated by shear stress in a time and magnitude dependent manner. VCAM-1 promoter contains strong NFB sites, an Sp1 site, intereferon response element, as well as two AP-1 sites. Interestingly, these AP-1 sites (both the distal and the proximal) are the elements that mediate VCAM-1 repression by shear stress. The exact proteins that bind to these site under shear stress have not been defined, although it is clear that c-fos is not involved in the complex. Whether these AP-1 sites are occupied by c-jun dimers solely, or additional proteins participate in the complex is yet to be defined. It is also not clear whether this AP-1 site mediates shear stress repression of additional genes such as PAI-1, ECE, ACE and MCP-1. A 400bp fragment has been localized (-2900 to -2500 up stream to transcription initiation) in the promoter of endothelin 1 which encodes for a yet to be defined element, responsible for the repression of this gene by shear stress (56). The characterization of promoter cis-element which positively or negatively mediates the regulation of endothelial genes by laminar shear stress (Table 7.3) lay the base for further investigation of the mechanism through which hemodynamic forces regulate endothelial gene expression. At the same time it also raised several interesting and unanswered questions which demonstrate the complexity of gene regulation by fluid shear stress. First, as discussed in this chapter, several positive SSREs are already defined, each of them was characterized in a particular gene. Sequence analysis of these promoters often reveals that each promoter contains the combination of various SSREs. Does the presence of these SSREs support the
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Table 7.3 Positive and negative shear response elements (SSREs) and their bound nuclear proteins
Table 7.3 presents defined positive and negative SSREs and the nuclear proteins that bind to them. Note that additional proteins may also participate in the formation of the complex with the SSREs.
responsiveness of the genes to shear stress through a cooperative mechanism (see Figure 7.1)? Second, the sequence of AP-1 and TRE resembles and similar transcription factors c-fos and c-jun bind to these elements, yet, TRE is a positive SSRE defined in MCP-1 and AP-1 is a negative SSRE defined in VCAM-1. Which are the parameters dictating the effect of these promoter elements on gene transcription—the promoter environment, namely the presence of binding sites to additional transcription factors, or the tissue origin of the endothelium? Finally,
Figure 7.1. Model of shear stress induced enhancer. The exposure of endothelial cells to shear stress lead to the enhanced binding of fos and jun dimers to a TRE site, displacement of Sp1 by Egr1, binding of phosphorylated SP1 (p-SP1) to its cognate site and enhanced binding of both NFB heterodimers and NFAT molecules to the PDGF-B/SSRE as well as additional yet be defined transcription factors (?). Some of these transcription factors like fos-jun and NFAT, Sp-1 and NFB, or fos-jun and NFB may interact with each other. As many promoters of shear stress responsive endothelial genes contain several SSREs, we suggest that these SSREs and their bound transcription factors form cooperatively a tertiary transcriptional complex, the shear stress induced enhancer which interacts as a unit with the basal transcriptional apparatus.
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PDGF-B SSRE is the only novel, non consensus SSRE defined so far, while all the other positive and negative SSREs are shared by many other genes, not all them are shear stress responsive. VCAM-1 for example contains in its promoter NFB, AP-1 and Sp-1 sites, but is suppressed by shear stress. PDGF-A contains in its promoter a dual binding site for Sp1 and Egr1, but under shear stress Sp-1 is displaced by Egr1 from its binding site and does not mediate shear stress responsiveness. In contrast, similar site in the tissue factor promoter is occupied solely by Sp1, under both static and shear stress conditions. Thus the complex temporal pattern of gene regulation described in the previous chapter, is amplified by the complexity of the mechanism (namely, promoter elements and their bound transcription factors) through which these genes are regulated. This complexity may be reflected by the synergism or antagonism of positive and negative promoter SSREs and their bound nuclear proteins, and may be dependent upon the tissue source of the endothelium and the growth conditions of the culture.
ACTIVATION OF TRANSCRIPTION FACTORS BY LAMINAR SHEAR STRESS The definition of promoter elements that mediate shear stress responsiveness, through the binding of nuclear proteins, lead to extensive studies on the nature of these proteins and the mechanism of their activation by shear stress. The regulation of transcription factors by shear stress was already suggested by Hsieh et al. (61) several years ago, studying the expression of immediate-early genes in endothelial cells exposed to shear stress. These genes included c-fos, c-jun and c-myc, which were demonstrated to be rapidly (minutes) induced by laminar shear stress, and rapidly return to their static level. Recently, egr1 (also an immediate early gene) was also demonstrated to be transcriptionally induced by shear stress (62). The immediate response of many shear stress responsive genes to shear stress, and the immediate binding of shear stress responsive nuclear proteins to the various SSREs, suggested that some of these factors may be regulated through mechanisms which do not involve transcription. Several transcription factors have been shown recently to be activated by shear stress through post translational modification or changes in their cellular localization. Among these factors is c-fos which rapidly translocates into the nucleus in endothelial cells exposed to arterial levels (25 dynes/cm2) of shear stress, in a process that involves protein kinase C, G proteins, phospholipase C and intracellular calcium. Likewise, the levels of Sp-1 are not affected by laminar shear stress but the phophorylation of the protein (and therefore the level of its activation) is enhanced by flow (54). Nuclear factor B (NFB)—a family of transcription factors that has been extensively studied in the past ten years, was first identified as a regulator of the kappa-light chain gene in B lymphocytes and was suggested to play a role in tumorogenesis inflammation, and cellular processes such as cell cycle and apoptosis (80, 81). NFB resides in the cytoplasm of non-activated cells most commonly as a heterodimer consisting of the two molecules p65 (rel A) and p50, but other members such as rel, relB, v-rel and p52 may also be part of the complex. In non-activated cells
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the complex is bound to an inhibitor—Inhibitor of kappa B (1B) which is also a multi-member family. The binding of the inhibitor to p65 masks its nuclear localization signal, therefor inhibiting the migration of the complex into the nucleus. The activation of NFB is a multi- step event, that includes the phosphorylation of IB by a complex of kinases only recently cloned (82), the addition of ubiquitin molecules to the phosphorylated IB, the degradation of the inhibitor by the proteasome complex and the entrance of the NFB complex into the nucleus. In the cytoplasm of non-activated cells p65 is bound in an additional complex with either p105, the precursor of p50, or p100 the precursor of p52. The binding of the precursors to p65 also masks its nuclear localization signal. Activation of the cells which results in enhanced phosphorylation of the precursors and their processing, unmasks p65 nuclear localization signal, leading to the migration of the processed complex into the nucleus. Various members of both NFB and IB are found in non-activated endothelial cells including p105/p50 p100/p52, p65, c-rel and relB, as well as, IBa, IBß and bcl3. In non activated endothelial cells p50 and p65 are found mainly in the cytoplasm and to some extent in the nucleus, unless the cells are grown in low serum conditions (for review see 83). IB and ß both reside in the cytoplasm while bcl-3 is found in the nucleus. Activation of endothelial cells with biochemical stimuli such as cytokines, LPS, poly d(I:C), oxidizing agents and others, lead to the activation of NFB and the nuclear accumulation of mainly p65/p50 heterodimers. This activation can be inhibited by anti-oxidation agents, as well as, proteasome inhibitors. As IB (but not IBß) contains an NFB binding sites in its promoter, the activation of NFB leads to enhanced transcription and translation of IB molecules and finally through a negative feedback mechanism, to the inhibition of NFB. Interestingly, it was recently demonstrated that the increased expression of IB, 60 minutes after the treatment of the endothelial cells with TNF, lead to the migration of IB molecules into the nucleus, resulting in the displacement of p65/p50 complexes from their DNA binding site. The migration of IB into the nucleus required protein synthesis and was dependent upon over expression of newly synthesized IB molecules (84). Accumulating evidences suggest that NFB plays a major role in the pathophysiology of blood vessels. Activated NFB molecules have been observed in various atherosclerosis models, as well as, various models of endothelial injury (85, 86). In vitro functional NFB binding sites are found in many genes whose expression is induced in endothelial cells at sites of inflammation and atherosclerosis (83). Although the activation of endothelial NFB by various biochemical stimuli is extensively studied, very little is known on the activation of this important family by hemodynamic forces. Recently, the activation of NFB by physiological levels of laminar shear stress has been demonstrated by Lan et al. using an NFB consensus binding site as a probe (87). The ability of p65/p50 heterodimers to bind to the PDGF-B/SSRE (77) suggested that NFB may play an important role in gene regulation by shear stress. Using immunofluorescence staining our group has demonstrated that as early as 10 minutes after the onset of flow both p65 and p50 migrate into the nucleus in both BAEC and HUVEC. The migration into the nucleus was accompanied by an increased binding of the heterodimer to either its cognate site or to the PDGF-B/SSRE, as demonstrated in EMSA and supershift assays. Interestingly, the intensity of NFB/DNA complexes
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in cells exposed to laminar shear stress is always weaker than in cells treated with TNF. To our surprise, we have found that in endothelial cells exposed to short intervals of shear stress (minutes) both IB and IBß migrate into the nucleus. IB molecules were also detected in the nuclei of endothelial cells subjected to shear stress, minutes after the onset of flow, by western immunoblotting. Unlike endothelial cells treated with cytokines, where IB rapidly degrades and can not be detected in the cytoplasm minutes after the treatment, cytoplasmic IB signals in cells exposed to laminar shear stress at various time points after the onset of flow, were stronger than the signals of static controls (manuscript in preparation). These results raised several questions—1. Is the rapid accumulation of IB molecules in the cytoplasm of endothelial cells exposed to shear stress the result of massive denovo IB synthesis, the lack of IB degradation or the combination of both? 2. Does the migration of IB molecules into the nucleus of sheared cells require denovo IB synthesis? And finally, 3. Does the presence of IB in the nucleus lead to the displacement of NFB complexes from their binding site? Using EMSA and supershift we were able to demonstrate that IB molecules participate in NFB/DNA complexes shortly (minutes) after exposing endothelial cells to shear stress, suggesting that this rapid binding of IB to NFB/DNA complexes may indeed displace NFB from its binding site. These results may explain the weak intensity of NFB/DNA signals in endothelial cells exposed to shear stress as compared to endothelial cultures treated with cytokins, suggesting that NFB molecules which exists in the nucleus after the onset of flow, may not be active in transcriptional regulation because of the presence of IB in the nucleus. Such situation may explain why genes like VCAM-1 and E-selectin, which contain strong NFB sites in their promoters are not upregulated by laminar shear stress. Further studies are ongoing in our group to identify the various stages of NFB activation/ IB degradation in endothelial cells exposed to laminar shear stress, to elucidate the rapid mechanism of IB induction by shear stress and finally to better understand the mechanism of IB rapid migration into the nucleus. These preliminary results thus imply that shear stress may regulate the activation of transcription factors via unique pathways. The studies of transcription factors activation by hemodynamic forces is only at its first stages. Results from these studies are crucial for better understanding of the mechanism through which biomechanical forces regulate endothelial gene transcription. Better understanding of the activation steps of these transcription factors and the various molecules and cellular components that play a role in them, may prove useful in our attempt to unravel the signaling pathways through which endothelial cells translate biomechanical forces into biochemical and genetic events.
ENDOTHELIAL GENE REGULATION BY COMPLEX SHEAR FORCES Throughout the previous chapters, our discussion was limited to the regulation of endothelial genes by continuous physiological laminar shear stress, applied to cultured endothelial monolayers. Although these simple in vitro systems have
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yielded new information and laid the firm base for the paradigm of gene regulation by biomechanical forces, they fail to mimic the complexities of the in vivo biomechanical environment in several respects: 1) Most of the experimental systems descried have used static endothelial cells that were abruptly move into flow environment. This transition from no-flow to flow, has only few analogues in vivo, in neonatal pulmonary circulation after the closure of the ductus arteriosis, or following surgical procedures. 2) Several studies have recently shown that prolonged exposure of endothelial cells to laminar shear stress lead to flow accommodation, namely the transcript levels of many genes, return to static levels after several hours exposure to shear stress (51, 56, 63, 70). Alternatively, experiments in which the flow has been turned on and off several times lead to endothelial gene induction. These results raise the question of the importance of preconditioning the culture to flow for extended periods (hours), and changes in shear stress magnitude within the experimental design. 3) The in vivo distribution of atherosclerotic lesions suggest that more complex shear forces are associated with the development of cardiovascular diseases. The in vitro generation of complex flow over endothelial monolayers may prove useful when studying the development of cardiovascular diseases. These complex shear stresses act in vivo in combination with additional hemodynamic forces such as pressure and cyclic strain. Very few studies have combined so far these forces in vitro, while studying the expression of endothelial genes. The present chapter thus attempts to summarize the few in vitro and in vivo studies that applied more complex flow patterns and tested the expression of endothelial genes under these conditions (See Table 7.4). Some of the results presented here are preliminary, but may point Table 7.4 Regulation of endothelial genes by complex shear stresses
Table 7.4 summarizes the pattern of regulation of endothelial genes by complex shear stresses. All experiments were carried out in large vessel endothelial cells. Shear stress gradients were created in vivo by carotid ligation using various animal models.
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to new important directions and serve as milestones in the field of gene regulation by complex hemodynamic forces. Endothelial cells were exposed to pulsatile flow and the expression of immediate early genes was tested and compared to the expression in endothelial cells exposed to laminar shear stress. While the induction of c-myc and c-jun was comparable under pulsatlie and laminar shear stress, c-fos, as well as PDGF-A expression was higher under pulsatile flow (61). Several genes responded similarly to laminar or turbulent flows among them are PDGF-B, TGF-ß, thrombomodulin and b-FGF (51, 56, 70). The expression of several large vessels endothelial genes was tested under oscillatory shear stress. The expression of heme-oxygenasel (HO-1) and Cu/Zn superoxide dismutase (SOD) was tested in HUVEC exposed to increasing magnitudes (5–25 dynes/ cm2) of oscillatory shear stress (67). The expression of HO-1 was induced by the oscillatory flow and remained elevated for more than 24 hours, while the expression of SOD was not affected by this flow pattern. This is in contrast to steady laminar shear stress, where HO-1 expression increased for short periods of time (up to 5 hrs) while SOD transcription increased for over 24 hours. Interestingly, VCAM-1 expression is down regulated by laminar shear stress (49), but is induced in HUVEC exposed to oscillatory shear stress (5 dynes/cm2), induction that was inhibited by pretreatment with antioxidants (72). The complexity of the flow in areas of bifurcation and curvatures which are predisposed to the development of atherosclerotic lesions was already discussed earlier in this review. These regions are characterized by the combination of shear stress gradients and changes in the flow pattern. The contribution of each of these flow components to the development of the lesion is not known, and in order to study this question two approaches have been taken. In the first experimental approach endothelial cells (mostly in vivo) are exposed to changes in the magnitude of shear stress, and the expression of various genes relevant to the development of atherosclerosis is tested. In the second approach in vitro models are used in which the cells are exposed to both spatial and temporal changes closely mimicking the pattern of flow in lesion— prone areas. Ligation of the carotid artery at one side, using various animal models (rabbits, ship, baboons) served to test the effect of changes in the magnitude of shear stress on gene expression. In these models endothelial cells are preconditioned to constant laminar shear stress, and are than transferred to either decreased or increased shear rates, depending on the position in the carotid artery. Endothelial cells in areas of low shear stress have shown an increase in lipid uptake and monocyte adhesion (89). Interestingly, this phenomenon was accompanied by an increase in the transcript levels of VCAM-1 (75), PDGF-A (90) and the transcription factor Egr-1 (91). VCAM-1 expression was also slightly induced by elevated levels of shear stress, in contrast to ICAM-1 expression which was solely induced by the transition to high levels of shear stress (75). Taken together these results point to the importance of preconditioning endothelial cells by uniform flow, and points to the fact that the pattern of endothelial genes expressed under constant laminar shear stress differs from that of the genes expressed under shear stress gradients. The combination of complex spatial and temporal shear stresses and its effect on endothelial gene expression was studied by several laboratories using the various in
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vitro flow models discussed in previous chapters. Connexin 43 is a gap junction molecule that has been shown to be upregulated by laminar shear stress. Using the model developed by DePaola et al. it has been shown that in areas of flow circulation connexin 43 expression increased 8–10 folds compared to the static controls an increase that lasted for more than 16 hours (41), a time point in which the expression of the molecule downstream to the bar (where the cells are exposed to constant laminar shear stress) decreased to static levels. Using the same shear stress model we have found that large vessels endothelial cells only at the region of flow circulation, also expressed elevated levels of VCAM-1 (40 and Resnick, N. unpublished results). The unique pattern of gene expression under complex flow patterns led to the hypothesis that different cis-element as well as transcription factors play a role in regulating gene expression in these regions. The activation of various transcription factors under disturbed-laminar shear stress was studied by Nagel et al. (40, 92) and compared to the activation of these molecules under static conditions or under uniform laminar shear stress. Four factors were tested, NFB (p65), Egr1, c-jun and c-fos, all of which are upregulated by laminar shear stress. Preliminary results have shown that all factors were further induced (total protein content in the cell) by non-uniform flow, with the most prominent induction at the reattachment point. Furthermore, nuclear localization of all four proteins was significantly higher in area of disturbed flow, where the most striking difference between laminar and disturbed shear stress observed for Egr1. It is important to emphasize that in this study only one time point was tested (30 minutes) after the onset of flow, and all factors tested are already known to be induced by uniform laminar shear stress. Based on the results obtained by Polaceck et al., demonstrating significant differences in time kinetics of gene induction by laminar and disturbed flow, it would be interesting to test the regulation of these transcription factors at more chronic time points following the onset of flow. The regulation of genes such as VCAM-1 by disturbed flow, that are unaffected (or down regulated) by laminar shear stress, suggests that attention should be given to additional transcription factors capable of binding to both VCAM-1 and connexin promoters and might play a role in transcriptional regulation by complex flow patterns.
CONCLUSIONS AND PERSPECTIVES The studies of endothelial gene regulation by uniform laminar shear stress have yielded new insights into how fluid mechanical forces generated in vivo by the flowing blood regulate endothelial gene transcription. In particular recent work from several laboratories focused our attention to several points: 1. Like biochemical stimuli, biomechanical forces affect the endothelium through the transcriptional regulation of endothelial genes. 2. This regulation is mediated by Shear Stress Response Elements (SSREs) located in the promoters of endothelial shear stress responsive genes.
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3. Transcription factors that are also regulated/activated by shear stress bind to these SSREs. The composition of the protein complex that binds to these SSREs determines the pattern of response to the force. 4. The definition of more than one SSRE together with the fact that more than one transcription factor is capable of binding to each SSRE, further demonstrate the complexity of transcriptional regulation by shear stress. 5. The development of new in vitro shear stress model systems which mimic more complex shear stresses, enables us to study the regulation of endothelial genes under conditions that are more relevant to the pathophysiology of the blood vessel. These studies further support in vivo observations, suggesting that a distinct set of genes is expressed under complex shear forces and is not affected (or down regulated) by uniform laminar shear stress. These exciting results in the field of gene regulation by hemodynamic forces raised additional questions that are waiting to be answered: 1. Does the regulation of a single gene by shear stress involve several SSREs and their bound transcription factors, via the formation of a higher order enhancer complex (see Figure 7.1), similar to the one formed in response to biochemical or viral stimuli? 2. Can unique activation pathways for certain transcription factors be determined in endothelial cells exposed to biomechanical forces? 3. Does the regulation of endothelial genes by complex shear forces is mediated by unique SSREs and novel transcription factors that bind to them? 4. How does a frictional force applied to the surface of an endothelial cell transmit its signals into the cell, signals that are translated into genetic regulatory events? 5. Does the tissue origin of the endothelium (blood vessels versus lymphatic vessels, arteries versus veins, large vessels versus capillaries) determine its response to shear stress, and the translation of this force into distinct genetic events? 6. Finally, how do endothelial cells integrate the various biomechanical and biochemical stimuli in their local environment? Future studies on the complexity of transcriptional regulation by uniform laminar shear stress, regulation of endothelial genes by complex shear forces and the combinatorial effect of shear stress and other hemodynamic forces, will hopefully help answering these questions. It is hoped that as the complexity of endothelial gene regulation by biomechanical forces is unraveled, we will arrive at a better understanding of some key physiological processes in the vascular system, as well as the pathogenesis of cardiovascular diseases. Resnick, N. wish to acknowledge the collaboration of C.F.Dewey Jr. And colleagues in the Fluid Mechanics Laboratory at MIT, as well as Tucker Collins, Levon Khachigian, William Atkinson, Tobi Nagel and Keith Anderson members of the Vascular Research Division at the Brigham and Women’s Hospital. Above all The author wish to thank Dr. MA Gimbrone Jr. for never ending support and insightful scientific discussions. Original research by the authors was supported by the Binational USA-Israel Science Foundation (BSF) (8123801) and the Israeli Ministry of Health (1000022).
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54. Lin, M.C., Almus-Jacob, F., Chen, H.H., Parry, G.C., McKman, N., Shyy, J.Y and Chien, S. (1997) Shear stress induction of the tissue factor genes. J. Clin. Invest., 99, 737–744. 55. Kawai, Y., Matsumoto, Y., Atanabe, K., Yamamoto, H., Satoh, K., Maruta, M., Handa, M. and Ikeda, Y. (1996) hemodynamic forces modulate the effect of cytokines on fibrinolytic activity of endothelial cells. Blood, 87, 2314–2321. 56. Malek, A.M., Greene, A.L. and Izumo, S. (1993) regulation of endothelin-1 gene by fluid shear stress is transcriptionally mediated and independent of protein kinase C and cAMP. Proc. Natl. Acad. Sci. USA, 90, 5999–6003. 57. Chun, T.H., Itoh, H., Masatsugu, K., Ogawa, Y., Kentaro, Y. and Inoue, M. (1997) Shear stress oppositely regulates the expression of two endothelial enzymes, prostacyclin synthase and endothelial converting enzyme. Circulation, 96, 1–349. 58. Nishida, K., Harrison, D.G., Naves, J.P., Fisher, A.A. and Docjerty, S.R. et al. (1992) Molecular cloning and characterization of constitutive bovine aortic endothelial cells nitric oxide synthase. J. Clin. Invest., 90, 2092–2096. 59. Reider, M.J., Carmona, R., Krieger, J.E., Protchard, K.A. and Greene, A.S. (1997) Suppression of angiotensin converting enzyme expression and activity by shear stress. Circ. Res., 80, 312–319. 60. Okahara, K., Kambayashi, J., Onishi, T., Fijiwara, Y., Kawasaki, T. and Monden, M (1995) Shear stress induces expression of CNP gene in human endothelial cells. FEBS Lett., 373, 108–110. 61. Hsieh, H.J., Li, N.Q. and Frangos, J.A. (1993) Pulsatile and steady flow induces c-fos expression in human endothelial cells. J. Cell. Physiol, 154, 143–151. 62. Khachigian, L.M., Anderson, K.R., Halnon, N.J, Gimbrone, M.A. Jr. Resnick, N. and Collins, T. (1997) Egr1 is activated in endothelial cells exposed to fluid shear stress and interacts with a novel shear stress response element in the PDGF-A chain promoter. Atreioscler. Thromb. Vasc. Biol., 17, 2280–2286. 63. Shyy, J.Y., Lin, M.C., Han, J., Lu, Y., Petrine, M. and Chien, S. (1995) the cis acting phorbol ester “12-o-tetradecanoylphorbol 13 acetate” responsive element is involved in shear stress induced MCP1 gene expression. Proc. Natl. Acad. Sci. USA, 92, 8069–8073. 64. Sterpetti, A.V., Cucina, A., Morena, A.R., Dinnas, D., Dangelo, L.S., Cavalero, A. and Stipa, S. (1993) Shear stress increases the release of IL-1 and IL-6 by aortic endotherlial cells. Surgery, 114, 911–914. 65. Ando, J., Tsuboi, H., Korenaga, R., Takahasi, K., Kosaki, K., Ishhiki, M., Tojo, T., Takada, Y. and Kamiya, A. (1996) Differential display and cloning of shear stress responsive mRNA in human endothelial cells. Biochem. Biophys. Res. Commun., 225, 347–351. 66. Inoue, N., Ramasamy, S., Fukai, T., Nerem, R.M. and Harrison, D.G. (1996) Shera stress modulates expression of Cu/Zn SOD in human aortic endothelial cells. Circ Res., 79, 32–37. 67. Kuelenaer, G.W., Chappel, D.C., Isizaka, N., Nerem, R.M., Alexander, W. and Griendling, K.K. (1997) Oscillatory and steady laminar shear stress differentially affect endothelial redox state. Circulation, 96, 1669. 68. Topper, J.N., Cai, J., Falb, D. and Gimbrone, M.A. Jr. (1996) Identification of vascular endothelial genes differentially responsive to fluid mechanical stimuli: Cox-2, MnSOD and ecNOS are selectively upregulated by steady laminar shear stress. Proc. Natl. Acad. Sci. USA, 93, 10417–10422. 69. Topper, J.N., Cai, J., Qui, Y., Anderson, K.R., Xu, Y.Y., Deed, J.D., Feeley, R. and Gimeno, C.J. et al. (1997) Vascular MAD: two novel MAD-related genes selectively inducible by flow in human vascular endothelial cells. Proc. Natl. Acad. Sci. USA, 94, 9314–9319. 70. Malek, A.M., Jackman, R.W., Rosenberg, R.D. and Izumo, S. (1994) Endothelial expression of thrombomodulin is reversibely regulated by fluid shear stress. Circ Res., 74, 852–860. 71. Takada, Y., Shinkai, F., Kondo, S., Yamamoto, S., Tsuboi, H., Korenaga, R. and Ando, J. (1994) Fuid shear stress increases the expression of thrombomodulin by cultured human endothelial cells. Biochem. Biophys. Res. Commun., 205, 1345–1352. 72. Varner, S.E., Chappel, D.C., Alexander, W.R., Medford, R.M. and Nerem, R.M. (1997) Endothelial VCAM-1 regulation by steady and oscillatory fluid shear stress. Atherosclerosis, 134, 288. 73. Korenaga, R., Ando, J., Kosaki, K., Ishhiki, M., Takada, Y. and Kamiya, A. (1997) Negative transcriptional regulation of the VCAM-1 gene by fluid shear stress in murine endothelial cells. Am. J. Physiol., C1506–1515. 74. Malek, A.M. and Izumo, S. (1995) Control of endothelial cells gene expression by flow. J. Biomechan., 28, 1515–1528. 75. Walpola, P.L., Gotlieb, A.I., Cybulsky M.I. and Lanigille B.L. (1995) Expression of ICAM-1 and VCAM-1 and monocyte adherence in arteries exposed to altered shear stress. Arterioscler. Thromb. Vasc. Biol., 15, 2–10. 76. Resnick, N., Sumpio, B.E., Du, W. and Gimbrone, M.A. Jr. (1995) Endothelial gene regulation by
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biomechanical forces. In: Atherosclerosis, edited by Woodfrod, F.P. Davignon, J. and Sniderman, A. pp. 838–843. 77. Khachigian, L.M., Resnick, N., Gimbrone, M.A. Jr. and Collins, T. (1995) Nuclear factor B interacts functionally with the PDGF-B chain shear stress response element in vascular endothelial cells exposed to fluid shear stress. J. Clin. Invest., 96, 1169–1175. 78. Resnick, N., Yahav, H., Khachigian, L.M., Collins, T., Andersn, K.R., Dewey, C.F. Jr. and Gimbrone, M.A. Jr. (1997) Endothelial gene regulation by laminar shear stress. In: Analytical and quantitative cardiology, edited by Sideman, S. and Bayer, R. pp. 155–164. 79. Sumpio, B.E., Chang, R., Xu, W.J. and Du, W. (1997) Regulation of tPA in endothelial cells exposed to cyclic strain: role of CRE, AP-2 and SSRE bindind sites. Am. J. Physiol, 273, C1441–1448. 80. Barnes, P.J. and Karin, M. (1997) Nuclear factor kB—A pivotal transcription factor in chronic inflammatory diseases. New Engl. J., 336, 1066–1071. 81. Baeuerle, P.A. and Baltimore, D. (1996) NFB: ten years after. Cell, 87, 13–20. 82. Verma, I.M. and Srevenson, J. (1997) IB kinases—beginning not the end. Proc. Natl. Acad. Sci. USA, 94, 11758–11760. 83. Collins, T., Read, M.A., Neish, A.S., Whitely, M.Z., Thanos, D. and Maniatis, T. (1995) Transcriptional regulation of endothelial cells adhesion molecules: NFB and cytokine- inducible enhancers. FASEB J., 9, 899–909. 84. Read, M.A., Neish, A.S., Gerritsen, M.E. and Collins, T. (1996) Postinduction transcriptional repression of E-selectin and VCAM-1 . J. Immunol, 157, 3472–3479. 85. Lindner, V. and Collins, T. (1996) Expression of NFB and IB alpha by aortic endothelium in an arterial injury model. Am. J. Pathol, 148, 427–438. 86. Page, S., Brandl, R., Neumeier, D. and Brand, K. (1997) Activation state of transcription factor NFB in acute aortic dissection. Atherosclerosis, 134, A282. 87. Lan, Q., Mercurius, K.Q. and Davies, P.F. (1994) Stimulation of transcription factors NFB and AP1 in endothelial cells subjected to shear stress. Biochem. Biophys. Res. Commun., 201, 950–956. 88. Mohan, S., Mohan, N. and Sprague, E.A. (1997) Differential activation of NFB in human aortic endothelial cells conditioned to specific flow environment. Am J. Physiol., 273, C5720578. 89. Zand, T., Majno, G., Nunneri, J.J., Huffman, A.M., Savilonis, B.J., MacWilliams, B. and Joris, I. (1991) Lipid deposition and intimal stress and strain. Am. J. Pathol., 139. 90. Krais, L.W., Geary, R.L., Mattsson, E.J., Vergels, A.V., Au, Y.P. and Clowes, A.W. (1996) Acute reduction in blood flow and shear stress induces PDGF-A expression in baboon prosthetic grafts. Circ. Res., 79, 45–53. 91. Bassiouny, H.S., Song, R.H., Kocharan, H., Vosicky, J., Glagov, S. and Weichselbaum, R.R. (1997) Egr-1 is upregulated by reduced flow in the injured rat carotid artery. Circulation, 96, 1548. 92. Nagel, T., Resnick, N., Dewey, C.F. Jr. and Gimbrone, M.A. Jr. (1999) Vascular Endothelial cells respond to spatial gradients in fluid shear stress by enhanced activation of transcription factors. Arterioscler. Thromb. Vasc. Biol., in press.
8 Shear Stress Mediated Gene Regulation Susan M.McCormick 1 and Larry V.McIntire2 Cox Laboratory for Biomedical Engineering, Rice University, P.O. Box 1892, Houston, TX 77251–1892, USA, 1E-mail: [email protected], 2E-mail: [email protected].
In this chapter the regulation of nine genes by shear stress and other regulatory factors is described in detail. The first four genes discussed are E-selectin, intercellular adhesion molecule1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1) and monocyte chemotactic protein1 (MCP-1). The proteins of these genes are involved in the immune system. The other five genes described, C-type natriuretic peptide, (CNP), endothelin-1 (ET-1), angiotensin converting enzyme (ACE), prostaglandin H synthase (PGHS)-1 and PGHS-2 all effect vascular tone and blood pressure. This chapter is written to allow comparisons to be made between how shear stress and other factors regulate gene expression. It demonstrates that shear stress has all the characteristics of biochemical regulatory factors and can be as significant as non-mechanical regulatory factors in controlling gene expression in endothelial cells. In order to facilitate this comparison, the regulation of the genes are described in detail including time points and magnitudes of changes in mRNA and protein levels where possible. KEYWORDS: Shear stress, gene regulation, inflammation, vascular tone.
INTRODUCTION It is now widely accepted that shear stress, a mechanical stimulus, regulates gene expression. There are a multitude of very good review papers covering this subject (Papadaki and Eskin, 1997, Resnick et al., 1995, Berk et al., 1995, Malek and Izumo, 1995, Malek and Izumo, 1994, Nollert et al., 1992). Signal pathways explaining how genes may be regulated by shear stress have been proposed in great detail and several shear stress regulatory elements have been identified. This chapter describes the effects of shear stress and other regulatory factors on gene expression in endothelial cells and how they compare with one another. This chapter also focuses attention on how mechanical and non-mechanical stimuli may interact with one another. To be able to intervene in the most intelligent way possible in biological systems we must understand how all parts of the system work in concert. We have chosen two general systems important in vascular biology as examples. The first is the immune/inflammation system in which the regulation of E-selectin, intercellular adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1) and monocyte chemotactic protein-1 (MCP-1) are detailed. The second system we present involves the regulation of vascular tone and blood pressure. We cover C-type natriuretic peptide, (CNP), endothelin-1 (ET-1), angiotensin converting enzyme (ACE), prostaglandin H synthase (PGHS)-1 and PGHS-2. For each gene there is a description of its biological function, the effect of 149
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shear stress on the expression of the gene and the effect of other stimuli on the gene’s expression. In many cases we have included time points and magnitudes of changes in gene expression to aide in the evaluation of the importance of the different stimuli and in how they may work together to regulate the different biological processes.
GENES REGULATED BY SHEAR STRESS IN THE IMMUNE SYSTEM AND INFLAMMATORY SYSTEM Biological Functions of E-selectin, ICAM-1 and VCAM-1 The initial interaction between leukocytes and endothelial cells under flowing conditions is normally mediated via the selectins expressed on them (Kostantopoulos and McIntire, 1996). The expression of P-selectin and E-selectin on the surface of endothelial cells is increased by inflammatory mediators such as interleukin-1 (IL-1) ß, histamine, and tumor necrosis factor- (TNF-). These two selectins recognize glycoproteins on leukocytes while L-selectin, which is constitutively expressed on the surface of leukocytes, recognizes glycoproteins on endothelial cells. E-selectin binds to neutrophils (Bevilacqua et al., 1989, 1987, Hession et al., 1990), monocytes (Hession et al., 1990) and lymphocytes (Picker et al., 1991, Shimizu et al., 1991). The interactions between the selectins and the glycoproteins can tether and slow down the leukocytes and cause rolling but not firm adhesion. Additional receptors, such as ICAM-1, ICAM2 and VCAM-1, which are expressed on the surface of endothelial cells, must also interact with the leukocyte integrins for firm adhesion. Once the leukocyte is attached to the endothelium it may transmigrate across the endothelium to the inflammatory site. Proteins such as platelet/endothelial cell adhesion molecule (PECAM-1) and chemotactic mediators 1-0-alkyl-2-acetyl-sn-glycero-3-phosphocholine (PAF), IL-8 and MCP-1 are involved in this step. Thus, there are many genes that must be expressed in order for leukocytes to reach sites of inflammation. Since this takes place in an environment that is constantly subjected to mechanical forces, it is reasonable to expect that these forces may be involved in the regulation of these genes.
Shear Stress Regulation of E-Selectin Gene Expression Several laboratories have observed that E-selectin gene expression does not change in endothelial cells exposed to shear stress in vitro. Using fluorescence immunobinding to measure the surface expression of E-selectin on human umbilical vein endothelial cells (HUVECs) (passage 2) that were cultured in the presence of shear stresses ranging from 2.5–46 dyn/cm2 for 4 to 48 hr in a cone and plate flow apparatus, Nagel et al. (1994) did not detect significant changes in expression. They also could not detect alterations in E-selectin mRNA levels in cells cultured in the presence of shear stress (10 dyn/cm2) for 2, 8 and 24 hr by Northern analysis. Morigi et al. (1995) found no changes in E-selectin expression when they cultured HUVECs in the presence of either laminar flow (8 dyn/cm2) or turbulent flow (8.6 dyn/cm2) with a cone and plate flow
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apparatus for 6 to 15 hr. These investigators assayed for E-selectin on the surface of the endothelial cells using indirect immunofluorescence with flow cytometry. Sampath et al. (1995) did detect a slight decrease in E-selectin mRNA levels in HUVECs cultured in the presence of shear stresses, 2 or 10 dyn/cm2 in a parallel plate flow chamber for 1 hr. At later time points no further changes in mRNA levels were detected.
Regulation of E-Selectin Gene Expression by Non-Mechanical Stimuli In contrast to the minimal effect shear stress has on E-selectin gene expression, IL-1, TNF- and lipopolysaccharide (LPS) cause dramatic increases in E-selectin mRNA levels and the expression of the protein on the surface of endothelial cells (Figure 8.1). E-selectin expression increases on the cell surface of HUVECs (passage 2–4) incubated with either IL-1, TNF- or LPS within 1 hr of treatment. The amount of E-selectin protein on the surface continues to increase over the next 3 to 6 hr to its maximum level. E-selectin protein levels then decrease to basal levels during the next 20 to 44 hr (Pober et al., 1986a, b, Bevilacqua et al., 1987, 1989). Interleukin-1 and TNF- induce a greater increase in E-selectin cell surface expression than LPS (Pober et al., 1986a, Bevilacqua et al., 1987). E-selectin protein and mRNA levels increase in HUVECs cultured in the presence of IL-1 at the same time E-selectin cell surface expression increases (Bevilacqua et al., 1989). Tumor necrosis factor- greatly increases E-selectin mRNA at 2 hr (Bevilacqua et al., 1989). Stimulation of E-selectin cell surface expression
Figure 8.1. The regulation of E-selectin gene expression in endothelial cells. Shear stress decreases E-selectin gene expression whereas, IL-1, TNF- and LPS increase expression. Interferon- enhances the effect of TNF- and IL-1 on E-selectin gene expression. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outward indicate that the factor decreases gene expression. Curved arrows indicate that the factor acts as an enhancer. The relative efficacy of the regulatory factors to increase or decrease E-selectin gene expression is represented by the sizes of the arrows.
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by IL-1, TNF- or LPS is inhibited by either actinomycin D or cyclohexamide, indicating that the increases in E-selectin cell surface expression caused by these agents is dependent upon mRNA and protein synthesis (Pober et al., 1986a, Bevilacqua et al., 1987). The qualitative pattern of E-selectin expression, with peak expression occurring 4–6 hr after stimulation and basal level expression returning in 24 hr is essentially independent of the stimulant’s concentration. Interferon- (IFN-) alone does not regulate E-selectin cell surface expression (Bevilacqua et al., 1987) or mRNA levels (Bevilacqua et al., 1989). However, IFN- does increase the effect of TNF- and IL-1ß on E-selectin surface expression in HUVECs at 4 hr (Doukas and Pober, 1990). In addition IFN- extends the time period during which E-selectin is expressed. There is a significant increase in the number of cells expressing the selectin at 24 hr in cell cultures incubated in medium containing IFN plus either IL-1ß or TNF-. Interferon- has a significantly greater effect on Eselectin expression in cells cultured in the presence of TNF- than in cells cultured in the presence of IL-1ß. Nearly 100% of HUVECs cultured in the presence of IL-1ß plus TNF- plus IFN- express E-selectin at 24 hr. It must be noted that none of these cytokine combinations increase the maximum number of E-selectin receptors on each cell over the number present on cells cultured in the presence of either IL-1ß or TNF. The interactions between IL-1ß and IFN- are dependent upon the presence of serum. Interferon- only increases E-selectin mRNA in the presence of TNF- at 6 hr. At 1 and 24 hr there is no difference in the level of E-selectin mRNA in cells cultured in the presence of TNF- or TNF- plus IFN-. The increase in E-selectin mRNA by IL-1ß is not enhanced by IFN- at 1, 6 or 24 hr. E-selectin expression is similarly regulated by the same factors in vivo. An intradermal injection of LPS into a baboon increases E-selectin expression within 2 hr (Brisoce et al., 1992). E-selectin expression is maximal at 2 hr, then begins to decrease at 9 hr but remains elevated compared to control values for at least 48 hr. The injection of TNF- also increases E-selectin expression in 2 hr but with this cytokine E-selectin expression is sustained at a higher level and for a longer period of time than when LPS is injected.
Regulation of ICAM-1 Gene Expression by Shear Stress Intercellular adhesion molecule-1, a member of the immunoglobin gene superfamily is expressed in hematopoietic cells; macrophages, leukocytes, dendritic cells and in non-hematopoietic cells; vascular endothelial cells, epithelial cells and fibroblasts (Dustin et al., 1986). It is interesting to note that the molecular weight of ICAM-1 isolated from different cell types is heterogeneous, indicating that glycosylation of the protein is differentially regulated in different cells. The regulation of ICAM-1 expression in vitro by shear stress seems to be affected by the methods used to culture and shear the cells as well as by the shear stress. Ando et al. (1994) saw no change in ICAM-1 cell surface expression in mouse lymph endothelial cells exposed to a shear stress of 1.5 dyn/cm2 for 6 hr. The same group reported that ICAM-1 cell surface expression increases 2.5 fold in HUVECs (passage 2–10) cultured in a parallel plate flow chamber in the presence of a shear stress of 15 dyn/cm2 for 4hr as measured by flow cytometry
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(Tsuboi et al., 1995). An increase in cell surface expression is first detected at 1 hr, is maximal at 8 hr and is maintained for at least 24 hr. Intercellular adhesion molecule1 expression is dependent upon the magnitude of the shear stress, increasing as the magnitude of the shear stress increases. Reverse transcription-polymerase chain reaction (RT-PCR) analysis showed that ICAM-1 mRNA levels increase after 1 hr of 15 dyn/ cm2 shear stress, and is maximal at 8 hr with a 8.3 fold increase and remains elevated at 24 hr. No change in ICAM-1 surface expression occurs on HUVECs (passage 1–5) seeded on polystyrene coated with 0.2% gelatin and exposed to a laminar flow with a shear stress of 8 dyn/cm2 for 3 hr but, after 6 and 15 hr there is a significant increase of 151 % and 145% respectively (Morigi et al., 1995). In contrast, if the flow is turbulent there is no change in ICAM-1 levels. Similarly Nagel et al. (1994) reported that intercellular adhesion molecule-1 surface expression on HUVECs (passage 2) seeded on polystyrene coated with 0.1 % gelatin exposed to a shear stress of 10 dyn/ cm2 using a cone and plate flow apparatus increases significantly after 8 hr. They observed a continual increase in ICAM-1 surface expression with time for at least 48 hr with cell surface expression increasing approximately 3 fold at 48 hr. Intercellular adhesion molecule-1 mRNA levels increase in cells exposed to a shear stress of 10 dyn/cm2 as early as 2 hr and are still elevated after 24 hr of exposure (Nagel et al., 1994). There is up to a 2.5 fold increase in ICAM-1 surface expression in HUVECs exposed to shear stresses ranging from 2.5 to 46 dyn/cm2 for 24 hr. However, the magnitude of the increase is independent of the magnitude of the shear stress. The expression of ICAM-1 in primary HUVECs seeded on glass treated with NaOH exposed to shear stress, 25 dyn/cm2 in a parallel plate flow chamber is biphasic (Sampath et al., 1995). Intercellular adhesion molecule-1 expression on the cell surface increases after 12 hr of flow and then decreases slightly below static levels at 24 hr. If the cells are exposed to shear stress in the presence of LPS there is no change in ICAM-1 cell surface expression compared to cells incubated in the presence of LPS with no shear stress. However, if the cells are pretreated with LPS for 18 hr and then exposed to shear stress in the absence of LPS or if the cells are subjected to shear stress for 24 hr in the absence of LPS and then cultured in the absence of shear stress but in the presence of LPS there is a significant increase in ICAM-1 cell surface expression compared to either 24 hr of shear stress in the presence of LPS or LPS in the absence of shear stress. There is an increase in ICAM-1 mRNA at 1 hr in cells cultured in the presence of shear stresses of 2, 10 and 25 dyn/cm2 whereas, at 6hr there is a slight decrease in ICAM-1 mRNA levels in comparison to ICAM-1 mRNA levels in cells cultured in the absence of shear stress. Regulation of ICAM-1 by shear stress has been suggested to involve reactive oxygen species (Chiu et al., 1997). An increase in reactive oxygen species occurs in HUVECs exposed to shear stress (20 dyn/cm2) as early as 15 min and peaks at 30 min after which it slowly decays over the next 5 hours but remains elevated compared to levels in cells cultured in the absence of shear stress. If the shear stress is removed reactive oxygen species return to their basal levels within 5 min. The increase in reactive oxygen species at 30 min is equivalent to the increase in cells cultured in the presence of phorbol 12-myristate 13 acetate (PMA). Antioxidant N-acetyl-cysteine (NAC) and the antioxidant enzyme catalase completely inhibit shear stress induced increases in ICAM-1 mRNA levels at 3 and 6 hr. Flow cytometry analysis showed
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that both NAC and catalase also inhibit the cell surface expression of ICAM-1 at 12 and 24 hr. Altered shear stress in vivo effects the expression of ICAM-1 too (Walpola et al., 1995). Intercellular adhesion molecule-1 expression increases in rabbit carotid arteries when the shear stress is increased by 170% from 11.3 to 30.5 dyn/cm2 for 5 days in comparison to expression levels in sham-operated carotid artery as detected by immunostaining. When shear stress is decreased by 73% from 12.1 to 3.26 dyn/ cm2 ICAM-1 expression decreases.
Regulation of ICAM-1 Gene Expression by Non-Mechanical Stimuli Proinflammatory cytokines IL-1, TNF- and IFN- regulate the cell surface expression of ICAM-1 (Figure 8.2, Dustin et al., 1986, Pober et al., 1986b). Interleukin-1 treatment of HUVECs (passages 3–8) increases ICAM-1 cell surface expression 2 and 3 fold at 6 and 24 hr, respectively. Similarly, TNF- increases ICAM-1 expression 4 and 6 fold at 6 and 24 hr of stimulation. Intercellular adhesion molecule-1 expression continues to increase for at least 72 hr, with TNF- causing the greatest change in expression. Interferon- increases ICAM-1 expression in a manner similar to IL-1 and TNF-a, except that changes in expression occur later and the induction is smaller (Pober et al., 1986b). Interferon- may be more of an enhancer than an inducer of ICAM-1 gene expression. Tumor necrosis factor- and IFN- both increase ICAM-1 cell surface expression on HUVECs (passage 4–8) although TNF- is much more effective than IFN- which creates only a minimal induction in ICAM-1 expression. Intercellular adhesion molecule-1 mRNA levels do not increase in cells incubated in the presence of IFN- until 24 hr. However, when IFN- is added with TNF- to the medium there is a synergistic increase in ICAM-1 expression at 24 hr but only an additive increase at 4 hr (Doukas and Pober, 1990). There is also a synergistic increase in the expression of ICAM-1 on the surface of human saphenous vein endothelial cells and human umbilical artery cells when the cells are cultured in the presence of IFN- plus TNF- (Paleolog et al., 1992). Interferon- has an additive effect with IL-1ß at 6 and 24 hr and neither cytokine effects ICAM-1 expression at 1 hr (Doukas and Pober, 1990). The expression of ICAM-1 mRNA by TNF- is greatly enhanced by IFN- at 6 and 24 hr but not at 1 hr. Interferon- has a slight effect on ICAM-1 mRNA levels in the presence of IL-1ß at 1 and 6 hr but none at 24 hr. In human synovial micro vascular endothelial (HSE) cells, IL-1 and TNF- after 24 hr have relatively small effects on ICAM-1 cell surface expression in comparison to their effect on expression in HUVECs (Gerritsen et al., 1993). Tumor necrosis factor- increases ICAM-1 expression 3.5 fold in HSE cells compared to 12.7 in HUVECs at 24 hr. Interferon- synergistically increases the effect of IL-1 and TNF at both the protein and mRNA levels in HSE but does so less effectively than it does in HUVECs. When TNF- is present in the medium the addition of IFN- increases ICAM-1 surface expression 25.2 fold on HUVECs compared to 10.4 on micro vascular endothelial cells. Intercellular adhesion molecule-1 surface expression increases on micro vascular cells cultured in the presence of either TNF- or IFN- at 4 hr and is maximal at 8 hr. Whereas, the maximum response in cells cultured in the presence of
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Figure 8.2. The regulation of ICAM-1 gene expression in endothelial cells. Intercellular adhesion molecule-1 gene expression is regulated by cytokines, shear stress and LPS in endothelial cells. Cytokines IFN- and IL-1 increase ICAM-1 gene expression more in the endothelial cells of large vessels panel A than in the endothelial cells of microvessels, panel B. In contrast, IFN- increases ICAM-1 gene expression more in microvascular cells. Arrows pointing inward indicate that the factor increases gene expression while curved arrows indicate that the factor acts as an enhancer. The relative efficacy of the regulatory factors to increase ICAM-1 gene expression is represented by the sizes of the arrows.
TNF-a plus IFN- occurs at 16 hr in HSE cells. Both IL-4 and LPS do not effect ICAM-1 mRNA levels in micro vascular cells but they do enhance IFN- induced increases in ICAM-1 cell surface expression. Interferon- inhibits the increase in ICAM1 cell surface expression induced by PMA. Cytokines also increase ICAM-1 mRNA levels in micro vascular endothelial cells. In HSE cells IFN- increases ICAM-1 mRNA levels more than TNF-, which increases ICAM-1 mRNA levels minimally. There is a synergistic increase in mRNA levels in HSE cells cultured in presence of TNF- and
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IFN-. The time course of ICAM-1 mRNA expression is approximately the same in the two different cell types. In addition to being regulated by cytokines ICAM-1 expression is also regulated by protein kinase C activators (Lane et al., 1989). Both PMA and Mezerein increase the surface expression of ICAM-1 on HUVECs (passage 2–4). The amount of protein on the cell surface peaks at 6–8 hr and then decreases over the next 16 to 18 hr. Protein kinase C (PKC) inhibitors block the stimulation of ICAM-1 expression by both PMA and Mezerein. Although both PMA and Mezerein stimulate ICAM-1 expression they are not as potent as IL-1. It is interesting to note that while IL-1 stimulation is maintained for at least 24 hr, both PMA and Mezerein stimulation decreases after 8 hr. Cell surface expression of ICAM-1 on keratinocytes is regulated by the phorbol ester 12-O tetradecanoylphorbol-13-acetate (TPA) and by IFN- (Griffiths et al., 1990). Similar to the stimulation of ICAM-1 in fibroblasts PKC inhibitors prevent stimulation of ICAM-1 by TPA but not by the cytokine IFN-. Intercellular adhesion molecule-1 cell surface expression is greater on cells cultured in the presence of IFN- than on the surface of cells cultured in the presence of TPA as determined by immunohistochemical staining. In contrast to the stimulation of ICAM-1 cell surface expression on HUVECs by phorbol esters, ICAM-1 cell surface expression on keratinocytes cultured in the presence of phorbol esters does not reach maximum level until day one and it remains at that level for 3 days. T-cells which express lymphocyte-function-associated antigen1 (LFA-1) bind to keratinocytes expressing ICAM-1. This interaction between the two cells is thought to be involved in many inflammatory skin diseases. Hydrogen peroxide, a phagocyte derived reactive oxygen radical increases ICAM1 cell surface expression in HUVECs in a dose and time dependent manner (Lo et al., 1993). In cells cultured in the presence of hydrogen peroxide ICAM-1 cell surface expression increases at 15 min increases 2 and 4 fold at 0.5 and 4 hr respectively, in comparison to initial ICAM-1 cell surface expression. Intercellular adhesion molecules1 mRNA is detectable at 30min in cells cultured in the presence of hydrogen peroxide. Maximum ICAM-1 mRNA levels are reached between 0.5–1 hr and at 2hr there is a slight decrease in ICAM-1 mRNA levels.
Regulation of VCAM-1 Gene Expression by Shear Stress Vascular cell adhesion molecule-1, a second member of the immunoglobin gene superfamily, is mainly expressed in the endothelium of post capillary venules and veins. The mature VCAM-1 transcripts predominantly consists of 7 immunoglobin (Ig) domains although in humans VCAM-1 mRNAs consisting of only 6 domains are sometimes present in low number. In addition, in rabbit there are mRNA species with 7 and 8 Ig domains and in mouse there are mRNA isoforms consisting of 7 and 3 Ig domains. Vascular cell adhesion molecule-1 cell surface expression on mouse lymph node endothelial cells cultured in a parallel plate flow chamber in the presence of shear stress of 1.5 dyn/cm2 decreases by 48% at 6 hr in comparison to VCAM-1 surface expression levels on cells cultured in the absence of shear stress (Ando et al., 1994). In cells cultured in the presence of shear stress, 1.5 dyn/cm2, VCAM-1 mRNA
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levels decrease at 1 hr. At 24 hr mRNA levels decrease 75% in comparison to VCAM1 mRNA levels in cells cultured in the absence of shear stress. Vascular cell adhesion molecule-1 mRNA levels also decrease in primary HUVECs cultured in the presence of shear stress in a parallel plate flow chamber (Sampath et al., 1995). In cells cultured in the presence of shear stress (25, 10, 2 dyn/cm2) VCAM-1 mRNA levels decrease at 1 hr and continue to decrease at 6 hr (Figure 8.3). In contrast, Nagel et al. (1994) observed no changes in VCAM-1 cell surface expression on HUVECs cultured in the presence of shear stresses ranging from 2.5–46 dyn/cm2 for 4–48 hr in a cone and plate flow apparatus. Tsuboi et al. (1995) also observed no changes in VCAM-1 surface expression on HUVECs (passage 2–10) cultured in the presence of shear stress, 15 dyn/cm2 in a parallel plate flow chamber at 4 hr. In rabbit carotid arteries where the shear stress had been increased by 170% or decreased by 73% for 5 days by surgical intervention VCAM-1 cell surface expression increases as detected by immunostaining (Walpola et al., 1995). Regulation of VCAM-1 Gene Expression by Non-Mechanical Stimuli In addition to being regulated by shear stress, VCAM-1 gene expression is also regulated by cytokines (Figure 8.3). Vascular cell adhesion molecule-1 cell surface expression surface expression on HUVECs incubated in the presence of TNF- increases at 2 hr, is maximal at 6–8 hr with a 13.6 fold increase and then remains
Figure 8.3. The regulation of VCAM-1 gene expression in endothelial cells. Cytokines IL-1 and TNF- increase VCAM-1 gene expression whereas, shear stress decreases its expression. Nitric oxide inhibits the ability of cytokines to increase VCAM-1 gene expression. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outward indicate that the factor decreases gene expression. Curved arrows indicate that the factor acts as an enhancer. Solid arrows indicate that the factor inhibits the induction of VCAM-1 gene expression by another agent. The relative efficacy of the regulatory factors to increase or decrease VCAM-1 gene expression is represented by the sizes of the arrows.
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constant for at least 48 hr in comparison to initial VCAM-1 cell surface expression levels (Rice and Bevilacqua, 1989). In HUVECs (passage 3–5) cultured in the presence of TNF- VCAM-1 mRNA levels dramatically increase at 2 hr and remain elevated for at least 72 hr (Osborn et al., 1989). Nuclear run-off experiments showed that increases in VCAM-1 mRNA levels in HUVECs and BAECs cultured in the presence of TNF- are due at least in part to increases in the rate of VCAM1 gene transcription (Neish et al., 1992). Interferon- does not significantly increases VCAM-1 expression on the surface of HUVECs, human umbilical artery endothelial cells or human saphenous vein endothelial cells at 24 hr (Paleolog et al., 1992). However, there is a synergistic increase in VCAM-1 cell surface expression in cells cultured in the presence of IFN- plus TNF-. Vascular cell adhesion molecule-1 cell surface expression increases 5.75 fold in cells incubated in the presence of TNF- (10 ng/ml) and 8.25 fold in cells incubated in the presence of TNF-a plus IFN- at 24 hr (5 U/ ml). Vascular cell adhesion molecule-1 cell surface expression on endothelial cells also increases synergistically in cells cultured in the presence of IL-4 plus TNF-. However, VCAM-1 cell surface expression is lower on cells cultured in the presence of TNF-a plus IL-4 than on cells cultured in the presence of TNF- plus IFN-. Interleukin-4, 100 U/ml increases VCAM-1 cell surface expression on the surface of endothelial cells after 72 hr and then only very mildly in comparison to the increase in VCAM-1 cell surface expression caused by TNF- (Iademarco et al., 1995). Neither TNF- at a concentration of 0.2 ng/ml nor IL-4 at a concentration of 100 U/ ml significantly induce VCAM-1 cell surface expression. However, there is a synergistic increase in VCAM-1 expression on HUVECs cultured in presence of TNF- plus IL4 at these concentrations at 16 hr which is maintained for at least 72 hr. Tumor necrosis factor- significantly increases VCAM-1 mRNA levels in a dose and time dependent manner. In cells cultured in the presence of TNF- VCAM-1 mRNA levels are maximum at 8 hr and then decrease. Vascular cell adhesion molecule-1 mRNA levels increase less in cells cultured in the presence of IL-4 than in the presence of TNF-a. In cells cultured in the presence of IL-4 plus TNF- VCAM-1 mRNA levels increase synergistically and continue to increase for at least 48 hr. The synergistic increase in VCAM-1 mRNA levels is at least in part due to the increase in the half-life of VCAM-1 mRNA in the presence of IL-4. The half-life of VCAM-1 mRNA is 5 hr in cells cultured in the presence of TNF- whereas, it is greater than 10 hr in cells cultured in the presence of TNF- plus IL-4. Vascular cell adhesion molecule-1 expression increases in the dermal microvessels of baboons injected intradermally with cytokines or LPS (Briscoe et al., 1992). In baboons injected with LPS VCAM-1 cell surface expression increases at 6 hr, is maximal at 9–12 hr, decreases at 24 hr and at 48 hr returns to baseline values as detected by immunocytochemistry. In contrast, in baboons injected with TNF-, VCAM-1 cell surface expression is detected at 6 hr, peaks at 9 hr and remains elevated for at least another 39 hr. Vascular cell adhesion molecule-1 cell surface expression is greater in the microvessels of baboons injected with TNF- than in the microvessels of baboons injected with LPS. VCAM-1 cell surface expression does not change in baboons injected with IL-4, but VCAM-1 cell surface expression increases synergisticly in the microvessels of baboons injected with TNF- plus IL-4. There is
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no increase in VCAM-1 cell surface expression in baboons injected with IL-4 plus LPS in comparison to VCAM-1 cell surface expression in baboons injected with LPS. Vascular cell adhesion molecule-1 is constitutively expressed at a significantly higher level in the synovial lining of rheumatoid arthritis and osteoarthritis patients than in normal subjects (Morales-Ducret et al., 1992). Increased levels of VCAM-1 are also present on the surface of the blood vessels and interstitial mononuclear cells of patients with arthritis. Nitric oxide inhibits the induction of VCAM-1 expression by cytokines in human saphenous vein endothelial cells (De Caterina et al., 1995). Nitric oxide donors sodium nitroprusside (SNP), 3-morpholino sydnonimine (SIN-1) and S-nitroso-glutathione (GSNO) inhibit IL-1a stimulation of VCAM-1 cell surface expression as measured by enzyme immunoassay and flow cytometry. S-nitroso-glutathione inhibits IL-1a induced expression 35–55% at 24 hr decreasing both the number of cells expressing VCAM1 and the density of VCAM-1 on each cell, but has no effect on VCAM-1 cell surface expression in unstimulated cells. Similarly GSNO inhibits the induction of VCAM-1 expression by IL-1ß, IL-4, TNF- and LPS. Cyclic GMP analogues have no effect on VCAM-1 expression. S-nitroso-glutathione decreases VCAM-1 mRNA levels in IL-1 stimulated cells but it does not change the half-life of VCAM-1 mRNA in the cells. The transcription rate of VCAM-1 in cells cultured in the presence of TNF- plus GSNO is less than the transcription rate of VCAM-1 in cells cultured in the presence of TNF-a but in the absence of GSNO. Nitric oxide donor diethylamine (DETA)-NO also inhibits TNF-a induced VCAM-1 expression by 65% on the surface of HUVECs and 52 on the surface of human dermal microvascular endothelial cells at 9 hr (Khan et al., 1996). It has a similar effect on VCAM-1 mRNA levels. NG-monomethyl-Larginine (L-NMMA) an inhibitor of NO synthesis increases VCAM-1 cell surface expression in cells cultured in the presence of TNF-. Monocyte Chemotactic Protein-1 Biological functions of monocyte chemotactic protein-1 Shear stress regulates the expression of cytokine MCP-1 in addition to regulating the expression of vascular adhesion molecules. Monocyte chemotactic protein-1 is also known as monocyte chemotactic activating factor (MCAF), tumor derived chemotactic factor (TDCF) and smooth muscle cell chemotactic factor (SMC-CF). In addition it is the human homologue to the mouse JE gene. It is expressed in a variety of cell types including fibroblasts, vascular endothelial cells, arterial smooth muscle cells, monocytes and cancer cell lines. Monocyte chemotactic protein-1 is a potent chemoattractant for monocytes in vitro (Valente et al., 1988, Matsushima et al., 1989, Yoshimura et al., 1989) and in vivo (Zachariae et al., 1990). Monocytes have approximately 1,600 to 1,700 high affinity binding sites for MCP-1 (Yoshimura and Leonard, 1990, Valente et al., 1991). In addition to facilitating the migration of monocytes across the endothelium to sites of inflammation, MCP-1 may also play a role in the formation of atherosclerosis. The adherence of monocytes to the endothelial cells lining the vascular system and their migration across the endothelium into the intima of the vessels are the first noticeable events in atherogenesis (Faggiotto et al., 1984, Gerrity et al., 1979).
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Monocyte chemotactic protein-1 has been localized to macrophage rich regions of rabbit and human atherosclerotic lesions by in situ hybridization and immunostaining (Yla-Herttuala et al., 1991). Monocyte chemotactic protein-1 may also be important in the prevention of tumors. Monocytes activated by MCP-1 inhibit the growth of tumor cells in vitro (Matsushim et al., 1989) and in vivo (Rollins and Sunday, 1991). Tumor forming cells, HeLa or Chinese hamster ovary (CHO) cells, do not form tumors in nude mice when coinjected with cells expressing high levels of MCP-1. In addition, histological examinations of tissues at sites where MCP-1 expressing cells or nonMCP-1 expressing cells were injected show that there are more monocytes at sites where MCP-1 expressing cells were injected than at sites where non-MCP-1 expressing cells were injected (Rollins and Sunday, 1991). Monocytes cytostatic activity against tumor cells may at least in part be due to the release of N-acetyl ß-D-glucosaminiase and superoxide by the cells upon their activation by MCP-1 (Zachariae et al., 1990). Regulation of monocyte chemotactic protein-1 gene expression by shear stress Shear stress has a biphasic effect on MCP-1 gene expression in human endothelial cells in vitro (Shyy et al., 1994, Figure 8.4). Northern analysis shows that MCP-1 message level increases quickly in HUVECs (<passage 4) cultured on a glass slide subjected to a shear stress of 16 dyn/cm2 in a parallel plate flow chamber. Monocyte chemotactic protein-1 mRNA levels peak after 1.5 hr increasing 2.5 fold compared to
Figure 8.4. The regulation of MCP-1 gene expression in endothelial cells. In addition to shear stress MCP-1 gene expression is also regulated by cytokines, lipids, growth factors and nitric oxide. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outward indicate that the factor decreases gene expression. Solid arrows indicate that the factor inhibits the induction of MCP-1 gene expression by another agent. The relative efficacy of the regulatory factors to increase or decrease MCP-1 gene expression is represented by the sizes of the arrows.
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the MCP-1 mRNA level in cells cultured in the absence of shear stress. Transcript levels return to constitutive level within 4 hr and after 5 hr MCP-1 mRNA can no longer be detected. It takes between 1.5 and 24 hr before the cells once again express MCP-1 message and are susceptible to regulation by shear stress. Nuclear run off experiments showed that shear stress increases MCP-1 mRNA at least in part by increasing the rate of MCP-1 gene transcription. Treatment of cells with cyclohexamide does not prevent MCP-1 mRNA levels from increasing in cells cultured in the presence of shear stress indicating that protein synthesis is not required for the transcription rate of the MCP-1 gene to increase. Both the genes quick response to shear stress and its lack of need for protein synthesis are features of immediately early response genes. It is also of interest that the regulation of the MCP-1 gene by shear stress is not cell type specific. Shear stress regulates MCP-1 gene expression in HeLa cells, glioma cells and fibroblasts in a manner similar to its regulation of MCP-1 in HUVECs. Analysis of the MCP-1 promoter has shown that the sequence GGTCTC, which is the closest sequence in the promoter to the reported shear stress regulatory element sequence, GAGACC, is not involved in the regulation of MCP-1 by shear stress (Shyy et al., 1995). Instead one of two divergent TPA shear stress elements that are present in the promoter is required for the shear response. Additional data indicate that shear stress and TPA share a common signal transduction pathway. Regulation of monocyte chemotactic protein-1 gene expression by nonmechanical stimuli Monocyte chemotactic protein-1 gene expression is regulated by many different agents in addition to shear stress (Figure 8.4). Oxidized low density lipoprotein (LDL), which is present in atherosclerotic lesions (Morton et al., 1986, Palinski et al., 1989) is thought to induce the formation of fatty streaks in arterial vessels by increasing the amount of MCP-1 and colony stimulating factors (CSF) secreted from endothelial cells (Liao et al., 1991). Minimally modified LDL (MM-LDL, LDL oxidized by storing it at 4°C for 3–6 months) is recognized by the LDL receptor on endothelial cells and increases the chemotactic activity of rabbit aortic endothelial cells (Berliner et al., 1990), human aortic endothelial cells and smooth muscle cells (Cushing et al., 1990). The adherence of monocytes to endothelial cells pretreated with MM-LDL for 2 hr increases significantly in comparison to the adherence of monocytes to endothelial cells not pretreated with MM-LDL. The adherence of monocytes to endothelial cells pretreated with MM-LDL for 24 hr increases 3 fold in comparison to the adherence of monocytes to cells not pretreated with MM-LDL. This increase in monocyte adherence is maintained for at least 48 hr. Cyclohexamide inhibits monocyte adherence indicating the need for protein synthesis prior to the binding of monocytes to endothelial cells (Berliner et al., 1990). Minimally modified-LDL increases the chemotactic activity of human aortic endothelial cells and SMCs (passage 6–9) by increasing MCP-1 gene expression (Cushing et al., 1990). Endothelial cells cultured with MM-LDL secrete approximately 7 to 10 fold more MCP-1 than cells cultured in the absence of MMLDL as measured by immunoprecipitation. The chemotactic activity of medium conditioned by endothelial cells incubated with MM-LDL is greater than the
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chemotactic activity of medium conditioned by endothelial cells which have not been pretreated with MM-LDL. The increase in MCP-1 mRNA levels is dose dependent for both SMCs and endothelial cells with mRNA levels increasing as the concentration of MM-LDL increases until it reaches 20–30 µg/ml. The increase in MCP-1 mRNA levels in smooth muscle cells cultured in the presence of MMLDL is transient. In SMCs cultured in the presence of MM-LDL MCP-1 mRNA levels increase at 1 hr and peaks at 4 hr increasing approximately 18 fold. Monocyte chemotactic protein-1 mRNA levels decrease at 8 and 20 hr. In contrast, MCP-1 mRNA levels in endothelial cells cultured in the presence of MM-LDL increase for 20 hr. In endothelial cells cultured in the presence of MM-LDL MCP1 mRNA levels increase at 1 hr and peak at 4 hr increasing 15 fold. Monocyte chemotactic protein-1 mRNA levels decrease slightly at 8 hr then at 20 hr mRNA levels increase approximately 41 fold in comparison to initial MCP-1 mRNA levels (Cushing et al., 1990). Minimally modified-LDL increases the expression of MCP1 in vivo. The injection of MM-LDL into mice increases MCP-1 mRNA levels in the liver, heart, lungs, spleen and kidneys as detected by Northern analysis (Liao et al., 1991). Navab et al. (1991) studied the effect of LDL on MCP-1 gene expression in SMCendothelial cell coculture. Medium conditioned by incubating media plus LDL with a coculture of SMCs and endothelial cells for 24 hr has a positive effect on the adherence of monocytes to endothelial cells. Monocytes adhere to endothelial cells cultured with media conditioned by SMC-endothelial cell cocultures in the presence of LDL 2.8 fold more than they do to cells incubated with medium conditioned by SMCendothelial cell cocultures in the absence of LDL. Coculture with LDL pre-conditioned medium also causes 7.1 fold more monocytes to transmigrate across the endothelium layer of cocultures than does medium preconditioned by cocultures in the absence of LDL. An MCP-1 antibody inhibits the transmigration of monocytes across the endothelial cells of the cocultures. Monocyte chemotactic protein-1 mRNA levels increase 7.2 fold in cocultures incubated with LDL for 24 hr compared to the sum of MCP-1 mRNA present in an equal number of endothelial and smooth muscle cells cultured separately with LDL. Monocyte chemotactic protein-1 protein levels increase 2.5 fold in cocultured cells in comparison to MCP-1 protein levels in either endothelial cells or SMCs cultured individually for 36 hr in the presence of LDL. Monocyte migration increases 5.8 fold in cocultures incubated with LDL for 48 hr compared to cocultures incubated in the absence of LDL. This effect of LDL on monocyte migration can be inhibited 91% by the addition of HDL to the media. Antioxidants also inhibit the induction of monocyte migration by LDL if they are added at the same time as LDL. If medium preconditioned in the presence of LDL is added to a coculture simultaneously with HDL or antioxidants the increase in monocyte migration is not inhibited. In addition to LDL, cytokines and endotoxin also regulate MCP-1 gene expression. Streiter et al. (1989) analyzed the effect of TNF-, IL-1ß and LPS on MCP-1 mRNA levels in HUVECs (passage 5) and human foreskin fibroblasts (passages 4–8) by Northern analysis (Figure 8.4). Monocyte chemotactic protein-1 mRNA levels are 7 fold greater in endothelial cells cultured in the presence of LPS at 1 hr than MCP-1 mRNA levels in cells cultured in the absence of LPS. At 8hr MCP-1 mRNA levels
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increase 11 fold after which they remain constant for at least the next 16 hours. Lipopolysaccharide has no effect on MCP-1 gene expression in human foreskin fibroblasts. Tumor necrosis factor-a and IL-1ß increase MCP-1 mRNA levels in both cell types in a dose and time dependent manner. At 8 hr MCP-1 mRNA levels increase 33 fold in fibroblasts cultured in the presence of TNF-a in comparison to initial MCP1 mRNA levels. In comparison, MCP-1 mRNA levels increase only 19 fold in HUVECs cultured in the presence of TNF- at 8 hr in comparison to initial MCP-1 mRNA levels. In HUVECs cultured in the presence of IL-1ß MCP-1 mRNA levels increase 5 fold in comparison to initial MCP-1 mRNA levels at 30 min (Figure 8.2). At 4hr MCP-1 mRNA levels increase 14 fold after which they remain constant until at least 24 hr. The effect of IL-1ß on MCP-1 mRNA levels is due at least in part to an increase in the transcription rate of the MCP-1 gene (Sica et al., 1990). Interleukin-1ß regulates MCP-1 mRNA expression in fibroblasts the same way it does in HUVECs. In addition to TNF-a and IL-1ß, IFN- also increases MCP-1 mRNA levels. However, the increase in mRNA levels due to IFN- is small in comparison to the increase induced by IL-1ß and TNF-a (Rollins et al., 1990). Rollins and Pober (1991) treated HUVECs with IL-4, a cytokine which can activate macrophages to prevent tumor growth (Tepper et al., 1989). In cells cultured in the presence of IL-4 MCP-1 mRNA levels increase in a dose and time dependent manner. Monocyte chemotactic protein-1 mRNA levels increase at 24 hr in cells cultured in the presence of IL-4 and at 72 hr MCP-1 mRNA levels in the cells are equivalent to MCP-1 mRNA levels in endothelial cells cultured in the presence of IL-1ß for 24 hr. Interleukin-4 does not effect MCP-1 mRNA levels in HUVECs cultured in the presence of either IL-1 or TNF-. In contrast to the increase in MCP-1 levels in the medium of HUVECs cultured in the presence of IL1 which occur within 24 hr, increases in MCP-1 protein levels in the medium of HUVECs cultured in the presence of IL-4 occur after 24 hr. However, at 72 hr MCP-1 protein levels in the medium of HUVECs cultured in the presence of IL-4 are equivalent to those in the medium of cells cultured in the presence of IL-1. Monocyte chemotactic protein-1 protein levels increase synergistically in cells cultured in the presence of IL-1 plus IL-4 at 72 hr. Shyy et al. (1993) cultured HUVECs (<passage 7) in the presence of human monocyte colony-stimulating factor (MCSF) which is a growth factor that stimulates the differentiation and proliferation of monocytic progenitor cells. Monocyte colonystimulating factor increases MCP-1 mRNA levels in a dose and time dependent manner. In cells cultured in the presence of MCSF, MCP-1 mRNA levels increase at 1 hr, peaks at 4 hr and remains elevated for at least 8 hr. Immuno-flourescence staining of HUVECs demonstrated that MCP-1 protein levels are also increased by MCSF. Nitric oxide regulates MCP-1 expression in vitro (Zeiher et al., 1995). In freshly isolated HUVECs incubated with complete media, MCP-1 mRNA levels increase at 3 hr and decrease at 6 hr. If NO production is inhibited by Ng-nitro-L-arginine (LNAG), MCP-1 mRNA levels increase slightly above MCP-1 mRNA levels in cells cultured in the absence of L-NAG for at least 6hr. Monocyte chemotactic protein1 mRNA levels in cells cultured in the presence of L-NAG for 24 hr increase 250% in comparison to MCP-1 mRNA levels in cells cultured in the absence of L-NAG. In addition to the increase in MCP-1 mRNA levels MCP-1 protein levels also
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increase in cells cultured in the presence of L-NAG as detected by western analysis. The chemotactic activity of medium conditioned by cells cultured in the presence of L-NAG is greater than the chemotactic activity of medium conditioned by cells cultured in the absence of L-NAG. An MCP-1 antibody decreases the chemotactic activity of the conditioned medium. The addition of SIN-1 and C87–3786, NO donors to the culture medium of HUVECs induces a dose and time dependent decrease in MCP-1 mRNA and protein levels. The decrease in MCP-1 expression that NO causes is not due to increases in cGMP level. The induction of MCP-1 mRNA levels in rabbit aortic smooth muscle cells by either oxLDL or LPS can be inhibited 50 to 75% by preincubating the cells with DETA-NO (Tsao et al., 1997). Transcription run-off experiments showed that LPS and NO regulate MCP-1 mRNA levels by changing the transcription rate of the MCP-1 gene. Nitric oxide may also regulate the expression of MCP-1 in vivo (Tsao et al., 1997). Experiments were done where rabbits were fed a normal diet, a diet rich in cholesterol both with or without drinking water containing nitro-L-arginine (LNA, a NOS antagonist) and a diet rich in cholesterol plus L-arginine, the precursor to NO. The rabbits were sacrificed after 2 weeks, total mRNA was isolated from the thoracic aorta and analyzed for MCP-1 message. Total RNA isolated from rabbits fed a normal diet contained no detectable MCP-1 mRNA. Rabbits fed L-arginine in conjunction with a high cholesterol diet had lower levels of MCP-1 mRNA than rabbits fed a cholesterol rich diet with no L-arginine supplement. MCP-1 mRNA levels were the highest in rabbits fed a normal diet supplemented with LNA. Growth factors are also capable of regulating the expression of MCP-1. Epidermal growth factor (EGF), a growth stimulatory peptide, increases JE mRNA in rat heart endothelial cells in a time dependent manner with maximal expression occurring at 2 hr (Takehara et al., 1987). Pretreatment of the cells with transforming growth factorß (TGF-ß), which has been shown to inhibit vascular endothelial cell growth, inhibits the induction of MCP-1 by EGF.
EXPRESSION OF GENES INVOLVED IN VASCULAR TONE AND BLOOD PRESSURE C-Type Natriuretic Peptide Biological functions of C-type natriuretic peptide C-type natriuretic peptide is a member of the natriuretic peptide family. The other two members of the family are atrial natriuretic peptide (ANP) and brain natriuretic peptide (BNP) both of which are expressed primarily in the heart and mainly regulate blood pressure and vascular tone. A few early observations regarding CNP led many to initially think that CNP was expressed only in the brain and acted exclusively as a neuropeptide in the central nervous system. The concentration of CNP in the human brain and in human cerebrospinal fluid is at least 10 times greater than the concentration of either ANP or BNP (Ogawa et al., 1992, Minamino et al., 1991, Kaneko et al., 1993). In addition CNP was not detected in the heart or in many other
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organs (Kojima et al., 1990, Minamino et al., 1991, Komatsu et al., 1991). However, other data suggest that CNP may be a paracrine factor which acts to regulate local vascular tone and inhibits smooth muscle cell proliferation in addition to being a neuropeptide. Both CNP and its mRNA are present in many different cell types including vascular endothelial cells. Natriuretic peptides act by increasing cyclic GMP levels in cells via their receptors, atrial natriuretic peptide receptor (ANPR)-A and ANPR-B, both of which contain an intracellular particulate guanylyl cyclase domain. C-type natriuretic peptide is a potent activator of ANPR-B whereas, activation by either ANP or BNP is minimal. In contrast, ANP has a greater affinity for ANPR-A than either CNP or BNP (Suga et al., 1992a, Roller et al., 1991). Cyclic GMP levels in rat aorta SMCs (passage 3–7) cultured in the presence of CNP increase 160-fold at 10 min (Furuya et al., 1990). The increase in cGMP levels coincided with an increase in particulate guanylate cyclase activity of the SMCs membrane fraction. Regulation of C-type natriuretic peptide gene expression by shear stress C-type natriuretic peptide gene expression in HUVECs is regulated by shear stress in a magnitude and time dependent manner (Okahara et al., 1995, Figure 8.5). C-type natriuretic peptide mRNA is not detectable in HUVECs seeded on collagen coated polystyrene cultured in the absence of shear stress or cultured in the presence of shear stress (24 dyn/cm2) for 1 hr by PCR analysis. C-type natriuretic peptide mRNA is present in HUVECs cultured in the presence of shear stress for 3 hr and the mRNA level continues to increase at 6 and 12 hr. C-type natriuretic peptide is not detectable in cells cultured in the presence of shear stresses below 3 dyn/cm2 for 12 hr, but a low
Figure 8.5. The regulation of CNP gene expression in endothelial cells. Shear stress TNF-a, TNF-b and LPS all greatly increase CNP gene expression in endothelial cells. Arrows pointing inward indicate that the factor increases gene expression while the sizes of the arrows indicates the relative efficacy of the regulatory factor.
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level is present in cells cultured in the presence of shear stress at 6 dyn/cm2 and the mRNA level increases as the magnitude of the shear stress increases to 24 dyn/cm2. Chun et al. (1997) were able to detect CNP mRNA in HUVECs seeded on glass slides cultured in the absence of shear stress by RT-PCR analysis. C-type natriuretic peptide mRNA levels increase 6 and 30 fold in HUVECs cultured in the presence of shear stress (15 dyn/cm2) for 4 and 24 hr, respectively in comparison to CNP mRNA levels in cells cultured in the absence of shear stress. Shear stress increases CNP mRNA levels in BAECs and mouse lymphoid endothelial cells too. In BAECs shear stress (15 dyn/cm2) increases CNP mRNA levels 2 and 3 fold at 4 and 24 hr, respectively. In mouse lymphoid endothelial cells shear stress (15 dyn/cm2) increases CNP mRNA levels 3 and 10 fold at 4 and 24 hr, respectively. In HUVECs shear stresses of 1.5 and 3 dyn/cm2 do not increase CNP mRNA levels. Whereas, shear stresses of 5 and 15 dyn/cm2 increase CNP mRNA levels 2.5 and 4.5 fold respectively at 6 hr. There is a corresponding increase in the amount of CNP released by the cells. The concentration of CNP in the medium of cells cultured in the absence of shear stress for 6 hr is below RIA detectable level, 0.4 fmol/ml. Whereas, the CNP concentration in the medium of cells cultured in the presence of shear stress (15 dyn/cm2) for 6hr is 2.2 fmol/ml. Regulation of C-type natriuretic peptide gene expression by non-mechanical stimuli There is a basal level of CNP expression in cultured bovine carotid artery endothelial cells (BCAECs)(passage 20–25, Suga et al., 1992b). The concentration of CNP increases in the medium of the cells at a fairly constant rate for approximately 18 hr after which CNP concentration increases very little. Most of the CNP synthesized by these cells is secreted, for at the end of 24 hr there is 100 times more CNP in the medium of the cells than there is in the cells. No ANP or BNP is detectable in BCAECs conditioned medium. There is no CNP in medium conditioned by smooth muscle cells. Transforming growth factor-ß (TGF-ß) increases the release of CNP by BCAECs in a dose dependent manner. Cells cultured in the presence of TGF-ß at a concentration of 10-9 M for 24 hr secrete 130 fold more CNP than cells cultured in the absence of TGF-ß. Whereas, cells cultured in media containing TGF-ß at a concentration of 10-11–10-10 M release 30–60 fold more CNP. Thrombin, basic fibroblast growth factor (BFGF), 8-bromo cGMP, 8-bromo cAMP, TPA and arginine-vasopressin (AVP) significantly increase the level of CNP in the medium of cells cultured in their presence 1.5 to 3 fold in 24 hr. Platelet-derived growth factor has no effect on the expression of CNP in BCAECs. C-type natriuretic mRNA levels in cells treated with TGF-ß (10-10 M) for 6 hr are almost equivalent to CNP levels in rat brain in vivo. Cytokines and endotoxin also regulate the expression of CNP in BCAECs (Figure 8.5). Interleukin-1ß and IL-1a have minimal effects on the expression of CNP in BCAECs increasing the amount of CNP in the medium of the cells at 24 hr 1.5 and 3 fold, respectively in comparison to the amount of CNP in the medium of cells cultured in the absence of cytokines (Suga et al., 1993). Lipopolysaccharide has a much greater effect on the secretion of CNP increasing the amount of CNP in the medium 26 fold at 24 hr. However, TNF-a is the most efficacious, increasing the amount of CNP in the medium of BCAECs 110 fold at 24 hr. While CNP levels stop
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increasing in the medium of BCAECs cultured in the presence of IL-1ß, IL-1 and LPS at 24 hr, CNP levels in the medium of cells cultured in the presence of TNF- continue to increase at a constant rate for 48 hr. All of the agents increase CNP levels in the medium of cells in a dose dependent manner. Interleukin-1a is 10 times more potent than IL-1ß with 10 ng/ml of IL-1 inducing the same stimulation as 100 ng/ml of IL-1ß. Tumor necrosis factor- is by far the most potent stimulant with a half maximal response of 15 ng/ml. Endothelial cells cultured in the absence of smooth cells secrete only a minute amount of CNP and SMC cultures release no CNP. After 12 hr CNP levels in the medium of cocultures of bovine carotid artery endothelial cells (passage 12–20) and rat thoracic aorta SMC (passage 11–15) are 60 fold greater than CNP levels in the medium of endothelial cells and SMC cultured separately at 48 hr (Komatsu et al., 1996).
Angiotensin-Converting Enzyme Biological functions of angiotensin-converting enzyme Angiotensin-converting enzyme (ACE) is part of the renin angiotensin system (RAS) which synthesizes angiotensin II. There are two different RASs. There is the circulating RAS where angiotensin II is released into the vascular system as an endocrine factor and there is the tissue RAS where angiotensin II acts locally as an autocrine/ paracrine factor. In the circulating RAS renin is synthesized in the kidney and released into the blood where it cleaves angiotensinogen, which is synthesized and secreted by the liver, to angiotensin I. The circulating angiotensin I is then hydrolyzed in the lung by ACE to produce active angiotensin II. Angiotensin II is a vasoconstrictor, decreases renal blood flow and glomerular filtration rate in the kidney, stimulates the secretion of mineralocorticoid aldosterone from the adrenal cortex and plays a role in chronic hypertension. Angiotensin II is also thought to be involved in vascular SMCs hypertrophic response to hypertension and their hyper-plastic response to balloon angioplasty. In the brain angiotensin II increases blood pressure and thirst. Angiotensinconverting enzyme also known as kininase II, inactivates bradykinin, a vasodilator. Nearly all of bradykinin is inactivated as it passes through the lung’s vasculature system. In addition to being present in the endothelial cells of the pulmonary vascular system, ACE is also located in vascular endothelial cells outside the pulmonary circulation, in epithelial cells, especially those of the kidney, and in body fluids. In vitro ACE metabolizes many peptides in addition to angiotensin I and bradykinin including enkephalins, ß-neoendorphin, dynorphins, chemotactic peptide N-formylMet-Leu-Phe, substance P, insulin and luteinizing hormone-releasing hormone (Ehlers and Riordan 1989). How important ACE is in each of these biological systems remains to be determined. Shear stress regulation of angiotensin-converting enzyme Angiotensin-converting enzyme gene expression is regulated in bovine pulmonary artery endothelial cells (passage 4–5) by shear stress (Figure 8.6, Reider et al., 1997).
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Figure 8.6. The regulation of ACE gene expression in endothelial cells. Shear stress decreases ACE gene expression whereas hormones and glucocorticoids increase its expression. Arrows pointing inwards the center indicate that the factor increases gene expression while arrows pointing outwards indicate that the factor decreases gene expression. Curved arrows indicate that the factor acts as an enhancer. Solid arrows indicate that the factor inhibits the induction of ACE gene expression by another agent. The relative efficacy of the regulatory factors to increase or decrease ACE gene expression is represented by the sizes of the arrows.
ACE activity increases in cultured cells after they reach confluency in a time dependent manner. There is no change in ACE activity in cells cultured for 18 hr in the presence of shear stress (20 dyn/cm2) 2 days post confluency. Whereas, in cells cultured in the presence of shear stress 4 days post confluency there is a 27% decrease in ACE activity in comparison to ACE activity in cells cultured in the absence of shear stress. In these cells, ACE mRNA levels decrease 86% in the presence of shear stress (20 dyn/cm2) for 18 hr. The effect of shear stress on ACE activity is dependent upon the magnitude of the force. In cells cultured for 18 hr in the presence of shear stress at 5 dyn/cm2 there is a slight, although not significant, increase in ACE activity. At shear stresses of 10 and 15 dyn/cm2 there is a slight, but not significant, decrease in ACE activity. It is not until the magnitude of the force reaches 20 dyn/cm2 that there is a significant decrease in ACE activity. The effect of shear stress on ACE activity is also time dependent. In cells cultured in the presence of shear stress (20 dyn/cm2) for 2 hr there is a significant (>150%) increase in ACE activity in comparison to cells cultured in the absence of shear stress. At 4 hr ACE activity decreases and by 8 hr there is a significant decrease. Angiotensin-converting enzyme activity continues to decrease in cells cultured in the presence of shear stress for at least 18 hr. In vivo ACE activity decreases 43% in rat abdominal aorta 5 days after the aorta is constricted so that the shear stress on the walls rises from 12 to 26.8 dyn/cm2. Regulation of angiotensin-converting enzyme by non-mechanical stimuli Del Vecchio and Smith showed that the expression of ACE in cultured calf pulmonary artery endothelial cells is dependent upon their state of proliferation (1981). Cells
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were seeded so that they were in log phase from day 2 through 6 after seeding. After day 6 they were confluent and entered stationary phase on day 7. From day 2 of culture to day 7 there is a steady decrease in ACE activity. On day 9 ACE activity increases. On day 12 there is a 6 fold increase in ACE activity compared to day 7 and a 8 fold increase on day 14 compared to day 7. Angiotensin-converting enzyme activity on days 12 and 14 is greater than ACE activity on day 2. Thus, ACE activity is inhibited in proliferating endothelial cells but not in stationary cells. The effect of cell growth on ACE activity is due to changes in ACE protein and mRNA levels in proliferating and nonproliferating cells (Shai et al., 1992). Bovine aorta endothelial cells were seeded so that in 2–3 days they were confluent. Six days post confluency ACE activity was 12 fold greater than it was 2 days preconfluency. During the same time period ACE mRNA levels increased 21.5 fold as measured by RNase protection analysis. Immunohistochemistry analysis showed that ACE protein levels are significantly greater in growth arrested cells than in proliferating cells. Glucocorticoids and thyroid hormones increases ACE activity in vascular endothelial cells (Krulewitz et al., 1984, Mendelsohn et al., 1982). Dexamethasone increases ACE activity 5 to 7 fold in cells and 2.8 fold in the medium. Both thyroid hormones, thyroxine (T4) and triiodothyronine (T3) increase ACE activity approximately 2 fold in cells and in medium. When dexamethasone is present in the culture medium with either T3 or T4 there is an additive effect on ACE activity. The presence of insulin in the culture medium decreases ACE activity by 30 and 40% in cells and medium respectively. Insulin also partially inhibits the effect of both dexamethasone and T3 on ACE activity. Phorbol esters and cAMP increase ACE activity in a dose and time dependent manner in HUVECs (passage 3–4, Iwai et al., 1987). Angiotensin-converting enzyme activity significantly increases in HUVECs cultured in defined medium plus PMA (100 nM) at 24 hr. At 48 hr there is approximately a 20 fold increase in ACE activity in the cells and a 3.7 fold increase in the medium in comparison to ACE activities in the medium and in HUVECs cultured in the absence of PMA. There is a 6 fold increase in ACE activity in cells cultured in the presence of Dibutyl-cAMP but no change in the ACE activity of the medium. Angiotensin II regulates ACE expression in vivo (Schunkert et al., 1993, Kohara et al., 1992). In one study, Angiotensin II was infused intravenously into rats at three different doses (100, 300 and 1000 ng/kg/min) for 3 days. Plasma angiotensin II levels were significantly increased only at the two higher doses. In the lung ACE activity decreased significantly at the 2 higher doses whereas, mRNA levels decreased significantly at all three doses. Although ACE mRNA levels decreased significantly at all three doses in the testis, ACE activity only decreased significantly at the lowest dose. Serum ACE activity significantly decreased (30%) at the high dose. Angiotensinconverting enzyme expression was also analyzed after inhibiting ACE activity for 3 days by adding quinapril to the drinking water. Plasma angiotensin levels decreased but not significantly. Serum and lung ACE activities decreased significantly but ACE mRNA increased significantly in the lung. No changes in ACE activity or mRNA levels occurred in the testis. Chronic hypertension can increase ACE activity. Angiotensin-converting enzyme activity increases in dogs who have induced chronic hypertension by partial
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occlusion of the renal artery for 8 months. Increases in ACE activity occur in jejunal, pulmonary and renal arteries aorta lung and cerebral cortex but not in the plasma (Miyazaki et al., 1987). C-type natriuretic peptide inhibits ACE activity in vivo (Davidson et al., 1996). The injection of CNP (500 pmol/min) with angiotensin I into the forearm of humans inhibits the decrease in blood flow rate which occurs when angiotensin I is injected without CNP by 57%. Coinjection of CNP with angiotensin II does not significantly effect the reduced blood flow which occurs when angiotensin II is injected with out CNP. Neither ANP nor CNP inhibit ACE activity in plasma in vitro. Endothelin-1 Biological functions of endothelin-1 Endothelin-1 is the most potent vasoconstrictor identified (Yanagisawa et al., 1988). It belongs to a family of vasoconstrictor peptides whose other members are ET-2 and ET-3, both of which differ from ET-1 by a couple of amino acids. The peptide can act as an autocrine, paracrine or an endocrine factor. Endothelin-1 is synthesized as a prepropeptide with a secretory signal sequence. PreproET-1 is cleaved by a dibasic pair-specific endopeptidase to release proET-1. The biological active form of the peptide, ET-1 is cut from the carboxyl terminal of proET-1 by endothelin converting enzyme. Endothelin-1 is expressed in a wide variety of tissues with high expression levels in lung and inner medulary renal vessels. Gene transcripts are present in bovine brain, heart, lung, stomach, intestines, spleen, renal cortex, renal medulla, ovary and urinary bladder (Imai et al., 1992). Of the three endothelins only ET-1 has been located in the medium of endothelial cells and in endothelial cells at both the protein and mRNA level. Endothelin-1 binds to at least two receptors and possibly a third which are located on the surface of cells in a variety of tissues including the heart, lung, uterus, brain, adrenal gland, eye, intestine and kidney. Endothelin receptor subtype A (ETA) preferentially binds ET-1 and ET-2 whereas, endothelin receptor subtype B (ETB) has an equal affinity for all three peptides. Both receptors are members of the seven transmembrane G-protein coupled receptor superfamily. Dependent upon the cell type, the ETA and ETB coupled G protein may activate ion channels, phospholipase C, phospholipase A2 and/or phospholipase D. The administration of ET-1 to animals usually causes an initial transient decrease in arterial blood pressure lasting only a few minutes followed by a more prolonged pressor action of several hours. The initial depressor activity is thought to be due to the transient increase in vasodilators prostacyclin (PGI2) and endothelium derived relaxation factor induced by ET-1 (De Nucci et al., 1988). Endothelin-1 is cleared from the blood mainly by the lungs and kidneys with a half-life of less than 7 min. Due to the low concentration of ET-1 in plasma, 1.5–3.7 pg/ml it is thought that a large portion of the ET-1 released from vascular endothelial cells is secreted abluminally where it functions as an autocrine factor on the endothelial cells and a paracrine on the SMCs. Endothelin-1 effects the heart, lung, anterior pituitary, adrenal glands, thyroid, parathyroid, kidney, liver, gastrointestinal tract and monocytes (McMillen and Sumpio, 1995). Endothelin-1 acts as vasoconstrictor, stimulates the release of eicosanoids, stimulates ANP, and
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increases catacholamine and aldosterone secretion by the adrenal gland. Endothelin is mitogenic for endothelial cells, smooth muscle cells, astrocytic glial cells, osteoblastic cells, melanocytes, mesangial cells and fibroblasts (Battistini et al., 1993). In many cases ET-1 will act synergistically with other mitogens such a EGF, TGF-, PDGF, vasopressin, TPA, forskolin and IGF-1. Endothelin-1 is a chemoattractant and activator of monocytes. In the kidney ET-1 increases urine flow rate and decreases glomerular filtration rate and renal blood flow (Simonson, 1993). Endothelin-1 is thought to play a role in the pathogenesis of many diseases including atherosclerosis, congestive heart failure, restinosis, arterial hypertension, pulmonary hypertension, acute and chronic renal failure, liver cirrhosis and ascites and in cyclosporine toxicity (Hocher et al., 1996). Regulation of endothelin-1 gene expression by shear stress Endothelin-1 gene expression increases in porcine thoracic aorta endothelial cells (passage 5–10) cultured in a cone and plate viscometer subjected to a low shear stress of 5 dyn/cm2 (Yoshizumi et al., 1989, Figure 8.7). PreproET-1 mRNA levels increase in cells cultured in the presence of shear stress at 1 hr and are maximal at 4 hr. PreproET-1 mRNA levels decrease at 12 hr and by 24 they have returned to basal
Figure 8.7. The regulation of ET-1 gene expression in endothelial cells. Endothelin-1 gene expression is controlled by a wide variety of regulatory factors in endothelial cells. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outwards indicate that the factor decreases gene expression. Curved arrows indicate that the factor acts as an enhancer. Solid arrows indicate that the factor inhibits the reduction of ET-1 gene expression by shear stress. The relative efficacy of the regulatory factors to increase or decrease ET-1 gene expression is represented by the sizes of the arrows.
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level. The same result was obtained for steady and pulsatile flow. Endothelin-1 protein levels increase 1.6 and 1.5 fold in the medium of cells cultured in the presence of shear stress (5 dyn/cm2) in comparison to ET-1 levels in the medium of cells cultured in the absence of shear stress at 24 and 48 hr. Thrombin causes an additional increase in preproET-1 mRNA levels when added in conjunction with the shear stress. PreproET-1 mRNA levels are greater in cells cultured in the presence of shear stress for 20 hr and then in the presence of shear stress (5 dyn/cm2) plus thrombin for 4 hr, than in cells cultured in the presence of shear stress (5 dyn/cm2) for 24 hr. Sharefkin et al. (1991) reported a striking decrease in ET-1 gene expression in primary HUVECs cultured in a parallel plate flow chamber exposed to a shear stress of 25 dyn/cm2 for 24 hr. PreproET-1 mRNA levels decrease to near undetectable levels by RT-PCR in cells cultured in the presence of shear stress. The rate of accumulation of ET-1 decreases 60–80% in the medium of the cells cultured in the presence of shear stress (25 dyn/cm2) in comparison to the rate of accumulation of ET-1 in the medium of cells cultured in the absence of shear stress. PreproET-1 mRNA levels also decrease in BAECs (passage 6–15) cultured in a cone and plate viscometer subjected to a shear stress of 15 dyn/cm2 (Malek and Izumo, 1992). Decreases in preproET-1 mRNA levels in cells cultured in the presence of shear stress first occur at 1 hr. At 2 hr ET-1 mRNA levels are 4–5 fold less than in cells cultured in the absence of shear stress. After 2hr preproET-1 mRNA levels remain fairly constant until 18 hr. The amount of endothelin in the medium of cells cultured in the presence of shear stress (15 dyn/cm2) also decreases with time. Endothelin-1 levels decrease 41 and 47% at 24 and 48 hr. Thus it appears shear stress regulates ET-1 mRNA levels in a magnitude dependent manner. In cells cultured for 4 hr in the presence of shear stress at 3 dyn/cm2 there is not a significant difference in preproET-1 mRNA levels compared to mRNA levels in cells cultured in the absence of shear stress. Whereas, shear stress of 8 and 15 dyn/ cm2 decrease mRNA levels by approximately 40 and 70%. Turbulent and pulsatile flow have the same effect on ET-1 gene expression as laminar flow. PreproET-1 mRNA levels increase in cells cultured in the presence of cyclohexamide for 1 hr (Malek et al., 1993). Similar to shear stress, PMA causes ET-1 mRNA levels to decrease in cells cultured in its presence at 1 hr and the message is undetectable by Northern analysis at 24 hr. Preincubation of cells with cyclohexamide does not completely block the reductions in ET-1 mRNA levels that occur in cells treated with PMA. However, cyclohexamide does completely block shear induced decreases in ET-1 mRNA levels, suggesting that shear stress and PMA may decrease ET-1 gene expression using, at least partially, separate signaling pathways. PMA stimulation of BAECs results in a transfer of PKC activity from the soluble to the particulate fraction in 15 min whereas, shear stress (20 dyn/cm2) does not. Nor does the PKC a isozyme shift from the soluble fraction to the particulate fraction of cells subjected to shear stress as it does in cells stimulated with PMA. In addition PKC inactivator calphostin does not inhibit the decrease in ET-1 gene expression that shear stress causes. Thus, the reduction in the expression of the ET-1 gene by shear stress in BAECs may not involve PKC activation. Kuchan and Frangos (1993) report that the early increase in ET-1 gene expression in primary HUVECs cultured in a parallel plate flow chamber exposed to shear
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stresses involves the activation of PKC. Cells cultured in the presence of shear stress, 10 dyn/cm2 for 1 hr or less secrete at least 2 fold more ET-1 than cells cultured in the absence of shear stress. Cells cultured in the presence of shear stress, 10 dyn/cm2 for 10 sec to 15min and then cultured in the absence of shear stress for 6hr secrete more ET-1 than cells never exposed to shear stress although the shorter their exposure to shear stress is, the more ET-1 they secrete. Protein kinase C inhibitor staurosporine and PKC/PKG inhibitor H-7 block the short term flow induced increase in ET-1 secretion. Whereas, HA 1004 a PKG/PKA inhibitor does not affect the increase. Kuchan and Frangos also report that shear stress regulates ET-1 gene expression in a bimodal magnitude dependent manner with low magnitude shear stresses increasing ET-1 gene expression and high magnitude shear stresses decreasing gene expression. In cells exposed to a shear stress of 1.8 dyn/cm2 there is a significant increase in the amount of ET-1 secreted from the cells at 6hr whereas, shear stresses of 12 and 25 dyn/cm2 cause significant decreases in the release of ET-1. Cells subjected to a shear stress of 1.8 dyn/cm2 for 16 hr secrete 1.7 fold more ET-1 than cells cultured in the absence of shear stress. Whereas, HUVECs secrete 40 and 70% less ET-1 when they are cultured in the presence of a shear stress of 12 and 25 dyn/cm2, respectively for 16 hr. The regulation of ET-1 gene expression by shear stress may be mediated by NO. LNNA, a competitive inhibitor of NOS blocks the shear stress induced decrease in ET1 secretion in a dose dependent manner. It may be that the NO induced increase in cGMP levels are responsible for the decrease in ET-1 gene expression, as the treatment of the cells with ANP or 8-bromo-cGMP cause similar decreases in ET-1 secretion similar to shear stress. In addition, L-NNA also inhibits flow induced increases in cGMP levels. The inability of HA 1004 to effect shear induced decreases in ET-1 secretion further suggest that this cGMP pathway does not involve the activation of PKG. Regulation of endothelin-1 gene expression by nonmechanical stimuli Endothelin-1 gene expression is regulated by many factors in addition to shear stress (Figure 8.7). Endothelin-1 gene is expressed in porcine aortic intima and in porcine aortic endothelial cells (PAECs) in vitro. Cultured PAECs secrete ET-1 at a constant basal rate for at least 9 hr. The release of ET-1 from these cells is regulated by thrombin in a dose dependent manner (Schini et al., 1989). Cells cultured in the presence of thrombin (10U/ml) secrete approximately 2.1 times more ET-1 than cells cultured in the absence of thrombin for 6 hr. Cyclohexamide inhibits the secretion of ET-1 from cells cultured in the absence and presence of thrombin. In PAECs cultured in the presence of thrombin or Ca2+ ionophore A23187 preproET-1 mRNA levels increase dramatically at 1 hr (Yanagisawa et al., 1988). Adrenaline also increases the rate of secretion of ET-1 from porcine aorta endothelial cells in a dose dependent manner (Kohno et al., 1989). Pulmonary artery endothelial cells cultured in the presence of adrenaline (10-5 M) secrete approximately 1.6 fold more ET-1 than cells cultured in the absence of adrenaline at 6 hr. Cyclohexamide (5 µg/ml) completely blocks the secretion of ET-1 by PAECs cultured in the presence of adrenaline. Cells cultured in the presence of adrenaline plus a-adrenergic blocker phentolamine secret the same amount of ET-1 as cells cultured in the absence of
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adrenaline whereas, cells cultured in the presence of adrenaline plus ß-adrenergic blocker propranolol secrete the same amount of ET-1 as cells cultured in the presence of adrenaline. This suggest that the regulation of ET-1 secretion by adrenaline is mediated by the -adrenergic receptor. Transforming growth factor-ß1 increases preproET-1 mRNA levels in adult PAECs (passage 5–10) in a dose dependent manner (Kurihara et al., 1989). PreproET-1 mRNA levels increase at 1 hr in cells cultured in the presence of TGF-ß1 in comparison to preproET-1 mRNA levels in cells cultured in the absence of TGF-ß1. At 2 hr preproET1 mRNA levels are maximal having increased 4 fold compared to mRNA level in unstimulated cells. PreproET-1 mRNA levels decrease to basal levels at 24 hr. The amount of ET-1 secreted by PAECs incubated in the presence of TGF-ß1 is 1.7 to 1.8 fold greater at 12 and 24 hr than the amount of ET-1 secreted by cells incubated in the absence of TGF-ß1. PreproET-1 mRNA levels are not altered by ADP, serotonin, 9,11-epithio-12-methano-thromboxane A2, an analogue of thromboxane A2 or PDGF in vitro. Endothelin-1 gene expression in PAECs is also regulated by cytokines in vitro (Yoshizumi et al., 1990). Interleukin-1 increases preproET-1 mRNA levels in a biphasic time dependent manner. In IL-1 stimulated cells preproET-1 mRNA levels increase 3.2 fold at 2 hr in cells cultured in the presence of IL-1 in comparison to preproendothelin mRNA levels in cells cultured in the absence of IL-1. PreproET-1 mRNA levels decrease to a minimum at 8 hr at which time they are 2 fold greater in IL-1 treated cells than in untreated cells. A second maximum is reached at 24 hr when preproET-1 mRNA levels are 4.8 fold greater in the cells cultured in the presence of IL-1 than in cells cultured in the absence of IL-1. PreproET-1 mRNA levels decrease in the cells at 24 but they remain elevated at 48 hr in comparison to mRNA levels in untreated cells. Interleukin-1ß has a similar effect on preproET-1 mRNA levels in endothelial cells. In contrast to the bimodal expression of preproET-1 mRNA in IL-1a treated cells, the secretion rate of ET1 by IL-1 treated PAECs is fairly constant with time. At 6 hr significantly more ET-1 accumulates in the medium of cells cultured in the presence of IL-1 than in the medium of cells cultured in the absence of IL-1. Both IL-1 and IL-1ß increase ET-1 gene expression in a dose manner. However, IL-1 is approximately 5 times more potent than IL-1ß. Endothelin-1 gene expression is regulated in HUVECs in vitro. Diacylglycerol homologue TPA and calcium ionophore ionomycin increase preproET-1 mRNA levels in primary HUVECs (Yanagisawa et al., 1989). PreproET-1 mRNA levels increase in 10 min in cells cultured in the presence of TPA in comparison to preproET-1 mRNA levels in cells cultured in the absence of TPA. In 20 min, preproET-1 mRNA levels are maximal after which the amount of preproET-1 mRNA in the cells begins to decrease. At 90min preproET-1 mRNA can no longer be detected in the HUVECs by Northern analysis. Ionomycin regulates preproET-1 mRNA levels in HUVECs in a similar manner. In ionomycin treated cells an increase in preproET-1 mRNA occurs within 10 min and the level peaks at 20 min. However, in ionomycin treated cells preproET1 mRNA levels decrease slower as preproET-1 mRNA is present in the cells at 4hr. Neither TPA nor ionomycin effect the short half-life (15 min) of preproET-1 mRNA in HUVECs. Cyclohexamide causes a super-induction of preproET-1 mRNA levels in
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cells incubated in the presence or absence of TPA and ionomycin at 20min. Forskolin which activates cAMP has no effect on preproET-1 mRNA levels in HUVECs. The release of ET-1 from HUVECs (passage 3–4) is regulated by LPS in a biphasic dose dependent manner. At a low dose, 1, 10 and 250 ng/ml LPS increases the release of ET-1 from HUVECs incubated for 4 hr in its presence. In high concentrations, 1 µg/ml, LPS decreases the secretion of ET-1 30% at 4 hr (Ros et al., 1996). Interestingly, LPS at concentrations of 1 and 10 ng/ml have no effect on the level of preproET-1 mRNA in HUVECs at 4 hr. Low oxygen tensions increase preproET-1 mRNA levels in primary HUVECs (Kourembanas et al., 1991). PreproET-1 mRNA levels increase 2 fold at 1 hr and 6 fold at 48 hr in HUVECs cultured in the presence of 1 % oxygen in comparison to preproET-1 mRNA levels in HUVECs cultured in the presence of 21% oxygen. The message level decreases to basal levels 24 hr after the cells are reexposed to 21% oxygen. The increase in the preproET-1 mRNA levels is not due to an increase in the half-life of ET-1 mRNA as it is the same whether the cells are cultured in the presence of 1% oxygen or 21% oxygen. The transcription rate of ET-1 gene increases 4–8 fold in cells cultured in the presence of 1% oxygen at 6, 24 and 48 hr as quantitated by nuclear run off analysis. The amount of ET-1 in the medium of the cells cultured in 1% oxygen also increases by 4–8 fold at 6 and 24 hr. Angiotensin II increases ET-1 protein levels in rat aorta, femoral artery and kidneys in vivo (Barton et al., 1997). In rats infused with buffered saline ET-1 protein levels in the aorta is 44 pg/gm tissue with and without the endothelium. In contrast, in the femoral artery ET-1 protein levels are 127 and 34 pg/gm total tissue with and without the endothelium indicating that the distribution of ET-1 protein in blood vessels is dependent upon the artery. In rats infused with angiotensin II for two weeks ET-1 protein levels increases 3 fold in the kidney, 1.6 fold in the femoral artery and 4.7 fold in the aorta in comparison to levels in rats infused with buffered saline. In the aorta with the endothelium removed ET-1 protein levels are 44 and 137 pg/mg total protein in control and treated rats increasing 3.1 fold. In the femoral artery with the endothelium removed ET-1 protein levels are 34 and 159 pg/mg total protein increasing 4.7 fold. Thus, angiotensin II effects ET-1 protein levels in SMCs more than in endothelial cells. PreproET-1 mRNA is not detectable in rat cardiac micro vascular endothelial cells (CMEC) by Northern analysis. However, preproET-1 transcripts are synthesized in CMECs cocultured with adult rat ventricular myocytes (ARVM) (Nishida et al., 1993). PreproET-1 mRNA is also present in CMECs cultured 48 hr in the presence of TGF-ß, angiotensin II, thrombin and conditioned media form CMEC-ARVM cocultures. In contrast, there is no preproET-1 mRNA present in cells cultured in the presence of media conditioned by ventricular myocyte cultures. The regulation of ET-1 expression in CMECs by TGF-ß is dose dependent. PreproET-1 mRNA levels are maximal in cells cultured in the presence of 500 pg/ ml of TGF-ß. At higher concentrations of TGF-ß preproET-1 mRNA levels decrease. Transforming growth factor-ß mRNA levels increase in CMECs after 3 days of coculture with AVRMs. That is one day before preproET-1 mRNA is at detectable levels in the cells. Transforming growth factor-ß and preproET-1 mRNA levels continue to increase in the CMECs until at least day 7. The increase in ET-1
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mRNA levels that occurs in the cocultures are blocked by the addition of antibodies to TGF-ß. Thus, it may be that it is the stimulation of TGF-ß gene expression in the cocultures that causes the increase in ET-1 gene expression.
Prostaglandin H Synthase 1 and Prostaglandin H Synthase 2 Prostaglandin H synthase 1 and prostaglandin H synthase 2 biological structures and functions Prostaglandin H synthase, also known as cyclooxygenase (COX), is encoded for in two genes, PGHS-1 and PGHS-2. The two genes are located on separate chromosomes. The human PGHS-1 gene is present on chromosome 9 and the PGHS-2 gene is located on chromosome 1. The human PGHS-1 gene is 22 kb in length consisting of 11 exons producing a mRNA 2.8 kb in length (Yokoyama and Tanabe, 1989). In contrast, the human PGHS-2 gene is only 7.5 kb in length and consist of 10 exons (Tazawa et al., 1994). Prostaglandin H Synthase-2 mRNAs are predominantly 4–4.5 kb in length with a long 3’ untranslated region containing 12 AUUUA motifs. However, shorter transcripts, 2.8 kb in length are synthesized using a polyadenylation site in the 3’ untranslated region of the gene different from the one used in the synthesis of the 4–4.5 kb transcripts. The AUUUA motif present in the 3’ untranslated region of the PGHS-2 gene transcripts is known to confer instability upon mRNA. The PGHS-1 promoter lacks a TATA box and is GC rich, both of which are characteristics of constitutively expressed genes. In contrast, the structure of the PGHS-2 gene, which includes a promoter with a TATA box, is that of an inducible gene. The PGHS-1 protein has a calculated molecular mass of 66 kDA and migrates as a single band on a SDS-PAGE gel as a 72 kDa protein. The difference in molecular weights is due to three N-linked high-mannose oligosaccharides on the protein. The PGHS-2 protein has a calculated molecular mass of 67 kDa and runs as a doublet on a SDS-PAGE gel as 72 and 74 kDA proteins. The 72 kDA protein has 3 N-linked oligosaccharides whereas, the 74 kDa protein has 4 N-linked oligosaccharides. The PGHS-1 gene is expressed constitutively in the majority of cell types whereas, the PGHS-2 gene is expressed constitutively in only a few tissues such as the brain. However, the expression of PGHS-2 is inducable in a wide variety of cell types by cytokines, endotoxins, phorbol esters, growth factors, prostaglandins and hormones. Prostaglandin H synthase catalyzes the rate limiting step in the synthesis of prostanoids from free arachidonic acid (AA). Arachidonic acid is converted to prostaglandin H2 (PGH2) by PGHS in a two step reaction. First the cyclooxygenase activity of PGHS converts AA to prostaglandin G2 (PGG2) and then PGG2 is reduced to PGH2 by the peroxidase activity of the enzyme. The cyclooxygenase active site on PGHS is separate and distinct from the peroxidase active site. Prostaglandin H2 is further metabolized to one of the biologically active prostanoids, prostacyclin (PGI2), prostaglandin E2 (PGE2), prostaglandin F2a (PGF2a), prostaglandin D2 (PGD2) or thromboxane A2 by their specific synthase. The particular prostanoid synthesized is dependent upon the cell type, for each cell type usually produces one prostanoid
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more abundantly than the other prostanoids. For example, PGI2 is the major prostanoid synthesized in many endothelial cells whereas, PGE2 is the predominant prostanoid in osteoblasts and thromboxane A2 predominate in platelets. The prostanoids are then released by the cells to act locally as either autocrine or paracrine factors. The biological functions of prostanoids are numerous. Prostanoids play a role in hemostasis, kidney functions, platelet aggregation, pain, fever, respiratory functions, reproduction, gastric functions, central nervous system processes and the immune system. During the synthesis of PGH 2, PGHS is irreversibly autoinactivated after approximately 1300 reactions. In endothelial cells, the majority of PGHS has a very short half-life of 10 min but, there is a pool of PGHS which has a half-life greater than 150 min. Prostaglandin H Synthase is biologically active as a homodimer embedded monotopically in the bilayer of the endoplasmic reticulum and the nuclear envelope. Prostaglandin H Synthase-1 is mainly located in the endoplasmic reticulum whereas, PGHS-2 is concentrated in the nuclear envelope. Both enzymes are inactivated by nonsteroidal antiinflammatory drugs. However, there is evidence indicating that it is the inactivation of PGHS-2 which is therapeutic, while some of the side effects of the drugs are due primarily to the inactivation of PGHS-1. Shear stress regulation of prostaglandin H synthase 1 and prostaglandin H synthase 2 gene expression The expression of PGHS in endothelial cells and in bone cells is regulated by shear stress. In HUVECs cultured in a parallel plate flow chamber exposed to shear stress PGHS-1 and PGHS-2 protein levels are effected by shear stress (unpublished results, McCormick and McIntire). In cells cultured in the presence of 4, 15 and 25dyn/cm2 PGHS-1 protein levels decrease 74, 63 and 69% at 10 min in comparison to PGHS-1 protein levels in HUVECs cultured in the absence of shear stress. Prostaglandin H synthase-1 protein level increased relative to their levels at 10 min, but at 4 and 25 dyn/cm2 they are still below PGHS-1 protein levels in cells cultured in the absence of shear stress at 30 min, whereas PGHS-1 protein levels in cells cultured in the presence of 15 dyn/cm2 are equivalent to PGHS-1 protein levels in cells cultured in the absence of shear stress. At 12 hr PGHS-1 protein levels increase and are 1.84, 2.68 and 2.49 fold greater in cells cultured in the presence of shear stress of 4, 15 and 25 dyn/cm2 respectively than PGHS-1 protein levels in cells cultured in the absence of shear stress. At 24 hr PGHS-1 protein levels are still elevated in cells cultured in the presence of 4, 15 and 25 dyn/cm2 at levels 2.49, 2.84 and 1.78 times greater than levels in cells cultured in the absence of shear stress. Prostaglandin H synthase-2 protein levels decrease in cells cultured in the presence of shear stress at 10 min similar to PGHS-1 protein levels although not as drastically as PGHS-1 protein levels. At 4, 15 and 25 dyn/cm2 PGHS-2 protein levels decrease 59, 44 and 58% at 10 min. At 30 min in cells cultured in the presence of 4 and 15 dyn/cm2 PGHS-2 protein levels are slightly greater than protein levels in cells cultured in the absence of shear stress. At 12 hr, PGHS-2 protein levels are greater than control levels similar to PGHS-1 protein levels. At 4, 15 and 25 dyn/cm2 protein levels increase 1.85, 1.42 and 2.54 fold above control levels. At 24 hr PGHS-2 protein levels decrease 50 and 60% compared to
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control levels at 15 and 25 dyn/cm2. At 4dyn/cm2 PGHS-2 protein levels are 2.22 fold greater than PGHS-2 protein levels in cells cultured in the absence of shear stress. PGHS-2 mRNA levels increase in HUVECs (passage 2–3) cultured in a cone and plate viscometer in the presence of laminar shear stress (10dyn/cm2) at 1, 6 and 24 hr but not in HUVECs cultured in the presence of turbulent shear stress (10 dyn/ cm2) in comparison to PGHS-2 mRNA levels in cells cultured in the absence of shear stress (Topper et al., 1996). Laminar shear stress, 10 dyn/cm2 for 24 hr also increases PGHS-2 protein levels in HUVECs whereas, nonlaminar shear stress, 10 dyn/cm2 has no effect on PGHS-2 protein levels. Transcription runoff assays showed that laminar shear stress increases PGHS-2 mRNA levels at least in part by increasing the transcription rate of the gene. In contrast to what we have found, Topper et al., did not observe any changes in PGHS-1 gene expression due to laminar shear stress. Regulation of prostaglandin H synthase 1 and prostaglandin H synthase 2 gene expression by non-mechanical stimuli Prostaglandin H synthase-1 and PGHS-2 gene expression is regulated in many cell types including fibroblasts, macrophages, monocytes, intestinal epithelial cells, chondrocytes, ovarian granulosa cells, amnion cells, decidual cells, mesangial cells, mast cells, SMCs, bone cells and endothelial cells. In these cells many factors regulate PGHS gene expression including cytokines, growth factors, endotoxin, serum and hormones (Herschman, 1994, 1995; Smith and Dewitt, 1996). In this chapter we will only discuss the regulation of PGHS in vascular endothelial cells. Prostaglandin H Synthase-1 and PGHS-2 gene expression are regulated by both cytokines and phorbol esters in endothelial cells in vitro (Figures 8.8 and 8.9). In HUVECs, IL-1a regulates PGHS-2 protein levels in a time dependent manner (Habib et al., 1993). In cells cultured in the presence of IL-1a plus serum, PGHS-2 mRNA levels increase 4.4, 6.5 and 22 fold in comparison to PGHS-2 levels in cells cultured in the absence of IL-1 but in the presence of serum at 2, 6 and 24 hr respectively. When the cells are cultured in presence of IL-1a plus BSA, in place of serum, PGHS2 levels increase by a factor of 8 and 12 in comparison to PGHS-2 levels in cells cultured in the presence of BSA at 2 and 6 hr respectively. The additional increase is accounted for by the increase in PGHS-2 mRNA levels in cells cultured in the presence of serum and the lack of induction of PGHS-2 gene expression in cells cultured in the presence of BSA. Interleukin-1a also increases PGHS-1 protein and mRNA levels in HUVECs in vitro although the increase is not as large as the increase IL-1a induces in PGHS-2 HUVECs cultured in the presence of IL-1a at 6hr and continue to increase for at least 15 hr in comparison to PGHS-1 protein levels in cells cultured in the absence levels (Maier et al., 1990). Prostaglandin H synthase-1 protein levels increase in of IL-1a. In addition to increasing PGHS-1 protein levels, IL-1a also increases PGHS-1 mRNA levels in HUVECs. Prostaglandin H synthase-1 mRNA levels increase at 2 hr and remain elevated for at least 24 hr in cells cultured in the presence of IL-1a. Pretreatment of the cells with cyclohexamide increases PGHS-1 mRNA levels. Actinomycin D inhibits
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Figure 8.8. The regulation of PGHS-1 gene expression in endothelial cells by shear stress and non-mechanical stimuli. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outward indicate that the factor decreases gene expression. The relative efficacy of the regulatory factors to increase or decrease PGHS-1 gene expression is represented by the sizes of the arrows. Microvascular cells (MVCs).
Figure 8.9. The regulation of PGHS-2 gene expression in endothelial cells by shear stress and non-mechanical stimuli. Arrows pointing inward indicate that the factor increases gene expression while arrows pointing outward indicate that the factor decreases gene expression. The relative efficacy of the regulatory factors to increase or decrease PGHS-2 gene expression is represented by the sizes of the arrows.
IL-1 induced increases in PGHS-1 mRNA levels indicating that IL-1a may increase PGHS-1 mRNA levels by increasing the transcription rate of the gene. In contrast to the sustained elevation in PGHS-2 protein levels induced by IL-1, PMA causes a transient increase in PGHS-2 protein levels in HUVECs (Habid et al., 1993). In cells cultured in the presence of PMA plus serum PGHS-2 protein levels
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increase 9.6 and 8.6 fold at 2 and 6 hr respectively but, at 24 hr the protein is almost undetectable by Western analysis. In cells cultured in the presence of PMA plus BSA, PGHS-2 levels increase by a factor of 12.7 and 38 at 2 and 6hr, respectively. PMA also transiently increases PGHS-2 mRNA levels in HUVECs (Hla and Neilson, 1992). In HUVECs cultured in the presence of PMA, PGHS-2 mRNA levels begin to increase at 1 hr with a 10.5 fold increase, peak at 6 hr with a 28 fold increase and decrease at 10 hr to a 13.8 fold increase in comparison to PGHS-2 mRNA levels in cells cultured in the absence of PMA. After 10 hr PGHS-2 mRNA levels remain constant for at least 24 hr. Although Habid et al. (1993) reported that PMA does not increase PGHS-1 protein levels, there are reports showing that PGHS-1 gene expression increases in HUVECs cultured in the presence of PMA in comparison to cells cultured in the absence of PMA (Hla and Neilson, 1992, Xu et al., 1996). Xu et al. (1996) reported that PGHS-1 protein levels increase in cells cultured in the presence of PMA at 1 hr and continue to increase at 6 hr. Two groups have reported similar results for PMA induced increases in PGHS-1 mRNA levels with slight variations in the amount of time it takes for maximal mRNA levels to be reached. Xu et al. (1996) reported maximal stimulation in PGHS-1 mRNA levels occurs at 4 hr with a 2 fold increase in mRNA levels when quantitated by Northern analysis and a 1.7 fold increase by quantitative RT-PCR analysis. After 4 hr PGHS-1 mRNA levels remain elevated for at least 24 hr. In comparison, Hla and Neilson (1992) reported that in HUVECs cultured in the presence of PMA, PGHS-1 mRNA levels begin to increase at 6 hr and reach maximum at 10 hr with a 2.8 fold increase in comparison to mRNA levels in cells cultured in the absence of PMA. The time differences may be due to different cell passage numbers 2 verses 4–8 or due to different culture conditions. Pretreatment of the cells for 4 hr with actinomycin D inhibits the PMA induction. Cyclohexamide pretreatment also inhibits PMA induced increase in PGHS-1 mRNA indicating that protein and mRNA synthesis is required for PMA regulation of PGHS-1 gene expression. Protein kinase C inhibitors staurosporine and calphostin prevent PMA induced increases in PGHS1 levels suggesting that PMA regulation of PGHS-1 gene expression requires the activation of PKC. Heparin binding growth factor-1 (HBGF-1), also known as acidic fibroblast growth factor, regulates the expression of the PGHS-1 gene in HUVECs. Prostaglandin H synthase-1 mRNA levels decrease 7 fold at 48 hr in HUVECs cultured in the presence of serum plus heparin plus HBGF-1 in comparison to PGHS1 mRNA levels in cells cultured in the presence of serum plus heparin but the absence of HBGF-1 (Hla and Macing, 1991). The regulation of PGHS-1 mRNA levels by HBGF-1 is dose dependent with 0.1ng/ml being capable of reducing PGHS-1 mRNA levels. Heparin binding growth factor-1 does not decrease PGHS-1 mRNA levels until 20 hr but mRNA levels remain depressed for at least 96 hr. Prostaglandin H synthase-1 protein levels also decrease in HUVECs cultured in the presence of HBGF1 plus heparin at 24 and 72 hr in comparison to protein levels in cells cultured in the absence of HBGF-1 plus heparin. In contrast to the decrease in PGHS-1 mRNA levels in HUVECs cultured in the presence of HBGF-1, PGHS-1 mRNA levels increase in microvascular endothelial cells cultured in the presence of HBGF-1 (Moatter and
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Gerritsen, 1992). Prostaglandin H synthase-1 protein synthesis increases 2 fold in rabbit microvascular endothelial cells cultured in the presence of HBGF-1 compared to PGHS-1 levels in microvascular cells cultured in the absence of HBGF-1 for 5 hr.
CONCLUSION Shear stress regulates gene expression in ways similar to other regulatory factors and in some cases can be just as important as biochemical modulators are in controlling gene expression in vascular endothelial cells. As other regulatory factors effect different genes differently, so does shear stress. Shear stress may either increase or decrease the expression of a gene. Early expression of ICAM-1, MCP-1 and CNP is increased by shear stress whereas, expression of E-selectin, VCAM-1 and ACE is decreased by shear stress. Even the qualitative type of expression modulated by shear stress is sometimes dependent upon the magnitude of the shear stress as it is for ET-1. Low shear stresses increase ET-1 gene expression whereas high shear stresses decrease its expression. Time can also be a critical factor in gene regulation by shear stress. MCP1 gene expression is initially increased by shear stress. However, after 5 hr MCP-1 mRNA levels are below control levels. Shear stress also has a biphasic effect on PGHS1 and PGHS-2 gene expression, except protein levels are first decreased and then increased. Like other regulatory factors, the degree to which shear stress regulates the expression of a gene in vascular endothelial cells varies from gene to gene. For some genes, shear stress has no or very little effect on the expression of the gene. However, there are also genes whose expression is modulated quite effectively by shear stress. In deciding the significance of shear stress in the regulation of a gene, the effect of other regulatory factors on the expression of the gene should be considered. There are genes whose expression is regulated by shear stress, but the change in gene expression is minute in comparison to that induced by more efficacious regulatory factors. However, there are also genes that are regulated equally by shear stress and other regulatory factors. E-selectin gene expression is only slightly modulated by shear stress whereas, cytokines and LPS cause large increases in E-selectin mRNA and protein levels in endothelial cells. The decrease in E-selctin expression is small in comparison to the increases caused by IL-1, TNF- and LPS. IFN- is similar to shear stress in that it does not significantly effect the expression of E-selectin alone. However, it does enhance the effect TNF- and IL-1 have on E-selectin gene expression. It is not known if shear stress acts similarly to either enhance or inhibit the actions of TNF- and IL-1 on Eselectin gene expression. Shear stress regulates ICAM-1 gene expression in a manner similar to that of IL-1. ICAM-1 protein levels increase approximately 1.5–2.5 fold in endothelial cells at 4– 8 hr. At 24–48 hr there is a 2.5–3 fold increase except for when the cells are grown on NaOH treated glass then cells surface expression decreases. These increases are similar to those caused by IL-1. In addition both the lag time before expression increases and the duration for which it lasts are nearly the same for shear stress and IL-1. IFN- acts to synergistically increase ICAM-1 expression with TNF- and there is an additive
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increase when it is combined with IL-1. The effect IFN- has when it is present with shear stress on ICAM-1 gene expression is unknown. Shear stress decreases VCAM-1 gene expression in endothelial cells whereas, cytokines and LPS increase its expression. Both cytokine and LPS effects are inhibited by NO. Since NO levels in endothelial cells are increased by shear stress it is plausible that shear stress may at least in part regulate VCAM-1 gene expression by increasing NO levels. In addition, shear stress probably decreases the effectiveness of cytokines and LPS in increasing VCAM-1 gene expression. Shear stress increases MCP-1 mRNA levels much less than cytokines and LPS. Shear stress increases MCP-1 mRNA levels maximally 2.5 fold after 1.5 hr. In comparison, IL-1 increases MCP-1 mRNA levels 5 fold after 30 min and LPS increases MCP-1 mRNA levels 7 fold after 1 hr and these are not maximum values. However, shear stress does drastically decrease MCP-1 expression at later time points. At 5 hr shear stress causes MCP-1 gene expression levels to decrease below detectable levels. Nitric oxide also has an inhibitory effect on the expression of MCP-1. Similar to its regulation of VCAM-1 gene expression, shear stress’s long term regulation of MCP-1 gene expression may be due to the increase in NO levels it causes. Shear stress increases CNP levels greater than IL-1, BFGF, cGMP, cAMP, TPA and AVP. The increase in CNP expression it causes is comparable to that caused by TGFß, LPS and TNF-. Whether or not shear stress will continue to stimulate CNP secretion past 24 hr as TNF- does or whether there will be a plateau in its expression similar to that induced by LPS and TGF-ß has not been determined so far. However, shear stress does appear to be as important as the other regulatory factors in controlling CNP gene expression. Shear stress is an effacious down regulator of ACE gene expression decreasing its expression more than insulin. Glucocorticoids and thyroid hormones increase ACE gene expression quite significantly. However, insulin inhibits these factors from increasing ACE gene expression. It is not known if shear stress acts similar to insulin and inhibits the regulation of ACE gene expression by glucocorticoids and thyroid hormones. Low magnitude shear stresses increase ET-1 gene expression to levels comparable with thrombin, adrenaline, TGF-ß, angiotensin II and IL-1 stimulation. Low oxygen tensions increase ET-1 protein levels approximately twice as much as low magnitude shear stress. The biphasic regulation of ET-1 by shear stress is similar to that of LPS. Low levels of LPS increase ET-1 protein levels comparable to low shear stress levels. The percent decrease in ET-1 protein levels is also similar for high LPS concentrations and high shear stress levels. However, low shear stresses also increase mRNA levels whereas, low LPS levels have no effect on ET-1 mRNA levels. The decrease in ET-1 secretion caused by shear stress is at least partially regulated by the synthesis of NO. The rapid initial decrease in PGHS-1 and PGHS-2 protein levels caused by shear stress are dramatic. Short term effects of IL-1 or PMA on PGHS-1 or PGHS-2 gene expression in endothelial cells is not known. HBGF-1 significantly decreases PGHS-1 protein levels but not until at least 20 hr. The long term effect of shear stress on PGHS-1 gene expression is similar to that of IL-1 and PMA. PGHS-1 protein and
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mRNA levels are increased slightly by all three regulatory factors and the levels remain elevated for at least 24 hr. Whether or not the effect of shear stress on PGHS-2 gene expression is comparable to that of IL-1 or PMA remains unclear. When a parallel plate flow chamber was used, the long term effect of shear stress on PGHS-2 protein levels was minimal compared to that of IL-1 and PMA. When a cone and plate apparatus was used, shear stress was as effective as IL-1 at increasing PGHS-2 gene expression. One of the major differences between the two apparatuses is that the media must be gradually replenished for the cone and plate viscometer where as the media is recirculated in a parallel plate flow chamber. Since fresh media increases PGHS-2 expression, the sustained increase in PGHS-2 gene expression observed using the cone and plate viscometer may be due in part to the fresh media that is being supplied to the cells. It is evident from the data presented here that a great deal more is known about the effects of cytokines, growth factors and hormones on gene expression and how these factors enhance and inhibit one another than is known about the effects shear stress has on gene expression and how shear stress effects gene expression in the presence of nonmechanical regulatory factors. Thus for many genes, before definite conclusions can be made regarding the importance of shear stress in regulating its expression, additional experiments need to be done. A particular need is experiments that include ascertaining the effect of shear stress on gene regulation in the presence of other regulatory factors. This is particularly important since most cells in vivo live and function in the presence of both mechanical and non mechanical stimuli.
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9 Flow-induced Endothelial Cell Activation and Gene Regulation by Mechanical Forces Eugene A.Sprague1, Antonio J.Cayatte2, Robert M.Nerem3 and Sumathy Mohan1,4 Department of Radiology, University of Texas Health Science Center at San Antonio, 7703 Floyd Curl Drive, San Antonio, TX 78284–7800, USA, Tel.: (210)567–5564, Fax: (210) 567–5541, E-mail: [email protected], 4E-mail: [email protected]. 2Vascular Biology, Department of Medicine, Boston University, 80 Concord Street, Boston, MA, USA, Tel.: (617) 638–7156, Fax: (210) 638–7263, E-mail: [email protected], 3Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, GA 30332–0363, USA, Tel.: (404) 894–2768, Fax: (404) 894–2291, E-mail: [email protected]. 1
This chapter examines the concept that flow patterns along the surface of the endothelium, like humoral mediators, can act either to enhance the typical antithrombogenic, tight junction endothelial phenotype or “activate” the endothelium in a manner analogous to the inflammatory cytokines. Moreover, this chapter puts forth the concept that the vascular endothelium exhibits a nonlinear response to fluid-imposed shear stress characterized by activation of vascular endothelial cells at low shear levels (0.5–4 dynes/cm2) relative to cells exposed to either no shear or shear levels exceeding 4 dynes/cm2. Evidence supporting the stimulatory influence of low shear stress on monocyte-endothelial interaction and expression of MCP-1 and VCAM-1 genes potentially involved in the recruitment and adhesion of blood monocytes to the endothelium is reviewed. The potential influence of low shear in mediating enhanced permeability of the arterial endothelium observed within arterial sites exposed to chronic low shear, reversing flow patterns is also discussed. Though much of the signal transduction pathway involved in transduction of the low shear signal into endothelial responses remains to be defined, evidence is presented indicating that longterm activation of the nuclear transcription factor, NF-B, is observed in cultured human aortic endothelial cells exposed to prolonged low shear stress and that this pattern of response parallels that of enhanced VCAM-1 and MCP-1 gene expression. In contrast, the influence of higher shear stress levels (12–15 dynes/ cm2) on endothelial cells to promote traits associated with a “healthy” endothelium are compared. Finally, the possible implications of low shear stress flow environments with regards to atherogenesis and restenosis are considered. KEYWORDS: Shear stress, cultured endothelial cells, transcription factors, MCP-1, VCAM1, atherogenesis.
INTRODUCTION The vascular endothelium now is well recognized as a tissue that acts as an exquisitely responsive sensor to its biochemical and mechanical environment and then transduces 189
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those stimuli to modulate both its own physiology as well as the physiology of the underlying vascular wall (Davies, 1995). Even the classic concept of the endothelium as a nonthrombogenic lining impermeable to macromolecules depends and is modulated by the nature and level of these environmental factors. The “normal”, physiologic endothelium maintains its nonthrombogenic and macromolecule barrier function primarily due to the predominant environment in which it dwells. Thus, the optimal blend of biochemicals in the blood plasma along with an active mechanical environment involving flow, stretch and pressure contributes to a healthy endothelium containing macromolecule impermeable, intercellular tight junctions, which still cannot be completely replicated in cell culture. Furthermore, the arterial endothelium exposed to blood flow-associated shear stress of 15–20 dynes/cm2 is known to dramatically increase its antithrombogenic nature through the synthesis and release of a variety of agents, including nitric oxide (NO) (Uematsu et al., 1995), prostacyclin (PGI2) (Frangos et al., 1985; Grabowski et al., 1985), and the fibinolytic agent, tissue plasminogen activator (t-PA) (Diamond et al., 1989). Though the vascular endothelium does typically function as an antithrombogenic barrier to macromolecules, alterations in its environment are capable of converting the endothelium to an “activated” state characterized by a change to a procoagulant, highly permeable tissue that has high affinity for leukocyte adhesion and transmigration (Pober et al., 1987). The most well documented example of this dramatic switch in tissue function is observed in response to inflammation. Specifically, endotoxins and cytokines such as interleukins and tissue necrosis factor-a and released at inflammatory sites induce the adjacent endothelial cells to express specific leukocyte adhesion molecules as well as molecules mediating the transmigration of these cells across the endothelium. For example, endothelial cells exposed to endotoxins or inflammatory cytokines exhibit enhanced gene and protein expression for the chemoattractant, monocyte chemotactic protein-1 (MCP-1), as well as the leukocyte adhesion molecules, E-selectin and vascular cell adhesion molecule-1 (VCAM-1) (Bevilacqua et al., 1985; Gamble et al., 1985; Pober et al., 1986; Bevilacqua et al., 1989; Springer, 1994). The objective of this chapter is to examine the concept that flow patterns along the surface of the endothelium, like humoral biochemical mediators, can act either to enhance the antithrombogenic, tight junction endothelial phenotype or to “activate” the endothelium in a manner analogous to the inflammatory cytokines. Moreover, this chapter puts forth the concept that the vascular endothelium exhibits a nonlinear response to fluid-imposed shear stress characterized by activation of vascular endothelial cells at low shear levels (0.5–4 dynes/cm2) relative to cells exposed to either no shear or shear levels exceeding 4 dynes/cm2. By way of contrast the effect of shear levels greater than 5 dynes/cm2 to promote endothelial antithrombogenicity, inhibition of leukocyte adhesion, and enhanced intercellular junctions will be reviewed.
Low Shear Stress Blood Flow as an Activating Factor of Vascular Endothelial Cells In vivo EC reside in a flow environment which may be characterized by the value of the local shear rate or shear stress, where some localized regions correspond to a low,
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oscillating shear and others to a high shear. Specifically, endothelial cells lining the straight segments of the arterial tree are exposed to a relatively high laminar flowassociated shear stress of 15–20 dynes/cm2 whereas endothelial cells residing in the outer aspects of arteries just distal to arterial branch points or in arterial wall areas opposite major branch sites are exposed to low oscillating or reversing laminar shear stresses of 0.5–2 dynes/cm2 (Ku et al., 1985; Karino et al., 1988). Over the last several years, many investigators have demonstrated that these arterial sites chronically exposed to oscillating low shear stress are coincident with arterial sites known to be prone to develop atherosclerotic lesions. Previous in vivo studies have demonstrated these atherosclerotic lesion-prone areas exhibit all the hallmarks of early atherogenesis such as enhanced endothelial permeability to proteins including lipoproteins (Bell et al., 1974; Gerrity et al., 1977), enhanced turnover of endothelial cells (Caplan and Schwartz, 1973), and an increased focal recruitment of blood monocytes (Gerrity et al., 1979). It is immediately obvious that these traits of endothelial cells residing in lesion prone sites parallel characteristics of activated endothelium at sites of inflammation. This parallel raises the possibility that the altered hemodynamic flow pattern of oscillating, low shear stress may play a key role in the development and expression of the lesion prone traits described above. The following discussion examines the studies available addressing the relationship of low shear to many of these endothelial cell activation traits.
Influence of Low Shear Stress on Genes Mediating Monocyte-Endothelial Cell Interaction The possibility that specific hemodynamic flow patterns may similarly alter endothelial functions is suggested not only by the enhanced monocyte recruitment and adhesion observed in lesion-prone arterial areas but also by a report demonstrating enhanced monocyte adhesion in vivo within a carotid artery surgically modified to create a low shear environment (Walpola et al., 1995). Furthermore, in this study, Walpola et al. observed an increase in both inducible cell adhesion molecule-1 (ICAM-1) and VCAM1 on the surface of arterial endothelial cells residing within the site exposed to low shear stress. As important as such in vivo studies have been, they suffer from an inability to define either the hemodynamic environment or the cellular responses to that environment. This is particularly true of experiments where the biologic endpoints represent changes taking place over a period of weeks. Thus, for such experiments one can only note comparative differences between regions qualitatively characterized as HS and LS. Furthermore, one cannot say whether the effect noted is truly due to wall shear stress, or related to some other feature of the hemodynamic environment such as differences in particle residence time. To probe the role of low shear stress in modulating key endothelial cellular and molecular mechanisms potentially involved in mediating in vivo events such as increased endothelial permeability, cell turnover, and recruitment of the blood monocytes potentially involved as early events in atherogenesis, our laboratory, along with many others, has developed in vitro flow systems incorporating cultured endothelial cells. Using a parallel plate flow chamber system to expose cultured bovine aortic endothelial cells (BAEC) to prolonged defined laminar flow shear stress levels, studies
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in this laboratory were designed to test the hypothesis that aortic endothelial cells conditioned to prolonged low shear would exhibit increased recruitment and adhesion of blood monocytes through the increased gene and protein expression of MCP-1 and VCAM-1. As a corollary to this concept, this altered monocyte recruitment would be further enhanced in vivo by prolonged residence time of monocytes trapped in the low shear recirculation patterns characteristic of lesion-prone arterial sites. Initial studies involved exposing BAEC to either low (0.5 dynes/cm2) or high (30 dynes/cm2) shear stress for 24 h prior to assaying subsequent adhesion of radio-labeled monocytes in a 30 min. static adhesion assay (Sprague et al., 1992). The circulating medium was identical to the culture medium (10% serum in Dulbeccos Modified Eagles Medium) with temperature maintained at 37°C and pH at 7.4. As illustrated in Figure 9.1, prolonged exposure of BAEC to low shear stress (24 hours) was associated with a significant increase in monocyte adherence (162.5±10.69%, p<0.01, n=9), relative to static cultured cells. Monocyte adherence to BAEC exposed to prolonged high shear stress was also significantly lower relative to cells exposed to LS and was not significantly elevated relative to that observed to cells exposed to no shear. To define the time course for the onset of this endothelial cell response to low shear stress, monocyte adhesion studies were performed after increasing periods of shear stress exposure. Endothelialized substrates were preconditioned in parallel to high
Figure 9.1. Monocyte adherence to BAEC preconditioned to different levels of shear stress for 24 h. Confluent BAEC residing on polyester substrates were preconditioned to either HS (30 dynes/cm2) or LS (0.5 dynes/cm2) for 24 h prior to removal and placement into multiwell manifolds for subsequent 51Cr-monocyte adherence studies. At the end of the monocyte incubation period, and thorough rinsing, each endothelialized well was excised and counted for radioactivity to quantitate adherent monocytes. LS results (# monocytes adherent/cm2) were normalized to HS values (100%) to allow comparison among individual experiments. Error bars represent SEM* and data compared (LS versus NS) using student’s paired-t analysis, n=9, p<.01.
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and low shear stress levels for increasing times ranging from 30 minutes to 6 hours prior to measuring monocyte adhesion. Figure 9.2 depicts the data from an individual time course experiment. Note that a significant difference between replicate LS and HS measurements was observed only after a minimum of 6 hours of exposure to the respective shear stress regimens. When data were compared among all of the time course experiments, this same time course was observed with significant increases (168.43±23.03%, p<.05, n=3) in monocyte adherence to low shear preconditioned BAEC relative to HS exposed cells at the 6 h time period. Although HS preconditioned cells also exhibited a small relative increase in monocyte adherence at 6 h relative to the shorter duration shear exposures, this difference failed to reach significance when compared among the time course experiments. To determine whether shear stress associated changes in monocyte adherence and chemotaxis might be paralleled by changes in MCP-1 and/or VCAM-1 gene expression within LS conditioned BAEC, cells exposed to either low or high shear stress for increasing times prior to extraction of total mRNA for northern blot analysis. As illustrated in Figure 9.3, MCP-1 and VCAM-1 levels were enhanced in BAEC within 3 hours after initiation of low shear stress exposure, relative to cells exposed to either high or no shear stress in BAEC. By densitometry, levels of both MCP-1 and VCAM-1 mRNA in LS treated cells were increased approximately 2-fold at 3 h relative to high shear stress exposed BAEC. Though a low level of VCAM-1 mRNA expression is often observed in static cultured NS cells (Figure 9.3), NS cells did not exhibit any consistent pattern of either VCAM or MCP-1 mRNA. As a positive control for both MCP-1 and VCAM-1 gene expression, an additional set of confluent BAEC residing on polyester was maximally stimulated
Figure 9.2. Time dependent increase in monocyte adherence to BAEC in response to exposure to low shear stress. Confluent BAEC were preconditioned to either LS (0.5 dynes/cm2) or HS (30 dynes/cm2) for increasing times prior to removal and placement into multiwell manifolds for subsequent 51Cr-monocyte adherence studies. Each data point represents the mean of 8 replicate measurements ±SEM. Data compared (LS versus HS) using student’s paired-t analysis, p<.005.
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Figure 9.3. Time-dependent expression of MCP-1 and VCAM-1 mRNA in BAEC exposed to different flow conditions. Top panel: Northern blot analysis of mRNA isolated from BAEC exposed to either high (HS, 30 dynes/cm2), low (LS, 0.5 dynes/cm2) or static, no shear (NS) stress flow conditions for times ranging from 1–3h. To provide a positive control, a separate set of confluent BAEC was exposed to lipopolysaccharide (10ng/ml) for 4h prior to RNA isolation and analysis by northern blot. BAEC expression of GAPD mRNA in the same cells is shown for comparison. Middle and bottom panels: Relative mean density measurements of the autoradiograms are presented for VCAM-1 and MCP-1 expression. These measurements were performed using an Apple Scanner combined with image analysis using the IMAGE program on a McIntosh II ci computer.
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with lipopolysaccharide (LPS) at a concentration of 10 ng/ml for 4h under static culture conditions at 37° (Lane 2, Figure 9.3). Note that a transient elevation in MCP-1 gene expression was measured in BAEC exposed to 2h high shear flow. This is consistent with previously reported observations by (Shyy et al., 1994). Not only was expression of these two genes elevated at 3 h but also, as shown in Figure 9.4, MCP-1 and VCAM-1 gene expression remained elevated after 51h exposure to LS. At this physiologic more relevant time period, note that both genes are completely downregulated in BAEC conditioned to high shear stress. No detectable levels of VCAM-1 mRNA expression were ever found in cells exposed to prolonged HS (>6 h), indicating a possible inhibitory influence of longterm HS exposure. This inhibitory effect of shear stress levels greater than 5 dynes/cm2 has now been well defined by several investigators. To provide evidence that the observed increase in gene expression correlated with expression of protein, additional experiments were performed to determine whether conditioned media obtained from BAEC subjected to flow regimens contained monocyte chemoattractant activity. As demonstrated in Figure 9.5, only media obtained from BAEC preconditioned to low shear for 24 h or incubated with LPS (10 ng/ml, 4h) exhibited monocyte chemoattractant activity significantly above (P<.001) the basal level seen in control media. To provide indirect evidence that VCAM-1 molecules expressed on the surface of LS conditioned cells might be involved in the observed increased monocyte adherence; a blocking antibody to human VLA-4 (clone P4G9, Life Technologies, Inc., Gaithersburg, MD), the monocyte surface integrin which acts as a ligand to VCAM-1, was employed. Pretreatment of
Figure 9.4. Relative MCP-1 and VCAM-1 gene expression in BAEC exposed to prolonged flow conditions. Northern blot analysis of mRNA isolated from BAEC exposed to either high (HS, 30 dynes/ cm2), low (LS, 0.5 dynes/cm2) or static, no shear (NS) stress flow conditions for 51h. Expression of GAPDH mRNA in the same cells is shown for comparison. These measurements were performed using an Apple Scanner combined with image analysis using the IMAGE program on a McIntosh II ci computer.
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Figure 9.5. Monocyte chemoattractant activity accumulated in media of BAEC exposed to prolonged (24h) flow conditions. Confluent BAEC were preconditioned to either LS (0.5 dynes/ cm2) or H S (30 dynes/ cm2) or no shear, static culture for 48h prior to collection of the circulating or culture media for chemotaxis assay. Flow conditioned, fresh control media (no cell exposure), or media obtained from BAEC maximally stimulated with lipopolysaccharide (10 ng/ml) was placed in the bottom well of a Boyden chamber. Freshly isolated human monocytes were added to the top well with a 5 pore polycarbonate filter separating the wells. Monocyte migration across the filter was measured by removing and Giemsa staining the filter and counting the cells in each of 5 light microscopic fields (400x magnification). These measurements were then averaged for each experimental data point (monocytes/hpf). *Data compared using student’s paired-t analysis, n=3, p<.001.
monocytes for 30 min. at 37° with anti-VLA-4 reduced monocyte adherence to LS preconditioned BAEC by 82±9 % but had no significant effect on monocyte adherence to either HS or NS conditioned cells. More recent studies employing an anti-VCAM-1 monoclonal antibody on human aortic endothelial cells (HAEC) have now confirmed increased VCAM-1 antigen on cells exposed to low shear stress for 6 h or longer. These results provide a plausible scenario for prolonged exposure of low shear stress to contribute to the sustained MCP-1 and VCAM-1 expression at human atherosclerotic lesion sites in addition to the contribution to that expression associated with macrophage-associated cytokines and oxidatively modified LDL. In this scenario, induction of MCP-1 and VCAM-1 on arterial endothelial cells in regions of low shear would stimulate the recruitment and adhesion of recirculating monocytes to the endothelial surface. These monocytes would then, in turn, migrate across the endothelium into the intima to transform into macrophages. Expression of cytokines by these macrophages could then further enhance activation of the overlying endothelium. If concomitant to these events, these low shear stress exposed arterial
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sites also exhibited enhanced permeability to macromolecules including LDL, then, particularly in the presence of hypercholesterolemia, the stage is set for interaction of all the key elements of atherogenesis; all of which are may contribute to endothelial activation.
Influence on Permeability of Vascular Endothelium Using the pig model, early studies demonstrated that low shear, lesion-prone arterial regions, as identified by the incorporation of Evans blue dye, exhibited an endothelium with a thinner glycocalyx as well as enhanced accumulation of albumin, fibrinogen, and LDL in comparison to the non-lesion prone, white or unstained regions, and increased monocyte recruitment (Bell et al., 1972; Bell, Gallus et al., 1974; Caplan et al., 1974; Gerrity, Naito et al., 1979). More recent evidence indicates that the decline in the macromolecule barrier function of the endothelium at low shear, lesion-prone sites is most likely linked to increased endothelial cell turnover in these areas. Specifically, Chen et al. (1995) have recently reported evidence demonstrating that high levels of endothelial turnover are associated with an enhanced ability of macromolecules to cross the endothelium via increased junctional gaps among cells in these areas (Chen et al., 1995). Accumulating evidence now indicates that the hemodynamic environment and elevated LDL levels likely act together to induce this dysfunctional state of the endothelium at these sites (Davies et al., 1995). As stated above, we postulate that the low shear component of the flow environment may simultaneously influence facets of endothelial cell biology that could enhance endothelial permeability as well as “activate” the endothelium to promote monocyte recruitment. In previous in vitro flow studies, we have demonstrated that endothelial cell proliferation rate, as measured by both cell number and thymidine uptake analyses, decreases with exposure to increasing shear stress levels (Levesque et al., 1990). Recently completed studies in this laboratory on the influence of prolonged low shear on an endothelial junctional protein, endothelial cell adhesion molecule (EndoCAM, CD31 or PECAM-1) may also provide insight as to how the local hemodynamic environment could affect endothelial physiology to enhance permeability of the endothelium (Sprague and Mowery, 1995). Specifically, our studies indicate that exposure of bovine aortic endothelial cells to prolonged steady low shear (0.5 dynes/ cm2) was associated with downregulation within 3 hr from the onset of flow of PECAM-1 gene expression as well as cell surface protein expression relative to static cultured cells or cells exposed to high shear stress (30 dynes/cm2). Interestingly, PECAM1 expression was significantly reduced again in cells exposed to a pulsatile reversing low shear flow regimen (2±4 dynes/cm2) relative to the level observed in endothelial cells exposed to a steady low shear stress of 2 dynes/cm2). In addition, this effect was maintained in cells exposed to flow for up to 96 hr. Whether a decrease in PECAM-1 present within endothelial cell to cell junctions might influence either endothelial cell turnover or passage of macromolecules between endothelial cells remains to be determined. Lending some support to this concept is a recent report by Stewart et al. (1996) indicating that a similar decrease in EndoCAM mRNA expression accompanied by a decrease in cell surface is also observed in human or bovine endothelial cells
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treated with the cytokines, TNF- or IFN-, which also act to increase endothelial permeability.
What are the Key Cellular Signaling Pathways Mediating the Influence of Low Shear Stress? Although insight into the cellular signaling molecules involved in the early response of endothelial cells to the onset of physiologic levels of arterial shear in the range of those in the straight arterial segments (12 dynes/cm2 or greater) has increased greatly in the past few years, we are only beginning to understand some of the key signals that participate in the response of the endothelial cell to low shear stress. The likely importance of the integrins and their associated intercellular molecules in the recognition and transduction of the high shear onset stimulus signal is thoroughly discussed in an excellent review of the subject by Berk et al. in a preceding chapter in this book. Other proposed possible shear stress recognition sites on the vascular endothelial cell include plasmalemmal potassium or chloride channels (Olesen et al., 1988), G-proteins (Gudi et al., 1996), or some yet to de defined specific shear receptor complex. At this time there is no definitive evidence whether any one or more of these are involved in the recognition of the low shear stimulus. Our efforts to begin to probe the mechanisms involved in transducing the low shear stimulus to gene activation have focused on examining the nuclear transcription factor, NFB. The basis for interest in this particular transcription factor lies in the fact that the promoter sequences for both MCP-1 and VCAM-1 contain NFB binding sites. In these studies, cultured human aortic endothelial cells (HAEC) were exposed to either steady flow low shear (2 dynes/cm2), oscillating flow low shear (2±2 dynes/cm2), steady flow high shear (15 dynes/cm2) or no shear, static culture control conditions for periods ranging from 30 min. to 24 h. using the parallel plate flow chamber system described above (Mohan et al., 1997). As illustrated in Figure 9.6, HAEC conditioned to high shear stress exhibited a biphasic response in NFB DNA binding activity, as measured by the electrophoretic mobility shift assay, characterized by an early, transient increase in activity sustained between 30 min. and 2h high shear exposure followed by a prolonged downregulation of NFB activity throughout the longer periods of high shear exposure up to 24 h, though a small secondary increase was consistently observed at the 16 h. time period. In contrast, HAEC exposed to either steady or pulsatile low shear stress exhibited significant, sustained elevations in NFB activity beginning at 30 min. and reaching a maximum plateau level at 16 h which was maintained throughout 24 h. Note, also that NFB activity in cells exposed to oscillating shear was consistently further augmented relative to cells exposed to steady low shear. This raises the possibility that this pattern of activation may be regulated both by the absolute shear level as well as the nature of the pulsatile flow pattern. Again, as described above for the responses of MCP-1, VCAM-1, and PECAM-1 to prolonged low shear stress levels, a nonlinear response of NFB activity to shear level is observed as demonstrated by a minimal activity present in static, no shear exposed cells, a sustained rise in activity at low shear exposure levels, and a pronounced downregulation under prolonged high shear levels. Not only does the general pattern of NFB activity follow that observed
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Figure 9.6. Kinetics of NF-B DNA binding activity in human aortic endothelial cells (HAEC) exposed to different flow regimens. HAEC were incubated in 2% serum containing media without growth factors prior to exposure to different shear stress levels for the indicated periods of time. Nuclear proteins were extracted and electrophoretic mobility shift assay analyses were performed. Lanes 1–4, NF-B activity in cells exposed to no, 15, 2, or 2±2 dynes/cm2, respectively. Each experiment was repeated independently at least 3 times. Arrows designate specific binding of NF- B. Below: Densitometric analyses of autoradiograms in top panel as analyzed using the NIH 1.58b19 image analysis software.
for MCP-1 and VCAM-1 gene expression but also the time course for activation of this transcription factor provides a logical basis for its possible involvement in mediating the low shear effects on expression of these two genes. Also, the early transient increase in NFB activation observed in our studies parallels results reported by Lan et al. (1994) indicating a maximal NFB activation response at 60 min. in human umbilical vein endothelial cells exposed to a shear stress of 12 dynes/cm2 over a 2 h. time course.
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This pattern of an early transient response to high shear followed by a prolonged downregulation has obvious similarities to the early increase reported for MCP-1 gene expression in endothelial cells exposed to the same level of high shear stress. Though the relationship of NFB activation to MCP-1 and VCAM-1 gene expression remains circumstantial, ongoing studies using inhibitors of NFB activation as well as cells transfected to overexpress the natural intracellular inhibitory factor, IB, should provide further insight to this possible linkage.
THE OPTIMAL FLOW ENVIRONMENT FOR THE ARTERIAL ENDOTHELIAL CELL Though the primary theme of this chapter is the activation of the endothelial cell by low shear stress hemodynamic flow patterns, the logical corollary to this theme is that vascular endothelial cells exposed to the more common arterial levels of higher shear stress (15–20 dynes/cm2) would not exhibit the hallmark traits of activated endothelial cells but would instead exhibit opposing characteristics. If hemodynamics and shear stress levels, in particular, play an important role in modulating the physiology and pathophysiology of the vascular endothelium, then one would expect that vascular endothelium exposed to the higher shear stress levels would exhibit enhanced antithrombogenic activity, impermeability to macromolecules, low cell turnover, and no expression of molecules involved in the recruitment and adhesion of circulating leukocytes. As is well known, this is the composite traits of a healthy, optimally functioning vascular endothelium. Though much of this information has been extensively reviewed in this book as well as elsewhere, this chapter will briefly survey the overall picture that is now coming together from many in vitro flow studies as well as in vivo reports.
Physiologic High Shear Upregulates Genes Responsible for a “Healthy” Arterial Endothelium A large number of laboratories have contributed to the development of the current state of knowledge in this area. Taken together these studies show that, for a confluent monolayer of cultured EC, the influence of an elevated shear stress is to cause: (i) an elongation in shape and an orientation of the cell’s major axis with the direction of flow (Dewey et al., 1981; Eskin et al., 1984; Levesque and Nerem, 1985); (ii) a rearrangement of the actin microfilaments into stress fibers aligned with the direction of flow, accompanied by a concomitant increase in cell stiffness (Sato et al., 1990); (iii) an influence on endocytotic processes, e.g. the enhancement of the receptormediated binding, internalization, and degradation of LDL (Sprague et al., 1987) and (iv) the increased expression of anti—thrombotic agents including PGI2, tPA, and thrombomodulin, not only at the product level but also at the gene expression level (Frangos, Eskin et al., 1985; Grabowski, Jaffe et al., 1985; Diamond, Eskin et al., 1989; Malek et al., 1994). It is noteworthy that these responses such as prostacyclin to elevated shear stress involve prolonged, sustained upregulated gene and protein
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expression that would be expected to be necessary to maintain critical ongoing endothelial functions in vivo. A similar sustained gene upregulation in response to elevated shear has also been reported for the nitric oxide synthase gene in a number of separate investigations (Kuchan et al., 1994; Berk et al., 1995; Ranjan et al., 1995; Uematsu, Ohara et al., 1995). It is well documented that flow modulated increases in endothelial nitric oxide play a critical role in such functions as vasodilation (Cohen, 1995), antithrombogenicity (Stamler et al., 1989; Irokawa et al., 1997), and suppression of underlying arterial wall smooth muscle cell proliferation (De Meyer et al., 1995; Groves et al., 1995; Chen et al., 1997). In addition, recent reports indicate that proteins such as PECAM-1 involved in endothelial cell to cell junctions may be upregulated in the presence of higher arterial shear stress levels. Specifically, PEC AM-1 gene expression levels as well as its activation through tyrosine phosphorylation appear to be enhanced upon exposure to shear stress (Osawa et al., 1997).
Flow Downregulation of Genes Potentially Involved in Arterial Endothelial Pathophysiology Extensive evidence now exists demonstrating that cultured vascular endothelial cells exposed to in vitro arterial-like shear stress levels from 12–15 dynes/cm2 exhibit downregulation of the VCAM-1 gene known to be associated with enhanced leukocyte adhesion. This pronounced downregulating effect of elevated shear has been observed in endothelial cells initially activated with inflammatory cytokines to up-regulate the VCAM-1 gene prior to exposure to shear (Sampath et al., 1995; Tsao et al., 1996) as well as in mouse endothelial cells isolated from lymphoid tissue that naturally exhibit high VCAM-1 expression without added cytokine stimulation (Ando et al., 1995). In this latter set of studies the extent of VCAM-1 down-regulation was directly related to increasing shear stress levels. As mentioned above, we have reported a similar pattern of downregulation in the DNA binding activity of NFB in human aortic cells exposed to prolonged shear stress levels of 15 dynes/cm2 (Mohan et al., 1997). Similarly, the MCP-1 gene appears to similarly downregulate in response to prolonged levels of elevated shear. In spite of the well-documented transient increase observed within the first 2 h. after the onset of applied in vitro shear stress (Shyy et al., 1994), endothelial cells exposed to prolonged elevated shear stress exhibit a prolonged downregulation of the MCP-1 gene that is sustained over time (Sprague et al., 1992).
IMPLICATIONS FOR ATHEROGENESIS AND RESTENOSIS Based upon the preceding discussion, the implications of exposure of endothelium to patterns of low shear, reversing flow with regards to atherogenesis are readily apparent. Both prolonged in vitro exposure of cultured endothelial cells to prolonged low shear stress flow regimens in the range of 0.5–2 dynes/cm2 (Sprague et al., 1992) as well as chronic exposure of arterial endothelium to a surgically created low shear environment in vivo (Walpola et al., 1993; Walpola et al., 1995) are associated with enhanced
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recruitment and adhesion of blood monocytes to the endothelial surface. Furthermore, this recruitment is facilitated by demonstrated increases in the cell surface monocyte adhesion molecule, VCAM-1, gene and protein expression both in vitro and in vivo. In vitro studies have shown that the monocyte chemoattractant, MCP-1, gene is also upregulated by prolonged in vitro flow exposure (Sprague et al., 1992) and transcripts have also been identified at atherosclerotic lesion sites (Li et al., 1993), though this could come from a number of cell types including macrophages or smooth muscle cells. As monocytes transmigrate across the arterial endothelium they begin to transform into macrophages as they enter the subendothelial space similar to the process observed when monocytes enter any other tissue. This represents one of the first key elements involved in atherogenesis (Schwartz et al., 1991). Secondly, the enhanced permeability associated with arterial areas chronically exposed to low shear reversing flow patterns along with the prolonged residence time of both blood borne cells and macromolecules in this type of flow pattern, opens the possibility for LDL, particularly when it is present in high concentrations in the plasma, to also cross the endothelium and begin to accumulate in the subendothelial space of the arterial intima. Much evidence is available indicating that LDL is oxidatively modified as it enters the arterial wall (Quinn et al., 1987). This oxidation of LDL is likely the result of attack of the molecule by reactive oxygen species and free radicals released primarily by the monocyte-derived intimal macrophages as well as some contribution from dysfunctional endothelial cells and intimal smooth muscle cells (Yla-Herttuala et al., 1994; Steinberg, 1995). Since the maturing monocyte-derived macrophage expresses increasing numbers of scavenger receptors that have a high affinity for oxidized LDL, these two transformations then set the stage for the first definable atherogenic event, formation of the foam cell or lipid-filled macrophage as a result of unregulated uptake of modified LDL. The formation of oxidized LDL not only serves to stimulate its uptake by monocyte-derived macrophages but also acts to further “injure” or activate the endothelium to enhance the synthesis and secretion of MCP-1 to recruit additional monocytes to the arterial wall (Berliner et al., 1993). In addition to its effects on macrophages and endothelial cells, oxidized LDL has been demonstrated to stimulate smooth muscle cells to secrete MCP-1, providing a chemotactic gradient to direct endothelial adherent monocyte migration into the arterial wall (Cushing et al., 1990; Yu et al., 1992). Obviously, then, the process of atherogenesis involves complex interactions among several factors, many of which can activate the arterial endothelium. The hemodynamic flow patterns in lesion prone arterial areas may be one of these factors. Restenosis or the reclosing of the arterial lumen at a site of vascular intervention performed to remove an atherosclerotic obstructive lesion is an increasing area of concern and investigation as the number and different types of interventional procedures rapidly increase. Since reopening a lesion-blocked artery necessarily involves an initial arterial injury either by balloon angioplasty, vascular bypass, atherectomy, or stent placement, one of the key issues involved in restenosis is the response of that injured arterial site to the imposed injury. The rate and extent of reendothelialization at these sites is one of the critical factors in limiting the extent of response of the arterial wall to injury. Previous studies indicate that relatively small injuries heal rapidly and completely (Lindner et al., 1989), whereas
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larger areas of endothelial loss may never completely be reendothelialized, though the initial phase of that partial endothelial restoration seems to occur fairly rapidly (Clowes et al., 1986). The overwhelming evidence derived from animal arterial injury models including the rat (Haudenschild and Schwartz, 1979), rabbit (Reidy, 1988), pig (Steele et al., 1985) and primate (Clowes et al., 1985) animal models indicates that the primary source of the endothelial cells responding to an acute injury to the arterial wall is the adjacent intact, healthy endothelium. In a quantitatve analysis of this reendothelialization response in vivo using the rat carotid model, Schwartz et al. (1978) employed a balloon catheter to create a defined injury and then examined the endothelial response using light and scanning electron microscopy along with thymidine autoradiography to identify areas of cell proliferation. These investigators observed that 1) coverage of the deendothelialized area occurred primarily by way of migration of adjacent endothelial cells, 2) after migration had already occurred a zone of proliferation was initiated removed by approximately 100 cells from the leading edge of migration, and 3) migration occurred along the axis of the vessel indicating that migration was influenced by blood flow. Several studies performed in vitro employing defined wounds to cultured endothelial cell monolayers have also now clearly demonstrated that the initial repair of the injured area occurs by way of migration rather than by cell proliferation (Sholley et al., 1977; Wall et al., 1978; Madri and Stenn, 1982; Schleef and Birdwell, 1982). Employing a parallel plate flow system to examine endothelial cell migration under different flow-related shear stress conditions, recently completed studies in this laboratory have demonstrated under arterial-like high shear stress flow conditions (15 dynes/cm2) that migration of endothelial cells from a confluent endothelialized surface onto a freshly implanted piece of stainless steel stent material occurs primarily along with the direction of flow, though some retrograde to flow migration was observed (Sprague et al., 1997). In contrast,, under low shear stress (2 dynes/ cm2) migration rate was much slower and no preferential direction of endothelial cell migration was observed. Interestingly, migration rate was increased nearly two-fold under flow conditions that we have shown previously to decrease endothelial cell proliferation rate (Levesque, 1990). In a recent in vivo study in which a defined wound was created in rabbit carotid arteries surgically modified to maintain a low flow rate and, thereby a low arterial wall shear stress; Vyalov et al. (1996) observed that endothelial repair was significantly impaired compared to a similar arterial wound created in an unaltered carotid with a higher shear stress environment. Thus, the flow conditions existing or created at an injury site could potentially greatly influence both the rate and extent of reendothelialization by migration as well as the general functional state of endothelium at or near that site.
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10 Hemodynamics and Endothelial Phenotype: New Insights into the Modulation of Vascular Gene Expression by Fluid Mechanical Stimuli James N.Topper# and Michael A.Gimbrone Jr.* Vascular Research Division, Department of Pathology, Brigham and Women’s Hospital, Harvard Medical School, 221 Longwood Ave., LMRC-401, Boston, MA 02115–5817, USA. #Corresponding author: Cardiovascular Division, Department of Medicine, Stanford University School of Medicine, Falk Cardiovascular Research Center, 300 Pasteur Drive, Stanford, CA 94305–5406, USA, Tel.: 650–725–6850, Fax: 650–725–1599, E-mail: [email protected].
*
Vascular endothelium is a dynamic interface, responsive to both local and systemic stimuli. Signals derived from blood flow are important determinants of vascular homeostasis and vascular disease pathogenesis. In vitro, fluid shear stresses can specifically and selectively regulate the expression of subsets of endothelial genes. We have employed gene discovery strategies in clutured human endothelial cells to identify a series of genes whose expression is modulated by defined fluid shear stresses in vitro. These genes fall into a variety of classes including enzymes, intracellular signaling molecules, cell-surface transporters, and receptors. A subset of these genes demonstrate a sustained upregulation in response to a steady laminar shear stress and are not upregulated by the non laminar stimulus turbulent shear stress. Interestingly, many of these genes manifest an endothelial-selective pattern of expression in vivo. These results have led us to propose that local differences in hemodynamic stimuli within the vascular tree may be responsible for regional differences in endothelial phenotype observed in vivo. Biomechanical stimuli such as shear stresses may be important determinants of endothelial phenotype in vivo.
HEMODYNAMICS AND VASCULAR ENDOTHELIAL BIOLOGY The involvement of vascular endothelium in disease processes such as atherosclerosis has been recognized for over a century, but a working knowledge of its relevant pathophysiology has been developed only recently, largely as a result of the application of modern cellular and molecular biological techniques. We now appreciate that this single-cell-thick lining of the circulatory system is, in fact, a vital organ whose health is essential to normal vascular physiology and that its functional phenotype can be dynamically regulated by various physiologic and pathophysiologic stimuli, including biochemical mediators such as inflammatory cytokines, growth factors, circulating hormones and bacterial products. 207
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Non-adaptive changes in this functional phenotype, so called “endothelial dysfunction” can be a critical factor in the pathogenesis of vascular diseases (Gimbrone, 1995; Gimbrone et al., 1995). In addition to biochemical stimuli, the endothelial lining of the cardiovascular system also is subjected to various types of biomechanical stimuli resulting from pulsatile blood flow. These stimuli include fluid shear stresses, cyclic strains and hydrostatic pressures. As the cellular layer directly in contact with blood, the endothelium in particular constantly bears the frictional forces (wall shear stress) derived from the flow of this viscous fluid. A number of in vivo observations suggest that these hemodynamic forces play a central role in vascular homeostasis and pathophysiology. For example, blood flow is an important modulator of vascular tone, acutely, and also can influence more chronic processes of vascular remodeling in both health and disease (Glagov et al., 1988; Langille and O’Donnell, 1986). Experimentally, many of these effects appear to result, at least in part, from the ability of the vascular endothelium to “sense” and transduce hemodynamic stimuli into changes in endothelial structure and function. Examples include the demonstration of increased macromolecular permeability, lipoprotein accumulation, endothelial cell damage and repair, leukocyte adhesion molecule expression and mononuclear leukocyte recruitment near branch points and bifurcations, i.e., in areas of complex, non-uniform disturbed laminar flows, as well as the topographical mapping of ellipsoidal endothelial cell (and nuclear) shape and axial alignment (in the flow direction) to laminar flow regions and the disruption of this orderly pattern in regions of disturbed flow. In addition, experimental alterations of vascular architecture (e.g., surgical coarctation or shunts) result in both acute and chronic vessel wall changes that appear to be, at least in part, endothelium dependent, and in the presence of hypercholesterolemia can result in lesions resembling atherosclerosis (Glagov et al., 1988; Langille and O’Donnell, 1986; Davies and Tripathi, 1993; Gimbrone, 1995). Taken together, these in vivo observations are consistent with either direct or indirect effects of one or more hemodynamic stimuli on endothelial function/dysfunction. Evidence of the direct action of specific hemodynamic forces on endothelial structure and function has come from in vitro studies in which cultured monolayers of endothelial cells (human and animal) have been subjected to defined fluid mechanical forces under controlled experimental conditions. Utilizing experimental apparatuses that can reproducibly generate defined flows in vitro (Bussolari et al., 1982; Sdougos et al., 1984), a variety of cell biological effects can be observed in endothelial monolayers in response to applied shear stresses. These include reorganization of actin-containing stress fibers as well as other cytoskeletal components, cell cycle effects, alterations in the metabolic and synthetic activities of the endothelial cells including the production of arachidonate metabolites (in particular, prostacyclin), growth factors (e.g., platelet-derived growth factor, PDGF), coagulation and fibrinolytic components, and vasoactive substances (Davies, 1995; Davies and Tripathi, 1993; Gimbrone et al., 1995). Certain of these more rapid shear-induced changes appear to involve regulation at the level of rate-limiting enzymes and/or substrate availability (e.g., arachidonic acid release by calciumsensitive phospholipases, NO production by nitric oxide synthase); however,
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especially in the case of delayed responses in which de novo protein synthesis is occurring, upregulation of gene expression appears to be occurring as a consequence of exposure to fluid mechanical forces (Davies, 1995; Gimbrone et al., 1997; Resnick and Gimbrone, 1995).
PATTERNS OF GENE EXPRESSION IN CULTURED ENDOTHELIAL CELLS IN RESPONSE TO HUMORAL AND BIOMECHANICAL STIMULI The observation that defined fluid mechanical forces could specifically influence endothelial gene expression has lead to the rapid elucidation of a growing, diverse set of endothelial genes, whose expression can be modulated by fluid shear stress in vitro. In the majority of cases these studies have involved the detailed investigation of relevant “candidate genes” (i.e., genes that had been discovered and/or studied previously in other contexts), and have utilized primarily uniform (e.g., steady laminar) flow conditions in vitro. These experimental approaches (which are reviewed in more detail in other chapters of this volume) have led to the appreciation that many of these genes are being regulated at the transcriptional level, a process involving definable, and in some cases discrete, cis-acting regulatory elements in the promoters of these genes, and the trans-acting protein factors that interact with these elements. Initially, our laboratory focused on the analysis of the promoter element(s) responsible for the transient upregulation of the PDGF-B gene in response to applied steady laminar shear stress (Resnick et al., 1993). These studies identified a specific 6 nucleotide motif, which was termed the shear stress response element (SSRE) that was both necessary and sufficient for the shearmediated upregulation of the PDGF-B promoter in vitro. This novel element, which is capable of interacting with the transcription factor NF-B, is able to confer shear stress responsiveness to a heterologous (normally non-shear responsive) promotor construct in vitro (Resnick et al., 1993; Resnick and Gimbrone, 1995). Subsequently, additional SSREs have been identified in the promoters of the MCP-1, PDGF-A, TGF-ß1 and VCAM-1 genes, consisting of a variety of cis-acting elements such as AP-1 and EGR-1/SP1 sites (Shyy et al., 1995; Khachigian et al., 1997; Ohno et al, 1995; Ando et al., 1994). These studies are reviewed in more detail in other chapters in this volume (see chapters by Resnick, Ando and others). More recently, we have begun to apply complementary strategies to study this process (Topper et al., 1996). These studies involve the use of gene techniques such as differential RT-PCR display and subtraction hybridization which do not require identification of “candidate genes” a priori, and can thus potentially yield unanticipated and unbiased insights into the patterns (e.g., the magnitude and complexity) of the endothelial response to defined fluid mechanical forces. In this chapter, we will summarize some of the results to date of these ongoing efforts, and discuss how our findings have provided some new insights into the role of hemo-dynamic forces in vascular pathobiology.
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IDENTIFICATION OF GENES DIFFERENTIALLY RESPONSIVE TO FLUID MECHANICAL STIMULI: THE ATHEROPROTECTIVE GENE HYPOTHESIS The observation that the early lesions of atherosclerosis are predominantly localized to regions of the arterial vasculature associated with complex, non-uniform flows (e.g., bifurcations and curvatures) while areas characterized by relatively uniform flows (in a time-averaged sense) tend to be spared; suggests that fluid mechanical stimuli are playing an important role in the atherogenic process (Cornhill and Roach, 1976; Glagov et al., 1988; Langille and O’Donnell, 1986). In collaboration with Dr. Dean Falb’s group at Millennium Pharmaceuticals, Inc. (Cambridge, MA), we have employed a high through-put differential display of mRNA transcripts (Liang and Pardee, 1992) to compare the patterns of genes that are up- (or down) regulated in human endothelial cells in response to a physiological level of steady laminar shear stress, a comparable level of turbulent (non-laminar) shear stress, and a soluble cytokine stimulus (recombinant human IL-1ß) at a maximally effective concentration. It is important to note that true turbulence is probably quite rare within the cardiovascular system, but as an in vitro fluid mechanical stimulus it represents a definably non-laminar stimulus that can be reproducibly established in vitro, and has been demonstrated to elicit distinct cell-biological responses in this experimental system (Davies et al., 1986). These studies have revealed distinctive patterns of gene expression not previously observed in endothelial cells, including a set of genes that appear to be upregulated, in a sustained fashion, by steady laminar shear stress, but not by turbulent shear stress (Figure 10.1 A). Certain of these differentially regulated transcripts reflect the upregulation of known genes, such as ecNOS (the endothelial isoform of nitric oxide synthase), COX-2 (the inducible isoform of cyclooxygenase), and Mn-SOD (manganese-dependent superoxide dismutase). Interestingly, both Mn-SOD and COX-2 had not previously been appreciated as flow-regulated species (Figure 10.1B) (Topper et al., 1996). These endothelial genes encode enzymes that can exert potent anti-thrombotic, anti-adhesive, anti-proliferative, anti-inflammatory, and anti-oxidant effects both within the endothelial lining and also interacting cells, such as platelets, leukocytes and vascular smooth muscle. Thus, the biological consequences of these steadylaminar-shear-stress upregulated endothelial genes would be predicted to be “vasoprotective” or “anti-atherogenic”. Other investigators, have independently demonstrated the upregulation of the Cu/Zn isoform of superoxide dismutase in cultured EC exposed to steady shear stress in vitro (Inoue et al., 1996). In addition, steady laminar shear stress has been demonstrated to inhibit the cytokine-induced expression of the mononuclear leukocyte adhesion molecule VCAM-1 in HUVEC, as well as down-regulating cell-surface VCAM-1 protein levels in EC that express this molecule constitutively in vitro such as murine microvascular EC (Ando et al., 1994; Marui et al., 1993). These findings have suggested a hypothesis, we have termed the “atheroprotective gene hypothesis” which can be stated as follows: The coordinated induction (or down regulation) of a subset of endothelial genes whose effects would be predicted to be anti-atherogenic, or vasoprotective, by the uniform shear stresses associated with
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Figure 10.1. Patterns of gene expression in response to fluid mechanical and cytokine stimuli in cultured endothelium. Confluent monolayers of cultured HUVEC were stimulated with steady laminar shear stress at 10 dyn/cm2 (LSS), the same time-averaged magnitude of turbulent shear stress (TSS) or 10 units/ml of recombinant human IL-1ß for varying amounts of time. (A) At least 6 distinct patterns of gene expression were observed in response to these stimuli. Two of the patterns, IV and V, are novel patterns demonstrating selective induction in response to LSS but not TSS stimulation, indicating that the endothelial cells can sense and respond distinctly to these two different fluid mechanical stimuli at the level of gene expression. (B) Examples of patterns of mRNA expression in response to LSS, TSS and IL-1ß cytokine stimulation in HUVEC. The figure shown is a semi-quantitative RT-PCR analysis of steady state message levels for the following genes identified by differential display, or chosen as controls; Mn-SOD, the manganesedependent isoform of superoxide dismutase; COX-2, the inducible isoform of cyclooxygenase (the rate limiting enzyme in prostanoid synthesis); COX-1 the major constitutive isoform of cyclooxygenase that is not regulated by fluid mechanical stimulation; ecNOS, also known as NOSIII, the endothelial isoform of nitric oxide synthase responsible for nitric oxide production; and GAPDH, a cytosolic enzyme involved in glucose metabolism. Reproduced with permission from (Topper et al., 1996).
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areas of vasculature that are typically protected from the early lesions of atherosclerosis, may represent a mechanism underlying the focal nature of atherosclerotic lesions (Topper et al., 1996). Stated another way, the differential regulation of endothelial genes by distinct fluid mechanical forces in vivo, may result in the ability of certain areas of the vasculature to more effectively resist the influence of systemic risk factors such as hyperlipidemia, and diabetes. (Figure 10.2). Although this hypothesis does not exclude a role for the disturbed flows (e.g., disturbed laminar, oscillatory or reversing flows) associated with locally complex geometries as “direct proatherogenic stimuli”, it represents an additional conceptual framework in which to approach this complex biological phenomenon. We are currently testing these hypotheses via a series of gene-targeting and overexpression studies, utilizing some of the putative atheroprotective genes recently identified, in atherosclerosis-prone strains of mice.
Figure 10.2. Schematic representation of the potential “atheroprotective” effects of steady laminar shear stress stimulation of vascular endothelium. Steady laminar shear stress (LSS) can selectively upregulate the genes encoding Mn-SOD, COX-2, and ecNOS, while down-regulating VCAM-1 expression in vascular endothelial cells. In vivo this would be predicted to have a variety of effects such as the inhibition of platelet aggregation via prostacyclin (PGI2) production, the inhibition of leukocyte adhesion and migration via PGI2 and NO (EDRF) production and a decrease in endothelial cell-surface VCAM-1 expression. Many of these effects could be potentiated by the induction of the important antioxidant enzyme, Mn-SOD within the endothelial cell, which could serve to buffer excess reactive oxygen species (ROS) generated in response to oxidant stresses such as hyperlipidemia and hyperglycemia, potentially leading to less inactivation of NO (EDRF) and less expression of proinflammatory effectors (e.g., VCAM1, MCP-1) via redox-sensitive mechanisms.
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A NEW CLASS OF SHEAR-REGULATED ENDOTHELIAL GENES: THE 12-MEMBRANE SPANNING TRANSPORTERS In addition to identifying patterns of gene expression, our gene discovery strategy provides the opportunity to identify individual novel genes that are responsive to flow. Among the genes we have identified are two members of the large superfamily of cell-surface, 12-membrane-spanning transporters that demonstrate selective responsiveness to fluid mechanical stimuli in cultured HUVEC (Topper et al., 1998; Topper et al., 1997b). The 12 membrane-spanners are a family of transporters whose members subserve diverse functions ranging from the transport of inorganic and organic ions to the regulation of macrophage activation and antimicrobial activity. In vascular endothelium, one member of this family, the electroneutral Na-K-2Cl cotransporter, is thought to function in the maintenance of a selective permeability barrier in certain vascular beds (e.g., brain), as well as in the preservation of endothelial homeostasis in the face of fluctuating osmotic conditions that may accompany certain pathophysiologic conditions (O’Donnell et al., 1995; Delpire et al., 1994). We identified BSC2 (bumetanide-sensitive cotransporter-2), one of the two major isoforms of electroneutral Na-K-2Cl cotransporters present in mammalian cells (and the major endothelial-expressed isoform), as a regulated gene selectively induced by steady laminar shear stress but not a turbulent shear stress stimulus in cultured HUVEC (Figure 10.3) (Topper et al., 1997b). Furthermore, utilizing a series of flow preconditioning regimens followed by step changes in the magnitude of the LSS stimulus, we could demonstrate that the induction of BSC2 mRNA in HUVEC was dependent upon the magnitude of the LSS stimulus, and in the absence of an LSS stimulus of sufficient magnitude (between 4 and 10 dyn/cm2) BSC2 message levels rapidly declined. These results suggest that the induction of BSC2 gene expression is dependent on both the nature as well as the magnitude of the applied fluid mechanical stimulus, and that this gene is not simply responding to the abrupt transition to flow from static (no flow) culture conditions, but rather that a sustained exposure to steady laminar shear stress is the effective stimulus. A second cDNA isolated from a HUVEC library which demonstrates selective upregulation in response to LSS in vitro also was found to encode a member of the 12-membrane spanning superfamily (Topper et al., 1998). This molecule, which is now known as hPGT (for human prostaglandin transporter) is capable of mediating the uptake of a variety of prostanoids including PGE1, PGE2, PGF2 and PGD2, but not Iloprost (a stable PGI2 analogue) when overexpressed in HeLa cells in vitro (Kanai et al., 1995; Lu et al., 1996). Interestingly, this gene demonstrates a very similar pattern of response to shear stress in cultured EC to that of the BSC2 gene described above, namely, it is selectively induced by a threshold magnitude of steady LSS between 4 and 10 dyn/cm2, but not by a comparable magnitude of TSS in vitro. Utilizing a antiserum generated against a peptide derived from this molecule, we have recently demonstrated expression of hPGT in the vascular endothelium of human arteries as well as the microvasculature of myocardial, renal and pulmonary tissues. These results implicate hPGT in the trafficking and/or metabolism of prostanoids within the cardiovascular system, and suggest that the modulation of hPGT gene expression by
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Figure 10.3. Upregulation of the gene encoding the bumetanide sensitive Na-K-2Cl cotransporter BSC2 in cultured HUVEC exposed to 24 hours of steady laminar shear stress. Monolayers of HUVEC were maintained in standard tissue culture conditions (static) or exposed to 24 hours of LSS at 10 dyn/cm2 (flow), and subsequently fixed, permeabilized, and immuno-stained with an antiserum generated against murine BSC2 (kindly provided by Dr. Eric Delpire, Children’s Hospital, Boston, MA) followed by a secondary antibody conjugated to Texas Red. A significant (approximately 5-fold) increase in immunoreactive BSC2 protein is observed within the HUVEC exposed to sustained LSS compared to static HUVEC. The BSC2 gene, together with the hPGT gene (see text) are two examples of a larger class of molecules, the superfamily of 12-membrane spanning transporters, that may be important components of the endothelial phenotype regulated by fluid mechanical stimuli. Reproduced with permission from (Topper et al., 1997b).
fluid mechanical stimuli demonstrated in cultured EC, may be relevant to its expression in vivo. In addition, hPGT and BSC2 may represent the first examples of this large class of molecules (i.e., the superfamily of cell-surface transporters) that in addition to other well characterized classes of endothelial genes such as the leukocyte adhesion molecules, will constitute important functional components of the endothelial phenotype modulated by biomechanical stimuli in vivo.
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VASCULAR SMADS: FLOW AND TGF-ß SIGNAL ING IN VASCULAR ENDOTHELIUM Recently, our differential display strategy has led to the discovery of two additional novel genes, Smad6 and Smad7, encoding members of the MAD-related family of molecules, that were found to be selectively induced in cultured human vascular endothelial cells by steady laminar shear stress stimulation (Topper et al., 1997a). MAD-related proteins (now known as Smad proteins) are a newly identified family of intracellular proteins that are thought to be essential components in the signaling pathways of the serine/threonine kinase receptors of the TGF-ß superfamily (Massague, 1996). Members of this family of proteins are thought to function as signaling intermediates by specifically interacting with ligand-activated receptor complexes at the cell surface and subsequently translocating to the cell nucleus where they are thought to modulate gene expression (a general mechanism of action that is thought to be analagous to that of the STAT family of signaling molecules). The Smad6 and Smad7 proteins, which we originally identified as flow responsive genes in HUVEC, were found to possess unique structural features (compared to the Smad proteins 1–5 that had been described to date). Surprisingly, transient overexpression of Smad6 or Smad7 in vascular endothelial cells resulted in inhibition of the activation of a TGF-ß responsive reporter gene, suggesting that these Smad proteins subserved functions unique from other known Smads. Indeed, subsequent work has demonstrated that both of these proteins can function as specific inhibitors of members of the TGF-ß superfamily of humoral cytokines, and has elucidated the molecular mechanisms responsible for these unique effects (Topper et al., 1997a; Hayashi et al., 1997). Interestingly, these “inhibitory Smads” also can modulate gene expression in response to fluid mechanical stimuli. Overexpression of either of these proteins in cultured endothelial cells significantly, and specifically, inhibits the induction of a reporter gene in response to laminar shear stress. In addition, both Smad6 and Smad7 appear to exhibit a selective pattern of expression in human vascular endothelium in vivo as detected by immunohistochemistry and in situ hybridization on human arterial specimens (Topper et al., 1997a). Thus, Smad6 and Smad7 appear to constitute a novel class of MAD-related proteins, which we have termed the Vascular MADs, that are induced by fluid mechanical forces and can modulate gene expression in response to both humoral and biomechanical stimulation in vascular endothelium.
INTEGRATION OF BIOMECHANICAL AND BIOCHEMICAL STIMULI: FLOW AS A CRITICAL MODULATOR OF ENDOTHELIAL PHENOTYPE With the growing appreciation of vascular endothelium as a central transducer and integrator of diverse stimuli within the cardiovascular system comes the realization that our ability to understand the physiology of this tissue will depend on our understanding of how this cell continuously performs these integrations in vivo. The demonstration that an increasing number of endothelial genes (and their products)
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are responsive to an array of physiologic and pathophysiologic stimuli in vitro has begun to provide a framework with which to examine this complex process. The ability of a uniform laminar shear stress stimulus to selectively induce the expression of a class of signaling molecules that affect the ability of the endothelial cell to respond to subsequent humoral or biomechanical stimuli (i.e., the vascular Smads), may represent an example of such an integration. The observation that many of the genes identified by differential display in cultured EC exposed to defined fluid mechanical stimuli, appear to manifest endothelial selective expression in vivo suggests that these findings are relevant to in vivo physiology. In particular, we hypothesize that the physiologic fluid mechanical stimuli to which endothelium are constantly exposed to in vivo, represent important “differentiative” stimuli, capable of (and potentially responsible for) modulating endothelial phenotype in vivo. The demonstration that the expression of a subset of endothelial-expressed genes appear to be modulated by fluid mechanical stimuli in vitro may in part be a consequence of the fact that EC in culture manifest low, or nonexistent levels of many important effectors (e.g., NOS III, hPGT, Smad6, etc.) as a consequence of maintenance under static (no flow) conditions. However, upon reconstitution of quasi-physiologic flow conditions, EC are able to reacquire, via specific alterations in gene expression, components of their in vivo phenotype that had been lost in culture. This hypothesis thus views applied fluid mechanical stimuli such as sustained laminar shear stresses, not as an acute stimulus (i.e., cytokine-like, capable of rapid modulations of cellular phenotype), but rather as tonic regulators of the differentiated state of the endothelium (and other cellular components of the vessel wall), capable of effecting the phenotype of vascular endothelium over longer time periods. A relevant analogy would be the ability of applied load (e.g., strain/ stretch) to induce patterns of contractile protein isoforms in cultured cardiac myoctes that more closely approximate those which exist in vivo rather than those typically found under standard (no load) culture conditions. A simple prediction of this hypothesis (in the context of our flow studies) would be that genes found to be upregulated by LSS in a sustained fashion in cultured HUVEC in vitro would be found to be expressed in the endothelial lining of umbilical veins (and arteries) in vivo. Indeed, as shown in Figure 10.4 by two color immunofluorescence, ecNOS (NOS III), hPGT, and Smad6 are all expressed at detectable levels in umbilical vessels, suggesting that the low levels of these proteins typically found in EC under standard (static) culture conditions may in fact be a consequence of the lack of sustained fluid mechanical stimulation.
CONCLUSION Over the last decade we have witnessed an explosion of interest in the vascular endothelial cell as a dynamically mutable interface, whose functional phenotype is responsive to various humoral substances including growth factors and cytokines. More recently, it has become apparent that the biomechanical environment of the vessel wall may be as important as the biochemical one (Gimbrone et al., 1997). The
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Figure 10.4. Three genes identified as LSS responsive genes in cultured HUVEC are expressed in the vascular endothelium of human umbilical vessels in vivo. Human umbilical cords containing both umbilical veins (panels A–C) and umbilical arteries (panels D–I) were obtained from normal deliveries (under established institutional protocols) and frozen sections (4–5 µn) were analyzed by immunofluor-escence for CD31 (PECAM-1), a marker of vascular endothelium (panels A, D, G, stained with Texas Red) or the human prostaglandin transporter (hPGT, panel B stained with FITC) ecNOS (NOSIII, panel E, stained with FITC) or Smad6 (panel H, stained with FITC). Panels C, F, I are the FITC and Texas Red images superimposed demonstrating the colocalization of these in vitro flow-responsive gene products with PECAM1 in umbilical vessel endothelium in vivo.
demonstration that specific fluid mechanical stimuli such as fluid shear stresses can directly influence the functional phenotype of the vascular endothelium has begun to provide a framework for a mechanistic understanding of endothelial-dependent processes ranging from remodeling of the vessel wall in response to changing hemodynamics, to the initiation and localization of chronic vascular diseases such as atherosclerosis. The specific fluid mechanical stimuli involved, their resultant phenotypic modulations, and the specific nature and function of the genes involved, promise to be fruitful areas of future investigation. ACKNOWLEDGMENTS The authors gratefully acknowledge the colleagial interactions of past and current members of the Vascular Research Division including Nitzan Resnick and Tucker Collins, as well as the important contributions of our collaborators, in particular, Forbes Dewey, Dean Falb, Jiexing Cai, Tobi Nagel, Keith Anderson, Maria DiChiara, Kay Case, William Atkinson, George Stavrakis, Barbara Sampson, and Fred Schoen. J.N.T. is a recipient of the Howard Hughes Medical Institute Fellowship for Physicians.
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This work was supported in part by grants from the National Institutes of Health (P50-HL56985, R37-HL51150) and an unrestricted gift from the Bristol Myers-Squibb Research Institute to M.A.G.
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Lu, R., Kanai, N., Bao, Y. and Schuster, V.L. (1996) Cloning, in vitro expression, and tissue distribution of a human prostaglandin transporter. Journal of Clinical Investigation, 98, 1142–1149. Marui, N., Offerman, M., Swerlick, R.C.K., Roxen, C.A., Ahmad, M., Alexander, R.W. and Medford, R.M. (1993) Vascular cell-adhesion molecule-1 (VCAM-1) gene transcription and expression are regulated through an antioxidant sensitive mechanism in human vascular endothelial cells. Journal of Clinical Investigation, 92, 1866–1874. Massague, J. (1996) TGFbeta signaling: receptors, transducers, and Mad proteins. Cell, 85, 947–950. O’Donnell, M.E., Brandt, J.D. and Curry, F.R. (1995) Na-K-CI cotransport regulates intracellular volume and monolayer permeability of trabecular meshwork cells. American Journal of Physiology, 268, C1067–1074. Ohno, M., Cooke, J.P., Dzau, V.J. and Gibbons, G.H. (1995) Fluid shear stress induces endothelial transforming growth factor beta-1 transcription and production. Modulation by potassium channel blockade. Journal of Clinical Investigation, 95, 1363–1369. Resnick, N., Collins, T., Atkinson, W., Bonthron, D.T., Dewey, C.F.J. and Gimbrone, M.A. Jr. (1993) Platelet-derived growth factor B chain promoter contains a cis-acting fluid shear-stressresponsive element [published erratum appears in Proc. Natl. Acad. Sci. USA 1993 Aug 15; 90(16): 79081. Proceedings of the National Academy of Science USA, 90, 4591–4595. Resnick, N. and Gimbrone, M.A. Jr. (1995) Hemodynamic forces are complex regulators of endothelial gene expression. FASEB J, 9, 874–882. Sdougos, H.P., Bussolari, S.R. and Dewey, C.F. (1984) Secondary flow and turbulence in a cone-and-plate device. Journal of Fluid Mechanics, 138, 379–404. Shyy, J.Y., Lin, M.C., Han, J., Lu, Y., Petrime, M. and Chien, S. (1995) The cis-acting phorbol ester “12-O-tetradecanoylphorbol 13-acetate”-responsive element is involved in shear stressinduced monocyte chemotactic protein 1 gene expression. Proceedings of the National Academy of Sciences of the USA, 92, 8069–8073. Topper, J.N., Cai, J., Falb, D. and Gimbrone, M.A. Jr. (1996) Identification of vascular endothelial genes differentially responsive to fluid mechanical stimuli: cyclooxygenase-2, manganese superoxide dismutase, and endothelial cell nitric oxide synthase are selectively up-regulated by steady laminar shear stress. Proceedings National Academy of Science, USA, 93, 10417–10422. Topper, J.N., Cai, J., Qiu, Y., Anderson, K.R., Xu, Y.Y., Deeds, J.D., Feeley, R., Gimeno, C.J., Woolf, E.A., Tayber, O., Mays, G.G., Sampson, B.A., Schoen, F.J., Gimbrone, M.A. Jr. and Falb, D. (1997a) Vascular MADs: two novel MAD-related genes selectively inducible by flow in human vascular endothelium. Proceedings of the National Academy of Sciences of the United States of America, 94, 9314–9319. Topper, J.N., Cai, J., Stavrakis, G., Anderson, K.A., Woolf, E., Falb, D. and Gimbrone, M.A. Jr. (1998) The Human Prostaglandin Transporter Gene (hPGT) is Regulated by Fluid Mechanical Stimluli in Cultured Endothelial Cells and Expressed in Vascular Endothelium In Vivo, Circulation, 98, 2396–2403. Topper, J.N., Wasserman, S.M., Anderson, K.R., Cai, J., Falb, D. and Gimbrone, M.A. Jr. (1997b) Expression of the bumetanide-sensitive Na-K-CI cotransporter BSC2 is differentially regulated by fluid mechanical and inflammatory cytokine stimuli in vascular endothelium. Journal of Clinical Investigation, 99, 2941–2949.
11 Differential Regulation of Endothelial Cell Surface Molecules by Diverse Hemodynamic Forces Peter I.Lelkes Laboratory of Cell Biology, Department of Medicine, University of Wisconsin Medical School, Sinai Samaritan Medical Center, P.O. Box 342, Milwaukee, WI 53201–0342, USA, Tel.: (414) 219–7753, Fax: (414) 219–7874, E-mail: [email protected].
Cell adhesion molecules of the immunoglobulin superfamily, such as intracellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1), regulate the interaction of the endothelium with circulating leukocytes and also determine their migration through the endothelial cell(EC) monolayer. Other EC surface proteins, such as tissue factor (TF), are pivotal for the maintenance of vascular hemostasis. The molecular mechanisms by which endothelial cell surface proteins, particularly TF, ICAM-1 and VCAM-1, are upregulated by inflammatory cytokines are fairly well characterized. In this communication we describe recent studies from our laboratory into the distinct mechanisms by which different hemodynamic forces, viz. cyclic strain, elevated pressure and pulsatile flow-induced shear stress, regulate the expression of the genes and the gene products for these EC surface molecules. Besides studying the differential effects of the various hemodynamic forces on the activation of nuclear transcription factors, we examine the role of endothelial cell heterogeneity, by describing differences (and similarities) between endothelial cells derived from various vascular beds (arterial vs. venous and large vessel vs. micro vessels) in terms of their susceptibility to activation by different mechanical forces. Furthermore we discuss the hypothesis that pathological modulations of “physiological” forces, such as shear stress induced by pulsatile perturbed rather than by steady laminar flow, might predispose endothelial cells to enhanced vascular injury.
Abbreviations EC HAEC HMVEC HUVEC ICAM-1 VCAM-1 MC540 PDTC RT-PCR TF TNFa
: endothelial cell(s) : human aortic endothelial cell(s) : human dermal microvascular endothelial cell(s) : human umbilical vein endothelial cell(s) : intercellular adhesion molecule-1 : vascular cell adhesion molecule-1 : Merocyanine 540 : pyrrollidone dithiocarbamate : Reverse transcription polymerase chain reaction : tissue factor : tumor necrosis factor
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INTRODUCTION Endothelial cells (EC) form the ubiquitous single-cell wide lining of the walls of the entire cardiovascular system. In the arterial tree, EC are exposed to pulsatile blood flow, whereas in the venous and in the lymphatic circulation, blood flow is largely steady laminar. Thus, depending on the anatomical site, EC sense and respond to a variety of flow fields which differ both qualitatively (e.g., steady, reversing or pulsatile, laminar vs. perturbed) as well as quantitatively, e.g., in terms of the force amplitudes. Recent studies suggest that the diverse hemodynamic forces associated with distinct local flow patterns might contribute to the functional basal heterogeneity of EC in vivo and also play a role in vascular pathophysiology (Gimbrone et al., 1997; Nakashima et al., 1998). The hemodynamic force of flowing blood is comprised of three components: fluid shear stress, pressure and tension/deformation of the vascular wall (Lelkes and Samet, 1991). Diverse model systems permit to explore, separately, the role of the individual components of the flowing blood at the cellular and molecular level under quasi physiological and/or pathophysiological conditions (Figure 11.1). Such “simple” dynamic systems have been pivotal in advancing our understanding of the importance of hemodynamic forces for the global regulation and maintenance of endothelial cell structure and function (Bussolari, Dewey and Gimbrone, 1982; Banes et al., 1982;
Figure 11.1. Schematic flow-chart of diverse in vitro systems for modeling various hemodynamic forces. Most past and current studies employ systems in which the cells are separately exposed to individual hemodynamic forces, viz. flow, strain and pressure. Recently more complex models are being developed, in which the cells are exposed to more realistic combinations of the separate hemodynamic forces.
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Acevedo et al., 1993). Numerous laboratories are growing EC in a “dynamic” in vitro environment and studying the effects of hemodynamic forces at the cellular and molecular level (Nerem et al., 1993; Patrick and McIntire, 1995; Resnick and Gimbrone, 1995; Papadaki and Eskin, 1997; Traub and Berk, 1998). More recently, EC cultures have been exposed to more realistic hemodynamic forces, featuring nonsteady (i.e. pulsating or oscillatory) flow, shear stress gradients or oscillatory pressure (Helmlinger et al., 1991; Thoumine, Nerem and Girard, 1995; Chappell et al., 1998; Kettlun, Samet and Lelkes, 1996; Tardy et al., 1997). As discussed below, some of the cellular responses to these complex hemodynamic forces are quite distinct from those obtained with the simplified systems (Chappell et al., 1998; Kettlun, Samet and Lelkes, 1996; Ziegler et al., 1998). Currently, model chambers are being developed that provide an even more complex (realistic?) model of the pulsating blood flow by combining pulsatile shear stress and cyclic strain and/ or (pulsatile) pressure in a well controlled dynamic environment (Busse and Fleming, 1998; Golledge et al., 1997). These systems will eventually yield a detailed picture about the differential contribution of the separate forces to the hemodynamic regulation of endothelial cells physiology and, in particular, pathophysiology. EC responses to flow-induced shear stress, depend on both the magnitude of the stress-force and the flow dynamics, i.e. whether the flow is steady, oscillatory or pulsatile. In the past, most studies on the effects of flow on endothelial cells were carried out mainly with steady laminar flow. In vivo, this most simple of all flow patterns is probably restricted to the venous and lymphatic circulation. Also, shear stress amplitudes of 50 dynes/cm2 or more that have been applied to cells in such laminar flow studies, appear to be somewhat un-physiological, certainly in mimicking blood flow in the larger veins (Nerem, 1981). Nevertheless, laminar flow studies were instrumental in defining the pivotal role that mechanical forces, in particular fluid shear stress, play in the maintenance of endothelial cell homeostasis. Cellular responses to applied fluid shear stress can be classified into early and late effects. Ion-channel activation, and changes in the intracellular signal transduction pathways, e.g., receptor tyrosine phosphorylation, activation of G-proteins and second messenger mobilization occur within seconds (Papadaki and Eskin, 1997; Olesen, Clapham and Davies, 1988; Osawa et al., 1997; Ando, Komatsuda and Kamiya, 1988). Activation of downstream targets of cellular signaling cascades (e.g., MAP kinase pathways) occurs within minutes and leads to activation of specific, mechanical force-sensitive genes. Co-ordinate cellular responses, such as changes in the rate of pinocytosis (Davies et al., 1984) or reorientation of the cells according to the flow patterns (Eskin et al., 1984; Levesque and Nerem, 1985; Dewey et al., 1981), occur within hours and require the interplay of both early and late events, as evidenced, e.g., by the rearrangement of the cytoskeletal proteins (Kim et al., 1989; Wechezak et al., 1989), a process which involves regulation of focal-adhesion kinase-associated proteins (Girard and Nerem, 1995). In recent years, several cis-acting shear-stress response elements (SSRE) have been identified, which mediate the mechanical-force induced activation or inhibition of gene-expression (Resnick et al., 1993; Khachigian et al., 1995, 1997; Korenaga, 1997). The by now “classical” SSRE (5’-GAGACC-3’) seems to suffice to confer flowsensitivity to select genes, such as PDGF-B and ICAM-1 (Resnick et al., 1993; Nagel et al., 1994). In general, shear stress response elements appear to be contained in/part
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of the binding sites for known nuclear transcription factors. For example, NF-B interacts functionally with the SSRE. Other transcription factors, such as egr-1, AP-1, and SP-1 have been implicated in the flow-induced upregulation of PDBF-A, MCP-1 and TF respectively, suggesting the existence of multiple cis-acting shear response motifs. (Khachigian, 1997; Lin et al., 1997; Gimbrone, Nagel and Topper, 1997). On the other hand, down-regulation of VCAM-1 expression by steady laminar flow involves the activation of two tandem AP-1 sites (Korenaga et al., 1997). Responsiveness to fluid shear stress (and to other mechanical forces) appears to be more complex than regulation of a single (or multiple) SSRE by steady laminar flow. For example, the promoter of the gene for Platelet/Endothelial Cell Adhesion Molecule1 (PECAM-1) contains binding sites for NF-B, egr-1, SP-1 as well as a sequence identical to that of the classical SSRE (Gumina et al., 1997; Almendro et al., 1996). Yet, while the expression of PECAM-1 is not known to be upregulated by fluid shear stress, flow (as well as other mechanical forces) induce rapid tyrosine phosphorylation of PECAM-1, suggesting that novel, alternative signaling pathways may be involved in mechano-sensing (Osawa et al., 1997; Masuda et al., 1997). In line with this notion, Topper et al. (1997a) recently identified two novel, flow-sensitive MAD-related proteins, which are believed to be essential regulators of the signaling cascade associated with serine/threonine kinases of the TGF-ß superfamily (Topper et al., 1997a). The EC shear stress-sensing machinery seems to be exquisitely sensitive to the nature of the hemodynamic forces, in particular the flow patterns. Recently, several genes have been identified, such as Mn-superoxide dismutase, cyclooxygenase-2 and BSC2 (one of the major Na/K/Cl cotransporters), which are selectively induced by laminar flow, but not by turbulent flow (Topper et al., 1996; Topper et al., 1997b). While some genes, such as ICAM-1, TF, and eNOS seem to be similarly affected by laminar or pulsatile/oscillatory flow (Chappell et al., 1998; Ziegler et al., 1998), the expression of some other genes is regulated by distinct flow patterns in an opposing fashion. For example, the expression of VCAM-1 is downregulated by laminar shear stress, while it is upregulated by both oscillatory and pulsatile perturbed flow (Chappell et al., 1998; Kettlun, Samet and Lelkes, 1996). Cyclic mechanical strain has been increasingly recognized as an important parameter which modulates EC biology. To mimic tensional deformations of the arterial vessel wall, both home made (Gorfien et al., 1989) and commercial cell-exercising/ growth systems (Flexercell®, Banes et al., 1987) are available which allow cyclic deformations, both convex and concave, of flexible, polymeric substrates according to a predetermined regime. These studies demonstrate that mechanical strain activates well described cellular signaling pathways and leads to alterations in gene expression in a manner which in some cases is similar to, in other cases quite distinct from the effects of fluid-shear stress. For recent reviews see (Mills, Cohen and Sumpio, 1993; Banes et al., 1995). Second messengers involved in the translation of the cyclic mechanical strain signal include calcium signaling as well as protein kinase C and cyclic AMP dependent pathways (Manolopoulos and Lelkes, 1993; Cohen et al., 1997; Evans et al., 1997). Also, in terms of the signaling pathways, cyclic strain activates both the erk and the JNK/SAPK pathways of the MAP kinase cascade (Reusch et al., 1997) and induces tyrosine phosphorylation of the focal adhesion kinase, pp125FAK (Yano, Geibel and Sumpio, 1996). Transcription factors activated by cyclic strain
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include NF-B as well as select members of the AP-1 family (Sumpio, Du and Xu, 1995). To date, no unique “Cyclic Strain Response Element” analogous to a flowsensitive SSRE has been discovered. Although nuclear proteins from strain-activated cells bound to the SSRE, the current belief is that the SSRE is neither necessary nor sufficient for gene activation by cyclic strain (Sumpio et al., 1998). In general, gene activation by cyclic strain alone appears to be less pronounced than that by fluid shear stress (Segurola et al., 1997; Silverman et al., 1996). Cyclic strain modulates gene expression both at the transcriptional and post-transcriptional level, the latter presumably by altering message stability. Furthermore, cyclic straininduced post-translational modifications, such as prenylation or farnesylation may constitute yet another mechanism by which hemodynamic forces may modulate EC protein expression and/or function. While the cellular effects of flow-induced shear stress and cyclic strain are under intense scrutiny, few studies have addressed the effects of elevated, time mean (hydrostatic) pressure on EC cultures in vitro. This is quite surprising, since throughout the vascular tree, ECs are exposed to different levels of steady (in the venous circulation) or pulsating (in the arterial tree) hydrostatic (blood) pressure. The few published reports imply a profound role for hydrostatic pressure in regulating the morphology and permeability characteristics of intact EC linings (Davies and Bowyer, 1975; Suttorp et al., 1989). Exposure of cultured EC to constant hydrostatic pressure was found to modulate cell proliferation, PGI2 synthesis (Tokunaga and Watanabe, 1987), and gene expression (Riley and Gullo, 1988). As yet another indicator for their heterogeneity, ECs derived form various tissues exhibit different pressure sensitivities. For example, prostacycline (PGI2) production in rat pulmonary ECs increases significantly at hydrostatic pressures exceeding 7.5 mm Hg (van Grondelle et al., 1984). On the other hand, PGI2 production in human micro vascular ECs seems to be inversely proportional to the applied pressure, being maximal at 0 mm Hg but very low at 80 mm Hg (Tokunaga and Watanabe, 1987). Recent studies suggest that ECs differentially sense and respond to constant and pulsatile pressure, respectively. For example, the rate of proliferation of bovine aortic EC is increased when the cells are exposed to low levels (11 mm Hg) of constant pressure, presumably due to the secretion of mitogenic growth factors (Acevedo et al., 1993). No growth inhibition is observed when the cells are exposed to higher levels (135 mm Hg) of constant pressure. By contrast, ambient pulsatile pressure (110/160 mm Hg) seems to trigger the release of an as yet undetermined humoral factor which inhibits EC proliferation (Vouyouka et al., 1998). Comparison of the effects of these three separate components of blood flow often reveals quite distinct, force-specific and sometimes opposing effects. For example, fluid-shear stress activates a potassium channel (Olesen, Clapham and Davies, 1988), whereas non-selective ion channels are activated by mechanical stretching (Naruse and Sokabe, 1993). DNA synthesis and cell proliferation in bovine aortic ECs is enhanced by mechanical stretching (Mills, Cohen and Sumpio, 1993), whereas, (turbulent) flow-induced shear stress reduces EC turnover (Davies et al., 1986). Protein synthesis is regulated in a complex manner by both flow and cyclic strain and is often cell type specific (Guptee and
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Frangos, 1990; Yoshizumi et al., 1989; Sumpio et al., 1988, 1990). For example, initiation of flow stimulates arachidonate metabolism, resulting in the transient release of PGI 2 (Nollert et al., 1989). PGI2 production increases also with increasing uniform-flow-induced shear stress (McIntire et al., 1987). By contrast, the effects of mechanical stretching on PGI 2 production are controversial (Sumpio and Banes, 1988; Upchurch et al., 1989). Moreover, EC align in parallel with the vector of flow, but perpendicular to the vector of mechanical deformation (Mills, Cohen and Sumpio, 1993). Finally, EC responses differ, on many occasions in a contradictory manner, at different pressure levels with and without perfusing flow (Ziegler et al., 1998; Ando, Komatsuda and Kamiya, 1988; Tokunaga and Watanabe, 1987). Understanding the complex regulation of EC physiology and pathophysiology by hemodynamic forces has several important clinical implications. Recent theories as to the origin of vascular injuries, such as atherosclerosis, clearly implicate “aberrant” hemodynamic forces as one of the major culprits, in conjunction with genetic pre-disposition and/or habitual risk factors, such as diet and smoking (Nakashima et al., 1998; Ross, 1993, 1995; Davies et al., 1995; Davies, 1995). Indeed, recent in vivo studies clearly demonstrate the early upregulation of leukocyte adhesion molecules at sites of atherosclerotic predilections (Nakashima et al., 1998; Davies et al., 1993). Thus, the onset/ development of atherosclerotic lesions might be prevented/reverted with proper pharmacological intervention that could avert the upregulation of these “atherogenic” molecules by hemodynamic forces. On the other hand, in trying to repair the damage to the cardiovascular system through bioengineering, numerous investigators have proposed to improve the hemocompatibility of artificial grafts and blood pumps by lining these devices with a monolayer of non-thrombogenic/ non-atherogenic ECs (Eskin et al., 1978; Ziegler and Nerem, 1994). The preclinical and clinical studies in this field have clearly demonstrated that the feasibility of this approach requires a thorough understanding of the effects of hemodynamic forces on the cells of the neointima. Our long-term goal is to generate non-thrombogenic, endothelial-cell lined artificial cardiovascular prostheses, e.g., total artificial blood pumps. A tissue-engineering approach towards reconstructing a “living”, bio-integrated blood-contacting surface in these devices recognizes that the functional properties of the endothelial cells are tightly controlled by the complex hemodynamic forces inside a beating cardiac ventricle (Lelkes and Samet, 1991). Over the past years, we have identified several crucial mechanical, force-sensitive parameters, such as the synthesis and assembly/ maturation of the subendothelial basement membrane, the anti-/procoagulant properties of the endothelial cell surface as well as the expression of leukocyte adhesion molecules. In this communication, we will, within the context of mechanical force-responsiveness and endothelial cell heterogeneity, compare the differential modulation by pulsatile non-laminar flow, cyclic strain and hydrostatic pressure of some of the endothelial cell surface molecules that mediate endothelial-leukocyte interactions (e.g., ICAM-1 and VCAM-1) and regulate blood coagulation, e.g., tissue factor.
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VASCULAR CELL ADHESION MOLECULES Leukocyte adhesion to vascular ECs plays an important role in the initiation and progression of numerous, heart-disease related pathophysiological conditions, such as atherosclerosis, hypertension, stroke, ischemic heart disease and cardiac allograft rejection (Bevilacqua, 1993; Albelda, Smith and Ward, 1994; Tedder et al., 1995). Similarly, in endothelialized artificial grafts, the integrity of the endothelial cell monolayer can be compromised by transmigrating leukocytes (Margiotta, Benton and Robertson, 1994; Herring, Emerick and Ashworth, 1987). The coordinated sequence of leukocyte adhesion and extravasation occurs via specific interactions between cell adhesion molecules (CAMs) expressed on the EC surface and their cognate receptors on various leukocytes(Bevilacqua et al., 1994). Heterotypic vascular cell adhesion molecules of the IgG super family, in particular intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1), play a pivotal role in leukocyte/endothelial cell interactions. The adhesion of circulating leukocytes to activated endothelial cells under pathophysiological conditions is regulated sequentially by an initial arrest, subsequent rolling and final attachment to the endothelial cell surface. ICAM-1, which is constitutively expressed on the EC surface and upregulated upon activation by, e.g., inflammatory cytokines, contributes to the firm adhesion and emigration of all classes of leukocytes. The expression of VCAM-1 is minimal in quiescent EC but dramatically upregulated by cytokines and other atherogenic factors. VCAM-1 is largely “responsible” for the adhesion and extravasation of mononuclear leukocytes, which subsequently form the foam cells in fatty streaks. Upregulation of CAMs, in particular of VCAM-1 in “high risk areas” for formation of atherosclerotic plaques, e.g., at bifurcations or in other regions of disturbed hemodynamic forces, is believed to be amongst the first signs of pre-atherosclerotic lesions and is observable prior to the intimal accumulation of extravasated monocytes (Nakashima et al., 1998; Cybulsky and Gimbrone, 1991). While VCAM-1 expression has been linked to the early onset of endothelial dysfunction leading to the beginning of atherosclerotic lesions, the EC lining of mature atherosclerotic plaques (fatty streak) does not express VCAM-1, but rather ICAM-1. Thus, the sequential induction and/or coordinate expression of these two adhesion molecules is important for the continued recruitment of leukocytes in inflammatory lesions of the vascular wall (Ross, 1995).
Differential Regulation of ICAM-1 and VCAM-1 by Cytokines Upregulation of ICAM-1 and VCAM-1 by cytokines, such as tumor necrosis factor (TNF), appears to differentially involve activation and nuclear translocation of the nuclear transcription factor NF-B (Roebuck et al., 1995; Marui et al., 1993). Although kappa B motifs are present in both the VCAM-1 and ICAM-1 promoters, pyrollidine dithiocarbamate (PDTC, a free oxygen radical scavenger and NF-B antagonist), reduces TNF-induced VCAM-1 but not ICAM-1 surface protein expression in human umbilical vein EC (HUVEC) (Collins et al., 1995). Since PDTC prevents NF-B mobilization by TNF, these results suggest that only VCAM-1 induction is controlled
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by NF-B. Recent studies indicate that both B sites in the VCAM-1 enhancer are required to optimally stimulate gene expression, but the enhancer is differentially regulated by specific combinations of NF-B subunits (Shu et al., 1993). In contrast to TNF, other cytokines, such as IL-1, or IL-4 appear to induce ICAM-1 and VCAM1 independent of NF-B activation (Marui et al., 1993; McCarty et al., 1995). Furthermore, ICAM-1 upregulation by H2O2 involves the activation of AP-1/Ets elements within the ICAM-1 promoter (Roebuck et al., 1995). Activation of CAM s in HUVEC by cytokines is also regulated, at least in part, by PKC activation, although this is not associated with the expected translocation of PKC from the cytosolic to the particulate fraction (Myers et al., 1992; Deisher et al., 1993; Sung, Arleth and Nambi, 1994). Selective inhibitors of tropoisomerase II, a nuclear target for PKC, selectively abrogated cytokine-induced VCAM-1, but not ICAM-1 surface expression (Deisher, Kaushansky and Harlan, 1993). Lysophosphatidylcholine, an atherogenic risk factor, elevates ICAM-1 in a PKC-independent fashion (Kume et al., 1995).
Regulation of 1C AM-1/VCAM-1 by Steady and Pulsatile Flow-induced Shear Stress Previous studies have used steady laminar flow conditions to reveal some of the differences and similarities in the mechanisms by which the expression of ICAM-1 and VCAM-1 is induced by various atherogenic risk factors and hemodynamic forces. Although VCAM-1 and ICAM-1 belong to the same IgG superfamily of CAMs, distinct activation pathways have been suggested which, depending on a particular agonist and cell-type, include diverse nuclear transcription factors (such as NF-B AP-1, SP1), protein phosphorylation via PKA, PKC, and receptor tyrosine kinases, as well as activation of phospholipase C (PLC) and the cyclooxygenase pathway. Endothelial cell activation through hemodynamic forces utilizes some of the same signaling pathways that are induced by cytokines. As described in detail in several chapters in this book, both fluid shear stress and cyclic mechanical stain activate protein phosphorylation, via PKC, PKA and receptor tyrosine kinases. Moreover, both these hemodynamic forces induce the nuclear translocation of distinct members of the families of NF-B and AP-1 transcription factors (Sumpio, Du and Xu, 1995; Lan, Mercurius and Davies, 1994). Yet, the literature clearly shows opposing in vitro responses of ICAM-1 and VCAM-1 to elevated laminar shear stress beyond a threshold of approx. 2 dynes/cm2: ICAM-1 is upregulated, while VCAM-1 is downregulated (Nagel et al., 1994; Ando et al., 1993). The possible reason is the presence of a shear stress-responsive element (SSRE) in the 5’ promoter region of ICAM-1 which is lacking in VCAM-1(Resnick and Gimbrone, 1995; Resnick et al., 1993; Nagel et al., 1994). The down-regulation of VCAM-1 by laminar shear stress has been attributed to the activation of a novel, inhibitory shear-regulatory element which corresponds to a tandem cis-acting AP-1 sequence in VCAM-1 promoter (Korenaga et al., 1997). These in vitro studies only partially reflect the in vivo situation. In animal models of atherosclerosis as well as in patients, both ICAM-1 and VCAM-1 are upregulated at sites of atherosclerotic predilections (Nakashima et al., 1998; Davies et al., 1993). To explain this dichotomy, we hypothesized that the expression of
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VCAM-1 might be sensitive to the “aberrant” flow patterns found at the sites of atherosclerotic lesions, such as the aortic arch, branching points or bifurcations. In order to study the effects of such non-laminar, pulsatile flow patterns on the regulation of EC adhesion molecules, specifically ICAM-1 and VCAM-1, we developed a novel circular flow chamber (Figure 11.2). In this system, pulsatile flow enters the chamber where covers slips lined with confluent EC monolayers are positioned at a various locations. The particular flow fields in each locale are such that in coverslips 1, 3, and 5 the cells experience a pulsatile, jet flow-like pattern. In positions 2 and 4, the flow fields represent a perturbed flow pattern similar to a recirculating eddy, which realistically mimics the flow patterns and the low-shear stress observed in the areas of flow perturbance such as the sites of atherosclerotic predilections (Dewey, 1979; Asakura and Karino, 1990). In line with previous reports by others, basal ICAM-1 mRNA levels in cultured human aortic endothelial cells (HAEC), exposed for 4 hours to laminar steady shear stress of 40 dynes/cm2, are upregulated by more than 20 fold, while VCAM-1 mRNA is down-regulated by approx. 2 fold, as assessed by quantitative RT-PCR (Davis and Lelkes, unpublished). Qualitatively similar results are obtained also at the protein level, using whole cell ELIS A (Kanda et al., 1998). By contrast, exposure for 5 hrs to pulsatile, pertubed flow upregulated VCAM-1 protein expression in both HUVEC
Figure 11.2. Schematic of the circular flow chamber used to model pulsatile non-laminar flow. Endothelial cells are grown on cover slips which are placed in positions 1 through 5. Cells in positions 1, 3, 5 are exposed to the pulsatile jet-stream, whereas in positions 2 and 4, the flow patterns simulate a perturbed flow comprising recirculating eddies.
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and HAEC (Kettlun et al., 1996), (Lelkes et al., 1998; Lelkes et al., manuscript in preparation). Upregulation of VCAM-1 was particularly pronounced (approx.7 fold over basal/static) at the sites of the flow perturbance/ recirculating eddies (positions 2,4), whereas at the sites of the jet stream (positions 1, 3, 5), the increase in VCAM1 levels was approx. 2 fold (Figure 11.3). These results are unique in that they show for the first time that non-laminar flow conditions indeed enhance and up-regulate the expression of VCAM-1 such as observed in situ. In confirming our findings, Chappell et al. (1998) recently showed that other non-laminar flow patterns, such as low-shear oscillatory flow, also stimulate VCAM-1 expression in cultured HUVEC. Concomitant with the up-regulation of VCAM-1 we also observed an up-regulation of ICAM-1 by pulastile-non-laminar flow (not shown). In contrast to VCAM-1, the Upregulation of ICAM-1 by pulsatile flow was not dependent on the flow pattern, viz., that is we observed similar degrees of up-regulation by low shear perturbed flow and by low shear jet flow. To further contrast the differential responsiveness of ICAM1 and VCAM-1, pulsatile flow-induced Upregulation of VCAM-1, but not of ICAM1 was abrogated by preincubation of the cells with 10 µM pyrollidine-dithiocarbamate (PDTC), an established scavenger of reactive oxygen species and inhibitor of the nuclear transcription factor NF-B (Figure 11.4). The induction of VCAM-1 protein expression by pulsatile, non-laminar flow was transcriptionally regulated as inferred from the inhibition of the flow-induced upregulation VCAM-1 mRNA and protein by actinomycin D and cyclohexamide, which disrupt transcription and translation respectively (not shown). In terms of the signaling pathway, it has been observed that laminar flow increases the expression of transcription factors such as AP-1 and NF-B. Indeed, the SSRE has been identified as one of the possible binding sites for the consensus sequence of NF-B. Interestingly, non-laminar perturbed flow does not activate NF-B (Figure 11.5). However, several
Figure 11.3. Upregulation of VCAM-1 expression by pulsatile non-laminar flow. Human aortic endothelial cells (HAEC), cultured in the flow chamber depicted in Figure 11.1, were exposed for 5 hrs to pulsatile flow (100 ml/min, 1 Hz). VCAM-1 protein expression in, as determined by whole-cell ELISA, is expressed relative to static controls. While VCAM-1 expression increases in all positions, the upregulation is particularly pronounced at the sites of perturbed flow (positions 2 and 4). Values represent means ± SEM.
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Figure 11.4. Pulsatile, non-laminar flow-induced upregulation of VCAM-1 but not of 1C AM1 is redox-sensitive. HAEC were exposed for 5 h to perturbed flow (positions 2 and 4) in the absence and presence of the reactive-oxygen species antagonists pyrollidine dithiocarbamate (PDTC). VCAM-1 and ICAM-1 expression were assessed by whole cell ELISA. The results are expressed relative to static controls. Values represent means ± SEM.
Figure 11.5. Differential activation of nuclear transcription factors by pulsatile, non-laminar flow. Following 30 min exposure of HAEC to perturbed flow (positions 2 and 4), the activation of nuclear transcription factors was assessed by electrophoretic mobility shift assays (EMSA). A: NF-B, B: AP-1. Lane 1: static control, basal lane 2: static control+10 ng/ml TNF; Lane 3: perturbed flow; lane 4: perturbed flow +10 ng/ml TNF.
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other transcription factors such as AP-1, egr-1 and SP-1 are differentially activated in model system (Figure 11.6). Our results indicate that endothelial cell perception of flow patterns is exquisitely sensitive to the nature of the flow fields. Since different cellular signaling mechanisms seem to be involved in the up-regulation of adhesion molecules by laminar vs. nonlaminar flow, we propose the existence of a “perturbed/pulsatile flow response element” in the promoter regions of genes like VCAM-1, which are selectively up-regulated by non-laminar flow patterns.
Differential Regulation of ICAM-1/VCAM-1 by Cyclic Strain The effects of cyclic mechanical strain are studied by culturing the cells in flexible bottom multiwell plates and using a commercially available strain/deformation system, the Flexcell®, as previously described (Banes et al., 1995; Mills, Cohen and Sumpio, 1993; Gilbert et al., 1994; Banes, 1993; Banes et al., 1990). Under basal/ static conditions, human EC of arterial origin express approx. twice as much ICAM-1 protein than EC of venous origin or micro vascular EC. By contrast, the basal protein levels of VCAM-1 protein are very low and indistinguishable in all EC1 types studied. Upon stimulation for 8 hr with 100 ng/ml TNF; (under static conditions), all cell types express similar levels of ICAM-1 and VCAM-1, respectively (Kanda et al., 1998).
Figure 11.6. Differential sensitivity of nuclear transcription factors to activation by distinct flow patterns. Nuclear translocation (Activation) of three putatively flow-sensitive transcription factors, AP-1, egr-1 and SP-1, was assessed by EMSA, as described in the legend to Figure 11.5. The data denote the increase in transcription factor activity relative to static controls, as determined densitometrically. Activation of AP-1 and SP-1 was virtually identical in all positions, while egr-1 was differentially induced by jet (positions 1, 3, 5) and perturbed flow (positions 2, 4). Note the similarity in the induction pattern of egr-1 and VCAM-1 (see Figure 11.3). Values represent means ± SEM.
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Cyclic strain-induced changes in the level of adhesion molecule expression were relatively small (<2 fold) and depended on the origin of the EC (Lelkes et al., 1996). For VCAM-1, exposure to cyclic strain (approx 20% maximal strain, 1 Hz) for 24 h induced an approximately 75% increase in VCAM-1 protein expression in HUVEC, but an approximately 20% decrease in HMVEC (Figure 11.7). Qualitatively similar, but smaller changes were observed for ICAM-1. In preliminary studies with human arterial endothelial cells, no cyclic-strain induced change in VCAM-1 protein expression was observed, whereas ICAM-1 levels were slightly elevated (Silverman and Lelkes, unpublished). Cyclic strain-induced upregulation of VCAM-1, but not of ICAM-1, was inhibited by pretreating HUVEC for 30 min with 10 µM PDTC (Figure 11.8), suggesting that, similar to distinct reposes to cytokine-stimulation, different cellular signaling pathways are involved in the activation of these adhesion molecules by cyclic strain. Taken together, our results suggest a distinct regulation of the expression of leukocyte adhesion molecules by cyclic strain. Based on our data, and in analogy to their response to fluid-shear stress, we suggest that the differences in the responsiveness of ICAM-1 and VCAM-1 to cyclic strain may, in part, be due to the presence of distinct differences in “cyclic-strain responsive elements” in the cis-acting promoter regions of these two adhesion molecules. The most striking finding, to date, is the down regulation of VCAM-1 protein in microvascular EC vs. an up-regulation in EC derived from large vessels. This result might reflect a cell-type-specific modulation of message stability by mechanical forces (Papadaki and Eskin, 1997). Alternatively, cyclic-strain-induced down-regulation of ICAM-1 and VCAM-1 in HMVEC might suggest either the
Figure 11.7. Differential induction of VCAM-1 and ICAM-1 expression in cultured human endothelial cells. Human umbilical vein endothelial cells (A) and dermal micro vascular cells (B) were exposed for 24 h to cyclic strain (15 % maximal strain, 1 Hz). VACM-1 and ICAM1 levels, as determined by whole cell ELISA (see Figure 11.3), are given as increase/decrease over static controls. Values represent means ± SEM.
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Figure 11.8. Differential redox sensitivity of cyclic-strain induced VCAM-1 and ICAM-1 expression in HUVEC. Confluent monolayers of HUVEC were exposed, in the absence or presence of 10 µM PDTC, for 5 hrs to cyclic strain. Relative VCAM-1 and ICAM-1 levels are expressed as increase over static controls. Values represent means ± SEM.
existence of distinct, cell type/tissue-specific cis-acting promoter elements and/or that the regulation of the functional protein expression occurs post-translationally. Tissue-specific gene expression in EC has recently been demonstrated (Aird et al., 1995, 1997). The 5’ flanking region of the ICAM-1 gene contains both tissue- and cytokine-specific responsive elements (Cornelius et al., 1993). Our data suggest that, in addition to the classical SSRE, the ICAM-1 promoter might entail also other/ novel mechano-sensitive response elements and that regulatory effects directed by such elements are likely to depend upon their cellular context. Future studies will have to address whether cyclic strain might also affect some of the post-translational mechanisms. Hydrostatic Pressure Does not Affect the Expression of ICAM-1/VCAM-1 Previous studies have shown a clear correlation between hypertension, atherogenesis and hypercoagulability (Susic, 1997; Glasser, Selwyn and Ganz, 1996; Alexander, 1995; Barbagallo et al., 1993; Daae et al., 1993). These pathophysiological states involve the upregulation of EC surface molecules. As a first step toward studying the effects of pressure on EC adhesion molecule expression, we exposed several types of human endothelial cells to constant hydrostatic pressure at physiological and pathophysiological levels of 100 and 170 mm Hg, respectively. The pressure system has been previously described (see chapter by Samet and Lelkes). In our hands, neither ICAM-1 nor VCAM-1 expression were affected by steady hydrostatic pressure of either 100 or 170 mm Hg (Kanda et al., 1996, Kanda et al., manuscript in preparation). The lack of any pressure effect was ascertained both at the protein as well as at the gene level. This finding is somewhat surprising since in
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the past it has been shown that exposure of endothelial cells to elevated static pressure induced morphological changes, increased proliferation (Sumpio et al., 1994), presumably due to an enhanced secretion of growth factors, such as bFGF (Acevedo et al., 1993). Similarly, ongoing studies in our laboratory suggest that endothelial cells exposed to continuous hydrostatic pressure for up to 3 days, are activated to ablumenally secrete significant amounts of vascular endothelial growth factor (VEGF), concomitant with enhanced basolaterally deposition of bFGF into the subendothelial basement membrane (Silverman et al., 1998). Both these growth factors are known EC “activators” which stimulate proliferation and migration. Both bFGF and VEGF reportedly affect cell adhesion molecule expression in angiogenic, i.e. proliferating/migrating EC, albeit in opposing directions: ICAM-1/ VCAM-1 expression is upregulated by VEGF and down-regulated by bFGF (Melder et al., 1996; Griffioen et al., 1996). In addition, hydrostatic pressure leads to a remodeling of the subendothelial extracellular matrix, including an enhanced deposition of basement membrane proteins, such as collagen type IV (Silverman, Samet and Lelkes, 1995). Collagen type IV has been shown to specifically inhibit the expression of VCAM-1 (Morisaki et al., 1995). We hypothesize that the apparent lack of a visible effect of static pressure on ICAM-1/VCAM-1 expression might reflect some compensatory mechanisms by which confluent EC monolayers are protected against activation by growth factors. Sustaining a monolayer configuration rather than the induction of migration will down-regulate pathways which might lead to the activation of adhesion molecules. Taken together, these findings might rule out a direct involvement of elevated steady pressure in the up-regulation of cell adhesion molecules and enhanced hypercoagulability which is observed for example in hypertension (Blann et al., 1994; Lip et al., 1995; McCarron et al., 1994; Komatsu et al., 1997; Arndt, Smith and Granger, 1993; Liu et al., 1996). Obviously, these results do not rule out the possibility that oscillatory pressure alone or in concert with any of the other parameters of the hemodynamic forces, such as cyclic strain and flow, may alter the expression of cell adhesion/cell surface molecules. Indeed, as shown below, elevated pressure transiently elevates the activity, but not, the expression of tissue factor.
TISSUE FACTOR Tissue Factor (TF) is a 46 kD, inducible transmembrane glycoprotein, which in concert with factor VIIa forms a proteolytically active complex on the surface of diverse cell types, including vascular ECs. This TF/VIIa complex is the major initiator of the extrinsic coagulation cascade (Camerer, Kolsto and Prydz, 1996; Carson and Brozna, 1993). The mechanisms of TF regulation both at the level of gene expression and biological activity of the gene product have been recently reviewed (Camerer, Kolsto and Prydz, 1996; Mackman, 1995). At the transcriptional level, regulation of TF expression is complex and cell type-specific. In some cells, such as monocytes and ECs, basal TF expression is very low, but is readily induced by, e.g., inflammatory cytokines or mitogens (Silverman et al., 1996; Oeth, Parry and Mackman, 1997; Terry and Callahan, 1996; Pettersen et al.,
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1992) in a calcium- and PKC-dependent mechanisms. In other cell types, such as fibroblasts, basal TF expression is several orders of magnitude higher than in EC and is further inducible in a PKC-independent fashion, e.g., following serum deprivation (Camerer, Kolsto and Prydz, 1996; Cui et al., 1994, 1996). Promoter deletion studies revealed the presence of a series of binding sites for diverse nuclear transcription factors: 2 AP-1 sites and 1 NF-B site, which are involved in cytokineand LPS-mediated TF up-regulation and 4 SP-1 sites and at least 3 partially overlapping egr-1 binding sites, involved in the serum-response (Camerer, Kolsto and Prydz, 1996; Mackman, 1995). In endothelial cells, there are additionally at least 2 negative regulatory elements and yet another NF-B binding site, which is involved in TF expression in response to advanced glycosylation end products in diabetes (Mackman, 1995; Moll et al., 1995). Transcriptionally, TF mRNA in EC is generally increased by agonists that elevated intracellular calcium and/or activate PKC (Terry and Callahan, 1996), whereas agents that increase intracellular cAMP cause a decrease in TF mRNA expression. In contrast to the established role of PKC, the involvement of growth factor receptor tyrosine kinase-mediated signaling in the transcriptional regulation of TF in EC is controversial (Clauss et al., 1996). Recent studies with protease inhibitors, the antioxidant pyrrolidine dithiocarbamate (PDTC) and aspirin established a strong correlation between the level of (in)activation of NF-B and the expression of TF, suggesting that the activation of NF-B may serve as the central integrator for cytokine/LPS/PMA/-dependent induction of TF in EC (Orthner, Rodgers and Fitzgerald, 1995). Remarkably, all of the above mentioned signaling pathways and/ or nuclear transcription factors are also induced/activated by hemodynamic forces (see below).
Effect of Cyclic Strain on TF Expression We used the Flexcell® system to study the effects of cyclic mechanical strain on TF activity in various cultured human endothelial cells (Silverman et al., 1996). In all EC types studied, basal TF activity, as detected by a two-step amidolytic assay, was barely detectable and increased significantly upon activation of the cells with cytokines. For example, in HAEC stimulated with 10 ng/ml TNFa, total cellular TF activity was approx. 35 fold higher than basal control. Of the total TF activity approx. 30% was cell surface associated. Exposure to 15% average cyclic strain for 5 hours caused a modest, yet statistically significant increase in TF activity in several human endothelial cell types, such as in HMVEC and in HAEC, but not in HUVEC (Silverman et al., 1996). The increase in cyclic strain-induced TF activity in HAEC is preceded (at two hours) by a pronounced, approx. 25 fold increase in mRNA expression, which is quantitatively similar to the cytokine-induced increase in TF mRNA (Wakita, Marumoto and Horiuchi, 1994). Yet, TF activity induced by cyclic strain is approx. one order of magnitude smaller than that induced by chemical agonists. This dichotomy raises interesting questions as to cyclic-strain-induced changes in mRNA stability, and/or possible post-translational modifications, all of which might affect the efficiency of the translation of mRNA into a functional protein.
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Cyclic-strain induced TF mRNA was not inhibited by the reactive oxygen antagonists PDTC (10 µM). This finding is somewhat surprising in light of the findings that cyclic strain activates the nuclear transcription factor NF-B (Du, Mills and Sumpio, 1995), which has been implicated in cytokine-induced TF activation (Mackman, 1997). On the other hand, this finding is in line with recent findings by Lin et al. (1997), as well as with our own data (see below) which clearly indicate that mechanical force (non-laminar flow)- induced activation of TF expression may be mediated by nuclear transcription factors distinct from NF-B. When HAEC were exposed to cyclic strain for up to 72 h, the elevation in TF expression was maintained for approx. 24 h and then gradually (by 48 h) returned to baseline levels. In long-term studies using HMVEC, continuous exposure of the cells to cyclic strain for up to 14 days initially, transiently elevated “basal” TF expression which subsequently (after approx. 4 days) decreased significantly below the static controls (Figure 11.9). This finding parallels our previous results showing that in the same HMVEC the production of plasminogen activator inhibitor type I (PAI-1) the main inhibitor of the fibrinolytic enzymes tPA and µPA, is sharply increased during the first 24 hours of exposure to cyclic strain, but then gradually declines to below baseline (Lelkes et al., 1992). By contrast, production of prostacyclin (PGI2), a hallmark for the antithrombotic EC phenotype, is dramatically increased over time, when the cells are continuously exposed to cyclic strain for up to 14 days (Lelkes et al., 1992). Taken together these observations suggest that while initial exposure to
Figure 11.9. Transient upregulation of tissue factor expression in HMVEC by cyclic strain. Exposure of confluent monolayers of HMVEC to cyclic strain leads to a transient upregulation in tissue factor (TF) expression, which peaks after approx. 1 day and then gradually returns to baseline after approx. 4 days. Continued exposure to cyclic strain results in TF levels significantly lower than those under static conditions. Values represent means ± SEM *: p<0.05.
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cyclic strain might contribute to a prothrombotic EC phenotype, prolonged exposure to the same mechanical force will passivate the cells and restore the nonthrombogenic phenotype. Effect of Elevated Pressure on TF Activity/Expression Exposure of HAEC for up to 48 hrs to elevated static pressure (170 mm Hg) induced TF activity by about 50%. Exposure to elevated pressure increased both basal and TNF-induced TF activity in HAEC in a similar fashion (Silverman et al., 1997). Surprisingly, the pressure-induced TF activity was not accompanied by an enhanced expression of TF, as confirmed at the gene level, using a highly sensitive quantitative RT-PCR assay as well as at the protein level, using both Western blotting and ELISA. Thus, we hypothesized that the observed increase in TF activity might be due to direct effects of the applied pressure on the physicochemical properties of the plasma membrane, either by damage (Carson, Perry and Pirruccello, 1994) or by changes in the fluidity of the phospholipid bilayer (Gudi and Frangos, 1994). We did not find any evidence for enhanced LDH-leakage, trypan blue uptake or enhanced pinocytosis/ endocytosis, as assessed by the uptake of fluorescein dextran (Berthiaume and Frangos, 1994), thus excluding pressure-induced damage to the cells. Also, in control experiments we established that the small alterations in the culture medium observed during pressurization for up to 48 h, viz., increase in the pH by 0.1 pH units, decrease in dissolved pCO2 by <10% and increase in dissolved pO2 by <20%, did not affect TF activity. To study the effects of pressure on membrane fluidity, we used a fluorescent dye Merocyanine 540 (MC540), which has been extensively studied as a probe for assessing membrane fluidity and lipid asymmetry (Lelkes, Bach and Miller, 1980; Williamson, Mattocks and Schlegel, 1983; McEvoy, Williamson and Schlegel, 1986; Lelkes and Miller, 1980; Stillwell et al., 1993). In support of our hypothesis we found that the pressure-induced increase in TF activity in HAEC was accompanied by an increase in membrane fluidity, as assessed by enhanced in MC540 incorporation (Figure 11.10). This result suggests that hydrostatic pressure may directly affect the physicochemical properties of the plasma membrane, thus altering the activity of TF.
Effects of Pulsatile, Non-Laminar Flow on TF Expression Previous studies suggested that steady, laminar flow induces an increase in TF activity, presumably through upregulation of TF gene expression concomitant with a down regulation of the tissue factor pathway inhibitor protein, TFPI (Lin et al., 1997; Grabowski and Lam, 1995). In order to study the effects of pulsatile, non-laminar low-shear (<1 dyne/cm2) on TF expression, we used our flow-chamber, (see Figure 11.2), exposing cultured HAEC to diverse local flow patterns (jet, recirculating eddy/ areas of stagnation). After 5 hours, TF activity was significantly (approx. 8-fold) elevated by pulsatile flow. This increase in TF activity amounted to approx. 25% of that observed upon stimulation of the cells with 1 ng/ml TNF. In contrast to VCAM-1, however, the upregulation of TF activity by pulsatile flow did not seem to
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Figure 11.10. Elevated constant pressure alters membrane fluidity in HAEC. Human aortic endothelial cells were exposed for 5 h to elevated ambient pressure (170mm Hg). The fluorescent dye Merocyanine 540 (MC540) was added during final 15 min of the experiment. MC540 incorporation/uptake is expressed as relative amount of cell associated fluorescence. Values represent means ± SEM *: p<0.05.
be affected by local flow patterns, but was essentially identical in all positions. In long-term experiments, pulsatile flow-induced TF activation was maintained at the maximal level for at least 48 hours, again in contrast to VCAM-1, which under the same conditions is transiently upregulated and returns to baseline after approx 48 h (Figure 11.11). Continued upregulation of TF by low-shear pulsatile flow in cultured EC is in line with the in vivo observations of enhanced thrombogenicity of the vascular wall at predilection sites of atherosclerosis (Grabowski and Lam, 1995). Preincubation of the cells with 10 µM PDTC for 30min completely abrogated the subsequent stimulation (for 5 h in the continued presence of PDTC) of TF activity by either pulsatile flow or, as also reported by others, by 1 ng/ml TNF (not shown). This finding might suggest the involvement of the PDTC-sensitive nuclear transcription factor NF-B in both activation processes (Marui et al., 1993; Lan, Mercurius and Davis, 1994; Orthner, Rodgers and Fitzgerald, 1995; Read et al., 1994; Gerritsen and Bloor, 1993; Shi et al., 1998). However, as described above, NF-B does not seem to be activated by pulsatile flow (see above, Figure 11.5). This finding, together with similar results obtained for the pulsatile-flow induced upregulation of VCAM-1, raises the question whether PDTC might affect other cellular (oxidative?) processes which are involved in gene activation, but independent of NF-B. On the other hand, pulsatile flow does activate some other transcription factors, such as egr-1 and SP-1. Indeed, the upregulation of egr-1, but not of SP-1 is positionally dependent (Figure 11.6). Since SP-1 has recently been shown to be involved in the steady laminar flow-induced upregulation of the TF gene (Lin et al., 1997), we propose that this transcription factor and the associated mechano-response element are also involved in the upregulation of TF by pulsatile, non-laminar flow.
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Figure 11.11. Pulsatile, non-laminar flow induces sustained increase in TF activity in HAEC. Human aortic endothelial cells were exposed to pulsatile flow, as described above. At various time points TF activity and VCAM-1 expression were assessed in parallel cultures amidolytically and by ELISA, respectively. The data are normalized to TF activity and VCAM-1 expression under static conditions as determined separately for each time point. Values represent means ± SEM.
Our studies suggest, that a number of different, perhaps novel, nuclear tran– scription factors and/or signaling mechanisms might be involved in the activation of TF expression/activity by the various hemodynamic forces. The distinct patterns by which the different hemodynamic forces activate TF expression/activity in HAEC are summarized in Table 11.1. These data further hint to a certain degree of diversity in terms of cellular mechanoreception and its post-“receptor” signaling mechanisms, which seem to be distinct for the different hemodynamic forces.
CONCLUSIONS Hemodynamic forces are some of the most effective regulators of the genotypic and phenotypic profile of vascular EC. The regulation of EC surface proteins in response to hemodynamic forces is complex and quite distinct from that following activation of the cells by cytokines. Hemodynamic forces can affect the function of endothelial cells through alterations at the gene level by inducing changes in the physicochemical state of the membranes, as well as through post-translational modifications. As new mechanical-force sensitive genes/gene products are discovered, the repertoire of novel “mechano-sensitive response elements” is expanding, while other, known cellular signaling mechanisms become an integral part of the mechanotransducing machinery.
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Table 11.1 Differential modulation of TF activity by hemodynamic forces
not determined in our preliminary studies TF mRNA is upregulated under laminar flow conditions (Line et al., 1997). bNF-B is upregulated under unidirectional, laminar flow-conditions (Lan Mercuricus and Davies, 1994). cNF-B is upregulated by cyclic strain (Du, Mills and Sumpio, 1995). * a
Mechanosensitivity seems to be conferred by a variety of signaling pathways involving distinct transcription factors, which are differentially activated by various hemodynamic forces. For example, in the case of tissue factor, cyclic-strain seems to act via NF-B, while flow, irrespective of whether steady or pulsatile, appears to activate SP-1. Regulation of other genes, such as VCAM-1, seems to be exquisitely sensitive to different flow patterns, while being only minimally, if at all, affected by other mechanical forces. Finally, for a given gene/gene product, the mechanisms of mechanoreception and -regulation depend not only on the type of hemodynamic force applied, but is also to a large degree tissue-specific, i.e. depends on the anatomical origin of the endothelial cells under investigation. ACKNOWLEDGMENTS Over the years, our work has been supported by grants from the Milwaukee Heart Research Foundation, the American Heart Association (National Center and Wisconsin Affiliate), the National Aeronautics and Space Administration, and Berlex Biosciences. I would like to acknowledge the longstanding collaboration with Brian R.Unsworth (Marquette University), the exciting co-operation with Gabor M.Rubanyi and Drew A.Sukovich (Berlex Biosciences), as well as the important contributions made by current and former members of my laboratory: Mishel Davis, G.Thomas Hayman, Keiichi Kanda, Claudia Kettlun, Vangelis Manolopoulos, Mark M.Samet, Matthew D.Silverman, Dawn M.Wankowski, John Wigboldus and Shaosong Zhang. REFERENCES Acevedo, A.D., Bowser, S., Gerritsen, M.E. and Bizios, R. (1993) Morphological and proliferative responses of endothelial cells to hydrostatic pressure: role of fibroblast growth factor. Journal of Cellular Physiology, 157, 603–614. Aird, W.C., Jahroudi, N., Weiler-Guettler, H., Rayburn, H.B. and Rosenberg, R.D. (1995) Human von Willebrand factor gene sequences target expression to a subpopulation of endothelial cells in transgenic mice. Proceedings National Academy of Science USA, 92, 4567–4571.
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Silverman, M.D., Manolopoulos, V.G., Unsworth, B.R. and Lelkes, P.I. (1996) Tissue factor expression is differentially modulated by cyclic mechanical strain in various human endothelial cells. Blood Coagulation and Fibrinolysis, 7, 281–288. Silverman, M.D., Samet, M.M, and Lelkes, P.I. (1996) Pressure modulates tissue factor expression in human aortic and vena cava endothelial cells. Journal of Vascular Research, 33, S93. Silverman M.D., Kanda, K., Hayman, G.T., Samet, M.M., Wigboldus, J. and Lelkes, P.I. (1997) Influence of hemodynamic forces on the expression of activation molecules in cultured human endothelial cells. The Japanese Journal of Artificial Organs, 26, S172 (Abstract). Silverman, M.D, Kanda, K., Hayman, G.T. et al. (1998) Effect of pressure on the expression of tissue factor activity, cell adhesion molecules, and fibroblast growth factor in cultured human endothelial cells. Sixth Biennial Meeting of International Society for Applied Cardiovascular Biology, 1–10 (Abstract). Stillwell, W., Wassall, S.R., Dumaual, A.C., Ehringer, W.D., Browning, C.W. and Jenski, L.J. (1993) Use of merocyanine (MC540) in quantifying lipid domains and packing in phospholipid vesicles and tumor cells. Biochimica et Biophysica Acta, 1146, 136–144. Sumpio, B.E. and Banes, A.J. (1988) Prostacyclin synthetic activity in bovine aortic endothelial cells undergoing cyclic mechanical deformation. Surgery, 104, 383–389. Sumpio, B.E., Banes, A.J., Buckley, M. and Johnson, G. Jr. (1988) Alterations in aortic endothelial cell morphology and cytoskeletal protein synthesis during cyclic tensional deformation. Journal of Vascular Surgery, 7, 130–138. Sumpio, B.E., Banes, A.J., Link, G.W. and Iba, T. (1990) Modulation of endothelial cell phenotype by cyclic stretch: inhibition of collagen production. Journal of Surgical Research, 48, 415–420. Sumpio, B.E., Widmann, M.D., Ricotta, J., Awolesi, M.A. and Watase, M. (1994) Increased ambient pressure stimulates proliferation and morphologic changes in cultured endothelial cells. Journal of Cellular Physiology, 158, 133–139. Sumpio, B.E., Du,W. and Xu, W. (1995) Exposure of endothelial cells to cyclic strain induces c-fos, fosB and c-jun but not jun B or jun D and increases the transcription factor AP-1. Endothelium, 2, 149– 156. Sumpio, B.E., Du,W., Galagher, G. et al. (1998) Regulation of PDGF-B in endothelial cells exposed to cyclic strain. Arteriosclerosis, Thrombosis, and Vascular Biology, 18, 349–355. Sung, C., Arleth, A.J. and Nambi, P. (1994) Evidence for involvement of protein kinase C in expression of intracellular adhesion molecule-1 (ICAM-1) by human vascular endothelial cells. Pharmacology, 48, 143–146. Susic, D. (1997) Hypertension, aging, and atherosclerosis—The endothelial interface. Medical Clinics of North America, 81, 1231–1240. Suttorp, N., Fuchs, T., Seeger, W., Wilke, A. and Drenckhahn, D. (1989) Role of Ca2+ and Mg2+ for endothelial permeability of water and albumin in vitro. Laboratory Investigation, 61, 183–191. Tardy,Y., Resnick, N., Nagel, T., Gimbrone, M.A. Jr. and Dewey, C.F. Jr. (1997) Shear stress gradients remodel endothelial monolayers in vitro via a cell proliferation-migration-loss cycle. Arteriosclerosis, Thrombosis, and Vascular Biology, 17, 3102–3106. Tedder, T.F., Steeber, D.A., Chen, A. and Engel, P. (1995) The selectins: vascular adhesion molecules. FASEB Journal, 9, 866–873. Terry, C.M. and Callahan, K.S. (1996) Protein kinase C regulates cytokine-induced tissue factor transcription and procoagulant activity in human endothelial cells. Journal of Laboratory and Clinical Medicine, 127, 81–93. Thoumine, O., Nerem, R.M. and Girard, P.R. (1995) Oscillatory shear stress and hydrostatic pressure modulate cell-matrix attachment proteins in cultured endothelial cells. In Vitro Cellular and Developmental Biology: Animal, 31A, 45–54. Tokunaga, O. and Watanabe, T. (1987) Properties of endothelial cell and smooth muscle cell cultured in ambient pressure. In Vitro, Cellular and Developmental Biology, 23, 528–534. Topper, J.N., Cai, J.X., Falb, D. and Gimbrone, M.A. Jr. (1996) Identification of vascular endothelial genes differentially responsive to fluid mechanical stimuli: Cyclooxygenase-2, manganese superoxide dismutase, and endothelial cell nitric oxide synthase are selectively up-regulated by steady laminar shear stress. Proceedings of the National Academy of Sciences USA, 93, 10417–10422. Topper, J.N., Cai, J., Qiu, Y. et al. (1997a) Vascular MADs: Two novel MAD-related genes selectively inducible by flow in human vascular endothelium. Proceedings National Academy of Science USA, 94, 9314–9319. Topper, J.N., Wasserman, S.M., Andersen, K.R., Cai, J., Falb, D. and Gimbrone, M.A. Jr. (1997b) Expression of the bumetanide-sensitive Na-K-Cl cotransporter BSC2 is differentially regulated by fluid mechanical and inflammatory cytokine stimuli in vascular endothelium. Journal of Clinical Investigation , 99, 2941–2949.
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12 Endothelium and Cyclic Strain Ira Mills and Bauer E.Sumpio* Department of Surgery, Yale University School of Medicine, 333 Cedar Street, P.O. Box 208062, New Haven, CT06510, USA. *Corresponding author: E-mail address: [email protected].
The objective of this chapter is to review studies conducted in our laboratory and others describing the effect of cyclic strain on endothelial cell biology. In particular, the focus of this review is on the delineation of putative mechanosensors, relevant signaling pathways, and strain-sensitive genes. We first report recent data from our laboratory supporting the involvement of focal adhesion plaques and integrins in the sensing of cyclic strain by endothelial cells. We also address recent studies performed in our laboratory examining the effect of cyclic strain on intracellular [Ca++] and it’s temporal relationship to IP3 generation as well as exciting new data of strain-induced ERK1/ERK2 activation. Shown are the distinct mechanisms of activation of strain-sensitive genes (e.g., PDGF-B and tPA) by cyclic strain versus shear stress. In addition, we provide recent, corollary data from our laboratory showing cyclic strain activation of keratinocyte cell biology. KEYWORDS: Endothelium, cyclic strain, focal adhesion molecules, gene promoter, transcription factors, keratinocytes.
INTRODUCTION Our laboratory and others have been rigorously studying the influence of mechanical forces on endothelial cell biology. It has been our contention that static conditions commonly utilized to study endothelial cells in culture may not reflect their in vivo milieu. Endothelial cells in vivo are subjected to a variety of flow-related forces including shear stress, hydrostatic pressure, and cyclic strain (1–2). Shear stress is the tangential stress applied across the endothelial cell surface due to the bulk flow of blood. Hydrostatic pressure is the normal stress acting radially on the vessel wall due to the propagation of the pressure wave. Cyclic strain represents the stress acting along the vessel wall due to circumferential deformation. Endothelial cells are exposed to all three forces whereas the underlying smooth muscle cells are exposed to hydrostatic pressure and cyclic strain. There are a several reviews of the effects of shear stress on endothelial cells in this text (see Chapters XX). Therefore, we will mention shear studies briefly and solely for comparison purposes to that of cyclic strain. Cyclic strain, along with the other forces listed above, play an important role in regulation of vascular tone, remodeling, and the genesis of atherosclerosis in vivo (3– 4). The mechanism by which cultured vascular endothelial cells perceive cyclic strain, utilizing in vitro devices to model this force, is the subject of this chapter. The strains described for the in vitro studies herein (~10% strain) are functionally significant 249
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based on in vivo models that replicate the major geometric features of blood vessels. These studies report a 5–6% wall excursion at peak systole under normal conditions, which can increase to 10% in the hypertensive state (5–7). Over the years, we have acquired a large body of data to suggest that the study of vascular cells challenged with mechanical stimuli is indeed more appropriate for studying their biology (8–21). This chapter represents an update of our earlier reviews of the literature describing the effect of cyclic strain on vascular cell biology (12)(22). Data obtained from our laboratory and that of others have begun to delineate the signaling pathways that may be involving in affecting vascular cell phenotype and is commented on in great detail in this review. We also discuss the recent discovery of strain-sensitive genes, in particular tPA and PDGF. These appear to be finely tuned in their ability to distinguish mechanical perturbations based on unique strain-sensitive or shear sensitive cis-elements in their promoter regions that respond to unique strainactivated or shear-activated transcriptional factors that either induce or repress gene induction. The nature of the mechanotransducer remains unsolved. However, recent studies from our laboratory and others are beginning to unravel integrins as likely candidates. In the “tensegrity model” as proposed by Ingber (23), any mechanical force acting on the cell is balanced between tensile actin filaments and the extracellular matrix anchoring proteins in the focal adhesion plaque. Since integrins play a pivotal role in maintaining this equilibrium, they represent mechanical transducers (see Chapter by Shyy and Chien in this volume (Chapter XX) and (24)). In support of this hypothesis, they have recently demonstrated the involvement of integrins in endothelial responses to shear stress. Likewise, we have accumulated recent data that also implicate integrins in endothelial responses to cyclic strain as outlined in the following chapter. Highlighted in this chapter are recent discoveries consistent with a role of integrins as mechanotransducers such as strain-induced integrin reorganization, focal adhesion kinase phosphorylation, and phosphorylation and activation of MAP kinase.
EFFECT OF CYCLIC STRAIN ON ENDOTHELIAL CELL BIOLOGY General Phenotype Previous studies have shown that cyclic strain can alter the general phenotype of endothelial cells (see Table 12.1). Strain-induced changes in endothelial cells include their proliferative rate (16)(25), morphology (9)(26–28) and the secretion of macromolecules such as prostacyclin (18)(29), endothelin (19), nitric oxide (30–31), tissue plasminogen activator (10)(21)(32) and plaminogen activator inhibitor-1 (PAI1) (33). Changes in endothelial cell phenotype are also modulated by shear stress (see Table 12.1). Changes include alterations in proliferation, morphology and the synthesis and secretion of many of the same proteins influenced by cyclic strain (34–36). However, the nature of the changed phenotype can be markedly different whether the perturbation is either shear stress or cyclic strain. For example, the orientation of endothelial cells in response to shear stress is parallel to the direction of flow (37). In
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contrast, cyclic strain causes a perpendicular alignment of endothelial cells in relationship to the strain vector (9). More recent studies performed in our laboratory support the involvement of focal adhesion plaques in the transmission of cyclic strain to the cell. Yano et al. (28) demonstrated phosphorylation of cytoskeletal proteins that reside at the cytoplamic face of the focal adhesion plaques, namely pp125FAK (Figure 12.1; taken from (28)) and paxillin (Figure 12.2; taken from (28)). The strain-induced phosphorylation of these proteins occurs on tyrosine residues and can be inhibited by specific tyrosine kinase inhibitors. The tyrosine phosphorylation of pp125FAK occurs earlier and is more robust than that observed with paxillin. However, after a four hour exposure to strain, both cytoskeletal proteins were found to align as shown by confocal microscopy Table 12.1 Effect of cyclic strain and shear stress on endothelial cell phenotype. Adapted from (34)
Figure 12.1. Changes in tyrosine phosphorylation of pp125FAK induced by cyclic strain. EC were subjected to strain for 0, 15s, and 1, 5, 10, and 30 min (a and b) or 0, 30 min, and 4 h (c and d). The pp125FAK was immunoprecipitated from cell lysates with anti-pp125FAK Mab, and immunoprecipitates were analyzed by immunoblotting with anti-pp125FAK Mab (a and c) or py-20 (b and b). Although the amount of pp125FAK was unchanged (a and c), the level of phosphotyrosine in pp125FAK increased after 30-min and 4-h strain.
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Figure 12.2. Changes in tyrosine phosphorylation of paxillin induced by cyclic strain. EC were stretched for 0, 15 s, and 1, 5, 10, and 30 min (a and b) or for 0, 30 min, and 4 h (c and d). Paxillin was immunoprecipitated from cell lysates with anti-paxillin MAb and was subjected to SDS-PAGE. Proteins were detected by immunoblotting with anti-paxillin MAb (a and c) and py-20 (b and d). Level of phosphotyrosine in paxillin slightly increased at 4 h after cells were exposed to cyclic strain (d). Arrowhead points to paxillin band. Bands under paxillin are immunoglobulin heavy chain bands.
(Figures 12.3 and 12.4; taken from (28)). This observation was in sharp contrast to their random distribution found in stationary controls. Tyrphostin A25, a tyrosine kinase inhibitor, blocked the strain-induced phosphorylation of pp125FAK and paxillin as well as their strain-induced alignment. Moreover, the importance of focal adhesion proteins on the mechanically induced gross morphological changes of endothelial cells was implicated by the ability of tyrphostin A25 to abolish both cellular alignment and migration caused by strain. Davies et al. (38) have studied the effect of shear stress on focal adhesion sites in bovine aortic endothelial cells. By tandem scanning confocal microscopy and digitized image analysis, they found a remodeling of focal adhesion sites in the direction of flow. In addition to this response, shear stress caused a redistribution of intracellular stress fibers, alignment of individual focal adhesion sites, and the coalescing of smaller sites to larger, and fewer focal adhesions per cell. Yano et al. (39) recently studied the involvement of different integrins in signaling induced by cyclic strain since integrins have been localized to focal adhesion sites. Furthermore, these focal adhesion sites are known to exhibit pp125FAK phosphorylation either by cyclic strain as well as by integrins. Cyclic strain of 4 hours led to a redistribution of and ß integrins in human umbilical vein endothelial cells (Figure 12.5; taken from (39)). In addition, ß1 integrin reorganized in a linear pattern parallel with the long axis of the elongated cells creating a fusion of focal adhesion plaques in cells plated on fibronectin (a ligand for 5ß1) or collagen (a ligand for 2ß1) coated plates. In contrast, the vitronectin
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Figure 12.3. Confocal immunofluorescent microscopy localization (A) and phase contrast images (B) of pp125FAK in EC. EC exposed to cyclic strain for various times with (f–h) or without (a–e) 100 µM tyrphostin were stained with anti-pp125FAK MAb. All cells except those in e, are cells in the periphery of the membrane. Cells in e, are those in the center of the membrane at 4 h of strain. After 30 min of strain (b), pp125FAK began to align with the same direction of the long axes of EC. After 4 h (c) and 24 (d) of exposure to strain, EC are elongated and pp125FAK is aligned to long axes of cells and was also observed in lamellipodia. Cells at the center of the membrane (e) and cells treated with tyrphostin and subjected to 4 and 24 h of strain (g and h, respectively) showed a similar pattern of distribution of pp125FAK observed in static EC (a). Static EC treated with trypohostin did not show any different pattern of distribution (f) compared with static EC without tyrphostin (a). Bar, 10 µM.
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Figure 12.4. Confocal immunofluorescent microscopy localization (A) and phase contrast images (B) of paxillin in EC. EC exposed to cyclic strain for various times with (f–h) or without (a–e) 100 µM tyrphostin were stained with anti-paxillin MAb. EC were exposed to strain for 30 min (b), 4 h (c, e and g), and 24 h (d and h). Static cells with tyrphostin (f) show no difference from static cells without tyrphostin (a). All cells except those in e are cells in periphery of membrane, although cells in e are cells in center of the membrane after they were subjected to 4-h strain. In static cells (a), paxillin reveals random speckle pattern. After 30 min of strain (b), in some cells paxillin began to align, although other cells show similar pattern of distribution in static cells. After 4 (c) and 24 h (d) of exposure to strain, EC are elongated and paxillin is aligned to long axes of cells. Cells in the center of the membrane (e) and cells treated with tyrphostin (f–h) showed a similar pattern of distribution observed in static. Bar, 10 µm.
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Figure 12.5. Spatial redistribution of a5 and a2 integrin induced by cyclic strain. HUVEC were seeded on fibronectin (A, C, E, G) or collagen I (B, D, F, H) coated membrane and were subjected to cyclic strain for 4 h (C, D, G, H) or maintained in stationary culture conditions (A, B, E, F). HUVEC were stained with anti-a5 (A–D) or anti-a2 integrin (E–H) and observed by confocal microscopy. In the static group, the cells cultured on fibronectin (A, E) or collagen (B, F) show a random speckle distribution of a5 and a2 integrin. After 4 h exposure to cyclic strain, cells were elongated and aligned perpendicular to the force vector (from upper left to bottom right). a5 integrin in the cells grown on fibronectin (C) and a2 integrin in the cells grown on collagen I (H) reorganized in a linear fashion, although a5 integrin in the cells on collagen (D) and a2 integrin in the cells on fibronectin (G) show a random punctate distribution similar to those in the static cells (A, B, E, F). Scale bar=10 µn.
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receptor, ß3 integrin, failed to redistribute in endothelial cells subjected to cyclic strain. Cyclic strain also caused a reorganization of a5 and a2 integrins in a linear pattern on cells seeded on fibronectin and collagen-coated plates, respectively. However, the expression of 5, 2, and ß1 was not influenced by 24 hours of cyclic strain as assessed by immunoprecipitation of these integrins. Strain-induced phosphorylation of pp125FAK occurred concomitantly with the reorganization of ß1 integrin. Thus, 5ß1 and 2ß1 integrins may play an important role in transducing mechanical stimuli into intracellular signals (39). Thoumine et al. (40) demonstrated effects on the organization and composition of the extracellular matrix upon subjecting endothelial cells to shear stress. They examined the pattern and levels of various extracellular matrix proteins including fibronectin, laminin, type IV collagen, and vitronectin. Of these, fibronectin was found to be the most affected by shear stress with a thickening and alignment of its fibril tracts. Moreover, the level of fibronectin increased after 1 or 2 days of shear stress after an earlier reduction at 12 hours. The extracellular matrix response to shear was found to be specific for fibronectin since both laminin and type IV collagen showed thickening of fibrils, but no alignment. Vitronectin was unaffected in any respect. Cell Signaling (see Table 12.2a and Table 12.2b) [Ca++]i, inositol phosphates and diacylglycerols Lansman et al. (52) demonstrated single stretch-activated non-selective cation channels in porcine aortic endothelial cells that has been supported by some studies (53–55) but not others (56). In our laboratory, previous studies demonstrated that cyclic strain causes a rapid but transient generation of 1, 4, 5 inositol trisphosphate (IP3), which peaked by 10 seconds of cyclic strain. To address the effect of cyclic strain on intracellular [Ca++] and its temporal relationship to IP3 generation, bovine aortic endothelial cells were grown on flexible membranes, loaded with aequorin and the membranes placed in a custom designed flow through chamber (57). The chamber was housed inside a photo multiplier tube and vacuum was utilized to deform the membranes. The initiation of strain caused a rapid increase in intracellular calcium of two components. The first component was a large initial peak 12 seconds after the initiation of stretch that closely followed the IP3 peak. The second component was a subsequent lower but sustained phase. Pre-treatment with 5 µM gadolinium for 10 minutes or nominally calcium-free medium for 3 minutes reduced the magnitude of the initial rise and abolished the sustained phase. Shear stress has been demonstrated to induce calcium transients in single endothelial cells as measured by fura-2 (58). Geiger et al. (58) studied the effect of shear stress at 30 dynes/cm2 in endothelial cells subjected to flow in parallel-plate flow chambers and glass capillary tubes. They found a rapid increase in [Ca++] that peaked by 30 seconds and decreased slowly to a plateau that persisted for greater than 5 minutes. Recent studies performed in our laboratory show that cyclic strain causes a biphasic increase in diacylglycerol (59). These data support a mechanism of phosphatidylcholine
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hydrolysis and diacylglycerol generation at both an early and late phase. Cyclic strain was shown to stimulate phospholipase C acutely and phospholipase D activity in an immediate and sustained manner (59). Bhagyalakshmi et al. (60) also showed stimulation of DAG by shear stress. In another study of shear stress-activated platelets, pathological levels of shear (i.e., 90 dynes/cm2) failed to elevate DAG despite increased PKC activity (61). Adenylyl cyclase Cyclic strain causes activation of the adenylyl cyclase/cyclic AMP/protein kinase A pathway in bovine aortic endothelial cells (15). In membranes deformed with 150 mm Hg (average 10% strain) vacuum at 60 cycles per minute (0.5 s strain; 0.5 s relaxation) for 15 minutes, there was a 1.5 to 2.2 fold increase in adenylyl cyclase, cAMP and PKA activity as compared to unstretched controls. The straininduced activation of this pathway appears to occur by exceeding a strain threshold since no change in adenylyl cyclase, cyclic AMP or PKA was observed in cells subjected to 37.5 mm Hg (average 6% strain). Further studies demonstrated an increase in cAMP response element activity as shown by gel shift analysis. Thus, cyclic strain may stimulate the expression of genes containing cAMP-responsive elements. We next tested the hypothesis that cyclic strain modulates G protein function, since G proteins are intimately involved in the activation of adenylyl cyclase (Mills, unpublished observations). To test this hypothesis, we examined the effect of acute cyclic strain on the immunoreactivity of the alpha subunits of the heterotrimeric G proteins, Gs and Gi, that promote stimulation and inhibition of AC activity, respectively. We observed a transient decrease in the immunoreactivity of the inhibitory G protein alpha subunits (Gi1,2). In contrast, immunoreactivity of Gs in bovine aortic endothelial cells and Gi1,2 and Gs in bovine aortic smooth muscle cells were unaffected by cyclic strain. Frangos and colleagues (62) have shown shear stress activation of G proteins, particularly Gq/11 and Gi3/o. This was determined by labeling of flowstimulated G proteins with a nonhydrolyzable GTP photoreactive analogue. The rapidity by which this response is obtained (1 sec) suggests a key role of G proteins in the mechanotransduction of shear stimuli. Previous studies by the same group further established a role of G proteins in the responsiveness of endothelial cells to shear stress. Both shear stress mediated prostacyclin and nitric oxide production were found to be mediated by pertussis-toxin sensitive and insensitive G proteins, respectively (63–64). Protein phosphatase activity As described above, we have reported previously that PKC in the membrane fraction of endothelial cell lysates is activated in response to cyclic strain (65). To better understand cellular responses that are dependent on the phosphorylated state of proteins, it is also important to study the role of protein phosphatases. In a recent study (66), we examined the effect of cyclic strain on protein phosphatase 1 and 2A activity in bovine aortic endothelial cells. Protein phosphatase 2A activity in the cytosol
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was decreased by 36.1% in response to 60 minutes of cyclic strain, whereas activity in the membrane was unchanged. Furthermore, treatment with a low concentration of okadaic acid (0.1 nM) enhanced proliferation of both static and stretched endothelial cells in 10% fetal bovine serum. These data suggest that protein phosphatase 2A acts as a growth suppresser and cyclic strain may enhance cellular proliferation by inhibiting protein phosphatase 2A (66). MAP kinase Extracellular signal regulated kinases 1/2 (ERK 1/2) play an important role in the transduction of cellular mitogenic and differentiation signals from the plasma membrane to the nucleus (67–68). Since endothelial cells exposed to cyclic strain demonstrate elevated proliferation and align perpendicular to the strain gradient, we examined whether cyclic strain phosphorylates and activates ERK 1/2 and also whether inhibition of this pathway could reverse these strain-dependent responses (69). In synchronized endothelial cells subjected to acute cyclic strain, we observed a time and strain-dependent phosphorylation and activation of ERK 1/2 with a peak phosphorylation at 10 min. Treatment of endothelial cells with BHQ to deplete inositol triphosphate-sensitive calcium storage and gadolinium chloride, a putative nonselective cationic blocker of activated Ca2+ channels, did not inhibit the cyclic strain induced activation of ERK 1/2. However, blockade of PKC by calphostin C partially inhibited strain-activated ERK 1/2, whereas inhibition of tyrosine kinase(s) with genistein completely inhibited strain-activated ERK 1/2. The significance of strain-activated ERK 1/2 in endothelial cells (69) and smooth muscle cells (70) remains unclear as blockade of this response with the MEK inhibitor, PD98059, failed to prevent strain-induced proliferative and morphological changes. Like cyclic strain, shear stress also causes phosphorylation and activation of ERK 1/2 and in Ca++-independent manner that may involve PKC activation (71–72) (see Chapter XX by Berk et al.). Berk et al. (73) proposed calcium-independent ERK 1/2 activation in bovine aortic endothelial cells subjected to shear stress. In addition, Tseng et al. (71) demonstrated that activation of ERK 1/2 by shear stress is dependent on PKC activation. Transcriptional Factor Activation (see Table 12.3a and 12.3b) In a recent study, Cheng et al. (33) demonstrated strain-induced activation of PAI-1 secretion. Although this effect could be accomplished directly on the secretory process, the authors postulate the upregulation of PAI-1 gene via a functional AP-1 binding site in its promoter regions. Interestingly, the strain-induced secretion of PAI-1 was shown to involve reactive oxygen species, a known activator of both AP-1 and NF-B transcription factors. Further studies will be required to test this hypothesis. However, studies performed in our laboratory support the hypothesis of increased activity of transcriptional activators in endothelial cells subjected to cyclic strain (77). Since previous studies demonstrated that cyclic strain stimulates protein kinase C activity in endothelial cells (65) we tested the hypothesis that downstream induction
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Table 12.2a Effect of cyclic strain on mechanosignaling in endothelial cells. Adapted from (34)
Table 12.2b Effect of shear stress on mechanosignaling in vascular endothelial cells. Adapted from (34)
of the fos and jun genes and the transcription factor activator protein-1 (AP-1) were also activated by strain. Sumpio et al. (20) showed that human umbilical vein endothelial cells subjected to cyclic strain for as short a duration as 30 minutes leads to the induction of c-fos, fosB and c-jun genes. In contrast, junB and junD were not altered by cyclic strain under identical conditions. We did detect an increase in AP-1 binding activity as measured by EMSA conducted after 2 hours of cyclic strain. It may be concluded that cyclic strain may be coupled to the endothelial cell response via activation of protein kinase C and elevated steady state levels of different fos and jun products, which may enhance the activity of the transcriptional activator, AP-1 (20).
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In more rigorous studies performed to characterize the induction of transcriptional factor activation, AP-1, CRE binding protein and NF-B binding activity were found to be influenced by cyclic strain in a species and vascular bed-dependent manner (77). In human umbilical vein endothelial cells subjected to cyclic strain, AP-1 activity was elevated as compared to unstretched controls (20). The onset of strain-induced AP-1 activity was at 2 hours, peaked by 4 hours, and returned to baseline levels by 24 hours. CRE levels were significantly stimulated in human umbilical vein endothelial cells in a biphasic manner with a 2-fold increase at 15 minutes and a nearly 5-fold increase at 24 hours. NF-B binding activity was stimulated in a monophasic manner in endothelial cells obtained from umbilical vein with a 4.6-fold increase at 4 hours. Similar findings were obtained in human aortic endothelial cells but not in bovine aortic endothelial cells (77). However, in a later study we did show evidence for strain-induced stimulation of CRE binding activity in bovine aortic endothelial cells (15). In this study, CRE binding activity was found by 30 minutes and diminished by 4 hours. Selective seeding of the endothelial cells to the high strain region (7–24% strain) led to a greater response than found in those cells grown in the center, low strain region (0– 7%). An increase in CRE binding activity was not observed in cells subjected to an average 6% strain. The reasons for observation of a strain response in stimulation of CRE binding activity of bovine aortic endothelial cells in this study (15) but not the earlier one (77) is unclear. However, it may suggest other variables aside from species and vascular bed diversity that remain to be uncovered. The nature of the mechanical force may also be critical in activating a particular transcription factor. For example, Lan et al (78) and Resnick et al. (79) have both shown NF-B activation in bovine aortic endothelial cells subjected to shear stress; a finding we have not been able to replicate in response to cyclic strain (see Chapter XX by Resnick et al.).
Table 12.3a Effect of cyclic strain on transcription factor activation in endothelial cells. Adapted from (34)
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Table 12.3b Effect of shear stress on transcription factor activation in endothelial cells. Adapted from (34)
Gene Expression PDGF In a recent study, we have demonstrated that cyclic strain stimulates PDGF-B expression in bovine aortic endothelial cells (81). In response to 4 hours of 10% strain, but not 6% strain, PDGF-B steady state mRNA levels by nearly 3-fold (Figure 12.6; taken from (81)). This strain-dependent activation of PDGF-B mRNA was via induction of new transcripts as determined by nuclear run-off transciption analysis. Immunoreactive PDGF-B protein was also found to be upregulated in a strain-dependent manner, such that cells resided in the high strain areas demonstrated more intense staining as compared to cells present in the center, low strain region of the membrane. We next examined the regulation of PDGF-B expression by cyclic strain, by conducting transfection studies with a construct containing a 450 bp fragment of the PDGF-B promoter coupled to chlorophenicol acetyltransferase (CAT) (81). A direct influence of cyclic strain on PDGF-B transcription was confirmed by our demonstration of 2-fold induction of PDGF-B promoter activity at 4 and 8 hours. Further transfection studies of bovine endothelial cells with a series of deletion constructs demonstrated a 55% drop-off in activity between position –313 and –153 with no induction of activity with the 101 bp minimal promoter (Figure 12.7 from (81)). Cyclic strain, as shown earlier for shear stress, stimulates the induction of an SSRE binding protein, however the binding site at position –125 does not appear to be necessary for the strain-induced activation of PDGF-B. This was determined by transfection of endothelial cells with a hybrid promoter construct containing the SV40 sequence promoter downstream of the SSRE or the –153 PDGF-B promoter sequence bearing a mutation in the SSRE. In neither case, was there any significant activation by either promoter. Further studies are required to delineate the critical strain-induced binding sites in the PDGF-B promoter, but a functional role for elements upstream of the SSRE is implicated by our findings. We propose that strain-inducible PDGF-B expression may involve cooperativity between factors interacting with upstream regions and those binding to the SSRE. Our premise is consistent with the complexity of transcriptional control and convergence and divergence of multiple signaling pathways.
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Figure 12.6. PDGF-B expression in EC exposed to either 10% (A) or 6% (B) average strain at 60 cycles/ min. The top panels are typical Northern blots of EC demonstrating PDGF-B expression after 2, 4, 8, 12, or 24 hours of cyclic strain or after exposure to 25 ng/ml of phorbol-12-myristate13-acetate (PMA) for 4 hours. GAPDH is the constitutive control for loading. The bottom graphs represent the average fold induction calculated from the densitometry values. The fold induction is the PDGF-B value divided by the GAPDH value for the different regimens divided by control, static conditions. (C) Nuclear run-off study demonstrating PDGF-B transcript levels increased significantly in response to cyclic strain, whereas GAPDH transcript levels were unaffected.
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Figure 12.7. (A) Fold induction of activity of a 450 bp human PDGF-B promoter fused to a CAT reporter gene transfected into endothelial cells which were subjected to up to 24 hours of 6% or 10% average strain at 60 cycles/min, and then analyzed for CAT activity. The promoterless CAT vector (N) is a negative control and a CMV-driven CAT construct (P) represents a positive control. The folds of induction are the reporter CAT activity (normalized for transfection efficiency) in the experimental cells (strain) compared with those in static (control) cells. The mean of 4–6 separate experiments, *p<0.05. (B) Fold induction of CAT activity generated from PDGF-B promoter deletion mutants fused to a CAT reporter gene transfected into EC which were subjected to 4 hours of 10% average strain at 60 cycles/min. The lengths of the 5'-flanking sequences of the PDGF-B promoter fused to the CAT reporter gene are shown on the x-axis. The folds of induction are the reporter CAT activity (normalized for transfection efficiency) in the experimental cells (strain) compared with those in static (control) cells. The mean of 4–10 separate experiments, *p<0.05.
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A SSRE was found to be necessary and sufficient for increased PDGF-B promoterdependent expression in cultured endothelial cells subjected to laminar shear stress (79). In fact, laminar shear stress induces a 10-fold increase in PDGF-A chain mRNA and a 2–3 fold increase in B chain mRNA expression in both human umbilical vein (82) and bovine aortic endothelial cells (79)(83–84). Thus, our studies demonstrate that cyclic strain, like shear stress, can increase PDGF-B gene expression in bovine aortic endothelial cells. However, the mechanotransduction processes involved in regulation of the PDGF-B promoter are dependent on the nature of the mechanical stimuli (see Chapter XX by Resnick et al.). Endothelin Cyclic strain stimulates endothelin-1 secretion and mRNA levels in human umbilical vein endothelial cells (85). Elevated expression of endothelin 1 mRNA was detected in cells subjected to 2 hours or longer duration of cyclic strain. This was abolished by treatment with actinomycin D. The strain-induced elevation of ET-1 mRNA synthesis was found to mediated by the activation of the PKC pathway and was shown to require extracellular Ca++. The involvement of the PKC pathway was determined since the PKC inhibitor, calphostin C, prevented strain-induced entothelin-1 expression. In contrast, the cAMP-dependent protein kinase inhibitors, KT5720 or KT5823, only partially blocked endothelin-1 expression stimulation by cyclic strain. A requirement of extracellular Ca++ was determined by blockade of strain-induced endothelin-1 mRNA by EGTA as well as a lesser effect by BAPTA/ AM, an intracellular calcium chelator (85). In our hands, we have observed strain-dependent reduction in ET-1 (J.Koo et al., unpublished data). Similar contradictory effects of shear stress have also been detected in measurement of ET-1 (50–51)(86). Malek et al. (86) have shown shear stressmediated down-regulation of ET-1 expression in bovine aortic endothelial cells conferred by a cis element between –2.5 kb and –2.9 kb of the 5'-upstream promoter region that does not involve the AP-1 or GATA-2-factor binding site. In contrast, Morita et al. (50)(87) have shown shear stress-induced ET-1 expression in porcine aortic endothelial cells that appears to be related to cytoskeletal disruption. Prostacydin Previous studies indicate that cyclic strain (18) and shear stress (46)(88) can increase PGI2 secretion by endothelial cells but the effect of these forces on prostacyclin synthase (PGIS) gene expression remains unclear. In a recent study, we examined this question by studying PGIS gene expression by Northern blot analysis and protein level by Western blot analysis (29). In addition, the effect of cyclic strain on the PGIS promoter was determined by the transfection of a 1kb human PGIS gene promoter construct coupled to a luciferase reporter gene into EC, followed by determination of luciferase activity. PGIS gene expression increased 1.7 fold in EC subjected to cyclic strain for 24 hours (29). Likewise, EC transfected with a pGL3B-PGIS(–1070/–10) construct showed an approximate 1.3 fold elevation in luciferase activity in EC subjected to cyclic strain
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for 2, 4, 8 and 12 hours. The weak stimulation of PGIS gene expression by cyclic strain was reflected in an inability to detect alterations in PGIS protein levels in EC subjected to cyclic strain for as long as 5 days. These data suggest that strain-induced stimulation of PGIS gene expression plays only a minor role in the ability of cyclic strain to stimulate PGI2 release in EC. These findings coupled with our earlier demonstration of a requisite addition of exogenous arachidonate in order to observe strain-induced PGI2 release (18), implicates a mechanism that more likely involves strain-induced stimulation of PGIS activity. In recent studies, Kito et al. (89) studied the effect of cyclic strain on cyclo-oxygenase expression and promoter activity in bovine aortic endothelial cells (89). Despite the key role of COX in the synthesis of prostacyclin (PGI2) and thromboxane A2, cyclic strain was found to cause only a mild stimulatory effect of inducible COX-2 promoter activity with minimal effect on mRNA expression. Moreover, constitutive COX-1 promoter activity and mRNA expression were unaffected by cyclic strain. In contrast, laminar shear stress upregulates COX-2 transcription and translation in endothelial cells (90). Tissue plasminogen activator (tPA) Previous studies showed an increase in tPA mRNA, immunoreactive tPA protein and tPA activity in the medium of cultured bovine aortic endothelial cells exposed to 10% average strain (10)(32). Previous studies documented that tPA expression is upregulated by shear stress (48). We have more recently examined the regulation of tPA gene expression in endothelial cells by cyclic strain (21). A functional analysis of the tPA promoter was performed by transfecting bovine aortic endothelial cells with a 1.4 kb construct of the human tPA promoter coupled to CAT. After 4 hours of cyclic strain, a nearly 3-fold increase in the activity of the 1.4 kb tPA promoter was detected (Figure 12.8; taken from (21)). A 60% drop-off in activity was found between position –145 and –105 by analysis of deletion mutants. DNase I protection analysis of the segment downstream of position –196 suggested involvement of AP-2 or CRE-like regions, which was confirmed by EMS A analysis. Site directed mutants of either the AP-2 or CRE-like regions resulted in a 65% decrease in activity compared to the wild type. In addition, double mutations abolished basal transcription and any strain-induced activity. A SSRE binding site was found to present at –945, but site directed mutants failed to show any drop in strain-induced activity as compared to the wild type. Overall, our studies demonstrate that cyclic strain regulates tPA gene transcription in bovine aortic endothelial cells by a mechanism similar to that shown previously for phorbol ester. Nitric oxide Awolesi et al. (31) demonstrated that cyclic strain can upregulate the expression of endothelial nitric oxide synthase (eNOS) in bovine aortic endothelial cells. At an average strain of 10%, eNOS expression in these cells was shown to be increased as compared to unstretched controls as determined by Northern blot analysis. A milder elevation in eNOS expression was measured in cells exposed to a lesser degree of strain of 6%. The strain-induced activation of eNOS expression was attributed to
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Figure 12.8. (A) Nuclear runoff study demonstrating tissue plasminogen activator (tPA) transcript levels increased 2.2-fold in response to 4 h of cyclic strain, whereas glyceraldehyde-3-phosphate dehydrogenase (GAPDH) transcript levels were unaffected. Representative experiment was performed in duplicate. Con, control. (B) representative experiment in which bovine aortic endothelial cells (EC) were transfected with a tPA promoter (–1452 to +308) fused to a chloramphenicol acetyltranferase (CAT) reporter gene and then analyzed for CAT activity. Top two bands (arrow) are the acetylated CAT products; bottom band is the unmodified CAT. A positive control was stimulated with 25ng/ml of phorbol 12-myristate 13-acetate (PMA), the promoterless CAT vector (N) is a negative control, and the cytomegalovirus-driven CAT construct (P) represents another positive control. (C) multiples of induction activity of a tPA promoter (–1452 to +308) fused to a CAT reporter gene and then analyzed for CAT activity. Multiples of induction were determined as reporter CAT activity (normalized for transfection efficiency) in the experimental cells (strain) compared with those in static (control) cells. Results are means of 4 separate experiments. *p<0.05.
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stimulation of transcriptional activity as shown by nuclear runoff assays. Western blot analysis and immunochemistry confirmed that eNOS protein levels were similarly increased in response to strain; again 10% strain led to a heightened response as compared to 6% strain. Awolesi et al. (31) showed that the strain-induced upregulation of eNOS expression in bovine aortic endothelial cells translates into an increase in functional activity. In this study, 10% cyclic strain for 24 hours led to an increase in both citrulline production and accumulated nitrite (Greiss reaction) as indices of eNOS activity. Specificity of this response was confirmed by blockade with EDTA and LNAME. As is the case for strain-induced expression, strain-induced eNOS functional activity was more pronounced at 10% average strain as compared to 6% strain. The effect of 10% strain was as potent as that observed with the calcium ionophore, A23187 (30). Shear stress has also been shown to increase nitric oxide gene expression in endothelial cells (47)(90–91). In a recent study, Uematsu et al. (47) reported 2– 3-fold elevations in eNOS by shear stress of 15 dynes/cm2 for up to 1 day. The involvement of K+ channels was suggested since shear stress induction of nitric oxide mRNA was blocked by tetraethylammonium chloride, a K + channel antagonist. MCP-1 Wang et al. (92) studied the effect of cyclic strain on monocyte chemotactic protein1 (MCP-1) expression in cultured human umbilical vein endothelial cells (HUVECs). They found a two-fold enhancement in MCP-1 expression as early as one hour after the initiation of strain. This response was maintained for at least 24 hours in the presence of strain, but was restored to baseline levels 3 hours after its release. Calphostin C was able to abolish strain-induced MCP-1 expression, implicating a role of PKC as a mediator of this response. In addition, a strong Ca++ requirement for strain-induced, as well as basal expression of MCP-1, was determined by blockade with either EGTA pretreatment, BAPTA/AM chelation, or verapamil addition to HUVECs (92). Shear stress-induced expression of MCP-1 has been shown to involve a cis-acting TRE-responsive element in its promoter (93). A construct of multiple copies of the TRE-responsive element coupled to a prolactin minimal reporter and luciferase gene was found to be sufficient to confer shear sensitivity in transfected bovine aortic endothelial cells (see Chapter XX by Shyy et al.). ICAM/VCAM In recent studies conducted in our laboratory, we have obtained evidence to suggest that ICAM and VCAM expression is downregulated by cyclic strain in human umbilical vein endothelial cells (Lee et al., unpublished observations). Ando et al. (94) also demonstrated down-regulation of VCAM mRNA and protein expression in cultured mouse endothelial cells (see Chapter XX by Ando). In contrast, ICAM-1 expression is upregulated in the same cell type under conditions of shear stress (90)(95).
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EFFECT OF CYCLIC STRAIN ON KERATINOCYTE BIOLOGY Cultured cell responsiveness to cyclic strain is not restricted to vascular cells. The study of mechanical forces on keratinocytes is of interest since tissue expansion has been widely used for breast reconstruction, craniofacial surgery and burn care in plastic reconstructive surgery (96). Tissue expansion techniques involve enlarging the skin surface area with an expandable balloon. To date the effect of tissue expansion on in vivo skin physiology and histology has been well studied, but the direct effect of cyclic strain on keratinocytes remains unknown. In a series of recent reports, we have characterized the in vitro effect of cyclic strain on the phenotype and relevant signaling pathways of human keratinocyte obtained from neonatal foreskins (97–99). The phenotype of keratinocytes was markedly affected by cyclic strain (97). Chronic cyclic strain (1–8 days) caused an increase in keratinocyte proliferation, DNA synthesis, elongation, and protein synthesis at a regimen of 150 mm Hg at 10 cycles/min as compared to static controls. In contrast, continuous strain of a similar magnitude and duration was a modest stimulator of keratinocyte proliferation. In addition, as previously demonstrated for vascular endothelial and smooth muscle cells, keratinocytes aligned perpendicular to the strain gradient. Cyclic strain was found to modulate keratinocyte phenotype and cAMP mediated signaling pathways in an inverse manner (97). After 30 minutes of cyclic strain, both cAMP and protein kinase A activity were decreased by 30 and 45%, respectively, as compared to unstretched controls. We also observed a nearly 60% decrease in PGE2 levels after 5 days of cyclic strain (97). Although the relevance of these signaling pathways to the changes in keratinocyte phenotype is not established, these studies support the hypothesis that keratinocytes may play a key role in the observed effects of tissue expansion and other strain-dependent responses present in skin. Since protein kinase C is known to play an important role in the regulation of keratinocyte growth and differentiation, we also examined the effect of the PKC signaling pathway as a mediator of strain modulation of the keratinocyte phenotype (98). We demonstrated that cyclic strain increases PKC activity coupled with translocation of PKC from the cytosolic to the membrane fraction in keratinocytes as compared to static controls. In addition, PKC- and , but not ß nor isoforms, were translocated from the cytosolic to the membrane fraction as demonstrated by Western blot analysis and confocal microscopy. Studies with specific PKC inhibitors such as calphostin C and staurosporine implicate PKC involvement in cyclic strain-induced keratinocyte proliferation, but did not block the effects of strain on cellular morphology or alignment. Thus, cyclic strain stimulates PKC activity and translocation in an isoform-specific manner that may selectively modulate strain-induced changes in keratinocyte phenotype. Since keratinocytes are a major source of IL-1 which acts as a key modulator of keratinocyte processes including inflammatory responses, growth, and differentiation, we also examined the effect of cyclic strain on IL-1 expression in human keratinocytes (99). Both IL-1 and IL-1ß mRNA were upregulated by cyclic strain in keratinocytes, with a peak at 12 hours. Interestingly, IL-1 and IL-1ß antibodies blocked straininduced keratinocyte proliferation as well as basal proliferation but had no effect on strain-induced morphological changes. The pronounced effect of IL-1 antibodies on
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basal proliferation obscures its role in strain-induced proliferation and the precise role of IL-1 in mediating strain-induced changes in keratinocyte biology remains to be determined.
SUMMARY In summary, recent data support the hypothesis that vascular endothelial cells and other cell types such as keratinocytes are keenly sensitive to mechanical perturbation such as cyclic strain. Our group and others have characterized strain-induced changes in the endothelial cell phenotype and have begun to delineate the relevant signaling pathways and strain-sensitive genes (see Figure 12.9).
Figure 12.9. Schematic cartoon of mechanotransduction in ECs subjected to cyclic strain. As described in this review, we have recently reported a prominent role of the integrin-MAP kinase signaling pathway as mechanotransduction molecules. In ECs grown on a fibonectin matrix coating, ECs exhibit marked phosphorylation of FAK. Acute cyclic strain also stimulates ERK 1/2 phosphorylation known to be critically involved in the proliferation and differentiation of ECs. Downstream events known to be activated include elevated binding activity of various transcriptional factors (e.g., AP-1, CRE, and NF-B). Strain-sensitive promoter sites such as AP-2 and CRE-like regions have been shown to be important, in particular for the strain activation of tPA. Other strain-induced genes include PDGF-B, NO, and ET-1. Phenotypic changes in ECs in response to cyclic strain include stimulated proliferation and morphological changes (i.e., elongation and alignment).
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54. Naruse, K. and Sokabe, M. (1993) Involvement of stretch-activated ion channels in Ca2+ mobilization to mechanical stretch in endothelial cells. Am. J. Physiol., 264, C1037–C1044. 55. Sigurdson, W.J. Sachs, F. and Diamond, S.L. (1993) Mechanical perturbation of cultured human endothelial cells causes rapid increases of intracellular calcium. Am. J. Physiol., 264, H1745–H1752. 56. Morris, C.E. and Horn, R. (1991) Failure to elicit neuronal macroscopic mechanosensitive currents anticipated by single-channel studies. Science, 251. 57. Resales, O.R., Isales, C.M., Barrett, P.Q., Brophy, C. and Sumpio, B.E. (1997) Exposure of endothelial cells to cyclic strain induces elevations of cytosolic calcium through mobilization of intracellular and extracellular pools. Biochem. J., 326, 385–392. 58. Geiger, R.V., Berk, B.C., Alexander, R.W. and Nerem, R.M. (1992) Flow-induced calcium transients in single endothelial cells: spatial and temporal analysis. Am. J. Physiol., 262, C1411–C1417. 59. Evans, L., Frenkel, L., Brophy, C.M. et al. (1997) Activation of diacylglycerol in cultured endothelial cells exposed to cyclic strain . Am. J. Physiol., 272, C650–C656. 60. Bhagyalakshmi, A., Berthiaume, F., Reich K.M. and Frangos, J.A. (1992) Fluid shear stress stimulates membrane phospholipid metabolism in cultured human endothelial cells. J. Vasc. Res., 29, 443–449. 61. Kroll, M.H., Hellums, J.D., Guo, Z. et al. (1993) Protein kinase C is activated in platelets subjected to pathological shear stress. J. Biol. Chem., 268, 3520–3524. 62. Gudi, S.R.P., Clark, C.B. and Frangos, J.A. (1996) Fluid flow rapidly activates G proteinse in human endothelial cells. Involvement of G proteins in mechanochemical signal transduction. Circ. Res., 79, 834–839. 63. Kuchan, M.J., Jo, H. and Frangos, J.A. (1994) Role of G proteins in shear stress-mediated nitric oxide production by endothelial cells. Am. J. Physiol., 267, C753–C758. 64. Berthiaume, F. and Frangos, J.A. (1992) Flow-induced prostacyclin production is mediated by a pertussis toxin-sensitive G protein. FEBS Lett., 308, 277–279. 65. Rosales, O.R. and Sumpio, B.E. (1992) Protein kinase C is a mediator of the adaptation of vascular endothelial cells to cyclic strain in vitro. Surgery, 112, 459–66. 66. Murata, K., Mills, I. and Sumpio, B.E. (1996) Protein phosphatase 2A in stretch-induced endothelial cell proliferation. J. Cell. Biochem., 62, 1–9. 67. Robinson, M.J. and Cobb, M.H. (1997) Mitogen-activated protein kinase pathways. Curr. Opin. Cell Biol., 9, 180–186. 68. Whitmarsh, A.J. and Davis, R.J. (1996) Transcription factor AP-1 regulation by mitogen-activated protein kinase signal transduction pathways. J. Mol. Med., 74, 589–607. 69. Ikeda, M., Takei, T., Mills, I. and Sumpio, B.E. (1999) Cyclic strain stimulates mitogen-activated protein kinase activation in cultured bovine aortic endothelial cells. Am. J. Physiol. (In Press). 70. Mills, I., Ikeda, M. and Sumpio, B.E. (1997) MAP kinase activation by cyclic strain in vascular smooth muscle cells. FASEB J., 11, A216. 71. Tseng, H., Peterson, T.E. and Berk, B.C. (1995) Fluid shear stress stimulates mitogen-activated protein kinase in endothelial cells. Circ. Res., 77, 869–878. 72. Pearce, M.J., McIntyre, T.M., Prescott, S.M., Zimmerman, G.A. and Whatley, R.E. (1996) Shear stress activates cytosolic phospholipase A2 (cPLA2) and MAP kinase in human endothelial cells. Biochem. Biophys. Res. Comm., 218, 500–504. 73. Berk, B.C., Corson, M.A., Peterson, T.E. and Tseng, H. (1995) Protein kinases as mediators of fluid shear stress stimulated signal transduction in endothelial cells: a hypothesis for calcium-dependent and calcium-independent events activated by flow. J. Biomech., 28, 1439–1450. 74. Rosales, O.R. and Sumpio, B.E. (1992) Changes in cyclic strain increase inositol trisphosphate and diacylglycerol in endothelial cells. American Journal of Physiology, 262, C956–62. 75. Nollert, M.U., Eskin, S.G. and McIntire, L.V. (1990) Shear stress increases inositol trispohosphate levels in human endothelial cells. Biochem. Biophys. Res. Comm., 170, 281–290. 76. Ohno, M., Cooke, J.P., Dzau, V.J. and Gibbons, G.H. (1995) Fluid shear stress induces endothelial TGF-ß1 transcription and production. Modulation by potassium channel blockade. J. Clin. Inv., 95, 1363–1369. 77. Du, W., Mills, I. and Sumpio, B.E. (1995) Cyclic strain causes heterogeneous induction of transcription factors, AP-1, CRE binding protein and NF-B, in endothelial cells: species and vascular bed diversity. Journal of Biomechanics, 28, 1485–91. 78. Lan, Q., Mercurius, K.O. and Davies, P.F. (1994) Stimulation of transcription factors NF-B and AP1 in endothelial cells subjected to shear stress. Biochem. Biophys. Res. Comm., 201, 950–956. 79. Resnick, N., Collins, T., Atkinson, W., Bonthron, D.T., Dewey, C.F. Jr. and Gimbrone M.A. (1993) Platelet-derived growth factor B chain promoter contains a cis-acting fluid shear-stress-responsive element. Proc. Natl. Acad. Sci., 90. 80. Khachigian, L.M., Resnick, N., Gimbrone, M.A. Jr. and Collins, T. (1995) Nuclear Factor-B interacts functionally with the platelet-derived growth factor B-chain shear-stress response element in vascular endothelial cells exposed to fluid shear stress. J. Clin. Inv., 96, 1169–1175.
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81. Sumpio, B.E., Du, W., Galagher, G. et al. (1998) Regulation of PDGF-B in endothelial cells exposed to cyclic strain. Arterioscler. Throm. Vasc. Biol., 18, 349–355. 82. Hsieh, H., Li, N. and Frangos, J.A. (1991) Shear stress increases endothelial platelet-derived growth factor mRNA levels. Am. J. Physiol., 260, H642–H646. 83. Malek, A.M., Gibbons, G.H., Dzau, V.J. and Izumo, S. (1993) Fluid shear stress differentially modulates expression of genes encoding basic fibroblast growth factor and platelet-derived growth factor B chain in vascular endothelium. J. Clin. Inv., 92, 2013–2021. 84. Mitsumata, M., Fishel, R.S., Nerem, R.M., Alexander, R.W. and Berk B.C. (1993) Fluid shear stress stimulates platelet-derived growth factor expression in endothelial cells. Am. J. Physiol., 265, H3–H8. 85. Wang, D.L., Wung, B.S., Peng, Y.C. and Wang, J.J. (1995) Mechanical strain increases endothelin-1 gene expression via protein kinase C pathway in human endothelial cells. Journal of Cellular Physiology, 163, 400–6. 86. Malek, A.M., Greene, A.L. and Izumo, S. (1993) Regulation of endothelin 1 gene by fluid shear stress is transcriptionally mediated and independent of protein kinase C and cAMP. Proc. Natl. Acad. Sci. USA, 90, 5999–6003. 87. Morita, T., Kurihara, H., Maemura, K., Yoshizumi, M., Nagai, R. and Yazaki, Y. (1994) Role of Ca2+ and protein kinase C in shear stress-induced actin depolymerization and endothelin 1 gene expression. Circ. Res., 75, 630–636. 88. Grabowski, E.F., Jaffe, E.A. and Weksler, B.B. (1985) Prostacyclin production by cultured endothelial cell monolayers exposed to step increases in shear stress. J. Lab. & Clin. M ed., 105, 36–43. 89. Kito, H., Yokoyama, C., Inoue, H., Tanabe, T., Nakajima, N. and Sumpio, B.E. (1997) Cyclooxygenase expression in bovine aortic endothelial cells exposed to cyclic strain. Endothelium, 28, 1–6. 90. Topper, J.N., Cai, J., Falb, D. and Gimbrone, M.A. Jr. (1996) Identification of vascular endothelial genes differentially responsive to fluid mechanical stimuli: cyclooxygenase-2, manganese superoxide dismutase and endothelial cell nitric oxide synthase are selectively up-regulated by steady laminar shear stress. Proc. Natl. Acad. Sci. USA, 93, 10417–10422. 91. Norris, M., Morigi, M., Donadelli. R. et al. (1995) Nitric oxide synthesis by cultured endothelial cells is modulated by flow conditions. Circ. Res., 76, 536–543. 92. Wang, D.L., Wung, B.S., Shyy, Y.J. et al. (1995) Mechanical strain induces monocyte chemotactic protein-1 gene expression in endothelial cells. Effects of mechanical strain on monocyte adhesion to endothelial cells. Circulation Research, 77, 294–302. 93. Shyy, Y.J., Lin, M.C., Han, J., Lu, Y., Petrime, M. and Chien, S. (1995) The cis-acting phorbol ester “12-O-tetradecanoylphorbol 13-acetate”-responsive element is involved in shear stress-induced monocyte chemotactic protein 1 gene expression. Proc. Natl. Acad. Sci., 92, 8069–8073. 94. Ando, J., Tsuboi, H., Korenaga, R. et al. (1995) Down-regulation of vascular adhesion molecule-1 by fluid shear stress in cultured mouse endothelial cells. Ann NY Acad Sci., 748, 148–156. 95. Nagel, T., Resnick, N., Atkinson, W.J., Dewey, C.F. Jr. and Gimbrone, M.A. Jr. (1994) Shear stress upregulates intercellular adhesion molecule-1 expression in cultured human vascular endothelial cells. J. Clin. Inv., 94, 885–891. 96. Takei, T., Mills, I., Arai, K. and Sumpio, B.E. (1998) Molecular basis for tissue expansion: Clinical implications for the surgeon. Plastic and Reconstructive Surgery, 102, 247–259. 97. Takei, T., Rivas-Gotz, C., Delling, C.A. et al. (1997) Effect of strain on human keratinocytes in vitro. J. Cell. Physiol., 173, 64–72. 98. Takei, T., Han, O., Ikeda, M., Male, P., Mills, I. and Sumpio, B.E. (1997) Cyclic strain stimulates isoform-specific PKC activation and translocation in cultured human keratinocytes. J. Cell. Biochem., 67, 327–337. 99. Takei, T., Kito, H., Du, W., Mills, I. and Sumpio, B.E. (1998) Cyclic strain induction of interleukin (IL)-1a and ß gene expression in human keratinocytes: Potential mediator of strain-induced keratinocyte proliferation. J. Cell. Biochem., 69, 95–103.
13 Effects of Hydrostatic Pressure on Endothelial Cells Eric A.Schwartz, Mary E.Gerritsen* and Rena Bizios† Department of Biomedical Engineering, Rensselaer Polytechnic Institute, Troy, NY 12180–3590, *Department of Cardiovascular Research, Genentech, Inc., South San Francisco, CA, 94080 USA. †Corresponding author: E-mail address: [email protected].
Vascular endothelial cells are exposed to a variety of mechanical forces in vivo, specifically, shear stress from blood flow, tensile stress from the compliance of the vessel wall, and hydrostatic pressure from containment of blood within the vasculature. The effects of hydrostatic pressure have received attention only recently, but results of several studies suggest that the responses of endothelial cells to hydrostatic pressure differ from those to shear and tensile stresses. These studies have shown that exposure of both bovine and human endothelial cells to sustained hydrostatic pressure stimulates cell proliferation and alters cell morphology, evidenced by cell elongation (without a predominant cell orientation) concomitant with cytoskeletal reorganization (Acevedo et al., 1993; Sumpio et al., 1987; Salwen 1994; Schwartz, 1999). Additionally, exposure of human endothelial cells to sustained pressure does not alter basal, surface expression of intercellular adhesion molecule-1 (ICAM-1), vascular cell adhesion molecule-1 (VCAM-1), E-Selectin, CD31, and p96 (Schwartz et al., 1998); these results are in contrast to the upregulation of ICAM-1 (Nagel et al., 1994) and the downregulation of VCAM1 and E-Selectin (Sampath et al., 1995) induced in endothelial cells by laminar shear stress. The mechanisms underlying the unique responses of endothelial cells to pressure are the subject of current investigations. KEYWORDS: Endothelium, hydrostatic pressure, mechanotransduction, integrins, angiogenesis.
INTRODUCTION In vivo, the vascular endothelial cell is exposed to a hydrostatic pressure which is the net sum of plasma osmotic and plasma oncotic pressures, tissue osmotic and tissue oncotic pressures, and a component of the “blood pressure” (as measured using sphygmomanometry). The blood pressure measured by a sphygmomanometer represents a stagnation pressure which is a conversion of the dynamic, unidirectional pressure which drives blood flow (and applies fluid shear stress to the endothelial cells) into a static, omnidirectional pressure (which contributes to the hydrostatic pressure to which the endothelial cells are exposed). Therefore, the hydrostatic pressure vascular endothelial cells are exposed to is proportional to, but not equal to, the sphygmomanometric blood pressure. For example, in the rat, abdominal aortic pressure ranges from 132–172 cm H2O but the range of tissue interstitial pressure in the kidney is from 4–29 cm H2O (Skarlatos et al., 1994). This relation also implies that variations in blood pressure (such as transient changes due to body posture and/ or vascular tone, or chronic changes due to hypertension and 275
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atherosclerosis) can alter the net hydrostatic pressure to which endothelial cells are exposed. Additionally, anatomical parameters (e.g., height of the vascular bed and rigidity of tissue support) expose endothelial cells at different sites of the vascular tree to different hydrostatic pressures; for example, in the upright human body, the veins of the neck are maintained at atmospheric pressure (Guyton, 1986), the veins of the skull are maintained at subatmospheric pressure (Guyton, 1986), and in the large vessels of the feet, pressure may exceed 100 cm H2O above atmospheric pressure (Guyton, 1986). Physiological and pathological conditions (e.g., postural change, exercise, injury, inflammation, hemorrhage, and fever) can also alter local blood and tissue pressures, and these pressures also contribute to small changes in the net hydrostatic pressure the endothelial cells at a particular site may be exposed to. Elevated hydrostatic pressure (in addition to altered shear and tensile stresses) and vascular endothelial cell dysfunction have been epidemiologically linked to a number of vascular disease processes including hypertension and atherosclerosis (Thubrikar and Robicsek, 1995), glaucoma-induced retinal microangiopathy (Aiello et al., 1994), rheumatoid arthritis (Ahlqvist et al., 1994), and malignant tumor growth (Nathanson and Nelson, 1994). Despite a correlation between physical forces (namely, pressure, shear, and tensile stresses) and vascular dysfunction, a causal link has not been established. Other chapters in this book discuss the potential roles of shear and tensile stresses in modulating endothelial cell function. The present chapter will review what is currently known about the effects of hydrostatic pressure on endothelial cell morphology, cytoskeletal organization, surface protein expression, and proliferation. Additionally, the similarities and differences in endothelial responses to hydrostatic pressure compared to other physical forces will be discussed.
EFFECTS OF HYDROSTATIC PRESSURE ON ENDOTHELIAL CELL MORPHOLOGY The morphology of a cell is a dynamic participant in, and indicator of, cellular function (Ingber, 1993); alterations in cell morphology are associated with and are necessary for cell differentiation, proliferation, migration, and cell-cell interactions (as reviewed in Sims et al., 1992). In vivo, vascular endothelial cells are elongated and aligned with the direction of blood flow (Langille and Adamson, 1981); however, at branching sites, where flow is disturbed, these cells are rounded and present a “cobblestone” morphology (Thubrikar and Robicsek, 1995). At the level of the capillary, the endothelial cells are flattened, and single cells wrap around to form the lumen of the vessel. In the process of angiogenesis, endothelial cells undergo several morphological alterations in order to form a new, small, blood vessel; specifically, cells at the site of a new capillary elongate, then migrate, invading the extracellular matrix which originally surrounded the vessel wall (as reviewed in Stromblad and Cheresh, 1996). Furthermore, initiation of either cell proliferation or apoptosis is regulated by the degree of cell extension (controlled by the size, shape, and spacing among protein-coated domains patterned on substrate
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surfaces where the cells are cultured), and is independent of the total area of focal adhesion plaques per cell; however, a synergistic effect exists between total cell area and select adhesive proteins on the substrate surfaces, as cells on surfaces micropatterned with fibronectin and collagen type I were more likely to undergo apoptosis than cells on surfaces micropatterned with vitronectin (Chen et al., 1997). Thus, in addition to being an important phenomenon in their own right, morphological changes that occur following exposure of endothelial cells to pressure may contribute to or even control cell functional changes, such as increased cell proliferation and changes in cellular protein expression. In vitro, exposure of bovine and human endothelial cells to sustained hydrostatic pressures of 1.5–109 cm H2O for 1–9 days results in cell elongation without a predominant cell orientation (Figure 13.1, frames A and B) (Acevedo et al., 1993; Salwen, 1994; Sumpio et al., 1994; Schwartz, 1999). These changes in cell shape are accompanied by concomitant reorganization of the cell cytoskeleton. Specifically, pressure-treated endothelial cells lose the characteristic peripheral actin band and reorganize the internal regions of the cytoskeleton from a web-like matrix (Figure 13.1, frame C) to an aligned array of thick, parallel stress fibers (Figure 13.1, frame
Figure 13.1. Exposure of bovine endothelial cells to sustained pressure alters cell morphology and cytoskeletal organization. Phase-contrast micrographs of bovine aortic endothelial cells that exhibit either the characteristic cobblestone morphology of endothelial cells maintained under control (0.3 cm H2O) pressure conditions (A) or the elongated morphology (without predominant cell orientation) of endothelial cells exposed to 15 cm H2O sustained hydrostatic pressure (B) for 7 days. Bar=250 µm. Fluorescent micrographs of bovine pulmonary artery endothelial cells that possess the characteristic peripheral actin band (but no stress fibers) of cells maintained under control pressure conditions (C) or that possess thick stress fibers (but no peripheral actin band) of cells exposed to 15 cm H2O sustained hydrostatic pressure (D) for 7 days and were subsequently stained with rhodamine phalloidin. Bar=22 µm. (Reprinted from Acevedo, A., et al., J. Cell. Phys., 157:603–614, 1993 Copyright (1993, John Wiley & Sons, Inc.), with permission by the publisher).
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D) (Acevedo et al., 1993; Sumpio et al., 1994; Salwen, 1994; Schwartz, 1999). In addition, the cytoskeleton reorganizes from a single plane in cells maintained under control conditions to a multilayered structure following exposure of the cells to pressure; the number of these layers (up to 5 distinct planes under 10cm H2O), as well as the thickness of individual fibers, depends on the magnitude (1.5–10 cm H2O) and duration (1–7 days) of the applied pressure (Salwen, 1994). The changes in cell morphology and cytoskeletal organization that occur while endothelial cells are exposed to sustained hydrostatic pressure are similar to changes observed for endothelial cells during angiogenesis and during other endothelial cell proliferation events; it is interesting to note that the cell elongation and cytoskeletal alignment observed in pressure-treated endothelial cells is similar to the elongation of capillary endothelial cells in the early stages of angiogenesis. Furthermore, the cell elongation (without predominant cell orientation) observed during exposure of endothelial cells to pressure is similar both to the morphological changes observed during exposure of these cells to turbulent fluid shear stress, a mechanical force also associated with increased cell proliferation (Levesque et al., 1990), and to morphological changes which result from controlled cell spreading (observed when cells are cultured on specially micropatterned surfaces) that inhibit apoptosis and initiate endothelial cell proliferation in vitro (Chen et al., 1997). Thus, the morphological changes that occur following exposure of endothelial cells to pressure suggest that these cells exhibit an active, proliferative phenotype.
EFFECTS OF HYDROSTATIC PRESSURE ON ENDOTHELIAL CELL SURFACE ANTIGEN EXPRESSION In addition to morphological and cytoskeletal changes, exposure of endothelial cells to sustained pressure may alter the basal or stimulated expression of cell surface antigens such as a5ß1 and E-selectin (CD62E), a cytokine-induced adhesion molecule which is thought to play a role in leukocyte trafficking (Bevilacqua et al., 1987) and in angiogenesis (as reviewed in Bevilacqua and Nelson, 1993; Nguyen et al., 1993; Koch et al., 1995). Integrin a5ß1 plays a role in cell-cell and cell-matrix interactions (as reviewed in Albeda and Buck, 1990), as well as in the transduction of mechanical stimuli in endothelial cells (Wang et al., 1993; Chen et al., 1994; Papadaki and Eskin, 1997). Exposure of bovine aortic endothelial cells to 52 cm H2O pressure for time periods of 12–48 hours results in increased clustering of vinculin, talin, and integrin a5ß1 in focal adhesion plaques, in thickening of laminin and collagen type IV fibrils in the extracellular matrix, and in increased deposition of fibronectin and laminin (but not of vitronectin) into the extracellular matrix (Thoumine et al., 1995). Exposure of human umbilical vein endothelial cells (HUVEC) to 4 cm H2O sustained pressure for 1 day does not affect the basal cell surface expression of intercellular adhesion molecule-1 (ICAM-1 or CD54), vascular cell adhesion molecule-1 (VCAM-1 or CD106), E-Selectin (CD62E), platelet-endothelial cell adhesion molecule (CD31), and p96 (Schwartz et al., 1998). However, exposure of HUVEC to pressure does appear to either prolong or enhance the expression of E-Selectin that occurs
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following exposure of these cells to 100 ng/mL Escherichia coli lipopolysaccharide (Schwartz et al., 1999; Table 13.1).
EFFECTS OF HYDROSTATIC PRESSURE ON ENDOTHELIAL CELL PROLIFERATION Most endothelial cells in vivo exhibit a very low turnover rate ( 1 cell division/year; Folkman and Shing, 1992). Endothelial cell proliferation, however, is observed under a number of normal physiological conditions including wound healing (Silver and Doillon, 1989), embryogenesis (Brooks et al., 1994), and ovulation (Iruela-Arispe and Dvorak, 1997). In addition, a number of pathological states, including glaucomainduced retinal microangiopathy (Aiello et al., 1994), rheumatoid arthritis (Ahlqvist et al., 1994), and solid tumor growth (Nathanson and Nelson, 1994) are associated with increased endothelial cell proliferation; interestingly, these pathologies (as well as wound healing) are also associated with localized elevations in hydrostatic pressure. The in vivo association between increased hydrostatic pressures and increased endothelial cell proliferation may be only circumstantial, although a causal relationship cannot be ruled out. A possible link is suggested by in vitro studies from several laboratories which have demonstrated that exposure of bovine, porcine, and human endothelial cells to sustained hydrostatic pressures in the range 1.5–260 cm H2O for time periods of 1–9 days stimulates endothelial cell proliferation and (at least in bovine endothelium) loss of contact-inhibited growth, leading to cell Table 13.1 Effects of sustained hydrostatic pressure and lipopolysaccharide (LPS) on the expression of constitutive surface antigens by endothelial cells. HUVEC were either maintained under control (0.2 cm H2O) pressure conditions (in the presence and absence of 100ng/mL LPS) or exposed to 4cm H2O sustained hydrostatic pressure (in the presence and absence of 100ng/mL LPS) for 24 hours. These cells were labeled with primary mouse monoclonal antibodies for CD54, CD106, CD62E, CD31, and p96; controls were cells without primary antibody labeling. All cells were subsequently stained with fluorescent rabbit anti-mouse antisera. Fluorescent intensities were read for 10,000 cells using a FACS® analyzer. Compared to endothelial cells exposed to LPS alone, the increased levels of E-Selectin in cells exposed to both LPS and pressure may reflect either a potentiation of the magnitude of the LPS response or a prolongation of E-Selectin expression. Note that E-Selectin expression in HUVEC is maximal at 4–6 hours after exposure to LPS and returns towards baseline levels at 24 hours (Bevilacqua et al., 1987).* Significantly different from LPS-treated group; p<0.05. (Reprinted from Schwartz, E.A., et al., in Pulmonary Edema, K.Weir and J.T.Reeves (eds.), 1998, Futura Publishing Company, Inc.,Copyright (1998, American Heart Association), with permisson by the American Heart Association)
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multilayering (Acevedo et al., 1993; Salwen, 1994; Sumpio et al., 1994; Tokunaga et al., 1989; Schwartz, 1999). An increase in proliferation in response to variations in sustained hydrostatic pressure is not a phenomenon restricted to endothelial cells. For example, vascular smooth muscle cells (Tokunaga and Watanabe, 1987), chondrocytes (Wright et al., 1992; Smith et al., 1996), and fibroblasts from a variety of tissues (Smith et al., 1996; Yousefian et al., 1995) demonstrate increased cell proliferation in response to changes in hydrostatic pressure. Thus, the proliferative response of cells to sustained hydrostatic pressure may be a fundamental process common to many cell types. A possible mediator of the proliferative response of endothelial cells to sustained hydrostatic pressure was suggested by the research results of Acevedo et al. (1993). This study demonstrated that exposure of naive bovine pulmonary artery endothelial cells to conditioned media from pressure-treated endothelial cells results in increased proliferation as well as in morphological and cytoskeletal changes identical to those observed when endothelial cells under fresh (non-conditioned) medium are exposed to pressure. The biological activity of the conditioned medium could be abolished by heating, freeze-thawing, binding to heparin-Sepharose, treatment with Suramin, and treatment with monoclonal antibodies to basic fibroblast growth factor (bFGF) (Acevedo et al., 1993). In the absence of sustained hydrostatic pressure, treatment of bovine endothelial cells with bFGF results in proliferative and morphological changes identical to those observed in pressure-treated and in conditioned-media treated cells. Additional evidence for a role of bFGF is provided by the observation that exposure of bovine pulmonary artery and of human umbilical vein endothelial cells to 1.5–15 cm H2O sustained hydrostatic pressure, for time periods up to 7 days, results in loss of immunoreactive bFGF from cellular and/or extracellular matrix stores without cell damage or cell death (Acevedo et al., 1993; Schwartz, 1999) (Figure 13.2). Both accumulation of bFGF in the media and loss of bFGF from the cells occurs without an apparent change in steady-state bFGF mRNA within 0.5–24 hours (Schwartz, 1999). Thus, the proliferative and morphologic responses of endothelial cells to sustained
Figure 13.2. Exposure of human endothelial cells to sustained pressure results in loss of immunoreactive intracellular basic fibroblast growth factor. Fluorescent micrographs of human umbilical vein endothelial cells that had been either maintained under control (0.3 cm H2O) pressure (A) or exposed to 4 cm H2O sustained hydrostatic pressure for 1 day (B) before immunostaining with monoclonal antibodies to bFGF and fluorescein-conjugated anti-mouse antisera. Exposure of endothelial cells to pressure results in loss of immunoreactive bFGF from cytoplasmic and nuclear stores.
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hydrostatic pressure may be mediated (at least in part) by autocrine stimulation with bFGF (Acevedo et al., 1993).
COMPARISON OF ENDOTHELIAL CELL RESPONSES TO PRESSURE, SHEAR, AND TENSILE STRESSES In addition to sustained hydrostatic pressure, endothelial cells respond to other mechanical forces, specifically, shear and tensile stresses (Akai et al., 1994; Buga et al., 1991; Carosi et al., 1994; Davies et al., 1986, Davies, 1993; Davies and Tripathi, 1993; Dewey et al., 1981; Diamond et al., 1989; Girard and Nerem, 1995; Langille and Adamson, 1981; Levesque et al., 1990; Nagel et al., 1994; Oluwole et al., 1997; Patrick and McIntire, 1995a; Patrick and McIntire, 1995b; Sampath et al., 1995; Sumpio et al., 1987; Thoumine et al., 1995). While some aspects of the responses of endothelial cells to these mechanical stimuli are similar, the details are unique to each particular mechanical force (Table 13.2). For example, exposure of endothelial cells to any one of the three stresses results in cell elongation and in formation of stress fibers aligned with the long axis of each cell. The unique difference is manifested in the relative orientation of these cells; specifically, exposure of endothelial cells to fluid shear and substrate tension for time periods of 3 hours to 2 days results in alignment of the cell long axes in the direction of fluid flow
Table 13.2 Comparison of the cellular-level effects of fluid shear, substrate tension, and sustained hydrostatic pressure on endothelial cells
References: a(Dewey et al., 1981); b(Ohshima and Ookawa, 1992); c(Sumpio et al., 1987); d(Acevedo et al., 1993); e(Sumpio et al., 1994); f(Salwen, 1994); g(Schwartz, 1999); h(Davies et al., 1986); i(Levesque et al., 1990).
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(Dewey et al., 1981), and perpendicular to the direction of stretch (Sumpio et al., 1987), respectively; in contrast, exposure of endothelial cells to sustained hydrostatic pressure for 4–9 days results in cell elongation without predominant cell orientation (Acevedo et al., 1993, Sumpio et al., 1994; Salwen, 1994). Exposure of endothelial cells to sustained pressure does not affect the surface expression of several endothelial cell-leukocyte adhesion molecules which are sensitive to fluid shear and substrate tensile stresses (e.g., ICAM-1 and VCAM-1). Furthermore, exposure of endothelial cells to sustained hydrostatic pressure does not alter steadystate mRNA levels of many of the shear-inducible (as reviewed in Patrick and McIntire, 1995) enzymes, such as nitric oxide synthase and cyclooxygenase-2 (Schwartz, 1999). A comparison of the effects of these three mechanical forces on various molecular events in endothelial cells is presented in Table 13.3. Exposure of endothelial cells to sustained pressure results in the release (Acevedo et al., 1993) (but not in altered steady-state mRNA levels; Schwartz, 1999) of basic fibroblast growth factor; in contrast, this growth factor is upregulated at the mRNA level by exposure of endothelial cells to fluid shear and substrate tensile stresses (Table 13.3). Exposure of endothelial cells to fluid shear and tensile stresses also results in secretion of a variety of vasoactive substances (such as endothelin-1 and prostacyclin) which, to date, have not been investigated for cells under hydrostatic pressure (Table 13.3). The proliferative responses of endothelial cells to sustained hydrostatic pressure, fluid shear, and substrate tensile stresses are similar, yet distinct from each other (Table 13.2). Specifically, exposure of endothelial cells to turbulent fluid shear stress results in increased cell proliferation (determined by 3H Thymidine incorporation), detectable as early as 3 hours (Davies et al., 1986); however, exposure of endothelial cells to either pulsatile, laminar shear stress or steady laminar shear stress has been reported in the literature to either decrease cell proliferation (determined by total cell number and 3H Thymidine incorporation) (Levesque et al., 1990; Nerem, 1993), or not affect cell proliferation (determined by total cell number and 3H Thymidine incorporation) over a time course of up to 8 days (Dewey et al., 1981; Davies et al., 1986; Thoumine et al., 1995). In addition, exposure of bovine endothelial cells to cyclic tension results in increased cell proliferation (determined by 3H Thymidine incorporation) within 1 day (Sumpio et al., 1987). In contrast, exposure of bovine (Acevedo et al., 1993; Sumpio et al., 1994) and human (Schwartz, 1999) endothelial cells to sustained hydrostatic pressure results in increased cell proliferation (determined by total cell number) over a time course of 1–9 days. More detailed comparison of cell cycle kinetics amongst the different force environments is not feasible at present due to the different methodologies used in the various studies reported in the literature (Dewey et al., 1981; Davies et al., 1986; Sumpio et al., 1987; Levesque et al., 1990; Nerem, 1993; Acevedo et al., 1993; Sumpio et al., 1994; Thoumine et al., 1995; Schwartz, 1999). The uniqueness of cell responses to each mechanical force may also result from differential sensitivities of endothelial cells to various types and durations of mechanical stimuli. For example, vascular endothelial cells in vitro sense and respond to levels of hydrostatic pressure as low as 1.5cm H2O (Acevedo et al., 1993). Compared to a physiological range (132–172 cm H2O) of mean “blood pressure” (Skarlatos et al., 1994), a pressure difference of 1.5 cm H2O seems quite low. However, this pressure
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Table 13.3 Comparison of the molecular-level effects of fluid shear, substrate tension, and sustained hydrostatic pressure on protein expression by endothelial cells
References: a(Patrick and McIntire, 1995a); b(Acevedo et al., 1993); c(Schwartz et al., 1998), (Schwartz, 1999), and (Schwartz et al., 1997); d(Papadaki and Eskin, 1997); e(Oluwole et al., 1997); f(Yano et al., 1996); g(Thoumine et al., 1995); h(Buga et al., 1991); i(Akai et al., 1994). *ND=not determined.
still exposes cells to a stress of 1472 dynes/cm2, which is actually greater than the 8– 20 dynes/cm2 laminar shear (Dewey et al., 1981) and 5–90 dynes/cm2 turbulent shear (Levesque et al., 1990) stresses which others have used in their investigations of the effects of mechanical forces on the endothelial cell phenotype. To date, most in vitro studies have examined cell responses to a single, isolated mechanical stimulus. Vascular endothelial cells in vivo, however, exist in a dynamic environment which exposes them to several mechanical forces simultaneously. The effects of a combined pressure-shear regime were investigated in vitro by Thoumine et al., 1995. In this study, and compared to control cells maintained under static conditions at atmospheric pressure, exposure of human endothelial cells in vitro to
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54cm H2O sustained hydrostatic pressure above atmospheric, concomitant with either steady (0.2 dynes/cm2) or pulsatile (0.2±15 dynes/cm2) fluid shear stress for time periods of 12–48 hours did not alter cell proliferation (Thoumine et al., 1995). In mixed-force stress regimes, the net response of endothelial cells to the combined mechanical forces may not merely be the sum of cell responses to each individual force, but some (as yet undetermined) novel cell response to the combination of mechanical forces. Nonetheless, these observations (Thoumine et al., 1995) infer that the effects of mixedforce stress regimes on endothelial cells need to be thoroughly investigated in the future.
MECHANOTRANSDUCTION OF SUSTAINED PRESSURE Endothelial cells respond to external stimuli (such as mechanical forces) through the sequential, parallel, or integrated activation of various signaling pathways which result in gene transcription and other cellular responses. Some details of the mechanotransduction cascades for fluid shear and tensile stresses are known; for example, cis-acting response elements within the gene promoter segments for plateletderived growth factor (Resnick et al., 1993) and monocyte chemotactic protein-1 (Shyy et al., 1995) are activated following exposure of endothelial cells to fluid shear, and roles for the transcription factors NF-B (Resnick et al., 1993), c-jun/ c-fos (AP1) (Shyy et al., 1995), and Egr-1 (Khachigian et al., 1996) have been implied. In addition, mechanotransduction mechanisms upstream of gene activation have been implicated for fluid shear and tensile stresses, including nitric oxide (NO) synthesis (Buga et al., 1991) and protein kinase C activation (Shyy et al., 1995). The sensory mechanisms by which endothelial cells respond to sustained hydrostatic pressure, however, have only received attention recently. There are a number of candidate mechanisms by which endothelial cells sense pressure including: pressure-gated ion channels (Kohler, 1996); changes in membrane tension and/or area (Schmid-Schoenbein et al., 1995), which may activate tension-gated ion channels (Kohler, 1996); and cytoskeletal mechanotransduction mechanisms (Ingber, 1993), including cell shape changes (Wang et al., 1993) and reduced repolymerization of F-actin during cytoskeletal turnover (Garcia et al., 1992). It is also possible that direct compressive effects on cellular protein and nuclear constituents could occur (Garcia et al., 1992; Nicolini et al., 1986). Some of these proposed pressure mechanosensory events are illustrated schematically in Figure 13.3. Some details of the mechanism(s) by which endothelial cells respond to sustained hydrostatic pressure have been experimentally determined; specifically, work by Acevedo et al., 1993, demonstrated that autocrine stimulation by bFGF mediated (at least in part) pressure-induced morphological, cytoskeletal, and proliferative changes in bovine endothelial cells (Acevedo et al., 1993). Protein kinase C (PKC) activation may also play a role in the response of endothelial cells to pressure; specifically, downregulation of PKC by pre-treatment of human umbilical vein endothelial cells with 100 nM tetradecanoyl phorbol acetate for 24 hours prevented pressure-induced increases in av integrin expression (Schwartz, 1999) (Figure 13.4). Upregulated expression and redistribution of integrins following exposure of endothelial cells to
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Figure 13.3. Proposed pressure-sensing mechanisms of endothelial cells. Endothelial cells may sense sustained hydrostatic pressure through a variety of mechanisms. Numbers on the figure represent the following proposed mechanisms: (1) pressure-gated ion channels; (2) compression of the cell membrane leading to closure of membrane tension-gated ion channels; (3) deformation of the cell body and/or nucleus (dotted lines); (4) invagination of the cell membrane, leading to reduction of cell surface area; (5) stretching of membrane regions over the rigid cytoskeletal framework; (6) compression of cytoskeletal elements leading to a decrease in actin repolymerization; (7) stress concentrations at junctions of cytoskeletal elements, which could enhance cytoskeletal mechanotransduction mechanisms; and (8) activation of integrin adhesion molecules by mechanical tension in response to changes in cell shape and/ or volume as the cell is exposed to pressure.
Figure 13.4. Expression of integrin v by endothelial cells exposed to pressure and phorbol ester. Proteins were extracted from HUVEC and analyzed by Western blot using polyclonal antibodies to integrin v. Compared to human umbilical vein endothelial cells maintained under control (0.2 cm H2O) pressure conditions (lane 1), cells that had been exposed to 4 cm H2O sustained hydrostatic pressure for 4 hours (lane 4) and 1 day (lane 5) exhibited increased immunoreactive integrin v levels. Similarly, cells that had been exposed to 100 nM tetradecanoyl phorbol acetate (PMA) under control pressure conditions for 4 hours (lane 2) and 1 day (lane 3) days exhibited increased integrin v levels. In contrast, cells that had been pre-exposed to 100 nM PMA for 24 hours (lanes 6 and 7) exhibited only a small increase in integrin v levels; when these cells were subsequently either maintained under control pressure conditions (lane 6) or exposed to 4 cm H2O pressure for 1 day (lane 7) in the absence of PMA, the pre-treated cells did not exhibit any pressure-induced increase in integrin ay levels.
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pressure also imply a role for these cell-cell/cell-matrix adhesion proteins in mechanotransduction. A hypothetical model integrating the potential roles of bFGF, PKC, and integrins in the mechanotransduction of sustained hydrostatic pressure is depicted in Figure 13.5. In this model, extracellular release of bFGF activates phosphatidylinositol 3' kinase and phospholipase-C (Chong et al., 1994; Sepp et al., 1994; Guo et al., 1995), which, in turn (through an increase in intracellular levels of diacylglycerol) activate protein kinase C (as reviewed in Woodgett, 1994). Activation of protein kinase C (by various agonists) has been shown to upregulate integrin synthesis (Swerlick et al., 1992; Tang et al., 1993b; Tang et al., 1993a; Sepp et al., 1994; Tang et al., 1995), and integrins have been implicated in the initiation and/or potentiation of cell proliferation (Brooks et al., 1994; Friedlander et al., 1995; Zheng et al., 1997).
Figure 13.5. Proposed hydrostatic pressure transduction mechanism(s) of endothelial cells. Some steps (namely, basic fibroblast growth factor autocrine stimulation and protein kinase C activation) in the mechanotransduction of sustained pressure by endothelial cells have been experimentally demonstrated; a hypothesis for the role of these steps in the pressure mechanotransduction pathway is illustrated in this schematic drawing. Exposure of endothelial cells to sustained hydrostatic pressure stimulates release of basic fibroblast growth factor, resulting in autocrine stimulation of the fibroblast growth factor receptor (FGFR). Active FGFR stimulates production of diacylglycerol (Guo et al., 1995), which in turn activates protein kinase C (PKC) (Tang et al., 1993b). PKC stimulates synthesis of integrins (Swerlick et al., 1992), which play a role in the control of endothelial cell proliferation (Brooks et al., 1994; Friedlander et al., 1995; Zheng et al., 1997). Morphological and cytoskeletal changes may result from either increased quantities of integrins in focal adhesion plaques (Swerlick et al., 1992), PKC activation (Tang et al., 1993b), or molecular mechanisms involved in cell proliferation (Nicolini et al., 1986).
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SUMMARY AND CONCLUSIONS In summary, exposure of endothelial cells (from various vascular beds and species) to sustained pressure results in increased cell proliferation, reorganization of the F-actin filaments of the cytoskeleton, morphological changes (elongation without a predominant cell orientation), secretion of soluble, bioactive molecules, and altered expression of integrins but not of endothelial cell-leukocyte adhesion molecules. These changes are similar to, but distinct from, those which occur during exposure of endothelial cells to either fluid shear or tensile stresses. In conclusion, sustained hydrostatic pressure affects endothelial cell morphology and function in a unique manner; these responses may serve as the causative link between elevated hydrostatic pressure and endothelial cell-related pathologies in vivo. ACKNOWLEDGEMENTS The authors wish to thank Dr. Marvin S.Medow and Ms. Lillian Kletter (New York Medical College, Valhalla, NY) for assistance with the PKC experiments and Dr. Keith Kelley (Bayer Corporation, West Haven, CT) for assistance with microscopy. The authors also wish to thank The Whitaker Foundation for a graduate fellowship to Eric A.Schwartz. REFERENCES Acevedo, A., Bowser, S., Gerritsen, M. and Bizios, R. (1993) Morphological and proliferative responses of endothelial cells to hydrostatic pressure: role of fibroblast growth factor. J. Cell Physiol., 157, 603–614. Ahlqvist, J., Harilainen, A., Aalto, K., Sarna, S., Lalla, M. and Osterlund, K. (1994) High hydrostatic pressures in traumatic joints require elevated synovial capillary pressure probably associated with arteriolar vasodilatation. Clin. Physiol., 14, 671–679. Aiello, L., Avery, R. and Arrigg, P. (1994) Vascular endothelial growth factor in ocular fluid of patients with diabetic retinopathy and other retinal diseases. New England Journal of Medicine, 331, 1480–1487. Akai, Y., Homma, T., Burns, K., Yasuda, T., Badr, K. and Harris, R. (1994) Mechanical stretch/ relaxation of cultured rat mesangial cells induces protooncogenes and cyclooxygenase. Am. J. Physiol., 267, C482-C490. Albeda, S. and Buck, C. (1990) Integrins and other cell adhesion molecules. FASEB J., 4, 2868–2880. Bevilacqua, M. and Nelson, R. (1993) Selectins. J. Clin. Invest., 91, 379–387. Bevilacqua, M., Pober, J., Mendrick, D., Cotran, R. and Gimbrone Jr., M. (1987) Identification of an inducible endothelial-leukocyte adhesion molecule. Proc. Natl. Acad. Sci. USA, 84, 9238–9242. Brooks, P., Clark, R. and Cheresh, D. (1994) Requirement of vascular integrin vß3 for angiogenesis. Science, 264, 569–571. Buga, G., Gold, M., Fukuno, J. and Ignarro, L. (1991) Shear stress-induced release of nitric oxide from endothelial cells grown on beads. Hypertension, 17, 187–193. Carosi, J., McIntire, L. and Eskin, S. (1994) Modulation of secretion of vasoactive materials from human and bovine endothelial cells by cyclic strain. Biotech. and Bioeng., 43, 615–621. Chen, C., Mrksich, M., Huang, S., Whitesides, G. and Ingber, D. (1997) Geometric control of cell life and death. Science, 276, 1425–1428. Chen, Q., Kinch, M., Lin, T., Burridge, K. and Juliano, R. (1994) Integrin-mediated cell adhesion activates mitogen-activated protein kinases. The Journal of Biological Chemistry, 269, 26602–26605.
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Chong, L., Traynor-Kaplan, A., Bokoch, G. and Schwartz, M. (1994) The small GTP-binding protein Rho regulates a phosphatidylinositol 4-phosphate 5-kinase in mammalian cells. Cell, 79, 507–513. Davies, P. (1993) Endothelium as a signal transduction interface for flow forces: cell surface dynamics. Thrombosis and Haemostasis, 70, 124–128. Davies, P., Remuzzi, A., Gordon, E., Dewey Jr., D. and Gimbrone Jr., M. (1986) Turbulent fluid shear stress induces vascular endothelial cell turnover in vitro. Proc. Natl. Acad. Sci., 83, 2114–2117. Davies, P. and Tripathi, S. (1993) Mechanical stress mechanisms and the cell: an endothelial paradigm. Circ. Res., 72, 239–245. Dewey, C., Jr., Bussolari, S., Gimbrone, M., Jr. and Davies, P. (1981) The dynamic response of vascular endothelial cells to fluid shear stress. J. Biomech. Eng., 103, 177–185. Diamond, S., Eskin, S. and McIntire, L. (1989) Fluid flow stimulates tissue plasminogen activator secretion by cultured human endothelial cells. Science, 243, 1483–1485. Friedlander, M., Brooks, P., Shaffer, R., Kincaid, C., Varner, J. and Cheresh, D. (1995) Definition of two angiogenic pathways by distinct v integrins. Science, 270, 1500–1502. Garcia, C., Amaral, J., Jr., Abrahamsohn, P. and Verjovski-Almeida, S. (1992) Dissociation of F-actin induced by hydrostatic pressure . Eur. J. Biochem., 209, 1005–1011. Girard, P. and Nerem, R. (1995) Shear stress modulates endothelial cell morphology and F-actin organization through the regulation of focal adhesion-associated proteins. J. Cell. Physiol., 163, 179–193. Guo, D., Jia, Q., Song, H.-Y., Warren, R. and Donner, D. (1995) Vascular endothelial cell growth factor promotes tyrosine phosphorylation of mediators of signal transduction that contain SH2 domains: association with endothelial cell proliferation. The Journal of Biological Chemistry, 270, 6729–6733. Guyton, A. (1986) Textbook of Medical Physiology, Saunders, Philadelphia. Ingber, D. (1993) Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton. J. Cell Sci., 104, 613–627. Iruela-Arispe, M. and Dvorak, H. (1997) Angiogenesis: a dynamic balance of stimulators and inhibitors. Thrombosis and Haemostasis, 78, 672–677. Khachigian, L., Lindner, V., Williams, A. and Collins, T. (1996) Egr-1-induced endothelial gene expression: a common theme in vascular injury. Science, 271, 1427–1431. Koch, A., Halloran, M., Haskell, C., Shah, M. and Polverini, P. (1995) Angiogenesis mediated by soluble forms of E-selectin and vascular cell adhesion molecule-1. Nature, 376, 517–519. Koehler, R., Hoyer, J. and Distler, A. (1996) Pressure activated cation channel (PAC) in intact endocardial endothelium (EE). FASEB J., 10, A626 (abstract). Langille, B. and Adamson, S. (1981) Relationship between blood flow direction and endothelial cell orientation at arterial branch sites in rabbits and mice. Circ. Res., 48, 481–488. Levesque, M., Nerem, R. and Sprague, E. (1990) Vascular endothelial cell proliferation in culture and the influence of flow. Biomaterials, 11, 702–707. Nagel, T., Resnick, N., Atkinson, W., Dewey, C. Jr. and Gimbrone, M.J. (1994) Shear stress selectively upregulates intercellular adhesion molecule-1 expression in cultured human vascular endothelial cells. J. Clin. Invest., 94, 885–891. Nathanson, S. and Nelson, L. (1994) Interstitial fluid pressure in breast cancer, benign breast conditions, and breast parenchyma . Ann. Surg. Oncol., 1, 333–338. Nerem, R. (1993) Hemodynamics and the vascular endothelium. Transactions of the ASME, 115, 510–514. Nguyen, M., Strubel, N. and Bishoff, J. (1993) A role for sialyl Lewis-X/A glycoconjugates in capillary morphogenesis. Nature, 365, 267–269. Nicolini, C., Belmont, A. and Martelli, A. (1986) Critical nuclear DNA size and distribution associated with S phase initiation. Cell. Biophys., 8, 103–117. Ohshima, N. and Ookawa, K. (1992) Changes in microstructure of cultured porcine aortic endothelial cells in the early stage after applying fluid-imposed shear stress. J. Biomech., 25, 1321–1328. Oluwole, B., Du, W., Mills, I. and Sumpio, B. (1997) Gene regulation by mechanical forces. Endothelium, 5, 85–93. Papadaki, M. and Eskin, S. (1997) Effects of fluid shear stress on gene regulation of vascular cells. Biotechnol. Prog., 13, 209–221. Patrick, Jr., C. and McIntire, L. (1995a) Shear stress and cyclic strain modulation of gene expression in vascular endothelial cells. Blood Purif., 13, 112–124. Patrick, Jr., C. and McIntire, L. (1995b) Fluid shear stress effects on endothelial cell cytosolic pH. Tissue Engineering, 1, 53–70. Resnick, N., Collins, T., Atkinson, W., Bonthron, D., Dewey Jr., C. and Gimbrone Jr., M. (1993) Plateletderived growth factor b chain promoter contains a cis-acting fluid shear-stress-responsive-element. Proc. Natl. Acad. Sci. USA, 90, 4591–4595. Salwen, S. (1994) MS Thesis. Department of Biomedical Engineering, Rensselaer Polytechnic Institute, Troy, NY, USA.
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Sampath, R., Kukielka, G., Smith, C., Eskin, S. and McIntire, L. (1995) Shear stress-mediated changes in the expression of leukocyte adhesion receptors on human umbilical vein endothelial cells in vitro. Annals of Biomedical Engineering, 23, 247–256. Schmid-Schoenbein, G., Kosawada, T., Skalak, R. and Chien, S. (1995) Membrane model of endothelial cells and leukocytes. A proposal for the origin of a cortical stress . J. Biomech. Eng., 117, 171–178. Schwartz, E. (1999) Mechanotransduction of sustained hydrostatic pressure by human umbilical vein endothelial cells. PhD Thesis, Department of Biomedical Engineering, Rensselaer Polytechnic Institute, Troy, NY, USA (In preparation). Schwartz, E., Bizios, R. and Gerritsen, M. (1997) Exposure of human endothelial cells to sustained pressure affected integrin-mediated cell functions. Annals of Biomedical Engineering, 25, 269. Schwartz, E., Bizios, R. and Gerritsen, M. (1998) Effects of sustained hydrostatic pressure on the expression of endothelial cell-leukocyte adhesion molecules. In Pulmonary Edema (Eds, Weir, E. and Reeves, J.) Futura Publishing Company, Armonk, NY, pp. 195–203. Sepp, N., Li, L., Lee, K., Brown, E., Caughman, S., Lawley, T. and Swerlick, R. (1994) Basic fibroblast growth factor increases expression of the alpha v beta 3 integrin complex on human microvascular endothelial cells. J. Invest. Dermatol., 103, 295–299. Shyy, J.-J., Lin, M.-C., Han, J., Lu, Y., Petrime, M. and Chien, S. (1995) The cis-acting phorbol ester “12– O-tetradecanoylphorbol 13-acetate”—responsive element is involved in shear stress-induced monocyte chemotactic protein 1 gene expression. Proc. Natl. Acad. Sci. USA, 92, 8069–8073. Silver, F. and Doillon, C. (1989) Biocompatibility, VCH Publishers, Inc., New York, NY, USA. Sims, J., Karp, S. and Ingber, D. (1992) Altering the cellular mechanical force balance results in integrated changes in cell, cytoskeletal, and nuclear shape. Journal of Cell Science, 103, 1215–1222. Skarlatos, S., Brand, P.H., Metting, P.J. and Britton, S.L. (1994) Spontaneous changes in arterial blood pressure and renal interstitial hydrostatic pressure in conscious rats. Journal of Physiology, 481, 743–752. Smith, R., Rusk, S., Ellison, B., Wessells, P., Tsuchiya, K., Carter, D., Caler, W., Sandell, L. and Schurman, D. (1996) In vitro stimulation of articular chondrocyte mRNA and extracellular matrix synthesis by hydrostatic pressure. J. Orthop. Res., 14, 53–60. Stromblad, S. and Cheresh, D. (1996) Cell adhesion and angiogenesis. Trends in Cell Biology, 6, 462–469. Sumpio, B., Banes, A., Levin, L. and Johnson, G. (1987) Mechanical stress stimulates aortic endothelial cells to proliferate. J. Vasc. Surg., 6, 252–256. Sumpio, B., Widmann, M., Ricotta, J., Awolesi, M. and Watase, M. (1994) Increased ambient pressure stimulates proliferation and morphologic changes in cultured endothelial cells. J. Cell Physiol., 158, 133–138. Swerlick, R., Brown, E., Xu, Y., Lee, K., Manos, S. and Lawley, T. (1992) Expression and modulation of the vitronectin receptor on human dermal microvascular endothelial cells. J. Invest. Dermatol., 99, 715–722. Tang, D., Diglio, C., Bazaz, R. and Honn, K. (1995) Transcriptional activation of endothelial cell integrin alpha v by protein kinase C activator 12(S)-HETE. J. Cell Sci., 108, 2629–2644. Tang, D., Grossi, I., Chen, Y., Diglio, C. and Honn, K. (1993a) 12(S)-HETE promotes tumor cell adhesion by increasing surface expression of vß3 integrins on endothelial cells. Int. J. Cancer, 54, 102–111. Tang, D., Timar, J., Grossi, I., Renaud, C., Kimler, V., Diglio, C., Taylor, J. and Honn, K. 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14 The Role of Hemodynamic and Mechanical Factors in Vascular Growth and Remodeling Olga Hudlicka*, Margaret D.Brown1 and Stuart Egginton Department of Physiology and1 School of Sport and Exercise Sciences, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK. *Corresponding author: E-mail address: [email protected]
This chapter explores how altered hemodynamics may control both the differentiation and physical extent of the vasculature. The differences between growth of new vessels and remodeling of existing vessels are emphasized, and an overview presented of the methods available for quantifying these processes. Concentrating mainly on skeletal and cardiac muscle, examples are given of how vessel architecture changes during development, while the diversity apparent from comparative studies illustrates the strategies which produce an efficient circulatory design. The response to variations in the major hemodynamic factors of blood flow and pressure are discussed, particularly their effects on conductance and wall thickness throughout different sections of the vascular bed. Physiological adaptations to increased skeletal muscle activity and altered heart rate are contrasted with the pathological changes seen in response to muscle hypertrophy and atrophy. The action of hemodynamics and mechanical factors, primarily through differential effects on shear rate and wall tension, lead to characteristic differences in the forms of angiogenesis, elicited by either luminal or abluminal signals. Finally, other humoral factors involved in vascular growth and remodeling are examined, in the context of possible mediation of transduction of hemodynamic and mechanical forces. KEYWORDS: Shear stress, wall, tension, stretch, capillaries, arterioles, large vessels.
INTRODUCTION Research into the role of mechanical factors in vitro in the growth of vascular endothelial and smooth muscle cells has expanded enormously during the last decade, manifested by the proliferation of knowledge of signal transduction mechanisms and molecular basis of these processes described in this book. The role of mechanical factors in vivo, however, is currently acknowledged widely in only a few situations, such as development or hypertension. Growth factors rather than mechanical factors are considered to be much more important in vivo regulators of growth. Yet increased or decreased blood flow or blood pressure are linked with growth of vessels or their regression under many circumstances. High blood flow occurs concomitantly with capillary growth in physiological conditions such as exercise or exposure to high altitude, and is also a feature of tumors, where its restriction has been used successfully to cause their regression (Denekamp et al., 1983). In wounds, vasodilation precedes capillary growth (Hughes and Dann, 1941) 291
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and it is an accompanying sign of other inflammatory diseases linked with angiogenesis such as arthritis or psoriasis. Folkman and Shing (1992) commented, nevertheless, that the role of vasodilation has been overlooked as an important initial step in the angiogenic cascade. Other mechanical factors such as deformation of the extracellular matrix or cell membrane occurring when tissues are stretched or when shear stress is elevated by increased velocity of blood flow may be necessary for the activation of various cytokines involved in angiogenesis (see Folkman, 1997). One reason why the role of mechanical factors in vascular remodeling in vivo has been rather neglected is the fact that the evidence is somewhat circumstantial, putative transduction mechanisms being difficult to study. What is known, however, can hopefully contribute to an understanding of their role in vessel growth or involution under both physiological and pathological circumstances. The first person to point out the relationship between blood flow and growth of vessels was probably John Hunter (1794) who observed enlargement of the carotid arteries in deer at the time of vascular growth in newly-formed antlers. This was confirmed almost 200 years later by direct blood flow measurements on the temporal artery by Hove and Steen (1978). Thoma in 1893 first established the principles of what he called ‘histomechanics’ while studying development, and later, pathological conditions (Thoma, 1911). These principles dictate that increases or decreases in the size of a blood vessel lumen depend on the rate of flow; decreased or increased vessel length is determined by the tension exerted on the vessel wall in a longitudinal direction by the surrounding tissue; increases or decreases in vessel wall thickness are dependent on blood pressure, wall tension being related to blood pressure and vessel radius. The formation of new capillaries is dependent upon capillary pressure and the pressure of the surrounding tissue; thus, capillaries grow more slowly at the surface of the brain where tissue expansion is limited by the dura mater. Clark (1918) observed in tadpole tails that capillary sprouting occurred preferentially from capillaries with high flow while those with low flow regressed (Figure 14.1). He and colleagues (Clark et al., 1931) showed a similar pattern in an implanted ear chamber in rabbits where they also demonstrated that capillaries with high flow gradually changed into venules or arterioles. Brånemark (1965) suggested that oscillatory movements of erythrocytes are important in the ‘opening up’ of newlyformed sprouts and such movements were indeed seen in capillary sprouts in skeletal muscles subjected to a long-term increase in contractile activity by electrical stimulation (Myrhage and Hudlicka, 1978). Rodbard (1971) also stressed the importance of mechanical forces not only in growth, but also in the regression of the vessels such as the ductus arteriosus or uterine vessels after parturition. There are obvious parallels between the mechanical forces acting on blood vessels in vivo and those used to study signal transduction in vitro. Increased blood flow, for example, presents a complex stimulus in vivo involving velocity of flow, viscosity and changes in vascular wall diameter and hence tension. These components can be isolated in experiments in vitro and the role of the individual factors—shear rate, shear stress or stretch—studied separately. In addition, cyclic strain applied to cells in vitro mimics in vivo changes in pulse pressure, which are sometimes more important than changes in shear stress. As will be shown later, the importance of these factors
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Figure 14.1. Dependence of the development of the capillary bed in tadpole tail on flow. Capillaries with fast flow enlarged and have new sprouts, capillaries without flow disappeared (marked by circles). Modified from Clark, 1918.
varies in vessels of different categories with wall tension being more important in the remodeling of arterioles while shear stress plays a role in growth of capillaries. The effects of stretch of surrounding tissue, so important for vascular growth and remodeling during development, can be reproduced by growing cells on flexible/ distensible substrata and cells can be exposed to variable pressures. In vivo, however, these factors affect the entire vascular wall, not just individual types of cells, and this is an important consideration when conclusions from in vitro experiments are applied to the situation in vivo or vice versa.
REMODELING VERSUS GROWTH OF BLOOD VESSELS The difference between remodeling and growth appears to be very simple. It is generally assumed that growth equals cell proliferation, while remodeling is connected with
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cell elongation, hypertrophy or rearrangement. Examples of capillary growth during development and changes in larger vessels exposed to increased pressure in different forms of hypertension show that it is not always possible to draw a line between these two processes simply on the basis of the above criteria. For instance, growth of microvessels in the rat brain during the earliest postnatal period proceeds by sprouting with abundant endothelial cell mitosis, but after the first week it continues by elongation of individual endothelial cells (Bär, 1983). Yet this elongation obviously requires some protein synthesis as it was attenuated in protein-deprived animals in small (<8 µm) but nor larger arterioles (Conradi et al., 1979). Another example of growth and remodeling occurring at the same time is the change in vasculature in rat incisors, one of the very few organs in adults in which vessel growth continues, along with growth of the teeth. Each artery supplies a section of arterioles which undergo cycles of proliferation at the branching point from the feed artery, elongation until they reach migrating capillaries and degeneration when the capillary network contacts another neighbouring arteriole (Moe, 1981). The number of arterioles supplying an individual section is dependent on the length and thus the growth rate of the tooth. The most clear cut difference between growth and remodeling is demonstrated by the arterial vasculature in hypertension. Growth of arteries implies both an increase in the thickness of the media due to hypertrophy or hyperplasia of smooth muscle cells (SMCs) and increases in the external diameter of vessels to accommodate the greater volume of these newly-grown cells, while remodeling involves realigning of existing tissue elements. By this definition, there is growth in small arteries and remodeling in large arteries in spontaneous hypertensive rats (SHR) (Heagerty et al., 1993) and remodeling but not growth in human essential hypertension (Schiffrin and Hayoz, 1997). Hypertension produced by treatment with a nitric oxide synthase inhibitor L-NAME resulted in remodeling with an increased media/lumen ratio, but not growth (Moreau et al., 1995) whereas both remodeling and growth were found in small arteries in animals with Goldblatt type hypertension (Bund et al., 1991). It is generally assumed that either growth or remodeling are due to increased pressure but experiments performed by Bund et al. (1991) and Mulvany (1992a) indicate that increased flow is as much, if not more, important, both these hemodynamic factors activating various protooncogenes by stretching the vascular wall. Vascular smooth muscle contraction or relaxation can play a significant role in the regulation of growth as shown by the fact that agonists inducing vasoconstriction promote growth of vascular smooth muscle cells while those producing dilatation inhibit it (Dzau, 1993). An important part is played in the remodeling of the vascular architecture by the activation of proteases and subsequent modification of the extracellular matrix (ECM) so that the growing cells, particularly capillary endothelium, can make passage towards neighbouring vessels (Pepper and Montesano, 1990; Tkachuk et al., 1996). The extent to which the proteases can be activated by hemodynamic factors in vivo is not known. Capillary growth can also occur in the absence of proliferation by ‘intussusceptive growth’ (Burri and Tarek, 1990; Patan et al., 1996) when columns of connective tissue gradually penetrate and occlude a capillary, dividing it into two. This type of
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growth occurs mainly during development and may be due to the fact that expansion of the connective tissue proceeds faster than proliferation of endothelial cells.
METHODS FOR STUDYING VASCULAR GROWTH AND REMODELING Appropriate methods for studying the vasculature in the whole organism will depend on whether growth per se and/or remodeling is to be quantified, on whether large or small vessels are the focus of attention, and on the nature of the tissue under examination. Figure 14.2 illustrates the different approaches that can be used. Visualisation of the vascular bed by injection of particulate dyes such as India ink and chemical clearing of tissue has been used in different organs since the last century (Spälteholz, 1888; Krogh, 1919) and, despite some uncertainty with degree of filling, is still helpful if the injectate is dialysed and filtered to permit only the smallest
Figure 14.2. Schema showing different methodologies appropriate for studying growth and remodeling of the vasculature depending on the imaging properties of the tissue, the size of the vessels and/or analytical resolution required. For translucent tissues—right—individual small vessels and their hemodynamics can be visualized directly, aided by the use of special dyes. Apart from the most superficial layers in thick tissues, indirect visualization is required—left— using intravascular dyes and penetrating illumination, or by producing replicas of the vascular structures. Functional correlates of the vascular bed can be assessed by flow measurements from entrapment of labelled particles. For all types of tissue—centre—cell proliferation can be quantified in samples taken after infusion of markers that are incorporated into nuclei during mitosis. Using the light or electron microscope, spatial distribution and cellular structure of large and small vessels can be quantified by application of stereological principles.
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particles to be used. For example, carmine dye in gelatin was used to demonstrate inflammatory angiogenesis in mouse granuloma (Colville-Nash et al., 1995). The size and topology of the whole vascular bed can be assessed by casting with curable latex or resin, which is allowed to harden and exposed by digestion of the surrounding tissue at high pH. Viscosity and perfusion pressure must be carefully controlled to ensure complete filling and avoid vessel rupture (Lametschwandtner et al., 1990). Cast volume (weight corrected for density) as a proportion of total organ volume gives an indication of changes in total vascular supply (e.g. Dawson and Hudlicka, 1989) but it is difficult to quantify any growth or remodeling in vessels of different size classes even though both large and small vessels can be observed with scanning electron microscopy (SEM). Gross structural features of the vascular bed have also been studied using angiography which shows visible filling of existing and newly-formed vessels, and is used mainly to assess the development of collateral circulation (e.g. Schaper, 1971). Again this method does not permit ready identification of different vessel categories to determine which are the new vessels. More recently, confocal microscopy, which uses a scanning raster of laser light to visualize ‘optical slices’ (0.2 µm thick) of tissue in which vessels have been labelled with e.g. fluorescent lectins, has been used for detailed examination of three-dimensional architecture of the vascular bed to establish patterns of growth (Hansen-Smith et al., 1996a, see Figure 14.10). In functional terms, the capacity of the whole vascular bed can be assessed in vivo by measuring the maximal, vascular conductance—maximal blood flow/blood pressure—(or minimum resistance) as an index of growth of arterioles and conduit arteries. In ischaemic myocardium, a good correlation existed between the increase in arteriolar density and cross-sectional area and rise in maximum conductance over time (White and Bloor, 1992). Standard histological techniques and light microscopy are used for identification of all vessels and evaluation of their growth, although clear distinction between arterial and venous vessels is not always possible at this magnification. Immunohistological staining for alpha-smooth muscle actin can distinguish arterial vessels down to quite small sizes and has been successfully used to show arteriolar growth in skeletal muscles subjected to electrical stimulation (Hansen-Smith et al., 1998) or chronic vasodilation (Price and Skalak, 1996). However, if larger vessels are analysed quantitatively to assess their growth in terms of numbers, their relatively low density compared to that of capillaries needs to be evaluated in a large enough sample area. Methods for evaluation of capillary growth are of fundamental importance, since most angiogenesis in the adult organism occurs by this route. The specificity of markers is critical because most quantitation is based on examination and counting vessels under the light microscope. Evaluation of capillary supply (and other tissue elements) can be made after staining of basement membrane for the periodic acidSchiff/amylase method which identifies carbohydrate moieties, silver methenamine, or with antibodies to laminin, fibronectin or Type IV collagen. Histochemistry of capillary endothelial enzymes such as alkaline phosphatase, serine protease, ATPase or carbonic anhydrase are suited for use on frozen tissue sections but staining pattern can vary among species, between organs, according to the state of ontogenetic
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development or in certain pathologies. For this reason, these methods should be validated against low power electron microscopy to confirm the same capillary counts (Egginton, 1990). Certain endothelial surface antigens such as CD31/PECAM and QBend10 can be demonstrated immunohistochemically, with Factor VIII-related antigen particularly suited for use in human tissue. Lectin staining of the sugar residues of cell surface glycoproteins is also widely used, again with species specificity e.g. lectin Ulex europaeus to depict capillaries in human tissues and Griffonia simplicifolia to react with vessels in many animal tissues (Holthofer et al., 1982; Hansen-Smith et al., 1988). Proper evaluation of growth and remodeling can be achieved if structural markers of large and/or small vessels, as described above, are combined with labelling of cells undergoing proliferation since this enables precise localization of sites and cell types and vessel classes actively involved in angiogenesis. Antibodies are available to nuclear components specific to periods of the cell division cycle, e.g. 3H-thymidine, bromodeoxyuridine (BrdU) and Ki-67 antigen during the S-phase of division, PCNA labelling for G1+S+G2 phases, the first two of these requiring pre-treatment by administration of the marker (Figure 14.2). Such methods have been used to demonstrate specific endothelial proliferation in hypo- and hyperthyroid hearts (Heron and Rakusan, 1995) or hearts with irradiation damage or infarction. Quantitation of vessel growth is often based on simple vessel counts. The effects of expansion of the surrounding tissue during development, hypertrophy or tumor growth must be accounted for because this will influence straightforward vessel counts by increasing vessel separation. In this case, it may be more useful to express capillary supply in terms of a ratio of numbers of vessels to units of tissue area (e.g. capillary/ fiber area ratio in skeletal or cardiac muscle). Three-dimensional aspects of growth and vessel tortuosity can only be evaluated using stereological analyses (Mall et al., 1987). This is useful since most capillary beds form a complex network with numerous interconnections, such that both effective capillary length and surface area are greater than those estimated from simple counts in tissue sections. An index such as length density, Jv, which evaluates the length of capillaries per unit volume of tissue (e.g. in km.cm-3), can be multiplied by tissue volume to determine the absolute volume of the capillary bed. The impact of vessel tortuosity on quantitative assessment varies considerably in importance in different organs, being highly significant in the brain where capillaries have an apparently random orientation. It may account for 1.6– 42% of length density even in skeletal muscle where capillaries lie alongside muscle fibers (Mathieu-Costello, 1993). If mean capillary dimensions are known then capillary volume and surface densities can be calculated from the product of Jv and capillary cross sectional area and perimeter, respectively. These, and other structural indices will give estimates of the maximal capacity of a micro-vascular network and permit more accurate evaluation of angiogenic growth, and have been examined in more detail elsewhere (Egginton, 1990). Electron microscopy provides a number of approaches to assessment of growth and remodeling, from analysis of capillary supply under low power magnification, to detailed ultrastructural observations of cells undergoing proliferation or expansion. Signs of initial endothelial activation during angiogenesis include increased organelles content (mitochondria, cytoplasmic vacuoles, endoplasmic reticulum and
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occasionally Golgi apparatus) or altered structure (greater electron translucency of the cytoplasm, increased luminal or abluminal surface roughness). Proliferation can be detected as increased numbers of cell profiles per capillary at high magnification and although mitotic nuclei may be rare by virtue of the low probability of fixing cells during the short time spent in this phase of the cell cycle, labelling after BrdU incorporation or for PCNA is possible by linking appropriate antibodies to an electron opaque stain (usually colloidal gold).The nature of interaction between vascular and perivascular cell types such as pericytes can also be established at high magnification. For example, the proportion of capillary outer surface covered by pericytes has been shown to be reduced during angiogenesis in skeletal muscle and heart (Egginton et al., 1996). Growth of small vessels—arterioles, venules and capillaries—can be investigated by intravital microscopic observations in living organisms (Clark, 1918, Figure 14.1) More recently, a classical description of growth of the microcirculation was presented by Rhodin and Fujita (1989) who combined intravital observations of vessels with electron microscopy in the rat mesentery during early postnatal development. Intravital observations have also been made of the patterns of capillary growth in tumors (e.g. Reinhold and Berg-Block, 1983) and capillary sprouts were demonstrated even in normal muscles exposed to chronically increased activity (Myrhage and Hudlicka, 1978). In addition to providing data on vessel diameters and lengths, intravital observations can allow quantification of changes in vascular branching pattern (internode length, number of branches of a given order, branching angle) particularly in thin tissues such as the mesentery or essentially planar muscles like the spinotrapezius. They also enable measurements of flow velocity and vessel diameters which are required for the estimation of the wall shear stress and tension and hence this method, in combination with quantitative evaluation of the microvascular supply, has provided much of the evidence upon which we base our understanding of the role of hemodynamic factors in growth and remodeling.
GROWTH AND REMODELING DURING PRE- AND EARLY POST-NATAL DEVELOPMENT The pattern of embryonic and neonatal growth of the vascular system in many different tissues and organs has been reviewed in depth by Hudlicka (Hudlicka, 1984; Hudlicka and Tyler, 1986). In the embryo, development of the vascular supply proceeds by initial differentiation of endothelial cells from precursor mesenchymal angioblasts (vasculogenesis) and by sprouting and/or elongation once endothelial cells are assembled (angiogenesis) (Noden, 1989; Beck and D’Amore, 1997). The time course of this process differs according to species and organ. For example, in the rat embryo, the coronary terminal vascular bed originates from sinusoids within the cardiac musculature (see Rakusan, 1984) with connections from outgrowths of endothelial cells in the region of the aortic sinus forming definite coronary vessels by day 17 (Blatt, 1973) and final maturation of capillaries after birth; in humans, the process is completed much earlier, in the first quarter of pregnancy (see Hudlicka and Tyler, 1986).
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The coordinated growth of a vascular network is often accomplished by vessel structures originating from different locations by both vasculogenesis and sprouting ultimately making connections. Lung vasculature in the mouse embryo forms from endothelial cell-lined spaces creating sinusoids at day 9 which connect with arteries and veins sprouting from the central pulmonary vascular trunks at days 13–14 (DeMello et al., 1997). Likewise, the vasculature of the avian limb bud proceeds by a combination of sprouting of vessels and recruitment of angioblasts (Brand-Saberi et al., 1995). Electron microscopic studies of the rat embryo reveal that at day 11.5 it is possible to distinguish the arterial from the venous types of endothelial cells which make up the capillary network on the basis of their morphology (Mensah-Brown, 1988). Differentiation of vascular smooth muscle cells can be characterized by alpha-smooth muscle actin at multiple sites throughout the vasculature of the mouse embryo between days 9.5–11.5 (Takahashi et al., 1996), and appear later in the rat embryonic kidney than in other vascular beds (Carey et al., 1992). Larger vessels are all formed from capillary plexuses (Woolard, 1922), and the origin of smooth muscles cells is assumed to be pericytes and/or fibroblasts which appose themselves to the capillary vessels and differentiate (Thayer et al., 1995). Many of the very early observations on embryonic vascular development related its modeling to mechanical factors connected with blood circulation (Roux, 1878; Thoma, 1893), as was reinforced by the work of Clark (1918) described previously (Figure 14.1). Despite this, there are, because of methodological considerations, relatively little data on measured hemodynamic parameters such as shear stress, wall tension and pressure which can be applied to embryonic vascular growth and remodeling. At this stage of development, current interest is more focused on regulation of vascularization by cell-cell interactions and by growth factors, aided by studies of genetically manipulated mice (Beck and D’Amore, 1997). Vascular endothelial growth factor (VEGF) has been identified as a key developmental determinant since lack of a single VEGF allele leads to abnormal blood vessel development and embryonic lethality (Carmeliet et al., 1996), and VEGF itself and its receptors, flk-1, flt-1, are expressed, together with the endothelial specific receptor tyrosine kinase tie, in mouse embryos from day 8.5 onwards (Korhonen et al., 1994; Dumont et al., 1995). Another growth factor, angiopoietin-1 and its endothelial cell receptor tie-2 are also essential for normal blood vessel development in the mouse, acting in balance with a naturally occurring tie-2 antagonist, angiopoietin-2 (Maisonpierre et al., 1997). mRNA for platelet-derived growth factor beta (PDGF-ß) receptor, was also expressed in blood vessel endothelium from day 9.5–16.5 in the mouse embryo (Shinbrot et al., 1994). Whether the expression of these factors is modified by hemodynamic stimuli once patent vasculature is established remains to be determined. Expression of VEGF is known, however, to be increased directly by hypoxia in adult tissues (Ladoux and Frelin, 1993; Minchenko et al., 1994). Meuer and Bertram (1993) studied red blood cell velocity and vessel length in the vitelline capillary network of 4 day old chick embryos and calculated that transit times were shorter than required for full blood oxygenation in one third of vessels, a situation conducive to hypoxia. Embryonic vessel growth is very sensitive to hypoxia, as shown by exposure of chick embryos to hypoxic conditions (12% oxygen) which
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significantly increased the total number of vessels in the CAM (Strick et al., 1991) and decreased the resistance of maximally dilated vasculature in 14–15 day old embryos (Adair et al., 1988). Hypoxia could therefore contribute to vascular development in the embryo by enhancing VEGF expression, until such time as adequate flow is established, or possibly by causing dilation of developing vessels and consequent flow-related stimuli. Dilation of vessels could also be modified by nitric oxide (NO) dependent relaxation. In fetal lambs, the NO-dependent relaxation of intra-pulmonary arteries and eNOS expression were attenuated by creation of pulmonary hypertension and were proposed to account for the excessive muscular development of the fetal pulmonary circulation in this condition (Shaul et al., 1997). The exact role of NO in vascular development is as yet unclear, but NO metabolizing neurones have been found in close apposition to blood vessels in human fetal brain (Yan and Ribak, 1997). The growth of arterial smooth muscle in the chick embryo coincides with the greatest relative increase in mean blood pressure (see Hudlicka and Tyler, 1986). The collagen and elastin content of the thoracic and abdominal aortic wall in sheep also increased most rapidly between 140 days gestation and 3 days postpartum, preceding a marked postnatal increase in arterial pressure (Bendeck and Langille, 1991). The content later decreased in the abdominal but not thoracic aorta, supposedly linked to the dramatic decrease in flow through this vessel upon loss of the placental circulation (Bendeck and Langille, 1991). In the chick embryonic yolk sac vasculature, pulsatile flow related to cardiac contraction was measured in larger vessels but not in the capillary bed (Gush et al., 1990) but it is not known if this is a cause or consequence of vessel size. One of the most potent factors regulating developmental vessel growth is contact with the extracellular environment and the stretch imposed by expansion of the surrounding tissue. This was clearly identified by Thoma (1893) as a determinant of vessel length, and there are many examples. Stretch applied directly by traction to the CAM in 6–8 day chick embryos increased vessel density (Yamashita et al., 1989) whereas in human hearts, casts of the coronary arteries showed that the length but not number of branches increased after birth in line with heart size (Reinecke and Hort, 1992). Total capillary length also increased in rat brain after birth (Bär, 1983) in growing rat skeletal muscles, the capillary network is formed from a plexus which is gradually stretched under tension from the elongating muscle fibers (Wolff et al., 1975). Contact between endothelial cells and the extracellular matrix constituents is clearly very important for this effect of stretch. Initial capillary networks are formed without a basement membrane, which develops later on (Ausprunck, 1982). Rongish et al. (1996) showed that fibronectin is deposited even before vasculogenesis on day 13 in the rat embryonic heart, with subsequent appearance of laminin which was linked to vessel tube formation and development of collagen IV, a component of the basement membrane. Fibronectin and laminin have also been shown to be crucial for adhesion between endothelial cells of existing vessels and mesenchymal cells in embryonic rat hind limb buds (Kanazawa et al., 1996) and the prenatal rabbit phallus (Hara et al., 1994). When embryonic day 12 avascular rat hearts were transplanted into the anterior eye chamber of adult hosts, the distribution of extracellular matrix components
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providing a scaffold for endothelial cells was not altered even though the hearts were hemodynamically unloaded (Rongish et al., 1996). Postnatal growth of the capillary bed in many tissues is by angiogenesis of new vessels from the existent network. In rats, 90% of the total number of capillaries found in adult hearts are formed postnatally (Rakusan, 1984) and intracortical capillary length density increases 5.5-fold between 6 and 20 days postpartum, while brain weight only doubles (Bär, 1983). The sequence of postnatal angiogenesis was described in detail for the rat mesentery from intravital observations by Rhodin and Fujita (1989), showing endothelial cell migration, proliferation and sprouting. However, other forms of postnatal growth are encountered in e.g. the heart, where division of existing vessels by splitting was seen in vascular casts (Van Groningnen et al., 1991) or in the lung, where ‘intussusceptive’ growth—penetration and division of a capillary by a column of connective tissue—occurs without proliferation (Burri and Tarek, 1990; Patan et al., 1996). Degradation of the ECMs necessary to accommodate the new vessels, and interstitial collagenase, capable of accomplishing this, was found in vessels undergoing sprouting and elongation in human fetal skin rather than gelatinase A and B which can break down Type IV collagen of the basement membrane (Karelina et al., 1995).
DIFFERENCES IN VESSEL ARCHITECTURE IN ADULT ORGANS WITH RESPECT TO HEMODYNAMIC FACTORS Studies of comparative physiology demonstrate the role of different mechanical and hemodynamic factors in the structure of the vascular bed, particularly capillaries. In skeletal muscles, differences in the size of fibers and their rates of growth are an important determinant in the development of the capillary network. The large fibers usually push capillaries apart and thus capillary density decreases. This basic relationship has been established in individual muscles in fishes (Egginton, 1992), birds (Snyder, 1990), rodents (Banchero, 1982) and humans (Ahmed et al., 1997). One unusual adaptation seen in fishes, however, is that the capillaries in muscles with larger fibers have a greater cross-sectional area (Egginton, 1992) which compensates for the reduction in capillary density. As their perfusion pressure appears to be maintained, shear stress in these capillaries will be lower and wall tension higher. In addition to fiber size, degrees of activity and the local environment can lead to specific adaptations of the capillary bed in, for example, skeletal muscles. Long-term exposure to low temperatures results in a high capillary density which could be due to increased shear stress resulting from a combination of high blood flow, described during acute cooling by Barcroft et al. (1955) and high viscosity (Heroux and St Pierre, 1957). In natural hibernators, like the ground squirrel and hamster, increased capillary density on cold exposure is also partly due to fiber atrophy (Steffen et al., 1991; Fairney and Egginton, 1994). In fish, in contrast, adaptation to a cold environment is linked with fiber hypertrophy as they often maintain locomotor activity, and capillary per fiber ratio (C:F) is increased (Egginton and Sidell, 1989). Coldinduced angiogenesis perhaps most clearly illustrates the role of hemodynamic and
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other mechanical factors within the same tissue. For instance, during seasonal acclimatization of trout there was an inverse linear relationship between environmental temperature and C:F which increased 20% when temperature changed from 18°C in summer to 11°C in the autumn, and was accompanied by higher blood flow and greater capacity for aerobic swimming. A similar increase in C:F between autumn and winter (4°C) was, however, accompanied by extensive fiber hypertrophy but reduced blood flow and impaired swimming performance (Egginton and Cordiner, 1997; Taylor et al., 1996), In contrast to cyclic adaptations to cold described in the above species, the extremely stable Antarctic environment has produced endemic fishes with a low hematocrit and a reduced blood hemoglobin concentration (Macdonald et al., 1987). This situation is taken to the extreme in the icefish (channichthyids) that possess only a few vestigial erythrocytes and no respiratory pigment. The low blood pressure and large cardiac output reported for icefish would best be accommodated by a relatively low afterload on the heart. All icefish species so far examined had a functional hypobranchial network (Rankin et al., 1987), an unusual anastomosing dual arterial blood supply to the main locomotory (pectoral) muscles. Comparison of vascular casts under SEM showed that all microvessels were around 3-fold wider than comparable vessels in rainbow trout (arterioles 54 v. 18 µm, capillaries 11 v. 3 µm and venules 54 v. 20 µm respectively, Egginton and Rankin, 1991). In contrast to other fishes, which exhibit relatively few connections between individual straight capillaries, capillaries from icefish show numerous capillary loops which represent true divergent and convergent capillary branches and not the arcade microvascular network observed in flat mammalian muscles. This high tortuousity in icefish maximises the contact area with muscle fibers, although the density of microvessels is much less than found in other teleosts (Fitch et al., 1984). The icefish is also an example of an animal with an extremely limited locomotor activity. Muscles adapted for unusually rapid and sustained activity have small mitochondriarich fibers with a high local capillary supply. Examples are the rattlesnake shaker muscle (Clark and Schultz, 1980) which is able to sustain a rate of contraction around 50Hz for at least three hours, fish swimbladder muscles where the contraction rate determines the fundamental frequency of the sound during displays, and bat or hummingbird pectoral muscle that structurally resembles myocardium (MathieuCostello, 1993). Whether tortuous capillary path lengths can be induced by remodeling in the face of sustained high flow rates or whether they exist as an adaptation in order to increase erythrocyte transit times remains unclear. Adaptation to the high hydrostatic pressure as experienced by the microcirculation in limbs of the giraffe have resulted in capillaries of very low permeability, necessary to avoid pathological edema (Hargen et al., 1987). A similar problem of high hydrostatic pressure in the lower part of the body in treeclimbing snakes is regulated physiologically, rather than morphologically, by localized vasoconstriction of vascular smooth muscles (Lillywhite, 1987) and a greater sensitivity to catecholamines (Conklin et al., 1995). In contrast, fish are exposed to minimal gravity effects and presumably have no regional differentiation among vessels which show a high permeability.
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INCREASED BLOOD FLOW AND PRESSURE: ROLE IN REMODELING OF THE VASCULAR BED Increased Blood Flow The effects of increased blood flow upon remodeling of large vessels and growth of capillaries have been widely studied, as have the effects of increased blood pressure on all type of vessels in different forms of hypertension. Folkow (1987) summarised the difference between these two stimuli as follows: increased blood flow in response to elevated metabolism or enlargement of a tissue mass results in increased lumen size in larger vessels and growth of capillaries, whereas increased blood pressure without any change in metabolism leads to a decrease in maximal flow capacity due to a high vessel wall media/lumen ratio and hence decreased vessel diameter but there is no change in the size of the capillary bed. Increasing the flow in a large artery such as the carotid by arterio-venous (A-V) shunting between the carotid artery and the external jugular vein resulted in a larger arterial radius (Kamiya and Togawa, 1980) (Figure 14.3). Langille (1993) referred to examples of remodeling of large arteries in response to increased flow, such as the diameter of the abdominal aorta increasing by 50% during development or by 20% during pregnancy to meet the high metabolic demands. Increased flow due to A-V shunt in patients on dialysis also increased the diameter of the relevant artery within about a month. Using an A-V shunt between the abdominal aorta and vena cava, BenDriss et al. (1997) found proliferation of endothelial cells but decreased thickness of the media in spite of increased wall tension and tensile stress. In contrast, Lehman et al. (1991) demonstrated smooth muscle cell hyperplasia in arteries exposed to increased blood flow in collateral circulation after ligation of common and internal carotid arteries. Increased blood flow is also an important stimulus for growth of capillaries although its causal role is not always recognized. This is demonstrated in the case of long-term exposure to hypoxia and capillary growth. The fact that myocardial capillary growth in animals exposed to high altitude hypoxia occurs only in the right but not the left ventricle (Turek et al., 1972) and that blood flow is increased in the right ventricle, which has to work harder to overcome the increased pulmonary resistance, but not in the left (Turek et al., 1975; Kasalicky et al., 1977) is usually ignored. Similarly, expansion of the volume of microvessels by elongation of existing cells in the brains of rats exposed to hypobaric hypoxia (Harik et al., 1995) can be explained on the basis of increased blood flow (Lamanna, 1996). There are numerous examples of direct evidence for the role of blood flow in vascular growth from studies showing capillary growth in response to long-term administration of vasodilating drugs in both normal and pathologically affected organs, reviewed by Hudlicka et al. (1992). For example, ethanol, which increases coronary blood flow (Abel, 1980), resulted in myocardial capillary growth (Mall et al., 1981, 1982). Likewise, adenosine or propentofylline, both of which increased blood flow in heart and skeletal muscle, led to increased capillary supply in both tissues after 3–5 weeks (Ziada et al., 1984), whereas prazosin, which did not increase flow in the heart but almost trebled flow in skeletal muscle, had no effect on the
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Figure 14.3. Changes in radius and flow ratios (after shunt/before shunt)—top—and in shear rate ratio % (after shunt/before shunt x 100)—bottom—in carotid arteries. Surgical shunt was created between the artery and jugular vein. Different symbols denote the number of days or weeks after the shunt was performed. Modified from Kamiya and Togawa, 1980, with permission from the authors and the American Physiological Society.
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former but increased capillary supply in the latter (Ziada et al., 1989; Dawson and Hudlicka, 1990a), and caused growth of arterioles (Price and Skalak, 1996). The type of capillary growth in skeletal muscle after prazosin treatment was unusual, occurring without the appearance of abluminal sprouting as is usually described in vitro in chorio-allantoic membrane (CAM) or in tumors (Ausprunck and Folkman, 1977), but by luminal longitudinal division (Zhou et al., 1998a). Propentofylline which increased blood flow in the brain also increased capillarization in the parietal cortex (Hudlicka et al., 1997) (Figure 14.4).
Figure 14.4. Blood flow in rabbit heart and brain during 3–4 hours i.v. administration of adenosine or the xanthine derivative propentofylline (HWA285) and in rat heart during i.v, prazosin, compared with control values before drug administration (=100%). Myocardial and brain capillary density (number of capillaries.mm-2) was measured after chronic administration of the drugs for 3–5 weeks. Drugs which increased coronary and brain blood flow (adenosine and HWA285) induced capillary growth; prazosin, which did not affect coronary flow, did not cause growth.
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In the heart, capillary growth after long-term administration of dipyridamole, evidenced by increased capillary supply, uptake of 3H-thymidine (Tornling, 1982) and increased total capillary length (Mall et al., 1987), was linked with dipyridamoleinduced dilation predominant in microvessels (Chilian et al., 1989). Growth of larger vessels did not occur in this model as minimal coronary resistance was not altered (Connel et al., 1990). Dipyridamole also stimulated growth of capillaries in skeletal muscle (Tornling et al., 1980a,b), albeit less than prazosin (Hudlicka, 1991). It is arguable that pharmacological vasodilators may stimulate growth of vessels by direct mitogenic action. Adenosine is taken up by coronary endothelium (Nees et al., 1980) and stimulates endothelial proliferation in tissue cultures (Meininger et al., 1988). Prevention of adenosine uptake into coronary micro vessel endothelial cells eliminated its positive effect on cell proliferation (Meininger and Granger, 1990) but dipyridamole had no direct effect on endothelial cells in cultures (Jakob et al., 1982). On the other hand, it significantly increased red blood cell velocity in cardiac microcirculation (Tillmans et al., 1977), dilated terminal arterioles (Tillmans et al., 1982) and slightly increased the pressure in venules (Tillmans et al., 1981) which could all increase both shear stress and capillary wall tension (Hudlicka et al., 1995). The stimulatory effect of other vasodilatory drugs can likewise be attributed to increases in shear stress or capillary wall tension. In pathological conditions, increased blood flow is equally effective in vascular remodeling. Long-term vasodilation induced by the calcium entry blocker nifedipine was shown to reverse capillary decrement in hypertrophic hearts of SHR rats (Turek et al., 1987), increase capillary length and diminish medial thickness in coronary arteries in the same species and increase coronary vascular reserve in hypertensive patients (Motz and Strauer, 1994). Coronary artery size was also greater in dogs with severe anemia and higher flow (Scheel and Williams, 1985). Villari et al. (1992) studied patients with aortic valve insufficiency, noting that the size of the left descending and circumflex coronary arteries was doubled even though coronary perfusion pressure remained unchanged. After valve replacement, coronary artery size regressed although not quite to the values of control subjects (Figure 14.5). This illustrates the plasticity of response of the larger arteries to the effects of flow.
Increased Blood Pressure Although it is difficult to separate the effects of increased pressure from those of increased flow, it is clear that pressure is the main stimulus for remodeling of large arteries and smooth muscle cells, either by hypertrophy or hyperplasia. This is also the case for veins used in coronary by-passes, shown experimentally by Goss (1978), and in the pulmonary circulation, where small arteries in the fetus have a much thicker media than three days after birth by which time pulmonary artery pressure has fallen (Reid, 1979). Capillary growth, on the other hand, is probably not stimulated by pressure alone. Remodeling of the vascular bed takes different forms in hypertension of various origins-SHR rats, hypertension of renal origin or spontaneous hypertension in man, or locally increased pressure due to stenosis of large arteries such as the aorta or pulmonary artery.
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Figure 14.5. Enlargement of the coronary arteries in patients with volume-overload cardiac hypertrophy due to aortic valve disease, and its partial regression after surgical valve replacement. Ordinate—cross sectional area of different branches of the coronary arteries. Left hand side— in control patients (C); middle and right hand side—patients before (AVD-pre) and after (AVDpost) operation. Reproduced from Villari et al., 1992, with permission of the American Heart Association and the authors.
Hypertension in spontaneously hypertensive rats (SHR) In the SHR rat, remodeling is different in large arteries from that in arterioles, with smooth muscle cell hypertrophy accounting for the increase in the media/lumen ratio in the former, while smooth muscle cell growth occurs in the latter (Heagerty et al., 1993; Table 14.1). There are also differences in remodeling among vascular beds, smooth muscle hyperplasia occurring in arterioles of the brain, mesentery and kidneys (with the exception of the afferent arterioles) (Daemen and deMey, 1995) while the media/lumen ratio is increased in skeletal muscle (Chen et al., 1981; Wang and Prewitt, 1990). Medial thickening could be attenuated by lowering blood pressure locally by partial occlusion of the abdominal aorta in rat hindquarters (Folkow, 1978) or by reduction of pressure and flow into the brain by partial clipping of the carotid artery (Baumbach et al., 1991), an effect attributed mainly to lowering of pulse pressure and vessel wall strain. This was also emphasized by experiments
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Table 14.1 Remodelling of the vascular bed in hypertension -=increased; ¯ =decreased; 0—no change. D=density; P=pressure; SMC=smooth muscle cells; EC=endothelial cells.
References: (1) Mulvany (1992b) (3) Williams et al. (1990) (5) Bohlen et al. (1977); Folkow (1978) (7) Chen et al. (1981); Wang and Prewitt (19901) (9) Gray (1988) (11) Bund et al. (1991) (13) Tornig et al. (1996)
(2) (4) (6) (8) (10) (12)
Shore and Tooke (1994) Henrich et al. (1998); Heagerty et al. (1993) Zweifach et al. (1981) Huttner and Gabiani (1982); Engler et al. (1983) Hansen-Smith et al. (1996b)
where the Ca2+-blocker isradipine, which was the only drug to decrease pulse pressure from a selection of blood pressure-decreasing drugs, decreased wall/lumen ratio in mesenteric arteries (Baumbach, 1991). In skeletal muscle, ACE inhibitors lowered pressure and diminished the media/lumen ratio, and at the same time prevented the reduction in the arteriolar density (Wang and Prewitt, 1990; Scheidegger et al., 1996) which had been described previously by Chen et al. (1981). This arteriolar rarefaction could be due to a combination of high pressure in arterioles (Zweifach et al., 1981) and their limited capacity to dilate (StruijkerBoudier et al., 1988). As a consequence, capillary perfusion could be lower, which may be the cause of the decreased capillary supply in SHR (Struijker-Boudier et al., 1988). On the other hand, a more extensive study in a number of muscles by Gray (1988) did not indicate any loss of capillaries. The finding that capillary pressure is no different from control values (Bohlen et al., 1977) could explain the maintenance of the capillary bed in the presence of fewer arterioles and diminished perfusion. Reduced arteriolar numbers in the hearts of SHR animals together with a decreased lumen/media ratio can explain the increase in the minimal coronary resistance at rest (Vitullo et al., 1993) and diminished capacity for increasing flow (Wangler et al., 1982; Peters et al., 1984). Renal hypertension Vascular remodeling in renal hypertension is not always exclusively due to altered pressure since hormonal factors such as angiotensin II can directly affect growth of smooth muscle and endothelial cells. Tornig et al. (1996) studied changes in thickening of the aorta and myocardial arterioles in animals with partial nephrectomy after treatment with three different drugs—an ACE inhibitor, a calcium entry blocker and a vasodilator—to lower blood pressure. All three diminished medial thickening in the aorta but only the first two prevented it in arterioles.
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The density of skeletal muscle arterioles, particularly the smallest, is decreased in renal hypertension (Prewitt et al., 1984; Greene et al., 1990), occurring prior to a reduction in perfusion as measured by laser-Doppler (Hernandez and Greene, 1995). Both the hypertension and decreased arteriolar density were accentuated by high salt intake which depresses angiotensin II activity. When normal levels of angiotensin II were restored by cessation of high salt intake (Rieder et al., 1997) or by chronic infusion, the rarefaction of arterioles was abolished, possibly by activation of PDGF , c-fos and c-myc stimulating growth of smooth muscle cells (Hernandez et al., 1992). This emphasizes once more that hemodynamic changes in renal hypertension are not the sole cause of vascular remodeling. There is no doubt that the loss of arterioles itself can be not only a consequence but a cause of hypertension since a smaller vascular bed would accommodate lower blood flow and thus increase pressure provided that cardiac output remains unaltered. Increased levels of angiotensin II, in addition to stimulating growth of smooth muscle cells and increasing media/lumen ratio, may also promote growth of endothelial cells as described in small vessels (Engler et al., 1983) and in the aorta (Hüttner and Gabiani, 1982). Proliferation and/or hypertrophy of endothelial cells was sufficient in small skeletal muscle arterioles to occlude the lumen (Hansen-Smith et al, 1996b). Perfusion would be diminished and conditions thus created for obliteration of arterioles. In contrast to vessels in skeletal muscles, the wall/lumen ratio of coronary arteries has not been found to increase in renal hypertension, neither was there any difference in total coronary resistance. Any limitation in the capacity to dilate was more a result of the increase in myocyte mass and possibly collagen (Tomanek et al., 1986, 1991). Capillary density (Anversa and Capasso, 1991) and the population of very small capillaries (<4µm) (Tomanek et al, 1991) are increased, indicating capillary growth. Nevertheless, severe renal hypertension leads to heart hypertrophy and hence a lower capillary density, and this was exacerbated by exercise training such that capillary density was 25% lower than in normotensive trained animals with a similar degree of heart hypertrophy (Marcus and Tipton, 1985). To what extent this diminished capillarization is due to reduced numbers of arterioles and lack of capillary perfusion is not known although exercise did not seem to further impair coronary blood flow in rats with renal hypertension (Buttrick et al., 1985). Essential hypertension in man In essential hypertension in man, the increase in the media in large vessels is due mainly to proliferation rather than hypertrophy of smooth muscle cells (Mulvany, 1992b) while the increase in the wall/lumen ratio in small vessels is caused by realignment of vascular smooth muscle cells without signs of either hyperplasia or hypertrophy (Heagerty et al., 1993). Rarefaction of arterioles has been described in the conjunctiva and of capillaries in the nailfold (Shore and Tooke, 1994) and in skeletal muscles (Henrich et al., 1988). In contrast to animals, capillary pressure is significantly increased (Williams et al., 1990). Treatment with the ß2 agonist salbutamol to lower blood pressure prevented the loss of arterioles although it is not known to what extent it changed the media/lumen ratio. It also emphasizes the
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importance of adrenergic mechanisms in the development of hypertension stressed by Folkow (1995). The coronary flow reserve in the heart is decreased due to the hypertrophy of the media in small coronary vessels (Strauer et al., 1991) even in the absence of coronary artery disease (Strauer, 1992, Figure 14.6). A contributory factor is also fibrosis in the extravascular space limiting arterial dilation (Schwartzkopf et al., 1993). Increased pressure due to aortic or pulmonary artery stenosis or banding This usually results in heart hypertrophy, the vascular remodeling of which will be discussed in a later section of this chapter. Both these conditions also affect the vessels below or above banding and result in hyperplasia rather than hypertrophy of smooth muscle cells in the large arteries exposed to the high pressure (Bevan et al., 1976; Berry, 1978). Berry (1978) suggested that strain applied across smooth muscle cells converts them from a contractile to a synthetic phase resulting in cell division but he acknowledged that it is difficult to estimate the specific contribution of either pressure, flow or tension. In addition to flow and pressure, the surrounding tissue also plays a part in vessel remodeling. It has been suggested that increased radius in small vessels affects the surrounding tissue pushing it aside to create space for the vessel expansion (Waxman, 1981).
DECREASED BLOOD FLOW AND PRESSURE: ROLE IN REMODELING OF THE VASCULAR BED Decreased Blood Flow Restricted blood flow has been studied extensively in the heart, limbs or brain in connection with vascular diseases and the development of collateral circulation.
Figure 14.6. Coronary reserve ratio (ratio between resistance at maximum dilation produced by dipyridamole to that at rest) in controls normal subjects, and patients with different types of heart disease. Based on data from Strauer, 1992.
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Numerous studies have dealt with the likely involvement of different growth factors which have recently been used as therapeutic agents. The extent to which changes in hemodynamics are responsible for the development of collateral circulation is still unclear. There are relatively few, sometimes controversial, quantitative studies on remodeling of different sections of the vascular bed—large arteries, arterioles or capillaries—in situations of decreased blood flow or pressure. Guyton and Hartley (1985) found that restriction of blood flow in juvenile rat carotid artery to 35% did not affect pressure but led to a 10% decrease in diameter with no changes in medial thickness. More recent experiments by Miyashiro et al. (1997) demonstrated that the decrease was less in mature than juvenile animals in response to a greater reduction of flow (by 93%). They concluded, not surprisingly, that remodeling in juvenile animals is more extensive than in adults, an observation made in rabbits by Langille et al. (1989). The latter authors showed that after ligation for one month in young animals, the 31 % smaller arterial diameter was due to decreased wall mass with loss of smooth muscle cells and elastin, but not collagen, and a reduced number of endothelial cells per cross section. By contrast, in adults the 21% decrease in diameter was not associated with any change in number of endothelial cells per cross section. The incorporation of BrdU into endothelial and smooth muscle cells was diminished in young rabbits after one week of reduced flow and cell death also contributed to remodeling (Cho et al., 1997). Constriction of the abdominal aorta increased the velocity of flow upstream where elongation of endothelial cells was observed 24 hours later and a 100-fold increase in endothelial cell turnover occurred after 30 days. Endothelial cells downstream, which were exposed to small irregular vortices of flow, became rounded and started to proliferate earlier but less extensively than those upstream (Langille et al., 1986). Increased shear stress upstream caused redistribution of f-actin filaments between central and peripheral parts of endothelial cells between 24–48 hours, with very slow regression after removal or constriction (Langille et al., 1991). The vascular remodeling response to decreased flow/pressure has been extensively investigated as development of collateral circulation after partial or total vessel occlusion. Many studies have discussed contributory factors, focusing especially on heart and skeletal muscle (see Schaper, 1971, 1995; Hudlicka and Tyler, 1986; Hudlicka et al., 1992; Hudlicka and Brown, 1994) but here, we will concentrate on the differences between types of remodeling, that is enlargement of existing vessels and/or growth of new vessels, and on which hemodynamic and mechanical factors could be involved. Coronary vascular remodeling The contrast between growth of new vessels and remodeling of preexisting ones is exemplified by a comparison of collateralization in dog and pig hearts in response to gradual constriction of the main coronary artery. In dogs, which have a preexisting network of anastomoses (about 40 µm in diameter) in the epicardial region, constriction caused their enlargement by proliferation of endothelium and smooth muscle cells. In pigs, where there are no preexistent anastomoses, growth was apparent in the form of vessels slightly larger than capillaries yet devoid of smooth
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muscle cells—presumably venules—arising in the vicinity of small regions of necrosis. These might have been stimulated by growth factors released from macrophages but not from ischemic myocardium. Arterial constriction would be likely to increase shear stress and activate endothelium throughout the remaining patent vasculature. At the same time, small arterioles, dilated possibly by hypoxia, became thin-walled due to rupture of the internal elastic lamina, or even apoptosis of some SMCs, while concomitant proliferation of SMCs eventually resulted in a much larger vessel with a new internal elastic lamina which was orientated longitudinally, thus causing the typical tortuosity of collateral vessels. Overproduction of fibronectin closes some vessels so that only about 20% of vessels remain functional (Schaper, 1991, 1995). Remodeling rather than growth of new vessels was also described in human hearts (Dimario et al., 1994) in which coronary vascular anatomy is much more similar to that of the pig than dog heart (Schaper, 1971). In dog hearts made ischaemic by multiple embolization, however, both processes occurred with growth of smaller vessels occurring within 1–3 weeks and development of epicardial collaterals at a later stage (Chilian et al., 1990) and proliferation of capillaries was also described in pig hearts under similar conditions (Zimmermann et al., 1997). Two to five days after partial constriction of the coronary artery, there was an increase in arteriolar density and some capillary sprouting (White et al., 1990). Whether the stimulus for this was ischemia or changed hemodynamics is not clear. Hearts with limited blood supply usually show a deficit in capillaries (Anversa et al., 1986) but this is mainly because the remaining viable tissue hypertrophies. Recent reports (Shammas et al., 1993; Xie et al., 1997) described angiogenesis in tissue close to the infarct attributable to the presence of either thrombin, a potent angiogenic factor, or to other growth factors rather than to hemodynamic changes. It is extremely difficult, particularly in the heart, to measure in vivo the relevant parameters such as velocity of flow and vessel diameters from which the involvement of hemodynamic factors could be calculated. Tillmans et al. (1974, 1981, 1982) measured diameters and pressures in epicardial arterioles and venules, and red blood cell velocities in capillaries, in normal hearts from different species but not during development of collateral circulation. Schaper (1971) considered tangential shear stress [t=PR/D where D is thickness of the wall, P pressure and R radius] an important factor in the development of collaterals in the heart, assuming that hypoxia rather than pressure gradients was the main cause of vessel dilation. The dilator role of hypoxia was also confirmed in studies showing better collateral development in patients experiencing angina (Schaper et al., 1981) or after ischemic preconditioning, i.e. repeated short duration occlusions of coronary arteries (Sasayama and Fujita, 1992; Tomoike, 1993). Recently Kern et al. (1993) showed that in patients with a partial or fully occluded coronary artery, velocity of flow measured distal to the occlusion in tissue supplied by collaterals was approximately 30% of that above the occlusion, but there are no comparable data on the diameters of these vessels from which the relative change in the shear stress could be calculated. Kass et al. (1992) reported a higher presence of anastomoses in patients whose coronary flow was high due to iron deficiency anaemia and concluded that increased flow is an important factors in remodeling, while ischemia and possible release of various growth factors are more important for growth of new vessels.
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Remodeling of skeletal muscle vascular bed In skeletal muscles the development of collateral vessels is related to the extent to which any existing collateralization can support flow after occlusion, and the speed with which either these and/or new vessels develop. Involvement of preexisting collaterals, identified by arteriograms, was shown in the cat and dog limbs after aortic and femoral occlusion respectively (Schaub et al., 1976; Conrad et al., 1971) and by autoradiographic studies in rabbits and rats after femoral or popliteal artery ligation (Jaya, 1980). Proliferation of new vessels was demonstrated by increased incorporation of 3H-thymidine in larger vessels and capillaries in dog calf muscles after surgical popliteal A-V anastomosis (Sewell and Roth, 1958) and by direct observation of the numbers of small arterioles in rat cremaster muscle 3 weeks after ligation of the main feeding vessel (Hogan and Hirschman, 1984). The development of collateral circulation seems to be better the further away from the investigated site the obstruction occurs. Ligation of the abdominal aorta caused very little damage in muscles of the lower hind limb (Hanzlikova and Gutmann, 1979) whereas common iliac artery ligation in rats decreased blood flow in the same region to about 30% of normal values at rest (Hudlicka and Price, 1990a) and the capacity for functional hyperemia was lost (Hughes and Hudlicka, 1992). Vascular casts showed incomplete vascularization even 5 weeks after iliac ligation (Hudlicka and Torres, 1990) with perfusion pressure in the femoral artery only 60% of systemic pressure (Hudlicka et al., 1994). Flow in the femoral artery was 60% lower than in the contralateral artery as long as 9 weeks after common iliac ligation (Rochester et al., 1994) and flow in contracting tibialis anterior, a mixed fast muscle, only returned to normal values after 12–18 weeks (Janda et al., 1974). The origin of collateral circulation after ligation of the common iliac artery could be traced from the contralateral iliac artery (Dawson and Hudlicka, 1990b). Ligation of the femoral artery resulted in the development of collateral circulation from preexisting vessels (Jaya, 1980), originating in the internal iliac and deep femoral artery (Ward and Angus, 1995). This progressed for up to 3 months with resting flow returning to control values within 3 weeks (Paskins-Hurlburt and Hollenberg, 1992) although flow during contractions was still much lower (Okyayuz-Baklouti, 1989). Vessels below a ligation were of smaller radius when visualized by casts but were more numerous indicating growth rather than remodeling (Conrad et al., 1971). This was recently confirmed by demonstration of proliferation in endothelial and SMCs below the site of ligation within 7 days (Ito et al., 1997a). On the other hand, conductance in collateral vessels was increased 6-fold (Ito et al., 1997b) indicating a degree of expansion in excess of that likely from growth alone, hence some remodeling must also occur. Ligation of the anterior tibial artery, which leaves little scope for the development of collateral circulation, produced necrosis of the distal third of the tibial muscle, although it was completely restored within 90 days by regeneration (Kaspar et al., 1969). There is comparatively little data available on hemodynamic changes in the microcirculation of muscles with decreased blood flow. Arteriolar diameters under resting conditions were unchanged over several weeks after iliac artery ligation (Dawson et al., 1990; Hughes and Hudlicka, 1993) but their ability to dilate during
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muscle contractions (Dawson et al., 1990; Hudlicka et al., 1994) or in response to vasodilators (Shiner et al., 1995) was considerably impaired. There was no change in diameter of venules (Hughes and Hudlicka, 1993) and no difference in the number of perfused capillaries although their intermittency of flow was greatly increased (Anderson et al., 1997). Maintenance of perfusion, albeit temporarily limited, obviously contributes to the preservation of the integrity of the capillary bed. There was no loss of capillaries in this model up to two (Hudlicka et al., 1994) and five weeks (Hudlicka and Price, 1990b) (Figure 14.7). Their diameters, however, were smaller, the volume of blood in individual capillaries was diminished by about 20%, and, with a slightly higher flow velocity (Dawson and Hudlicka, 1990b), shear stress would be increased as a stimulus to prevent the disappearance of capillaries. Maintenance of the integrity of capillary supply in patients with peripheral vascular diseases was reported by Hamarsten et al. (1980), Henriksson et al. (1980) and Esbjornson et al. (1993). Extra-vascular mechanical factors may also be important for maintenance of a normal vascular bed. For example, when tension in the cremaster skeletal muscle was reduced by orchidectomy, flow decreased as did the diameter of all arterioles and the number of terminal arterioles (Wang and Prewitt, 1993). Whether hemodynamic factors can explain such extensive vascular remodeling or growth in skeletal muscle vasculature in situations of decreased blood flow is a matter of debate. It has been suggested that decreased pressure below the site of an obstruction would result in dilation and increased wall tension (John and Warren,
Figure 14.7. Left—capillary/fiber ratio (C/F) in rat lower hind limb muscles (EDL=extensor digitorum longus, TA=tibialis anterior, glycolytic cortex and oxidative core). Right—percentage changes in C/F from values in control muscles after ligation of the common iliac artery for 7, 14 and 35 days. Despite low flow, there was no significant deficit or increment in C/F in the ischemic muscles.
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1961; Liebow, 1963) and that pressure differentials across the site of occlusion contribute to expansion of the arterial tree and can possibly stimulate growth by increasing wall tension (Ito et al., 1997b). Factors which modify remodeling under conditions of decreased flow The question of whether development of collateral circulation be enhanced by interventions such as exercise training or drugs is unresolved (see Hudlicka et al., 1992; Tomanek, 1994, for reviews). Training studies are equivocal as to improvements or not in collateral development. Although recent data show that only very vigorous training can protect against coronary heart disease (Morris, 1994), mortality in patients with non-fatal myocardial infarction is lower possibly because exercise slows the progression of arteriosclerosis and leads to adaptations in skeletal muscles which reduce demands on cardiac output rather than improving blood supply to the heart (McKirnan and Bloor, 1994; Libonati et al., 1997). Heaton et al. (1978) explained improved flow close to the infarcted zone in trained dogs as a consequence of training-induced bradycardia. Certain drugs which lower heart rate indeed increase flow in collaterals when administered acutely (Daemmgen et al., 1985; Gross and Daemmgen, 1987) but as they have not been utilized on a chronic basis it is impossible to say whether long-term bradycardia (which certainly has many mechanical effects of the coronary vascular bed, see below) stimulates growth of collaterals. Although improved muscle fatigue resistance, particularly in the lower hind limb, was attributed to better collateral circulation after exercise training (Yang et al., 1995), most studies do not support the concept that it improves collateral development (Sanne and Silverston, 1968; Mathien and Terjung, 1986, 1990, 1992; Roberts et al., 1997). On the other hand, a more specific increase in muscle activity induced by chronic electrical stimulation applied for 10–15 minutes seven times per day for two weeks not only improved blood flow but also stimulated growth of capillaries in ischemic muscles. It also increased perfusion pressure below the site of ligation indicating inflow from either preexisting or newly-grown collaterals (Hudlicka et al., 1994). Formation of collaterals in ischaemic muscles, as demonstrated by casts, was also enhanced by administration of the drug torbafylline (Hudlicka and Torres, 1990), which increased capillary perfusion and restored the capability of arterioles to dilate during muscle contractions (Dawson et al., 1990).
REMODELING UNDER PHYSIOLOGICAL CIRCUMSTANCES—INCREASED ACTIVITY AND CHANGES IN HEART RATE Capillary supply is considerably higher in athletic animals than in sedentary animals of related species. This has been shown in skeletal muscle by comparison of the hare and the rabbit (Wachtlova and Parizkova, 1972), or greyhound and thoroughbred horse with other dog and horse species (Gunn, 1981) and also in the heart, where it is linked with a lower heart rate (Wachtlova et al., 1965, 1967). Greater physical
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activity is therefore linked with a higher capillary supply, and remodeling and/or growth of the vascular bed in these tissues as a result of increased activity through exercise training has been studied mainly in connection with capillary growth. There are also some data on the expansion of the whole vascular bed, measured as its total capacity under conditions of maximal dilatation. More recently, the growth of arterioles relative to that of capillaries has been investigated to assess to what extent coordination of expansion throughout the vascular bed takes place. It is remarkable that in skeletal muscles endurance exercise training results in capillary growth which is not always accompanied by growth of arterioles, whereas the reverse seems to hold for the heart.
Skeletal Muscle Vascular Bed—Responses to Exercise Training In skeletal muscle, the extent of capillary growth varies with the length and type of training (see Hudlicka et al., 1992, for review). The importance of hemodynamic factors in training-induced capillary growth depends on which muscle fibers are recruited and thus supplied preferentially with blood according to the type of training undertaken. In man and animals, growth of capillaries usually takes weeks or months (Andersen and Henrikson, 1977; Ingjer, 1979; Saltin and Gollnick, 1983; Hudlicka, 1990), appearing in the most active muscles (Hudlicka, 1990). These will obviously have the highest blood flow, but distribution of blood flow is different even within one muscle depending which muscle fibers are recruited by training. Slow muscle fibers are almost permanently active, and slow muscles such as the postural soleus have a higher blood flow than mixed muscles (Hudlicka, 1975). They also have much higher capillary supply than fast muscles (Romanul, 1965; Gray and Renkin, 1978). Fast oxidative fibers are recruited first in endurance training, while fast glycolytic muscles are recruited only during training for speed. The distribution of blood flow within muscles composed of different fiber types both during acute exercise and in endurance-trained animals was increased specifically in the parts of muscles preferentially recruited in the particular movement (Armstrong and Laughlin, 1983, 1984). The finding that capillarity is increased around fast oxidative fibers in endurance-trained animals (Mai et al., 1970) or people (Ingjer, 1979) is consistent with a role for increased perfusion as a stimulus. Increased capillarization was also reported in muscles composed of fast glycolytic fibers in sprint-trained animals (Gute et al., 1994), related to the changes in blood flow in this type of training (Laughlin et al., 1987). Although there is no direct evidence for capillary proliferation using the labelling index of endothelial cells for 3H-thymidine, PCNA or BrdU in training studies, those mentioned here used either electron microscopy or staining specific for capillary endothelium to show all capillaries present in muscles, so that growth is clearly shown by an increased capillary density or C/F ratio. Despite plentiful studies on capillary growth, data on changes in larger vessels are much scarcer. Several studies have shown enhanced maximal vasodilator capacity in trained human muscles (Snell et al., 1987; Martin et al., 1987; Sinoway et al., 1986), indicating potential growth of all vessel classes and definitely of the resistance vessels. It is, however, impossible to ascertain whether numerical vessel density or remodeling
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leading to increased diameter of individual vessels had occurred. In conduit arteries in endurance trained rats, medial thickness was increased with no change in lumen diameter (Segal et al., 1993). Koller et al. (1995a) found that arterioles from muscles of endurance-trained rats had larger diameters, but there was no difference in the maximal capacity to dilate suggesting an increase in total vascular conductance would be more likely due to increased numbers of arterioles. Increased arteriolar density and shorter intervals between individual branches were described in spinotrapezius muscle after training (Lash and Bohlen, 1992) with only a moderate growth of capillaries and a decrease in minimum vascular resistance (Sexton and Laughlin, 1994) (Table 14.2). Coronary Vascular Bed—Responses to Exercise Training The effect of training on growth of the coronary vascular bed has recently been thoroughly reviewed (Laughlin and McAllister, 1992; Hudlicka et al., 1992; Laughlin, 1994; Tomanek, 1994; Hudlicka and Brown, 1996). On the basis of the higher myocardial capillarization observed in athletic animals, referred to above, exercise training could be expected to promote capillary growth. Capillary proliferation has been demonstrated by incorporation of 3H-thymidine into nuclei of capillary endothelial cells in the trained heart using electron microscopy with autoradiography (Mandache et al., 1973; Unge et al., 1979; Ljungqvist et al., 1984) but most studies show that increases in capillary supply are very limited (Figure 14.8) and happen almost exclusively in younger animals (Tomanek, 1970; Anversa et al., 1983; Mattfeldt et al., 1986). Endurance training does, however, stimulate growth of arterioles and enlargement of larger coronary arteries by either remodeling or growth (reviewed by Laughlin and McAllister, 1994). The size of the whole coronary vascular bed, shown by casts in rats, is increased (Tepperman and Pearlman, 1961; Ho et al., 1983), but only by relatively moderate exercise (Stevenson et al., 1964; Haslam and Stull, 1974). An increase in the number of arteriolar profiles, mainly small (20 µm) vessels, was reported in rats (Rakusan and Wicker, 1990) and pigs (Breisch et al., 1986; White et al., 1987). Table 14.2 Capillary and arteriolar supply in rat spinotrapezius muscle (Lash and Bohlen, 1992) and maximum blood flow and conductance, measured during exercise in conscious rats (Armstrong and Laughlin, 1984) and minimum vascular resistance, measured in maximally dilated rat hind limb muscles (Sexton and Laughlin, 1994) after endurance training. *P<0.05 trained v. control sedentary
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Dimensions of large coronary arteries
Figure 14.8. The effects of endurance exercise training on different sections of the coronary vascular bed. Size of large coronary arteries and arteriolar density increase, but capillary density does not. Based on data for capillary density and size of the coronary arteries from Wyatt and Mitchell, 1978, and for density of arterioles from Breisch et al., 1986. *P<0.05 trained v. untrained.
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In the latter species, smaller arterioles (21–40 µm) density increased after 16 weeks of training while larger arterioles had increased diameters but not numbers. At this time, capillary density was not changed (Breisch et al., 1986; White et al., 1998) but it was transiently increased after 1–3 weeks of training (White et al., 1998); thus the new arterioles could arise by transformation from capillaries, a process described in developing skeletal muscles (Price et al., 1994). The size of the vascular bed (number of vessels times cross sectional area) increased by 50% with only 11% of this accounted for by greater numbers of vessels and White and Bloor (1995) concluded that the expansion of the vascular bed resulting from training is accomplished to a very small extent by growth and much more by remodeling of the existing vessels by dilation. This has been shown in large coronary arteries whose diameter was increased with training (Wyatt and Mitchell, 1978; Tharp and Wagner, 1982) (Figure 14.8). Maximal coronary conductance (or minimal resistance) which is an indicator of the size of the whole coronary vascular bed has also been reported to be increased in rats, dogs and pigs (see Laughlin and McAllister, 1992; Tomanek, 1994). The relationship between hemodynamic changes in the coronary circulation upon exercise training and the differential remodeling/growth of larger vessels yet lack of capillary growth are difficult to assess. Although capillary density is often reported to be unchanged, this does not necessarily point to a complete absence of growth. The fact that the density is already very high in the heart may preclude further expansion of this section of the vascular bed but in the face of training-induced mild heart hypertrophy and larger fibers, the maintenance of capillary density implies occurrence of a certain degree of growth. There is no doubt that blood flow is increased in the heart during exercise and possible mechanical factors affecting capillaries could arise from changes in microcirculation such as velocity of flow and capillary pressure and wall tension. In addition, the extramural pressure exerted by contracting myocytes during exercise training-induced changes in chronotropic (decreased heart rate) and inotropic (increased force of contraction) state of the heart could act on the abluminal side of capillaries by disturbing the extracellular matrix and possibly the basement membrane. Vascular Remodeling in Skeletal Muscle—Responses to Chronic Electrical Stimulation From training studies, there is, as yet, no direct evidence based either on electron microscopy or intravital observation which would confirm the individual steps in angiogenesis—disturbance of the basement membrane, migration and proliferation of endothelial cells and formation of sprouts in heart or skeletal muscle. The complexities of exercise-induced hemodynamic changes in the microcirculation in vivo also make it difficult to study individual parameters as stimuli for vascular growth/ remodeling. In order to analyse factors in detail separately, we have developed methods whereby capillary growth can be achieved relatively rapidly by increasing activity by chronic electrical stimulation in skeletal muscle and by long-term bradycardial pacing in the heart (see below). Increasing activity of skeletal muscles by electrical stimulation to promote capillary growth and improve performance is becoming increasingly important in
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connection with cardiomyoplasty. In this procedure, skeletal muscles are ‘conditioned’ to make them less fatigable and therefore suitable as auxiliary hearts in cases of heart failure. The only cure for severe heart failure is heart transplantation, and in view of the limited number of donors, ‘wrapping’ a non-fatiguable skeletal muscle around the myocardium to help cardiac function is increasingly of interest. Alternatively, skeletal muscles are wrapped around the aorta to create ‘secondary’ ventricle (Greer et al., 1996). Original work exploring the effect of chronic electrical stimulation on capillary growth in skeletal muscle was based on the findings of Adrian and Bronk (1929) who recorded electrical activity from nerves to slow and fast muscles. They established that while slow postural muscles were continuously activated by low-frequency (10Hz), activity in nerves to fast muscles was in short bursts of 60–100Hz. This finding was utilized by Salmons and Vrbova (1969) to transform fast into slow muscles by stimulating the supplying nerves continuously at 10Hz. Both this and lower (5Hz) activity, applied for 8 hours per day, were subsequently shown to increase C:F ratio by 20% within 4 days, with a 50% increase in muscles stimulated for 14 days and doubling of the capillary supply within 28 days (Brown et al., 1976) (Figure 14.9). Capillary growth started within 2 days (Hudlicka et al., 1982) in the vicinity of the fast glycolytic fibers which are activated first during stimulation (the opposite of recruitment in voluntary exercise). Capillary proliferation was shown by an increased labelling index in capillary-linked nuclei at this time (Pearce et al., 1995) and in muscles stimulated for longer periods of time (Joplin et al., 1987). Growth was preceded by increased capillary perfusion (Hudlicka et al., 1984) indicating the role of flow. Since the velocity of flow in capillaries in contracting muscles increases more than their diameter (see Figure 14.13), the shear stress is higher which could represent a stimulus for capillary growth. Stimulated muscles have not only increased flow but the capillaries are also exposed to continuous distortion due to repetitive changes in sarcomere length. Similar length changes can be elicited by muscle overload (stretch) induced by extirpation of the agonist muscle e.g. extirpation of tibialis anterior causes overload of extensor digitorum longus, removal of gastrocnemius and soleus causes overload of plantaris. Although blood flow in these overloaded muscles is not increased (Egginton and Hudlicka, 1992) there are more numerous capillaries (James, 1981; Frischknecht and Vrbova, 1992; Degens et al., 1992; Egginton and Hudlicka, 1992). The greater sarcomere length (Egginton et al., 1998) may cause disruption of the extracellular matrix and hence capillary growth, verified by abluminal sprouting and presence of mitosis (Zhou et al., 1998b). The appearance of growth, however, was much later (after 2 weeks) than in stimulated muscles suggesting that a combination of increased blood flow and changes in muscle length in the latter are a more potent angiogenic stimulus. There was also ultrastructural evidence in stimulated muscles for growth by both sprouting and luminal division (HansenSmith et al., 1996a; Zhou et al., 1997a) producing an incredibly complicated network pattern (Figure 14.10). Increasing skeletal muscle activity by chronic electrical stimulation increases the number of arterioles (Figure 14.9). This occurs possibly by arteriolarization of capillaries because there is a greater increase in vessels of smaller diameters (Hansen-
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Figure 14.9. Changes in the vascular bed in rat hind limb muscles chronically stimulated for different durations up to 7 days. Data for capillary per fiber (C:F) ratio from Brown et al., 1995; for arteriolar density from Hansen-Smith et al., 1998 and for maximal conductance form Hudlicka and Egginton, 1994. Full line=terminal (=10 µm), interrupted line=preterminal (>10 µm) arterioles.
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Figure 14.10. Capillary network in rat extensor hallucis proprius muscle stained with Griffonia simplicifolia I lectin and viewed by confocal microscopy. Top—control muscles; bottom— muscle stimulated for 7 days. Modified from Hansen-Smith et al., 1996a. Bar=100 µm.
Smith et al., 1998) or by dilation of vessels of larger calibres (Adair et al., 1995). Increased density of veins and arteries after a very brief electrical stimulation (5 minutes daily for 3–4 days) was described long ago by Vanotti and Pfister (1933). In their experiments, it might be that vessels were dilated since they were only visualized by inspection of the muscle surface, but the authors also described more frequent anastomoses and more vigorous pulse pressure.
Coronary Vascular Bed—Responses to Long-term Bradycardia It is very well known that endurance training results in a decrease in heart rate, although the mechanisms—increased vagal activity or intrinsic changes in the sinoatrial node—are not clear (Badeer, 1975). Blood flow in coronary vessels is higher during diastole than during systole (Hoffman and Buckberg, 1976) and decreased heart rate would therefore be expected to enhance coronary vascular bed perfusion which could stimulate capillary growth similar as do coronary vasodilators. Moreover, the ‘domain’ area of myocardium supplied by any individual capillary is greater in systole than in diastole (Batra and Rakusan, 1992) so that the
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faster beating myocardium with shorter duration of diastole would be relatively hypoxic in addition to lower perfusion. All these considerations have prompted experiments with bradycardial drugs in an attempt to improve coronary perfusion and possibly function. Indolfi et al. (1989) found that after brief coronary artery occlusion, the bradycardic drug ULFS49 increased flow in the subendocardium, which is usually more vulnerable during ischemia, and improved left ventricular function of the ischaemic region (Indolfi et al., 1991). A similar effect was achieved with the ß agonist atenolol which also reduced heart rate (Guth et al., 1991). Whether these particular drugs would promote vessel growth if given on a long-term basis has not been investigated. Chronic treatment with another ß agonist propranolol, which reduced heart rate, increased the volume density of myocardial capillaries in young rabbits (Tasgal and Vaughan Williams, 1981) and volume fraction of arterioles (Acad et al., 1988). Capillary growth was also achieved in rats by the bradycardic agent alinidine in both normal and hypertrophic (renal hypertension) hearts (Brown et al., 1990). The effect of decreased heart rate on the growth of coronary vessels and the mechanism of this growth was investigated in experiments where bradycardia was induced by either chronic electrical pacing eliminating every second heart beat in rabbits and reducing cardiac frequency by about 55% (Wright and Hudlicka, 1981) or by dual chamber telemetric pacemakers in pigs (Brown et al., 1994a) lowering heart rate by about 35% (Figure 14.11). The resultant increase in capillary density was related to the decrease in heart rate in all three species (Brown et al., 1994b; Figure 14.11). Heart rate reduction also elicited capillary growth in hypertrophic hearts in rabbits (Wright et al., 1989) and in infarcted hearts in pigs (Brown et al., 1995). Although coronary blood flow per beat was increased (Hudlicka et al., 1989) neither resting nor maximal flow or conductance was altered (Hudlicka and Brown, 1994; Brown et al., 1994a). From measured heart rate in these experiments and published data on capillary velocity of flow and diameters during the cardiac cycle (Tillmans et al., 1974), shear stress was calculated to be unchanged (Hudlicka, 1994) but it is possible that capillary pressure was higher. Since capillary diameters are larger in diastole than in systole (Tillmans et al., 1974), capillary tension was estimated to be higher as well (Hudlicka et al., 1995). In bradycardially paced hearts, an increase in stroke work and thus force of contraction occurred before increased capillarization (Hudlicka and Brown, 1993). Another possibility is therefore that the increased force of contraction exerted by the heart during systole and the stretch of myocytes in the prolonged diastole could initiate capillary growth by modifying the extracellular matrix. In order to ascertain the role of these inotropic factors, rabbits were treated for two weeks with a ß agonist dobutamine which increased stroke work without altering either blood pressure or heart rate. Coronary blood flow was also not significantly different from controls yet capillary density increased to a similar extent as in paced hearts (Brown and Hudlicka, 1991). Bradycardia therefore stimulates growth of capillaries but not larger vessels—maximal coronary conductance was not changed—by a combination of changes in microcirculation and mechanical forces acting on the vessels from abluminal side.
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Figure 14.11. Top left—diagram to show electrode placement in pigs hearts for electrical pacing to produce bradycardia. Top right—electrocardiogram to show heart rate before (A) and after (B) 4 weeks of pacing. Bottom—relative decrease in heart rate (as % of control values) achieved in rabbits (ref. Wright and Hudlicka, 1981) and pigs (ref. Brown et al., 1994a) by electrical pacing and in rats by the bradycardic drug alinidine (ref. Brown et al., 1990) against the increase in myocardial capillary density (as % of control values).
REMODELING DURING PATHOLOGICAL CONDITIONS IN SKELETAL AND CARDIAC MUSCLE Hypertrophy and Atrophy of Skeletal Muscles Most data on capillary supply in muscle hypertrophy relates to functional over-load in mammals and birds, or cold-acclimation and training in fishes, and shows that C:F
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increases approximately linearly with fiber area. Stretch is an important factor involved in capillary growth in skeletal muscle hypertrophied by exposure to functional overload. Increased capillarization was seen in avian muscles over-loaded by increased weight on the wings (Holly et al., 1980; Snyder and Coelho, 1989) and in mammalian skeletal muscles stretched by overload due to agonist removal (Frischknecht and Vrbova, 1992). In the latter muscles, blood flow was not increased while sarcomere length was (Egginton et al., 1998). Acute muscle stretch decreased capillary tortuosity in indirect proportion to sarcomere length (Ledvina and Segal, 1995), and capillary diameters and Vrbc were decreased in muscles with longer sarcomeres (Poole et al., 1997). Muscle stretch also caused constriction of arterioles and feed arteries as a result of increased activity of the sympathetic nerve fibers (Welsh and Segal, 1996). The stimulus for capillary growth during hypertrophy by stretch-induced overload is therefore unlikely to be related to hemodynamic factors associated with blood flow. That it is largely mechanical in origin arising from tension applied by the expanding fibers acting via the abluminal surface of capillaries is indicated by the fact that growth is by sprouting, with electron microscopic evidence of disturbance to the basement membrane rather than the luminal endothelial surface (Zhou et al., 1998b). Capillary proliferation following compensatory overload in rat plantaris was not enhanced by the additional stimulus of endurance training (Degens et al., 1992), which would provide hemodynamic stimuli associated with increased flow. In the essentially high resistance exercise accompanying fish swimming, which causes a 20fold increased in muscle blood flow (Wilson and Egginton, 1994), trout slow muscle fiber area was nearly doubled yet the increase in C:F was relatively modest, 10% (Johnston and Moon, 1980). Responses to hemodynamic factors may therefore be attenuated or even superseded by other stimuli. When muscle fiber size is reduced by atrophy, a similar trend is evident in the relationship between C:F and fiber area, albeit with less of a decline in C:F relative to fiber size. Changes in the vascular bed appear to proceed less quickly than in muscle fibers themselves. The maintenance or even increase in capillary supply after tenotomy (Josza et al., 1988) or denervation (Hassler and Stroinska-Kusinova, 1976) may be because blood flow in muscles atrophied by these procedures is relatively high (Hudlicka and Renkin, 1968; Hudlicka, 1967; Eisenberg and Hood, 1994), possibly due to the release of vasodilating metabolites from degenerating muscle fibers. Similarly, preservation of capillary supply following unloaded atrophy as a result of unilateral limb suspension in man (Hather et al., 1992) or rats following space flight (Roy et al., 1996) is presumably due to maintenance of normal blood flow. Other signals may play a permissive role as capillary sprouting following crush injury occurred more rapidly and to a greater extent in mobilised than immobilised limbs (Jarvinen, 1976). Increased capillarity was also noted in dystrophies of mice (Atherton et al., 1982) and human (Carry et al., 1986), as well as over-wintering atrophy in rats (Heroux and St. Pierre, 1957), but the cause is unknown. In more generalised pathologies and other dystrophies (e.g. Duchenne; Carry et al., 1986), diabetes (Sexton et al., 1994), and even ischemia (Yamaguchi et al., 1994) substantial reductions in capillary supply parallel often severe reductions in fiber size. As with hypertrophy, the few data available suggest a coordinated remodeling of the
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microvasculature, e.g paraplegia-induced disuse atrophy is accompanied by increased venous resistance (Hopman et al., 1994). Hypertrophy of Cardiac Muscle Recent reviews of coronary vascular remodeling in hypertrophic hearts (Tomanek, 1990; Tomanek and Torry, 1994; Hudlicka and Brown, 1994, 1996) summarize the fact that growth and remodeling are variable, depending upon factors such as age, model and duration of cardiac hypertrophy. Hypertrophy resulting from pressure overload due to spontaneous or renal hypertension is usually associated with decreased capillary density (see Rakusan, 1987), increase in minimal coronary resistance, and hence decrease in coronary reserve, indicating insufficient growth of larger resistance vessels. Long-term inhibition of NO synthase also results in hypertension, cardiac hypertrophy and capillary rarefaction, typical of the response to pressure overload (Sladek et al., 1996). If, however, pressure overload is initiated in the young, there is proportional growth of capillaries and larger vessels in relation to cardiac size. This has been shown after aortic constriction in rabbits (Rakusan et al., 1967), sheep (Flanagan et al., 1991) and humans (Rakusan et al, 1992). Capillary growth was also shown in young SHR by an increased labelling index for 3Hthymidine incorporation (Tomanek et al., 1982) or morphometric capillary supply (Anversa et al., 1984). The duration of pressure overload is also a factor in vascular remodeling. Tomanek and colleagues found in dogs that renal hypertension decreased capillary supply and minimal coronary resistance and increased arterial medial thickness after 6 weeks (Tomanek et al., 1986) but after 7 months, arterial densities, minimal coronary resistance (Tomanek et al., 1989) and capillary density (Tomanek et al., 1991) were virtually normal, indicating growth. White et al. (1992) also demonstrated 3Hthymidine incorporation in arterial and capillary vessels indicating proliferative capacity in pressure-overloaded pig hearts, and in the same model, decreased resistance was attributed to an increase in the numbers of arterial vessels, even though their diameter had decreased (Kassab et al., 1993). Pressure overload not only leads to enlargement of the cardiac mass but to increased perfusion pressure in coronary vessels, which may be responsible for the thickening of the medial wall. Once hypertrophy is established, all capillaries are perfused at rest (Henquell et al., 1977; Marsicano et al., 1977) and there would be no reserve for recruitment during increased cardiac work. Bishop et al. (1996) found left ventricular arteriolar density to be the same as control in dogs one year after aortic banding when puppies, yet coronary reserve was lower. As the dilator capacity of resistance vessels is compromised, possibly by fibrosis (Strauer, 1992), increased intercapillary distances (Kayar and Weiss, 1992) together with absence of a capillary reserve produce hypoxia, particularly in subendocardial regions. Whether hypoxia induces expression of growth factors such as VEGF under these conditions is not known, but it is unlikely that hemodynamic factors associated with increased flow are responsible for any growth. The situation for hypertrophy by volume overload is different. Aorto-caval shunts produce hypertrophy of both ventricles, and while previous studies reported a decreased
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myocardial capillary density (Rakusan et al., 1980; Thomas et al., 1984; Camilleri et al., 1984), more recent work showed that capillary growth paralleled the magnitude of hypertrophy (Batra et al., 1991; Chen et al., 1994) and that the length density and diameters of arterioles were the same as controls (Chen et al., 1994). Volume overload elicited by anemia from iron deficiency (Poupa et al., 1964) or iron and copper deficiency (Olivetti et al., 1989) led to increased capillary density and 60% greater total length of capillaries respectively, but when produced by aortic valve lesion, capillary density decreased (Wright et al., 1989). When hypertrophy was induced by thyroxine treatment, capillary supply was significantly increased (Chilian et al., 1985) and maximal perfusion normal even in old animals (Tomanek et al., 1995). Some of these differences can be accounted for by e.g. that fact that in anemia, hypertrophy is due to hyperplasia rather than hypertrophy of individual fibers (Poupa et al., 1964) so that capillary growth occurred concomitant with that of new fibers. In contrast to pressure overload, increased blood flow could explain the increased capillarization in animals exposed to thyroxine or anemia when cardiac work and hence coronary flow would also be increased. Also the stimulus of stretch of myocytes due to increased stroke volume and concomitant increase in force of myocyte contraction must be considered. When end diastolic ventricular dimensions are increased, longitudinal stretch of vessels would occur, and in combination with high blood flow, would be a potent stimulus for vascular growth in several of the models of volume overload. The capacity for vascular growth may also be modified by alterations in myocardial structure, in particular the extracellular matrix, which responds independently to hypertension and hypertrophy (Brilla et al., 1990). For example, interstitial collagen content is increased in cardiac hypertrophy due to pressure overload (Weber et al., 1992) but not during volume or thyroxine-induced hypertrophy (Michel et al., 1986). Hypertrophic interstitial fibrosis would be obstructive to capillary growth, as demonstrated by the fact that its regression in the hypertrophic heart following captopril treatment (Rossi and Peres, 1992) is accompanied by an increase in capillary density (Canby and Tomanek, 1989). INDIVIDUAL HEMODYNAMIC AND MECHANICAL FACTORS IN VASCULAR REMODELING The participation of individual hemodynamic and mechanical factors in vascular remodeling differs according to the nature of the stimulus and the vessel categories affected. While it is generally believed that increased pressure results in an increase in wall thickness and diminished lumen (Mulvany, 1992a), flow results in an increased velocity and shear stress and increased size of the lumen (Langille, 1993), by complex activation of ion channels and humoral factors (Gibbons and Dzau, 1994) (Figure 14.12), which eventually returns the shear stress to normal values (Kamiya et al., 1984).
Shear Rate and Shear Stress The pattern of blood flow is important for the orientation of the endothelial cells in large vessels. In the aorta, endothelial cells are aligned with the long axis of the vessel
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Figure 14.12. Schematic representation of hemodynamic and mechanical factors affecting microcirculation in skeletal/cardiac muscle. Modified from Hudlicka, 1998.
in segments where flow is laminar but they are rounded, with no obvious orientation, in places where flow is disturbed or stagnant such as the origin of renal artery branches or the division of the aorta into iliac arteries (Reidey and Langille, 1980). The rounded cells also have a higher labelling index for 3H-thymidine than cells in regions with laminar flow (Fry, 1968). Changes in flow also play a role in the remodeling of the vascular wall after endothelial injury. While low flow resulted in a greater number of smooth muscle cells in the injured intima, high flow prevented intimal thickening and preserved or even enlarged the size of the vessel lumen (Kohler and Jawien, 1992). The effects of flow and wall shear stress are also important in the development of endothelial lesions at branching point in large arteries during the development of arteriosclerosis (Nerem, 1981). Shear rate (8.Vrbc/d, where d is a diameter of the vessel) and shear stress (shear rate · viscosity) are highest in small arterioles and capillaries (Nerem, 1981) and it is therefore likely that changes in either of these would be important factors in their remodeling. Increased flow implies that red blood cell velocity (Vrbc) is also increased, but this may not occur uniformly in different regions of the vascular tree.
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For instance, total flow during electrically evoked skeletal muscle contractions increased several fold due to dilation of arterioles and conduit vessels but there was no change in the velocity of flow in the smaller arterioles (Hester and Duling, 1988; Berg et al., 1997) yet Vrbc in capillaries increased (Dawson et al., 1987). Wall shear rate in arterioles is therefore either smaller or similar to that under resting conditions but it increases in capillaries (Figure 14.13). Since venular diameters in contracting muscles were the same as at rest while Vrbc increased, shear rate was also increased in venules (Hester and Duling, 1988). Whether similar changes occur in skeletal muscle vasculature when blood flow is repeatedly increased by exercise is not known. When blood flow was increased in skeletal muscles by chronic electrical stimulation or by prolonged administration of the vasodilator prazosin, capillary Vrbc under resting conditions was greater in both situations. Capillary diameters were increased only slightly with stimulation and even decreased after prazosin so that shear stress was calculated to be higher (Dawson and Hudlicka, 1993). It was proposed that increased shear rate plays an important role in the stimulation of capillary proliferation in these models, supported by the finding that administration of compound GP1794 which normalised Vrbc in stimulated muscles prevented capillary growth (Dawson and Hudlicka, 1990c). Evidence that increased shear rate or shear stress could damage
Figure 14.13. Velocity of red blood cells (Vrbc) and diameters of arterioles and capillaries in skeletal muscles are shown at rest and either immediately after tetanic contractions or following administration of adenosine. Data on arterioles after contractions taken from Hester et al., 1988; data on adenosine from Sweeney and Sarelius (1989); data on capillaries after contractions are unpublished observations from Dawson and Hudlicka. Calculated shear rate in capillaries increased in contracting muscles but decreased in arterioles both after contractions and dilation with adenosine.
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the capillary endothelial inner surface was seen in muscles stimulated for only two days in which the layer of endothelial glycocalyx, depicted by staining with ruthenium red, was severely disturbed (Brown et al., 1996). This could make capillary endothelial cells more sensitive to proteases present in the blood stream resulting in the disturbance of the capillary basement membrane as the first step in angiogenesis. To date there are no data on the role of shear stress in the growth of arterioles in chronically stimulated muscles (Hansen-Smith et al., 1998) but Skalak and Price (1996) suggested that growth of arterioles after prazosin treatment is due to circumferential wall stress rather than to shear stress. Vasodilation due to dipyridamole also increased significantly the Vrbc in capillaries and venules in the heart (Tillmans et al., 1982), as well as shear stress (Hudlicka et al., 1995), and this could explain capillary growth described in the heart after its long-term administration (Tornling, 1982). Muscle contractions and/or dilatation also increase capillary hematocrit (Damon and Duling, 1987) and hence viscosity. As this factor contributes to shear stress, the latter could be elevated even if Vrbc is not changed and could account for the increased density of microvessels in the hearts, brains and skeletal muscles of animals with polycythemia (Miller and Hale, 1970). When, however, hematocrit was increased experimentally from 45 to 60% by repeated administration of CoCl2 in rats, Vrbc in skeletal muscle capillaries was not changed and as capillary diameters were slightly smaller, there was no increase in shear stress and no capillary growth (Dawson and Hudlicka, 1993). That hematocrit, by modifying shear stress, does not play a significant role capillary growth was also demonstrated in the hearts where anaemia, induced either by iron (Poupa et al., 1964) or iron and copper deficient diets (Olivetti et al., 1989), actually resulted in capillary growth despite the reduced hematocrit. Red blood cells may, nevertheless, play a role in capillary growth by other means. Pulsatory movements of red cells occur in capillaries with changes in blood pressure (Smaje et al., 1980), and it was suggested that such movements observed in the capillaries in healing wounds could continuously irritate the existing endothelium to proliferate (Branemark, 1965). A similar explanation was proposed for the formation of capillary sprouts in chronically stimulated muscles (Myrhage and Hudlicka, 1978). Finally, the role of blood flow, already referred to in the remodeling of the vascular bed in the experiments on tadpole tails and rabbit ears by Clark (1918) and Clark et al. (1931), was recently reinforced by observations of capillaries in the pupillary membrane in rat pups (Meeson et al., 1996). Absence of flow caused apoptosis of 90% of endothelial cells while fewer (30%) underwent apoptosis when flow was oscillatory and only 10% with free flow.
Wall Tension Changes in wall tension can result from alterations of either flow or pressure (T=P · r, where r is the vessel radius). In the heart, pressure is higher during systole and wall tension greater in large vessels, arterioles, capillaries and even venules (Nellis and Liedtke, 1982). In large vessels and arterioles, circumferential wall stress ( =P · r/h)
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is inversely proportional to the thickness of the wall, h (Skalak and Price, 1996). Wall tension and circumferential wall stress would not only decrease along the vascular tree but vary according the status of the vessel. For example, dilation in skeletal muscles caused by papaverine affected arterioles but not venules thereby increasing post-capillary resistance (Fronek and Zweifach, 1975; Ballard et al., 1991); capillary pressure, and wall tension increased in consequence. Isoproterenol decreased resistance in both arterioles and venules with less effect on capillary pressure. The heterogeneous responses of individual vessel segments which determine capillary pressures must therefore be acknowledged when considering the effects of drugs which elicit capillary growth. There is, unfortunately, often little data on how such drugs alter pressures and diameters in pre- and post-capillary vessels except in a few instances. In the case of dipyridamole and dobutamine, which both stimulate coronary capillary growth, it is known that they increase pressure in venules in the heart (Tillmans et al., 1981). Together with the increased inflow from arterioles, this would cause a rise in capillary pressure and hence wall tension, contributing to their angiogenic action. As an alpha1 antagonist, prazosin would have little effect on skeletal muscle venules which have relatively sparse noradrenergic innervation (Marshall, 1982) but it dilates arterioles thereby increasing capillary pressure. The diameters of capillaries in animals treated with prazosin were only marginally smaller than in controls but the velocity of flow was almost doubled, implying that capillary wall tension may be less important as a stimulus for capillary growth in prazosin-treated skeletal muscles than shear stress (Dawson and Hudlicka, 1993). It could, however, be more significant for capillary growth in chronically stimulated muscles in which slightly enlarged capillary diameters (Myrhage and Hudlicka, 1978; Dawson and Hudlicka, 1989) are combined with increases in capillary pressure from 16mmHg at rest to 32mmHg during muscle contractions (Mellander and Bjornberg, 1992). Even after as little as two days of chronic stimulation, arteriolar diameters were larger at rest with no change in venular diameters (Pearce and Hudlicka, 1994), emphasizing that increased capillary wall tension may play an important part as a stimulus for angiogenesis. Capillary wall tension can also be identified as a likely factor in capillary growth in the heart subjected to long-term bradycardia. Capillaries are wider during diastole than during systole (Tillmans et al., 1974) and, based on this data and on pressure measurement by Nellis and Liedtke (1982), calculated capillary wall tension is higher (Hudlicka et al., 1995) although calculated shear stress is not (Hudlicka, 1994). Bradycardia prolongs the duration of diastole more than the duration of systole, and hence the diastolic period during which wall tension is higher is longer, increasing the possibility of disturbance of the basement membrane. Increased wall tension in small venules has also been suggested as an underlying cause of retinopathies of varying origins (Stefanson et al., 1982). Previous mention has been made that increased luminal pressure reduces the luminal diameter and causes thickening of the vessel wall. The increase in pressure precedes wall thickening, leading Price and Skalak (1994) to suggest that it is the resulting increase in circumferential wall stress that can cause transformation of some capillaries into arterioles and this can occur in both chronically stimulated muscles (Hansen-
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Smith et al., 1998) or in animals treated with prazosin (Price and Skalak, 1996). In stimulated muscles, it is also possible that the permanent increase in flow and capillary pressure cause transformation of pericytes into smooth muscle cells (Hansen-Smith et al., 1998). In prazosin treated animals, increased flow could lead to either dedifferentiation of some smooth muscle cells in terminal arterioles and their migration towards adjacent capillaries or proliferation of fibroblasts and their transformation into smooth muscle cells (Skalak and Price, 1996). In hypertension or aging, increased pressure results in thickening of the vessel wall in large arteries and a decreased lumen, but the relationship between the individual components is such that circumferential wall stress changes relatively little (Wolinsky, 1972). Re-structuring in small arteries in the same situations is due not only to hemodynamic factors, but also to the release of factors from endothelium e.g. endothelin which further decrease the vessel lumen by constriction and also cause proliferation of smooth muscle cells (Mulvany and Aalkjoer, 1990). In addition to the increase in the mean pressure, pulsatile pressure is very important in remodeling of arteries during hypertension and in veins. Only drugs which reduced pulse pressure as well as lowering mean blood pressure were found to be effective in reducing the media/lumen ratio in SHR rats (Baumbach, 1991; Christensen, 1991). The wall of the jugular vein exposed to pulsatile pressure developed a much thicker media (Liebow, 1963) and smooth muscle cells isolated from the saphenous vein showed increased incorporation of 3H-thymidine and increased cell numbers when exposed to pulsatile stretch in culture while those from the mammary artery did not (Predel et al., 1992). Pulsatile stretch applied to smooth muscle cells in culture also caused their perpendicular alignment to the direction of stretch, similar to their circumferential alignment in vivo (Kanda and Matsuda, 1993). These observations provide an explanation for the remodeling of the vascular wall which takes place without proliferation. Remodeling of individual components of the vascular wall can also occur in response to changes in transmural pressure. Upon exposure to increased transmural but not intraluminal pressure, increased incorporation of 14C cholesterol into the media of carotid arteries was due to reorganisation of the endothelial cell cytoskeleton and increased permeability Herman et al. (1987). Rearrangement of cytoskeleton and elongation were also observed in endothelial cells isolated from pulmonary arteries exposed to increased hydrostatic pressure (Acevedo et al., 1993). Stretch Other mechanical factors involved in vessel growth or remodeling which have been studied in vivo include stretch exerted by the surrounding tissues and disruption of the integrity of the extracellular matrix. The role of stretch as a stimulus for growth of larger vessels during development was demonstrated more than 100 years ago by Thoma (1893) as described above. Recent studies emphasized the role of stretch in the development of the vascularization in hearts transplanted in oculo and thus accessible to direct observation. Campbell and Rakusan (1991) attributed the better development of capillaries in beating than non-beating transplanted hearts to the stretch of myocytes. They also described the sequence of gradual vascular development with capillaries appearing at the end of the first week after transplantation and small
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arteries and veins after three weeks. Rongish et al. (1994) used a similar model, transplanting embryonic hearts before vascularization was established. Its development, studied by electron microscopy and immunohistochemistry, was similar to that found in utero, indicating that vascular development is dependent on the contractile activity of myocytes but not on the work performed by the heart to overcome the resistance of the whole vascular bed. Using specific vessel markers, it was established that PECAM-1 identified presumptive endothelial cells while fibronectin provided an environment which facilitated migration of endothelial cells and their precursors in the myocardium (Rongish et al., 1996). The appearance of laminin coincided with tube formation and was closely followed by appearance of collagen IV in the newlyformed basement membrane, while collagen I and III were components of the arteriolar and venular wall. Corrosion casts of capillary bed in the heart demonstrated changes in the configuration of capillaries during systole and diastole (Hossler et al., 1984) with a greater tortuosity during the former. Such constant length change might disturb the anchorage of capillaries to the extracellular matrix, and could be one of the stimuli for developmental capillary growth which is much more intensive in the heart than in most other organs. Later in life, however, the capillary wall may become refractory to this continuous change in length, although long-term stretch exerted, for instance, by increased volume of the ventricle when the duration of diastole is prolonged during persistent bradycardia still seems to be an important stimulus for capillary growth (Hudlicka and Brown, 1994; 1996). In a similar fashion, deformation of the conduit coronary arteries during increased ventricular filling resulted in an increase in proteosynthesis in those vessels (ramus interventricularis) which were stretched by the greater ventricular volume but not in the branch (ramus circumflexus) which was not affected (Gerova et al., 1995). The mechanism by which stretch can stimulate capillary growth and/or remodeling of larger vessels can be explained partly by its direct action on signal transduction (see the following section), and partly by possible disruption of the extracellular matrix and anchorage of vessels to it. Such a mechanism could be operative in capillary growth not only for stretched muscles but also to a certain degree in chronically stimulated or trained muscles. In the heart, collagen struts tether capillaries to myocytes and adopt different angular configurations depending on whether the heart was arrested in systole or diastole (Caulfield and Borg, 1979; Robinson et al., 1988). They could act as mechanical couplers between myocytes and vessels signalling mechanical forces to endothelial cells. Similarly, fibronectin, another extracellular matrix component located close to vessels (Casscells et al., 1990), could be involved in communicating physical stresses to endothelial cells (Courtoy and Boyles, 1983) and therefore play a role in myocardial angiogenesis in response to mechanical stimuli. Remodeling of the vascular bed and capillary growth is different depending on the type of stimulus and that includes not only the different effect of flow or pressure but also whether the stimulus acts from the luminal or abluminal side of the vessels, particularly true in case of capillaries. It is generally assumed that capillary growth starts by the disturbance of the basement membrane induced by increased activity of serine or metalloproteinases, followed by endothelial cell migration and proliferation
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in the cells closest to the original endothelial lumen (Ausprunck and Folkman, 1977; Beck and D’Amore, 1997). Our recent studies indicate that this general description is not valid in all situations of capillary growth. A similar degree of capillary growth could be induced in extensor digitorum longus muscles whether chronically stimulated, overloaded by stretch or exposed to chronic vasodilation by prazosin, albeit with differing time courses. Nevertheless, when the stimulus for capillary growth was acting from the luminal side of the vessels, as in prazosin treatment, capillary growth occurred without breakage of the basement membrane (Zhou et al., 1998a) or proliferation of endothelial cells (Brown et al., 1996) but by longitudinal splitting of existing vessels by luminal protrusions which were seen with both electron (Zhou et al., 1997b) and confocal (Hansen-Smith et al., 1996a) microscopy. In stretched overloaded muscles, where blood flow was not higher and the likely stimulus was acting from the abluminal side, capillary growth occurred by abluminal sprouting with disrupted basement membrane at the sprout tips (Zhou et al., 1998b). In stimulated muscles, increased blood flow represents a stimulus acting from the luminal side and continuous contraction and relaxation of muscle fibers disturbs the anchorage of capillaries to the extracellular matrix from the abluminal surface. In this situation, signs of both luminal protrusions similar to those in prazosin treated animals and breakage of the basement membrane were observed (Hansen-Smith et al., 1996a), preceded by endothelial cell proliferation (Pearce et al., 1995). Thus different stimuli induce different forms of angiogenesis which are not the same as those described in capillary growth under pathological circumstances.
MEDIATORS OF VASCULAR REMODELING The signal transduction mechanisms involved in growth of either endothelial or smooth muscle cells exposed to shear rate, shear stress, cyclic strain or stretch are discussed in detail in other chapters of this volume. To what extent they are applicable in vivo still remains to be elucidated but certainly deformation of endothelial cells by flow can induce a variety of changes, elegantly summarized by Davies (1995, 1997) Growth Factors Disturbance of the extracellular matrix in stretched and to a certain extent in chronically stimulated or trained muscles can, of course, activate growth factors, an aspect which has received surprisingly little interest in this context. Evidence for the involvement of basic fibroblast growth factor (bFGF) in stimulated (Brown et al., 1998) or stretched muscles was negative (Brown et al., 1992; Egginton et al., 1998), but increased levels of a small molecular endothelial stimulating growth factor (ESAF) (Weiss et al., 1984) were found in both models of angiogenesis (Brown et al, 1995; Egginton et al., 1998). Vascular endothelial growth factor (VEGF) is known to play an important role in the formation of vessels during development (Beck and D’Amore, 1997) and VEGF mRNA was found to increase after a single bout of muscle exercise (Breen et al., 1996) and in chronically stimulated muscles (Hang et al., 1995) where the peptide could be demonstrated in muscles stimulated for 4 but not 2 days (Jeal et al., 1997).
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Growth factors are considered to be important for vascular remodeling during development of the collateral circulation but information about which particular factors are significant is still under discussion. Both mRNA and protein ß-ECGF are found in developing collateral vessels after gradual occlusion of the coronary artery, with mRNA for VEGF and TGFß1 in the myocytes (Sharma et al., 1992). Bernotat-Danielowski et al. (1993) from the same laboratory demonstrated the presence of acidic but not basic fibroblast growth factor (FGF) in pig hearts with a complete coronary artery occlusion. FGFs and VEGF have both been used to stimulate the development of collateral circulation in heart (Banai et al., 1994) and skeletal muscle (Pu et al., 1993) and improve the training-induced enhancement of collateral flow in ischemic myocardium (Firoozan and Forfarr, 1996). Basic FGF is also important in vascular remodeling after endothelial damage of large arteries where upon infusion it stimulated proliferation of vascular smooth muscle cells (Agrotis and Bobik, 1996). An issue regarding the role of growth factors in vascular remodeling which is unresolved as yet is whether they are activated by changes in the hemodynamic forces or whether they themselves exert an influence by producing hemodynamic effects. Both basic FGF and acidic FGF have been reported to have a vasodilator action when infused intravenously, manifested by a decrease in arterial pressure (Cuevas et al., 1991). Basic FGF also dilated pial (Rosenblatt et al., 1994), hamster cheek pouch (Brown et al., 1996) and cremaster muscle arterioles (Mac Wu et al., 1996). It was localized on arterioles, but not capillaries, in chronically stimulated muscles (Brown et al., 1998) so it is plausible to assume that it may be involved as a vasodilator, thus contributing to other factors of hemodynamic origin as a stimulus for capillary growth. Integrins The role of integrins in cellular responses to mechanical stress and cell deformation was summarized recently (Ingber, 1997; Shyy and Su-Chien, 1997) and although their role in angiogenesis under pathological circumstances and in models of angiogenesis induced by growth factors has been demonstrated (Polverini, 1996), it is not known if they are involved in mechanically-induced growth of vessels in vivo. The significance of integrin actions is shown by the fact that disruption of the bond between vß3 integrin and the extracellular matrix by antibodies prevented vessel formation in chicken chorioallantoic membrane and in growth-factor induced angiogenesis in mouse cornea or retina, and that patients lacking ß3 integrin have problems with wound healing (Stromblad and Cheresh, 1996). vß3 integrin has been co-localized with metalloproteinase MMP-2 on sprouting capillaries in tumors (Brooks et al., 1996) which may explain its involvement in the process of the breakage of capillary basement membrane and endothelial cell migration. 2 and v integrins are also present on capillary sprouts in neonatal skin (Enenstein and Kramer, 1994). A possible link between flow-dependent mechanical factors and integrins was studied by Muller et al. (1997) whereby shear stress-dependent dilatation in isolated coronary arteries was inhibited by GRGDPN peptide—an antagonist of integrin binding to ECM.
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Prostaglandins and Nitric Oxide Two humoral agents activated by hemodynamic factors which have been shown to contribute substantially to vascular remodeling and growth are prostaglandins and nitric oxide (NO). Increased shear stress results in the release of both prostaglandins and NO from endothelium in arteries and arterioles (see Koller and Kaley, 1996) due to increased velocity of flow and/or viscosity. In both large arteries (Melkumyants et al., 1989) and skeletal muscle arterioles (de Wit et al., 1997), increased viscosity, achieved by increased hematocrit or exchange of plasma for high molecular weight dextran respectively, caused NO release, only if the endothelium was intact. This NO production leads to dilatation which either normalizes or decreases the shear stress; as a result, there is no relationship in the long-term between increased flow and shear stress e.g. in the carotid artery exposed to high flow due to A-V fistula (Tronc et al., 1996). Nevertheless, in this situation, increased flow resulted in a considerable proliferation of smooth muscle cells which was blocked by the nitric oxide synthase inhibitor L-NAME (Tronc et al., 1996). The release of NO in response to increased shear stress is greater in large than in small arterial vessels in skeletal muscle (Hester et al., 1993) and the heart (Kuo et al., 1995). In man, increased blood flow results in a greater release of NO from the coronary vascular bed than from forearm vessels (Tagawa et al., 1997) indicating a heterogeneous response to flow of vessels in different vascular beds even though the basal levels of NO are similar in all vessels (Mesaros and Grunfeld, 1997). The release of NO from the coronary vessels is increased by acute bouts of exercise both in animals (Bernstein et al., 1996) and man (Niebauer and Cooke, 1996). Exercise training upregulates endothelial NO synthase (eNOS) in rats (Sun et al., 1994), dogs (Shen et al., 1995) and pigs, in which the increased levels return to normal after 7–10 days of training (Laughlin, 1995). The flow-dependent endothelial release of prostaglandins (Koller and Kaley, 1990; Koller et al., 1994) is associated with deformation of microtubules in the endothelial cells (Sun et al., 1995) and tyrosine phosphorylation (Fleming et al., 1996). While it is quite clear that both prostaglandins and NO are involved in dilation of arterioles due to increased shear stress, this effect can be only very short-lasting as increased diameter normalizes shear stress very quickly, and, even if hematocrit and velocity of flow are increased during dilation, shear stress is either not changed or decreases. This was demonstrated by Sarelius (1993) and is shown in Figure 14.14. In contrast, increased blood flow due to muscle contractions or administration of vasodilators does not result in a great change in capillary diameters but the velocity of red blood cells and consequently shear stress are increased (see Figure 14.13). Under these conditions, prostaglandins and NO released from capillary endothelium could be involved in capillary growth. The exact role of NO or prostaglandins in vascular growth is still a matter of controversy. In tissue culture, NO has been described as an inhibitor as well as a stimulator of growth of endothelial and smooth muscle cells but the inhibitory effects could be due to the status of the cells to which it was applied and the dosage of NO donors used. For example, 3H-thymidine incorporation in vascular smooth muscle was inhibited only in cells in the synthetic phase (Assender et al., 1991) and
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Figure 14.14. Changes in wall shear stress, diameter and hematocrit in arterioles (1–4 represent different branching orders, the fourth being the most distal in the vascular network of hamster cremaster muscle) at rest and during adenosine-induced dilatation. Although hematocrit increased, wall shear stress was never higher than at rest because dilation did not result in a proportional increase in the velocity of flow. Based on data from Sarelius, 1993, with the author’s permission.
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by relatively high doses of NO donor (Sarkar et al., 1997), whereas low doses stimulated 3H-thymidine incorporation (Oconor et al., 1991) and potentiated basic FGF-induced mitogenesis (Hassid et al., 1994). High doses of NO donors (1 mM sodium nitroprusside) also inhibited proliferation of bovine aortic endothelial cells (BAEC) (Gooch et al., 1997) but increased levels of NOS were found in growing cells (Arnal et al., 1994) and promoted proliferation of subconfluent BAEC (Guo et al., 1995). In vivo, NO donors inhibit angiogenesis in CAM and tumors (Pilipi-Synetos et al., 1993, 1995) yet potentiate substance P- or VEGF-induced angiogenesis in the cornea (Ziche et al., 1994; Ziche et al., 1997). Both substance P and VEGF increase blood flow which of itself may generate NO. In chronically stimulated muscles where capillary shear stress was increased (Dawson and Hudlicka, 1993), capillary growth and proliferation of capillary-linked nuclei, demonstrated by BrdU incorporation, were inhibited by the NOS inhibitor L-NNA (Hudlicka et al., 1996). Apart from the work referred to above on the carotid artery by Tronc et al. (1996) there are no other data on the role on NO in remodeling or growth of larger vessels. The role of prostaglandins in vessel growth and remodeling has been studied by their implantation into the rabbit cornea to induce growth of vessels (Ben Ezra, 1978; Ziche et al., 1982). They also stimulated 3H-thymidine incorporation into cultured vascular smooth muscle cells (Pasricha et al., 1992), potentiated growth of vessels induced by basic FGF on CAM (Spisni et al., 1992) and caused growth of small vessels when applied in the rat femoral vein periadventitial tissue. Indomethacin, a cyclooxygenase and thus prostaglandin synthesis inhibitor, inhibited growth of vessels in tumors (Peterson et al., 1984). Although it has been well established that hemodynamic factors like shear stress release prostaglandins from the vessel endothelium (Koller and Kaley, 1990, 1996), the relationship between prostaglandins and hemodynamic changes in vessel growth or remodeling has been little studied. Inhibition of prostaglandin synthesis by indomethacin inhibited capillary growth and proliferation of capillary-linked nuclei in chronically stimulated muscles (Pearce et al., 1995) which was assumed to be linked with changes in the micro-circulation occurring in those muscle as a result of long-term increase in activity (Pearce et al., 1994). The mechanism of stimulatory effects of either prostaglandins or nitric oxide on capillary proliferation is still a matter of conjecture. Topper et al. (1996) demonstrated that cyclooxygenase and endothelial cell nitric oxide synthase are upregulated by steady laminar shear stress in large vessel endothelium but it is not clear whether the higher shear stress in capillaries in muscles with increased blood flow results in a similar upregulation. Mitchell and Tyml (1996) reported release on NO from capillaries in response to increase velocity of flow but the molecular mechanism between this release and growth still has to be elucidated. CONCLUSIONS Other chapters in this book show the importance of mechanical factors in the modification of endothelium in vitro, and elucidate the transduction mechanism involved. It is still not clear, however, to what extent the same mechanisms are valid
Figure 14.15 Scheme to show the effect of shear stress on intracellilar events leading to changes in vascular tone and possible proliferation of endothelial/smooth muscle cells.
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in the in vivo situation. The characteristics of endothelial cells differ in vivo not only with respect to size of vessels and organs, but many of their properties are lost when the cells are placed in tissue culture (Shima et al., 1995). Moreover, in vitro, changes due to the effect of mechanical factors have been studied over minutes and hours, while in vivo they may take weeks, months or even years. Furthermore, although interactions between individual cell types of different origins such as endothelial cells, pericytes, smooth muscle cells etc. have been investigated in tissue cultures, they may respond very differently within the complexity of the vascular wall. The combined effects of mechanical and hemodynamic factors cannot be separated from the multifactorial effects in vivo of growth, humoral or hormonal factors, nor can their influence on vascular cells be considered without acknowledgement of possible effects upon the surrounding tissues. The role of hemodynamic factors appears to vary according to the type of vessel, with shear stress and pulse pressure being most important in remodeling of the large arteries, while the effect of pressure is predominant in veins, small arteries and arterioles. Growth of arterioles is also regulated by wall tension, but this factor is of less importance in capillaries (except in the heart) than shear stress, and possibly mechanical factors such as stretch, which are important stimuli for growth. The growth pattern in capillaries also differs depending on whether the mechanical stimuli act from the luminal side, as in case of chronic pharmacologically-induced increases in blood flow, or from the abluminal side e.g. stretch of the surrounding tissue. From our own work, the example of skeletal muscle demonstrates that capillary growth in the former situation occurs by longitudinal splitting without endothelial cell proliferation, without breakage of the basement membrane, whereas growth in the latter develops by abluminal sprouting with proliferation. When both stimuli are combined, as in the case of long-term increases in muscle activity, both patterns of growth are observed. Although the transduction signals involved in the vessel growth in vivo remain to be fully elucidated, one possible scheme is presented in Figure 14.15.
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INDEX
1,4,5 inositol trisphosphate (IP3) 256 8-bromo cGMP 166 9-anthracene carboxylic acid (9-AC) 42 A23187 267 A431 cells 80 actin 39, 64, 277, 280, 284, 287 actinomycin D 230, 264 activator protein-1 (AP-1) 77, 119, 121, 135–137, 224, 225, 228, 232, 258–260, 284 activator protein-2 (AP-2) 265 adenosine 303, 306, 337 adenylyl cyclase 257 adhesion molecules 285, 287 adrenomedulin (AM) 112, 114 aequorin 256 amino acid pathway 38 anaemia 306, 327, 330 annexin II 40 P-11 heteromers 40 angiogenesis 276, 278, 292, 296, 312, 319 brain 294, 305 CAM 338 coronary vascular bed 312, 317–319 skeletal muscle 301, 316, 320 angiography 296, 313 angiotensin II 169, 308 converting enzyme (ACE) 112, 114, 149, 167, 181 ACE inhibitors 308 proliferation of endothelial cells 309 proliferation of smooth muscle cells 294 anion permeation through channels 38 anti-malaria drugs 42 anti-VCAM-1 196 anti-VLA-4 195–196 antisense oligonucleotides 19, 64 aorta 300, 310, 311, 313 aortic arch 229 apoptosis 277, 278, 311, 330 arachidonic acid (AA) 35, 265 arginine-vasopressin 166 arteries 8 carotid 292, 303, 304, 311, 336 remodeling 304
coronary 306, 310, 311, 318, 319, 333 femoral 313 iliac 313 pulmonary 300, 306, 310, 332 arterioles circumferential wall stress 329, 330, 331, 337 density 312, 317, 318, 327 hypertension 308, 309 rarefaction 308, 309 shear stress 306, 329, 330, 337 stimulated muscles 320, 321 arthritis 276, 279 artificial cardiovascular prostheses 226 aspirin 236 atherogenesis 191–192, 196–197, 201–202, 226, 234 atheroprotective 210 atherosclerosis 55, 127, 128, 138, 141, 207, 217, 275, 276 atherosclerotic lesions 226 plaques (fatty streak) 227 predilection sites 226, 228 adenosyl triphosphate (ATP) 61 ATP Binding Casette (ABC) transporters 47 atrophy—skeletal muscle 301, 325 barium 61 BAPTA/AM 264 basement membrane 226, 325, 333, 334 biaxial tensile stress 34 bicarbonate 37 bifurcation 128, 141, 229 blood 5 flow 55, 56 coronary 323 decreased 291, 293, 310–315 effect on growth of arterioles 330 effect on growth of capillaries 305 in skeletal muscle 313 growth of large vessels 311 heart 3, 12, 315 361
362
increased 291, 293, 303, 316, 327 limited in skeletal muscle 313, 314 pressure 15 role in growth of arteries 303 role in growth of capillaries 303 role in growth of smooth muscle cells 294 role in growth of veins 315 vessels 8 curvature 13, 14, 128, 141 distension 11 growth—quantification 322 mechanical properties 8 regression 292, 293 structure 8 wall tension 306, 307, 314, 319, 330 wall shear stress, see shear stress 11, 12 viscosity 6 viscous friction in 11 bovine arterial endothelial cell 58 carotid artery endothelial cells 166 bradycardia and growth of vessels drug induced 315, 322–324 electrical pacing 322–324, 331, 333 exercise 322 in myocardial infarction 323 bradykinin 61 brain capillary growth in development 294 induced by increased flow 305 bumetanide-sensitive cotransporter-2 213 C-3 exoenzymes 40 c-Crk 73 c-fos 113, 127, 131, 132, 135, 136, 137, 140, 141, 142, 259 fosB 259 c-jun 113, 259 c-jun N-terminal kinase (JNK) 74, 77 c-myc 113 c-src 71, 74, 78 C-type natriuretic peptide (CNP) 112, 114, 149, 164, 170, 181 regulation by chemical agonists 166 by shear stress 165 calcium entry 35 intracellular 61, 256, 264, 267 ionophore 267 release-activated calcium channels (CRAC) 35
Index
calphostin C 258, 264, 267, 268 capillary 276 diameters and growth 325, 329 pressure 308, 320, 333, 334 proliferation 320 red blood cell velocity 319, 328–330 shear stress 323, 328, 329, 331 splitting 320, 333, 334 sprouts 292, 293, 312, 320 supply (density, capillary/fiber ratio) 314, 326, 327 brain 300, 305 cold exposure 302 estimation 296, 297 heart 318, 319, 327 index of growth 309 limited blood flow 312–315 long-term dilatation 303, 306, 314, 330 skeletal muscle 315, 317, 320, 321 training 316–319 various species 302 wall tension 323, 328, 331 cardiac cycle 7 efficiency 7 cardiomyoplasty 320 cardiovascular system 5 cation channels 256 pressure activated 34 strech activated 34 caveolae 48 CD31 (PECAM-1) 57, 197–198, 201 cell adhesion 57, 58, 60, 63–65 morphology 276–278, 280 signaling 256 substrate attachment 65 channels, see Ion channels chloride channels 33 blockers 42 dependent kinase 47 reversal potential 38 chlorophenicol acetyltransferase (CAT) 261 chondrocyte 280 chorioallantoic membrane (CAM) 338 chromones 42 chronic hypertension 169–170 CIC-family 48 circulation collateral heart 311, 312 skeletal muscle 311, 313, 315 coagulation 235
Index
colcemide 39 colchicine 39 collagen 252 type I 277 type IV 235, 256, 278 cone and plate viscometer 10, 19, 95, 129, 130 contact inhibition 279 corkscrew 62 cotransporter 213 curvature, see vessel curvature cyclic AMP (cAMP) 224, 236, 257, 268 dependent protein kinase inhibitors 264 response element (CRE) 260, 265 cyclic strain, see also stretch 19, 208, 221, 224, 232, 236, 237, 249, 265, 292 IP3 256 mechanosignaling 256–261 PAI-1 258 sensitive genes 250, 269 cycloheximide 230 cyclooxygenase (COX) 210, 265 cyclooxygenase-2 (COX-2) 114, 224, 282–283 cytokines 207, 210, 228 cytoskeletal proteins 223, 252 cytoskeleton 39, 60, 64, 276, 277, 278, 280, 285, 286, 287, 339 dean number 4, 14 deletion constructs 261 development—growth of vessels 298–299, 301, 332, 333 dexamethasone 169 diabetes 212 diacylglycerols 256 dibutyeryl-cAMP 169 DIDS 42 differential display 122, 209, 210 dobutamine 323, 331 drugs, vasodilating effect on capillary growth 303, 306, 314, 330 E-selectin 114, 133, 149, 150, 181, 278–279 IL-1 151 interferon-␥ 152 lipopolysaccharide 151 shear stress 150, 151, 181 TNF-␣ 151 EDHF 44 EDTA 267 Egr-1 79, 127, 131, 132, 135, 136, 137, 140, 141, 142, 224, 232, 236, 239, 284
363
EGTA 264 elastic modulus 17 embryogenesis 279 electrogenesis 42 electrophoretic mobility shift assay (EMSA) 119, 120, 257, 259, 265 endoCAM (PECAM-1) 57, 197, 198, 201 endothelial cells 8, 11, 18, 56–58, 60–64, 190–203, 275–287 activation 297 capillaries 316, 317 collaterals 315 dysfunction 208 elongation 294 genes 216 heterogeneity 221 injury 328 large vessels 317 migration 203, 333 monocyte adhesion 190–196 nitric oxide synthase (eNOS) 80, 210, 265 permeability 190–191, 197, 201–202 proliferation 46, 291, 293, 320, 333 proliferation rate 197–203 stimulating angiogenic factor (ESAF) 334 ultrastructure 297, 298, 334 endothelin (ET) 112, 149, 170, 181, 250, 264, 282, 283 adrenaline 173 eNOS 224 IL-1␣ 174 ionomycin 174 oxygen tension 175 shear stress 171–173, 181, 182 thrombin 172 TPA 174 transforming growth factor-ß1 174 entrance length 13 epidermal growth factor 283 epidermal growth factor receptor (EGFR) 70, 80 exercise growth of arterioles 316, 317 exocytosis 45 extracellular matrix 256, 276, 278, 286, 294, 300, 327, 332, 333 extracellular signal regulated kinase (ERK), see mitogen activated protein kinases (MAPKs) fibroblast growth factor (FGF) 280–281, 282–283, 284, 286, 334, 335, 338 basic fibroblast growth factor (bFGF) 113, 166, 235
364
fibroblasts 236, 280, 332 fibronectin 252, 256, 277, 278, 333 flexcell® 232, 236 flow chamber circular 229, 238 parallel plate 20, 93, 129 fluid flow 56, 57, 60, 61, 63, 64 arterial 10 boundary layer 13 disturbed 10 entry 13 fully developed 12 Hagen-Poiseuille 11 laminar 3, 238 non-steady 223 non-uniform 2 oscillatory 223 patterns aberrant 226 local 239 pulsatile 4, 229, 300 low-shear 239 non-laminar 230 secondary, in curved vessels 4, 14 sensing 56 steady 3, 221 turbulent 3 uniform 2 unsteady 3 velocity 293, 314, 319, 323, 328 venous 10 shear stress, see shear stress 55–57, 60, 208 newtonian 2 non-newtonian 2 fluidity 238 fluorescein dextran 238 focal adhesion kinase (pp125FAK) 57, 70, 77, 223, 250, 251 focal adhesion plaque 251, 277, 286, 339 friction factor 14 fura-2 256 fyn 71 G protein 19, 223, 257 gadolinium 61 GDPßS 40 gene expression 190–191, 193–195, 197, 200–201, 209, 215, 261 gene transcription 118, 284 genistein 20, 40 glaucoma 276, 279 glycine 45 glycocalyx 330
Index
granulocyte/macrophage colony stimulating factor (GM-CSF) 122 growth factor receptor binding protein-2 (Grb2) 73, 77 growth factors 207, 335 GST fusion protein 62 GTPase-activating protein of Ras (RasGAP) 74 GTP␥S 40 heart 6 capillary density bradycardia 315, 322–324, 331, 333 hypertrophy 323, 326, 327, 310 long-term vasodilation 315, 322–324 myocardial infarction 312, 315, 323 training 322 HEK293 cell 60 hematocrit 330, 337 hemocompatibility 226 hemodynamic forces 2, 11, 19, 127, 128, 129, 143 aberrant 226 heparin-binding epidermal growth factor (HB-EGF) 113 herbimycin A 20 high shear 200–201 histamine 61 hormones 207 human dermal microvascular endothelial cells (HMVEC) 221, 233, 237 prostaglandin transporter 213 umbilical vein endothelial cells (HUVEC) 58, 112, 221, 233 hydrostatic pressure 208, 275–287 endothelial cell morphology 276 proliferation 279 surface antigens 278 tissue factor 238 membrane fluidity 238 hypercoagulability 234 hyperlipidemia 212 hyperosmotic 60, 63 hyperplasia 306, 308, 327 hypertension 234, 275, 276 arteries 294, 332 arterioles 308, 309 pressure overload 310 renal 308–309, 326 smooth muscle cells, hyperplasia 294 smooth muscle cells, hypertrophy 294 spontaneous (SHR) 294, 307, 308 hypertrophy
Index
capillary supply heart 323, 326, 327 capillary supply skeletal muscle 320, 324–325 heart—pressure overload 310, 326 heart—volume overload 326 hyposmotic 60 hypoxia 300, 303, 312, 326 IgE receptor 60 IB 138, 139 kinase complex (IKK) 79 immunocomplex (IP) kinase assays 71 immunoglobulin superfamily 59 inflamation 149, 150 inositol phosphates 256, 339 insulin 169 integrins 250, 252, 278, 284–286, 287, 335 ␣vb3 78 ␣2ß1 256 ␣5 256 intercellular adhesion molecule-1 (ICAM-1) 113, 131, 132, 133, 141, 149, 150, 181, 267, 278, 221, 227, 233, 282–283 antioxidants 153 hydrogen peroxide 156 IL-1 154 IL-4 155 interferon-␥ 154 lipopolysaccharide (LPS) 153 PMA, TPA 156 shear stress 152–154, 181 tumor necrosis factor a (TNF-a) 154 interleukin-1(IL-1) 228, 268 IL-1␣ 166, 167 IL-1ß 166, 167, 210 IL4 228 intracellular [Ca++], see Calcium intussusceptive capillary growth 294, 301 ion channels 33, 256, 285 activation 223 anion magnesium 39 permeation 38 calcium-release activated calcium channels CRAC) 35 cation pressure activated 34 strech activated 34 chloride channels 33 blockers 42 inactivation 39 non-selective ion 225 potassium 61, 225, 267 inwardly rectifying 34
365
ROM 34 small conductance (SKCa) 34 shear stress activated channels 34 stretch-activated 61 volume-regulated anion channels (VRAC) 37 ionic strength 41 ionophore 61 ischemia, heart 312, 315 ischemia, skeletal muscle 313, 314 JNK/SAPK 18, 225 JNKK 75 jun 127, 131, 132, 135, 136, 137, 140, 141 142, 259 junB 259 junD 259 keratinocyte 268 KT5720, KT5823 (cAMP protein kinase inhibitors) 264 L-NAME 267, 294 labeling of proliferation 3H thymidine, BrdU or PCNA 297, 313 arterioles 330 capillaries 313, 317 endothelial cells 327 smooth muscle cells 332 lactate 37 laminar flow, see shear stress 56 laminin 256, 278, 296, 333 laplace’s law 17 lesion-prone areas 191–192, 197, 201–202 leukocyte 278, 282, 287 adhesion molecules 226 lipopolysaccharide (LPS) 166–167, 279 loading, see also cyclic strain biaxial 26 circumferential 26 dynamic of arteries 18 time-dependent 17 uniaxial 25 low shear stress 190–199, 201–202 luciferase assay 118 lysophosphatidylcholine 228 MAD-related proteins 215 manganese-dependent superoxide dismutase 210, 224 MAP kinase see mitogen activated protein kinase mast cell 60
366
Index
MDR1 P-glycoprotein 47 mechanical force 65 signal 55, 65 stimuli 60, 62, 63 stress 63, 64 mechanoreception 240 mechanosensing 56, 64, 65, 241 mechano-sensitive response elements 240 mechanostimulation 33 ion channel activation 35 mechanotransduction 56, 57, 64, 65, 250, 278, 284–286 membrane, see also plasma membrane potential hyperpolarization 44 resting potential 44 proteins 56 unfolding 36 merocyanine 540 (MC540) 221, 238 metalloproteinases 333, 335 mibefradil 44 microscopy confocal 296, 322 electron 297, 319, 320, 333 intravital 298, 319 mitogen-activated protein kinases (MAPKs) 62, 63, 76, 223, 225, 250, 258 extracellular signal regulated kinase (ERK) 63, 64, 74, 225 ERK 1/2 13, 258 ERK5 18 JNK/SAPK 18, 225 JNKK 75 MAP kinase kinase kinase (MEKK) 75 MAP kinase kinases (MEKs) 64, 74 MEK inhibitor (PD98056) 258 MEK1 18 p38 kinase 18, 74 model systems 222 monocyte chemotacting protein-1 (MCP-1) 77, 113, 120, 131, 132, 133, 134, 135, 136, 190, 192–196, 198– 202, 209, 267, 283–284 epidermal growth factor 164 IL-1ß 162–163, 149, 150, 159–160, 181 IL-4 163 interferon-g 163 HDL 162 LDL 161–162 LPS 162–163, 164 monocyte colony-stimulating factor 163 nitric oxide 163–164 shear stress 160–161, 181
TNF-␣ 162–163 transforming growth factor-b 164 monocytes 57, 227 recruitment 190–196, 202 morphology 250 mRNA stability 122 multilayering 278, 279–280 multiple signaling pathways 261 myocardial infarction 312, 315, 323 Navier-Stokes equations 11 neutrophils 57 Newton’s law 2 NFAT 127, 134, 136 nitric oxide (NO) 15, 44, 112, 250, 265, 294, 300, 336, 338 nitric oxide synthase (NOS) 80, 113, 208, 282–283, 294, 336, 338 non-laminar flow patterns 232 nonreceptor PTKs 70 NPPB 42 nuclear factor kappa B (NF-B) 79, 121, 127, 134, 136–139, 142, 198, 199, 200,209, 224, 225, 227, 236, 237, 260, 284 nuclear transcription factors, see also individual transription factors 121, 127, 131, 132, 137, 142, 143, 198–200, 221, 224, 241 nuclear run-off transcription analysis 261 okadaic acid 79 oncotic pressure 275 osmotic pressure 275 ovulation 279 p21RhoA 40 p38 kinase 18, 74 p70/p85 S6 kinase (pp70S6K) 80 p96 278 p130cas 73 p-Glycoprotein 47 pICln-channel 47 parallel plate apparatus 8, 19, 20, 129, 130 passivate 238 paxillin 251 pBPB 42 PD 98059 6, 258 PECAM-1(CD31) 60–64 pericytes 298, 332 pertussis-toxin 257 PGE2 268 pH regulation 37 phorbol ester 265
Index
phorbol myristate acetate (PMA) see also tumor promoting agent (TPA) 61, 169, 265 phosphatidylinositol 3-kinase (PI 3-kinase) 74, 282 phospholipase A2 35 phospholipase C 257, 339 phospholipase C-␥ 74, 286 phospholipase D 257 phosphorylation 13, 250 pinocytosis 223 plasma membrane 56–58, 60, 285 plasminogen activator inhibitor-type1 (PAI-1) 237, 250 platelet 57, 60, 62 platelet derived growth factor (PDGF) 113, 120, 121, 250, 261, 284 PDGF-A 131, 132, 135, 136, 137, 140, 141, 209, 264 PDGF-B 131, 132, 133, 134, 136, 137, 139, 140, 141, 264 Platelet-derived growth factor receptor (PDGFR) 70 Platelet endothelial cell adhesion molecule-1 (PECAM-1) 57–65, 81, 197–198, 201, 224, 278 Poisson’s ratio 17 prazosin 303, 329, 330, 331, 334 pre-atherosclerotic lesions 227 pressure 9, 11 capillary 308, 319, 323, 331 dynamic 15 elevated 19, 23, 221, 225, 238 hydrostatic 15, 23, 225, 234, 235, 238, 249, 275 lateral 16 oscillatory 223 perfusion 308, 315, 326 potential energy 15 pulsatile 18, 225, 307, 308, 332 transmural 17, 332 pressurized cell culture system 23 proliferation 46, 276, 277, 278, 279, 280, 281, 282, 284, 286, 298, 313 proliferative rate 250 promoter deletion studies 236 promoters 209 prostacyclin (PGL2) 237, 250, 264, 282–283 prostacyclin synthase (PGIS) 264 prostaglandin H synthase-1 149, 176–177, 181 heparin binding growth factor-1 180–181 IL-1␣ 178–179
367
PMA 180 shear stress 177–178, 181, 182 prostaglandin H synthase-2 149, 176–177, 181 IL-1␣ 178 PMA 179–180 serum 178 shear stress 177–178, 181, 182 prostaglandins 336 prostanoids 213 protein kinase A (PKA) 268 protein kinase C (PKC) 19, 20, 61, 76, 224, 228, 236, 258, 259, 264, 267, 268, 284, 286, 339 protein phosphatase 257 protein phosphorylation 69, 228 protein tyrosine kinases (PTKs) 40, 70 nonreceptor PTKs 70 phosphatases (PTP) 40 protein tyrosine phosphatases (SH-PTP1 and SH-PTP2) 74 PTP1D 62 PTP2C 62 pulsatile flow 9, 15, 198–199 pyrrollidine dithiocarbamate (PDTC) 221, 227, 230, 237 raf-1 20 ras 19, 73, 76, 78 rasN17 76 re-circulating eddy 238 reactive oxygen species (ROS) 258 receptor tyrosine kinases (RTKs) 70 receptor tyrosine phosphorylation 223 red blood cells endothelial cell interaction 308 velocity 306, 314, 319, 320, 323, 328, 329 reendothelialization 202–203 remodeling 127, 128, 217, 293, 294, 304,308, 311, 315, 319, 320, 327, 328, 332 reorientation 223 response element cyclic strain 260 perturbed flow 232 shear stress, see also SSRE 121, 136 restenosis 203 Reynolds number 3, 11 serine phosphorylation 60 serine/threonine protein kinases 18, 70, 224 shc 74 shear rate 304, 327, 328 shear stress disturbed 130, 140, 141, 142
368
Index
flow induced 1, 2, 34, 35, 69, 112, 165–168, 181, 182, 189–203, 207, 223, 249, 256, 264, 267, 275, 276, 278, 281– 284, 287, 292 effects on angiotension converting enzyme 167–168, 181, 182 C-type natriuretic peptide 165–166, 181, 182 E-selectin 150–151, 181 endothelin-1 171–173, 181, 182 gene activation 257 ICAM-1 152–154, 181 ion-channels 34 MCP-1 160–161, 181 prostaglandin H synthase-1 177–178, 181, 182 prostaglandin H synthase-2 177–178, 181, 182 tissue factor 79, 238 VCAM-1 156, 181, 182 in arterioles 312, 336, 337 capillaries 306, 314, 320, 323, 328 large vessels 394, 328 venules 329 gradients 223 laminar 129, 130, 131, 132, 137, 207, 264, 265, 281–283 magnitude high 200–201 low 190–199, 201–202 oscillatory 140, 141 pulsatile 128, 130, 140, 141, 221 response element (SSRE) 79, 120, 121, 127, 134, 135, 136, 137, 139, 142, 209, 223, 228, 243, 261, 264, 265 turbulent 3, 140, 207, 278, 281–282 signal transduction 56, 57, 62, 65, 198– 200, 216, 223 SITS 42 skeletal muscle electrically stimulated 319–322 models of angiogenesis 328 smad proteins smad6 215 smad7 215 smooth muscle cell 8, 280 proliferation 307, 332, 336, 339 son of sevenless (sos) 73, 77 SP-1 79, 127, 135, 136, 137, 224, 232, 239 sprouts abluminal 325 capillary 293, 325, 333 luminal 320, 333, 334
src 20, 62, 64 src-homology domain-2 (SH2) 62, 64, 73 SH3 73 SHP2 62, 63 SH-PTP2 62, 63, 64 SSRE, see shear stress response element stagnation 238 staurosporine 268 steady laminar 207 strain, see cyclic strain stress, see also shear stress biaxial tensil 26, 34 compressive 16 circumferential 17 fibers 252, 277–278, 281 normal 11 stretch, see also cyclic strain 256, 300, 323, 325, 327, 332 ion channels 61 tensile 16 substraction hybridization 209 superoxide dismutase (SOD) 114 syp 62 T lymphocyte 57 talin 278 tamoxifen 42 taurin 45 taxol 39 tensegrity model 250 tensile stress 276, 281–283, 287 biaxial 34 tetradecanoyl phorbol acetate (TPA) see also PMA 61, 166, 169, 284–285 tetraethyl ammonium ion (TEA) 61, 267 thrombin 41, 61, 166 thrombomodulin (TM) 114 thyroid hormone 169 tissue factor (TF) 79, 115, 120, 131, 132, 135, 136, 221, 235, 238, 239 tissue plasminogen activator (tPA) 113, 250, 265, 283 tissue-specific 241 gene expression 234 total artificial blood pumps 226 transcriptional regulation 230 transforming growth factor ß (TGF-ß) 166, 215, 224, 283 TGF-ß1 114, 209 transmigration of white blood cell 58 transport 45 transporters 213 tropoisomerase II 228 tumor 276, 279
Index
tumor necrosis factor (TNF␣) 167, 221, 236 tumor promoting agent (TPA) 166 see also PMA TPA response element (TRE) 77, 134, 135, 136, 267 tumors—vascular supply 291, 335 turbulence 10, 210 turbulent shear stress 3, 207, 278, 281–282 tyrphostin 40 tyrophostinsA25 252 tyrosine kinase 20, 62, 63 phosphatase 62, 64 phosphorylation 56, 57, 60–64, 251 umbilical vein 216 vascular cell adhesion molecule-1 (VCAM-1) 114, 115, 131, 132, 133, 135, 141, 142, 149, 150, 181, 190, 192–196, 198–202, 209, 210, 221, 227, 233, 267, 278, 282–283 IL-1 159 IL-4 158–159 interferon-␥ 158 LPS 158–159 nitric oxide 159 shear stress 115, 156, 181, 182 TNF-␣ 157–159
369
vascular casts 296, 313, 317, 333 conductance 296, 316, 317, 321, 323 endothelial cells (ECs) 69, 207 endothelial growth factor (VEGF) 64, 235, 299, 326, 334, 445, 338 homeostasis 207 remodeling 208 tone 149 vasculogenesis 299 vasodilation see blood flow increased vasoprotective 210 VEGF, see vascular endothelial cell growth factor vein discovery 207 veins 8, 306, 315, 322 venules 306, 314, 331 verapamil 267 vessel, see blood vessel vinculin 278 viscosity, apparent 6 viscous damping 18 vitronectin 252, 256, 277 volume regulated anion channels (VRAC) 37 regulation 37 Womersley number 5, 15 wound healing 279 angiogenesis in 291, 292 ZO-1 58
