Tight Junctions, Second Edition

TIGHT JUNCTIONS Second Edition

TIGHT JUNCTIONS Second Edition

Edited by

MARCELINO CEREIJIDO Department of Physiology Biophysics and Neuroscience Cinvestav, Mexico

JAMES ANDERSON Yale School of Medicine New Haven, Connecticut

CRC Press Boca Raton London New York Washington, D.C.

Library of Congress Cataloging-in-Publication Data Tight junctions / edited by Marcelino Cereijido, James Anderson.--2nd ed. p. cm. Includes bibliographical references and index. ISBN 0-8493-2383-5 (alk. paper) 1. Tight junctions (Cell biology). I. Cereijido, Marcelino. II. Anderson, James, 1952-. QH603.C4 T54 2001 571.6--dc21

2001035330

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Preface A major function of polarized epithelia is to separate tissue spaces and regulate the exchange of material between them. The tight junction (TJ) creates the paracellular component of this barrier. In the past, the TJ was considered to be a permanent structure whose only function was to block the passage of substances between the cells. We now know the barrier is regulated and highly variable among different types. Its regulation occurs in response to physiologic, pharmacologic, and pathologic conditions. The first edition of this book, published a decade ago, was the first attempt to systematically review all aspects of the biology of TJs. Since its publication there has been an unprecedented expansion of information, including enumeration of the TJs’ molecular components. This has provided new insights into the regulation of permeability, cell signaling, and disease. This new information is coming from researchers in traditionally separate fields; thus we felt this was an appropriate time and valuable effort to again collect these many viewpoints in a second edition. The first part of this book (Chapters 1 through 9) serves as a general introduction to the structure and physiology of the tight junction. This material is best appreciated in the context of the biology of epithelial cells; consequently, we include discussion of cell polarity, protein sorting, and other types of intercellular junctions. We felt it was also important to present the relevant theoretical and technical issues required to study tight junctions. The second part of the book (Chapters 10 through 21) reviews more focused aspects of the TJ, but those that can still be generalized to TJs in all cell types. This includes discussion of its molecular components, and cellular mechanisms for regulating permeability and cell signaling. The final part (Chapters 22 through 33) deals with the role of tight junctions in pathologic conditions or issues that are unique to specific organs. Among other topics, this includes genetic diseases and the effects of bacterial toxins and inflammation. Within this last section are chapters addressing the practical question of whether paracellular permeability could be manipulated for therapeutic purposes such as to enhance drug absorption. We have convened a group of the world's leading experts to address these topics. A few are veterans of the first edition. They attacked their assignments with enthusiasm and we are humbled by and grateful for their outstanding efforts. We would like to thank all the authors and Ms. Dorothy Franco and Elizabeth del Oso for their success in keeping all 69 of us on track and on time.

The Editors Marcelino Cereijido, M.D., Ph.D., is Professor of Physiology and Biophysics at the Center for Research and Advanced Studies in Mexico City, Mexico. He is also a Career Investigator of the National Research System, Mexico, since 1984, and an Endowed Chair, since 1994. He received his Doctor of Medicine degree from the University of Buenos Aires in 1957 and his Ph.D. in Physiology in 1961. His postdoctoral studies were conducted at the Biophysical Laboratory, Harvard Medical School (1961–1964). Dr. Cereijido has also held professorial positions at a variety of international universities, including the University of Buenos Aires, the Albert Einstein Center of Medical Research, and the New York University School of Medicine. He has written extensively on the science of politics and the social impact of science. His current professional focus is in the area of biological membranes. James Melvin Anderson, Ph.D., M.D., is Professor of Internal Medicine and Cell Biology and Chief of the Section of Digestive Diseases at the Yale University School of Medicine in New Haven, Connecticut. He received his Ph.D. degree in Biology at Harvard University (1979) and his M.D. from the Harvard Medical School (1983) in Boston, Massachusetts. His research group cloned the first tight junction protein, ZO-1, and has provided numerous insights into the basic cell and molecular biology of the tight junction. He has written widely on tight junctions with an emphasis on the interface between basic research and its implications for human disease.

Contributors James Melvin Anderson, Ph.D., M.D. Yale University School of Medicine New Haven, Connecticut Antonia Avila, M.S. Department of Physiology, Biophysics, and Neurosciences Center for Research and Advanced Studies Mexico City, Mexico Maria S. Balda, Ph.D. Department of Cell Biology Institute of Ophthalmology University College of London London, United Kingdom Neal Beeman, B.S. Department of Physiology University of Colorado Health Sciences Center Denver, Colorado Gaëlle Benais-Pont Department of Cell Biology Sciences III University of Geneva Geneva, Switzerland

Vera Bonilha, Ph.D. Margaret Dyson Vision Research Institute Weill Medical College of Cornell University New York, New York Alan R. Burns, Ph.D. Department of Medicine, Section of Cardiovascular Sciences and Department of Pediatrics, Section of Leukocyte Biology Baylor College of Medicine Houston, Texas Marcelino Cereijido, M.D., Ph.D. Center for Research and Advanced Studies Mexico City, Mexico Keith A. Choate Departments of Genetics, Cell Biology, and Medicine Howard Hughes Medical Institute Yale University School of Medicine New Haven, Connecticut

Yehuda Ben-Shaul, Ph.D. Department of Cell Research and Immunology George S. Wise Faculty of Life Sciences Tel Aviv University Tel Aviv, Israel

Sandra Citi, M.D., Ph.D. Department of Molecular Biology University of Geneva Geneva, Switzerland and Department of Biology University of Padova Padova, Italy

Abigail Betanzos, M.S. Department of Physiology, Biophysics, and Neurosciences Center for Research and Advanced Studies Mexico City, Mexico

David Cohen, M.D., Ph.D. Margaret Dyson Vision Research Institute Weill Medical College of Cornell University New York, New York

Ruben Gerardo Contreras, Ph.D. Department of Physiology, Biophysics, and Neurosciences Center for Research and Advanced Studies Mexico City, Mexico Pamela Cowin, Ph.D. Departments of Cell Biology and Dermatology New York University School of Medicine New York, New York Judith Eckert, Ph.D. School of Biological Sciences University of Southampton Southampton, United Kingdom Rachel Eelkema Departments of Cell Biology and Dermatology New York University School of Medicine New York, New York Alan S. Fanning, Ph.D. Department of Internal Medicine Yale University School of Medicine New Haven, Connecticut Alessio Fasano, M.D. Department of Pediatrics, Medicine, and Physiology School of Medicine University of Maryland Baltimore, Maryland Irina Fesenko, Ph.D. School of Biological Sciences University of Southampton Southampton, United Kingdom Tom P. Fleming, Ph.D. School of Biological Sciences University of Southampton Southampton, United Kingdom

Michael Fromm, M.D. Department of Clinical Physiology Universitätsklinikum Benjamin Franklin Freie Universität Berlin Berlin, Germany Lorenza González-Mariscal, Ph.D. Department of Physiology, Biophysics, and Neurosciences Center for Research and Advanced Studies Mexico City, Mexico Alexander Gow, Ph.D. Center for Molecular Medicine and Genetics/Departments of Pediatrics and Neurology Wayne State University School of Medicine Detroit, Michigan Gail Hecht, M.D. Section of Digestive and Liver Diseases Department of Medicine West Side VA Medical Center University of Illinois Chicago, Illinois Dr. Nanette Kälin Department of Cell Biology and Histology Academic Medical Center University of Amsterdam Amsterdam, The Netherlands Karl J. Karnaky, Jr., Ph.D. Department of Cell Biology and Anatomy and the Marine Biomedical and Environmental Sciences Program Medical University of South Carolina Charleston, South Carolina and the Mt. Desert Island Biological Laboratory Salisbury Cove, Maine

Olga N. Kovbasnjuk, Ph.D. Department of Medicine Division of Gastroenterology Johns Hopkins University School of Medicine Baltimore, Maryland Geri Kreitzer, Ph.D. Margaret Dyson Vision Research Institute Weill Medical College of Cornell University New York, New York Lukas Landmann, Ph.D. Department of Anatomy University of Basel Basel, Switzerland Nancy J. Lane, Sc.D. Department of Zoology University of Cambridge Cambridge, England Chao-Pin Lee, Ph.D. Drug Delivery Systems GlaxoSmithKline Collegeville, Pennsylvania Simon A. Lewis, Ph.D. Department of Physiology and Biophysics University of Texas Medical Branch Galveston, Texas Richard P. Lifton, M.D., Ph.D. Departments of Genetics, Medicine, and Molecular Biophysics and Biochemistry Howard Hughes Medical Institute Yale University School of Medicine New Haven, Connecticut Yin Lu, Ph.D. Department of Genetics Howard Hughes Medical Institute Yale University School of Medicine New Haven, Connecticut

Robert D. Lynch, Sc.D. Department of Biological Sciences University of Massachusetts Lowell, Massachusetts James L. Madara Department of Pathology Emory University School of Medicine Atlanta, Georgia James A. Marrs, Ph.D. Department of Medicine Indiana University School of Medicine Indianapolis, Indiana Karl Matter, Ph.D. Department of Cell Biology Institute of Ophthalmology University College of London London, United Kingdom Bruce A. McClane, Ph.D. Department of Molecular Genetics and Biochemistry University of Pittsburgh School of Medicine Pittsburgh, Pennsylvania Laura L. Mitic, Ph.,D. Department of Internal Medicine Yale University School of Medicine New Haven, Connecticut Bruce A. Molitoris, M.D. Department of Medicine Indiana University School of Medicine Indianapolis, Indiana Collin G. Murphy, Ph.D. Department of Ophthalmology University of California San Francisco, California

Anne Müsch, Ph.D. Margaret Dyson Vision Research Institute and Department of Biochemistry Weill Medical College of Cornell University New York, New York

Enrique Rodriguez-Boulan, Ph.D. Margaret Dyson Vision Research Institute and Department of Cell Biology Weill Medical College of Cornell University New York, New York

Margaret C. Neville, Ph.D. Department of Physiology University of Colorado Health Sciences Center Denver, Colorado

Eveline E. Schneeberger, M.D. Department of Pathology Massachusetts General Hospital and Harvard Medical School Boston, Massachusetts

Duy-Ai D. Nguyen, Ph.D. Department of Physiology University of Colorado Health Sciences Center Denver, Colorado

Jörg-Dieter Schulzke, M.D. Department of Clinical Physiology Universitätsklinikum Benjamin Franklin Freie Universität Berlin Berlin, Germany

R.-Marc Pelletier, Ph.D. Départment de Pathologie et Biologie Cellulaire, Faculté de Médicine Université de Montréal Montreal, Quebec, Canada

Bhavwanti Sheth, Ph.D. School of Biological Sciences University of Southampton Southampton, United Kingdom

Ilana Ophir, Ph.D. Department of Cell Research and Immunology George S. Wise Faculty of Life Sciences Tel Aviv University Tel Aviv, Israel

Liora Shoshani, Ph.D. Department of Physiology, Biophysics, and Neurosciences Center for Research and Advanced Studies Mexico City, Mexico

Luis Reuss, M.D. Department of Physiology and Biophysics University of Texas Medical Branch Galveston, Texas

Usha Singh, Ph.D. Department of Molecular Genetics and Biochemistry University of Pittsburgh School of Medicine Pittsburgh, Pennsylvania

Lawrence J. Rizzolo, Ph.D. Departments of Surgery and of Ophthalmology and Visual Science Yale University School of Medicine New Haven, Connecticut

C. Wayne Smith, M.D. Department of Pediatrics Section of Leukocyte Biology Baylor College of Medicine Houston, Texas

Philip L. Smith, Ph.D. Drug Delivery Systems GlaxoSmithKline Collegeville, Pennsylvania

Jerrold R. Turner, M.D. Department of Pathology Wayne State University School of Medicine Detroit, Michigan

Cherie M. Southwood Brookdale Center for Development and Molecular Biology Mount Sinai School of Medicine New York, New York Kenneth R. Spring, D.M.D., Ph.D. Laboratory of Kidney and Electrolyte Metabolism National Heart, Lung and Blood Institute National Institutes of Health Bethesda, Maryland

Johnnie L. Underwood, Ph.D. Department of Ophthalmology University of California San Francisco, California Christina M. Van Itallie, Ph.D. Department of Internal Medicine Yale University School of Medicine New Haven, Connecticut

Bruno Stieger, Ph.D. Division of Clinical Pharmacology Department of Internal Medicine University Hospital Zurich, Switzerland

Prof. Gerrit van Meer Department of Cell Biology and Histology Academic Medical Center University of Amsterdam Amsterdam, The Netherlands

Fay Thomas School of Biological Sciences University of Southampton Southampton, United Kingdom

David C. Walker, Ph.D. Department of Pathology University of British Columbia Vancouver, British Columbia, Canada

Steven D. Wilt, Ph.D. Department of Biology Kentucky Wesleyan College Owensboro, Kentucky

Table of Contents Chapter 1 Introduction: Evolution of Ideas on the Tight Junction............................................1 James Melvin Anderson and Marcelino Cereijido Chapter 2 Ultrastructure and Immunolabeling of the Tight Junction .....................................19 Eveline E. Schneeberger and Robert D. Lynch Chapter 3 Tight Junctions in Invertebrates ..............................................................................39 Nancy J. Lane Chapter 4 Tight Junction Permeability to Ions and Water ......................................................61 Luis Reuss Chapter 5 The Relationship between Structure and Function of Tight Junctions ..................89 Lorenza González-Mariscal, Antonia Avila, and Abigail Betanzos Chapter 6 General Themes in Cell–Cell Junctions and Cell Adhesion ................................121 Rachel Eelkema and Pamela Cowin Chapter 7 Protein Targeting Pathways and Sorting Signals in Epithelial Cells ...................145 Enrique Rodriguez-Boulan, Geri Kreitzer, David Cohen, Vera Bonilha, and Anne Müsch Chapter 8 Biogenesis of Epithelial Polarity and Tight Junctions..........................................165 Liora Shoshani and Ruben Gerardo Contreras Chapter 9 Optical Methods for the Study of Tight Junctions ...............................................199 Olga N. Kovbasnjuk and Kenneth R. Spring

Chapter 10 Occludin and Claudins: Transmembrane Proteins of the Tight Junction.............213 Laura L. Mitic and Christina M. Van Itallie Chapter 11 The Cytoplasmic Plaque Proteins of the Tight Junction ......................................231 Sandra Citi Chapter 12 Organization and Regulation of the Tight Junction by the Actin–Myosin Cytoskeleton ..........................................................................................................265 Alan S. Fanning Chapter 13 Developmental Assembly of the Tight Junction ...................................................285 Tom P. Fleming, Bhavwanti Sheth, Fay Thomas, Irina Fesenko, and Judith Eckert Chapter 14 Tight Junctions and Cell Surface Lipid Polarity ..................................................305 Nanette Kälin and Gerrit van Meer Chapter 15 Physiological Regulation of Tight Junction Permeability by Na+-Nutrient Cotransport.............................................................................................................333 Jerrold R. Turner and James L. Madara Chapter 16 Extracellular Macromolecules Modulate Epithelial Permeability........................349 Simon A. Lewis Chapter 17 Intracellular Signaling in Classical and New Tight Junction Functions ..............367 Gaëlle Benais-Pont, Karl Matter, and Maria S. Balda Chapter 18 Regulation of Tight Junction Permeability in the Mammary Gland....................395 Duy-Ai D. Nguyen, Neal Beeman, and Margaret C. Neville Chapter 19 Unique Aspects of the Blood–Brain Barrier.........................................................415 Steven D. Wilt and Lawrence J. Rizzolo

Chapter 20 Teleost Chloride Cell Tight Junctions: Environmental Salinity and Dynamic Structural Changes.................................................................................................445 Karl J. Karnaky, Jr. Chapter 21 Tight Junctions and Proteases ...............................................................................459 Yehuda Ben-Shaul and Ilana Ophir Chapter 22 Claudins Mediate Specific Paracellular Fluxes in Vivo: Paracellin-1 Is Required for Paracellular Mg2+ Flux.....................................................................483 Keith A. Choate, Yin Lu, and Richard P. Lifton Chapter 23 Microbial Pathogens That Affect Tight Junctions ................................................493 Gail Hecht Chapter 24 Interactions between Clostridium perfringens Enterotoxin and Tight Junction Proteins....................................................................................................517 Bruce A. McClane and Usha Singh Chapter 25 Ischemia-Induced Tight Junction Dysfunction in the Kidney..............................533 James A. Marrs and Bruce A. Molitoris Chapter 26 Tight Junctions in Intestinal Inflammation ...........................................................553 Jörg-Dieter Schulzke and Michael Fromm Chapter 27 Tight Junctions in Liver Disease...........................................................................575 Lukas Landmann and Bruno Stieger Chapter 28 The Tight Junctions in the Testis, Epididymis, and Vas Deferens .......................599 R.-Marc Pelletier Chapter 29 Relationship Between Tight Junctions and Leukocyte Transmigration ...............629 Alan R. Burns, David C. Walker, and C. Wayne Smith

Chapter 30 Ocular Tight Junctions in Health, Disease, and Glaucoma..................................653 Johnnie L. Underwood and Collin G. Murphy Chapter 31 Implications of Transport via the Paracellular Pathway on Drug Development ..........................................................................................................685 Philip L. Smith and Chao-Pin Lee Chapter 32 Pathological and Therapeutical Implications of Macromolecule Passage through the Tight Junction ....................................................................................697 Alessio Fasano Chapter 33 Functions of OSP/Claudin-11-Containing Parallel Tight Junctions: Implications from the Knockout Mouse ...............................................................723 Cherie M. Southwood and Alexander Gow Index......................................................................................................................747

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Introduction: Evolution of Ideas on the Tight Junction James Melvin Anderson and Marcelino Cereijido

CONTENTS 1.1 A Mere Terminal Bar.......................................................................................1 1.2 Not Always So Tight........................................................................................4 1.3 What Is a Tight Junction?................................................................................7 1.4 Tight Junctions and Apical/Basolateral Polarity .............................................9 1.5 Biosynthesis and Assembly of the Tight Junction ........................................11 1.6 New Roles in Signaling .................................................................................11 1.7 Tight Junctions in Special Situations ............................................................12 1.8 The Role of Tight Junctions in Human Disease ...........................................12 1.9 Concluding Remarks......................................................................................13 References................................................................................................................14

1.1 A MERE TERMINAL BAR The ability of transporting epithelia to act as diffusion barriers between compartments with different composition and to withstand steep chemical and electrical gradients requires a seal between the cells (Figures 1.1 and 1.2); otherwise, these gradients would dissipate through the intercellular space. This seal was expected to be located at the very limit between the lumen and the intercellular space; otherwise, it would interfere with the exchange of nutrients between the cells and the internal milieu. Furthermore, since the physiology of these epithelia was initially — and for many years — studied in the frog skin, where fluxes through the intercellular space are practically negligible, this seal was also expected to be impermeable. Therefore, it is understandable that for almost a century the anatomical formations detected with light microscopy at the outermost end of the intercellular space received names such as “Schlussleisten,” “terminal bars,” “bandelettes de fermeture,” “hoops,” “occluding junctions,” “tight junctions,” “gaskets,” and “attachment belts” (Bizzozero, 1870; Bonnet, 1895; Dahlgren and Kepner, 1925; Schaffer, 1927; Fawcett and Selby, 1958; Bennett et al., 1959; Palay and Karlin, 1959; Fawcett, 1961).

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FIGURE 1.1 Epithelial surface as observed from the apical side, in a freeze-fracture replica of three adjacent cells from frog skin. These cells adapt their lateral borders and occlude the intercellular space, thus conferring to the epithelium the property of a diffusion barrier. (Courtesy of Prof. A. Martínez-Palomo.)

The introduction of electron microscopy in the late 1950s permitted observation of the plasma membrane stained with OsO4 as a sequence of three layers, [dark]–[light]–[dark], which corresponded to the [cytoplasmic polar groups]–[hydrophobic chains]–[external polar groups] of biochemical models. With this approach,

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FIGURE 1.2 Clockwise from top left: Schematic representation of a kidney tubule. Six epithelial cells form tight junctions that severely reduce the escape of substances from the lumen (LUM). Two adjacent epithelial cells form tight junctions (TJs), intermediate junctions (IJs), desmosomes (D), and gap junctions (GJs). Lateral side of an epithelial cell: while the filaments of the tight junction form a continuous belt that completely surrounds the cells at the apical/basolateral border, desmosomes and gap junctions only occur at a discrete spot. Strands represented at a higher magnification to illustrate hypothetical channels can be in an open or a closed state. Current may only flow through those channels that are open at a given moment.

the region of the tight junctions (TJs), where the plasma membranes of two neighboring cells come into contact, did not appear to have six layers (three from each membrane), but only five, because the single dark layer in the middle results from the fusion of the external dark bands of the two membranes (Robertson, 1958; 1960; Moe, 1960; Karrer, 1960a,b; Millington and Finean, 1962). Such images, resulting from the early fixation techniques, fostered the erroneous notion that the outer leaflets were physically fused and that the TJ was a fixed and impermeable barrier. The finer resolution of electron microscopy also demonstrated that the “terminal bar” is in fact a complex of different types of specialized intercellular junctions (see Figure 1.2), which received the names of tight junction (zonula occludens), intermediate junction (zonula or fascia adhaerens), and desmosome (macula adhaerens) (Farquhar and Palade, 1963). In fact, desmosomes were well known from earlier studies (Palade and Porter, 1954). Today, it is known that neighboring cells may also

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establish gap junctions, which may sometimes be intercalated between the tight junctions (Robertson, 1963; Robertson et al., 1963; Loewenstein, 1981; Ramon and Rivera, 1986). Later on, freeze-fracture studies of both epithelia and endothelia revealed that the TJ consists of a distinctive reticular pattern or meshwork of fibrils embedded in the plane of the membrane (Figures 1.2 and 1.3) (Staehelin et al., 1969; Chalcroft and Bullivant, 1970; Staehelin, 1973; 1975). Junctions of similar appearance have also been documented in invertebrate species, although it is not resolved whether they are biochemically similar. Current information on the anatomy of tight junctions in vertebrates is reviewed by E. Schneeberger and R. D. Lynch (Chapter 2) and in invertebrates by N. J. Lane (Chapter 3).

1.2 NOT ALWAYS SO TIGHT When the study of the permeability of water and solutes was extended from single cells to epithelia, it seemed natural to assume that in these preparations permeation occurs across cell membranes and not through the intercellular space (KoefoedJohnsen and Ussing, 1953; Palay and Karlin, 1959; Miller, 1960; Farquhar and Palade, 1961; Peachey and Rasmussen, 1961; Kaye and Pappas, 1962a,b; Muir and Peters, 1962). In fact, the suggestion that the occluding junction constitutes an essentially tight seal (Bonnet, 1895; Zimmerman, 1911) was supported by the demonstration that the diffusion of macromolecules that can be detected by transmission electron microscopy, such as hemoglobin, is stopped exactly at the level of these junctions (Miller, 1960; Kaye and Pappas, 1962a,b) (Figure 1.4). However, it was later found that while plasma membranes have pores with a radius of around 4 Å (Sidel and Solomon, 1957; Paganelli and Solomon, 1957; Goldstein and Solomon, 1960), epithelia like the intestinal mucosa have pores with radii of some 30 to 40 Å (Lindemann and Solomon, 1962), suggesting that water and small solutes do not cross the epithelium through a transcellular route but through an extracellular one. Those permeability studies had their electrical counterpart. Until the early 1960s, attention was focused on epithelia with large short-circuit currents (in the order of 100 µA · cm–2) and high electrical resistances (above 1500 Ω · cm2). Typically, these epithelia were mounted as flat sheets between two Lucite chambers and studied for several hours until their short-circuit current and electrical resistance decreased, at which moment they were supposed to have exhausted their metabolic resources and were consequently discarded. On the contrary, epithelia like those of the small intestine or the gallbladder exhibit no current and have a comparatively low resistance from the beginning of the experiment, and were considered to be too delicate to withstand dissection and mounting. It was the reign of what Jared M. Diamond called “the paradigm of tight epithelia,” as opposed to “leaky” ones. Nevertheless, in spite of these “disadvantages,” leaky epithelia did show a robust ability to transport water and solutes in vitro (Diamond, 1962; 1968; 1971; Diamond and Wright, 1969; Moreno, 1975a,b). The electrical resistance across leaky epithelia is much lower (around 20 to 80 Ω·cm2) than the resistance across the plasma membrane of their cells (several

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FIGURE 1.3 Freeze-fracture replica of the epithelium of a mouse small intestine, showing the belt of junctional strands separating the apical (upper right) from the lateral side (bottom left). (Courtesy of Prof. A. Martínez-Palomo.)

thousand Ω · cm2) (Lundberg, 1957; Windhager et al., 1966; Hoshi and Sakai, 1967; Boulpaep, 1971; Boulpaep and Seely, 1971; Fromter and Diamond, 1972; Frömter, 1972; Frizzell and Schultz, 1972), indicating that current circumvents the transcellular route. It was eventually concluded that the leakiness of these epithelia is by

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FIGURE 1.4 Transmission electron microscopy of two adjacent cells from the epithelium of a mouse mammary gland. Lanthanum hydroxide (black) added to the basolateral side freely diffuses through the intercellular space, until it reaches the apical end and is stopped by the tight junction. Notice that the apical side (above) is free from the marker.

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no means an abnormal condition resulting from dissection and mounting, but an essential property due to a large extracellular permeation route (Diamond et al., 1970). Those observations prompted studies with extracellular markers, which indicated that small hydrophilic solutes could penetrate through the junctional complex. There was an upper limit to the size of solutes permitted to traverse the TJ, adding to the growing speculation that the barrier contained channels of defined size. When cells are experimentally shrunk by hypertonic media, the pathway is dilated so that relatively large molecules can also permeate across the epithelium (Ussing, 1965; 1971; Whittembury and Rawlins, 1971; Machen et al., 1972; Tisher and Yarger, 1973; Martínez-Palomo and Erlij, 1973). It was additionally observed that the presence of hypertonic media on the mucosal side caused the formation of blisters in the junctions (DiBona and Civan, 1972; 1973; Wade et al., 1973). While the interpretation of this observation remains in dispute, it did suggest both that water and solutes traverse the TJ contacts and that the contacts were points of cell-to-cell adhesion. Eberhard Frömter (1972) applied an electric current across the Necturus gallbladder, and mapped with a glass microelectrode the points of current flow over the apical border, thus identifying the intercellular space as the route of low resistance. When the TJ barrier is characterized by its overall electrical resistance, the measurement reflects the conductance for the principal ions in the solution, usually Na+ and Cl–. It was subsequently discovered, using methods described in Chapter 4, that the TJ could discriminate among different ions. Most displayed a preferential permeability for cations and even a small discrimination between cations of similar size and charge density, such as Na+ and K+ (Moreno, 1975a; Powell, 1981). Investigators observed that as the solution pH is lowered there was a point at which the TJ switches preference from cations to anions (Wright and Diamond, 1968). The specific conductance for different ions and this so-called isoelectric point of TJ varied among epithelia (Powell, 1981). Together these observations led several highly prescient investigators to speculate that the TJ contained aqueous protein-lined channels of defined size. The biochemistry of the amino acid side chains and the proteins lining these channels might explain the charge selectivities. These channels probably differ among epithelia (Wright and Diamond, 1968). Further insight into the molecular basis for these properties would have to wait almost 40 years. This combination of electrical, permeability, and electron microscopy studies demonstrated the importance of the paracellular route limited by the TJ. Since this route may account for up to 90% of the total movement of substances across some epithelia, it is now obvious that the name of “tight” junction is somewhat misleading. However, a century of usage has consecrated this nomenclature. Chapter 4 reviews current information on TJ permeability to ions and water and Chapters 10 and 22 the probable molecular basis.

1.3 WHAT IS A TIGHT JUNCTION? In freeze-fracture replicas the TJ appears as a flat meshwork of anastomosing filaments grouped in a narrow belt, which surrounds the cell on the basolateral side

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at the outermost limit of the intercellular space (see Figures 1.2 and 1.3). This structure suggested that each strand behaves as a resistive element: the larger the number of filaments, the higher the electrical resistance of the TJ (Claude and Goodenough, 1973). However, when the electrical resistance of the paracellular route of different epithelia is plotted against the number of strands in their TJs, it is observed that the increase in resistance with each additional strand is not linear — as expected from the addition of resistors in series — but exponential, suggesting that strands are spanned by flickering pores (Claude, 1978) and that each segment of strand is electrically isolated from neighboring segments (see Figure 1.2) (Cereijido et al., 1988). In Chapter 5, González-Mariscal and her colleagues offer a review of the relationship between structure and function of the TJ. The molecular nature of the TJ strands was for several decades in dispute. The extreme tightness of some TJs and their tubular appearance in freeze-fracture images led some investigators to propose that each strand is a cylindrical micelle in which the polar heads of phospholipid molecules are oriented toward the axis (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982). Subsequent studies on the polarized distribution of lipid molecules did not seem to support this view, as discussed in Chapter 14. Since inhibitors of protein synthesis, such as cycloheximide and puromycin, impair the development of the TJ (Cereijido et al., 1978b; 1981; Griepp et al., 1983), an alternative hypothesis was that each strand of the TJ consists of a row of proteins. Consistent with this, when cell samples were fractured without prior fixation in glutaraldehyde, the strands appeared as rows of distinct 10-nM particles, visually reminiscent of the particles formed by several well-characterized transmembrane proteins. Ultimately, the molecular nature of the strands appears now to have been resolved in a spectacular series of publications from the Tsukita laboratory, in which two types of transmembrane proteins were characterized within the strands. These discoveries are reviewed by Mitic and Van Itallie in Chapter 10. These proteins are occludin (Furuse et al., 1993; Ando-Akatsuka et al., 1996) and a large family of proteins called the claudins (Furuse et al., 1998a; Morita et al., 1999). Of these, the claudins appear to be the major structural and functional elements of the strands. They can form strands when expressed in fibroblasts and influence the electrical resistance when overexpressed in cultured epithelial cells (Furuse et al., 1998b; Inai et al., 1999; McCarthy et al., 2000). Individual claudins show highly restricted patterns of tissue expression consistent with a potential role in creating cell-specific differences in ion and solute permeability (Rahner et al., 2001). Southwood and Gow, in Chapter 33, provide further support for this function in describing the results of deleting claudin-11 from mice though homologous recombination, and Choate et al., in Chapter 22, describe the clinical phenotype of humans with mutations in claudin-16 (paracellin-1). The latter studies strongly suggest claudin-16 is a Mg2+selective, or at least cation-selective, channel in the TJs of renal tubules. The strands of the TJ afford the main resistive element; yet this function is dependent on the presence of other intercellular junctions, principally the cadherinbased adherens or intermediate junctions (see Figure 1.2). Although both occludin and the claudins show homophilic intercellular adhesion, antibodies that specifically

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block cadherin result in the widening of the space between the cells and in the opening of the TJ. The weight of present evidence is consistent with the idea that cadherin initiates intracellular signaling events required to establish and maintain the TJ and apical-basolateral cell polarity. General themes in cell junctions and adhesion are reviewed in Chapter 6, the relationship between development of cell polarity and establishment of TJ in Chapters 7 and 8, and the special case of de novo development of TJs in the mammalian embryo in Chapter 13. The structural integrity of the TJ and its degree of tightness are affected by the status of the actin cytoskeleton (see Chapters 12 and 15) and vary in response to intracellular signals involving protein kinase C, phospholipase C, adenylate cyclase, calmodulin, nonreceptor tyrosine kinases, and G protein receptors (see Chapter 17). Further insight into these events has come in the last 10 years with the discovery and characterization of a very large number of proteins under the strand contacts (Chapter 11). This cytoplasmic plaque contains numerous signaling proteins as well as scaffolding proteins that organize the junction and physically couple the strand proteins to actin. The exact role of all these signaling proteins is still unresolved; yet the implication is that they play a part in adjusting the permeability of the paracellular route to a variety of physiological conditions (Chapters 15 through 17). Because of the complex network of mechanisms affecting the TJ, this structure is highly dynamic, to the point that it may disassemble temporarily to allow the passage of leukocytes (Chapter 29), may suffer drastic changes with age, or may be present only during specific stages of development, such as in the myocardium of the fetal heart (Navaratnam, 1987). TJs within the salt glands of fish can even adjust their tightness in response to salinity (Chapter 20).

1.4 TIGHT JUNCTIONS AND APICAL/BASOLATERAL POLARITY Vectorial transport of substances across epithelia is due to the asymmetric distribution of pumps, carriers, channels, and receptors in the plasma membrane of their cells. This asymmetry is often attributed to the barrier constituted by the TJ, which would prevent diffusion and mixing of membrane components over the whole cell surface. However, even though the TJ may help to prevent mixing of membrane molecules that are already distributed asymmetrically, in particular when these are free to diffuse as in the case of lipids (see Chapter 14), it remains unclear to what degree it participates in achieving such a distribution. Indeed, while there is abundant evidence for polarization of membrane components achieved in the presence of wellestablished tight junctions, there are also clear examples of changes in polarization that occur by endocytosis and exocytic fusion of membrane proteins that elude the fence offered by the TJ (Figure 1.5). The recognition that some cell types, such as neurons, lack TJs yet clearly have a polarized distribution of plasma membrane components contributed to doubt that the TJ is actively involved in generating polarity. However, the identity of several of the recently described cytoplasmic plaque proteins suggests this possibility must be revisited. Among these is a protein called ASIP, which is the homologue of a protein found in the nematode Caenorhabditis elegans

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FIGURE 1.5 Membrane proteins may achieve an apical/basolateral polarity in spite of the TJ: (a) Na, K-ATPase (circles) is not polarized in a renal epithelial cell cultured in Ca-free medium. Ca2+ addition triggers the formation of TJ and traps some Na, K-ATPases on the apical (wrong) side. The cell then removes apical Na, K-ATPases (open arrow) and inserts new enzyme in the basolateral membrane (arrow) until its typical polarity is achieved. (b) An epithelial cell from the intestinal mucosa first exhibits aminopeptidase N in its basolateral (wrong) membrane, but gradually displaces this enzyme toward the apical pole. (c) When G protein of VSV virus is fused to the apical (wrong) membrane using liposomes, the cell removes it from this location and reinserts it in the basolateral membrane. (d) Receptors occupied by IgG are removed from the basolateral position and transferred to the opposite pole of the cell. (e) In a thyroid cell of a follicle suspended in medium without collagen or serum, the apical side faces the medium. Upon addition of collagen or serum the cell reverses its polarity and relocates its TJ.

where it is required for defining early cell polarity in the embryo (Izumi et al., 1998). Others, such as the Sec6/8 complex (Grindstaff et al., 1998), RAB 3B (Weber et al., 1994), and VAP33 (Lapierre et al., 1999) may play roles in the targeting of the vesicles delivering plasma membrane proteins that show a polarized distribution on

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one or the other side of the TJ. Chapters 7 and 8 review the mechanisms responsible for establishing and maintaining cell polarity and the interrelationship between TJs and polarity.

1.5 BIOSYNTHESIS AND ASSEMBLY OF THE TIGHT JUNCTION Several epithelial cell lines are able to establish TJs in culture. Much of the recent information about TJs comes from study of such epithelial model systems (see Figure 1.5). When cells are harvested with trypsin-EDTA, they lose their junctions, but they resynthesize and reassemble them in a few hours on replating the cells at confluence in the presence of Ca2+. When cultures are made on permeable supports (Misfeldt et al., 1976; Cereijido et al., 1978a b; 1988; Gonzalez-Mariscal et al., 1985), the process of junction formation can be followed through: (1) the development of transepithelial electrical resistance; (2) the decrease in permeability to extracellular markers such as inulin and mannitol; (3) transmission and freezefracture electron microscopy using ruthenium red or horseradish peroxidase; and (4) immunofluorescent techniques to observe the distribution of junction-associated proteins. Cell culture models have been used extensively to study intracellular signals that alter TJs (Chapter 17). Extracellular materials can also alter the assembly and permeability characteristics of the TJ. The wide range of substances capable of affecting the TJ is reviewed in Chapter 16. Some of these are physiological and others not. Curiously, proteases such as trypsin and pronase may induce de novo formation of TJs in natural epithelia (Orci et al., 1973; Metz et al., 1977), as well as in epithelial cell lines (Polak-Charcon et al., 1978). This phenomenon is discussed in Chapter 21. A striking example of cyclic assembly and disassembly of TJs within the body is observed in epithelia of the mammary gland, and this is discussed in Chapter 18.

1.6 NEW ROLES IN SIGNALING Until very recently, the principal function attributed to the TJ was that of a paracellular barrier. Although it was appreciated that cellular signaling pathways could regulate assembly and barrier properties, it was not generally thought that the TJ itself might regulate cellular signals, gene transcription, or the state of cellular differentiation. This viewpoint is changing. Among the cytoplasmic plaque proteins ZO-1, ZO-2, and ZO-3 are members for the MAGUK (membrane-associated guanylate kinase) family (Anderson, 1996). Other members of this protein family are tumor suppressors (Woods and Bryant, 1991) or are required for developmental cell fate decisions in invertebrates (Kaech et al., 1998). Alternative RNA splicing of ZO-2 has been observed in several human cancers (Chlenski et al., 2000), ZO-1 binds a transcription factor capable of regulating differentiation-specific genes (Balda and Matter, 2000), and expression of mutant forms of ZO-1 in cultured epithelial cells represses epithelial marker genes and increases tumorgenic cell behaviors (Reichert et al., 2000). Although at present these are poorly understood events, their implication

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is that TJs participate in the regulation of differentiation. Such a role should not be surprising given that other cell–cell and cell–substrate junctions are well known to participate in these events.

1.7 TIGHT JUNCTIONS IN SPECIAL SITUATIONS TJs play special roles in some systems, and these have systemically been reviewed in individual chapters. Thus, Chapter 28 is devoted to TJs between Sertoli cells. These TJs suffer dramatic changes with age, can be traversed by germ cells, and contribute to form a secluded compartment where the necessary conditions for spermatogenesis are achieved. Novel properties of TJs in the eye are reviewed in Chapter 30. TJs are not only found in epithelia, but also between endothelial cells of blood vessels (Robertson, 1960; Muir and Peters, 1962). As in the case of epithelia, the comparison of the relatively low permeability of the plasma membrane with the relatively high one of the capillary wall suggested long ago that most of the transendothelial flux of water and small solutes occurs through the intercellular space (Landis, 1934; Wilbrant, 1946). The TJ also limits this pathway, with perhaps the only exception being the endothelium of microvessels in hemopoietic tissues (Tavassoli and Shaklai, 1979). The tightness of endothelial TJs may be very low, as in the spleen and endocrine glands, or very high, as in the brain and the retina. The number and arrangement of the strands in endothelia also varies from arteries and veins to small vessels (Simionescu et al., 1975; 1976). Unique aspects of the endothelial blood–brain barrier are reviewed in Chapter 19. As mentioned above, in spite of their tightness and fine selectivity, under certain circumstances TJs can be traversed by whole germ cells. They may also be traversed by leukocytes migrating toward the site of infection. This process seems to be quite regulated, as the seal is reestablished after the leukocyte reaches the opposite side. The intrinsic mechanism of this phenomenon is not well understood at present, but the information available is reviewed in Chapter 29. TJs may even be found between cells that are neither epithelial nor endothelial, such as those of the glia (Gray, 1961; Peters, 1962), muscle fibers (Karrer, 1960a,b; Dewey and Barr, 1962), and fibroblasts (Davis and James, 1962), and may even be present between two regions of the same cell (see Chapter 33).

1.8 THE ROLE OF TIGHT JUNCTIONS IN HUMAN DISEASE Disruption of the TJ barrier contributes to disease principally by increasing the backdiffusion of ions, solutes, and water across transporting epithelia thereby reducing the electrical and osmotic gradients that drive absorption and secretion. In endothelia breakdown of the TJ accounts for inappropriate water movements, causing tissue swelling. Pathological consequences are specific to each organ and these are reviewed for the kidney (Chapter 25), the intestine (Chapter 26), the liver (Chapter 27), and the eye (Chapter 30). Insights are also emerging about specific pathological effectors for the TJ, including bacterial toxins (Chapter 23 and 24). A

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very significant recent insight into how TJs are involved in pathology comes from recognition that mutations in claudin-16 (Paracellin-1) are the basis for failure of paracellular magnesium transport through TJs in the kidney (Chapter 22). It seems entirely likely that other human diseases or disease predispositions will result from mutations in other claudins or other TJ proteins. Although disruption of the TJ can contribute to pathology, it seems possible that the barrier could also be manipulated for therapeutic purposes, such as to enhance the delivery of medicinal compounds. This possibility is explored for both the general case of how researchers in the pharmaceutical industry analyze paracellular drug transport (Chapter 31) and the case of a specific substance that may reversibly open TJs and enhance drug absorption (Chapter 32). The understanding of how the TJ participates in specific disease states and the possibility for therapeutic manipulation of the TJ will certainly grow with the understanding of its molecular structure and regulation.

1.9 CONCLUDING REMARKS Almost a century and a half after it first attracted the attention of light microscopists, the TJ is no longer considered a static, almost inert seal, whose only role is that of a mechanical barrier to the passage of substances. Today, the “kiss” of the TJ observed by transmission electron microscopy appears to be the tip of an iceberg, where adhesion molecules form selective paracellular channels attached to a cytoplasmic plaque of scaffolding and regulator proteins. Together they work to create barriers with differing properties and responding to physiological stimuli and situations. New roles for the TJ include a possible role in signaling, influencing gene transcription, and serving as an active zone for vesicle targeting and active generation of cell polarity. Understanding of the molecular basis for all these properties is growing rapidly. The first edition of Tight Junctions a decade ago was the first attempt to bring together a broad range of current information on TJs. The goal was to assemble the wisdom of experts in a wide range of biologic areas, using different approaches to study TJs. However, the rapid growth of knowledge and the expanding range of methods used to study the TJ now threaten to fragment the field. Consequently, the current goal is to reunify and focus inquiry by assembling opinions for this updated volume that cover the expanding breath of knowledge about TJs while encouraging the contributors to make their writing approachable and relevant for readers from different disciplines. The editors hope this edition will be valuable to students and researchers and that it will provide a broadly synthetic and useful compendium. We extend gratitude to our colleagues who again helped to define the subjects and create the limits, and also contributed to this volume. We hope their chapters, both updated and new to this edition, will continue to encourage and expand our knowledge and understanding of TJs. Above all, we thank Mrs. Dotty Franco for her courteous yet efficient insistence in keeping our authors on schedule and maintaining the organization of this project. Thanks are also given to Prof. Adolfo Martínez-Palomo of the Center of Research and Advanced Studies (CINVESTAV) for kindly providing the micrographs of

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Figures 1.1, 1.3, and 1.4 and for providing the cover image for the book. Our own work in the subject was supported by the National Research Council (M.C.), and the National Institutes of Health of the United States (M.C. and J.M.A.).

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Schaffer, J. 1927. Das Epitheligewebe. von Mollendorff, W., Ed. Handbuch der mikeroskopische Anatomie des Menschen. Part 1. Julius Springer, Berlin, 35. Sidel, V. W. and Solomon, A. K. 1957. Entrance of water into human red cells under an osmotic pressure gradient. J. Gen. Physiol., 41:243. Simionescu, M., Simionescu, N., and Palade, G. E. 1975. Segmental differentiation of cell junctions in the vascular endothelium. The microvasculature. J. Cell Biol., 67:863. Simionescu, M., Simionescu, N., and Palade, G. E. 1976. Segmental differentiations of cell junctions in the vascular endothelium. Arteries and veins. J. Cell Biol., 68:705. Staehelin, L. A. 1973. Further observations of the fine structure of freeze-cleaved tight junctions. J. Cell Sci., 13:763–786. Staehelin, L. A. 1975. A new occludens-like junction linking endothelial cells of small capillaries (probably venules) of rat jejunum. J. Cell Sci., 18:545. Staehelin, L. A., Mukherjee, T. M., and Williams, A. W. 1969. Freeze-etch appearance of the tight junctions in the epithelium of small and large intestine of mice. Protoplasma, 67:165. Tavassoli, M. and Shaklai, M. 1979. Absence of tight junctions in endothelium of marrow sinuses: possible significance for marrow cell egress. Br. J. Haematol., 41:303. Tisher, C. C. and Yarger, W. E. 1973. Lanthanum permeability of the tight junctions (zonula occludens) in the renal tubule of the rat. Kidney Int., 3:238. Ussing, H. H. 1965. Relationship between osmotic reactions and active sodium transport in frog skin epithelium. Acta Physiol. Scand., 63:141. Ussing, H. H. 1971. Introductory remarks to discussion on active transport of salts and water in living tissues. Philos. Trans. R. Soc. B, 262:85. Wade, J. B., Revel, J. P., and DiScala, V. 1973. Effect of osmotic gradient on intercellular junctions of the toad bladder. Am. J. Physiol., 224:407. Weber, E., Berta, G., Tousson, A., St. John, P., Gree, M. W., Gopalokrishnam, U., Jilling, T., Sorscher, E. J., Elton, T. S., Abrahamson, D. R., and Kirk, K. L. 1994. Expression and polarization of a Rab3 isoform in epithelial cells. J. Cell Biol., 125:583–594. Whittembury, G. and Rawlins, F. A. 1971. Evidence of a paracellular pathway for ion flow in the kidney proximal tubule: electronmicroscopic demonstration of lanthanum precipitate in the tight junction. Pflugers Arch., 330:302. Wilbrant, W. 1946. Physiologie der Zellund kapillar Permeabilitat. Helv. Med. Acta, 13:143. Windhager, E. E., Boulpaep, E. L., and Giebisch, G. 1966. Proc. 3rd International Congress on Nephrology, Vol. 1, Karger, New York, 35. Woods, D. F. and Bryant, P. J. 1991. The discs-large tumor suppressor gene of Drosophila encodes a guanylate kinase homolog localized at septate junctions. Cell, 66:451–464. Wright, E. M. and Diamond, J. M. 1968. Effects of pH and polyvalent cations on the selective permeability of gall-bladder epithelium to monovalent ions. Biochim. Biophys. Acta, 163:57–74. Zimmerman, J. W. 1911. Zur Morphologie der Epithelzellen der Saugetierniere. Arch. Mikrosk. Anat. Entwicklungsmech., 78:199.

2

Ultrastructure and Immunolabeling of the Tight Junction Eveline E. Schneeberger and Robert D. Lynch

CONTENTS 2.1 2.2 2.3

Introduction ....................................................................................................19 Components of the Junctional Complex .......................................................20 Ultrastructure of the Tight Junction and Immunolocalization of Tight Junction Proteins ............................................................................................21 2.4 Morphology of Tight Junctions in Freeze-Fracture Replicas and Immunolocalization of Tight Junction Proteins in Tight Junction Strands .....23 2.5 Tight Junctions and Their Lipid Environment ..............................................26 2.6 Ultrastructure of Developing Epithelial Tight Junctions ..............................30 2.7 Structure–Function Correlations ....................................................................32 2.8 Summary ........................................................................................................32 References................................................................................................................33

2.1 INTRODUCTION Epithelia form vital cellular barriers that generate and maintain a very different fluid and solute composition between adjacent tissue compartments. This requires that epithelial cells be selectively permeable to molecules that are either secreted or absorbed, a process that is mediated by asymmetrically distributed cellular transport mechanisms via a route known as the transcellular pathway. However, solutes may also diffuse between cells, via the paracellular route, whose selective permeability is regulated, in part, by the tight junction (TJ) (Schneeberger and Lynch, 1992; Anderson and Van Itallie, 1995). The TJ forms a circumferential, selective seal in the intercellular space and maintains a polarized distribution of subclasses of lipids (Simons and van Meer, 1988) and integral membrane proteins (e.g., water channels, ion pumps, and ion channels) (Yeaman et al., 1999) in the plane of the plasma membrane. The TJ, therefore, has at least two functions: (1) it forms a regulated barrier in the intercellular space and (2) it maintains a fence between apical and basolateral domains of the plasma membrane. More recent data suggest that some components of the TJ may also be important in the regulation of cell growth and 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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differentiation (Tsukita et al., 1993; Balda and Matter, 2000; Reichert et al., 2000). This chapter examines the ultrastructural organization of the TJ in light of recent new data pertaining to its composition and its lipid environment. The principal emphasis will be on the TJs of epithelial cells; however, reference will also be made to those between endothelial cells.

2.2 COMPONENTS OF THE JUNCTIONAL COMPLEX Light microscopic observations made over a century ago revealed the presence of a barlike area of condensation, measuring approximately 1 to 2 µm in depth near the apex of epithelial cells where they are joined; it was designated the “terminal bar” (Bizzozero, 1870). Although its composition and functional significance were not known, it was assumed to have a sealing function. It was only after the introduction of the electron microscope that it became possible to obtain a more-detailed description of the terminal bar. In 1963, Farquhar and Palade, using electron microscopy of thin sectioned tissues, recognized that the terminal bar in fact consists of at least three components, which they defined as the “junctional complex” (Farquhar and Palade, 1963). The most apical member of this complex is the TJ or zonula occludens and just below that is the intermediate junction or zonula adhaerens. The desmosome or macula adhaerens, which functions as a focal adhesion site, is the most basal member of the complex (Figure 2.1). Gap junctions or nexi may be closely associated

FIGURE 2.1 Diagrammatic representation of the junctional complex in a pair of cuboidal epithelial cells. The TJ is the most apical member of the junctional complex. Its network of TJ strands forms a continuous, gasket-like band, sealing the intercellular space. Below the tight junction is the E-cadherin-rich intermediate junction that also forms a continuous band around the perimeter of the cell. The desmosomes form discrete disklike adhesion sites. The gap junction, while formally not included in the junctional complex, and usually found below it, is in fact present within the network of the TJs of arterial and venous endothelial cells as well as in fetal epithelial TJs.

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with TJs of some cells, including those of arterial and venous endothelium, but are not considered as part of the junctional complex. The gap junction has the important function of providing regulated communicating channels between the cytoplasmic compartments of adjacent cells.

2.3 ULTRASTRUCTURE OF THE TIGHT JUNCTION AND IMMUNOLOCALIZATION OF TIGHT JUNCTION PROTEINS At low magnification the TJ appears as an apical zone, between 0.1 to 0.7 µm in depth, where adjacent plasma membranes are in close apposition (Figure 2.2a). At a higher magnification, the TJ is seen as discrete punctate sites of close membrane contact where the outer lipid leaflets of the adjacent plasma membrane appear to merge and the extracellular space is obliterated (Figure 2.2b). These correspond to the complex of strands in the TJ network that are revealed by freeze-fracture techniques (discussed below). A major advance in the understanding of the protein composition of the TJ sealing elements was achieved with the identification of occludin (Furuse et al., 1993) and later the claudin family of TJ proteins (Furuse et al., 1998; Simon et al., 1999) and their immunolocalization on ultrathin sections to TJ strands. Interestingly, both are tetra-span proteins with cytoplasmic N and C termini and two extracellular loops, but they lack any homology to each other. The claudin family now has grown to 24 members, and immunolocalization studies indicate that some of these have specific cell and tissue distributions (Simon et al., 1999; Tsukita and Furuse, 2000), as well as unique functions (Gow et al., 1999). A third protein, the junctional adhesion molecule (JAM), belonging to the immunoglobulin superfamily, has also been immunolocalized to both epithelial and endothelial tight junctions (MartinPadura et al., 1998), where it interacts with ZO-1, cingulin, and occludin (Bazzoni et al., 2000). Whether JAM is an integral part of the TJ strands themselves, however, has not been determined. It is important in promoting monocyte migration through the TJ. Given the unique permeability properties of TJs in different tissues, it is likely that additional members of the claudin family and/or other TJ-related proteins will be discovered. Aggregates of fine fibrillary material are present on the cytoplasmic side of the TJ (Figure 2.2a). These, when incubated with S1 fragments of heavy meromyosin, become decorated with characteristic S1 arrowheads indicating that actin filaments are associated with the TJ (Madara, 1987). In addition to actin filaments, immunolocalization studies indicate that there are a growing number of cytoplasmic proteins that are localized to a submembranous plaque at the tight junction (Table 2.1). Some of these are unique to the tight junction (cingulin, ZO-2, ZO-3) (Jesaitis and Goodenough, 1994; Haskins et al., 1998; Cordenonsi et al., 2000), others are localized to both the TJ and the nucleus (ZO-1, symplekin, ZONAB) (Gottardi et al., 1996; Keon et al., 1996; Balda and Matter, 2000) and still others are not restricted to TJs (AF-6, VAP-33, monomeric and heterotrimeric GTPases, PKC) (Weber et al., 1994; Zahraoui

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FIGURE 2.2 (a) Epithelial junctional complex in the lung of a fetal lamb. At the level of the TJ the intercellular space is focally obliterated by punctate close approximations of the plasma membrane of adjacent cells. At the intermediate junction (IJ) the adjacent plasma membranes are slightly approximated and the intercellular space contains electron-dense E-cadherin rich fibrillary material. In the desmosome (D), the intercellular space is filled with electron-dense desmoglein-rich fibrillary material. On the cytoplasmic side of each of these junctions there are variably dense aggregates of cytoplasmic fibrillary material that represent the cytoplasmic plaques associated with each of these junctions. (b) Higher magnification of the TJ shows several punctate sites of close membrane approximation. Original magnification (a) ×75,000, (b) ×100,000. (From Schneeberger, E. E. et al., J. Cell Sci., 32:307–324, 1978. With permission.)

et al., 1994; Stuart and Nigam, 1995; Yamamoto et al., 1997; Saha et al., 1998; Lapierre et al., 2000). Several of these TJ-associated proteins (ZO-1, ZO-2, ZO-3, and cingulin) form links between integral TJ proteins and the actin cytoskeleton (Fanning et al., 1998; Itoh et al., 1999; Wittchen et al., 1999; Cordenonsi et al., 2000). The contractile activity of the actin cytoskeleton may be responsible, in part, for regulating TJ permeability (Hecht et al., 1996; Turner et al., 1999). Recent studies suggest that other TJ plaque proteins may contribute to the regulation of cell growth and differentiation by participating in the transduction of signals from the surface of the cell to the nucleus (Balda and Matter, 2000).

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TABLE 2.1 TJ-Associated Cytoplasmic Proteins Protein ZO-1 ZO-2

Properties MAGUKa (Nucleus and TJ) MAGUK

ZO-3 Cingulin Symplekin 7H6 AF6 ASIP

MAGUK Phosphoprotein Nucleus and TJ of epithelia Phosphoprotein Ras interacting protein Atypical PKCb iso-type-specific interacting protein Vesicle targeting protein Vesicle targeting protein Serine kinase ErbB-2 transcription factor Monomeric GTPase Monomeric GTPase Heterotrimeric G protein Protein kinase

Sec6/8 VAP-33 ZAK ZONAB Rab 3B Rab 13 Gαi2 PKC a b

Ref. Stevenson et al., 1986 Beatch et al., 1996; Gumbiner et al., 1991; Jesaitis and Goodenough, 1994 Haskins et al., 1998 Citi et al., 1988; Cordenonsi et al., 2000 Keon et al., 1996 Zhong et al., 1993 Yamamoto et al., 1997 Izumi et al., 1998 Grindstaff et al., 1998 Lapierre et al., 2000 Balda et al., 1996 Balda and Matter, 2000 Weber et al., 1994 Zahraoui et al., 1994 Saha et al., 1998 Stuart and Nigam, 1995

MAGUK = membrane-associated guanylate kinase. PKC = protein kinase C.

2.4 MORPHOLOGY OF TIGHT JUNCTIONS IN FREEZEFRACTURE REPLICAS AND IMMUNOLOCALIZATION OF TIGHT JUNCTION PROTEINS IN TIGHT JUNCTION STRANDS With the development of freeze-fracture techniques, it became possible to examine the unique TJ strand network in greater detail. Briefly, working under vacuum and at low temperatures (–115°C), freeze fracturing involves making a platinum-carbon replica of the fractured surface of either fixed or unfixed frozen cells. Characteristically, at low temperatures the fracture plane tends to proceed along the weak, interior hydrophobic plane of cell membranes, thereby exposing integral transmembrane proteins, including those of the TJ. The freshly fractured surface is covered with a film of platinum and strengthened with a layer of carbon. The cells underlying the replica are then removed by the application of sodium hypochlorite, and the cleaned replica, representing the three-dimensional relief of the fractured surface, is examined in the electron microscope (Figure 2.6) (Bullivant, 1973). Viewed by this technique, the TJ forms a continuous network of interconnected strands on the protoplasmic fracture face (P face) and complementary empty grooves on the exoplasmic fracture face (McCarthy et al., 1996) (E face) that encircles the apical zone of the lateral membranes of epithelial cells (Figure 2.3). Fixation may

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FIGURE 2.3 Freeze-fracture replica of a TJ between the ciliated cells of human nasal mucosa. The TJ forms a complex network of continuous TJ strands on the P face (P) and particle-free grooves on the E face (E). Original magnification ×61,000.

have a significant effect on how TJ strands partition during freeze fracturing. In well-fixed epithelia, the strands are generally intact and continuous and tend to remain on the P face. This has been attributed to strengthened cross-linking between tight junctional and cytoskeletal proteins by aldehyde fixatives. In nonfixed or lightly fixed tissues, the TJ consists of rows of particles or short segments of strands, some of which partition onto the E face, leaving gaps in the strands on the P face (Revel, 1982). To avoid the pitfall of attributing low transepithelial electrical resistance to such TJ strand defects, analysis of mirror-image double replicas will usually show that the segment missing on the P face is present on the E face (Schneeberger et al., 1978). It should be noted, however, that regardless of the degree of fixation, the TJ particles of endothelial cells partition preferentially onto the E face (Simionescu and Simionescu, 1976a,b; Schneeberger, 1981). This suggests that fixation alone cannot explain the difference in partitioning behavior of TJ proteins in different tissues. An interesting new light was shed on this phenomenon when Furuse introduced cDNAs of claudins 1, 2, 3 into fibroblasts, cells that do not form TJs. Freeze-fracture replicas revealed that these cells indeed formed TJ strands; however, the TJ particles of claudins 1 and 3 preferentially partitioned onto the P face, whereas those of claudin2 partitioned onto the E face (Furuse et al., 1999). This difference in partitioning behavior of these three claudins cannot be directly attributed to a difference in their ability to bind to the ZO proteins, since claudins 1 to 8 have all been shown to bind to ZO-1, ZO-2, and ZO-3 (Itoh et al., 1999). Furthermore, the composition of the TJ strands of tissue epithelial and endothelial cells is more complex. A given claudin may oligomerize with occludin and/or with other types of claudins present in the

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FIGURE 2.4 (a) TJ and gap junctions between adjacent pulmonary arterial endothelial cells of a rat lung. In the area of these junctions the adjacent membranes are closely apposed and the two types of junctions cannot be resolved. (b) Freeze-fracture replica of a TJ similar to that shown in Figure 2.4a. Note that the TJ particles (arrowhead) preferentially partition onto the E face (E) leaving particle-poor ridges (arrow) on the P face (P). Within the TJ network, foci of tightly packed gap junction particles are present. Original magnification (a) ×82,000, (b) ×75,000. (From Schneeberger, E. E., Circ. Res., 49:1101–1111, 1981. With permission.)

TJs, a property that may also affect the manner in which these proteins partition into the hemi-leaflets of the plasma membrane during freeze fracturing. TJs between arterial endothelial cells are unusual in that they are intimately associated with numerous gap junction particles (Figure 2.4a) that are tightly packed within the meshwork of the TJ strands. These gap junction particles partition onto the P face, whereas the particulate TJ strands partition preferentially onto the E face (Figure 2.4b). TJs between venous endothelial cells are also associated with gap junctions, but these are considerably smaller and fewer in number (Simionescu and Simionescu, 1975; Schneeberger, 1981). Gap junctions are largely absent from the TJs of capillary endothelial cells. Instead, they are simple in structure consisting of two to three particulate, sparsely interconnected strands that preferentially partition onto the E face (Simionescu and Simionescu, 1975; Schneeberger and Karnovsky, 1976). Immunogold labeling techniques have been used to advantage to localize proteins by electron microscopy in ultrathin sections (Roth, 1983). This has enabled investigators to immunolocalize ZO-1 and cingulin to the cytoplasmic side of the tight junction (Stevenson et al., 1986) and occludin (Furuse et al., 1993) or claudin-1 and claudin-5 to the TJs of epithelium and endothelium of the lung (Figure 2.5a and b). However, this approach does not provide definitive information regarding which

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FIGURE 2.5 (a) TJ between two alveolar type I cells of a mouse lung, immunogold labeled for claudin-1. The alveolar space is at the top and the adjacent capillary lumen is at the bottom of the figure. (b) Endothelial TJ of a pulmonary vein in a mouse lung immunogold labeled for claudin-5. The gold particles line up specifically in the area of the TJ; their association with TJ strands cannot, however, be determined. Original magnification ×59,000.

proteins are integral to the TJ strands. For this purpose, morphologists have devised strategies that combine immunogold labeling with freeze-fracture techniques. By modifying earlier methods (Fujimoto and Pinto da Silva, 1992), Fujimoto developed a fracture labeling technique that made it possible to localize integral TJ proteins to TJ strands (Fujimoto, 1995). Briefly, this involves either rapidly freezing (Fujimoto, 1995) or lightly fixing cells or tissue with 1% paraformaldehyde (Mitic et al., 1999). The sample is then freeze-fractured and coated with platinum/carbon. The apolar domains of the fractured, split membranes with their contained membrane proteins are protected by the platinum/carbon layer from the sodium dodecyl sulfate (SDS)-induced micelle formation (Figure 2.6). By contrast, the SDS detergent readily solubilizes the nonfractured membranes and the cytoplasmic components. Applied antibodies that bind to native epitopes or epitope tags engineered onto the cytoplasmic domains of the membrane protein of interest are then detected using the appropriately coated gold particles and the immunolabeled replica is examined by electron microscopy. Since most of the available anti-occludin and anti-claudin antibodies are directed against epitopes in the cytoplasmic C terminus, they are well suited for detection of these proteins in TJ strands (Figure 2.7a and b) (McCarthy et al., 2000). Furthermore, the use of antibodies generated in different species makes it possible to label more than one protein in TJ strands of a given replica (Furuse et al., 1999).

2.5 TIGHT JUNCTIONS AND THEIR LIPID ENVIRONMENT While a growing number of proteins are associated with and/or form TJ strands, a role for plasma membrane lipids in the structure and/or function of TJ should not be overlooked. The effect of changing the membrane lipid environment on TJ

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FIGURE 2.6 Diagrammatic representation of the fracture labeling process. The lipid bilayer is preferentially cleaved at low temperatures along the weak hydrophobic plane of the lipid bilayer (A). The cleaved lipid bilayers, including the short segment of uncleaved membrane (B), are covered with a layer of platinum and carbon (C). Note that the exposed hydrophobic acyl chains are protected by the platinum/carbon layer, but the short, uncleaved segment of the membrane is not (C). Fragments of cytoplasmic proteins and uncleaved membranes are solubilized by SDS and the integral membrane proteins of interest are labeled by immunogold techniques (D).

structure and function (Lynch et al., 1993; Stankewich et al., 1996; Francis et al., 1999), and the presence of lipid-modified signaling molecules found at the TJ (Jou et al., 1998) have stimulated renewed interest in the role of plasma membrane lipids in TJ biology (Nusrat et al., 2000). Analyses of purified membrane fractions show that there are no qualitative differences in the lipid composition between apical and basolateral domains of the plasma membrane of epithelial cells (Simons and van Meer, 1988). However, quantitative differences between these two domains are substantial and this difference is maintained, in part, by the presence of the TJ. At the level of the plasma membrane lipid bilayer, the observed asymmetry of the phospholipid composition is generated by the activity of translocases (Bevers et al., 1999). An interesting feature of eukaryotic

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FIGURE 2.7 Freeze-fractured epithelial Madin–Darby canine kidney (MDCK) cell TJs immunogold labeled for (a) occludin and (b) for claudin-1. As seen by this technique, the gold particles are aligned with the TJ strands. Original magnification ×70.000.

cells is that as much as 90% of the cholesterol of a cell resides in the plasma membrane (Lange et al., 1989). Lateral phase separation of lipids, together with either the spontaneous or externally induced clustering of selected membrane proteins within the plasma membrane, produces cholesterol/sphingolipid-rich subdomains, that are typically Triton X-100 insoluble at 4°C (Brown and London, 1998; 2000; Smart et al., 1999). These have been variably termed detergent-insoluble glycolipid-enriched membranes (DIGs), detergent-resistant membrane fragments (DRMs), or glycolipid-enriched membranes (GEMs). The liquid-ordered structure of these rafts is largely due to the long, saturated acyl chains of the contained sphingolipids that facilitate their close packing with cholesterol. The tetra-span TJ proteins occludin and the claudins are presumed to form highly oligomerized complexes within the TJ strands which, in turn, reside in the cholesterol-enriched lipid environment of the plasma membrane. That changes in this lipid environment profoundly affect both the structure and barrier function of the TJ was shown by a series of studies in which cell cholesterol content was reduced either by treating cells with Lovastatin, a 3-hydroxy-3-methylglutaryl-CoA (HMG CoA) reductase inhibitor, or by rapidly removing cholesterol using methyl-β-cyclodextrin (Lynch et al., 1993; Stankewich et al., 1996; Francis et al., 1999). In Lovastatintreated MDCK cells, formation of TJs, assessed by measuring transepithelial electrical resistance (TER) occurred more rapidly and reached higher peak values following a calcium switch than did their untreated controls (Lynch et al., 1993). On the other hand, in confluent MDCK cell monolayers with well established TJs,

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reduction of plasma membrane cholesterol content by 60 to 75% using methyl-βcyclodextrin resulted in an initial rise in TER followed later by a decline below initial values (Stankewich et al., 1996; Francis et al., 1999). To examine the distribution of cholesterol relative to the TJ and the structural changes in TJ morphology induced by the removal of cholesterol, MDCK cells were treated with filipin, a polyene antibiotic that forms complexes with cholesterol (Miller, 1984). In control monolayers, numerous filipin–cholesterol complexes formed on the apical and basolateral membranes, but were absent from the network of the TJ (Figure 2.8a). That filipin is excluded not only from TJs, but also from

FIGURE 2.8 (a) Freeze-fracture replica of control MDCK cells that were treated with the polyene antibiotic filipin that forms complexes with cholesterol. Note that there are numerous filipin–cholesterol complexes on both the apical and basolateral cell membranes, but no complexes are detected within the meshwork of the TJ. (b) In control MDCK cells the TJ strands preferentially partition onto the P face (P). (c) Freeze-fracture replica of MDCK cells that were treated for 2 h with 10 mM methyl-β-cyclodextrin to remove 60% of the cholesterol of the cell. They were then reacted with filipin and freeze-fractured. Note the marked reduction in the number of filipin–cholesterol complexes. (d) Following this treatment, the TJ strands are particulate and tend to partition onto the E face (E). Original magnification ×62,500. (From Francis, S. A. et al., Eur. J. Cell Biol., 78:473–484, 1999. With permission.)

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intermediate and gap junctions has been extensively documented in native epithelia (Stetson and Wade, 1983). Thus, the absence of filipin–cholesterol complexes in the TJ network does not indicate the absence of cholesterol in this region of the plasma membrane (see below). When up to 60% of the cells’ cholesterol was removed, using methyl-β-cyclodextrin, the number of filipin–cholesterol complexes was markedly reduced in both apical and basolateral plasma membrane domains (Figure 2.8b) and no filipin–cholesterol complexes were observed in the TJ network. Interestingly, in these cholesterol-depleted epithelial cells, the TJ strands partitioned preferentially onto the E face rather than the P face (compare Figures 2.8c and d). In addition, phalloidin-labeled F-actin formed prominent aggregates at the tricellular regions and occludin content appeared to be reduced in the TJs of these cells, as determined by immunofluorescence microscopy (Francis et al., 1999). Thus, removal of significant amounts of cholesterol from the plasma membrane may alter interactions of the underlying cytoskeletal proteins with integral TJ proteins, thereby affecting their partitioning between E and P fracture faces and decreasing TER. Since a number of lipid-modified signaling proteins have been localized to the TJ (Mitic and Anderson, 1998), there has been increased interest in the composition of the membrane lipid environment of the TJ. A recent study suggests that the TJ protein occludin resides in a raftlike cholesterol/glycolipid-enriched environment. Following extraction in cold Triton X-100, both hyperphosphorylated occludin and ZO-1 were found in the low-density fraction of a sucrose density gradient that was also enriched for cholesterol and sphingomyelin (Nusrat et al., 2000). The presence of the cytoplasmic protein ZO-1 in this fraction is somewhat surprising; presumably ZO-1 remained bound to occludin during the isolation procedure. Ongoing studies indicate that claudin-1, like occludin and caveolin-1, is also found in the low-density fraction of an Opti-prep gradient (Francis, McCarthy, Lynch, and Schneeberger, unpublished observations).

2.6 ULTRASTRUCTURE OF DEVELOPING EPITHELIAL TIGHT JUNCTIONS Numerous studies dealing with the ultrastructure of TJs in a variety of fetal tissues were conducted more than two decades ago (Montesano et al., 1975; Revel and Brown, 1975; Schneeberger et al., 1978; Suzuki and Nagano, 1979). Only those structural features that are common to the TJs from a variety of fetal tissues will be summarized here. The reader is referred to the lucid studies of Fleming described in Chapter 13 in which the expression of known TJ proteins are carefully mapped during early embryonic development. The earliest morphological evidence of TJ formation is the appearance of a series of raised segments of particle-poor ridges on the P face. This is followed later by the appearance of short, isolated segments of rows of particles on ridges on the P face. The E face grooves likewise contain numerous TJ particles, suggesting that binding of the transmembrane TJ proteins to the underlying actin cytoskeleton is tenuous and not yet fully developed (Figure 2.9). In some of these developing TJs,

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FIGURE 2.9 Freeze-fracture replica of an epithelial TJ from a fetal lamb lung at 39 days of gestation (full term is at 120 days). Note the particulate appearance of the TJ strands and grooves on the P and E face, respectively. A disconnected segment of a TJ strand is indicated (arrow). Two small desmosomes are shown (arrowhead). Original magnification ×82,500. (From Schneeberger, E. E. et al., J. Cell Sci., 32:307–324, 1978. With permission.)

FIGURE 2.10 At 10 h after adding the mitogen-activated protein kinase kinase (MEK1) inhibitor PD98059 to a confluent monolayer of Ras-transformed MDCK cells, short segments of TJ strands appear on the P face. Note the gaps on the P face strands (arrowheads) indicating that some of the particles have partitioned onto the E face (not shown). Original magnification ×62,500. (From Chen, Y. H. et al., Mol. Biol. Cell., 11:849–862, 2000. With permission.)

aggregates of particles, resembling gap junction particles, are observed within the meshwork of the TJ strands. These disappear as the TJ matures. A similar sequence of TJ strand formation was recapitulated in a recently described series of experiments using Ras-transformed MDCK cells (Chen et al., 2000). These transformed epithelial cells adopt a fibroblastic morphology, they do not express occludin, claudin-1, or ZO-1, and they lack TJs. After treatment with the mitogen-activated protein kinase (MEK1) inhibitor PD98059, which blocks the activation of mitogen-activated protein kinase (MAPK), occludin, claudin-1 and ZO-1 were recruited to the cell membrane. TJs were assembled first as short segments of particulate strands (Figure 2.10) that merged and became interconnected to form a TJ network, by a series of steps similar to that observed in fetal tissues.

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2.7 STRUCTURE–FUNCTION CORRELATIONS The complexity, the number of parallel strands, and the measured transepithelial resistance can vary widely among TJs in different tissues. This led Claude and Goodenough (1973) to postulate that there was a direct relationship between the number of parallel TJ strands and the transepithelial electrical resistance. Further analysis of the data, however, indicated that the number of parallel strands in the TJ was proportional to the logarithm of the transepithelial electrical resistance. Based on this observation, Claude suggested that the TJ strands are not inert barriers, but instead contain pores that have a finite probability of fluctuating between an open and closed conformation (Claude, 1978). The physical characteristics of these so-called TJ pores have been deduced from physiological studies. These indicate, with few exceptions, that TJs in most epithelia are cation selective (Diamond and Wright, 1969; Cereijido et al., 1981; Powell, 1981), a property that can be altered by the passage of current (Finn and Bright, 1978) and changes in pH (Diamond, 1978). Such observations implicate the presence of fixed negative charges, e.g., COO– within the TJ pore. The integrity of the TJ barrier is calcium dependent; removal of calcium results, indirectly, in the disassembly of the TJ strands (MartínezPaloma et al., 1980). This effect involves Ca2+-dependent E-cadherin adhesion at the adherens junction (Takeichi, 1991), and/or other intracellular Ca2+-dependent proteins, and excludes the claudins that mediate cell–cell adhesion activity via a calcium-independent mechanism (Kubota et al., 1999). Proteases may be involved in the pathological alterations of the TJ barrier. For example, although resistant to trypsin (Lynch et al., 1995), both occludin and claudins are susceptible to digestion by a cysteine protease identified in the fecal pellets of house mites (Wan et al., 1999). This may facilitate the access of allergens to underlying cells of the immune system, including dendritic cells, thus stimulating an allergic immune response (Gong et al., 1992). Metalloproteinases, implicated in cell migration including that of tumor cells, are also able to cleave occludin and presumably also claudins, thereby compromising the barrier function of TJs (Wachtel et al., 1999).

2.8 SUMMARY With the application of molecular techniques, identification of proteins integral to the TJ strands as well as those associated with the cytoplasmic TJ plaque has been greatly facilitated. It is likely that other TJ-related proteins will be identified. The task at hand will be to determine at the molecular level which integral TJ proteins contribute to the formation of the pores predicted by physiological studies, how these proteins are oligomerized to form TJ strands, and to determine how the plaque proteins regulate the activity of these pores. The role of membrane lipids in the activity of the TJ and the role of lipid-modified signaling molecules remains a fertile area of investigation. Finally, emerging data strongly suggest that, in addition to its traditional fence and gate functions, the TJ plays an important role in the regulation of cell growth and differentiation.

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REFERENCES Anderson, J. M. and Van Itallie, C. M. 1995. Tight junctions and the molecular basis for regulation of paracellular permeability. Am. J. Physiol., 269, G467–G475. Balda, M. S. and Matter, K. 2000. The tight junction protein ZO-1 and an interacting transcription factor regulate ErbB-2 expression. EMBO J., 19, 2024–2033. Balda, M. S., Anderson, J. M., and Matter, K. 1996. The SH3 domain of the tight junction protein ZO-1 binds to a serine protein kinase that phosphorylates a region C-terminal to this domain. FEBS Lett., 399, 326–332. Bazzoni, G., Martinez-Estrada, O. M., Orsenigo, F., Cordenonsi, M., Citi, S., and Dejana, E. 2000. Interaction of junctional adhesion molecule with the tight junction components ZO-1, cingulin and occludin. J. Biol. Chem., 275, 20520–20526. Beatch, M., Jesaitis, L. A., Gallin, W. J., Goodenough, D. A., and Stevenson, B. R. 1996. The tight junction protein ZO-2 contains three PDZ (PSD-95/Discs-Large/ZO-1) domains and an alternatively spliced region. J. Biol. Chem., 271, 25723–25726. Bevers, E. M., Comfurius, P., Dekkers, D. W., and Zwaal, R. F. 1999. Lipid translocation across the plasma membrane of mammalian cells. Biochim. Biophys. Acta, 1439, 317–330. Bizzozero, T. 1870. Osservazion: sulla struttura degli epitheli pavimentosi stratificati. Rend. Ist. Lomb. Accad. Sci. Lett. Ser. II, 3, 675. Brown, D. A. and London, E. 1998. Functions of lipid rafts in biological membranes. Annu. Rev. Cell Dev. Biol., 14, 111–136. Brown, D. A. and London, E. 2000. Structure and function of sphinolipid and cholesterolrich membrane rafts. J. Biol. Chem., 275, 17221–17224. Bullivant, S. 1973. Freeze-Etching and Freeze-Fracturing. Springer-Verlag, New York. Cereijido, M., Meza, I., and Martínez-Paloma, A. 1981. Occluding junctions in cultured epithelial monolayers. Am. J. Physiol., 140, C96–C102. Chen, Y. H., Qun, L., Schneeberger, E. E., and Goodenough, D. A. 2000. Restoration of tight junction structure and barrier function by down-regulation of the mitogen activated protein kinase pathway in Ras-transformed Madin–Darby canine kidney cells. Mol. Biol. Cell, 11, 849–862. Citi, S., Sabanay, H., Jakes, R., Geiger, B., and Kendrick-Jones, J. 1988. Cingulin, a new peripheral component of tight junctions. Nature, 333, 272–276. Claude, P. 1978. Morphological factors influencing transepithelial permeability: a model for the resistance of the zonula occludens. J. Membr. Biol., 39, 219–232. Claude, P. and Goodenough, D. A. 1973. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. J. Cell Biol., 58, 390–400. Cordenonsi, M., D’Atri, F., Hammar, E., Parry, D. A. D., Kendrick-Jones, J., Shore, D., and Citi, S. 2000. Cingulin contains globular and coiled-coil domains and interacts with ZO-1, ZO-2, ZO-3 and myosin. J. Cell Biol., 147, 1569–1581. Diamond, J. M. 1978. Channels in epithelial cell membranes and junctions. FASEB J., 37, 2638–2644. Diamond, J. M. and Wright, E. M. 1969. Biological membranes: the physical basis of ion and non-electrolyte permeability. Annu. Rev. Physiol., 31, 581–646. Fanning, A. S., Jameson, B. J., Jesaitis, L. A., and Anderson, J. M. 1998. The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J. Biol. Chem., 273, 29745–29753. Farquhar, M. G. and Palade, G. E. 1963. Junctional complexes in various epithelia. J. Cell Biol., 17, 375–412.

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Finn, A. L. and Bright, J. 1978. The paracellular pathway in toad urinary bladder: permselectivity and kinetics of opening. J. Membr. Biol., 44, 67–83. Francis, S. A., Kelly, J. M., McCormack, J. M., Rogers, R. A., Lai, J., Schneeberger, E. E., and Lynch, R. D. 1999. Rapid reduction of MDCK cell cholesterol by methyl-βcyclodextrin alters steady state transepithelial electrical resistance. Eur. J. Cell Biol., 78, 473–484. Fujimoto, K. 1995. Freeze-fracture replica electron microscopy combined with SDS digestion for cytochemical labeling of integral membrane proteins. Application to the immunogold labeling of intercellular junctional complexes. J. Cell Sci., 108, 3443–3449. Fujimoto, K. and Pinto da Silva, P. 1992. Fracture-flip Triton-X reveals the cytoplasmic surface of human erythrocyte membranes. Acta Histochem. Cytochem., 25, 255–263. Furuse, M., Hirase, T., Itoh, M., Nagafuchi, A., Yonemura, S., Tsukita, S., and Tsukita, S. 1993. Occludin: a novel integral membrane protein localizing at tight junctions. J. Cell Biol., 123, 1777–1788. Furuse, M., Fujita, K., Fujimoto, K., and Tsukita, S. 1998. Claudin 1 and 2: novel integral membrane proteins localizing at tight junctions with no sequence similarity to occludin. J. Cell Biol., 141, 1539–1550. Furuse, M., Sasaki, H., and Tsukita, S. 1999. Manner of interaction of heterogeneous claudin species within and between tight junction strands. J. Cell Biol., 147, 891–903. Gong, J. L., McCarthy, K. M., Telford, J. R., and Schneeberger, E. E. 1992. Intraepithelial airway dendritic cells: a distinct subset of pulmonary dendritic cells obtained by microdissection. J. Exp. Med., 175, 797–807. Gottardi, C. J., Arpin, M., Fanning, A. S., and Lovard, D. 1996. The junction-associated protein, zonaula occludens-1, localizes to the nucleus before the maturation and during the remodeling of cell-cell contacts. Proc. Natl. Acad. Sci. U.S.A., 93, 10779–10784. Gow, A., Southwood, C. M., Li, J. S., Pariali, M., Riordan, G. P., Brodie, S. E., Danias, J., Bronstein, J. M., Kachar, B., and Lazzarini, R. A. 1999. CNS myelin and sertoli cell tight junction strands are absent in Osp/claudin-11 null mice. Cell, 99, 649–659. Grindstaff, K. K., Yeaman, C., Anandasabapathy, N., Hsu, S. C., Rodriguez-Boulan, E., Scheller, R. H., and Nelson, W. J. 1998. Sec6/8 complex is recruited to cell–cell contacts and specifies transport vesicle delivery to the basal-lateral membrane in epithelial cells. Cell, 93, 731–740. Gumbiner, B., Lowenkopf, T., and Apatira, D. 1991. Identification of a 160-kDa polypeptide that binds to the tight junction protein ZO-1. Proc. Natl. Acad. Sci. U.S.A., 88, 3460–3464. Haskins, J., Lijie, G., Wittchen, E. S., Hibbard, J., and Stevenson, B. R. 1998. ZO-3, a novel member of the MAGUK protein family found at the tight junction, interacts with ZO-1 and occludin. J. Cell Biol., 141, 199–208. Hecht, G., Robinson, B., Koutsouris, A., Tripuraneni, J., Lorimer, D. D., Nowack, G., Guerriero, J. V., Elson, E. L., and Lanerolle, P. D. 1996. Expression of the catalytic domain of myosin light chain knase increases paracellular permeability. Am. J. Physiol., 271, C1678–C1684. Itoh, M., Furuse, M., Morita, K., Kubota, K., Saitou, M., and Tsukita, S. 1999. Direct binding of three tight junction-associated MAGUKs, ZO-1, ZO-2 and ZO-3, with the COOH termini of claudins. J. Cell Biol., 147, 1351–1363. Izumi, Y., Hirose, T., Tamai, Y., Hirai, S., Nagashima, Y., Fujimoto, T., Tabuse, Y., Kemphues, K. J., and Ohno, S. 1998. An atypical PKC directly associates and colocalizes at the epithelial tight junction with ASIP, a mammalian homologue of Caenorhabditis elegans polarity protein PAR-3. J. Cell Biol., 143, 95–106.

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Jesaitis, L. A. and Goodenough, D. A. 1994. Molecular characterization and tissue distribution of ZO-2, a tight junction protein homologous to ZO-1 and the Drosophila discs-large tumor suppressor protein. J. Cell Biol., 124, 949–961. Jou, T. S., Schneeberger, E. E., and Nelson, W. J. 1998. Structural and functional regulation of tight junctions by RhoA and Rac1 small GTPases. J. Cell Biol., 142, 101–115. Keon, B. H., Schafer, S., Kuhn, C., Grund, C., and Franke, W. W. 1996. Symplekin, a novel type of tight junction plaque protein. J. Cell Biol., 134, 1003–1018. Kubota, K., Furuse, M., Sasaki, H., Sonoda, N., Fujita, K., Nagafuchi, A., and Tsukita, S. 1999. Ca2+-independent cell-adhesion activity of claudins, a family of integral membrane proteins localized at tight junctions. Curr. Biol., 9, 1035–1038. Lange, Y., Swaisgood, M. H., Ramos, B. V., and Steck, T. L. 1989. Plasma membranes contain half the phospholipid and 90% of the cholesterol and sphingomyelin in cultured human fibroblasts. J. Biol. Chem., 264, 3786–3793. Lapierre, L. A., Tuma, P. L., Navarre, J., Goldenring, J. R., and Anderson, J. M. 1999. VAP33 localizes to both an intracellular vesicle population and with occludin at the tight junction. J. Cell Sci., 112, 3723–3732. Lynch, R. D., Tkachuk, L. J., Ji, X., Rabito, C. A., and Schneeberger, E. E. 1993. Depleting cell cholesterol alters calcium-induced assembly of tight junctions by monolayers of MDCK cells. Eur. J. Cell Biol., 60, 21–30. Lynch, R. D., Tkachuk, L. J., McCormack, J. M., McCarthy, K. M., Rogers, R. A., and Schneeberger, E. E. 1995. Basolateral but not apical application of protease results in a rapid rise of transepithelial electrical resistance and formation of aberrant tight junction strands in MDCK cells. Eur. J. Cell Biol., 66, 257–267. Madara, J. L. 1987. Intestinal absorptive cell tight junctions are linked to cytoskeleton. Am. J. Physiol., 253, C171–C175. Martinez-Paloma, A., Meza, I., Beaty, G., and Cereijido, M. 1980. Experimental modulation of occluding junctions in a cultured transporting epithelium. J. Cell Biol., 87, 746–754. Martin-Padura, I., Lostaglio, S., Schneemann, M., Williams, L., Romano, M., Fruscella, P., Panzeri, C., Stoppacciaro, A., Ruco, L., Villa, A., Simmons, D., and Dejana, E. 1998. Junctional adhesion molecule, a novel member of the immunoglobulin superfamily that distributes at intercellular junctions and modulates monocyte transmigration. J. Cell Biol., 142, 117–127. McCarthy, K. M., Skare, I. B., Stankewich, M. C., Furuse, M., Tsukita, S., Rogers, R. A., Lynch, R. D., and Schneeberger, E. E. 1996. Occludin is a functional component of the tight junction. J. Cell Sci., 109, 2287–2298. McCarthy, K. M., Francis, S. A., McCormack, J. M., Lai, J., Rogers, R. A., Skare, I. B., Lynch, R. D., and Schneeberger, E. E. 2000. Inducible expression of claudin-1-myc but not occludin-VSVG results in aberrant tight junction strand formation in MDCK cells. J. Cell Sci., 113, 3387–3398. Miller, R. G. 1984. The use and abuse of filipin to localize cholesterol in membranes. Cell Biol. Int. Rep., 8, 519–535. Mitic, L. L. and Anderson, J. M. 1998. Molecular architecture of tight junctions. Annu. Rev. Physiol., 60, 121–142. Mitic, L. L., Schneeberger, E. E., Fanning, A. S., and Anderson, J. M. 1999. Connexinoccludin chimeras containing the ZO-binding domain of occludin localize at MDCK tight junctions and NRK cell contacts. J. Cell Biol., 146, 683–693. Montesano, R., Friend, D. S., Perrelet, A., and Orci, L. 1975. In vivo assembly of tight junctions in fetal rat liver. J. Cell Biol., 1975, 310–319.

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Nusrat, A., Parkos, C. A., Verkade, P., Foley, C. S., Liang, T. W., Innis-Whitehouse, W., Eastburn, K. K., and Madara, J. L. 2000. Tight junctions are membrane microdomains. J. Cell Sci., 113, 1771–1781. Powell, D. W. 1981. Barrier function of epithelia. Am. J. Physiol., 241, G275–G288. Reichert, M., Mueller, T., and Hunziker, W. 2000. The PDZ domains of zonula occludens-1 induce an epithelial to mesenchymal transition of Madin–Darby canine kidney I cells. J. Biol. Chem., 275, 9492–9500. Revel, J. P. 1982. Intercellular Communication. Josiah Macy Foundation, New York. Revel, J. P. and Brown, S. S. 1975. Cell junctions in development. Cold Spring Harbor Symp. Quant. Biol., 40, 433–355. Roth, J. 1983. The Colloidal Gold Marker System for Light and Electron Microscopic Cytochemistry. Academic Press, New York. Saha, C., Nigam, S. K., and Denker, B. M. 1998. Involvement of Gαι2 in the maintenance and biogenesis of epithelial cell tight junctions. J. Biol. Chem., 273, 21629–21633. Schneeberger, E. E. 1981. Segmental differentiation of endothelial intercellular junctions in intra-acinar arteries and veins of the rat lung. Circ. Res., 49, 1102–1111. Schneeberger, E. E. and Karnovsky, M. J. 1976. Substructure of intercellular junctions in freeze-fractured alveolar-capillary membranes of mouse lung. Circ. Res., 38, 404–411. Schneeberger, E. E. and Lynch, R. D. 1992. Structure, function and regulation of cellular tight junctions. Am. J. Physiol., 262, L647–L661. Schneeberger, E. E., Walters, D. V., and Olver, R. E. 1978. Development of intercellular junctions in the pulmonary epithelium of the foetal lamb. J. Cell Sci., 32, 307–324. Simionescu, M. and Simionescu, N. 1975. Segmental differentiations of cell junctions in the vascular endothelium. The microvasculature. J. Cell Biol., 67, 863–886. Simionescu, M. and Simionescu, N. 1976a. Segmental differentiations of cell junctions in the vascular endotheium. Arteries and veins. J. Cell Biol., 68, 705–723. Simionescu, N. and Simionescu, M. 1976b. Galloylglucoses of low molecular weight as mordant in electron microscopy. J. Cell Biol., 70, 608–621. Simon, D. B., Lu, Y., Choate, K. A., Velazquez, H., Al-Sabban, E., Praga, M., Casari, G., Bettinelli, A., Colussi, G., Rodriguez-Soriano, J., McCredie, D., Milford, D., Sanjad, S., and Lifton, R. P. 1999. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science, 285, 103–106. Simons, K. and van Meer, G. 1988. Lipid sorting in epithelial cells. Biochemistry, 27, 6197–6202. Smart, E. J., Graf, G. A., McNiven, M. A., Sessa, W. C., Engelman, J. A., Scherer, P. E., Okamoto, T., and Lisanti, M. P. 1999. Caveolins, liquid-ordered domains and signal transduction. Mol. Cell. Biol., 19, 7289–7304. Stankewich, M. C., Francis, S. A., Vu, Q. U., Schneeberger, E. E., and Lynch, R. D. 1996. Alterations in cell cholesterol content modulate of calcium induced tight junction assembly by MDCK cells. Lipids, 31, 817–828. Stetson, D. L. and Wade, J. B. 1983. Ultrastructural characterization of cholesterol distribution in toad bladder using filipin. J. Membr. Biol., 74, 131–138. Stevenson, B. R., Siliciano, J. D., Mooseker, J. D., and Goodenough, D. A. 1986. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J. Cell Biol., 103, 755–766. Stuart, R. O. and Nigam, S. K. 1995. Regulated assembly of tight junctions by protein kinase C. Proc. Natl. Acad. Sci. U.S.A., 92, 6072–6076. Suzuki, F. and Nagano, T. 1979. Morphogensis of tight junctions in the peritoneal mesothelium of the mouse embryo. Cell Tissue Res., 198, 247–260.

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Takeichi, M. 1991. Cadherin cell adhesion receptors as a morphogenetic regulator. Science, 251, 1451–1455. Tsukita, S. and Furuse, M. 2000. Pores in the wall: claudins constitute tight junction strands containing aqueous pores. J. Cell Biol., 149, 13–16. Tsukita, S., Itoh, M., Nagafuchi, A., Yonemura, S., and Tsukita, S. 1993. Submembranous junctional plaque proteins include potential tumor suppressor molecules. J. Cell Biol., 123, 1049–1053. Turner, J. R., Angle, J. M., Black, E. D., Joyal, J. L., Sacks, D. B., and Madara, J. L. 1999. PKC-dependent regulation of transepithelial resitance: roles of MLC and MLC knoase. Am. J. Physiol., 277, C554–C562. Wachtel, M., Frei, K., Fontana, A., Winterhalter, K., and Gloor, S. M. 1999. Occludin proteolysis and increased permeability in endothelial cells through tyrosine phosphatase inhibition. J. Cell Sci., 112, 4347–4356. Wan, H., Winton, H. L., Soeller, C., Tovey, E. R., Gruenert, D. C., Thompson, P. J., Stewart, G. A., Taylor, G. W., Garrod, D. R., Cannell, M. B., and Robinson, C. 1999. Der p 1 facilitates transepithelial allergen delivery by disruption of tight junctions. J. Clin. Invest., 104, 123–133. Weber, E., Berta, G., Tousson, A., St. John, P., Green, M. W., Gopalokrishnan, U., Jilling, T., Sorscher, E. J., Elton, T. S., and Abrahamson, D. R. 1994. Expression and polarized targeting of a Rab 3 isoform in epithelial cells. J. Cell Biol., 125, 583–594. Wittchen, E. S., Haskins, J., and Stevenson, B. R. 1999. Protein interactions at the tight junction. J. Biol. Chem., 274, 35179–35185. Yamamoto, T., Harada, N., Kano, K., Taya, S., Canaani, E., Matsuura, Y., Mizoguchi, A., Ide, C., and Kaibuchi, K. 1997. The Ras target AF-6 interacts with ZO-1 and serves as a peripheral component of tight junctions in epithelial cells. J. Cell Biol., 139, 785–795. Yeaman, C., Grindstaff, K. K., and Nelson, W. J. 1999. New perspectives on mechanisms involved in generating epithelial cell polarity. Physiol. Rev., 79, 73–98. Zahraoui, A., Joberty, G., Arpin, M., Fontaine, J. J., Hellio, R., Tavitian, A., and Louvard, D. 1994. A small rab GTPase is distributed in cytoplasmic vesicles in nonpolarized cells but colocalizes with the TJ marker ZO-1 in polarized epithelial cells. J. Cell Biol., 124, 101–115. Zhong, Y., Saitoh, T., Minase, T., Sawada, N., Enomoto, K., and Mori, M. 1993. Monoclonal antibody 7H6 reacts with a novel tight junction-associated protein distinct from ZO-1, cingulin and ZO-2. J. Cell Biol., 120, 477–483.

3

Tight Junctions in Invertebrates Nancy J. Lane

CONTENTS 3.1 3.2 3.3

Introduction ....................................................................................................39 Distribution of Tight Junctions among Invertebrates ....................................40 Fine Structural Features of Tight Junctions ..................................................41 3.3.1 In Thin Sections .................................................................................41 3.3.2 In Replicas after Conventional Fixing and Cryoprotection ..............41 3.3.3 In Replicas after Rapid Freezing with No Fixation..........................44 3.3.4 Coexistence with Other Junctions .....................................................45 3.3.5 Cytoskeletal Associations ..................................................................46 3.4 Physiological Roles........................................................................................47 3.4.1 Permeability Barriers .........................................................................47 3.4.2 Cell–Cell Adhesion ............................................................................48 3.5 Models of Invertebrate Tight Junctions .........................................................48 3.6 Invertebrate Groups That Possess Tight Junctions........................................49 3.7 Other Junctions Peculiar to the Invertebrates with Tight Junction-Like Characteristics ................................................................................................49 3.7.1 Smooth Septate Junctions ..................................................................49 3.7.2 Reticular Septate Junctions................................................................51 3.7.3 Retinular Junctions.............................................................................51 3.8 Comparisons between Tight Junctions in the Invertebrates, Lower Chordates, and Vertebrates.............................................................................51 3.9 Assembly of Arthropod Tight Junctions during Development .....................54 3.10 Biochemistry of Invertebrate Tight Junctions ...............................................54 3.11 Conclusions ....................................................................................................55 Acknowledgments....................................................................................................55 References................................................................................................................56

3.1 INTRODUCTION When tight junctions (TJs) were originally observed under the electron microscope (EM), it was assumed they were restricted to vertebrates, since septate junctions, found only in invertebrates, were thought to form the equivalent function to TJs in those organisms, as the morphological basis of the observed permeability barriers. 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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Studies commencing in the 1970s, however, revealed that TJs were in fact to be found in the lower organisms, but that not all phyla possessed them. The majority of TJ-containing tissues in the invertebrates are found within the Arthropoda, in groups such as the arachnids and the insects, where these junctions are responsible for forming blood–brain, blood–eye, and blood–testis barriers. TJs in invertebrates exhibit some differences from those encountered in vertebrate tissues, although they share many common attributes, as is manifest in their thin section and freeze-fracture appearance and the transmembrane migration of their intramembranous particles (IMPs) during junctional assembly. One of the major differences seems to be that very simple tight junctional ridges, combined with tortuous membrane interdigitations, are often found in the tissues of insects rather than the complex network of anastomosing tight junctional ridges that is characteristic of many vertebrate tissues, as well as of arachnid tissues. Their function seems very alike in both groups, however, in that they form the morphological basis of permeability barriers and help maintain cell–cell adherence via junctional membrane-associated cytoskeletal components.

3.2 DISTRIBUTION OF TIGHT JUNCTIONS AMONG INVERTEBRATES TJs exist in the tissues of invertebrates, but only in certain tissues of particular groups. Initially, it was thought that zonulae occludentes, as originally described in 1963 and 1965 in vertebrate epithelia by Farquhar and Palade, were absent in tissues from nonchordate organisms (Satir and Gilula, 1973). Those species possessing these sealing junctions were thought to include ones from all the conventional vertebrate phyla (see Claude and Goodenough, 1973; Staehelin, 1974) together with such lowly chordates as the invertebrate-like tunicates (Lorber and Rayns, 1972; Georges, 1979; Lane et al., 1986). These tunicate TJs have been shown to be highly complex (Martinucci et al., 1988; Burighel et al., 1992; Lane et al., 1994). Until relatively recently, the remaining invertebrate groups appeared to lack any structurally comparable junctions and the physiologically analogous structures responsible for producing permeability barriers were thought to be the septate junctions (Satir and Gilula, 1973; Green et al., 1979; Noirot-Timothée and Noirot, 1980). TJs are identifiable in thin sections as close appositions of adjacent cell membranes, but these can sometimes be confused with gap junctions. The technique of freeze fracturing, however, demonstrates their unequivocal existence and their distribution; such studies showed that definitive TJs occur in the tissues of certain groups of arthropods. Both insects and arachnids, such as the spiders and scorpions, have been found to possess TJs. The first demonstration of such junctions was in the central nervous system (CNS) of cockroaches (Lane and Treherne, 1972). Earlier tracer studies had revealed that a permeability barrier existed to the entry of exogenous molecules (Lane and Treherne, 1970), while physical disruption of the cellular basis of the barrier (Lane and Treherne, 1969; 1970), the outer glial cell layer round the CNS, the perineurium, destroyed this “barrier” effect. When the fine structure of the insect perineurium is examined carefully, the cell borders are found to interdigitate in a highly complex way, with basally positioned punctate tight junctional

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appositions (Lane and Treherne, 1972; Lane, 1972; Lane et al., 1977). The eye, testis, and CNS of insects, which also exhibit a blood–eye (Shaw, 1978), blood–testis (Szollosi and Marcaillou, 1977; Toshimori et al., 1979), and blood–brain barrier (Lane et al., 1977), possess both TJs and septate junctions.

3.3 FINE STRUCTURAL FEATURES OF TIGHT JUNCTIONS 3.3.1 IN THIN SECTIONS TJs were originally found in thin sections of the insect perineurium that ensheathes the CNS and the larger peripheral nerves (Lane, 1972; Lane and Treherne, 1972; 1973). In all cases their presence could be correlated with the presence of a permeability barrier measured both electrophysiologically (Treherne et al., 1970; Treherne and Pichon, 1972) and by impedance to the free entry of tracer molecules (Lane and Treherne, 1972; Lane, 1972). Subsequent studies corroborated the existence of TJs in the CNS of a range of insects (Leslie, 1973; McLaughlin, 1974; Lane et al., 1977; Lane and Swales, 1978a,b; 1979) and further investigations on other tissues, such as the testis and eye, where barriers exist, have all revealed a number of punctate appositions between cells in thin sections. The perineurial layer in the CNS of arachnids, such as in spiders (Lane and Chandler, 1980; Lane, 1981a) and scorpions (Lane et al., 1981), also can be seen to possess TJs in thin sections. The characteristic appearance of TJs in arthropod tissues is essentially similar to that reported for zonulae occludentes in vertebrate material. The membranes of the adjacent cells associated by TJs are fused together in a number of punctate appositions so that the intercellular cleft, normally 10 to 20 nm in width, may be totally obliterated at these points (Figures 3.1 and 3.2). When arthropod tissues are incubated in saline to which electron opaque tracers have been added, the inward penetration of these exogenous substances is restricted. The lateral borders between adjacent cells in many arthropod epithelia are enormously interdigitated and the TJs are at the basal surface of the perineurial cell layer in insects and nearer the outer surface in spiders. In both cases the tracers may be seen to move into the intercellular clefts and are then stopped by the fused membrane appositions (Figures 3.3 and 3.4).

3.3.2 IN REPLICAS AFTER CONVENTIONAL FIXING AND CRYOPROTECTION Freeze-cleaved replicas have an advantage over thin sections for visualizing junctional structures in that en face views of membranes are revealed and the intramembranous patterns assumed by junctional particles may then be studied. The tight junctional particles are aligned in ridges that are shared, by fusion of the plasma membranes of adjacent cells, thereby leading to closure of the intercellular space and sealing of the cleft. There are variations in the degree of complexity of the arrangements of the component intramembranous ridges. In freeze-fracture replicas, fracture face P (P face or PF) is the cytoplasmic or inner membrane half, and fracture

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FIGURES 3.1 AND 3.2 Thin sections of tight junctions from the perineurial sheath of the CNS of a spider. Note that the highly interdigitating glial borders are associated by both TJ (arrows) and gap junctions (GJ). Note the tricellular contact (circle) in Figure 3.1. Original magnification: Figure 3.1, ×112,000; Figure 3.2, ×87,000. FIGURE 3.3 When tissues are incubated in solutions containing electron opaque tracer molecules, the tracers can impregnate the intercellular clefts up the point of the punctate tight junctional appositions, beyond which further penetration is blocked (at arrow). Original magnification: ×146,000. FIGURE 3.4 When the intercellular cleft is infiltrated with tracers, in this case lanthanum, the lines of fusion of adjacent cell membranes, at the tight junctional appositions, are indicated by unstained fibrils (arrows) against the dense, stained background of the intercellular space. Clearly here the lines of fusion are incomplete, so that the tracer can migrate around the discontinuities and move further on into the intercellular cleft. Original magnification: ×142,500.

face E (E face or EF) is the extracellular or outer membrane half. The TJs in arthropods are usually characterized by single intramembranous P face ridges or E face grooves, which are often discontinuous and variable in length. In some cases two or three may lie in parallel with one another, occasionally in fairly close proximity. They are clearly moniliform and hence appear to derive from linear arrays of IMPs, which are fused laterally (Figure 3.5). The degree of this fusion is variable

Tight Junctions in Invertebrates

43

FIGURES 3.5 AND 3.6 Freeze-fracture replicas of arthropod TJs show networks of ridges, which may be either on the P or E face; these are mainly in the form of beadlike, moniliform alignments (Figure 3.5) when fixed. They may, however, in some places take on the appearance of smooth ridges or cylinders (Figure 3.6) depending on the angle of shadowing. The gap junctions (GJ) that are found in arachnids are in the form of plaques of IMPs on the E face (EF) or macular arrays of pits on the P face (PF). Original magnifications: Figures 3.5, ×50,500; Figure 3.6, ×67,000.

and ranges over a spectrum from the distinctly beadlike, to ridges displaying partial fusion, to smooth strands (Figure 3.6). More complex TJs are arranged as a distinct circumferential belt around the cells forming the permeability barriers; they are composed of a network of moniliform ridges on one fracture face, usually the PF, and a complementary interconnecting array of grooves, usually on the EF. These preferential fracturing faces are found consistently in insect and certain arachnid tissues independent of the pretreatment, that is, whether the material is frozen without fixing or after chemical fixation. However, in other arachnid tissues, notably the scorpions, the fracture face varies depending on whether or not the tissue has been fixed; in such cases, the unfixed cryoprotected material reveals the conventional P face ridges and EF grooves,

44

Tight Junctions

whereas fixed, cryoprotected tissue possesses EF ridges. In possessing EF ridges they resemble the tight junctional system found in some vertebrate tissues, notably that in the blood–testis barrier (Nagano and Suzuki, 1976) and the vascular endothelium (Simionescu et al., 1978). TJs are also found between perineurial cells in the imago and adult forms of dipteran flies and are spatially associated with septate junctions both in the CNS proper and in the adult compound eye (Carlson et al., 2000); here many heterocellular TJs also form between glial cells and neurons (Chi and Carlson, 1981; Saint-Marie et al., 1984). When the fracture plane cleaves across a face transition in invertebrate TJs, the PF ridges, although offset, are coincident with the EF grooves, thereby indicating their complementary nature (Figures 3.7 and 3.8). The component IMPs that comprise these fibrils or ridges are about 8-10 nm in diameter like those that make up vertebrate zonulae occludentes. This contrasts with the gap junctional particles, which are somewhat larger (13 nm in diameter), in arthropod tissues and consistently fracture onto the E face. These may be arrayed in plaques or be more loosely aggregated depending on their stage in the assembly process (Figure 3.9).

3.3.3 IN REPLICAS

AFTER

RAPID FREEZING

WITH

NO FIXATION

The TJ fibrils in these freeze-fracture replicas can appear either as a continuous cylinder or as a row of particles that partition to either the P or the E face, consistently leaving a continuous furrow in the complementary fracture face. This appearance of particles, or, alternatively, of continuous cylinders has been attributed to the different conditions used in tissue preparation, such as strength of fixation (van Deurs and Luft, 1979) or, if unfixed and uncryoprotected, freezing rate (Kachar and Reese, 1982). From a careful analysis of unfixed fractured tissue in arthropods examined at high resolution, a substructure in the tight junctional fibrils can sometimes be seen as a periodicity (Figure 3.10). This periodicity suggests the existence of fibril subunits, which appear to be asymmetric (Kachar et al., 1992). It is possible that there may be indirect interaction of fibril components from the cytoplasm into the cytoplasmic halves of the interacting bilayers. In rapidly frozen tissues, the PF and EF structures are always offset with respect to one another at PF/EF transitions (Figure 3.10a), although the offset fibril may be hidden by the fractured membrane face. In this regard they reveal the fracturing characteristics observed by Bullivant (1978) in TJs in fixed tissues from which he constructed the offset-two fibril model for TJs. The tight junctional fibrils of arthropods, after fast freezing, exhibit one of two quite different appearances in freeze-fractured replicas; these vary with the fracture face that is revealed by the cleaving process. This could be interpreted as demonstrating an underlying asymmetry whereby alternative aspects of asymmetric subunit structures in the tight junctional fibrils are selectively exposed, depending on the unpredictable pathway of the fracture plane (Kachar et al., 1992). The intramembranous fibrils themselves are thought to be attached to the cytoskeleton via a hypothetical linking molecule, which could be claudin or occludin.

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45

FIGURES 3.7, 3.8, AND 3.9 Replicas from arachnids showing tight junctional networks making up the blood–brain barrier in spider and scorpion CNS. The E face (EF) grooves are in register with the P face (PF) ridges (arrows in Figure 3.8), but the grooves and ridges are offset with respect to one another (arrows in Figure 3.8 and in insert). The tight junctional grooves and ridges are in intimate spatial association with plaques of gap junctional (GJ) E face connexons (insert, Figure 3.7) or P face pits (Figure 3.7), which may be, in embryonic tissues, in the process of assembling (connexons, at arrows in Figure 3.9) into clusters, apparently directed, in part, into position by the tight junctional fusions (Figure 3.9). Original magnifications: Figure 3.7, ×32,000; Insert, ×70,000; Figure 3.8, ×55,000; Insert, ×112,000; Figure 3.9, ×29,000.

3.3.4 COEXISTENCE

WITH

OTHER JUNCTIONS

In arthropods, TJs coexist with a variety of other junctional types. The most common is the gap or communicating junction. In replicas, gap junctional IMP clusters occur in the interstices of, or close to, the network of ridges that comprises the zonulae occludentes (Lane and Chandler, 1980; Lane et al., 1981) (see Figure 3.7). Since the gap junctional IMPs fracture onto the E face in arthropod tissues (Flower, 1972) and since they are larger than the tight junctional particles (Lane, 1981c), they are readily distinguished as a separate class of IMP. This feature has been extremely

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FIGURE 3.10 Rapidly frozen TJs from the spider, after slamming against a liquid heliumcooled cooper block. Tissue near the periphery (a) shows junctional fibrils in the form of cylinders on the P face (PF) that exhibit a periodic substructure, but as grooves with only occasional irregular, plastic-deformed particles on the E face (EF). Membranes of cells in tissue some distance from the copper block surface are less well preserved, and may show moniliform ridges (as in b). The insert in a shows that the cylinders and grooves are offset with respect to one another. Original magnifications: (a), ×176,000; (b), ×64,000; insert to (a), ×177,000.

useful in developmental studies where stages in the formation of the two categories of junction can clearly be separated, demonstrating that each junctional type originates from a different class of precursor particle (Lane, 1981c) and that the TJs may assemble early, well before the final clustering of gap junctional connexons (see Figure 3.9). TJs also, in certain rare cases, coexist with the septate junctions of arthropods, except for, as far as can be ascertained, the arachnids.

3.3.5 CYTOSKELETAL ASSOCIATIONS In thin sections, the TJs of arthropods can often be seen to be associated with fibrils (Lane, 1991), particularly after tannic acid staining enhancement; these often appear as an unstructured fuzz (Figures 3.11 through 3.13). The nature of these can be established after treatment with the S1 subfragment of heavy meromyosin (HMM); this produces the characteristic arrowhead or corkscrew labeling on these fibrils as is found when actin is bound by HMM (Figure 3.14). In some cases, microtubules lie underneath, at right angles to the arrays. The actin fibrils may be tethered onto these microtubules, as is the case with arthropod septate and intermediate junctions (Lane and Flores, 1988; 1990). The evidence for the intimate relationship of cytoskeleton to the TJs in vertebrate tissues has been summarized (Stevenson et al., 1988) with the suggestion that epithelia regulate paracellular permeability through tensile forces generated via the cytoskeletal components that are found adjacent to the TJs. A comparable situation would appear to exist in invertebrates.

Tight Junctions in Invertebrates

47

FIGURES 3.11, 3.12, AND 3.13 In thin section, the punctate appositions of the arthropod tight junctions often show faint microfibrils (arrows) projecting into the cytoplasm at the points of membrane fusion, but obvious associations with other cytoskeletal elements are not apparent (Figure 3.11). Original magnifications: Figure 3.11, ×105,000; Figure 3.12, ×155,000; Figure 3.13, ×177,000. FIGURE 3.14 When tight junctions are glycerinated and incubated with the S1 subunit of heavy meromyosin, the subunits bind to the actin fibrils projecting from the tight junctional membrane appositions; here (at arrow) they appear as corkscrews, rather than arrowheads, in section, as they are being viewed tangentially or end on, rather than side on; Original magnification: ×207,500.

3.4 PHYSIOLOGICAL ROLES 3.4.1 PERMEABILITY BARRIERS TJs are considered to provide a seal between epithelial cells, thereby producing a diffusion barrier to the intercellular movement of ions and molecules. In arthropods, the TJs that have been observed are rarely found in situations that permit the measurement of transepithelial resistance or the analysis of movement of tracers in both directions across the TJ-bearing cell layers. The CNS in insects is avascular; hence, ions and molecules are only able to gain access to the nerve cells by dint of moving across the sheath of modified glial cells that surround them. There is also a perineurium around the CNS in such arachnids as spiders and scorpions (Lane et al., 1981), but in these organisms it appears that the ganglia are vascularized (see Lane and Treherne, 1980). Blood sinuses and intracerebral channels have been reported in all these arachnids, and their cellular linings, modified glial cells, possess intercellular TJs, like the endothelial junctions in the lining of mammalian CNS blood vessels. Electrophysiological experiments originally demonstrated the insect blood–brain permeability barrier. Tracer studies confirmed this since the entry of exogenous substances was restricted to the perineurial clefts and they did not invade the underlying nervous tissue (Lane and Treherne, 1970; 1972; Lane, 1972). Ionic lanthanum penetrated both septate and gap junctions, so the remaining punctate appositions or

48

Tight Junctions

TJs found between adjacent perineurial cells were presumed to be the morphological basis for the observed physiological barrier. Subsequent freeze-fracture studies revealed that the basal perineurium of insects possesses a simple tight junctional system of ridges and grooves (Lane et al., 1975; 1977; Lane and Swales, 1978a,b). The perineurium in such arachnids as spiders and scorpions provides a blood–brain barrier in the same way as it does in the insect CNS (Lane and Chandler, 1980; Lane et al., 1981). There have been a number of reports on a blood–retinal barrier (Shaw, 1978) in insect compound eyes where complex, vertebrate-like TJs are to be found by freeze fracture (Lane, 1981d; Carlson et al., 2000). A permeability barrier has also been found in the testis of insects. A basal compartment containing the differentiating spermatids has been shown, by using a variety of tracers, to be closed with unequivocal tight junctional ridges lying below septate junctions in the cyst envelope of the silkworm (Toshimori et al., 1979).

3.4.2 CELL–CELL ADHESION Junctions between cells serve in a dual capacity: With TJs, in creating a diffusion barrier by punctate cell-to-cell fusions, the cells are held firmly together both by the junctions and their associated cytoskeleton and by the extensive lateral cellular infoldings, and, in so doing, they maintain the integrity of the tissue in which they occur. In arachnid CNS, TJs form a broad apical girdle, which could be extremely effective as an adhesive device, whereas in insects they are not nearly so extensively distributed. When insect TJs become disrupted in the normal process of early pupation (Lane and Swales, 1978b), the perineurial cells and glia become dissociated from one another fairly promptly.

3.5 MODELS OF INVERTEBRATE TIGHT JUNCTIONS A model has been proposed to account for the features of invertebrate TJs (Lane, 1981a; 1990), notably that for spider TJs and a modification of it for the scorpion occluding junctions (Lane et al., 1981). Although there is a difference in complexity between insect and spider TJs, they both have comparable fracturing characteristics; that is, they tend to have PF ridges and EF grooves in both fixed and unfixed tissues. In scorpions, which have “leaky” TJs, however, the ridges fracture onto the EF in fixed tissue and onto the PF only if the material is unfixed, although in fixed tissues of both scorpions and some spiders they may fracture partly onto the EF and partly onto the PF (Lane et al., 1981). This suggests that there may be local variations in the way that the tight junctional particles are associated with either side of the membranes or laterally with each other. The model favored in vertebrates is the offset double-fibril model (Bullivant, 1978), which also seems appropriate for the arthropod TJs, since the data from replicas (Lane, 1981a; 1990) show the arthropod fibrils to be offset with respect to one another (Figure 3.17).

Tight Junctions in Invertebrates

49

3.6 INVERTEBRATE GROUPS THAT POSSESS TIGHT JUNCTIONS TJs have been reported in the Crustacea, Insecta, and Arachnida. The structures observed are not precisely the same in all cases, and indeed this diversity is a feature of arthropod zonulae occludentes. The evidence derives from several sources: (1) the occurrence in thin sections of punctate appositions between adjacent cell membranes that obliterate the intercellular space, (2) the existence in freeze-fracture replicas of a junctional belt composed of IMPs with complementary grooves, and/or (3) the presence of a permeability barrier where these junctions are found, demonstrated either electrophysiologically or by the impedance of tracer entry. In certain instances in the various arthropod classes, only one or two rather than all these criteria have been shown to be fulfilled. In the arthropods, which exhibit many physiological differences from vertebrate species, TJs have only been found in tissues known to possess permeability barriers: these include blood–brain barriers, blood–testis barriers, and blood–retinal barriers. Thus far, TJs have only been found in tissues in insects, crustacea, spiders, and scorpions, which are the more highly evolved arthropods, and not in the Onychophora (Peripatus) (Lane and Campiglin, 1987), the Acarina (mites and ticks) (Binnington and Lane, 1980), the Myriapoda (centipedes and millipedes) (Dallai et al., 1990), or Xiphosaura (horseshoe crabs) (Lane, 1989). There have been a few reports of TJs in the mollusca, with TJ-like ridges described in the glial cells of opisthobranch gastropods (see references in Lane, 1981b), and apparent singlestranded tight junctional ridges between the adaxonal Schwann cells in the cephalopod nervous system (Villegas et al., 1987; Zwahlen et al., 1988), but there are no physiological data to indicate their functional significance.

3.7 OTHER JUNCTIONS PECULIAR TO THE INVERTEBRATES WITH TIGHT JUNCTION-LIKE CHARACTERISTICS The ability to distinguish true occluding junctions from other junctions that may bear a superficial resemblance to TJs, particularly in replicas, is crucial. There are several other kinds of junction found in arthropods that are not occluding, but which either have been or may be misinterpreted as such. These either have been thought to have a sealing action or are structurally sufficiently similar to TJs in one way or another to have been considered by some investigators as indistinguishable from them (Figure 3.18).

3.7.1 SMOOTH SEPTATE JUNCTIONS The structural features of septate junctions differ dramatically from those of the zonulae occludentes, for they consistently exhibit a regular 15 to 20 nm cleft wherein exist undulating septal ribbons that, cut transversely in thin sections, appear as ladderlike structures (Lane, 1981a). However, after freeze fracturing they possess

50

Tight Junctions

FIGURES 3.15 AND 3.16 Arthropod septate junctions are often mistaken for TJs in freezefracture replicas; their intercellular cleft (C) is 18 to 20 nm wide, rather than nonexistent, as in TJs, but the membrane faces show rows of aligned particles (arrows) that look moniliform or ridgelike in chemically fixed or cryoprotected preparations (Figure 3.15) and which in unfixed, rapidly frozen tissues can also clearly be seen to exhibit ridges made up of individual particles (arrows) on the E face (EF) with complementary grooves on the P face (PF) (Figure 3.16). Original magnifications: Figure 3.15, ×67,000; Figure 3.16, ×109,000.

intramembranous PF particles that lie in rows that sometimes resemble tight junctional IMP rows. The pleated septate junctions are less like TJs, for they have IMPs in rows that are separated from one another by fairly consistent distances; these lie in undulating tracts over the membrane fracture faces and do not form networks (Figure 3.15). Structurally, then, they do not resemble TJs at all in thin sections and very little in freeze-fracture replicas in spite of the similarity of the two in their localization near the apical border of epithelia. However, smooth septate (continuous) junctions are rather a different matter. When these continuous junctions were first described (Noirot and Noirot-Timothée, 1967) they were called zonula continua and attention was drawn to the observation that in freeze fracture, they looked structurally very similar to

Tight Junctions in Invertebrates

51

FIGURE 3.17 Model of arthropod TJs as viewed side on at the points of membrane junctional ridge fusion on the left, and as the resultant replicas of the E face (top) or P face (bottom) on the right. This is based on studies of rapidly frozen unfixed spider and other tissues after impact with a liquid helium-cooled copper block (Kachar et al., 1992). The possible indirect association of the intramembranous fibrils with cytoskeletal components, such as ZO-1, cingulin, and actin, is indicated.

vertebrate TJs. This was due to the fact that their intramembranous composition is that of particles fused laterally into extensive fibrils or moniliform ridges (Figure 3.16).

3.7.2 RETICULAR SEPTATE JUNCTIONS Reticular “septate” junctions may also be confused with TJs as a result of their freeze-fracture appearance; they are composed of rows of fused IMPs which appear as ridges arrayed in a semireticular fashion and are found over the face of the membranes lying on either side of the stacked scalariform junctional arrays in the dipteran rectal papillae (Lane, 1979a; Flower and Walker, 1979). The intercellular space, which is 15 to 20 nm or greater, seems to contain not septa, but faint striations, which may be columns in some regions. The physiological role of these junctions is still fairly speculative, but it seems unlikely to be an occluding one.

3.7.3 RETINULAR JUNCTIONS Retinular junctions (Lane, 1979b), also called R cell junctions (Chi and Carlson, 1980b), are found between retinular (and glial) cells in the regions of the photoreceptor cell axons or the rhabdomere region of dipteran compound eyes. Again, as with continuous and reticular “septate” junctions, they look like TJs proper by virtue of their freeze-fracture appearance, which is that of PF ridges or EF grooves arrayed at angles to one another. These, however, need not be complementary and the intercellular cleft is never obliterated.

3.8 COMPARISONS BETWEEN TIGHT JUNCTIONS IN THE INVERTEBRATES, LOWER CHORDATES, AND VERTEBRATES In insects, the TJs lie at the inner border of the cells that they link, but in the arachnids, they are found toward the outer border, as they are in vertebrates (Claude and Goodenough, 1973) and lower chordates (Georges, 1979; Lane et al., 1986; Martinucci et al., 1988; Burighel et al., 1992). These layers are similar to the TJ-containing

FIGURE 3.18 Models of junctions that have occasionally been confused with TJs, mainly because of the tight junctional-like intramembranous ridges and grooves that are seen in freeze-fracture replicas. In each case, the lipid bilayers comprising the two plasma membranes of the adjacent cells that form the junction are indicated; the E fracture face is shown on the top left followed by the intercellular cleft components of septal ribbons or columns, with the P fracture face indicated on the bottom right. The tight junction exhibits a pinching together of the two membranes, to occlude the cleft, whereas the smooth septate junction, the retinular or reticular septate junction, and the linker junction all exhibit a cleft that is either unreduced in width or only slightly so.

52 Tight Junctions

Tight Junctions in Invertebrates

53

myelin sheath of vertebrates (Schnapp and Mugniani, 1975; Shinowara et al., 1977) in not being interposed between two extensive fluid compartments as vertebrate epithelia layers often are. There are also differences in the fracturing characteristics of TJs from different sources after comparable treatment. The forces binding the IMPs to the cytoplasmic half of the membrane or to each other can therefore presumably not be completely due to cross-linking induced by glutaraldehyde fixation, as has been suggested for many vertebrate tissues (van Deurs and Luft, 1979). However, the situation in other vertebrate, including lower chordate, tissues, which have been chemically fixed, may be rather different. TJs with E face particle rows are found in the Sertoli cells of the testis (Nagano and Suzuki, 1976), in the endothelial cells of small blood vessels (Simionescu et al., 1978), and in some of the species of tunicates studied (Georges, 1979). In lower chordates such as the tunicates, TJs have been observed in pharyngeal, intestinal, and epidermal epithelia, arranged in strands of IMPs forming a circumferential network at the apical part of cells (Georges, 1979; Lane et al., 1986; 1994; Martinucci et al., 1988). In cephalochordates such as Amphioxus, however, no TJs have been observed at all (Lane et al., 1987). The simple insect ridges tend to lie parallel to the outer surface of the CNS, whereas the more complex arachnid ones are more open meshworks, often more so than the parallel, interconnected alignments that characterize many vertebrate TJs. Tunicate TJs have particle rows arranged in a loose network in a beltlike position but their apical parallel strands are often underlain by a network of ridges, which increase in looseness with the distance from the luminal surface (Lane et al., 1986). The capacity of TJs to coexist with other intercellular junctions seems a universal characteristic. In both vertebrate and invertebrate systems, the TJs are to be found along the same cell border as gap junctions and desmosomes, and, in some insect examples, along with septate junctions. The first of these junctional types is the most common partner of the TJs. Indeed, their persistent coexistence has led to the possibility of making studies in arthropods of their concurrent developmental changes. In young spider hatchlings, the different preferential fracture planes to which the particles of tight (8 to 10 nm, PD) and gap (12 to 14 nm, EF) junctions adhere enable one to follow their distinct patterns of insertion into the presumptive junctional membrane and their subsequent differentiation via translateral intramembranous migration. This, combined with their distinctive sizes, makes it possible to distinguish them unequivocally during development (Lane, 1981a; 1990). There are also differences in insect vs. vertebrate physiology. Insects are remarkable organisms in that they appear to absorb passively and with little selectivity through the septate junction–laden cell wall of their gut almost anything they consume, which tends to lead to fluctuating and unpredictable ionic proportions and concentrations in their hemolymph. Malpighian tubules and rectal tissues are then active in regulating the hemolymph composition. Seemingly in response to this, any particularly important and potentially vulnerable systems have become sealed off from the circulating hemolymph by TJ-based permeability barriers, such as the blood–brain or blood–eye barriers. Lacking septate junctions, the lower chordates (except for the cephalochordates; Lane et al., 1987) similar to the higher vertebrates,

54

Tight Junctions

exhibit a TJ-based barrier along the whole of their gut tract (Lane et al., 1986) as well as in their branchiae (Martinucci et al., 1988).

3.9 ASSEMBLY OF ARTHROPOD TIGHT JUNCTIONS DURING DEVELOPMENT Analysis of the development of junctions in arthropods can normally occur in vivo only during embryonic and very early hatchling stages or during the regeneration of injured tissues (Blanco and Lane, 1990). In some groups, however, such as the holometabolous insects, metamorphosis presents another possible occasion for the formation of junctions during cellular reassembly from disrupted larval tissues into organized adult systems. The first studies to be performed on developing TJs in arthropods were in the holometabolous dipteran blow fly Calliphora. Here the TJs of the developing perineurial layer, the cellular basis of the blood–brain permeability barrier, were studied during the stages of their assembly in embryonic and early life (Lane and Swales, 1978a) as well as during pupal metamorphosis (Lane and Swales, 1978b). In Calliphora the advent of a barrier to the entry of tracer through the perineurial clefts can be correlated with the appearance of simple nonanastomosing tight junctional ridges (Lane and Swales, 1978a). Observations of the CNS during pupal metamorphosis revealed that in the first few days of pupal life the blood–brain barrier broke down. Tracers leaked in around the nerve cells and only fragments of ridges were to be found in fractured perineurial membrane faces (Lane and Swales, 1978b). Toward the middle of pupal life, ridges reformed by apparent reassembly of IMPs and fragments into TJs; tracers were once more excluded from the CNS by the end of pupal life. Developmental studies on TJs in the holometabolous moth Manduca sexta were also made (Lane and Swales, 1979); various stages in apparent junctional assembly were followed in the late embryonic life of the larval form. These stages involved IMPs becoming aligned into short parallel ridges, which ultimately fused into lengthier ridges. Junctional assembly has also been studied in arachnids, in spider CNS where the perineurial layer is associated on its lateral borders by gap junctions, intimately associated with rather more complex TJs. Stages in the assembly of these junctions have been followed in hatchlings wherein the blood–brain barrier is not yet completely formed (Lane, 1981a,c). In this system the 8 to 10 nm tight junctional particles are found first as free PF IMPs; they then begin to become aligned into short and then increasingly long ridges. Ultimately, the fibrils form a reticular network over much of the lateral cell borders.

3.10 BIOCHEMISTRY OF INVERTEBRATE TIGHT JUNCTIONS The TJs in arthropods appear to be associated with cytoskeletal components. As in the vertebrates (Meza et al., 1980; Stevenson and Paul, 1989), there is a suggestion of actin fibrils associated with the cytoplasmic face of the punctate tight junctional

Tight Junctions in Invertebrates

55

appositions in the epithelial layer surrounding insect and arachnid CNS (Lane, 1992). There are, in vertebrates, certain other proteins, also associated with the periphery of tight junctional contacts; these are cingulin and ZO-1. The peripheral proteins, cingulin (108 to 140 kD mol wt; Citi et al., 1988; 1989; Cordenonsi et al., 1999) and ZO-1 (225 kD mol wt; Stevenson and Goodenough, 1984; Stevenson et al., 1986; 1988) have been shown to be specific to tight junctional contacts in vertebrates. In support of this contention, there is a rapid assembly of TJs after mammalian cell–cell contact occurs (Anderson et al., 1989) and at this time the ZO-1 protein levels increase markedly. There is also a report of a 192-kD protein associated with vertebrate TJs (Chapman and Eddy, 1989). The actinlike filaments near the junctions are assumed to be linked to this or to cingulin and/or ZO-1, because ZO-1 is closer to the junctional membrane (Stevenson et al., 1989; Stevenson and Keon, 1998) and, thence, directly or indirectly, to the proteins within the membrane of the TJs. Arthropod TJs also have associated microfilaments; these also appear to be actin, since after S1 labeling, the TJs in spider CNS show the characteristic arrowhead labeling with a corkscrew configuration when viewed end on (Lane, 1992). In any case this cytoskeletal complex presumably permits flexibility and modifies the permeability of the occluding junctions (see Gumbiner, 1987) in arthropod systems as well as vertebrate ones. More recent studies have revealed the existence of intramembranous TJ proteins — occludin and claudin; investigations on occludin (Furuse et al., 1993) and the claudins I and II (Tsukita and Furuse, 1999) show them to be transmembrane TJ components. Preliminary work suggests they may also exist in arachnid TJs (Kachar and Lane, unpublished observations).

3.11 CONCLUSIONS A good deal is known about the basic morphology of invertebrate TJs, by thinsection and freeze-fracture criteria, but there is still much to be determined concerning their biochemistry. Their structural features and apparent association with cytoskeletal elements suggests that there may be a fundamental similarity in physiology to the vertebrate TJ but the subtleties of their functional regulatory mechanisms have yet to be elucidated. The technical difficulties encountered in making the appropriate physiological measurements make this task a formidable one, while the relative paucity of TJs in arthropod tissues makes their successful biochemical isolation problematic. More feasible, and one hopes fruitful, studies involve further immunocytochemical investigations, as well as genetic analysis looking for sequences homologous with vertebrate tight junctional proteins.

ACKNOWLEDGMENTS This author is grateful to the E.M.P. Musgrave Fund for support during the preparation of this manuscript.

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REFERENCES Anderson, J. M., Van Itallie, C. M., Peterson, M. D., Stevenson, B. R., Carew, E. A., and Mooseker, M. S. 1989. ZO-1 mRNA and protein expression during tight junction assembly in Caco-2 cells. J. Cell Biol., 109: 1047–1056. Binnington, K. C. and Lane, N. J. 1980. Perineurial and glial cells in the tick Boophilus microplus (Acarina: Ixodidae): freeze-fracture and tracer studies. J. Neurocytol., 9: 343–362. Blanco, R. E. and Lane, N. J. 1990. Changes in intercellular junctions during peripheral nerve regeneration in insects. J. Neurocytol., 19: 873–882. Bullivant, S. 1978. The structure of tight junctions, in Electron Microscopy, Vol. III. State of the Art Symposia. Sturgess, J.M., Ed., Imperial Press, Canada, 659–672. Burighel, P., Martinucci, G. B., Lane, N. J., and Dallai, R. 1992. The junctional complexes of the branchia and gut of the tunicate, Pyrosoma atlanticum. Cell Tissue Res., 267: 357–364. Carlson, S. D., Juang, J.-L., Hilgers, S. L., and Garment, M. B. 2000. Blood barriers of the insect. Annu. Rev. Entomol., 45: 151–174. Chapman, L. M. and Eddy, E. M. 1989. A protein associated with the mouse and rat hepatocyte junctional complex. Cell Tissue Res., 257: 333–341. Chi, C. and Carlson, S. D. 1980a. Membrane specializations in the first optic neuropil of the housefly (Musca domestica L.). I. Junctions between neurons. J. Neurocytol., 9: 429–449. Chi, C. and Carlson, S. D. 1980b. Membrane specializations in the first optic neuropil of the housefly (Musca domestica L.). II. Junctions between glial cells. J. Neurocytol., 9: 451–469. Chi, C. and Carlson, S. D. 1981. The perineurium of the adult housefly. Ultrastructure and permeability to lanthanum. Cell Tissue Res., 217: 373–386. Citi, S., Sabanay, H., Jakes, R., Geiger, B., and Kendrick-Jones, J. 1988. Identification and isolation of cingulin, a new cytoplasmic component of tight junctions. Nature (London), 333: 272–275. Citi, S., Sabanay, H., Kendrick-Jones, J., and Geiger, B. 1989. Cinglin: characterization and localization. J. Cell Sci., 93: 107–122. Claude, P. and Goodenough, D. A. 1973. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia. J. Cell Biol., 58: 390–400. Cordenonsi, M., D’Atri, F., Hammar, E., Parry, D. A. D., Kendrick-Jones, J., Shore, D., and Citi, S. 1999. Cingulin contains globular and coiled-coil domains and interacts with ZO-1, ZO-2, ZO-3 and myosin. J. Cell Biol., 147: 1569–1581. Dallai, R., Bigliardi, E., and Lane, N. J. 1990. Intercellular junctions in myriapods. Tissue Cell, 22: 359–369. Farquhar, M. G. and Palade, G. E. 1963. Junctional complexes in various epithelia. J. Cell Biol., 17: 375–412. Farquhar, M. G. and Palade, G. E. 1965. Cell junctions in amphibian skin. J. Cell Biol., 26: 263–291. Flower, N. E. 1972. A new junctional structure in the epithelia of insects of the order Dictyoptera. J. Cell Sci., 10: 683–691. Flower, N. E. and Walker, G. D. 1979. Rectal papillae in Musca domestica: the cuticle and lateral membranes. J. Cell Sci., 39: 167–186. Furuse, M., Hirase, T., Itoh, M., Nagafuchi, A., Yonemura, S. et al. 1993. Occludin: a novel integral membrane protein localizing at tight junctions. J. Cell Biol., 123(6): 1777–1788.

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Georges, D. 1979. Gap and tight junctions in tunicates: study in conventional and freezefracture techniques. Tissue Cell, 11: 781–792. Green, C. R., Bergquist, P. R., and Bullivant, S. 1979. An anastomosing septate junction in endothelial cells of the phylum Echinodermata. J. Ultrastruct. Res., 68: 72–80. Gumbiner, B. 1987. Structure, biochemistry and assembly of epithelial tight junctions. Am. J. Physiol., 253: C749–C758. Kachar, B. and Reese, T. S. 1982. Evidence for the lipidic nature of tight junction strands. Nature (London), 296: 464–466. Kachar, B., Reese, T. S., and Lane, N. J. 1992. Structural domains of the TJ intramembranous fibrils. Tissue Cell, 24, 291–300. Lane, N. J. 1972. Fine structure of a lepidopteran nervous system and its accessibility to peroxidase and lanthanum. Z. Zellforsch., 131: 205–222. Lane, N. J. 1979a. Freeze-fracture and tracer studies on the intercellular junctions of rectal tissues in insects. Tissue Cell, (3)11: 481–506. Lane, N. J. 1979b. A new kind of tight junctional-like structure in insect tissues. J. Cell Biol., 83: 82A. Lane, N. J. 1981a. Tight junctions in arthropod tissues. Int. Rev. Cytol., 73: 243–318. Lane, N. J. 1981b. Invertebrate neuroglia: junctional structure and development. J. Exp. Biol., 95: 7–33. Lane, N. J. 1981c. Evidence for two separate categories of junctional particle during the concurrent formation of tight and gap junctions. J. Ultrastruct. Res., 77: 54–65. Lane, N. J. 1981d. Vertebrate-like tight junctions in the insect eye. Exp. Cell Res., 132: 482–488. Lane, N. J. 1989. Novel arthropod cell junctions with restricting intercellular “linkers.” J. Neurocytol., 18: 661–669. Lane, N. J. 1990. Intercellular junctions, structure and cytoskeletal associations, in Structure and Function in Zoology, Proc. Symp. Bormio, Lanzavecchia, G., Ed. Selected Symposia and Monographs, 0.2.1, 5, Mucchi, Modena, 87–102. Lane, N. J. 1992. Anatomy of the tight junction: invertebrates, in Tight Junctions, Cereijido, M., Ed., CRC Press, Boca Raton, FL, 23–48. Lane, N. J. and Campiglia, S. 1987. The lack of a structured blood–brain barrier in the onychophoran Peripatus acacioi. J. Neurocytol, 16: 93–104. Lane, N. J. and Chandler, H. J. 1980. Definitive evidence for the existence of tight junctions in invertebrates. J. Cell Biol., 86: 765–774. Lane, N. J. and Flores, V. 1988. Actin filaments are associated with the septate junctions of invertebrates. Tissue Cell, 20: 211–217. Lane, N. J. and Flores, V. 1990. The role of cytoskeletal components in the maintenance of insect junctions. Cell Tissue Res., 262: 373–385. Lane, N. J. and Swales, L. S. 1978a. Changes in the blood–brain barrier of the central nervous system in the blowfly during development, with special reference to the formation and disaggregation of gap and tight junctions. I. Larval development. Dev. Biol., 62: 389–414. Lane, N. J. and Swales, L. S. 1978b. Changes in the blood–brain barrier of the central nervous system in the blowfly during development, with special reference to the formation and disaggregation of gap and tight junctions. II. Pupal development and adult flies. Dev. Biol., 62: 415–431. Lane, N. J. and Swales, L. S. 1979. Intercellular junctions and the development of the blood–brain barrier in Manduca sexta. Brain Res., 169: 226–245. Lane, N. J. and Treherne, J. E. 1969. Peroxidase uptake by glial cells in desheathed ganglia of the cockroach. Nature (London), 223: 861–862.

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Lane, N. J. and Treherne, J. E. 1970. Uptake of peroxidase by the cockroach central nervous system. Tissue Cell, 2: 413–425. Lane, N. J. and Treherne, J. E. 1972. Studies on perineurial junctional complexes and the sites of uptake of microperoxidase and lanthanum in the cockroach central nervous system. Tissue Cell, 4: 427–436. Lane, N. J. and Treherne, J. E. 1973. The ultrastructural organization of peripheral nerves in two insect species. Tissue Cell, 5: 703–714. Lane, N. J. and Treherne, J. E. 1980. Functional organization of arthropod neuroglia, in Insect Biology in the Future, V.B.W. 80, Locke, M. and Smith, D. S., Eds., Academic Press, London, 765–795. Lane, N. J., Skaer, H. leB., and Swales, L. S. 1975. Junctional complexes in insect nervous systems. J. Cell Biol., 67: 233A. Lane, N. J., Skaer, H. leB., and Swales, L. S. 1977. Intercellular junctions in the central nervous system of insects. J. Cell Sci., 26: 175–199. Lane, N. J., Harrison, J. B., and Bowerman, R. F. 1981. A vertebrate-like blood–brain barrier, with intraganglionic blood channels and occluding junctions, in the scorpion. Tissue Cell, 13: 557–576. Lane, N. J., Dallai, R., Burighel, P., and Martinucci, J. 1986. Tight and gap junctions in the intestinal tract of tunicates: a freeze-fracture study. J. Cell Sci., 84: 1–18. Lane, N. J., Dallai, R., Martinucci, G. B., and Burighel, P. 1987. Cell junctions in Amphioxus (Cephalochordata): a thin-section and freeze-fracture study. Tissue Cell, 19: 399–411. Lane, N. J., Dallai, R., Martinucci, G., and Burighel, P. 1994a. Electron microscopic structure and evolution of epithelial junctions, in Molecular Mechanisms of Epithelial Cell Junctions: From Development to Disease. Citi., S., Ed., R.G. Landes, Austin, TX, 23–43. Leslie, R. A. 1973. A comparison of the fine structure of interganglionic connectives in newly hatched and adult stick insects. Z. Zellforsch., 145: 299–309. Lorber, V. and Rayns, D. G. 1972. Cellular junctions in the tunicate heart. J. Cell Sci., 10: 211–227. Martinucci, G., Dallai, R., Burighel, P., and Lane, N. J. 1988. Different functions of tight junctions in the ascidian branchial basket. Tissue Cell, 20 (1): 119–132. McLaughlin, B. J. 1974. Fine structural changes in a lepidopteran nervous system during metamorphosis. J. Cell Sci., 14: 369–387. Meza, I., Ibarra, G., Sabanero, M., Martinez-Paloma, A., and Cereijido, M. 1980. Occluding junctions and cytoskeletal components in a cultured transporting epithelium. J. Cell Biol., 87: 746–754. Nagano, T. and Suzuki, F. 1976. The postnatal development of the junctional complexes of the mouse Sertoli cells as revealed by freeze-fracture. Anat. Rec., 185: 403–415. Noirot, C. and Noirot-Timothée, C. 1967. Un nouvenu type de jonction intercellulaire (zonula continua) dans l’intestin moyen des insectes. C.R. Acad. Sci. Ser. D, 264: 2796–2798. Noirot-Timothée, C. and Noirot, C. 1980. Septate and scalariform junctions in arthropods. Int. Rev. Cytol., 63: 97–140. Saint Marie, R. L., Carlson, S. D., and Chi, C. 1984. The glial cells of insects, in Insect Ultrastructure, Vol. 2, King, R. C. and Akai, H., Eds., Plenum, New York, 437–475. Satir, P. and Gilula, N. B. 1973. The fine structure of membranes and intercellular communication in insects. Annu. Rev. Ent., 18: 143–166. Schnapp, B. and Mugnaini, E. 1975. The myelin sheath: EM studies with thin section and freeze-fracture, in Golgi Centennial Symp. Proc., Santini, M., Ed.. Raven Press, New York, 209–233. Shaw, S. R. 1978. The extracellular space and blood–eye barrier in an insect retina: an ultrastructural study. Cell Tissue Res., 188: 35–61.

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Shinowara, N. L., Beutel, W. D., and Revel, J. P. 1977. Tight junctions in peripheral myelin. J. Cell Biol., 75: 62A. Simionescu, N., Simionescu, M., and Palade, G. E. 1978. Open junctions in the endothelium of the postcapillary vesicles of the diaphragm. J. Cell Biol., 79: 27–44. Staehelin, L. A. 1974. Structure and function of intercellular junctions. Int. Rev. Cytol., 39: 191–283. Stevenson, B. R. and Goodenough, D. A. 1984. Zonulae occludentes in junctional complex enriched fractions from mouse liver: preliminary morphological and biochemical characterization. J. Cell Biol., 98: 1209–1221. Stevenson, B. R. and Keon, B. H. 1998. The tight junction: morphology to molecules. Annu. Rev. Cell Dev. Biol., 14: 89–109. Stevenson, B. R. and Paul, D. L. 1989. The molecular constituents of intercellular junctions. Curr. Opin. Cell Biol., 1: 884–891. Stevenson, B. R., Siliciano, J. D., Mooseker, M. S., and Goodenough, D. A. 1986. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J. Cell Biol., 103: 755–766. Stevenson, B. R., Anderson, J. M., and Bullivant, S. 1988. The epithelial tight junction: structure, function and preliminary biochemical characterization. Mol. Cell. Biochem., 83: 129–145. Stevenson, B. R., Heintzelman, M. B., Anderson, J. M., Citi, S., and Mooseker, M. S. 1989. A comparison of the localizations of the tight junction proteins ZO-1 and cingulin. J. Cell Biol., 109: 45A. Szollosi, A. and Marcaillou, C. 1977. EM study of the blood–testis barrier in an insect. J. Ultrastruct. Res., 59: 158–172. Toshimori, K., Iwashita, T., and Oura, C. 1979. Cell junctions in the cyst envelope in the silkworm testis, Bombyx mori Linne. Cell Tissue Res., 202: 63–73. Treherne, J. E. and Pichon, Y. 1972. The insect blood–brain barrier, in Advances in Insect Physiology, Vol. 9, Treherne, J. E., Berridge, M. J., and Wigglesworth, V. B., Eds., Academic Press, New York, 257–313. Treherne, J. E., Lane, N. J., Moreton, R. B., and Pichon, Y. 1970. A quantitative study of K+ movements in the CNS of Periplaneta americana. J. Exp. Biol., 53: 109–136. Tsukita, S. and Furuse, M. 1999. Occludin and claudins in tight junction strands: leading or supporting players? Trends Cell Biol., 9: 268–273. van Deurs, B. and Luft, J. H. 1979. Effects of glutaraldehyde fixation on the structure of tight junctions. A quantitative freeze-fracture analysis. J. Ultrastruct. Res., 68: 160–172. Villegas, G. M., Lane, N. J., and Villegas, J. 1987. Freeze-fracture studies on the giant axon and ensheathing Schwann cells of the squid. J. Neurocytol., 16: 11–21. Zwahlen, M. J., Sandri, C., and Greeff, N. G. 1988. Transglial pathway of diffusion in the Schwann sheath of the squid giant axon. J. Neurocytol., 17: 145–159.

4

Tight Junction Permeability to Ions and Water Luis Reuss

CONTENTS 4.1 4.2

Introduction ....................................................................................................62 Electrical Resistance of Tight Junctions........................................................63 4.2.1 The Junctional Electrical Resistance Is a Good Measure of Junctional Ion Permeability ...............................................................63 4.2.2 Two Kinds of Epithelia, Tight and Leaky, Can Be Distinguished from the Ratio of the Paracellular to the Transcellular Conductance .......................................................................................66 4.2.3 The Paracellular Pathway May Have Several Roles in Leaky Epithelia .............................................................................................69 4.2.3.1 Electrical Coupling of the Cell Membranes ......................69 4.2.3.2 Passive (Electrodiffusive) Ion Transport ............................69 4.2.3.3 Osmotic Water Transport....................................................71 4.2.3.4 Solvent Drag .......................................................................71 4.2.4 The Junctional Location of the High-Conductance Pathway in Leaky Epithelia Was Demonstrated with Electrophysiological and Morphological Techniques..........................................................71 4.3 Junctional Ion Selectivity...............................................................................72 4.4 Mechanisms of Junctional Ion Permeation ...................................................74 4.4.1 The Junctional Ion Permeation Pathway Consists of Pores .............74 4.4.2 The Junctional Permeability Depends on Solute Charge and Size .....................................................................................................75 4.4.3 Relationship of Junctional Depth and Junctional Permeability ........77 4.5 Junctional Water Permeability .......................................................................77 4.5.1 Relationship between Junctional Ion and Water Permeability..........78 4.5.2 Junctional Permeation of Large Hydrophilic Solutes .......................79 4.5.3 Estimates of Junctional Water Permeability......................................79 4.5.4 Solute–Solvent Coupling and Electrokinetic Phenomena.................81 4.6 Conclusions ....................................................................................................82 References................................................................................................................83

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4.1 INTRODUCTION The issue of the chemical composition of the tight junctions (TJs) appears to be near resolution. Recent studies carried out principally by Tsukita’s group (reviewed in Tsukita et al., 1999; Tsukita and Furuse, 2000) indicate that several hydrophobic proteins are components of the junctional strands. The claudins (integral membrane proteins of molecular mass ~28 kDa, cytoplasmic amino and carboxy termini, four transmembrane domains, over 20 isoforms described; Furuse et al., 1998a,b; see also Chapter 10) are essential. Occludin (integral membrane protein of molecular mass ~60 kDa, similar topology to occludins, one known gene product), the first integral junctional protein to be identified (Furuse et al., 1993; Ando-Akatsuka et al., 1996), proved nonessential in gene-knockout and heterologous-expression experiments (Saito et al., 1998; Furuse et al., 1998b). In addition, the junctions contain numerous peripheral proteins (see Chapter 11) that appear to play roles in targeting the junctional proteins themselves, association of TJs with the actin cytoskeleton, intracellular signaling, and vesicle targeting. A second recent advance in this field was the discovery of paracellin, a claudin isoform expressed on the junctions of thick-ascending limb of Henle’s loop, in the kidney (Simon et al., 1999). Paracellin-1/claudin-16 accounts for the high permeability of these junctions to divalent cations and thus underlies Mg2+ and Ca2+ reabsorption in this nephron segment. This important discovery provides the first identification of a molecular component of the paracellular pathway for transepithelial ion transport. The relationship between the emerging chemistry of the TJs and their morphology has not been established. In other words, the parts of this organelle (all or most of them) are known, but not their detailed arrangement in space and time. It follows that since the morphology is not fully understood, it is difficult to choose a structural basis on which to interpret biophysical and physiological data. TJs are formed by strands roughly parallel to the cell surface, arranged as multiple barriers in series, with variable degree of branching and anastomoses. The strands contain claudins and occludin, and perhaps other proteins and specific lipids as well. Biophysical evidence supports the view that ion permeation is via pores in or between the strands, but this has not been proved. It has also been proposed that the pores fluctuate between open and closed states (Claude, 1978; González-Mariscal et al., 1984; Cereijido et al., 1989). In contrast to cell membranes, which have low permeability for hydrophilic solutes and are highly permeable to lipophilic molecules, the paracellular pathway of all epithelia so far studied behaves as an aqueous pathway restricting solute permeation on the basis of size and charge. Studies of junctional permeability generally involve measurements of fluxes (of ions, water, or nonelectrolytes), total ionic permeability (or electrical conductance), or transepithelial-voltage changes upon exposure to solutions of different composition. Because of their ease, accuracy, and time resolution, electrophysiological techniques are generally preferred to other methods when studying junctional ion permeability. This chapter aims to emphasize the bases of the major techniques used to study junctional ion and water permeability, pitfalls associated with the interpretation of the results, examples of their application,

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and a framework for interpretation of the physiological data at the structural level. Issues pertaining to TJ regulation and TJ-associated pathology are discussed elsewhere in this book (Chapters 15, 16, and 21 through 32).

4.2 ELECTRICAL RESISTANCE OF TIGHT JUNCTIONS 4.2.1 THE JUNCTIONAL ELECTRICAL RESISTANCE IS A GOOD MEASURE OF JUNCTIONAL ION PERMEABILITY Electric-current flow in aqueous solutions and across biological membranes is caused by ion flow. Thus, the electrical resistance (or its reciprocal, the conductance) of TJs provides a direct assessment of junctional ion permeability. Ion conductance and ion permeability are related (Schultz, 1980), but not equivalent. Permeability is an intrinsic property of the membrane, in principle independent of ion concentration, whereas ion conductance is concentration dependent. To assess the electrical resistance of TJs one must consider the complex equivalent circuit that represents an epithelium. So far, it has not been possible to measure junctional permeability directly, either in situ or in a purified and reconstituted system. Junctions and cell membranes constitute parallel transepithelial current (ion) pathways. Transepithelial ion fluxes and current flow include, in principle, contributions of both pathways (Figure 4.1, left). To estimate the conductance of the junctions one needs to perform equivalent-circuit analysis. The simplest model for such analysis is depicted in Figure 4.1, right. Steady-state resistances are measured. The unknowns are the cell-membrane electrical resistances (apical and basolateral: Ra and Rb, respectively) and the paracellular electrical resistance (Rs). In monolayered epithelia of one cell type, Rs denotes the resistance of the junctions and the lateral intercellular spaces (in series), but this may not be the case if there are other parallel conductive pathways, such as edge or focal damage of the epithelium. This circuit is an oversimplification. In the cell membranes, capacitance and equivalent electromotive forces (EMF, caused by electrodifussion of permeant ions) should be added to the resistances, as well as rheogenic (i.e., current-generating) ion pumps. Provided that the active components of the circuit — zero-current voltage (EMF) and current (I) — are constant, the circuit can be analyzed by DC and by AC techniques. Both approaches are difficult: DC studies are based on simple, perhaps oversimplified, equivalent circuits; AC methods depend critically on the model chosen and may have poor time resolution. The remainder of this section briefly discusses the principles of the measurements. To assess the junctional resistance one starts with the passive equivalent circuit shown in Figure 4.1, right. The lateral and basal portions of the basolateral membrane of the epithelial cells are “lumped,” implying a small lateral-intercellular-space resistance (Rlis). In other words, the entire space would be isopotential during current application. The equivalent resistor depicted is correct only under this condition, but if Rlis is significant, then the circuit is distributed, and calculations based on the lumped circuit yield overestimates of the basolateral membrane resistance (Rb). For further discussion, see Boulpaep and Sackin (1980). The lumped equivalent circuit of Figure 4.1, right has been solved by several approaches:

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FIGURE 4.1 Passive equivalent electrical circuit for an epithelium. As shown on left, ions can permeate through TJ and lateral intercellular space (paracellular or “shunt” pathway) or through cells (cellular pathway), either across apical and basal membrane or across apical membrane and lateral membrane, the latter in series with lateral space. On right, Ra = equivalent electrical resistance of apical membrane, Rb = equivalent resistance of basolateral membrane and lateral intercellular space, and Rs = shunt resistance. M, C, and S stand for mucosal solution, cell, and serosal solution, respectively. (From Reuss, L., in Membrane Transport in Biology, Vol. IVB, Giebisch, G. et al., Eds., Springer-Verlag, New York, 1979. With permission.)

1. Experimental alteration of a cell-membrane conductance and measurement of the effect on the transepithelial conductance (Gt). Yonath and Civan (1971) used this method in toad urinary bladder epithelium. Antidiuretic hormone (ADH) was used, and it was assumed that the only effect of the hormone was to increase Ga. In the toad urinary bladder exposed to identical bathing solutions the short-circuit current (Isc = transepithelial current required to make the transepithelial voltage 0 with identical bathing solutions on both sides) is almost exactly equivalent to the net transepithelial Na+ transport. ADH increases the Isc via an increase in the apicalmembrane Na+ conductance and a stimulation of the basolateral Na+,K+ATPase secondary to Na+ entry. A plot of Gt vs. time yields the straight line described by Equation 4.1: Gt = Gs + Isc ENa

(4.1)

where Gs = paracellular conductance (= 1/Rs) and ENa is the driving force for active Na+ transport, a complex term involving apical and basolateral membrane parameters. Gs is given by the y-axis intercept of the linear fit of the data to Equation 4.1.

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2. Complete passive circuit analysis: determination of the three resistors (Ra, Rb , and Rs) by cable analysis and measurements of Rt and Ra /Rb (Frömter, 1972). These three measurements yield values for the three unknowns, Ra , Rb, and Rs (Frömter, 1972; Reuss and Finn, 1975a). 3. Measurements of Rt and Ra /Rb before and after altering experimentally either Ra or Rb . Without cable analysis, it is possible to obtain four independent measurements (values of Rt and Ra /Rb) before and after the change in a cell-membrane resistance, e.g., from Ra to Ra′. From these four measurements the four unknowns (Rs, Rb, Ra, and Ra′) can be calculated solving simultaneous equations (Reuss and Finn, 1974). This or similar analyses have been used in a variety of epithelia (Reuss and Finn, 1974; Lewis et al., 1977; Frömter and Gebler, 1977; Lewis and Wills, 1982; Reuss et al., 1983). 4. Measurements of the changes in Vt and a membrane voltage, e.g., Vb produced by altering experimentally the permeability of the contralateral cell-membrane domain (in this case, apical). The current flow elicited by this experimental perturbation causes a loop flow in the epithelium and voltage drops across Rs and Rb. Combining these measurements with determinations of Rt and Ra /Rb, Ra, Rb, and Rs can be calculated. This or similar analyses have been used by Frömter and Gebler (1977), Lewis and Wills (1982), and Reuss et al. (1983). The method of Yonath and Civan hinges on the assumption that only the apical membrane properties are affected by ADH. Using a similar approach, Erlij (1976) exposed toad urinary bladder epithelium to amiloride to block completely the apicalmembrane Na+ conductance, and assumed that the remaining conductance was entirely paracellular and insensitive to amiloride. However, in at least two epithelia amiloride reduces the Na+ flux from basolateral to apical solution, suggesting that the paracellular Na+ permeability falls (O’Neil and Helman, 1976; Sansom and O’Neil, 1985). To overcome the limitation imposed by the length of time required for a twodimensional cable analysis experiment, a two-point cable analysis method was designed. Current is injected in one cell and the elicited voltage is measured in the same cell and in another (distant) cell (Petersen and Reuss, 1985; Stoddard and Reuss, 1989; Segal and Reuss, 1990). This method, combined with determinations of Ra /Rb, allows for an assessment of the direction of changes in Ra and/or Rb with excellent time resolution. As explained above, the interpretation of the value of Ra /Rb requires the choice of an equivalent circuit for the lateral membrane and the intercellular space. If the circuit is distributed instead of lumped, then the value determined is an underestimate of the true ratio, an effect inversely proportional to the width of the spaces (Boulpaep and Sackin, 1980). Intracellular-microelectrode studies suggest that this is not the case in Necturus gallbladder epithelium (Stoddard and Reuss, 1988). Selective modifications of the properties of one cell-membrane domain require the investigator to rule out paracellular effects as well as indirect effects mediated by changes in cell volume and/or composition and voltage dependence of other

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pathways. Significant changes in cell composition may be prevented by reducing the time of exposure to the experimental solution (see Frömter and Gebler, 1977; Reuss et al., 1983). Voltage-dependent conductances can be ruled out by the use of appropriate channel blockers (Stoddard and Reuss, 1988). When the value of Rs (or Gs) is calculated by one of the above methods, the meaning of that value remains uncertain, for three main reasons. First, the conductance may be in part artifactual; in vitro studies can result in considerable damage at the edge of the preparation, yielding a spuriously large value of Gs (Helman and Miller, 1973; Higgins et al., 1975; Lewis and Diamond, 1976). Certain epithelia become leaky in vitro because of errors in bathing-solution composition. For example, the rabbit gallbladder incubated in phosphate-free solutions (Barry et al., 1971) and the Necturus gallbladder exposed to low-Ca2+ bathing medium (Hill and Hill, 1978b; Diamond, 1979) undergo progressive and irreversible increases in paracellular conductance in vitro. Both problems can be identified and corrected (Helman and Miller, 1973; Higgins et al., 1975; Lewis and Diamond, 1976). Second, other high-conductance pathways can be confused, e.g., a particular cell type, dead or damaged cells. To demonstrate the intercellular location of the parallel pathway, other approaches are needed (see the next section). Third, the relative contributions of junctions and lateral spaces to the paracellular resistance are many times unclear. In AC studies in Necturus gallbladder it was found that there is a high resistance in the lateral spaces and thus that the equivalent circuit is distributed (Kottra and Frömter, 1984a,b). However, DC intracellular microelectrode studies did not support this interpretation. Changes in space width were elicited by the “transport number effect” (Barry and Hope, 1969a,b), passing large transepithelial currents. Current flow from apical to basolateral solution widens the spaces because of accumulation of NaCl and water; current flow in the opposite direction narrows the spaces by depletion of NaCl and water. In the latter condition, Ra and Rb in parallel, estimated from two-dimensional cable analysis, did not change significantly, whereas Rt underwent large changes (Stoddard and Reuss, 1988). This result suggests that the changes in lateral space width are dominant in their apical ends, but a voltage-dependence TJ conductance is also possible. It may be of interest to compare the effects on Rt and Ra /Rb of closure of the spaces by transepithelial osmotic gradients. A discussion of AC-analysis methods is beyond the scope of this chapter but can be found in several excellent reviews (Clausen et al., 1979; 1993; Clausen, 1989; Lewis et al., 1996).

4.2.2 TWO KINDS OF EPITHELIA, TIGHT AND LEAKY, CAN BE DISTINGUISHED FROM THE RATIO OF THE PARACELLULAR TO THE TRANSCELLULAR CONDUCTANCE The first coherent explanation for the mechanism of transepithelial ion transport, formulated by Ussing and co-workers in the 1950s (Koefoed-Johnsen and Ussing, 1958), is the so-called two-membrane hypothesis, based on studies of Na+ absorption across frog skin epithelium, and later applied to many other epithelia. As illustrated in Figure 4.2, there are two barriers in series, the outer (apical) and inner (basolateral) cell-membrane domains that have different transport properties: the apical membrane

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FIGURE 4.2 Transport mechanism in Na+-absorptive epithelia. The two-membrane hypothesis for Na+-absorbing “tight” epithelia (Koefoed-Johnsen and Ussing, 1958). At the apical cell membrane, Na+ entry is via a Na+-channel blockable by amiloride. At the basolateral cell membrane, Na+ extrusion is mediated by a Na+, K+-ATPase inhibitable by ouabain. K+ “recycles” across the basolateral cell membrane via a K+ channel. (From Reuss, L., in Handbook of Physiology, Section 14, Cell Physiology, Hoffman, J. E. and Jamieson, J., Eds., Oxford University Press, New York, 1997. With permission.)

is Na+ permeable; whereas the basolateral membrane is K+ permeable and contains the active Na+ transporter, i.e., the Na+,K+-ATPase. The two-membrane hypothesis rests on three experimental pillars: first, the measurement of the Na+ flux by the short-circuit current technique (Ussing and Zerahn, 1951; see Figure 4.2); second, the assessment of the ionic permeabilities of the apical and basolateral membranes in skins mounted in the Ussing chamber (Koefoed-Johnsen and Ussing, 1958), and, third, the finding of the basolateral location of the Na+,K+-ATPase (Koefoed-Johnsen, 1957). The short-circuit current technique allowed Ussing and co-workers to demonstrate that only Na+ was actively transported across the epithelium. The permeability studies, based on cation substitutions in one of the solutions (after replacing Cl– with the impermeant SO42– or adding the Cl– transport blocker Cu2+) and observing the changes in transepithelial voltage, demonstrated the location of the leak pathway for Na+ to the apical membrane. The location of the Na+,K+-ATPase was initially based on the inhibition of Na+ transport by removal of K+ from basolateral-membrane bathing solution. The later use of pump inhibitors such as ouabain and metabolic poisons confirmed this interpretation (Koefoed-Johnsen, 1957; Huf et al., 1957). Although clearly valid for a large number of epithelia, the two-membrane hypothesis does not explain ion transport in epithelia with properties very different from those of the frog skin epithelium. For example, in the fish gallbladder (Diamond, 1962a,b,c) there is a very small transepithelial voltage during transport and

68

Tight Junctions

a transepithelial ion permeability (or conductance) one or two orders of magnitude higher than that of the frog skin. The result is near-electroneutral transport. The gallbladder may absorb Na+ at higher rate than the frog skin, but the short-circuit current is near zero. From these and other studies (Windhager et al., 1967; Frömter and Diamond, 1972) two important new ideas were developed. First, there may be a paracellular or “shunt” pathway in parallel with the cells, first suggested by Ussing and Windhager (1964) from studies in frog skin epithelia treated with hyperosmotic solutions. Second, in some epithelia the ion-leak pathway may be electroneutral, totally or in part (Nellans et al., 1973; Frizzell et al., 1975). The entry step would not be via ion channels, but via carriers, later identified as Na+-Cl– or Na+-K+-2Cl– – cotransporters, or Na+/H+ and Cl–/HCO3 parallel exchangers. Ion transport mediated by these transporters does not involve net-charge translocation. The first demonstrations of paracellular ion permeation were obtained by electrophysiological studies in amphibian renal proximal tubules and gallbladder (Windhager et al., 1967; Frömter, 1972; Frömter and Diamond, 1972). Ultrastructural studies demonstrated high tight junctional permeability to lanthanum in epithelia from small intestine and gallbladder (Machen et al., 1972), in contrast to the impermeability of native frog skin epithelium (Ussing and Windhager, 1964). The concepts of “tight” and “leaky” epithelia emanated from this work. The components of the transepithelial resistance in a simple epithelium with one cell type, homogeneous junctions, and lack of edge or artifactual damage are the transcellular and the paracellular pathway. These are in parallel (see Figure 4.1), so that the transepithelial resistance (Rt) depends on the transcellular (Ra + Rb) and paracellular (Rs) resistances according to the following equation 1 1 1 = + Rt Ra + Rb Rs

(4.2)

where the subscripts denote: t = transepithelial, a = apical, b = basolateral, and s = paracellular (shunt). Resistance values are usually expressed per unit area of epithelium. This equation can also be written for the conductances, which are reciprocal to the resistances (Gt = Gc + Gs, where the subscript c denotes the cellular pathway). The latter equation indicates that the high-conductance pathway determines the value of Gt (or its reciprocal, Rt). The degree of epithelial leakiness is best assessed by the ratio Rc /Rs (where Rc is the transcellular resistance, = Ra + Rb) or the equivalent conductance ratio Gs /Gc. The larger this ratio, the leakier the epithelium. In most cases, low values of Rt can be explained by low values of Rs. However, an exception is the salivary duct, a lowRt epithelium with a high-conductance transcellular pathway and a low-conductance paracellular pathway (Augustus et al., 1977). True leaky epithelia have low Rs and low Vt, and cannot establish or maintain large concentration differences of permeant ions between the two solutions. The leakiest epithelium is the mammalian proximal renal tubule (Lutz et al., 1973; Frömter, 1982), and the tightest the mammalian urinary bladder epithelium (Lewis and Diamond, 1976). Examples of leaky and tight epithelia are listed in Table 4.1.

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69

TABLE 4.1 Electrical Resistances of Some Epithelia Ω · cm2 Epithelium PT (Necturus) GB (Necturus) Colon (rabbit) UB (toad) UB (rabbit)

Rt 260 310 330 3,800 10,000–40,000

Ra 3,400–10,900 4,000 1,100 4,000 9,000–43,000

Rb 2,400 2,500 100 3,200 1,000–1,800

Rs 270 350 500 13,200 ≥31,000

Ref.a 1 * * 2 3

Abbreviations: PT = proximal renal tubule, GB = gallbladder, UB = urinary bladder. Rt = transepithelial resistance, Ra = apical cell membrane resistance, Rb = basolateral cell membrane resistance, Rs = paracellular resistance. a

References not given (*) can be found in Powell (1981). 1. Guggino et al. (1982); 2. Reuss and Finn (1974); 3. Lewis et al. (1977).

4.2.3 THE PARACELLULAR PATHWAY MAY HAVE SEVERAL ROLES IN LEAKY EPITHELIA The functional role of the paracellular pathway in leaky epithelia remains controversial. The main issues are whether or not it contributes significantly to ion and water transport, and the magnitude and mechanisms of these contributions. 4.2.3.1 Electrical Coupling of the Cell Membranes A paracellular path of relatively low resistance to ion flow couples the cell membranes because it allows for circular current flow in the epithelium if the zero-current voltages of the cell membranes are different (Figure 4.3; see Schultz, 1979). Generally, the K+ selectivity of the basolateral membrane is greater than that of the apical membrane, and therefore the basolateral-membrane zero-current voltage is greater. Thus, the intraepithelial current flow causes hyperpolarization of the apical membrane, which increases the driving force for inflow of positively charged solutes (e.g., Na+–organic substrate complexes). This happens in the renal proximal tubule and the small intestine, where the epithelial cells express electrogenic Na+-nutrient cotransporters, but not in other leaky epithelia. In all leaky epithelia, a consequence of intraepithelial current flow is apical-membrane hyperpolarization that tends to reduce K+ loss to the lumen via apical-membrane K+ channels. 4.2.3.2 Passive (Electrodiffusive) Ion Transport Passive ion transport is driven by the gradient in electrochemical potential across the junctions. The driving force depends on the valence of the ion and the transjunctional differences in ion concentration and/or electrical potential. If transcellular ion transport is electrogenic, it may cause a transepithelial voltage, which in turn may drive paracellular transport of other ion(s). There are several well-studied

70

Tight Junctions

FIGURE 4.3 (A) “Active” steady-state equivalent circuit of an epithelium with one cell type and a paracellular pathway of finite conductance. Each element in the circuit (a: apical membrane, b: basolateral membrane, s: paracellular pathway) is represented by a Thévenin electrical equivalent, i.e., an EMF, E, in series with a resistance, R. The Thévenin equivalent is a representation of any combination of linear electrical elements at the steady state. (B) Membrane voltages for an epithelium with the indicated values of EMFs (in mV) and resistances (in Ω·cm2). Polarities referred to basolateral solution (transepithelial values) or to adjacent solution (cell membrane values). Note that all voltages differ from the respective EMF values. (From Reuss, L., in Handbook of Physiology, Section 14, Cell Physiology, Hoffman, J. E. and Jamieson, J., Eds., Oxford University Press, New York, 1997. With permission.)

examples of this phenomenon. In Cl– transporting epithelia, both absorptive (such as the thick ascending segment of the loop of Henle) and secretory (such as enteric crypts), electrogenic Cl– transport generates a transepithelial voltage (negative polarity on the side into which Cl– is transported). This causes a parallel Na+ flux in the same direction as the Cl– flux, across the paracellular pathway (Silva et al., 1977; Greger and Schlatter, 1983; Greger, 1985; Sullivan and Field, 1991; Reeves and Andreoli, 1992). An instance in which transcellular transport causes changes in

Tight Junction Permeability to Ions and Water

71

substrate concentration that drive paracellular ion transport is anion transport in the late renal proximal tubule. In the early proximal tubule, an important fraction of – Na+ reabsorption is linked to HCO3 transport (Gottschalk et al., 1960; Rector et al., 1965). In the cell, the catalytic action of carbonic anhydrase causes production of H2CO3, a weak acid that partially dissociates in H+ and HCO3–. The H+ is secreted across the apical membrane in exchange for Na+ (Na+/H+ exchanger, NHE) and the HCO3– is cotransported with Na+ across the basolateral membrane. The reabsorption of NaHCO3 is accompanied by water reabsorption by osmosis and less-transported anions, such as Cl–, are thus concentrated in the lumen fluid. The increased [Cl–] generates a driving force for passive Cl– reabsorption via the paracellular pathway. 4.2.3.3 Osmotic Water Transport Net transepithelial solute transport reduces the osmolality of the fluid in the cis side and increases the osmolality of the fluid in the trans side. This difference in osmolality may cause water flow via the cell membranes and the junctions, according to their respective osmotic water permeabilities. The magnitude of the paracellular osmotic water flow in leaky epithelia is controversial (see below). 4.2.3.4 Solvent Drag In solvent drag (House, 1974; Finkelstein, 1987) there is net solute flux in the same direction as the net water flow, the latter driven by hydrostatic and/or osmotic forces. The coupling between solvent and solute fluxes is thought to be by frictional interaction between solute and solvent particles because of the Newtonian (viscous) nature of the fluid. Solvent drag is a consequence of net water flow and occurs only when the pathway has a finite permeability for water and solute. It can enhance transepithelial net solute transport because active salt transport can cause osmotic water flow and the latter can cause additional (passive) solute transport in the same direction. The magnitude of solvent drag in epithelia is uncertain, because the presence of unstirred fluid layers at the fluid–membrane interfaces makes such determinations difficult (House, 1974.).

4.2.4 THE JUNCTIONAL LOCATION OF THE HIGH-CONDUCTANCE PATHWAY IN LEAKY EPITHELIA WAS DEMONSTRATED WITH ELECTROPHYSIOLOGICAL AND MORPHOLOGICAL TECHNIQUES The electrophysiological analysis outlined above does not demonstrate the anatomic location of the paracellular pathway; i.e., its location could be particular cell type(s) or damaged portions of the epithelium, for example, at the edge of the epithelial sheet mounted in vitro. Direct demonstration that the junctions have high ionic permeability was obtained by electrophysiological and morphological techniques. The electrophysiological approach was to measure the current density normal to the epithelium in the proximity of the apical membranes and the junctions when large transepithelial currents were applied. Frömter (1972) demonstrated that in the epithelium of Necturus gallbladder the junctions are current sinks. Cereijido and co-workers confirmed these observations in Necturus gallbladder epithelium

72

Tight Junctions

(Cereijido et al., 1982) and extended them to epithelial monolayer cultures (Cereijido et al., 1980; 1981). The morphological approach was to test for junctional permeation of large molecules. Machen et al. (1972) showed that ionic lanthanum permeates the junctional complexes in mammalian gallbladder and small intestine. Ussing and Windhager (1964) showed that the frog epidermis, a tight epithelium, develops a high junctional conductance when exposed to hyperosmotic solutions on the apical side. Under these conditions, BaSO4 precipitated in junctions and lateral spaces when Ba2+ was added to the apical bathing solution and SO42– to the basolateral one. Transepithelial fluxes of hydrophilic nonelectrolytes are frequently used as an index of junctional leakiness (Mandel and Curran, 1972; Dawson, 1977; Madara and Dharmsathaphorn, 1985). In most cases the mannitol fluxes correlate well with junctional ion permeability, but the size of the junctional “pores” can vary among epithelia and it is possible that there are pores of different sizes in the same epithelium.

4.3 JUNCTIONAL ION SELECTIVITY The ion selectivity of tight junctions, i.e., the differential permeability of these structures to ions, is best studied by measuring the effects of unilateral ion substitutions on the voltage generated at the junctions. In leaky epithelia, this can be done by assessing transepithelial diffusion potentials. In tight epithelia, the preferred approach has been to measure transepithelial tracer fluxes. The diffusion potentials used to quantify the relative ion permeabilities of TJs and many other biological membranes are dilution potentials and bi-ionic potentials, both illustrated in Figure 4.4. The diagrams at the bottom depict how the potential differences are generated. The formulas relating concentrations, permeabilities, and voltage are model dependent (see Barry and Diamond, 1971; Barry, 1989). An equation describing the dilution potential for a single salt (monovalent cation and anion) is

[

]

Vdil = − [ RT F ] (uc − ua ) (uc + ua ) ln (C1 C2 )

(4.3)

where Vdil is the dilution potential (V1 – V2), uc and ua are the cation and anion relative mobilities, respectively (assuming identical activity coefficients), C1 and C2 are the salt concentrations on sides 1 and 2, respectively, and R, T, and F have their usual meanings (Barry, 1989). The bi-ionic potential, also for the case of monovalent ions is

[

Vbi = − [ RT F ] ln (uc + ua )1 (uc* + ua )2

]

(4.4)

where Vbi is the bi-ionic potential (V1 – V2), the cations are c on side 1 and c* on side 2. The same anion and identical salt concentrations are present on both sides (Barry, 1989). Figure 4.4 depicts measurements of dilution and bi-ionic potentials across frog gallbladder epithelium. In the panel on the left (dilution potential), the [NaCl] in the apical bathing solution was reduced at constant osmolality (sucrose replacement). This

M

Na Cl

+ _

Cl Na S

10 s

M Ch Cl

+ _

Cl Na S

20 mV

FIGURE 4.4 Typical records of transepithelial diffusion potentials in frog gallbladder. Continuous tracing of both records is transepithelial voltage (Vms). The tissue was bathed on both sides with Ringer’s solution, except during the intervals delimited by the arrows, during which the mucosal bathing solution was changed. Vertical voltage deflections are Vms changes (∆Vms) produced by transepithelial DC pulses. Current density was kept constant in each record; thus, changes in magnitude of ∆Vms denote roughly proportional changes in transepithelial resistance (Rt). (Left) 2:1 NaCl dilution potential. At first arrow, the mucosal solution was rapidly replaced with a solution in which [NaCl] was reduced to 50%, with other ion concentrations remaining constant, and sucrose added to maintain osmolality. Upward (mucosa-positive) change in Vms is accompanied by a slight increase in Rt. Changes are reversible upon return to control solution (second arrow). Since Na+ and Cl– are the only ions whose concentrations differ across the tissue, the polarity of the change in Vms indicates that PNa > PCl. Diagram below record is a simplified explanation of Vms change. (Right) Cho1ine-Na+ bi-ionic potential. Replacement of all Na+ in mucosal solution with choline (first arrow) is followed by a large Vms change (mucosa-positive) and a pronounced increase in Rt. Polarity of Vms change indicates that PNa > Pcholine. In the bottom parts of the figure, segmented lines denote smaller ion fluxes (see text for further details). (From Reuss, L., in Membrane Transport in Biology, Vol. IVB, Giebisch, G. et al., Eds., Springer-Verlag, New York, 1979. With permission.)

10 mV

Tight Junction Permeability to Ions and Water 73

74

Tight Junctions

causes an apical-positive change in transepithelial voltage (Vt). Circuit analysis (not shown) indicates that the change in Vt is predominantly originated at the junctions themselves and the polarity of the change indicates that junctional PNa > PCl. The magnitude of the voltage change is proportional to PNa /PCl. A quantitative estimate of this ratio requires circuit analysis based on simultaneous measurements of cell membrane voltages, Rt , and Ra /Rb (Reuss and Finn, 1975a,b). In the panel on the right, Na+ replacement with the impermeant cation choline caused a mucosa-positive change in Vt , i.e., the junctions are more permeable to Na+ than to choline ions. Again, calculation of relative permeabilities involves circuit analysis (Reuss and Finn, 1975a). An important issue in the measurement of diffusion potentials is that liquidjunction potentials occur at the interfaces between the voltage electrodes and the bathing solutions. These voltages do not reflect tissue properties, and must be minimized and/or corrected for (Barry and Diamond, 1970). In particular, the use of static Ringer–agar or 3 M KCl–agar bridges results in errors in the measurement or calculation of liquid-junction potentials because the compositions of these bridges are not constant when the solutions are changed. The author prefers flowing, saturated KCl junctions to minimize the junction potentials. The KCl dominates diffusion at the solution–bridge interface and the junction is stable and virtually independent of the ionic composition of the external solution. To prevent excessive KCl leak, Reuss and Costantin (1984) reduced the cross-sectional area of the tip of the bridge, kept the pressure difference (bridge–solution) low, and maintained a high rate of replacement of the bathing solutions. The changes in bathing solutions must be fast and brief, to prevent time-dependent secondary changes in ion composition in the cells or the contralateral extracellular compartment. Most leaky epithelia are cation selective (mammalian and amphibian gallbladder, mammalian small intestine and choroid plexus). In contrast, the amphibian renal proximal tubule is anion selective. Relative ion permeabilities of leaky epithelia are summarized in Table 4.2.

4.4 MECHANISMS OF JUNCTIONAL ION PERMEATION Extensive studies carried out in gallbladders of several species during the early 1970s provided a detailed characterization of the biophysics of ion permeation in leaky epithelia (for a review, see Moreno and Diamond, 1975b). Some of the major conclusions will be briefly summarized here, as well as a permeation hypothesis based on the recent discoveries of integral junctional proteins.

4.4.1 THE JUNCTIONAL ION PERMEATION PATHWAY CONSISTS OF PORES Junctional pores (aqueous communications between the solutions in the apical and lateral-space compartments) have not been directly demonstrated, either morphologically or electrophysiologically. However, this conclusion is based on solid indirect experimental observations. First, cation-selective leaky epithelia have permeability ratios for alkali metal cations typically within one order of magnitude. In contrast, pores

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75

TABLE 4.2 Paracellular Ionic Permeability Ratios of Epithelia Epithelium PT (rabbit) PT (dog) MDCK (dog) GB (rabbit) GB (Necturus) Jejunum (rat) Ileum (rabbit) UB (toad) Skin (frog)

PK /PNa — 1.1 1.2 2.3 1.8 1.2–1.6 1.1 1.4 1.3

PCl /PNa 0.8 0.7 0.1 0.3 0.3 0.1–0.2 0.4 0.7 4.5

Ref.a 1 * 2 * * * * * *

Abbreviations: PT = proximal renal tubule, MDCK = Madin–Darby canine kidney cells, GB = gallbladder, UB = urinary bladder. Note: Free-solution mobility ratios are PK /PNa = 1.47 and PCl /PNa = 1.52. a

References not given (*) can be found in Powell (1981). 1. Berry et al. (1978); 2. Cereijido et al. (1981).

in cell membranes have much greater selectivity. This result suggests that the permeation pathway consists of rather large aqueous pores. Second, the current–voltage relationship (measured at short times after the current or voltage pulses) is linear over a large range. This is unlikely for ion transport mediated by carriers or narrow pores. Third, junctional permeability to hydrophilic nonelectrolytes is graded (van Os et al., 1974), consistent with transport via a hydrated pathway that discriminates on the basis of size. The critical questions are the location and molecular identity of the pores. A definitive answer will not be available until the structure of the TJs at the level of atomic resolution is ascertained. Current knowledge is that claudins are the main components of the linear polymers identified in freeze-fracture studies as TJ strands, with an apparently secondary presence of occludin. (See Chapter 10.) This suggests that the barrier for transjunctional ion flux is formed by the claudin and occludin molecules contained in the strands. The pores could be hydrophilic pathways in between these molecules (see below and Figure 4.5). This hypothesis is particularly attractive because it suggests that specific claudin isoforms could determine the degree of tightness of the junctions and also their selectivity (see below).

4.4.2 THE JUNCTIONAL PERMEABILITY DEPENDS AND SIZE

ON

SOLUTE CHARGE

That TJs discriminate on the basis of size is based on determinations of permeability coefficients of hydrophilic nonelectrolytes and nitrogenous cations carried out in the

76

(A)

Tight Junctions

(B)

FIGURE 4.5 Proposed molecular bases of TJ ion permeability and selectivity. (A) Threedimensional view of TJ structure. The TJs are formed by strands consisting of linear polymers of integral membrane proteins (claudins, occludin, and perhaps others) and possibly lipids. The strand proteins bind laterally with the homologous structure of the adjacent cell, thus forming a continuous belt near the apical pole of each cell. In this diagram two parallel strands are shown. The strands are thought to contain aqueous pores that underlie paracellular permeation. Transcellular and paracellular pathways are depicted by long arrows (compare Figure 4.1A). (B) Two-dimensional scheme depicting the association of TJ strands at the molecular level. Three claudin isoforms are shown (Cld-1, Cld-2, and Cld-3). On the left, the putative TJ is formed by Cld-1 and Cld-3, yielding a structure without pores. On the right, the association of Cld-1 and Cld-2 generates a TJ with pores. The large number of claudin isoforms supports the view that the specific composition of the tight junctions can account for numerous permeability patterns, including a case of highly selective permeability to divalent cations (see text). (From Tsukita, S. and Furuse, M., J. Cell Biol., 149, 13, 2000. With permission.)

1970s (van Os et al., 1974; Moreno and Diamond, 1975a). In the latter studies it was shown that permeability decreases with increasing molecular size and increases with the number of donor protons; i.e., permeability is favored by hydrogen-bond formation. Extracellular acidification, elevation of [Ca2+], and the presence of polyvalent cations (Th4+, La3+) decrease the amplitude of dilution potentials across epithelia with cation-selective junctions. This effect results from both a decrease in cation conductance and an increase in anion conductance (Wright and Diamond, 1968; Machen and Diamond, 1972; Moreno and Diamond, 1974). The results are well explained by pores lined with dipoles with titratable negative sites oriented toward the lumen. The molecular basis of the selectivity of the putative TJ pores is not known, but a possibility is that the presence of different claudin isoforms in the strands results in the formation of pores of different diameters and charges (see Figure 4.5). This would account for the widely diverse selectivities of TJs of different epithelia. The discovery of paracellin-1/claudin-16 is the main argument in favor of this notion,

Tight Junction Permeability to Ions and Water

77

because this molecule is present in the TJs of cortical ascending loop of Henle and it is clearly responsible for the permeability of these junctions to divalent cations (Simon et al., 1999). What has not yet been demonstrated, however, is that paracellin-1/claudin-16 forms a channel. An interesting point when considering hypotheses of TJ structure is the width of the strands, ~10 nm. As pointed out by Tsukita and Furuse (1999) this size, by analogy with connexons (gap-junctional hemichannels), is consistent with claudin hexamers, complicating the simplified scheme shown in Figure 4.5. Resolution of this point will require the high-resolution structure of the TJs.

4.4.3 RELATIONSHIP OF JUNCTIONAL DEPTH AND JUNCTIONAL PERMEABILITY Claude and Goodenough (1973) first suggested that the junctional ion permeability is inversely correlated with the number of junctional strands encountered by the permeating ions in the transepithelial direction, although exceptions exist (MartínezPalomo and Erlij, 1975). As explained above, the paracellular resistance does not depend only on the specific resistance of the TJs. Other factors are the length of the network of intercellular clefts per unit area of epithelium, a parameter related to the cross-sectional area of the cells, as well as the length of the lateral intercellular spaces and the degree of interdigitation of the adjacent cells in the junctional region. Claude (1978) found that the value of Rt is an exponential function of the number of junctional strands. This relationship cannot be explained by an increase in the number of series resistors, but is consistent with strands containing channels that fluctuate between open and closed states (Claude, 1978; González-Mariscal et al., 1984; Cereijido et al., 1989; Reuss, 1992). The large effects of temperature on the transepithelial resistance of monolayers of MDCK cells (González-Mariscal et al., 1984) are consistent with this view. A requirement of this model is that in each strand the channels must be insulated from their neighbors. This insulation would be provided by the anastomoses between parallel strands (Cereijido et al., 1989; Reuss, 1992).

4.5 JUNCTIONAL WATER PERMEABILITY The transepithelial osmotic water permeability varies considerably among epithelia. In water-tight epithelia, e.g., thick ascending loop of Henle, renal collecting duct in absence of antidiuretic hormone (ADH), both the apical membrane and the junctions are virtually impermeable to water. ADH causes insertion of pores (aquaporin-2; Fushimi et al., 1993) in the apical membrane; thus its water permeability increases, but the junctions remain impermeable to water (Nielsen et al., 1995). An experimental result that was used to argue for a high junctional water Pos was the fact that in epithelia Pos is much higher than the diffusional water permeability (Pdw), i.e., Pos /Pdw  1. This result supports the conclusion that water permeation is via pores (Finkelstein, 1987), but does not indicate that their location is junctional. The discovery of the aquaporins (Agre et al., 1993), water pores expressed in plasma membranes, solved this issue. Certainly, demonstration of large-diameter pores would still be a good argument for junctional location.

78

Tight Junctions

As a rule, leaky epithelia have high transepithelial osmotic water permeabilities and transport salt and water in near-isosmotic proportions. The mechanism of water transport is probably local osmosis (Reuss, 1997; Spring, 1998) although there are other views (Loo et al., 1999). According to the local-osmosis theory, salt transport results in a difference in the osmolalities of restricted fluid compartments on both sides of the epithelium, and this difference elicits water transport in the same direction as salt transport. Water could in principle flow across the cells, the paracellular pathway, or both. Direct support for a role of the lateral intercellular spaces was obtained from microscopic measurements of the width of the spaces in the transporting Necturus gallbladder epithelium (Spring and Hope, 1979). For example, when NaCl was removed from the apical-bathing solution, the lateral intercellular spaces collapsed. However, demonstrating that the spaces are part of the transepithelial fluid transport pathway does not indicate that the TJs are water permeable. Part of the water moving via the cells could flow across the lateral membranes into the spaces. An argument against a significant junctional contribution to water transport is that the total surface area of the junctions is extremely small compared with that of the cell membranes (Spring, 1991). In addition, recent fluorescence microscopy studies suggest that there is no sizable paracellular water transport across confluent monolayers of MDCK cells (a renal cell line) (Kovbasnjuk et al., 1998), but this might not be the best experimental system to resolve this issue. Although local osmosis is favored by many, it has not been proved to be the water transport mechanism in epithelia. Several alternatives have been proposed (for reviews, see Reuss, 1997; Spring, 1998). More than 20 years ago, Diamond (1979) pointed out the missing information needed to understand the mechanism of epithelial water transport: (1) Measurements of transepithelial water fluxes and permeabilities with adequate time resolution and without unstirred-layer artifacts. (2) Assessment of relative values of transcellular and paracellular water fluxes. (3) Direct determinations of solute concentrations in the lateral spaces. Spring and associates (Spring, 1998) have made major efforts to address these questions using quantitativefluorescence and other sophisticated microscopic approaches (see Chapter 9).

4.5.1 RELATIONSHIP BETWEEN JUNCTIONAL ION AND WATER PERMEABILITY The high junctional ion permeability in leaky epithelia suggests that the junctions are also highly permeable to water. As summarized above, there is strong evidence that junctional ion permeation occurs via aqueous pores. If the pathways for ions and water were the same, then the measurement of the junctional electrical resistance would correlate with the junctional water permeability. Unfortunately, this is not the case. A first problem is that the TJs and the lateral intercellular spaces are in series, and both contribute to Rs (Kottra and Frömter, 1993). Although difficult, a distinction between junctions and spaces can be made electrophysiologically during experimental changes of Rs (Stoddard and Reuss, 1988), but quantification of these data is difficult. Another issue is that in renal tubules there is no correlation between the transepithelial resistance and the osmotic permeability coefficient (Pos ) (Berry, 1983). An extreme

Tight Junction Permeability to Ions and Water

79

case is that of the thick ascending loop of Henle, which has a high paracellular conductance (Hebert et al., 1984) and a vanishingly low Pos (Rocha and Kokko, 1973).

4.5.2 JUNCTIONAL PERMEATION

OF

LARGE HYDROPHILIC SOLUTES

The demonstration that hydrophilic nonelectrolytes such as mannitol permeate leaky epithelia (e.g., Madara and Dharmsathaphorn, 1985) is one of the most convincing arguments for the existence of junctional pores. Mannitol, because of its relatively large size and low oil/water partition coefficient, cannot permeate cell membranes. If the paracellular pathway for mannitol were the same as for ions and water, then the three fluxes would be linearly related and the problem of junctional permeability would be resolved from the biophysical point of view. However, several arguments indicate that this is not the case. First, in cell membranes there is no necessary correlation between ion and water permeation (see above). Second, detailed studies of nonelectrolyte permeability in leaky epithelia are not consistent with a single pore radius, but support the idea of pore heterogeneity, with abundant small-radius and scarce large-radius pores (van Os et al., 1974). As discussed above in the context of junctional permeation of ions, pore heterogeneity in size and charge can be a direct result of what claudin isoforms contribute to forming each pore.

4.5.3 ESTIMATES

OF JUNCTIONAL

WATER PERMEABILITY

To assess cellular vs. paracellular water fluxes, a rather obvious approach is to measure the Pos before and after blocking specifically one of the pathways. No demonstrably specific blockers have been found, however, so studies with mercurials to block cell-membrane water pores and polyvalent cations to block paracellular water flow should be considered inconclusive (for further discussion, see Spring, 1998). With the recent advances in knowledge of the chemical composition of TJs, there is great hope that specific blockers will be developed. In the absence of adequate blockers, no direct measurements of junctional water permeability are available (Spring, 1998). Indirect estimates are based on a circuit analysis that involves comparisons between the transepithelial and transcellular permeabilities and assumes that the junctions are the main barrier for paracellular water flow. Inasmuch as the cells and the junctions are in parallel, the transepithelial osmotic water permeability (Post ) is given by Post = Posc + Posp

(4.5)

where Posc = Pos of the cellular pathway and Posp = Pos of the paracellular pathway. If Post and Posc are experimentally determined, then Posp can be calculated from Equation 4.5, i.e., Posp = Post – Posc . Values of Post and Posc in renal proximal tubule and in Necturus gallbladder epithelium are presented in Table 4.3. The hydraulic water permeability (Lp) can be assessed from the transepithelial water flow (Jv) in response to a known difference in effective osmotic pressure (σ∆π), where σ is the reflection coefficient of the solute and ∆π is the difference in total osmotic pressure:

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TABLE 4.3 Cell Membrane and Transepithelial Osmotic Water Permeabilities Pos , cm·s–1 Epithelium PCT (rat) PCT (rabbit) PST (rabbit) RC (rabbit) PT (Necturus) PT (Ambystoma) GB (Necturus)

Transepithelial 0.12–0.30 0.04–0.35 0.20–0.70 — 0.02–0.03 — 0.04

Apical — — 0.13 0.40 — 0.007 0.04–0.06

Basolateral — 0.23–0.60 0.14–0.32 0.50–0.60 — 0.008 0.05–0.12

Ref.a * * *,** ** * 1 2–4

Abbreviations: PT = proximal renal tubule, PCT = proximal convoluted tubule, PST = proximal straight tubule, RC = renal cortex vesicles (brush border or basolateral membranes), GB = gallbladder. Note: Membrane vesicle Pos is expressed per unit membrane area, whereas all other Pos estimates are relative to nominal basal surface area (not corrected for membrane foldings). a

References not given can be found in Berry, 1983 (*) or Reuss and Cotton, 1988 (**). 1. Tripathi and Boulpaep (1988); 2. Persson and Spring (1982); 3. Zeuthen (1982); 4. Cotton et al. (1989).

Jv = Lpσ∆π

(4.6)

Lp [cm2 s–1 (osmol/kg)–1] is related to Pos by the following relationship: Pos = Lp

RT Vw

(4.7)

– where R and T have their usual meanings and Vw is the partial molar volume of water. The main difficulty in measuring Post is the existence of unstirred layers in series with the epithelium (Barry and Diamond, 1984). It is difficult to eliminate them, correct for their effects, or make rapid measurements of transepithelial water transport. Although the anatomical thickness of these layers is small in isolated perfused tubules, their effect is exacerbated when the presence of membrane folds (such as microvilli and basolateral membrane infoldings) causes funneling of water flow, increasing the flow velocity (Barry and Diamond, 1984). In planar epithelia, such as gallbladder, the errors introduced by unstirred layers are large. Therefore, reported Post values are quite uncertain. The cell-membrane Pos has been estimated from net water fluxes (i.e., volume changes) upon changing external osmolality. Three approaches have been used.

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1. Optical techniques, such as light scattering, in membrane vesicle preparations (Verkman, 1989); 2. Optical measurements of cell height or, using optical sectioning, reconstruction of cell volume in epithelia mounted in vitro (Persson and Spring, 1982); 3. Electrophysiological measurements of the intracellular concentration of ions, which are essentially impermeant and therefore behave as volume markers (Zeuthen, 1982; Reuss, 1985; Cotton and Reuss, 1989; Cotton et al., 1989). In the latter approach, the change in cell volume and the osmolality of the solution at the apical surface of the cell were assessed simultaneously. Results in renal tubule segments and in amphibian gallbladder epithelium are summarized in Table 4.3. In Necturus gallbladder, given the uncertainty in the determination of Post , the results were deemed consistent with a predominant transcellular water permeation pathway (Cotton et al., 1989), but this does not rule out a sizable paracellular Pos. In proximal renal tubule, it has been argued that Post is significantly higher than Posc , suggesting that Posp is significant (Hill, 1980; González et al., 1984; Tripathi and Boulpaep, 1988). Again, because of the uncertainties of the experimental methods, this conclusion is uncertain. Kobvasnjuk et al. (1998) performed fluorescence microscopy studies in MDCK cells that allowed the measurement of flow-velocity profiles in the lateral intercellular spaces. The flow velocity was near zero in the vicinity of the junctions. These results do not support the notion of sizable paracellular water transport. A problem, however, is that this may not be the most appropriate epithelium to test the hypothesis, i.e., a clearly leaky epithelium with high-permeability junctions should be considered. At this time, the issue of the degree and significance of junctional water transport remains unresolved (Reuss, 1997; Spring, 1998).

4.5.4 SOLUTE–SOLVENT COUPLING ELECTROKINETIC PHENOMENA

AND

Water flow via large aqueous pores is expected to “drag” permeant solutes by frictional interaction between solvent and solute. This phenomenon is solvent drag, and has been claimed to exist in both proximal tubule (Whittembury et al., 1980) and gallbladder (Hill and Hill, 1978a,b). In both preparations there were positive correlations between the water and solute fluxes. Also, the solute fluxes are inversely proportional to molecular size. An alternative explanation for these results is “pseudo-solvent drag” (Diamond, 1979; Barry and Diamond, 1984). If the osmotic water flow causes changes in solute concentrations in the unstirred layers (increase on the cis and decrease on the trans side), then simple diffusion could explain a net solute flux via a separate pathway. Solvent drag and pseudo-solvent drag could be distinguished by measuring fluxes of hydrophilic and lipophilic solutes under the same experimental conditions (Diamond, 1979). Pseudo-solvent drag would elicit fluxes of both kinds of solutes, whereas true solvent drag would cause the flux of

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hydrophilic solutes but not of lipophilic solutes, because the latter flux is predominantly transcellular. Apparent demonstrations of solvent drag do not prove that the TJs have a high Pos. Streaming potentials are another effect of frictional coupling between water and ion fluxes. In the interior of membrane pores bearing net charge, the concentration of counterions is higher than the concentration of co-ions. Osmotic water flow drags counterions and elicits a change in membrane voltage. For cation-selective pores, the hyperosmotic side becomes electrically positive (Pidot and Diamond, 1964). An alternative explanation of this experimental observation is that it corresponds to a “pseudo-streaming potential.” Transepithelial osmotic water flow causes changes in salt concentration in the fluid layers adjacent to the epithelium. The resulting change in transepithelial voltage is thus a “dilution potential” and not a true streaming potential (Wedner and Diamond, 1969). The author’s group (Reuss et al., 1992a,b) carried out detailed studies of the changes in transepithelial electrical potential elicited by sucrose osmotic gradients across amphibian gallbladder epithelium (apparent streaming potentials). The time course was compared with that of a biionic potential obtained from isosmotic NaCl substitution using a salt of similar diffusion coefficient to that of sucrose. These studies revealed that the apparent streaming potential is of paracellular origin, and that its time course is too slow compared with that of the diffusion potential, indicating that it results from transepithelial ion-concentration gradients secondary to the osmotic water flow (pseudostreaming potential). This work described a novel biophysical method to distinguish true and pseudo-streaming potentials. In summary, recent estimates of the values of cell membrane Pos in leaky epithelia clearly indicate that both apical and basolateral membranes have high water permeabilities and constitute an important, if not dominating, pathway for transepithelial osmotic water flow. Nevertheless, the contribution of junctional water permeation to transepithelial spontaneous water transport or to the water flow induced by imposed osmotic gradients remains unresolved. Calculations of Posp , from Post and Posc are inconclusive, and indirect arguments based on assessment of apparent coupling between solvent and solute fluxes are difficult to interpret because of complications imposed by the presence of anatomical and functional unstirred layers. It appears that the problem of the water permeability of the TJs will not be solved until direct measurements are made using new techniques.

4.6 CONCLUSIONS Biophysical studies of TJ permeability have succeeded in providing a reasonable functional picture of these important structures. The junctions are transepithelial permeation barriers of diverse degree of tightness, as well as different permeabilities to inorganic ions and larger solutes. There is no strict correlation between ion and water permeabilities, suggesting that at least in some cases different molecules may underlie ion and water permeation. The recent progress in the identification of integral junctional proteins has opened up the possibility of studying the basis of junctional permeability at the molecular level. Finding out the high-resolution structure of TJs is the main challenge ahead.

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Tsukita, S. and Furuse, M. 1999. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol., 9, 268. Tsukita, S. and Furuse, M. 2000. Pores in the wall: claudins constitute tight-junction strands containing aqueous pores. J. Cell Biol., 149, 13. Tsukita, S., Furuse, M., and Masahiko, I. 1999. Structural and signalling molecules come together at tight junctions. Curr. Opin. Cell Biol., 11, 628. Ussing, H. H. and Windhager, E. E. 1964. Nature of shunt path and active solute transport path through frog skin epithelium. Acta Physiol. Scand., 61, 484. Ussing, H. H. and Zerahn, K. 1951. Active transport of sodium as the source of electric current in the short-circuited isolated frog skin. Acta Physiol. Scand., 23, 110. van Os, C. H., de Long, M. D., and Slegers, J. F. G. 1974. Dimensions of polar pathways through rabbit gallbladder epithelium. The effect of phloretin on nonelectrolyte permeability. J. Membr. Biol., 15, 363. Verkman, A. S. 1989. Mechanisms and regulation of water permeability in renal epithelia. Am. J. Physiol., 257, C837. Wedner, H. J. and Diamond, J. M. 1969. Contributions of unstirred-layer effects to apparent electrokinetic phenomena in the gall-bladder. J. Membr. Biol., 1, 92. Whittembury, G. and Reuss, L. 1992. Mechanisms of coupling of solute and solvent transport in epithelia, in The Kidney: Physiology and Pathophysiology, Seldin, D. W. and Giebisch, G., Eds., Raven Press, New York, chap. 13. Whittembury, G. et al. 1980. Solvent drag of large solutes indicates paracellular water flow in leaky epithelia. Proc. R. Soc. Lond., 211, 63. Windhager, E. E., Boulpaep, E. L., and Giebisch, G. 1967. Electrophysiological studies in single nephrons, in Proceedings of the Third International Congress on Nephrology, Schreiner, G. E., Ed., Washington, D.C., 1966, Vol. 1, Karger, New York, 35–47. Wong, V. and Goodenough, D. A. 1999. Paracellular channels. Science, 285, 62. Wright, E. M. and Diamond, J. M. 1968. Effects of pH and polyvalent cations on the selective permeability of gallbladder epithelium to monovalent ions. Biochim. Biophys. Acta, 163, 57. Yonath, I. and Civan, M. M. 1971. Determination of the driving force of the Na+ pump in toad bladder by means of vasopressin. J. Membr. Biol., 5, 366. Zeuthen, T. 1982. Relations between intracellular ion activities and extracellular osmolarity in Necturus gallbladder epithelium. J. Membr. Biol., 66, 109.

5

The Relationship Between Structure and Function of Tight Junctions Lorenza González-Mariscal, Antonia Avila, and Abigail Betanzos

CONTENTS 5.1 5.2

5.3

5.4 5.5

5.6

5.7

Introduction ....................................................................................................90 Ultrastructural Features of Tight Junctions ...................................................90 5.2.1 Thin Section .......................................................................................90 5.2.2 Freeze Fracture...................................................................................91 The Molecular Nature of Tight Junction Strands..........................................93 5.3.1 Occludin .............................................................................................95 5.3.1.1 Structure–Function Properties of Occludin........................95 5.3.2 Claudins..............................................................................................98 5.3.2.1 Structure–Function Properties of Claudins ........................98 The Molecular “Fence” and Paracellular “Gate” Functions of Tight Junctions.........................................................................................................99 The Gate Function of Tight Junctions.........................................................100 5.5.1 The Passage of Tracers through the Paracellular Pathway .............100 5.5.2 The Electrical Resistance of the Tight Junction..............................100 5.5.3 Classification of Epithelia as “Tight” or “Leaky”...........................102 5.5.4 Morphological Aspects That Affect TER ........................................103 5.5.5 The Relationship between the Specific Resistance of the Tight Junction (Rj lp) and the Number of Its Strands ................................104 Presence of Pores or Channels Within the Tight Junction Strands ............107 5.6.1 Nephrin, a Porous Filter of the Podocyte Slit Diaphragm..............108 5.6.2 Paracellin, a Mg2+/Ca2+ Channel of Tight Junctions .......................108 The Electric Circuit Analysis as a Tool for Predicting Epithelial Resistance.....................................................................................................109 5.7.1 In a Natural Epithelia Formed by Different Types of Cells ...........109 5.7.2 Between Epithelia Derived from Different Animal Species ...........110

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

89

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5.7.3 5.7.4

During the Establishment of Epithelial Monolayers.......................110 Between Cells That Significantly Differ in Their TER in Spite of Belonging to the Same Cell Line................................................110 5.7.5 In Mixed Monolayers Formed by Cells Derived from Different Animal Species ................................................................................111 5.8 The Permeability of the Tight Junction Can Be Modified Experimentally without Concomitant Changes in the Arrangement of the Strands ............113 5.9 Changes in the Phosphorylation State of Tight Junction Components May Affect the Permeability of the Epithelium..........................................113 5.10 Concluding Remarks....................................................................................113 Acknowledgment ...................................................................................................114 References..............................................................................................................114

5.1 INTRODUCTION The establishment of particular environments in opposing compartments separated by epithelial or endothelial sheets is made possible by the presence of tight junctions (TJs). The function of the TJ is now envisioned as double: a “fence” that prevents the free diffusion of proteins and lipids between apical and basolateral cell surfaces and a “gate” that regulates the passage of ions and molecules through the paracellular pathway. In the last decade, the discovery of several molecules that make up the TJ has dramatically changed the perception of this structure. To date, several peripheral (ZO-1, ZO-2, ZO-3, cingulin, rab3b, symplekin, 7H6, AF6, and ASIP) and integral (claudins, occludin, and JAM) membrane proteins have been identified as TJ constituents. However, the relationship between transepithelial electrical resistance (TER), paracellular permeability, the expression of TJ proteins, and the number and distribution of TJ strands is not simple, and several elements need to be considered. This chapter reviews the TJ structure–function relationship, considering the TJ as a part of an electric circuit, and takes into account the new information provided by the proteins that constitute it.

5.2 ULTRASTRUCTURAL FEATURES OF TIGHT JUNCTIONS 5.2.1 THIN SECTION The TJ appears in thin sections, as a zone in which the plasma membranes are closely apposed (Faquhar and Palade, 1963). This area, 100 to 800 nm in depth, surrounds the cell like a belt at the limit between the apical and the lateral membrane. At the TJ, the lateral membranes appear to fuse at certain points, informally known as kisses. OsO4-stained plasma membranes appear by transmission electron microscopy (TEM) as a structure with dark–light–dark layers, that correspond, respectively, to the polar heads, hydrophobic tails, and polar heads of the phospholipids that constitute the lipid matrix of biological membranes. However, at the kiss region of the TJs, five and not six layers (three from each membrane) are distinguishable, because at these

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points the external polar groups are so close together, that they appear to fuse and exclude the extracellular space. These kisses constituted by the strands and grooves seen on freeze-fracture replicas impose a rate-limiting barrier to passive permeation through the intercellular space. In fact, when electron-opaque exogenous tracers such as lanthanum, horseradish peroxidase, and ruthenium red are added, their diffusion is stopped by the TJ kisses (Reese and Karnovsky, 1967; Machen et al., 1972; Martínez-Palomo and Erlij, 1973; González-Mariscal et al., 1985) (Figure 5.1A). Alterations in the thin section appearance of TJs can be observed under physiological conditions, during the process of intestinal absorption of sugars, peptides, and amino acids after a meal. The high concentration of nutrients in the intestinal lumen activates the sodium coupled transport of glucose and amino acids into the cell. The increased activity of intracellular Na+ activates the Na+, K+-ATPase, which in turn enhances the extrusion of Na+ to the intercellular lateral space, thus providing the osmotic force for absorption of fluid through the paracellular pathway. Sodiumcoupled solute transport also triggers contraction of the perijunctional actinomyosin ring, resulting in increased permeability of TJ and expansion of lateral spaces, providing optimal conditions for transport of luminal nutrients involved by solvent drag. By thin section, this physiological change in TJ permeability correlates with the appearance of numerous intrajunctional dilatations or blebs, which range from less than 0.1 to 0.5 µm in width. These blebs correspond to expanded interstrand compartments in freeze-fracture replicas (Figure 5.2) (Madara and Pappenheimer, 1987; Pappenheimer and Reiss, 1987).

5.2.2 FREEZE FRACTURE In the freeze-fracture procedure membranes break along the weakest point, i.e., the central hydrophobic plane. Therefore, freeze-fracture images do not show a true membrane surface but the fractured membrane face. TJs studied with this technique appear as a series of strands and grooves, which anastomose to form a meshwork that circumvents the cell below the microvilli (Figure 5.1B). The fracture face associated with the cytoplasmic leaflet of the plasma membrane (P face) generally reveals strands, whereas the exoplasmic leaflet (E face) usually reveals grooves (Bullivant, 1978). Strands and grooves are correspondent, as revealed by studies of complementary replicas (van Deurs and Koehler, 1979). The appearance of TJs by freeze fracture is dependent on the methods utilized to prepare the tissues. Thus, without aldehyde fixation TJ strands appear more as rows of E face particles, than as continuous P face strands (van Deurs and Luft, 1979). It has been hypothesized that there is an important functional role for the P face association of TJ strands, and thus the cytoplasmic anchoring of TJ particles. For example, endothelial cells that form the blood–brain barrier exhibit a high electrical resistance and their TJ strands associate with the P face (Wolburg et al., 1994), whereas peripheral nonbarrier endothelial cells that display a low resistance have TJ particles associated with the E face (Table 5.1) (Simionescu et al., 1988; Mühleisen et al., 1989). Linkage of TJ particles to the membrane appears to be environmentally regulated since cAMP and astrocyte-conditioned medium induce association of particles to the P face in cultured brain endothelial cells (Wolburg et al., 1994).

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FIGURE 5.1 TJ morphology. (A) Transmission electron microscopy of two adjacent epithelial cells (MDCK). The TJ (arrow) stops free diffusion of ruthenium red added to the apical side. (B) Freeze-fracture replica of an MDCK monolayer shows the belt of junctional strands that separates the apical from the lateral side.

Several distinct models of TJ organization have been proposed (Figure 5.3): 1. The Chalcroft and Bullivant (1970) two-fibril model, in which fibrils composed of particles in the membrane of one cell are in direct register with similar fibrils in the opposing membrane. As seen in Figure 5.3 the fracture plane is presumed to pass around the fibril in one membrane, but

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FIGURE 5.2 Schematic representation of a TJ dilatation (arrow) induced by perfusion with glucose. N = nucleus.

not to include the adjoining fiber in the neighboring membrane. Thus, TJ particles are considered to be more strongly bound to the P halves of their membrane than to their partners. 2. The Wade and Karnovsky (1974) single-fibril model, which proposes that both membranes share a single set of fibrils. Therefore, the fracture plane includes the fibril from the opposing membrane, leaving a large strand on the P face. 3. The Staehelin model (Staehelin, 1973), which considers that the row of particles in one membrane is directly in register with a similar row in the opposing membrane. The particles are proposed to be so strongly bound to their partners that they stay together as a unit upon fracture, appearing as a unit on the P face. However, in TJs opened by treatment with hypertonic saline or EGTA, the height of the P face strands appeared similar to that of sealed junctions (Hirokawa, 1982), thus supporting the two fibril model and providing evidence against Staehelin’s model, which predicted that the height of the P face strands in closed TJs should be twice as high as those on the P face of split junctions. 4. The single-fibril model was later modified by Bullivant (Bullivant, 1978), whose evidence proposed that the TJ fibrils are not in direct register, but slightly offset. The discovery in the last decade of several TJ strand components at the membrane of both cells that establish the junctional seal has further validated the two fibril model.

5.3 THE MOLECULAR NATURE OF TIGHT JUNCTION STRANDS The biochemical nature of TJs remained elusive for a long time. Almost 20 years ago, cylindrical inverted lipid micelles were proposed as the main constituents of TJ strands (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982). However, the detergent stability shown by the strands suggested instead a main proteic nature

94

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TABLE 5.1 Relationship between the Freeze-Fracture Appearance of TJs and TER in Cells Subjected to Different Experimental Procedures That Modify the TJ (P and E indicate the membrane fracture face)

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of TJ filaments (Stevenson and Goodenough, 1984). Furthermore, when the total composition of phospholipids, sphingolipids, cholesterol, and the content of fatty acids was experimentally changed in MDCK cells, no significant alteration in TER or the structure of the TJ strands was detected (Schneeberger et al., 1988; Calderón et al., 1998). Recently phospholipids derivatives have been successfully employed to enhance the paracellular absorption of drugs across epithelia (Dong-Zhou et al., 1999a,b). However, rather than acting upon the strands, these phospholipids seem to alter the intracellular mechanisms that regulate the junctional complex. Conventional freeze fracture provided crucial knowledge on the appearance of TJ strands; however, it could not offer information regarding their biochemical nature. The recent development of SDS-digested freeze-fracture replica labeling (SDS-FRL) permitted instead immunocytochemical detection of transmembrane molecules, and therefore, for the first time, provided the opportunity to detect proteins in TJ strands (Fujimoto, 1997).

5.3.1 OCCLUDIN In 1993, Tsukita’s group identified occludin, the first integral membrane protein of the TJ. This molecule has four transmembrane domains, two extracellular loops, and a long COOH-terminal cytoplasmic tail (Furuse et al., 1993). When this protein was overexpressed in baculovirus-infected insect Sf9 cells, multilamellar bodies bearing TJ-like structures accumulated in the cytoplasm. Analysis of these structures by SDS-FRL showed intense immunogold labeling of these TJ-like strands with an antioccludin antibody (Furuse et al., 1996). Moreover, this technique allowed the detection of occludin in the TJ strands of hepatocytes, thus demonstrating for the first time the proteic nature of TJ filaments (Fujimoto, 1995; Furuse et al., 1996). 5.3.1.1 Structure–Function Properties of Occludin Occludin has fence and gate properties. The former, for clarity, will be discussed in Section 5.4; the latter are supported with the following evidence: 1. Transfected occludin, either full length or COOH-terminally truncated, increases TER and the paracellular flux of small-molecular-weight tracers (for explanation of this apparently paradoxical result, see Section 5.5.5) (Balda et al., 1996; McCarthy et al., 1996; Chen et al., 1997). 2. A peptide corresponding to the second extracellular loop (Wong and Gumbiner, 1997) and a mutant occludin lacking both the cytoplasmic N terminus and both extracellular loops decrease TER and augment the paracellular passage of tracers (Bamforth et al., 1999). 3. In endothelial cells treated with anticancer polyunsaturated fatty acids, an increased tightness of TJs is coupled to occludin upregulation (Jiang et al., 1998). 4. Occludin mRNA is high in tissues with well-developed TJ or epithelioid phenotype but not detectable in fibroblasts (Saitou et al., 1997) and differentiated astrocytes (Bauer et al., 1999).

96

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FIGURE 5.3 Models of the intramembrane structure of TJs revealed by freeze-fracture analysis. Thick dashed lines indicate the fracture plane. The membrane domains above the thick line are removed during freeze fracture and a replica of the surface of the remaining region (speckled area) is obtained. The arrowheads indicate the transitional area, where the fracture passes from a protoplasmic (P) to an exoplasmic (E) face. M = membrane; i = intercellular space.

5. Endothelial cells of non-neural tissues, with a considerable number of leaky junctions, express a significantly lower amount of occludin than those of the blood–brain barrier, with tighter TJs. Moreover, occludin expression is developmentally coupled to maturation of brain endothelia (Hirase et al., 1997). 6. Occludin in endothelial monolayers is more concentrated in arterial than in venous junctions, correlating with the arterial lower permeability (Kevil et al., 1998). 7. The increase in TER, from the proximal to the collecting duct of the mammalian nephron, is paralleled by an enhanced occludin expression (González-Mariscal et al., 2000). 8. In the small intestine, 73% of the paracellular conductance along the cryptvillus is attributable to the crypt that contains comparatively fewer TJ

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97

FIGURE 5.3 (continued).

strands (Marcial et al., 1984). In conformity, occludin appears as puncta at the borders of the crypt epithelium and as a honeycombed network around the villus cells (Westcarr et al., 1999). 9. In epithelial cells transfected with oncogenic Raf, introduction of the occludin gene results in reacquisition of a monolayer phenotype, formation of functionally intact TJs, relocation of other TJ proteins to the cell borders, and inhibition of anchorage-independent growth (Li and Mrsny, 2000). Therefore, there is compelling evidence to support the importance of occludin in TJs. However, the following recent and unexpected results have led to reconsideration of the role that this molecule plays as a structural component of TJ strands: 1. Occludin-deficient embryonic stem cells differentiated into polarized epithelial cells with well-developed TJ strands (Saitou et al., 1998). 2. Hepatocytes treated with an actin depolymerizing agent display normal TJ strands in freeze-fracture replicas; yet actin and occludin have disappeared from their cell borders (see Table 5.1) (Kojima et al., 1999).

98

Tight Junctions

The presence of apparently normal TJ strands without occludin immediately suggested the existence of other integral proteins capable of forming strand structures, and assigned occludin a more regulatorial than structural role at the TJ.

5.3.2 CLAUDINS In 1998, claudin-1 and claudin-2 were identified as components of TJ strands. These 23-kDa integral membrane proteins, with four transmembrane domains, bear no sequence similarity to occludin, and when introduced into epithelial cells, they incorporated into preexisting TJ strands (Furuse et al., 1998a). When claudin-1 and claudin-2 were introduced into fibroblasts, they formed well-developed networks of strands, morphologically similar to native TJs. In contrast, fibroblasts transfected with occludin formed only small numbers of short and straight TJ strands (see Table 5.1). In cells double-transfected with occludin and claudin-1, both molecules were admixed along the strands, thus suggesting their ability to copolymerize into TJ strands (Furuse et al., 1998b). Similarity searches through databases identified the existence of several sequences similar to claudin-1 and claudin-2, thus giving rise to the so-called claudin family. All the members have similar hydrophilicity plots, and when claudin-3 to claudin-8 were introduced into MDCK cells they concentrated at preexisting TJ strands (Morita et al., 1999a). Based on sequence similarity, seven more cDNAs have tentatively been designated as claudins 9 to 16 (Tsukita and Furuse, 1999; Simon et al., 1999). 5.3.2.1 Structure–Function Properties of Claudins As stated above, a correlation between the number of TJ strands, TER, and the level of expression of occludin has been found. However, with claudins, a more complex picture is emerging, since each tissue expresses a different set of claudins. Thus, for example, while the kidney has most types of claudins, the testis and the central nervous system (CNS) have a very low expression level of claudins 1 to 8. These tissues, on the other hand, are particularly enriched in claudin-11/oligodendrocytespecific protein (OSP). This claudin is the main component of the TJ strands in oligodendrocytes and Sertoli cells (Morita et al., 1999b). As expected, OSP null mice exhibit neurological deficits, are sterile, and no TJs are found on the freezefracture replicas of their CNS myelin and Sertoli cells (Gow et al., 1999). These results therefore assert that claudins constitute the main framework of TJ strands. The amount of claudins in a particular epithelia appears to be related to its tightness. Thus, in MDCK cells overexpression of claudin-1 significantly increases TER and reduces paracellular flux of dextrans (Inai et al., 1999). Furthermore, in MDCK cells, treatment with Clostridium perfringens enterotoxin (CPE) destroys TER with a concomitant increase in paracellular flux, apparently through its deleterious action over claudin-4. The direct involvement of claudins in the gate functions of TJs is further demonstrated in fibroblasts where CPE disintegrates TJ strands constituted by transfected claudins 3 or 4 (Sonoda et al., 1999). In addition to the amount of claudins, the tightness of a particular tissue might also depend on the species of claudins involved and their mixing ratio within the

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strands. It is noteworthy that strands formed with claudins 1 and 3 are largely associated with the P face as continuous structures, whereas claudins 2 and 5 (an exclusively endothelial claudin) are discontinuous at the P face with complementary grooves at the E face occupied by chains of particles (see Table 5.1) (Furuse et al., 1998b; 1999; Morita et al., 1999c). Since tight and leaky TJs show P and E face association of particles, respectively, it is tempting to speculate that TJ strands consisting of claudins 1 and/or 3 are tighter than those with claudins 2 and/or 5. Different claudin species can interact within and between TJ strands, except in certain combinations (Furuse et al., 1999). For example, claudin-1 strands do not interact with claudin-2 strands, although they copolymerize within a strand in transfected fibroblasts. Therefore, this result suggests the existence of a relative degree of partnership selectivity between claudins and possibly indicates that not all claudin mixtures actually exist in natural epithelia. Claudins lacking their COOH terminal cytoplasmic domain, necessary for ZO-1, ZO-2, and ZO-3 binding (Itoh et al., 1999), still bear well-developed networks of strands (Furuse et al., 1999). Thus, interaction between claudins and peripheral membrane proteins is apparently not required for the formation of TJ strands and their face association. However, the ability of these deleted claudins to act as a gate has not yet been explored.

5.4 THE MOLECULAR “FENCE” AND PARACELLULAR “GATE” FUNCTIONS OF TIGHT JUNCTIONS Although the gate and fence functions of TJs appear simultaneously during epithelial morphogenesis, it has been possible to separate them with an energy depletion procedure. Although this treatment abolishes gate function, discerned by a dramatic decrease in TER, it leaves the fence function intact as determined by the blockade in the diffusion toward the basolateral membrane of lipid probes introduced in the apical region (Mandel et al., 1993) However, lipid probes inserted into the inner membrane leaflet (van Meer and Simons, 1986) and others able to flip-flop (Dragsten et al., 1981; Spiegel et al., 1985), can diffuse to the opposite surface domain; the former presumably by free diffusion while the latter by shunting the TJ diffusion constraint through a flip-flop to the inner membrane leaflet. These results imply that the TJ may form a “fence” to the passage of lipids only in the outer membrane leaflet. The functional gate–fence separation of the TJ emerges as a result of two different types of molecular interactions: the fence function dependent on the connection between the particles forming the strands within each cell, while the gate relies on the contact between strands located on apposing cells. Molecular manipulation of TJ proteins has also supported the dual gate–fence activity of the junction. Epithelial cells transfected with a COOH-terminally truncated occludin incorporate this mutant occludin into TJs in a discontinuous pattern and display an increased TER. However, the structural properties of TJs responsible for the electric seal or gate function appear to be different from those required for the intramembrane fence, since cells expressing this altered occludin are no longer capable of preventing the diffusion of a fluorescent lipid from one cell surface domain

100

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to the other. Expression of this carboxyl-truncated occludin does not affect the polarity of proteins, indicating that lipid polarity is more sensitive to alterations in the fence function of the TJ (Balda et al., 1996). This might be due to the fact that interactions between membrane proteins and the cytoskeleton restrict lateral mobility and that many mobile membrane proteins possess bulky extracellular domains, whose passage through the paracellular pathway will be restricted by the extracellular TJ domains. Therefore, occludin emerges as a structural component of the TJs that physically forms the intramembrane diffusion fence.

5.5 THE GATE FUNCTION OF TIGHT JUNCTIONS 5.5.1 THE PASSAGE OF TRACERS THROUGH THE PARACELLULAR PATHWAY The transport of soluble tracers such as [H3]-mannitol, horseradish peroxidase, inulin, or fluorescent dextran across epithelia occurs along a transcellular route (i.e., transcytosis) and by passive transport along a concentration gradient through the paracellular pathway. The relative contribution of each route to the total transepithelial flux depends on the physical characteristics of the particular marker. Since TJ “pores” have charge and size selectivity (see below), small neutral tracers are chosen for measurements of paracellular permeability. This experimental approach has been used to study junction formation triggered by calcium (González-Mariscal et al., 1990) and to determine the role that claudins (Inai et al., 1999; Sonoda et al., 1999) and occludin (Balda et al., 1996; McCarthy et al., 1996) play on the regulation of the paracellular flux.

5.5.2 THE ELECTRICAL RESISTANCE

OF THE

TIGHT JUNCTION

In 1978, Claude (1978) proposed a model (Figure 5.4) in which the total transepithelial resistance (Rt) is represented by two resistances in parallel: (1) the transcellular resistance (Rc) and (2) the paracellular resistance (Rp). The transcellular pathway is in turn resolved in two resistances in series, one due to the apical membrane (Ram) and the other due to the basolateral surface (Rbm). The paracellular pathway is also formed by two resistances in series: (1) the resistance of the TJ itself (Rj) and (2) the resistance of the intercellular space (Ri). In both tight and leaky epithelia Ram and Rbm are usually very high (Reuss and Finn, 1974). Using the whole-cell clamp method, the electrical resistance across the plasma membrane of a single MDCK cell (Rm) is around 2 GΩ (Stefani and Cereijido, 1983; Bolivar and Cereijido, 1987). If this resistance were assumed to be homogeneously distributed (Figure 5.5), a fraction Fi of cell membrane would have an electrical resistance of Electrical resistance of Fi =

Rm Fi

(5.1)

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FIGURE 5.4 Electric circuit diagram of an epithelial monolayer, showing the paracellular and transcellular pathways in parallel. The junctional and intercellular resistances in series give rise to the paracellular pathway resistance.

FIGURE 5.5 Schematic representation of a hypothetical epithelial cell. Rm is the resistance of the whole membrane (left). To make a minimum estimate of the resistance across the transcellular route (right), it is assumed that Rm is homogeneously distributed over the cellular membrane, so that fractions a and b have a resistance of Rm/Fa and Rm/Fb, respectively. (From González-Mariscal, L. et al., J. Membr. Biol., 107, 43, 1989. With permission.)

The electrical resistance of the transcellular route of a single cell is given by the resistance of the apical fraction (Fa) plus the resistance of the basolateral fraction (Fb) as follows: Transcellular resistance of a single cell =

Rm Rm + Fa Fb

(5.2)

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The addition of the two fractions constitutes the entire membrane: Fa + Fb = 1

(5.3)

Therefore, Fb can be replaced in Equation 5.2 by 1 – Fa. If the value of Rm found experimentally for MDCK cells (2GΩ) is now introduced, Equation 5.2 becomes Transcellular resistance of a single cell =

2 GΩ Fa − Fa2

(5.4)

This equation has some interesting consequences. For example, if the apical fraction constitutes 50% of the membrane (Fa = Fb = 0.5), Equation 5.4 predicts that the transcellular route across a single cell will have a resistance of 8 GΩ. If, at the most, 200,000 cells occupy a square centimeter of an MDCK monolayer and act as resistors arranged in parallel, they would therefore offer a total transcellular resistance of 40,000 Ω·cm2. In the circuit diagram of Figure 5.4 TER is given by: 1 1 1 = + TER Rtranscellular Rparacellular

(5.5)

Even using the highest value of TER measured in MDCK cells (5000 Ω·cm2 in strain I; Stevenson et al., 1988), the resistance through the transcellular route of this epithelial monolayer is seven times higher, indicating that the paracellular pathway (Rp) is much more conductive than the transcellular route. It may be noticed that, if the electrical resistance of the cell membrane was not evenly distributed and the fractions occupied by the apical and the basolateral sides were not identical (as has been assumed in the example discussed above), the resistance of the transcellular route will afford a higher percentage of the resistance, and the paracellular route will generate a much lower one. Therefore, the value of TER across monolayers of MDCK cells reflects mainly the resistance offered by the TJ.

5.5.3 CLASSIFICATION

OF

EPITHELIA

AS

“TIGHT”

OR

“LEAKY”

The TER may be very low in tissues, such as the small intestine, gallbladder, and proximal tubule of the kidney, that transport large quantities of water and solutes (e.g., 6 Ω·cm2 at the proximal tubule of the kidney; Boulpaep and Seely, 1971). Conversely, other tissues, such as the frog skin, the urinary bladder, and the stomach mucosa, are much more restrictive to the paracellular passage of water and other molecules (e.g., 12,000 Ω·cm2 at the urinary bladder; Claude, 1978). This led to the categorization of epithelia as leaky or tight (Diamond, 1974). In the former, the transcellular resistance is larger than the paracellular, whereas in the latter, the paracellular resistance is ~100 times higher than the transcellular one. This bimodal

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FIGURE 5.6 Morphological parameters that affect TER. The linear amount of junction per square centimeter (lp) varies from epithelium to epithelium and depends on: (1) (top) the length of the intercellular cleft: the longer the cell perimeter the larger lp; (2) (bottom) the size of the cells in the epithelium: the smaller the diameter of the cells, the larger the lp .

classification of epithelia as tight or leaky is a practical oversimplification, since the ratio of junctional to cellular resistance can assume any value along a continuum.

5.5.4 MORPHOLOGICAL ASPECTS THAT AFFECT TER The amount of current that traverses the paracellular pathway depends not only on the resistance of this route, but also on how much pathway is available per epithelial surface. The linear amount of paracellular route per square centimeter of epithelium (lp) varies from epithelium to epithelium and depends on two factors: (1) the size of the cell in the epithelium: the smaller the diameter of the cells, the larger lp (Figure 5.6); and (2) the tortuosity of the cell borders: cells with wavy interdigitated profiles have a much larger lp than cells of the same size with smooth borders (Figure 5.6). For example, the villus surface of guinea pig ileum is covered by polygonal absorptive cells with an estimated width of 10 µm, while the crypt epithelial cells, which are also polygonal in shape, have instead widths of only 3.5 µm. Morphological evaluations have shown mean lp values for villi and crypt of 21.8 and 76.8 m/cm2, respectively. In this case the high linear junctional density (lp) in the crypt is not due to tortuous cell borders, but rather to the smaller cell widths (Marcial et al., 1984). The above considerations may serve to explain the change in TER values of MDCK monolayers as a function of their age. Figure 5.7 shows that MDCK monolayers exhibit an initial rapid increase in TER followed by a decrease to a stable level. This variation may be associated to the number of cells in the monolayer: as the density increases, the length of the intercellular space per unit area of monolayer also increases. This increase may be responsible for the reduction in the value of TER. This possibility is supported by the observation that monolayers treated with thymidine to arrest cell growth show no decrease in TER as a function of monolayer age, while the intrinsic properties of their TJ are similar to control monolayers (Rabito, 1986).

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FIGURE 5.7 Cellular density (--) and electrical resistance across monolayers (-●-) of MDCK cells as a function of time after plating. Cellular density was counted in the same disks used for the electrical measurements. (From Cereijido, M. et al., J. Exp. Biol., 106, 205, 1983. With permission.)

As stated above, part of the resistance of the paracellular pathway is due to length, narrowness, and tortuosity of the intercellular cleft. Claude (1978) has given the following expression for intercellular resistance: Ri = ρL ωlp

(5.6)

Where ρ is the resistivity of the bulk solution (assuming that intercellular solution has the same resistivity as the bulk solution), L is the height of the interspace, ω is the width of the interspace, and lp is the linear amount of paracellular route per square centimeter of epithelium. In Necturus proximal tubule, Claude (1978) calculated an Ri value of 6 Ω·cm2, i.e., less than 10% of Rp. In most tissues Ri is even lower because the cells are smaller so L decreases while the value of lp increases. However, if the lateral cell membranes are tightly apposed (interspace width < 20 nm), as occurs under certain experimental situations such as the presence of osmotic gradients, the resistance of the intercellular space (Ri) may increase greatly and contribute substantially to the paracellular resistance (Rp). Yet, when the intercellular width is larger than 0.5 µm, the intercellular resistance Ri is very small. Estimates for a large variety of epithelia show that Ri is very small with respect to Rp, so that the resistance of the junction (Rj) may be taken as an approximate value of the paracellular one (Rp).

5.5.5 THE RELATIONSHIP BETWEEN THE SPECIFIC RESISTANCE OF THE TIGHT JUNCTION (RJlP) AND THE NUMBER OF ITS STRANDS Claude (1978) calculated Rj for a number of epithelia and adjusted it for lp, the amount of junction per unit area, by (1) using the data available in the literature for

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105

FIGURE 5.8 Specific functional resistance as a function of the number of TJ strands. (Upper) The graph shows that once the electrical resistance of diverse epithelia is corrected for length per square centimeter of epithelium, it increases with the number of strands in an exponential manner, and not in a direct fashion as would be expected for a sum of resistors in series. This is attributed to the existence of channels that fluctuate from an open to a closed state. (Bottom) A segment of TJ with two strands shows the four possible states of the channels. The segment becomes conductive only when the two channels coincide in the open state (last condition on the right). (From Cereijido, M. et al., NIPS, 4, 72, 1989. With permission.)

Rt and Rc, (2) estimating lp on the basis of cell diameters, and (3) assuming a negligible Ri. When Claude plotted Rj lp against the number of junctional strands in each epithelium, she did not obtain a linear relationship as would be expected by the sum of resistors in series, but a logarithmic one (Figure 5.8). On these bases, Claude suggested that strands may contain labile porelike structures for the movement of small ions that can be in either an open or a closed state. The local resistance of each strand is then related to the probability of the strand having an open pore in that region. In the case of TJs constituted by many strands, a conductive state is only achieved when each strand in that area presents an open pore. The bottom part of Figure 5.8 depicts the four possible configurations of a TJ with two strands, each with one flickering channel with an open probability of 0.5 (e.g., the channel spends the same amount of time in the open or closed state). Only the one on the right-hand side, where both channels are simultaneously open, is in a conductive state. Therefore, a TJ constituted by two strands would not have a TER twofold higher than a TJ with a single strand (as expected from the sum of two resistors in series), but would have a TER fourfold higher. However, as pointed out by Cereijido et al. (1989), the model does not apply to long strands that surround the entire cell

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FIGURE 5.9 Compartmentalized flickering channels. In a junctional belt, which has no anastomoses (A), current that has flown through one strand may use any open channel in the next strand at a given time. Therefore, the conductance of a two-stranded TJ is one half the conductance of a one-stranded TJ. If channels were instead compartmentalized (B), so that current flowing through one segment can only cross the next strand if it also has an open channel, conductance will be markedly restricted. Thus, because of branching and anastomosing of its ridges, a TJ may exhibit the exponential relationship between the number of strands and TER shown in Figure 5.8, and an epithelium may offer a high TER with only a few strands in its TJs. (From Cereijido, M. et al., NIPS, 4, 72, 1989. With permission.)

and therefore have a large number of such channels, but is only true for short segments of strands having a single channel. In a strand with many channels (Figure 5.9A), current would flow through any open channel and, therefore, a TJ with two strands would have a TER twofold (instead of fourfold) higher than a single-stranded one. To explain the exponential relationship between number of strands and electrical resistance on the basis of channels, it is necessary to assume that the segments of the strands in which they are contained are electrically isolated. Cereijido et al. (1989) suggested that electrical compartmentation may be afforded by the frequent anastomoses of TJ strands that are observed in freeze-fracture replicas (Figure 5.9B). High values of TER and low paracellular flux (and vice versa) have generally been considered to go together. However, in MDCK cells whose linoleic acid content has been enriched (Calderón et al., 1998) or that overexpress chick occludin (Balda et al., 1996; McCarthy et al., 1996), a functional dissociation of paracellular permeability from electrical resistance has been found. These apparent contradictory findings can be explained on two bases: (1) that TJs may contain carriers and (2) that TJs possess compartmentalized channels. It should be considered that in the mechanism known as “exchange diffusion,” carriers translocate thousands of ions per second without an electrical manifestation. During their cyclic work they undergo

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107

changes in configuration to deliver ligands to the opposite sites of the membrane, which are particularly sensitive to modifications in the viscosity of the lipid matrix. Therefore, replacing oleic acid with linoleic acid would allow the carrier to operate in a more fluid environment, thus increasing a paracellular flux without a concomitant change in electrical resistance. Moreover, it should be considered that while TER is an instantaneous measurement that reflects the permeability at a given point in time, the paracellular flux is an indicator for the permeability over a period of, usually, minutes or hours. Thus, when a flux of tracer starts to enter into the junction, it can go through, provided the first pore is open, and remains in this subcompartment until, at a later time, the channel in the next strand opens. By this time, the pore in the first barrier might already be closed again. As a consequence, the tracer could migrate one compartment at a time through TJs, which as a unit were electrically sealed during the entire process.

5.6 PRESENCE OF PORES OR CHANNELS WITHIN THE TIGHT JUNCTION STRANDS The existence of flickering pores within the strands implies that in one state certain ions may pass and in the other state they may not. Since the permeability of ions through pores or channels depends on the electric charges in or near the pore wall, slight modifications in the environment of the strand or in its structure may exert a profound effect on the permeability. TJs of leaky epithelia are selectively permeable to cations over anions, thus behaving as if they were negatively charged at physiological pH. This perception is confirmed by the observation that an acidic milieu triggers a reversible inversion of selectivity. Furthermore, changes in pH elicit distinct effects on the specific permeability of each ionic species (Wright and Diamond, 1968; Moreno and Diamond, 1974; Cereijido et al., 1978). In fact, the use of low-pH buffer in cerebral endothelia increases the penetrance of normally excluded polar compounds through the blood–brain barrier (Oldendorf et al., 1994). The proposal of TJs with tritable negative charges is further supported by the observation that treatments with polycations such as 2,4,6-triaminopyrimidine (Moreno, 1974) and Th4+ (Machen and Diamond, 1972) abolish paracellular cation selectivity, and that chitosan, a cationic polysaccharide, enhances hydrophilic drug penetration through the paracellular pathway (Junginger and Verhoef, 1998). However, the action of other cationic compounds on paracellular conductance has been more difficult to understand. For example: 1. In Necturus gallbladder, protamine reversibly increases TER and decreases the conductance of cations through the TJ (Fromm et al., 1985). 2. In the glomerular epithelia, both protamine and puromycin aminonucleosides induce a nephrosis in which the slit diaphragms are displaced by occluding-type junctions (Kerjaschki, 1978; Kurihara et al., 1992). 3. In a patient with Fanconi’s syndrome, a condition characterized by the abnormal presence of TJs between glomerular foot processes, a cationic kappa light-chain protein isolated from urine increased both TER and TJ depth when added to a Necturus gallbladder (Alavi et al., 1983).

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The mechanism by which polycations are such powerful stimulators of TJ formation remains unknown. However it is tempting to speculate that as polycations neutralize the net negative charges between adjacent membranes, charge repulsion disappears, the cells come into closer proximity, and consequently form TJs. In fact, in patients with minimal-change nephrosis, the loss of fixed negative charges in glomerular structures is coupled to a depressed clearance of dextrans (Carrie et al., 1981). An explanation for the paradoxical condition in which some polycations promote junction formation while others, like chitosan, increase paracellular permeability is difficult to find. However, the answer may lie in the particular chemical structure of each polycationic molecule and the biological system under study.

5.6.1 NEPHRIN, A POROUS FILTER SLIT DIAPHRAGM

OF THE

PODOCYTE

At the glomeruli, podocytes are normally maintained wide open, facilitating the passage of plasma from the fenestrated endothelium. Podocyte foot processes are interconnected through slit diaphragms that evolved during renal development from true TJs. In slit diaphragms ZO-1 is coexpressed with α, β, and γ-catenin (Schnabel et al., 1990; Reiser et al., 2000). Instead of occludin (Kwon et al., 1998) or claudin, slit diaphragms display the transmembrane proteins P-cadherin and nephrin. The latter regulates the permeability and selectivity properties of the slit diaphragm complex, by forming negatively charged “pores” (for review, see Tryggvason, 1999) (Figure 5.10), therefore, explaining why, when the gene for nephrin is mutated, a serious congenital nephrotic syndrome develops (Kestilä et al., 1998).

5.6.2 PARACELLIN, A MG2+/CA2+ CHANNEL

OF

TIGHT JUNCTIONS

Although all TJs so far studied in tight and leaky epithelia possess a common background of peripheral proteins (ZO-1, ZO-2, ZO-3, cingulin, etc.), distinct claudins are expressed in different epithelia, thus suggesting that variations in both TJ structure and permeability properties might reside on the particular combination of claudins found on each tissue. Employing positional cloning, Simon (Simon et al., 1999) identified a new member of the claudin family (paracellin-1/claudin 16, PCLN-1), whose mutations cause hereditary renal hypomagnesemia in humans. Differently from other cations such as Na+, K+, and Ca2+, resorption of Mg2+ is largely due to transport through the paracellular pathway driven by an electrochemical gradient across the epithelium of the thick ascending limb of Henle (TAL). PCLN-1 is exclusively found in the TJs of the TAL where it apparently constitutes a highly selective Mg2+/Ca2+ channel. PCLN-1 therefore constitutes the first TJ protein that seems to form an intercellular pore. The first extracellular domain of PCLN-1 contains ten negatively charged residues and a net charge of –5, thus complying with the selectivity of cations over anions previously detected in TJs. A distinguishable feature between PCLN-1 pores and conventional transmembrane channels rests on their orientation, as the former are oriented parallel and not perpendicular to the plane of the membrane and do not cross the membrane lipid bilayer as regular channels do (Goodenough and Wong, 1999).

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FIGURE 5.10 Hypothetical assembly of nephrin into a porous filter of the podocyte slit diaphragm. Nephrin molecules extending toward each other from two adjacent foot processes are likely to interact homophilically in the slit through their Ig repeats (1 to 6). This interaction would constitute the central filament of the slit observed in TEM. Surrounding it, slit membrane “pores” would appear, limited by the region between the fibronectin domain and the zippered Ig repeats of nephrin.

Since claudins are a heterogeneous family, each species or their combination could form channels at the TJ, with different permeability and selectivity properties, that might be responsible for the variation in paracellular conductance among epithelia.

5.7 THE ELECTRIC CIRCUIT ANALYSIS AS A TOOL FOR PREDICTING EPITHELIAL RESISTANCE The electric circuit analysis of epithelia, based on Claude’s equations (Claude, 1978), has been tested for its capacity to predict TER under different conditions.

5.7.1 IN OF

A

NATURAL EPITHELIA FORMED

BY

DIFFERENT TYPES

CELLS

Histological analysis shows that ileum and colon are formed by crypt and villi cells. In ileum, the total surface area occupied by crypt and villus are 13 and 87%, respectively. Since the specific junctional resistance (Rj lp) calculated for the crypt is an order of magnitude lower than that of the villus, electric circuit analysis predicts that 73% of the paracellular conductance is attributable to the crypt. This makes the crypt region responsible for the majority of net ileal paracellular conductance (Marcial et al., 1984).

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Tight Junctions

In the colon, while the crypt surface is 1.2 times bigger than the villus area and the resistance of the former is more than threefold higher, the paracellular resistances are not different. This paradoxical result suggests the presence of a higher density of ion channels in the apical membrane of villus cells (Gitter et al., 2000).

5.7.2 BETWEEN EPITHELIA DERIVED ANIMAL SPECIES

FROM

DIFFERENT

As mentioned above, early comparisons of TJs and TERs in different epithelia (e.g., mammalian ileum vs. toad urinary bladder), based solely on mean TJ strand counts, indicated that the number of TJ strands is apparently unrelated to TER (MartínezPalomo and Erlij, 1975). To analyze this discrepancy, Claude’s morphometric parameters were evaluated in both of these epithelia, and results were used to predict their TERs. Theoretical TER for toad urinary bladder was found to be two orders of magnitude higher than for mammalian ileum (Marcial et al., 1984). These predictions agree with experimental measurements of TER, thus showing the validity of this analysis to study the degree of permeability of a given epithelium.

5.7.3 DURING

THE

ESTABLISHMENT

OF

EPITHELIAL MONOLAYERS

Monolayers of epithelial cells develop as their TJs become assembled and sealed. In cultured cells this process can be studied from the moment of plating to the establishment of steady-state TER values. The morphometric analysis of TJ described above allowed Madara and Dharmsathaporn (1985) to predict correctly increments in TER of monolayers of T84 cells as a function of time after plating.

5.7.4 BETWEEN CELLS THAT SIGNIFICANTLY DIFFER IN THEIR TER IN SPITE OF BELONGING TO THE SAME CELL LINE Clones of the same cell line may express remarkable variations in their steady-state values of TER. Yet, in two clones of MDCK cells that express significantly different TERs, morphometric analysis indicated that they have similar values of (1) amount of junction, (2) specific resistance of the TJ, and (3) theoretical resistance (Table 5.2) (Stevenson et al., 1988; González-Mariscal et al., 1989). This discrepancy between TJ structure and TER may be explained by assuming that, at a given time, the strands in high-resistance MDCK cells contain a smaller number of channels, contain an equal number of channels but that they preferentially remain in the closed state, or express a different set of claudins. When high- and low-resistance MDCK strains are cocultured, the resultant mixed monolayers develop TERs whose values are very close to the theoretical ones, thus suggesting that low and high resistance cells work in the same monolayer like a parallel circuit (Table 5.3) (González-Mariscal et al., 1989). In these mixtures, the intercellular contacts between cells from dissimilar strains are atypical, in the sense that although they contain E-cadherin, ZO-1, and occludin, they are located at the bottom and not at the uppermost region of the lateral membrane (Figure 5.11) (Collares-Buzato et al., 1998).

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TABLE 5.2 Comparison of Theoretical and Experimental TER in Wild and High Resistance (HR) MDCK Cells Specific Junctional Resistance (Ω·cm) 2.64 × 104 2.64 × 104

Cell Type MDCK MDCK (HR) a

Linear Amount of TJ per Unit Area (cm/cm2) 765 ± 23 792 ± 41

TER n 111 135

Theor.a 35 29

Exp. (Ω·cm2) 185 ± 17 (23) 625 ± 32 (22)

Theoretical TER was calculated as follows: 1 TER

theor

=

(

linear amount of TJ per unit area cm cm specific junctional resistance (Ω ⋅ cm )

2

)

TABLE 5.3 Theoretical vs. Experimental TER in Monolayers of Mixeda Types of MDCK Cells TER (Ω·cm2) 236 ± 12 (8) 625 ± 32 (22) 306 ± 44 332 ± 18 (5)

Cell Type Low resistance High resistance Mixed (theoretical)b Mixed (experimental) a b

Mixed in a 50:50 ratio. Rtheor was calculated as:

F F 1 = 1 + 2 R R R theor 1 2 where F1 and F2 are the fractions (0.5) of each cell type in the mixture, and R1 and R2 are their experimental values of TER Source: González-Mariscal, L. et al., J. Membr. Biol., 107, 43, 1989. With permission.

5.7.5 IN MIXED MONOLAYERS FORMED FROM DIFFERENT ANIMAL SPECIES

BY

CELLS DERIVED

Cells from different animal origins cocultured in monolayers can make sealed TJs, suggesting the conserved nature of this structure. The experimental TER values obtained in these mixed monolayers closely resemble the theoretical estimations for the culture as a parallel circuit (Table 5.4). TJs cannot be established if one of the partners does not normally express them (González-Mariscal et al., 1989). Accordingly, neither ZO-1

112

Tight Junctions

FIGURE 5.11 Schematic representation of strains I and II MDCK cocultured monolayers. Cells from the same MDCK strain form apical intercellular junctions (arrow), whereas contacts between different cell types are abnormally located at the bottom of the cells (empty arrowheads). Strain II and not strain I MDCK cells display microvilli strongly stained with horseradish peroxidase–conjugated peanut agglutinin (HRP-PNA) (full arrowhead).

TABLE 5.4 Theoretical vs. Experimental TER in Monolayers of Mixed Cell Types Cells in the Mixture, % Cell Line

Ptk2 MDBK

MK2 LLC-RK1 MA-104

LLC-PK1

CPA52 VERO

a 35 b

Plated 50 75 50 25 50 50 75 50 25 75 50 25 50 75 50 25

Exp.a 47 ± 1 (6) 72 ± 3 (4) 54 ± 1 (8) 34 ± 2 (4) 45 ± 1 (8) 55 ± 1 (8) 72 ± 3 (4) 54 ± 1 (8) 34 ± 2 (4) 77 ± 3 (4) 51 ± 2 (4) 23 ± 2 (4) 51 ± 2 (6) 55 ± 3 (6)

TER (Ω·cm2) Theor.b 127 ± 8 22 ± 1 28 ± 2 47 ± 2 36 ± 5 53 ± 8 47 ± 5 58 ± 5 80 ± 6 174 ± 10 186 ± 9 202 ± 8 15 ± 3 7±3 10 ± 2 18 ± 1

Exp. 119 ± 15 (7) 35 ± 2 (8) 31 ± 2 (19) 82 ± 9 (7) 31 ± 3 (8) 84 ± 4 (8) 35 ± 2 (8) 31 ± 2 (19) 82 ± 9 (9) 184 ± 16 (21) 185 ± 16 (22) 239 ± 22 (15) 9 ± 3 (8) 20 ± 6 (9) 36 ± 5 (15) 55 ± 9 (10)

Percentage of cells experimentally found to attach, measured with S-methionine at the moment of TER determination. Resistance was calculated as explained in Table 5.3.

Source: González-Mariscal, L. et al., J. Membr. Biol., 107, 43, 1989. With permission.

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113

nor occludin has been found at the boundaries between epithelial and fibroblastic cells in coculture (Cereijido, unpublished observations), supporting the idea that to form TJs each neighbor must contribute its moiety.

5.8 THE PERMEABILITY OF THE TIGHT JUNCTION CAN BE MODIFIED EXPERIMENTALLY WITHOUT CONCOMITANT CHANGES IN THE ARRANGEMENT OF THE STRANDS To minimize the uncertainties of comparing TJ structure and function between epithelia formed by several cell types and derived from different animal species, some studies resorted to changes in the environment that alter the permeability of TJs of monolayers formed by single cell types. Thus, when the incubation temperature of monolayers of MDCK was lowered from 37 to 4oC, there occurred no detectable change in the number and distribution of TJ strands, yet the value of TER increased by 305% in a reversible way (González-Mariscal et al., 1984). These data also support the existence of ion channels in the strands, and suggest that the open and closed states possess temperature-sensitive gates. An alternative explanation is that the low temperature decreases the ionic mobility near the strands.

5.9 CHANGES IN THE PHOSPHORYLATION STATE OF TIGHT JUNCTION COMPONENTS MAY AFFECT THE PERMEABILITY OF THE EPITHELIUM Once the first TJ molecules were identified, an effort was made to relate their degree of phosphorylation to the junctional function. Starting with the submembranous TJ protein ZO-1, it was determined that a low-resistance strain contained approximately twice as much phosphorylated ZO-1 as a high-resistance strain (Stevenson et al., 1989). When occludin was later discovered, it became clear that TJ formation is accompanied by the insolubilization and serine phosphorylation of this protein (Sakakibara et al., 1997). However, during the early development of Xenopus laevis, a correlation between occludin dephosphorylation and TJ assembly was found (Cordenonsi et al., 1997). A number of studies have recently shown that TJ biogenesis is accompanied by changes in tyrosine phosphorylation. Thus, it seems that the level of ZO-1 and occludin tyrosine phosphorylation relates with the value of TER achieved (Kurihara et al., 1995; Tsukamoto and Nigam, 1999; Chen et al., 2000). This difference in the phosphorylation state of the TJ suggests that the permeability of the junction may in principle be regulated by biochemical processes that cannot be distinguishable by structural criteria.

5.10 CONCLUDING REMARKS The study of the relationship between structure and function of TJs began with the expectancy that the value of TER would be linearly related to the number of strands observed by freeze-fracture replicas, as expected from resistors arranged in series.

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Tight Junctions

The information since obtained, especially with the identification of TJ-specific molecules (ZOs, claudins, and occludin), has not supported this simplistic view, and has led to the incorporation of the following additional aspects: 1. The percentage of junctional segments with 1, 2, 3, … n strands; 2. The length of junctional cleft, given in turn by the number of cells per unit area and the interdigitation of their borders; 3. The length, width, and tortuosity of the intercellular space; 4. The specific molecular composition of the TJ strands (e.g., type of claudins involved); 5. The existence within the strands of channels (formed by claudins?); 6. The compartmentation of channels, afforded by the frequent anastomoses between junctional strands; 7. The biochemical state of junctional components (e.g., phosphorylation); and 8. The control afforded by its relationship to submembranous molecules, the cytoskeleton, G proteins, cAMP, etc. The new discoveries and the necessary modifications of conceptual frameworks afford a more complex view of the relationship between structure and function of TJs.

ACKNOWLEDGMENT The authors thank Dr. Marcelino Cereijido from CINVESTAV, Mexico, for his reading and helpful comments on the manuscript. Antonia Avila and Abigail Betanzos are recipients of doctoral fellowships from the Mexican National Council on Science and Technology (CONACYT: 90147 and 95736, respectively). This work was supported by CONACYT Grant 28083N.

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Simon, D. B., Lu, Y., Choate, K. A., Velazquez, H., Al-Sabban, E., Praga, M., Casari, G., Bettinelli, A., Colussi, G., Rodriguez-Soriano, J., McCredie, D., Milford, D., Sanjad, S., and Lifton, R. P., 1999. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption, Science, 285, 103. Sonoda, N., Furuse, M., Sasaki, H., Yonemura, S., Katahira, J., Horiguchi, Y., and Tsukita, S., 1999. Clostridium perfringens enterotoxin fragment removes specific claudins from tight junction strands: evidence for direct involvement of claudins in tight junction barrier, J. Cell Biol., 147, 195. Spiegel, S., Blumenthal, R., Fishman, P. H., and Handler, J. S., 1985. Gangliosides do not move from apical to basolateral plasma membrane in cultured epithelial cells, Biochim. Biophys. Acta, 821, 310. Staehelin, L. A., 1973. Further observations on the fine structure of freeze-cleaved tight junctions, J. Cell Sci., 13, 763. Stefani, E. and Cereijido, M., 1983. Electrical properties of cultured epithelial cells (MDCK), J. Membr. Biol., 73, 177. Stevenson, B. R. and Goodenough, D. A., 1984. Zonulae occludentes in junctional complexesenriched fraction from mouse liver: preliminary morphological and biochemical characterization, J. Cell Biol., 98, 1209. Stevenson, B. R., Anderson, J. M., Goodenough, D. A., and Mooseker, M. S., 1988. Tight junction structure and ZO-1 content are identical in two strains of Madin–Darby canine kidney cells which differ in transepithelial resistance, J. Cell Biol., 107, 2401. Stevenson, B. R., Anderson, J. M., Braun, I. D., and Mooseker, M. S., 1989. Phosphorylation of the tight junction protein ZO-1 in two strains of Madin–Darby canine kidney cells which differ in transepithelial resistance, Biochem. J., 263, 597. Tryggvason, K., 1999. Unraveling the mechanisms of glomerular ultrafiltration: nephrin a key component of the slit diaphragm, J. Am. Soc. Nephrol., 10, 2440. Tsukamoto, T. and Nigam, S. K., 1999. Role of tyrosine phosphorylation in the reassembly of occludin and other tight junction proteins, Am. J. Physiol., 276, F737. Tsukita, S. and Furuse, M., 1999. Occludin and claudins in tight junction strands: leading or supporting players? Trends Cell Biol., 9, 268. van Deurs, B. and Koehler, J. K., 1979. Tight junctions in the choroid plexus epithelium, a freeze-fracture study including complementary replicas, J. Cell Biol., 80, 662. van Deurs, B. and Luft, J. H., 1979. Effects of glutaraldehyde fixation on the structure of TJs, J. Ultrastruct. Res., 68, 160. van Meer, G. and Simons, K., 1986. The function of tight junctions in maintaining differences in lipid composition between the apical and the basolateral cell surface domains of MDCK cells, EMBO J., 5, 1455. Wade, J. B. and Karnovsky, M. J., 1974. The structure of the zonula occludens: a single fibril model based on freeze-fracture, J. Cell Biol., 60, 168. Westcarr, S., Farshori, P., Wyche, J., and Anderson, W. A., 1999. Apoptosis and differentiation in the crypt-villus unit of the rat small intestine, J. Submicrosc. Cytol. Pathol., 31, 15. Wolburg, H., Neuhaus, J., Kniesel, U., Krauss, B., Schmid, E. M., Öcalan, M., Farrell, C., and Risau, W., 1994. Modulation of tight junction structure in blood–brain barrier endothelial cells. Effects of tissue culture, second messengers and cocultured astrocytes, J. Cell Sci., 107, 1347. Wong, V. and Gumbiner, B. M., 1997. A synthetic peptide corresponding to the extracellular domain of occcludin perturbs the tight junction permeability barrier, J. Cell Biol., 136, 399. Wright, E. M. and Diamond, J. M., 1968. Effect of pH and polyvalent cations on the selective permeability of gallbladder epithelium to monovalent ions, Biochem. Biophys. Acta, 163, 57.

6

General Themes in Cell–Cell Junctions and Cell Adhesion Rachel Eelkema and Pamela Cowin

CONTENTS 6.1 6.2

Introduction ..................................................................................................121 Molecular Structure and Assembly of Adherens Junctions ........................122 6.2.1 Cadherins..........................................................................................122 6.2.2 The Role of the Adherens Junction in Initiating Cell–Cell Contact .............................................................................................123 6.2.3 Stage 1: Lateral Cadherin cis-Dimer Formation in the Plane of the Membrane..............................................................................123 6.2.4 Stage 2: Adhesive trans-Dimerization of Cadherins.......................125 6.2.5 Stage 3: The Role of the Cadherin Juxtamembrane Domain and p120 in Lateral Dimerization and Clustering...........................125 6.2.6 Stage 4: The Role of Catenins in Harnessing the Cytoskeleton.....128 6.2.7 Stage 5: Actin Remodeling and Cell Compaction ..........................129 6.2.8 Regulation of Cell Adhesion............................................................129 6.3 Desmosomes.................................................................................................131 6.3.1 Desmosomal Cadherins....................................................................131 6.3.2 Desmosomal Cytoplasmic Interactions ...........................................132 6.3.3 Plakoglobin.......................................................................................133 6.3.4 Intermediate Filament Binding Proteins..........................................133 6.3.4.1 Desmoplakins....................................................................134 6.3.4.2 Plakophilins.......................................................................134 6.4 A Role for Junctional Proteins in Signal Transduction and Gene Regulation ....................................................................................................135 6.5 Conclusion....................................................................................................136 References..............................................................................................................136

6.1 INTRODUCTION Cell–cell adhesive junctions bind cells together and connect the cytoskeleton to the plasma membrane in cell-type-specific patterns that greatly influence tissue contour and resilience. Several components of these junctions also participate in signal 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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transduction pathways that regulate gene expression. Adhesive junctions may be considered therefore as epicenters for signal reception, transduction, and response to local cellular patterning cues. There are two major categories of cell–cell adhesive junction: desmosomes, which bind intermediate filaments and are largely restricted to epithelial cells and cardiac myocytes; and the more ubiquitous actin-associated adherens junctions, which are additionally seen in endothelia and fibroblasts. Rare examples of complexus adhaerens composed of desmosomal plaque proteins and adherens junction adhesive proteins are found in certain endothelial cells. Adhesive junctions are frequently found in specific spatial relationships with other cell junctions. Classical examples of this include the terminal bar complex of polarized epithelia and the intercalated disks of heart (Farquhar and Palade, 1963). The presence of adherens junctions in each of these junctional combinations reflects their essential role in initiating cell adhesion (Gumbiner et al., 1988; Vasioukhin et al., 2000).

6.2 MOLECULAR STRUCTURE AND ASSEMBLY OF ADHERENS JUNCTIONS 6.2.1 CADHERINS The adhesive components of adherens junctions are formed by classical type I cadherins (calcium-dependent adhesive proteins), which belong to the larger cadherin superfamily comprising classical and desmosomal cadherins, protocadherins, and cadherin-related proteins (Yagi and Takeichi, 2000). Classical type I cadherins, of which E-cadherin is the prototype, consist of five tandem extracellular 110 amino acid repeats (EC1-5), a transmembrane domain, and a highly conserved cytoplasmic region. In general, classical cadherins mediate highly specific homophilic adhesion, although there are a few reports of weak heterophilic interactions between the closely related N- and R-cadherins, as well as between E-cadherin and αEβ7-integrin (Karecla et al., 1996; Nose et al., 1988; Shan et al., 2000). Cadherins are expressed in complex temporal and spatial patterns and determine cell recognition events that govern the morphogenesis of embryonic tissues and the homeostasis of adult tissues. As a result, the consequences for the organism of aberrant expression or dysfunction of cadherins are usually dire. Loss of E-cadherin expression occurs in many carcinomas and has been shown to hasten tumor progression (Berx et al., 1995; Birchmeier, 1995; Perl et al., 1998). Intriguingly, cadherin type confers not only adhesive preference on a cell, but also determines cell polarity and migratory properties. For example, cells lacking E-cadherin or gaining N-cadherin expression become highly motile and show molecular evidence of acquired invasive capabilities (Birchmeier, 1995; Hazan et al., 2000; Nieman et al., 2000). Given the critical importance of proper cadherin expression for tissue homeostasis, surprisingly little is understood about cadherin gene regulation. This has been due to difficulty in locating the promoters upstream of the exceptionally large first introns that are present in most cadherin genes. Recently, several exciting observations have been made in this area of research. It has been shown that Snail, a transcription factor that regulates epithelial–mesenchymal transitions, represses

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E-cadherin expression (Batlle et al., 2000; Cano et al., 2000). β-Catenin/plakoglobin and Wnt signal transduction pathways also transcriptionally and post-translationally regulate several cadherins (Bradley et al., 1993; Huber et al., 1996; Stewart et al., 2000; Yanagawa et al., 1997). Many cadherin genes are clustered and therefore may be subject to coordinated regulation (Yagi and Takeichi, 2000). The most extreme example of this clustering is found among the proto-cadherins where the exons encoding the ectodomain of each member are arranged in tandem followed by a single exon encoding the common cytoplasmic domain to which they are all spliced (Wu and Maniatis, 1999).

6.2.2 THE ROLE OF THE ADHERENS JUNCTION IN INITIATING CELL–CELL CONTACT In simple epithelia, the cadherin transmembrane components of adherens junctions are diffusely spread on the surface of noncontacted cells. They coalesce, however, forming puncta at points of labile cell–cell contact (Adams et al., 1996; Angres et al., 1996). Stronger and more stable contact ensues when cadherins become immobilized within the puncta by association with thin actin cables. Puncta are then swept into plaques that concentrate at the margins of the cell–cell contact region, by a process involving cortical actin remodeling. This reorganization results in compaction of the cells and formation of a circumferential actin cable (Adams et al., 1996; Angres et al., 1996). Studies in epidermal keratinocytes have shown that, in this cell type, contact is initiated by calcium-induced, actin-dependent propulsion of filopodia into apposing cells (Vasioukhin et al., 2000). Cadherins cluster at the filopodial tips forming a double row of puncta, termed the adhesion zipper. The puncta provide foci for the recruitment of Vasp/Mena, which promote actin polymerization and remodeling. These processes force the intervening membranes together (Vasioukhin et al., 2000). While transcriptional regulation of cadherins directs the dramatic changes in cell–cell interactions that occur during embryonic development, post-transcriptional regulation of junctional assembly is an equally important mechanism for providing rapid, reversible and subtle changes in cell adhesion (Gumbiner, 2000). Adherens junctions assemble in several clearly defined stages.

6.2.3 STAGE 1: LATERAL CADHERIN CIS-DIMER FORMATION IN THE PLANE OF THE MEMBRANE A considerable body of data from crystallographic, mutagenesis, and in vitro binding studies supports the view that lateral cis-dimerization of cadherins, occurring in the plane of the membrane, is a prerequisite for the subsequent trans (adhesive) cadherin interactions (Brieher et al., 1996). The mode of cis-dimer formation, however, remains controversial (Figure 6.1). Crystal structures of the EC1 and EC1/2 domains of E- and N-cadherin show that each EC domain adopts an immunoglobulin-like fold forming a beta-barrel (Shapiro et al., 1995; Overduin et al., 1995; Nagar et al., 1996; Pertz et al., 1999). The five EC domains become rigidified into a rodlike structure by the articulation

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FIGURE 6.1 Stage 1: cis-Dimerization. Stage 2: trans-Dimerization.

of calcium ions at the base of each beta-barrel. One model, based on crystal structures of a single N-cadherin EC1 domain, posits that cis-dimerization results from the reciprocal insertion of Trp-2 side chains into hydrophobic pockets on neighboring cadherin strands (Shapiro et al., 1995). This model is supported by mutational analysis of N–R-cadherin heterodimers and fractionation studies using tagged and mutated E-cadherin, in which cis- and trans-dimers can be distinguished (Chitaev and Troyanovsky, 1998; Shan et al., 2000). N-cadherin EC1/2 crystals, however, show an X-like cis-dimer that is splayed in a manner that is incompatible with Trp-2 exchange and is similar to structures observed in E-cadherin EC1 and EC1/2 crystals (Tamura et al., 1998). In the latter structures the cadherin strands interface at the calcium-binding region and Trp-2 is either disordered or found buried in the hydrophobic pocket of its own strand (Nagar et al., 1996; Pertz et al., 1999). A second model based on E-cadherin structures has suggested that high calcium induces cisdimerization and promotes burial of Trp-2 into the hydrophobic pocket of its own strand in a manner that alters the conformation of the underlying adhesive face (Koch et al., 1999). The involvement of calcium in progressive rigidification of the cadherin rod and cis-dimerization is supported by electron microscopy images of multimerized E-cadherin immobilized on beads and chemical cross-linking studies (Tomschy et al., 1996; Pertz et al., 1999). At the present time, however, there are data from mutagenesis experiments both for and against involvement of Trp-2 and calcium in cis-dimerization, and thus current models remain conflicted (Chitaev and Troyanovsky, 1998; Pertz et al., 1999; Shan et al., 2000).

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6.2.4 STAGE 2: ADHESIVE

TRANS-DIMERIZATION OF

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CADHERINS

The structural basis of the high specificity of cadherin homophilic adhesion is equally contentious. Mutagenesis, domain-swap, and antibody interference studies have identified sequences in EC1 that are responsible for adhesion and homophilic specificity (Nose et al., 1990). Although adhesive contacts are not observed in most crystal structures, antiparallel arrangements seen in N-cadherin EC1 crystals have suggested that the outer surface of the cis-dimer represents the adhesive interface (Shapiro et al., 1995). The physiological relevance of this observation is supported by the fact that this face contains amino acids, flanking the conserved HAV motif, which have been shown by mutagenesis to govern the specificity of cadherin adhesion (Blaschuk et al., 1990; Nose et al., 1990). Shapiro et al. (1995) have proposed a model in which strand dimers adhere in an anti-parallel fashion to form an adhesion zipper. The width of the cadherin cis-dimers forming the teeth of this zipper corresponds well with electron microscopy observations of the related desmosomal cadherins. The predicted length of an EC1-bonded trans-dimer, however, is too great to be accommodated within the intercellular space of adherens junctions. Moreover, recent direct-force measurements of immobilized monolayers of recombinant C-cadherin have shown that EC1 overlap is insufficient for adhesion and that the maximum adhesive strength is obtained when the opposing cadherins fully overlap (Sivasankar et al., 1999). Taken together with earlier observations that adhesiondisrupting antibodies map to epitopes in EC4 and EC5 as well as in EC1, these data suggest that EC1 interact with additional EC domains on the trans-cadherin partner to bring about adhesion (Nose et al., 1990; Ozawa et al., 1990; Sivasankar et al., 1999).

6.2.5 STAGE 3: THE ROLE OF THE CADHERIN JUXTAMEMBRANE DOMAIN AND P120 IN LATERAL DIMERIZATION AND CLUSTERING Although adhesive associations of the ectodomain occur spontaneously, many experiments have shown that the cytoplasmic region is required to support sustained adhesion. The prevailing view is that the cytoplasmic domain governs two experimentally separable steps: clustering of cadherins into cooperative zipperlike arrangements and the tethering of cadherins to the cytoskeleton. The cytoplasmic region of cadherins is highly conserved and can be divided into the juxtamembrane domain that regulates clustering (Figure 6.2) and the C-terminal domain that is required to harness the actin cytoskeleton (Figure 6.3). The juxtamembrane domain has been shown to play both positive and negative roles in cadherin-mediated adhesion. For example, a C-cadherin mutant containing the juxtamembrane domain but lacking the C-terminal domain can sustain strong cell adhesion to immobilized extracellular fragments of cadherin and shows clustering capability despite a lack of cytoskeletal association (Yap et al., 1998). In contrast, other reports have shown that E- or N-cadherins lacking the juxtamembrane region are nevertheless functional, and indeed improve the adhesion of several tumor cells (Ozawa and Kemler, 1998b; Aono et al., 1999). The juxtamembrane domain binds directly to p120, Figure 6.2, a distant relative (22% identity) of plakoglobin and

FIGURE 6.2 Stage 3: Clustering.

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FIGURE 6.3 Stage 4: Cytoskeletal assocation.

β-catenin, that comprises ten central Arm repeats and alternatively spliced N- and C-terminal domains (Reynolds et al., 1992; Anastasiadis and Reynolds, 2000). p120 is a close relative (45% identity) of a further subgroup of Arm proteins, including ARVCF, delta-catenin, p0071, and plakophilins, each of which binds cadherins and localizes to the nucleus (Reynolds et al., 1994; Anastasiadis and Reynolds, 2000). In an attempt to reconcile the opposing findings on the function of the juxtamembrane domain, one recent model has proposed that p120 operates as a switch that promotes clustering of cadherins when “activated,” but inhibits their coalescence when in the “inactivated” state (Anastasiadis and Reynolds, 2000). Although the nature of the activating/inactivating cues remains to be determined, phosphorylation or conformational change in p120 and regulation of Rho A activity, which is required early in cell contact for junction formation, are likely candidates (Braga et al., 1997). p120 is ideally positioned to receive tyrosine phosphorylation signals from inside and outside the cell. It associates with the tyrosine kinase Fer, binds to protein tyrosine phosphatase (PTP) µ, and is a potent substrate for v-src (Reynolds et al., 1989; Arregui et al., 2000; Zondag et al., 2000). Indirect support for the hypothesis that

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serine phosphorylation causes p120 to adopt an adhesion “inactivating” conformation comes from studies on tumor cells that show improved cell adhesion when expressing cadherins lacking the juxtamembrane domain. These cells also respond positively to treatment with the serine kinase inhibitor staurosporine as well as to light trypsinization that leaves cadherins intact but removes a putative signaling receptor (Aono et al., 1999). Significantly, each adhesion-enhancing treatment reduced the mobility of p120 in a manner consistent with limited dephosphorylation. Deletion analysis has suggested that the N terminus of p120 contains a site that is required for it to assume an adhesion-inactivating function (Aono et al., 1999). It has been suggested that p120 regulates lateral cis-dimerization as well as the specificity of cadherin localization within endothelial cell junctions (Navarro et al., 1998; Ozawa and Kemler, 1998b). However, studies investigating the ability of Chinese hamster ovary (CHO) cells expressing mutant cadherins to adhere to immobilized cadherin, have shown that the juxtamembrane domain is required for adhesion-dependent clustering, thus implicating p120 in this process (Yap et al., 1998). Cells expressing minimally altered mutants of E-cadherin that are uncoupled from p120 adhere weakly, but fail to reorganize the circumferential cortical actin filaments, an event that was also interpreted to reflect a requirement for p120 to promote the preceding clustering step (Thoreson et al., 2000). The mechanism by which p120 promotes clustering is obscure, as it does not self-associate in in vitro binding assays (Anastasiadis and Reynolds, 2000). However, recent studies have suggested that cytosolic p120 inhibits Rho A activity by preventing GDP dissociation. Rho A inhibition is relieved when cadherins associate with p120 and it has been suggested that local activation of this released Rho A by neighboring Rho exchange factors stimulates cadherin clustering (Anastasiadis et al., 2000).

6.2.6 STAGE 4: THE ROLE THE CYTOSKELETON

OF

CATENINS

IN

HARNESSING

The C-terminal domain is the most highly conserved and best-studied region of cadherins. Early experiments demonstrated that this domain forms a stable complex with three major proteins, α-catenin, β-catenin, and plakoglobin, which form a bridge to the actin cytoskeleton (Ozawa et al., 1989) (Figure 6.3). Plakoglobin and β-catenin share 65% amino acid identity and a similar organization, comprising 12 central 42 amino acid Arm repeats flanked by N- and C-terminal domains (Franke et al., 1989; McCrea et al., 1991). Structural studies on β-catenin have shown that the Arm repeats fold and stack together to form an elongated, righthanded superhelical core (Huber et al., 1997). A charged convex groove spirals around the core forming a putative binding surface for interacting partners (Huber et al., 1997). Plakoglobin and β-catenin bind directly to the cadherin C-terminal domain, through a central block of arm repeats, immediately after cadherin synthesis (Ozawa and Kemler, 1992). It has recently been suggested that this association is required for cadherins to exit from the endoplasmic reticulum and be transported through the secretory pathway (Chen et al., 1999). Plakoglobin has also been demonstrated to have topogenic capability, targeting fused proteins to cell junctions (Chitaev et al., 1996). The major function of β-catenin and plakoglobin within adherens junctions is

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to form a modulatable link, via a site in their head and first repeat to α-catenin (Aberle et al., 1996; Witcher et al., 1996). They also participate outside of the junctional complex in Wnt signal transduction, a topic that will be dealt with later. α-Catenin joins the cadherin–catenin complex late, coincident with the proteolytic processing of these cell adhesion proteins to their mature form and their appearance on the cell surface (Ozawa and Kemler, 1992). Although capable of homodimerization in the cytoplasm, α-catenin uses its N-terminal self-association site to bind to β-catenin or plakogobin and thus joins the cadherin–catenin complex as a monomer (Pokutta and Weis, 2000). Its major function is to stabilize cell adhesion through its ability to connect to the actin cytoskeleton both directly, via its C-terminal actin-bundling domain, and indirectly, via association of its central domain with the actin bundling protein α-actinin (Knudsen et al., 1995; WatabeUchida et al., 1998). Recent studies, however, suggest that α-catenin may participate in the regulation of cell proliferation vis suppression of the ras-MAPK pathway (Vasioukhin et al., 2001).

6.2.7 STAGE 5: ACTIN REMODELING

AND

CELL COMPACTION

While cadherin–catenin complexes are bound to α-actinin and the actin cytoskeleton along the entire length of apposing cell membranes of polarized cells, association of α-catenin with its cousin vinculin occurs only at adherens junctions (Figure 6.4). The 90-kD head domain of vinculin has the potential to homodimerize and also binds to the central domain of α-catenin and the C-terminal spectrin repeat of αactinin (Knudsen et al., 1995; Watabe-Uchida et al., 1998; Pokutta and Weis, 2000). The tail domain of vinculin has actin-bundling activity when released from autoinhibitory association with the head domain. α-Actinin and vinculin additionally bind zyxin, Vasp, and Mena, which stimulate the actin polymerization and remodeling that provide the force necessary to achieve cell compaction (Vasioukhin et al., 2000). Thus, vinculin may further consolidate lateral clustering within the junctional plaque and provides additional actin tethering. Intriguingly, the C-terminal domain of vinculin also binds to the tight junction protein ZO-1 and is required for organization of the honeycomb pattern of the tight junction strands (Watabe-Uchida et al., 1998). Vinculin, therefore, contributes to the organizing of the entire junctional complex of the terminal bar.

6.2.8 REGULATION

OF

CELL ADHESION

Dynamic rearrangement of the cadherin–catenin complex is an essential mechanism by which cells modulate their adhesion. The first three stages of cell adhesion are relatively labile, but subsequent cytoskeletal harnessing stabilizes the adherens junction. As all three mechanisms to harness the cytoskeleton are directly, or indirectly, dependent upon α-catenin, most factors that modulate cell adhesion act by regulating α-catenin association with the cadherin complex. For example, in the absence of cell contact, α-catenin is prevented from associating with the cadherin complex by the occupation of its binding site on the N-terminal domain of β-catenin by IQGAP (Kuroda et al., 1998; Fukata et al., 1999b). When activated by Tiam 1, a nucleotide exchange factor that localizes to the adherens junction, Cdc42 and Rac 1 sequester

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FIGURE 6.4 Stage 5: Cytoskeletal harnessing and actin remodeling.

IQGAP at the membrane, thereby permitting α-catenin to join the cadherin complex, see Figure 6.3 (Hordijk et al., 1997; Fukata et al., 1999a). Thus G proteins, their regulatory factors, and targets constitute a switch that regulates cell adhesion (Braga et al., 1997; Fukata et al., 1999b). Phosphorylation also plays a role in regulating linkage of the cadherin–catenin complex to the cytoskeleton. Several studies have shown that poor cell adhesion correlates with phosphorylation of catenins by src kinases and can be abrogated by treatment with kinase inhibitors (Matsuyoshi et al., 1992). Similarly, ErbB2 and EGF-receptor activity has been shown to result in loss of cytoskeletal association of the cadherin–catenin complex and poor adhesion (Hoschuetzky et al., 1994; Shibamoto et al., 1994). ErbB2 and EGF-receptor bind the Arm repeats of β-catenin and/or plakoglobin and phosphorylate their termini in a ligand-dependent manner (Hoschuetzky et al., 1994; Kanai et al., 1995). Treatment of cells with tyrosine phosphatase inhibitors, such as orthovanadate, leads to loss of α-catenin from the cadherin complex, again supporting the view that phosphorylation regulates β-catenin–α-catenin association (Ozawa and Kemler, 1998a). A number of phosphatase

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receptors are associated with elements of the cadherin–catenin complex and are appropriately positioned to counter the effects of kinases and stabilize cell adhesion (Brady-Kalnay et al., 1995; Balsamo et al., 1996; Sap, 1997). In addition to kinases, phosphatases, and G proteins, several other proteins are found at the adherens junction that regulate cadherin adhesion in as yet poorly understood ways. For example, Ep-CAM, an adherens junction protein comprising two extracellular EGF repeats, downregulates cadherin function and expression by causing a dissociation of the cadherin–catenin complex from the cytoskeleton (Litvinov et al., 1997). Moreover, a parallel calcium-independent adhesion system composed of nectin, an Ig-CAM of the polio-receptor related family that binds via its short cytoplasmic domain to afadin, a PDZ-containing F-actin binding protein, appears to be superimposed in these junctions (Takahashi et al., 1999).

6.3 DESMOSOMES Adherens junctions are responsible for initiating cell adhesion; however, the requirement for desmosomes to further fortify cell adhesion in many tissues is without question. Compelling evidence for this role is provided by several examples of devastating diseases, characterized by massive tissue disruption, that are linked to mutation, loss, or immunological attack on desmosomal components. The ability of desmosomes to provide strong tissue cohesion also explains their high frequency in heavily stressed stratified epithelia, such as epidermis as well as heart.

6.3.1 DESMOSOMAL CADHERINS Desmosome adhesion is also a calcium-dependent process mediated by two unique families of cadherins, desmogleins and desmocollins (Goodwin et al., 1990; Collins et al., 1991; Koch et al., 1991; Mechanic et al., 1991; Buxton et al., 1993). The intron/exon borders of desmoglein and classic cadherin genes are absolutely conserved, emphasizing their common evolutionary relationship (Puttagunta et al., 1994). Desmogleins and desmocollins are each encoded by three genes, which are clustered on chromosomes 18q12 and 24q21 and differentially expressed (Buxton et al., 1993; Solinas-Toldo et al., 1995). Currently little is known about the transcriptional regulation of desmosomal genes. Intriguingly, Dsg mRNA levels respond to E-cadherin expression and desmosomes are disassembled in response to expression of the Slug transcription factor (Jou et al., 1995; Savagner et al., 1997). Desmogleins and desmocollins show sequence similarity to classic cadherins and a similar organization of their ectodomain (Goodwin et al., 1990; Collins et al., 1991; Koch et al., 1991; Mechanic et al., 1991). The precursor segment and the fifth EC repeat are, however, truncated in desmoglein and the latter has been subject to homologous recombination events that have produced variation in its sequence (Koch et al., 1991; Puttagunta et al., 1994). Dsg1 and 3 are the target antigens of two devastating autoimmune blistering diseases, pemphigus foliaceus, also known as fogo selvagem, and pemphigus vulgaris, respectively (Stanley, 1989). Pemphigus antibodies bind to sequences in the EC1 domain of their respective desmogleins and are presumed to block adhesion

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sterically (Amagai et al., 1994). Pemphigus foliaceus causes dry crusty blisters in the upper layers of the epidermis. Pemphigus vulgaris is more frequent, and causes weeping blisters, most frequently in the oral mucosa (Stanley, 1989). Dsg3-null mice show similar oral blistering and cyclical balding due to loss of desmosomes at the base of the hair shaft that serve to stabilize the hair (Koch et al., 1997; 1998). Antibodies to desmocollins are also found in some pemphigus sera and monovalent guinea pig antibodies raised against these proteins have been shown to inhibit desmosome formation when added to cultured simple epithelial cells (Dmochowski et al., 1993; Cowin et al., 1984). Although the adhesive function of desmogleins and desmocollins is clear from these observations, reconstitution of the desmosomal adhesive mechanism in nonadhesive cells has not been convincingly demonstrated (Kowalczyk et al., 1999a). Studies of tagged desmosomal cadherins introduced into desmosome-forming cells however, suggests that desmoglein and desmocollins form heterophilic trans-dimers (Chitaev and Troyanovsky, 1997).

6.3.2 DESMOSOMAL CYTOPLASMIC INTERACTIONS Desmosomal cadherins show marked differences from classic cadherins and from each other in their cytoplasmic regions (Koch et al., 1990; Collins et al., 1991; Mechanic et al., 1991) (Figure 6.5). The desmocollin cytoplasmic domain contains a juxtamembrane domain and an alternatively spliced C-terminal region producing a large and small isoform designated “a” and “b,” respectively. The desmoglein

FIGURE 6.5 Desmosomal protein interaction.

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cytoplasmic region is very long and contains a juxtamembrane domain, a prolinerich region, a central domain, a variable number of globular 29 amino acid repeats, and an elongated glycine-rich C terminal region of unknown function. Significantly, the larger C-terminus of the Dsc a and the central domain of desmoglein contain sequence with homology to the C-terminal catenin-binding domain of classic cadherins. It is these domains that interact with plakoglobin, the only component found in both adherens junctions and desmosomes (Cowin et al., 1986; Mathur et al., 1994; Witcher et al., 1996).

6.3.3 PLAKOGLOBIN Plakoglobin binds avidly and in large amounts through a discrete site on its N-terminal repeats to desmoglein (Witcher et al., 1996) (Figure 6.5). It binds with much lower stoichiometry through a central block of repeats to desmocollin (Witcher et al., 1996). In addition, plakoglobin interacts with the N-terminal domain of desmoplakin and is proposed to form a bridge between desmoglein and this cytoskeletal linker protein, thereby functioning in a manner analogous to its role in the adherens junction (Kowalczyk et al., 1997b). A role for plakoglobin in lateral associations within the desmosome has been suggested on the basis that cells expressing C-terminally deleted plakoglobin produce enlarged junctions (Palka and Green, 1997). Plakoglobin is known to form dimeric complexes in the cytosol, and self-association of the repeat region has been described and could provide a mechanism for clustering adherin complexes (Kapprell et al., 1987; Troyanovsky et al., 1996). The N- and Cterminal domains have been implicated in specifying the interactions of the central repeats in a manner that could regulate junctional assembly (Witcher and Cowin, unpublished data). Last, plakoglobin has been shown to have topogenic potential and may be involved in targeting delivery, desmosomal cadherins to junctional sites (Chitaev et al., 1996). Despite this multiplicity of proposed roles for plakoglobin within the desmosome, desmosome-like junctions have been observed in plakoglobin-null mice (Ruiz et al., 1996). They are, however, reduced in number and mixed with adjacent adherens junction components (Bierkamp et al., 1996). However, desmosome function is severely compromised in plakoglobin-null mice, which die because the heart bursts at E12, the time at which this organ first begins to beat (Ruiz et al., 1996). The few embryos that survive beyond this stage show extensive sloughing of the epidermis (Bierkamp et al., 1996). Recently a human syndrome, Naxos disease, has been linked to mutations in the plakoglobin gene that truncate the plakoglobin C terminus and result in cardiac arrhythmias and woolly hair (McKoy et al., 2000).

6.3.4 INTERMEDIATE FILAMENT BINDING PROTEINS Desmosomes associate with intermediate filaments creating a supracellular tensile web that is vital for the resilience of many tissues and stratified epithelia in particular. Intermediate filament attachment is achieved through two families of desmosomal proteins: desmoplakins and plakophilins.

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6.3.4.1 Desmoplakins Desmoplakins are large proteins, comprising globular N- and C-terminal domains flanking a central rod domain, that dimerize to form a parallel coiled coil (Green et al., 1992). The C-terminal domain binds to intermediate filaments (IF) and has sequence similarity to the IF 1B rod domain. A similar domain is found in all members of the plakin family (Stappenbeck and Green, 1992). This family includes the hemidesmosomal IF-binding counterparts, bullous pemphigoid antigen 1 (BPAG1) and plectin, as well as envoplakin and periplakin, which are found in the keratinocyte cornified envelope (Klymkowsky, 1999). A number of other members of this family, such as ACF7, Kakapo, and BPAG1n, have been described recently, each of which has the fascinating property of cross-linking multiple elements of the cytoskeleton into interdependent systems (Karakesisoglou et al., 2000). The N terminus of desmoplakin binds to the juxtamembrane region of desmocollin-a and to desmoglein via its association with plakoglobin (Kowalczyk et al., 1997a; Karakesisoglou et al., 2000). It also connects to plakophilin (Kowalczyk et al., 1999b). Thus, desmoplakins provide critical direct and indirect linkage between the desmosomal cadherins and the IF cytoskeleton, as well as lateral associations within the plaque. Genetic data are consistent with this function. Mutations in the desmoplakin gene, found on chromosome 6p21, which result in either null allele and consequent haploinsufficiency, or a C terminally deleted protein, produce a striate subtype of palmoplantar keratoderma (Norgett et al., 2000; Armstrong et al., 1999). The phenotype of this disease involves disruption of desmosomes with consequent loosening of keratinocyte contact. Fissure of the skin elicits a compensatory linear thickening of the skin on the palms and soles. Mutations resulting in C-terminal truncated desmoplakin also lead to heart failure and woolly hair (Norgett et al., 2000).Desmoplakin-null mice exhibit very early embryonic lethality, dying far earlier than keratin-null or plakoglobin-null mice (Gallicano et al., 1998). Again, desmoplakinnull mice show severe impairment of desmosome assembly and stability, suggesting that desmoplakin may have additional regulatory functions besides serving as a cytoskeletal linker (Gallicano et al., 1998). 6.3.4.2 Plakophilins Plakophilins are Arm proteins and close relatives of p120. They are encoded by three genes and are produced as multiple alternatively spliced isoforms that localize to desmosomes and the nucleus (Hatzfeld and Nachtsheim, 1996; Paffenholz and Franke, 1997; Schmidt et al., 1997; 1999; Ide et al., 1999; Mertens et al., 1999). Plakophilin 1 binds to desmosomal cadherins, desmoplakins, and IFs via its unique N-terminal domain (Kapprell et al., 1988; Mathur et al., 1994; Hatzfeld et al., 2000). In addition to providing a second major linkage to IFs, it is thought to play a role in conjunction with plakoglobin in desmosome assembly and clustering (Kowalczyk et al., 1999b). There is strong genetic evidence for the importance of this protein. Null mutations in plakophilin 1 underlie several skin fragility and ectodermal dysplastic diseases involving skin erosions, dystrophic nails, sparse hair, and thickened and cracked palms and soles (McGrath et al., 1997). Genetic defects in plakophilin cause

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more dramatic and extensive diseases than those described for desmoplakin. However, plakophilin mutant heterozygotes show no phenotype, and therefore desmoplakin appears to be more critical than plakophilin for desmosome function.

6.4 A ROLE FOR JUNCTIONAL PROTEINS IN SIGNAL TRANSDUCTION AND GENE REGULATION As is the case for tight junctions, both adherens junctions and desmosomes contain a number of multifunctional proteins that participate in cytoplasmic signal transduction pathways and the transcriptional regulation of patterning and proliferation. For some, such as plakophilins, these additional roles are poorly understood at present but are inferred from their prominent cytoplasmic and nuclear localization. Cytoplasmic p120 binds to the VAV2 Rho exchange factor and causes an increase in cdc42 and Rac1 and a decrease in RhoA activity, thereby promoting cell migration (Anastasiadis et al., 2000; Noren et al., 2000). Nuclear p120 binds to Kaiso, a member of the poxvirus and zinc finger family of transcription factors, which recruit histone deacetylase complexes and repress transcription (Daniel and Reynolds, 1999). The best-described dual-function protein is β-catenin. Several Wnt signaling cascades operate by elevating cytosolic β-catenin levels or otherwise activating this protein and promoting its nuclear entry (Nusse and Varmus, 1992; Gumbiner, 1995). Once in the nucleus, β-catenin forms a bipartite transcription factor with Lef/Tcf proteins (Behrens et al., 1996; Molenaar et al., 1996) and modulates the expression of an array of genes that encode proteins involved in transcription, cell cycle regulation, apoptosis, and matrix remodeling.* Wnts are proposed to achieve this effect by binding to specific members of the Frizzled transmembrane receptor family, resulting in recruitment of cytosolic disheveled (Dvl) proteins to the plasma membrane and inactivation of glycogen synthase kinase (GSK-3β) (Bhanot et al., 1996; Wang et al., 1996; He et al., 1997). GSK-3β normally acts as part of a protein complex that promotes a series of post-translational modifications that target cytoplasmic β-catenin for proteosomal degradation (Rubinfeld et al., 1996; Yost et al., 1996; Aberle et al., 1997; Orford et al., 1997; Zeng et al., 1997). Thus, Wnt-inactivation of GSK-3β causes β-catenin to accumulate by preventing its degradation. Perturbation of β-catenin protein levels produces dramatic effects in both embryonic and adult tissues and has been found in many types of tumor (Funayama et al., 1995; Haegel et al., 1995; Gat et al., 1998; Harada et al., 1999; Polakis, 1999; Imbert et al., 2001). Experiments in rodent mammary and neuropheochromocytoma cells have shown that Wnt-1 expression upregulates both plakoglobin and β-catenin (Bradley et al., 1993; Hinck et al., 1994). The late embryonic-lethal phenotype of plakoglobin-null mice and the inability of endogenous plakoglobin to rescue the early embryoniclethal phenotype of β-catenin-null mice suggest that plakoglobin does not play a * For current list of target genes, references, and pathway model, see: http:www.stanford.edu/~rnusse/pathways/targets.html.

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significant role in Wnt pathways governing early development (Haegel et al., 1995; Bierkamp et al., 1996). However, recent data have shown that plakoglobin can mimic the effects of Wnt-3 and Dvl-2 when overexpressed in epidermis (Millar et al., 1999; Charpentier et al., 2000). Significantly, plakoglobin, Wnt-3, and Dvl-2 all suppress hair growth whereas overexpression of β-catenin results in additional hair follicles and formation of follicle tumors (Gat et al., 1998; Millar et al., 1999; Chan et al., 1999; Charpentier et al., 2000). Therefore, although both plakoglobin and β-catenin are enhanced by Wnts, they exert opposite effects and the general theme emerging is that plakoglobin and β-catenin function as tumor suppressor and oncogene, respectively (Aberle et al., 1995; Simcha et al., 1996; Charpentier et al., 2000). Whether plakoglobin antagonizes β-catenin transcription by sequestering transcriptional partners, promoting β-catenin degradation, silencing β-catenin’s target genes or more simply activates its own set of target genes remains a matter of debate (Zhurinsky et al., 2000).

6.5 CONCLUSION In the last decade cDNAs and genes encoding each of the major components of desmosomes and adherens junctions have been cloned, and the interactions of their protein products have been determined. Strong genetic evidence for the importance of these junctional proteins in cell adhesion has been gained from analyses of null mice and a variety of human skin and heart diseases. The focus of the field is now on understanding the precise roles of individual junctional components within the junctional assembly process as well as their fascinating additional signal transduction and transcriptional capacities that serve to coordinate cell adhesion, motility, proliferation, and shape.

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Protein Targeting Pathways and Sorting Signals in Epithelial Cells Enrique Rodriguez-Boulan, Geri Kreitzer, David Cohen, Vera Bonilha, and Anne Müsch

CONTENTS 7.1 7.2

Generation and Maintenance of Polarity in Epithelial Cells ......................145 Intracellular Sorting and Polarized Delivery of Proteins to the Cell Surface..........................................................................................................146 7.3 Apical and Basolateral Sorting Signals Directing Polarized Protein Trafficking ....................................................................................................147 7.4 Apical Sorting Signals Interact with Specialized Lipid Microdomains (Rafts) or Sorting Receptors ........................................................................150 7.5 Basolateral Sorting Signals Interact with Specialized Cytoplasmic Adaptors .......................................................................................................151 7.6 Role of Microtubule Motors in Exit from the TGN and Transport to the Plasma Membrane .............................................................................152 7.7 Role of the Actin Cytoskeleton in Golgi Exit and Arrival at the Cell Surface..........................................................................................................153 7.8 Control of Polarity by Small GTPases ........................................................155 7.9 The Tight Junction/Zonula Adherens: A Hot Area for Surface Targeting? .....................................................................................................156 7.10 Passive Mechanisms That Account for the Polarized Distribution of Membrane and Cytoskeletal Proteins ..........................................................157 Acknowledgments..................................................................................................158 References..............................................................................................................158

7.1 GENERATION AND MAINTENANCE OF POLARITY IN EPITHELIAL CELLS A large body of knowledge has been accumulated in the last two decades on the nature of the mechanisms responsible for the polarization of epithelial cells (RodriguezBoulan and Nelson, 1989; Rodriguez-Boulan and Powell, 1992). Three fundamental 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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types of mechanisms have emerged that contribute to epithelial polarity (Figure 7.1): (1) intracellular sorting and polarized delivery of proteins and lipids to the cell surface (Figure 7.1, center); (2) “trapping” interactions with domain-restricted cytoskeleton elements (Figure 7.1, right); and (3) diffusive restriction by the tight junctions (Figure 7.1, left). This chapter deals primarily with the first type.

7.2 INTRACELLULAR SORTING AND POLARIZED DELIVERY OF PROTEINS TO THE CELL SURFACE Proteins destined to reside on apical and basolateral membrane domains share a common site of origin: the rough endoplasmic reticulum (ER) and travel together through the Golgi complex (Rindler et al., 1984; Fuller et al., 1985). Segregation of these proteins occurs by packaging into distinct populations of transport intermediates that arise from the trans-Golgi network (TGN), in the trans region of the Golgi apparatus (Wandinger-Ness et al., 1990). In addition to the TGN, a group of basolateral endosomes functionally associated with the basolateral membrane and apical membrane also act as a major site of vesicular sorting in the recycling and biosynthetic pathways (see Figure 7.1). Different epithelial cell types vary extensively in the degree they rely on these different sorting organelles, resulting in very different sorting phenotypes. MDCK cells sort nearly all apical and basolateral proteins in the TGN and deliver these molecules vectorially to the appropriate plasma membrane domain (direct pathway) (Simons and Wandinger-Ness, 1990; Rodriguez-Boulan and Powell, 1992). In striking contrast, hepatocytes target all surface proteins from the TGN to basolateral endosomes, from where apical proteins are transcytosed to the apical (biliary) pole and basolateral proteins are cycled basolaterally (Hubbard et al., 1989). Only one protein, the polymeric immunoglobulin receptor (polyIgR), follows a transcytotic pathway to the apical surface in MDCK cells (Mostov, 1994). Transcytosis, basolateral recycling, and apical recycling of membrane proteins involve separate early endosomes associated with the apical and basolateral surfaces and a “common endosome” located deeper in the cytoplasm (Mostov et al., 2000). Apical proteins and transcytosing Poly-IgR may also utilize a novel endosomal compartment, called the apical recycling endosome (see Figure 7.1) (Apodaca et al., 1994). The substantial “flexibility” of the targeting pathways in different epithelial cell types clearly depends on the cell type, and in certain cases on the extent of maturation of the epithelium (Keller and Simons, 1997). The same protein, dipeptidylpeptidase IV (DPPIV), can arrive at the apical surface via transcytotic, direct, or mixed routes, according to whether it is expressed in hepatocytes, MDCK, or intestinal (Caco-2) epithelial cells (Le Bivic et al., 1990; Matter et al., 1990a; Casanova et al., 1991b). Strikingly, in a thyroid epithelial cell line (FRT), DPPIV is targeted via the indirect route during the first few days after the monolayer is established but shifts to a direct pathway in mature monolayers (Zurzolo et al., 1992). Retinal pigment epithelium (RPE) cells appear to have an intermediate sorting phenotype between MDCK cells and hepatocytes. Influenza HA utilizes a transcytotic route whereas p75NTR follows

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a direct apical route in RPE (Bonilha, 1997); these two proteins are vectorially targeted to the apical surface of MDCK cells (Misek et al., 1984; Le Bivic et al., 1991). It was initially believed that a main difference between polarized and nonpolarized cells was the existence of two post-Golgi routes to the cell surface. However, different lines of evidence now indicate that two routes biochemically similar to the “apical” and “basolateral” routes of epithelial cells exist in fibroblasts and other nonpolar cells (Musch et al., 1996; Yoshimori et al., 1996). Indeed, at least two postGolgi routes to the cell surface are also found in yeast. As a matter of fact, it is likely that several apical and several basolateral routes exist, each one guided by a specific targeting signal. An important question is, therefore, how these different routes are focused on apical or basolateral domains as cells form a confluent, polarized monolayer. This process depends on the establishment of specific cell–substrate and cell–cell interactions mediated, respectively, by integrins and E-cadherin, but the details of these mechanisms are largely unknown (Yeaman et al., 1998).

7.3 APICAL AND BASOLATERAL SORTING SIGNALS DIRECTING POLARIZED PROTEIN TRAFFICKING Several sorting signals that control polarized protein trafficking have been identified (Table 7.1). The following general principles emerge: Apical sorting signals are in the great majority of cases studied located on the luminal or transmembrane domains of apical membrane proteins. Luminal apical signals are N-glycans (Scheiffele et al., 1995), O-Glycanated regions (Yeaman et al., 1997) and unidentified proteinaceous motifs (Rodriguez-Boulan and Gonzalez, 1999). Apical sorting signals in the membrane-bound domain of apical proteins include glycosylphosphatidylinositol (GPI) (Lisanti et al., 1988; 1990; Ali and Evans, 1990). Both native and recombinant GPI proteins, including growth hormone, a secretory protein lacking N-glycans, are sorted apically in MDCK cells, suggesting that the GPI anchor is sufficient to direct apical sorting (Brown et al., 1989; Lisanti et al., 1989; Powell et al., 1991). Apical targeting information has been identified in the transmembrane domains of influenza HA and neuraminidase (Kundu et al., 1996; Scheiffele, 1997) as well as in the fourth transmembrane domain of H,K-ATPase (Dunbar et al., 2000). Interestingly, an apical sorting signal has been recently identified in the cytoplasmic domain of rhodopsin; yeast two-hybrid screening identified a crucial interaction of this signal with a dynein light chain (Tai et al., 1999). Basolateral sorting signals are localized on the cytoplasmic domain of the protein. They fall into three different groups: tyrosine-based motifs of the type YxxΦ, dileucine motifs, and pleomorphic motifs distinct from the previous two motifs. Examples of proteins utilizing the YxxΦ motif are the LDL receptor, VSV G protein, and mutant forms of p75 neurotrophin receptors (p75) and influenza HA, where a tyrosine residue was incorporated to the cytoplasmic domain (Brewer and Roth, 1991; Casanova et al., 1991a; Le Bivic et al., 1991; Matter et al., 1992). A basolateral protein with the LL motif is the Fc receptor (FcRII-B2) (Hunziker and Fumey, 1994).

FIGURE 7.1 Three mechanisms responsible for cell polarity. (Left) Tight junctions act as a passive barrier (fence) to the lateral diffusion of transmembrane proteins and lipids present in the inner leaflet of the bilayer. Lipids in the outer leaflet diffuse freely across the tight junction. (Center) Post-Golgi endocytic and exocytic pathways in MDCK cells. After synthesis in the ER and passage through the Golgi cisternae (G), proteins and lipids exit the TGN via vesicles and tubules (see Figure 7.2) helped by actin and microtubule-associated motorproteins. The direct route to the apical surface (1) utilizes syntaxin 3 and possibly SNAP 23. The direct route to the basolateral surface (2) is docked via the exocyst and syntaxin 4; rab 8 is

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TABLE 7.1 Sorting Signals in Epithelial Cells Type of Signal Apical Sorting Signals GPI anchor N-Glycans O-Glycan stem region Transmembrane domain

Proteinaceous signals

Example Decay Accelerating Factor Placental Alkaline Phosphatase Gp80 Erythropoietin P75 Neurotrophin Receptor Influenza virus hemagglutinin Influenza neuraminidase H,K-ATPase (α subunit) Rhodopsin (C-terminal domain) Hepatitis virus antigen

Basolateral Sorting Signals Tyrosine motifs Vesicular stomatitis virus G protein, influenza virus hemagglutinin (Tyr mutant), LDL receptor Dileucine motifs FcRII-B2 Other basolateral signals Neural cell adhesion molecule, 140-kD and 180-kD forms RET-PE2 antigen (EMMPRIN) Polymeric Ig Receptor

Location Membrane Luminal Luminal Membrane

Cytoplasmic Luminal

Cytoplasmic Cytoplasmic Cytoplasmic

FIGURE 7.1 (continued) also involved somewhere in this pathway. Some proteins may be initially targeted to a common endosome (CE) (8) before delivery to the basolateral (5) or to the apical surface via the apical recycling endosome (ARE). The polymeric Ig receptor, after reaching the basolateral surface via route 2, is internalized into early basal endosomes (EBE) and reaches the apical surface after stops in the CE and ARE (route 3). Many basolateral receptors, such as those for transferrin and LDL, are internalized into EBE and are recycled basolaterally from the CE (route 5). A recycling route for apical membrane proteins involves the early apical endosome (EAE) and the ARE (route 4). Some membrane proteins recycling from the apical or basolateral surface may cycle through the TGN from the CE or ARE (8). Internalized fluid or ligands dissociated from their receptors in the EAE or EBE may reach a common late endosome on their way to the lysosome (route 6, 7). (Right) Trapping of plasma membrane proteins by associations with the submembrane cytoskeleton. At the lateral membrane, E-cadherin-mediated cell–cell contacts promote the formation of an ankyrin/fodrin submembrane cytoskeleton that immobilizes Na,K-ATPase. The apical microvilli of certain epithelial cells (e.g., retinal pigment epithelium; Bonilha et al., 1999) express ezrin, a linker protein with binding sites for actin and for certain transmembrane receptors, such as cd44, ICAM-1, and ICAM-2. Ezrin also interacts with ezrin-binding protein of 50-kD (EBP-50), one of whose PDZ domains can immobilize CFTR (Bretscher, 1999).

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The third group of basolateral signals is heterogeneous and includes those of the neural cell adhesion molecule (NCAM) (Le Gall et al., 1997) and the extracellular matrix metalloprotein inducer (EMMPRIN) (Marmorstein et al., 1998). The only transcytotic signal characterized in detail is that of the polymeric IgA receptor (pIg-R). Activation of this signal relies both on the binding of ligand at the basolateral surface and on phosphorylation of a critical cytoplasmic serine residue (as reviewed in Mostov (1994).

7.4 APICAL SORTING SIGNALS INTERACT WITH SPECIALIZED LIPID MICRODOMAINS (RAFTS) OR SORTING RECEPTORS Apical proteins with signals in the luminal or transmembrane domain are thought to require association with specialized membrane microdomains, also called “rafts,” formed by glycosphingolipids, sphingomyelin, and cholesterol, for incorporation into transporting vesicles or tubules destined to the apical surface (Simons and Ikonen, 1997). Lipid rafts appear to be absent from the ER, and are likely assembled in the Golgi complex. The affinity of GPI for rafts is based on the possession of long unsaturated fatty acid chains. Mutation of certain transmembrane domain amino acids prevents the association of influenza HA with rafts, and its apical targeting (Scheiffele et al., 1997). It not yet clear how N-glycans and O-glycans promote apical sorting. Putative lectin receptors have been postulated, but none has yet been identified that fulfills all the properties of a sorting receptor (e.g., Golgi localization, involvement in polarized sorting). The authors have recently proposed the alternative possibility that N-glycans and O-glycans might be required for structural purposes, to prevent aggregation or facilitate incorporation into rafts (Rodriguez-Boulan and Gonzalez, 1999), rather than as ligands for lectin receptors. Whether apical sorting is carried out by rafts or by sorting receptors, the luminal or transmembrane sorting information must somehow be transduced to the cytoplasmic face of the TGN for recruitment of specific adaptors and coat proteins necessary for the assembly of apical transport vesicles or for the interaction with actin or microtubule-associated motors. A puzzling aspect of apical targeting signals is the growing number of exceptions to their ability to target proteins apically. For example, N-glycans are not always sufficient to act as apical targeting signals (Rodriguez-Boulan and Gonzalez, 1999). Similarly, GPI, by itself, is in some cases not sufficient to target all proteins attached by this mechanism in MDCK cells (Benting et al., 1999) and does not act as an apical targeting mechanism in the thyroid cell line FRT (Zurzolo et al., 1993). Furthermore, attachment via rafts is also not sufficient to target proteins to the apical surface (Benting et al., 1999). There is not enough information yet to evaluate the contribution of O-glycanated structures, such as the juxtamembrane domain of the apical marker p75 neurotrophin receptor. A possible explanation of the available data is that apical sorting of membrane proteins is a cooperative event that requires in some cases two apical targeting signals.

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Using adenovirus vectors to overexpress two apically secreted soluble glycoproteins, Marmorstein et al. (2000) have recently shown that the apical pathway can be saturated, as previously shown for basolateral proteins (Matter et al., 1994). Interestingly, when both proteins were expressed simultaneously, overexpression of one protein resulted in missorting of the other, suggesting competition for common sorting receptors. Since tunicamycin did not affect the apical sorting of one of the glycoproteins under study, the authors suggested that the saturable apical receptor was not a lectin. To date, no “apical sorting receptor” has been identified.

7.5 BASOLATERAL SORTING SIGNALS INTERACT WITH SPECIALIZED CYTOPLASMIC ADAPTORS The striking generalization has emerged that certain basolateral targeting signals are structurally highly related to endocytic sorting motifs. For example, both tyrosine and dileucine motifs appear to be effective in promoting the proteins that possess them to be endocytosed via coated pits and targeted basolaterally from the TGN. Endocytic and basolateral signals are not identical, however, as analysis of these signals by alanine mutagenesis detects subtle differences in the structural requirements for endocytosis and basolateral targeting. A satisfying explanation for these observations is provided by the analysis of a family of heterotetrameric adaptor proteins, AP1, AP2, AP3, and AP4, that recognize these signals at specific subcellular compartments. Whereas the mechanisms that target APs to specific cellular locales are still unknown, progress has been made in the characterization of their interaction with sorting signals (Bonifacino and Dell’Angelica, 1999). Tyrosine motifs bind the µ subunit of the APs in yeast 2 hybrid assays (Ohno et al., 1995), whereas the dileucine motif reportedly binds the β subunit of AP2 (Rapoport et al., 1998). The cocrystallization of the signal-binding C-terminal domain of µ2 with the endocytic signal peptides of EGF receptor and the TGN protein TGN38 has yielded a wealth of information on the structural details of these interactions (Owen and Evans, 1998). Because the key amino acids involved in the interactions with the signal peptide are highly conserved in the various µ subunits, it is possible to test the participation of a given AP in a specific sorting process using a dominant negative approach (Nesterov et al., 1999). A subset of AP1 characterized by the possession of a different µ subunit, µ1B, is found exclusively in epithelial cells in contrast to the ubiquitous µ1A (Folsch et al., 1999). The pig kidney epithelial cell line LLCPK lacks µ1B and displays defects in the sorting of certain basolateral proteins, such as LDL and transferrin receptors and the alpha subunit of Na,K-ATPase (Rousch et al., 1998; Folsch et al., 1999). Transfection of µ1B corrects the sorting defect. It is interesting to mention that not all basolateral proteins possessing tyrosine motifs are missorted by LLCPK cells, indicating that a multiplicity of mechanisms participates in the sorting of basolateral proteins. A completely sequenced human genome augurs a new era in which all candidate adaptor proteins involved in sorting will be known. Indeed, a new family of TGN

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adaptors with homology to the γ subunit of AP1 has been recently identified through BLAST (Dell’Angelica et al., 2000; Hirst et al., 2000).

7.6 ROLE OF MICROTUBULE MOTORS IN EXIT FROM THE TGN AND TRANSPORT TO THE PLASMA MEMBRANE The role that microtubules (MTs) and their associated motor proteins play in the post-TGN trafficking and efficient delivery of proteins to the plasma membrane is controversial. There is overwhelming evidence that MTs are required for longdistance transport of membrane-bound organelles and vesicles, e.g., to the tips of neuronal axons. Interference with microtubules or with the anterograde microtubulemediated transport by antibody injection or by antisense treatment causes the accumulation of synaptic vesicles in the cell body (Caceres and Kosik, 1990). However, the role of MTs and MT-associated motors in post-Golgi transport in non-neuronal cells, such as fibroblasts, endothelial, and epithelial cell types, is considerably less clear. Initial experiments in fibroblasts have shown that disruption of microtubules by colchicine or nocodazole does not impair transport of plasma membrane proteins to the cell surface (Rogalski et al., 1984). In epithelial cells, the effect of microtubule disruption on transport to the cell surface appears to depend on the cell type and on the protein under study. Some in vivo studies with microtubule depolymerizing agents have detected alterations in the polarized distribution of certain plasma membrane proteins, mainly apical ones, and a delay in their direct or transcytotic delivery to the apical cell surface (Rindler et al., 1987; Breitfeld et al., 1990; Hunziker et al., 1990; Matter et al., 1990b; Gilbert et al., 1991a; Saunders and Limbird, 1997). Other studies failed to detect any alteration in the polarized distribution of surface proteins although they noticed a delay in apical surface delivery (Salas et al., 1986; van Zeijl and Matlin, 1990). Furthermore, in vitro assays in cells permeabilized with SLO have implicated the motors kinesin and dynein in apical transport, and kinesin in basolateral transport (Lafont et al., 1994). An important shortcoming of the studies using microtubule depolymerizing agents is that these agents also cause intense fragmentation and redistribution of mini Golgi apparatus to peripheral cell regions while keeping them close to the widely distributed ER. Under these conditions, cargo proteins might be transported through a much shortened ER → Golgi → plasma membrane route using exclusively the actin cytoskeleton surrounding the Golgi complex (see below) and underlying the plasma membrane, which would account for the insensitivity to microtubule disrupting agents. Recent evidence supports a key role for microtubules and associated motors in post-Golgi transport under physiological conditions, with an intact microtubule system and a central Golgi complex. Live imaging experiments with apical and basolateral marker proteins coupled to green fluorescent protein (GFP) have shown that exit from the Golgi complex toward the plasma membrane occurs via two types of transport intermediates, vesicles and tubules of variable length (1 to 3 µm) (Hirschberg et al., 1998; Toomre et al., 1999; Kreitzer et al., 2000). In spread cells,

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where the microtubules have a centrosome origin, both vesicles and tubules are transported centrifugally through the cytosol along linear microtubule paths with speeds of ~1 µm/s, compatible with kinesin (Figure 7.2). Importantly, block of kinesin function with antibodies completely inhibits exit of GFP-tagged p75 neurotrophin receptor (p75-GFP) (Kreitzer et al., 2000). High-resolution microscopy demonstrates that the formation of tubules, but not the release of vesicles, is inhibited by kinesin antibodies; these vesicles only undergo Brownian movements and are never seen to displace over long distances. Interestingly, dominant negative forms of dynamin II, a molecule that participates in the fission of coated vesicles from the plasma membrane, completely blocks the release of vesicles and tubules, but does not affect the formation of tubules, which extend and retract like normal tubules but fail to undergo fission.

7.7 ROLE OF THE ACTIN CYTOSKELETON IN GOLGI EXIT AND ARRIVAL AT THE CELL SURFACE The discovery of actin-binding proteins, notably spectrin, ankyrin, and several myosins (I, II, V, and VI), at the Golgi and TGN has led to the speculation that budding and fusion events at this organelle are also regulated by an actin-based cytoskeleton (De Matteis and Morrow, 1998; Heimann et al., 1999; Valderrama et al., 2000). However, in contrast to cortical actin microfilaments, Golgi-associated actin filaments are not readily detectable when cells are probed with phalloidin, suggesting that a Golgi-based actin network might be highly dynamic. Indeed, labeling of perforated cells with fluorescent actin demonstrates a perinuclear actin network that surrounds the Golgi complex (Müesch et al., 2001). This network appears to be regulated by cdc42. Activated forms of cdc42 promote disappearance of the periGolgi actin network and thickening of the cortical network. Surprisingly, however, there is little functional evidence for the involvement of actin in the constitutive secretory pathway. Disruption of the actin cytoskeleton failed to affect secretion in several constitutively secreting cell lines (Griffin and Compans, 1979; Genty and Bussereau, 1980; Salas et al., 1986) and did not alter the kinetics of post-Golgi transport to the cell surface when only this leg of the secretory pathway was analyzed (Babia et al., 1999; di Campli et al., 1999). However, since most of these studies focused on multistep pathways, the negative data obtained might reflect opposite roles of the actin cytoskeleton at the level of the individual steps (e.g., budding, transport, fusion) of the pathway. Indeed, there is evidence that actin filaments are positively involved in vesicle fusion and fission events in cortical areas but represent a barrier for the association of vesicles with the plasma membrane and for the expression of membrane plasticity (Aunis and Bader, 1988; Bretscher, 1991; Muallem et al., 1995). Recently developed imaging techniques that follow the exit of GFP-tagged proteins from the Golgi in vivo revealed that the exit of both apical and basolateral proteins from the TGN is facilitated by microfilaments, since actindisrupting agents delayed the exit of VSVG-GFP from the TGN in fibroblasts (Hirschberg et al., 1998) and of LDLR-GFP and p75-GFP in MDCK cells (Müsch et al., submitted).

FIGURE 7.2 Trafficking of tubular and vesicular post-Golgi transport intermediates containing apically targeted p75-GFP. Tubular and vesicular transport structures containing an apical membrane protein coupled to green fluorescent protein (p75-GFP) emerge from the Golgi complex and follow linear paths toward the cell periphery at rates consistent with kinesin-driven, microtubule-dependent motility. The protein was accumulated in the TGN of MDCK cells by microinjection of its cDNA into the nucleus and successive incubations of the cells at 37°C (1 h) and 20°C (2.5 h). The four top panels show the extension, fission, and linear movement of a single post-Golgi tubule (white arrow) and the extension of a second tubule (star). The five middle pannels show the movement of a single vesicle (circle); the cumulative movements along a linear path are shown on the right panel of the series. The bottom panel shows the loss of p75-GFP fluorescence from the Golgi complex as a function of time (triangles) and the block in TGN exit promoted by injection of an antikinesin antibody (squares) or by expression of a GTPase-deficient form of dynamin (circles). Scale bar represents 2 µm. (From Kreitzer, G. et al., Nat. Cell Biol., 2, 125, 2000. With permission.)

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FIGURE 7.2 (continued.)

7.8 CONTROL OF POLARITY BY SMALL GTPASES The GTPases of the rho-family rhoA, rac1 and cdc42, are key regulators of cortical actin polymerization and modulate various exocytic and endocytic vesicle transport steps at the plasma membrane (Price et al., 1995; Lamaze et al., 1996; Gasman et al., 1997; 1999; Brown et al., 1998; Jou et al., 2000). Interestingly, Erickson et al. (1996) have reported the presence of cdc42 in the Golgi complex, where it binds in a Brefeldin A (BFA)-dependent manner characteristic of coat proteins that generate transport vesicles. A similar BFA-sensitive association with the Golgi has also been shown for the actin-binding proteins spectrin (Beck et al., 1994), ankyrin (Godi et al., 1998), and myosin IIA (p200) (Narula et al., 1992). A recent report (Kroschewski et al., 1999) has implicated cdc42 in the regulation of apical/basolateral polarity in MDCK cells. The authors observed a reversal in the polarity of a basolateral protein (VSV G protein coupled to GFP) in cells microinjected with cDNAs of VSVG-GFP and an activated form of cdc42. Although these experiments suggest that cdc42 might be involved in protein sorting in the secretory pathway,

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no specific mechanisms were uncovered. The GTPase could regulate the sorting of apical and basolateral proteins at the TGN or, alternatively, it might control the specificity of vesicle fusion at the cell surface. Indeed, it has been shown that rac1, a downstream effector of cdc42, regulates cytoskeletal complexes at cell–cell adhesion sites, including tight junctions (Jou and Nelson, 1998), that provide spatial cues for the basolateral vesicle docking machinery (Grindstaff et al., 1998). Recent work indicates that the TGN exit of apical glycoproteins is enhanced by activated and dominant-negative forms of cdc42, whereas the TGN exit of basolateral proteins is slowed (Müsch et al., 2001). These experiments suggest a role of cdc42 in the exit pathways from the Golgi complex. Since most, if not all, effector cascades of cdc42 result in a reorganization of the actin cytoskeleton, the regulation of microfilaments appears to be important for protein exit from the TGN. In agreement with this idea, in vitro experiments suggest the participation of myosin II in the exit of basolateral but not apical proteins from the Golgi complex (Müsch et al., 1997). Other myosins with a Golgi localization, such as Myosin I, V, and VI, could also be involved in specific exit steps for certain transported proteins (reviewed in Olkkonen and Ikonen, 2000).

7.9 THE TIGHT JUNCTION/ZONULA ADHERENS: A HOT AREA FOR SURFACE TARGETING? The process of vesicle docking and fusion of post-Golgi vesicles and tubules with the plasma membrane involves a large number of membrane and cytosolic components, some of which have been identified. The core fusion machinery is thought to consist of the v-SNAREs in vesicles or tubules and the t-SNAREs in the target apical and basolateral membranes (Rothman and Warren, 1994). Different t-SNARES are found on apical and basolateral surfaces; syntaxin 3 is apical, syntaxin 4 is basolateral, and syntaxin 2 and syntaxin 11 as well as snap23 are found on both surfaces (Weimbs et al., 1997). Although the SNAREs are thought to account for the specificity of vesicle fusion with the right surface domain, it is clear that additional factors organize vesicle fusion in space and time. In yeast, SNAREs are distributed all over the cell surface; yet vesicle fusion is restricted to the bud site. Small GTPases of the rab-family (Brennwald, 2000) and a protein complex called the exocyst (TerBush et al., 1996) have been characterized to regulate vesicle fusion. Recently, yeast and neuronal homologues of the Drosophila tumor suppressor lethal (2) giant larvae (Jacob et al., 1987; Lehman et al., 1999) emerged as proteins involved in post-Golgi vesicle transport and synaptic vesicle fusion (Wodarz, 2000). Interestingly, Drosophila L(2)gl is localized at the lateral membrane of epithelial cells and essential for the proper development of larval epithelia. These data raise the possibility that lgl may be involved in establishing epithelial polarity by spatially organizing basolateral secretion. Studies are under way to test this hypothesis in cell culture models. Rab proteins and the exocyst have been identified and characterized in the regulation of basolateral vesicle fusion with the plasma membrane in MDCK cells (Huber et al., 1993; Grindstaff et al., 1998). Rab8 is specifically associated with basolateral postGolgi transport vesicles and inhibition of rab8 function affects basolateral exocytosis.

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Addition of antibodies against mammalian exocyst components sec6/sec8 to permeabilized MDCK cells selectively blocks the delivery of basolateral protein LDLR but does not affect the delivery of the apical marker p75. Sec6/sec8 localizes intracellularly in subconfluent MDCK cells but quickly redistributes to the zonula adherens region of the lateral cell surface when MDCK cells reach confluency. The junctional localization of the exocyst suggests that delivery of basolateral transport carriers to the cell surface might occur at that region of the cell. However, no direct evidence is available on the site of initial docking and fusion of post-Golgi transport intermediates in polarized cells. An indirect approach used by Louvard (1980) attempted to define the site of surface insertion of an apical membrane protein. When the apical hydrolase leucine amino peptidase was removed from the cell surface by exposure to cross-linking antibodies, the internalized protein reappeared at sites of cell–cell contact. However, live imaging data with evanescent field microscopy, which provides a high-resolution view of events within 100 nm of the basal cell membrane lying on the coverslip, show that, in subconfluent cells, vesicular and tubular transporters carrying VSV G protein coupled to GFP fuse at high rates with the basal surface of fibroblastic cell lines (Schmoranzer et al., 2000; Toomre et al., 2000). The authors have recently shown that this is also true for GFP-tagged apical (p75) and basolateral (NCAM) proteins in subconfluent MDCK cells. However, when the cells become confluent, the number of basolateral protein fusion events is reduced dramatically, even though the protein is still delivered to the cell surface (as demonstrated by the increase in lateral fluorescence). In contrast, the rate of fusion of the apical protein with the basal surface, although reduced, continues at a significant rate. The experiments are consistent with the establishment of a fusion site for basolateral transport away from the basal surface when the cells establish contacts with their neighboring cells. Indirect evidence described above suggests that the tight junction/zonula adherens area is a hot area for docking and fusion of basolateral and apical post-Golgi vesicles and tubules. In addition to the proteins mentioned above, a variety of molecules that are involved in the positioning of tight junctions and zonula adherens, such as E-cadherin and the mammalian equivalents of Drosophila proteins scribble, lethal giant larvae, crumbs, discs large, discs lost might be involved in the setting of a “targeting patch” for post-Golgi vesicles and tubules (Yeaman et al., 1998). Current intense research in this area is bound to uncover exciting mechanisms involved in this process.

7.10 PASSIVE MECHANISMS THAT ACCOUNT FOR THE POLARIZED DISTRIBUTION OF MEMBRANE AND CYTOSKELETAL PROTEINS Figure 7.1 (left) demonstrates the role of tight junctions in the maintenance of cell polarity. Once apical and basolateral membrane proteins and lipids are delivered to the apical or basolateral surfaces by the mechanisms discussed above, the tight junction acts as a fence that prevents their intermixing (Cereijido et al., 1989). This fence is effective for lipids present in the inner leaflet but not for those in the outer leaflet. Figure 7.1 (right) illustrates two cytoskeletal “traps” for membrane proteins.

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The actin-binding protein ezrin, a member of the ERM (Ezrin, Radixin, Moesin) protein family, can bind and immobilize membrane proteins directly or via an adaptor protein (Bretscher, 1999) (see Figure 7.1, legend). Similarly, the Na,K-ATPase is immobilized at the lateral membrane by an actin–fodrin cytoskeleton assembled upon establishment of E-cadherin-mediated cell–cell contacts (Nelson, 1992).

ACKNOWLEDGMENTS Supported by grants from the National Institutes of Health and a Jules and Doris Stein award from the Research to Prevent Blindness Foundation to E.R.B. and an NRSA award to G.K.

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8

Biogenesis of Epithelial Polarity and Tight Junctions Liora Shoshani and Ruben Gerardo Contreras

CONTENTS 8.1 8.2 8.3

8.4

8.5 8.6

8.7 8.8

Introduction ..................................................................................................166 Ubiquity of Polarity .....................................................................................166 Polarity-Maintaining Mechanisms...............................................................166 8.3.1 Selective Targeting ...........................................................................167 8.3.2 Selective Stabilization ......................................................................167 Epithelial Cell Adhesion ..............................................................................168 8.4.1 Cell–Cell Contacts ...........................................................................168 8.4.1.1 Adherens Junctions...........................................................168 8.4.1.2 Tight Junctions..................................................................168 8.4.2 Cell–Extracellular Matrix Contacts .................................................169 8.4.3 Cell–Cell Contacts and Signaling....................................................169 Cytoskeleton.................................................................................................170 Biogenesis of Epithelial Cell Polarity .........................................................171 8.6.1 External Signals Induce Cell Surface Asymmetry and the Axis of Polarity.........................................................................................171 8.6.1.1 Ca2+-Activated Cell–Cell Contacts ...................................171 8.6.1.2 Interactions with Extracellular Matrix Components........172 8.6.1.3 Other Extracellular Associations ......................................173 8.6.2 Assembly of Cytoskeleton Proteins and Signaling Complexes at Sites of Cell Contacts ..................................................................175 8.6.2.1 Cytoskeleton Assembly at Focal Adhesions ....................175 8.6.2.2 Role of Cytoskeleton Assembly in Cell–Cell Adhesion ...........................................................................176 8.6.3 Structures That Specify Targeting and Retention of Membrane Proteins Synthesized de Novo..........................................................177 8.6.3.1 Targeting Patches ..............................................................177 8.6.3.2 Membrane Skeleton ..........................................................178 Polarity of Tight Junctions...........................................................................179 Polarity of Adherens Junction Proteins .......................................................181

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8.9 Dynamics of Cell Membrane Polarity.........................................................182 8.10 Relationship Between Tight Junctions and Adherens Junctions.................182 8.10.1 The Establishment of Structural and Molecular Asymmetry at the Cell Surface Requires an External Signal............................182 8.10.2 The Role of Sec6/8 Complex in the Initial Stage of Cell–Cell Contact ............................................................................................183 8.10.3 Cell Junction Proteins Are Recruited to Initial Lateral Membrane .......................................................................................185 8.10.4 Membrane Skeleton Stabilizes Membrane Proteins ......................186 8.10.5 TJ Is a Targeting Patch ...................................................................187 8.10.6 Selective Targeting and Stabilization Maintain Epithelial Contacts and Polarity......................................................................187 8.11 Concluding Remarks....................................................................................188 References..............................................................................................................188

8.1 INTRODUCTION Vectorial transport of substances across epithelia rely on the polarization of the plasma membrane and on the formation of tight junctions (TJs). Until a few years ago it was thought that, since the TJ marks the limit between apical and basolateral domains, it would be responsible for the polarized distribution of membrane components. Apart from the fact that, as pointed out by Cereijido et al. (2000), the TJ cannot sort membrane components, it is now realized that it is just the other way around: the TJ is itself a product of an overall polarizing process, as its assembly results from precise targeting of its molecular component and, once assembled, the TJ can even act as a transitory station for proteins en route to a final, polarized distribution in the membrane. This chapter reviews the generation of epithelial polarity and its relationship with the TJs.

8.2 UBIQUITY OF POLARITY In a broad sense, polarity is not an exclusive feature of the cell membrane, but is reflected in the position of the nucleus, the Golgi apparatus, microvilli, mitochondria, flagella, dendrites, axons, microtubules, microfilaments, and the composition of the extracellular matrix, the basal lamina, etc. Actually, a certain degree of asymmetry, or at least “regionality” in the distribution of membrane components is found in most cells, including some that do not even have a TJ. Thus neurons, spermatozoa, yeast, skeletal muscle fibers, osteoclasts, T cells, etc. have pumps, channels, carriers, receptors, bud daughter cells, and bind viruses in restricted domains of the membrane (Miledi, 1960; Poindexter, 1964; Shapiro, 1985; Baron et al., 1985; Stowers et al., 1995). However, this chapter is restricted to the polarity of the cell plasma membrane of epithelial cells.

8.3 POLARITY-MAINTAINING MECHANISMS Differences in the distributions of membrane proteins between apical and basolateral plasma membrane domains are maintained by protein sorting from intracellular

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compartments also known as selective targeting and selective retention or stabilization in the membrane domain (Rodriguez-Boulan and Nelson, 1989; Yeaman et al., 1999; Matter, 2000).

8.3.1 SELECTIVE TARGETING Newly synthesized proteins are sorted and subsequently packaged into membranebound carriers for delivery to the cell surface in the trans-Golgi network (TGN) (Griffiths and Simons, 1986; Geuze et al., 1987) and in endosomes (Mellman, 1996). To be selectively sorted, proteins must carry polarity determinants that are recognized by specific sorting machinery in both organelles. Apical targeting has been attributed to a number of different types of sorting signals, including glycosylphosphatidylinositol (GPI) anchors (Lisanti et al., 1989; Rodriguez-Boulan and Gonzalez, 1999), O-linked carbohydrates, specific protein transmembrane, lumenal or cytoplasmic determinants, and N-linked carbohydrates, although in this case, it probably results indirectly by association with lectins (Rodriguez-Boulan and Powell, 1992; Rodriguez-Boulan and Gonzalez, 1999). Targeting determinants for basolateral sorting are continuous amino acid sequences in the cytoplasmic domains of membrane proteins. They often rely on critical tyrosinedependent or dileucine-dependent motifs, frequently followed by a cluster of acidic residues (Rodriguez-Boulan and Powell, 1992; Matter and Mellman, 1994; Matter, 2000). Recently, Folsch et al. (1999) have found that the subunit µ1B of the clathrin adaptor complex AP-1, known to interact with tyrosine-based sorting signals, is implicated in the basolateral sorting of membrane proteins in epithelial cells. Basolateral targeting determinants are generally dominant over apical sorting signals because they interact earlier or more avidly with the sorting machinery (Matter and Mellman, 1994). In the absence of specific sorting signals, transmembrane proteins accumulate in the Golgi apparatus, suggesting that none of the routes to the cell surface is an efficient default pathway (Gut et al., 1998; Wittchen et al., 1999).

8.3.2 SELECTIVE STABILIZATION Some membrane proteins are stabilized through association to the submembrane cytoskeleton. A scaffold of cytosolic proteins bind membrane proteins and associate them to the signaling machinery required for proper function. Specialized membrane domains like synapse or epithelial cell junctions are thus formed and maintained. Src homologous (SH2 and 3), proline-rich, PDZ (PSD-95/Dlg/ZO-1), and ankyrinbinding modules are relevant for epithelial cells (Fanning and Anderson, 1999a). Thus, receptors (Yamada et al., 1999), ion channels (Nehring et al., 2000), ion pumps (Kim et al., 1998; Zhang et al., 1998), and cell–cell junction proteins (Buchert et al., 1999) are expressed in a polarized manner by selective stabilization. In contrast to the numerous examples in higher eukaryotes, no PDZ proteins have been implicated in membrane polarization in yeast, suggesting that this family of proteins has evolved specifically for the maintenance of multiple membrane domains (Shulman and St. Johnston, 1999).

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Selective targeting and stabilization are not mutually exclusive. Thus, newly synthesized Na+,K+-ATPase is polarized by delivery to the lateral membrane, where it clusters to the ankyrin/fodrin cytoskeleton (Caplan et al., 1986; Nelson and Veshnock, 1987; Nelson et al., 1990; Dunbar et al., 2000).

8.4 EPITHELIAL CELL ADHESION Cell adhesion to neighboring cells as well as to the extracellular matrix is central in some of the most fundamental properties of multicellular systems. Selective adhesion sorts embryonic cells into germ layers, guides cell migration and response to extracellular cues, and underlies both differentiation and stability of the differentiated state. Loss or misregulation of adhesion leads to disease processes such as metastatic cancer. However, cell adhesion is a reversible and dynamic process. Cells attach and detach from each other as tissues are formed during development or as cells metastasize in malignant transformation. Cell–cell and cell–substratum contacts are specified by adhesion receptor proteins.

8.4.1 CELL–CELL CONTACTS Adhesion between epithelial cells is mediated mainly by adherens junctions, desmosomes, and TJs. Junctions are constituted by a cluster of transmembrane and intracellular proteins that anchor the cytoskeleton. 8.4.1.1 Adherens Junctions In this junction the attaching transmembrane protein is E-cadherin, a classical Ca2+dependent homophilic cell adhesion molecule (Takeichi, 1991). E-cadherin is characterized by five extracellular structural repeats (EC1 to EC5) and a highly conserved cytoplasmic domain that associates with several cytoplasmic proteins, most prominently the catenins (Ozawa et al., 1989). Catenins are capable of associating with the actin cytoskeleton (Knudsen et al., 1995; Watabe-Uchida et al., 1998; Gumbiner, 2000). There is compelling evidence that protein interactions mediated by both the cadherin ectodomain and cytoplasmic domain participate in cell adhesion (Yap et al., 1997). E-cadherin is a well-known polarity inducing protein that is able to polarize Na+,K+-ATPase in fibroblasts (McNeill et al., 1990). 8.4.1.2 Tight Junctions The TJ is a belt of anastomosing strands that surrounds the lateral membrane of epithelial cell, and seals the outermost end of the intercellular space. Three types of integral proteins constitute TJs strands: occludins (Furuse et al., 1993), claudins (Furuse et al., 1998), and junctional adhesion molecule (JAM) (Martin-Padura et al., 1998). The first two are protein families with four transmembrane segments, two extracellular loops, and a long COOH-terminal cytoplasmic tail. JAM has only one transmembrane segment. Occludin and claudin contribute to the barrier function of the TJs (Balda et al., 1996; McCarthy et al., 1996). Occludin is also involved in the

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fence function of TJs (Inai et al., 1999). Membrane proteins of the TJs have specific binding domains for the TJ cytoplasmic plaque proteins ZO-1, ZO-2, and/or ZO-3 (Furuse et al., 1994; Itoh et al., 1999; Bazzoni et al., 2000). These proteins belong to the membrane-associated guanylate kinase (MAGUK) family of proteins, characterized by having an SH3, a GUK, and as many as three PDZ domains that participate in specific protein/protein associations. Other intracellular proteins are also components of the plaque, like cingulin (Cordenonsi et al., 1999), although its role on TJ functions is unknown, while some others, i.e., G proteins, protein kinase C (PKC), Ras-binding protein AF6, Sec6/8 are signaling proteins (Dodane and Kachar, 1996; Yamamoto et al., 1997; Izumi et al., 1998) or components of the secretory machinery (Grindstaff et al., 1998). Intracellular proteins associate the membrane TJ proteins to the actin cytoskeleton (Itoh et al., 1997; Fanning et al., 1998). The whole complex is a structural and functional membrane microdomain (Fanning and Anderson, 1999b) that establishes cell–cell contacts, participates in the regulation of paracellular transport, and plays an important role in the generation and maintainance of epithelial cell polarity (see below).

8.4.2 CELL–EXTRACELLULAR MATRIX CONTACTS The attachment of cells to the extracellular matrix (ECM) is of crucial importance in the maintenance of tissue structure and integrity. In stratified epithelia, such as in skin as well as in other complex epithelia, multiprotein complexes called hemidesmosomes are involved in promoting the adhesion of epithelial cells to the underlying basement membrane. Adhesion to ECM or focal adhesion is mediated by integrins, a family of adhesion receptors involved in a diverse array of cellular processes including migration, polarity, survival, growth, and differentiation (Burridge and Chrzanowska-Wodnicka, 1996). The α and β chains of integrin are single transmembrane proteins that assemble in a variety of heterodimers expressed differentially during development. The extracellular domain binds to the three-peptide RGD, present in many proteins of the ECM. The short intracellular domain of the β chain couples with cytoplasmic proteins that nucleate the formation of large protein complexes containing both cytoskeletal (talin, vinculin, paxilin, α-actinin, tensin, and actin) and catalytic signaling (focal adhesion kinase, Src, CAS) proteins (Giancotti and Ruoslahti, 1999; Critchley, 2000).

8.4.3 CELL–CELL CONTACTS

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SIGNALING

Accumulating evidence suggests that cell–cell and cell–substrate contacts function as signal transduction centers (Hynes, 1999). A major feature of these signaling pathways is tyrosine phosphorylation/dephosphorylation of proteins that link the adhesion molecules to the cytoskeleton. In the case of adherent junctions, the catenins are target molecules of this apparently phosphotyrosine-regulated assembly/disassembly (Gumbiner, 2000). In the case of TJs there is evidence indicating that the phosphorylation of occludin (in Ser and Thr residues) is an important step in regulating TJ formation and permeability (Sakakibara et al., 1997; Contreras et al., 1999). On the other hand, there is evidence that tyrosine phosphorylation regulates

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the TJ assembly (Balda et al., 1991; 1993; 1996). In the case of integrin-mediated adhesion there is a chain of tyrosine phosphorylation, in which FAK and Src are primary actors, that regulates the assembly of focal adhesion (Schaller et al., 1994; Giancotti and Ruoslahti, 1999). Cell–cell and cell–substrate adhesion functions are expected to be coordinated during development. In a recent work (Arregui et al., 2000), Fer, a nonreceptor tyrosine kinase, was found to mediate the cross talk between adherent junctions (N-cadherin) and focal adhesions (β1-integrins). The accumulated evidence suggests that the components of the junctional plaque function as signaling proteins that control processes that occur outside the junctions, i.e., the nucleus. Interestingly, β-catenin and plakoglobin, members of the armadillo family, participate in wingless/Wnt signaling pathway (Papkoff et al., 1996). Moreover, β-catenin and plakoglobin interact with the transcription factor LEF-1 and this complex associates with the 5′ end of E-cadherin gene (Behrens et al., 1996; Huber et al., 1996). These findings strongly suggest that junctional proteins can regulate the expression of target genes. Moreover, there is significant evidence for a possible role of catenins in tumor supression (Waltzer and Bienz, 1999; Hinoi et al., 2000; Seidensticker and Behrens, 2000). Interestingly, in Caenorhabditis elegans the functions of β-catenins in adhesion and in signaling are performed by separate proteins BAR-1 and HMP-1, respectively (Korswagen et al., 2000).

8.5 CYTOSKELETON Submembrane scaffolding and cross-linking proteins play an important role in establishing and maintaining the polarized distribution of some membrane proteins. The cytoskeletal elements participate in several important aspects of the life of the cell, including cell shape, cell motility, cell division, and signal transduction in addition to their involvement in cell polarity. The mammalian cell cytoskeleton consists of a diverse group of dynamic fibrillar elements that consists of three highly abundant major protein families: microfilaments (MF), microtubules (MT), and intermediate filaments (IF), as well as a growing number of associated proteins. The prototype members of these three protein families are actins, tubulins, and keratins, respectively. The actin cytoskeleton formation and remodeling underlies the fundamental processes of cell motility and shape determination. To serve these roles, different compartments of the actin cytoskeleton engage in forming specific coupling sites between neighboring cells and with the underlying matrix, which themselves serve signal transducing functions. Microtubules are bundled and stabilized by various microtubule-associated proteins. Many differentiated cells including polarized epithelial cells display a nonradial, apico-basal microtubule array. While microtubule arrays in cells are often focused at the centrosome, a variety of cell types contain a substantial number of noncentrosomal MTs. The Rho family of small GTPases plays important roles in regulating actin cytoskeleton organization and cell adhesion (Takaishi et al., 1997; Hall, 1998; Nobes et al., 1998).

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8.6 BIOGENESIS OF EPITHELIAL CELL POLARITY Like embryonic patterning, which begins with the polarization of the body axes, patterning at the single-cell level starts with the specification of an axis of cell polarity. In some cases, these two processes are identical: in organisms such as C. elegans, Drosophila and Xenopus, the main body axis of the animal is defined by the polarity of the single-cell zygote. Thus, cell polarity presents cell biologists with many of the same conceptual challenges as developmental patterning but on a much smaller scale. Cells not only polarize, but they polarize in a specific direction, and therefore they must respond to asymmetric cues, which can be either intrinsic or extrinsic to the cell.

8.6.1 EXTERNAL SIGNALS INDUCE CELL SURFACE ASYMMETRY AND THE AXIS OF POLARITY The generation of epithelial cell polarity begins with the establishment of both physical and molecular asymmetry in the cell surface. It is well established that extracellular contacts, either with neighboring cell or with ECM (extrinsic cues), initiates the polarized segregation of membrane domains into apical and basolateral domains, initially defined as contacting and noncontacting surface domains. 8.6.1.1 Ca2+-Activated Cell–Cell Contacts TJ formation and apical/basal polarization depend on Ca2+ activated cell contacts (Gumbiner and Simons, 1986; Gumbiner et al., 1988; Gumbiner, 2000). Cells plated at subconfluence in media with normal Ca2+ concentrations or at confluence but in low Ca2+ do not develop TJs, nor do they polarize membrane components like the Na+,K+-ATPase and voltage dependent ion channels, but maintain them in intracellular pools (Figure 8.1). When cells are incubated in conditions that restore Ca2+activated cell–cell contacts, TJs form rapidly (Gonzalez-Mariscal et al., 1985) and start to express Na+,K+-ATPases and ion channels in a polarized fashion (Contreras et al., 1989). Ca2+ is needed primarily on the extracellular side (Gonzalez-Mariscal et al., 1990; Contreras et al., 1992). Contact formation promoted by Ca2+ activates a cascade of intracellular reactions, which includes PKC, phospholipase C (PLC), and calmodulin (CaM) (Gonzalez-Mariscal et al., 1990; Balda et al., 1991), that participates on formation of TJs (Figure 8.2) as well as restoration of the plasma membrane and on the polarized expression K+ ion channels (Ponce et al., 1991a; b; Ponce and Cereijido, 1991). Ca2+ is also important for the maintenance of the TJ, as these can be opened and resealed by removal and restoration of Ca2+ in diverse experimental conditions (Martinez-Palomo et al., 1980; Citi, 1992). The cell-membrane receptor for external Ca2+ is E-cadherin (Yap et al., 1997). Experiments with Madin–Darby canine kidney (MDCK) monolayers cultured in Ca2+-depleted medium or at conditions avoiding cell–cell contacts (Contreras et al., 1989) demonstrate that cell–cell adhesion and specifically adherens junction formation are required to restrict the localization of basolateral membrane proteins in the plasma membrane.

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FIGURE 8.1 Polarized expression of Na+,K+-ATPase depends on Ca2+-activated cell–cell contacts. MDCK cells were plated on glass coverslips for 24 h, fixed, and processed by immunofluorescence with antibodies against the Na+,K+-ATPase β subunit. Subconfluent cells cultured in medium with Ca2+ (A). Confluent cells cultured without (B) or with Ca2+ (C). Cells cultured on plastic dishes were incubated at confluence (left column, D and E), at confluence without Ca2+ (center column, D and E). Cells at confluence with normal Ca2+ (right column, D and E) and membrane (D) and total Na+,K+-ATPase (E), were detected by 3H-ouabain binding or immunoblot, respectively (Contreras et al., 1989; 1999).

8.6.1.2 Interactions with Extracellular Matrix Components Integrin-mediated cell adhesion to ECM is particularly important for organizing the apicobasal axis of epithelial cell polarity. Apical orientation toward the outside of epithelial cells grown in suspension and forming a clump is inverted by the addition of collagen (Chambard et al., 1981; Barriere et al., 1988). Thus, the orientation of apical and basolateral membrane domain is relative to the biological compartments

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separated by the epithelium, hence the direction of ion and solute transport. Interestingly, intercalated cells of the collecting tubule exist in a spectrum of types. Whereas the alpha form secretes acid by an apical H+-ATPase and a basolateral anion exchanger which is an alternatively spliced form of the red cell band 3 (kAE1), the beta form secretes HCO3– by having these transporters on the opposite membrane (Al Awqati et al., 2000). In a clonal cell line of the beta form, seeding density causes this conversion. A protein, termed hensin, a member of the scavenger receptor family, was deposited in the ECM of high-density cells that reversed the polarity of the transporters. Hensin also induced the expression of the microvillar protein, villin, and caused the appearance of the apical terminal web proteins cytokeratin 19 and actin, all of which led to the development of an exuberant microvillar structure (Hikita et al., 1999). In addition, hensin causes beta cells to assume a columnar shape. All these studies demonstrate that the conversion of polarity in the intercalated cell, at least in vitro, represents terminal differentiation and that hensin is the first protein in a new pathway that mediates this process (Hikita et al., 1999; Al Awqati et al., 2000). 8.6.1.3 Other Extracellular Associations Interactions of Ca2+-independent cell adhesion molecules with ligand on adjacent cells could also account for cell polarity generation. Plasma membrane proteins in nonpolarized cells are randomly distributed. Nevertheless, expression of exogenous adhesion proteins in fibroblasts results in concentration of these proteins in the contacting plasma membrane domain (Nagafuchi et al., 1987; McNeill et al., 1990; Van Itallie and Anderson, 1997). Interestingly, the endothelial cell adhesion molecule-1 (PECAM-1/CD31), which localizes to cell–cell contacts in endothelia, keeps this localization even when transfected into COS or 3T3 cells, suggesting that recruitment of PECAM-1 to cell–cell borders is an intrinsic property of the molecule (Zocchi et al., 1996). Sun et al. (2000) recently described an extracellular mechanism that stabilizes the localization of PECAM-1 molecule at endothelial cell–cell borders. They studied the localization to cell–cell contacts of mutants and chimeric constructs transfected in non-PECAM-expressing cells, identifying the protein domains that are sufficient to direct efficient localization of the molecule to the cell–cell border. Interestingly they found that only constructs that support PECAM-1-mediated adhesion localize to cell–cell borders. Therefore, they suggest that PECAM-1 movement in the cell membrane occurs relatively freely until the stabilized extracellular domain of the molecule encounters its ligand on an adjacent cell. When this occurs, the complex is captured at the cell–cell interface, leading to localization at cell–cell borders. The authors propose a similar mechanism for Na+,K+-ATPase, a typical lateral membrane protein in epithelial cells. As described above, this protein is randomly distributed in nonpolarized MDCK cells kept in low Ca2+ concentration in the culture medium, but when this ion is added and cell–cell contacts are established, it is concentrated in the lateral membrane of adjacent MDCK cells. Interestingly, when MDCK cells contact other epithelial cell types, such as Ma-104, the lateral membrane protein no longer concentrates in this mixed cell border (Contreras et al., 1995a; Cereijido et al., 1998; 2000) (Figure 8.3). Since the β2-subunit of Na+,K+-ATPase

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functions as an adhesion molecule in glia cells (Gloor et al., 1992) and since Na+, K+-ATPase distribution in neuroepithelia depends on specific cell–cell contacts (Rizzolo, 1999), one can hypothesize the existence of a third polarization mechanism for this ion pump, which depends on anchorage to an extracellular molecule. In the case of the Na+, K+-ATPase anchoring is probably provided by the same β subunit.

8.6.2 ASSEMBLY OF CYTOSKELETON PROTEINS AND SIGNALING COMPLEXES AT SITES OF CELL CONTACTS Cadherin- and integrin-mediated adhesions induce the localized assembly of specialized proteins at the contacting membranes. Association of actin cytoskeleton with both cadherin and integrin adhesion receptors might serve to reinforce the spatial cues provided by extracellular contacts. 8.6.2.1 Cytoskeleton Assembly at Focal Adhesions The cytoplasmic face of focal adhesions provides an attachment site for bundles of actin filaments known as stress fibers. Clustering of integrins and the association with actin microfilaments induce the assembly of a network of signaling proteins that transmits the spatial information to the interior of the cell. Current models postulate that once integrins bind ECM components, they form complexes with signaling proteins, such as tyrosine kinases, through the cytoplasmic tail. Consequently, a signaling cascade is initiated. One of the phosphorylated proteins is FAK, which localizes to focal adhesions and serves to recruit several other signaling proteins including nonreceptor tyrosine kinases, members of the Ras–MAPK pathway, as well as small GTP-binding proteins (Miyamoto et al., 1995). In vitro assays for integrin association with cytoskeleton and with actin-binding proteins, such as vinculin, paxillin, and tensin, reveal that actin filaments may be linked to integrins directly through α-actinin and talin, or indirectly through vinculin and tensin (Clark and Brugge, 1995; Yamada and Miyamoto, 1995).

FIGURE 8.2 Initial steps in the assembly of the TJ during a calcium switch. In the absence of Ca2+ the two molecules of E-cadherin (bottom) are inactive. The addition of this ion (small dots) elicits two main effects: it binds to the extracellular repeats, and promotes complexing of E-cadherin with α- and β-, or with α- and γ-catenins (β and γ seem to be exchangeable, upper right). In turn, the complex so formed binds to p120, vinculin, α-actinin, and, indirectly, to the cytoskeleton of actin. Ca2+ also causes clustering of E-cadherins in the plane of the membrane, which acquire the capacity to bind to E-cadherin in the neighboring cell, through their outermost repeat. ECCDI, an antibody that binds specifically to the first extracellular E-cadherin repeat, blocks TJ formation and opens already sealed TJs. A putative contact receptor activates PLC through two different G proteins. PLC then splits PiP2 into IP3 and diacylglycerol (DAG). DAG activates PKC. La3+ prevents penetration of Ca2+ into the cells (upper right), but does not impair E-cadherin triggering of TJ formation. Cd2+, instead, has affinity for both the mechanisms that translocate Ca2+ across the cell membrane and for the extracellular moiety of E-cadherin. The affinity of Cd2+ for this moiety is enough to block Ca2+ effect, but not enough to replace this ion in the triggering of TJ formation.

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FIGURE 8.3 Fluorescence image of monolayers formed by mixed populations of MDCK and Ma104 (monkey) cells. Ma104 cells were labeled beforehand with 50 µM CMTMR (A, B). Na+,K+-ATPase β subunit from MDCK cells stained with a monoclonal antibody (C, D) is only observed in MDCK–MDCK borders (filled arrow) but not in MDCK borders in contact with Ma104 cells (empty arrow). Images obtained by confocal laser scanning microscopy from the X–Y (A, C) and X–Z (B, D) planes (Contreras et al., 1995b).

8.6.2.2 Role of Cytoskeleton Assembly in Cell–Cell Adhesion The actin cytoskeleton is central for intercellular adhesion. Yonemura et al. (1995) and Adams et al. (1996; 1998) described the formation of initial cell–cell contacts as dots or “puncta” in which E-cadherin and adherent junction molecules are concentrated. As the region of intercellular contact grows, new puncta are added. Timelapse imaging and electron microscopy studies suggest that puncta are spatially coincident with membrane attachment sites for actin filaments that branch from the cortical actin cytoskeleton. As cell–cell adhesion proceeds, the actin cytoskeleton remodels, concomitantly with changes in puncta distribution (Adams and Nelson, 1998). The function and mechanism underlying this rearrangement remain to be elucidated. However, several reports suggest that small GTPases of the Rho family may be involved (Braga et al., 1997; Braga, 1999; Kodama et al., 1999). Nevertheless, little is known about the mechanisms of this interesting puncta formation and actin dynamics associated with intercellular adhesion. Mouse epidermal keratinocytes stimulated with calcium form filopodia that penetrate and embed into neighboring cells forming an “adhesion zipper,” and initiate adherens junction formation (Vasioukhin et al., 2000). Ultrastructural analysis of filopodia reveal that each filopod contains densely packed cytoskeleton filaments composed almost exclusively of actin, and that adherens junctions are formed at the tips. Adhesion zipper formation depends on actin cytoskeleton (Adams et al., 1996; 1998; Adams and Nelson, 1998). Thus, zippers do not assemble in the presence of cytochalasin D, an inhibitor of actin polymerization, and even though E-cadherin was concentrated at the tips of contacting cells, contacts do not seal and epithelial sheets do not form. Furthermore, immunofluorescence analysis shows that each punctum was associated with a cellular actin filopodium where actin polymerizes

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actively (Vasioukhin et al., 2000). Further investigations revealed that VASP, Mena, zyxin, vinculin, and α-actinin, all known proteins that participate in actin reorganization and polymerization (Machesky et al., 1999), are localized in puncta in the same stage of epithelial sheet formation. Furthermore, α-catenin was found to be essential for actin organizing proteins as well as for adhesion zipper and epithelial sheet formation.

8.6.3 STRUCTURES THAT SPECIFY TARGETING OF MEMBRANE PROTEINS SYNTHESIZED

RETENTION DE NOVO AND

Once plasma membrane asymmetry is generated by cell–cell and cell–ECM contacts, apical and basolateral proteins synthesized de novo must be inserted into the correct plasma membrane domain. At least three membrane-associated structures are suggested to realize this function: (1) targeting patches for docking and fusion of vesicles carrying apical or basolateral proteins (Yeaman et al., 1999); (2) membrane skeleton that helps to direct the retention and accumulation of specific proteins in different membrane domains (Nelson and Veshnock, 1987); and (3) tight junctions, providing a physical barrier that prevents the intermixing of membrane proteins as well as lipids in the outer leaflet of the bilayer (Dragsten et al., 1981; van Meer et al., 1986). 8.6.3.1 Targeting Patches A mechanism to ensure polarized delivery of transport vesicles to contacting and noncontacting membrane is likely to be established immediately after the onset of cell adhesion. Since recognition, docking, and fusion of vesicles must be a very specific and accurate process, it is proposed that as a consequence of the initial cellcontact formation, distinct vesicle targeting sites are assembled on each membrane domain. These patches specify and enhance the efficiency of vesicular trafficking to the correct surface domain and prevents docking and fusion with the incorrect membrane domain. Knowledge about the mechanism that results in such specific interactions between TGN-derived transport vesicles and the plasma membrane of polarized cells is scarce. Fusion of intracellular membranes in eukaryotic cells involves several protein families including SNAREs, Rab proteins, and Sec1/Munc-18 related proteins (Jahn and Sudhof, 1999). SNAREs reversibly assemble into tightly packed helical bundles, named the core complexes. Assembly is thought to pull the fusing membranes closely together, thus inducing fusion. A basic tenet of the SNARE hypothesis is that the minimal machinery for membrane fusion is a cognate set of v-SNAREs and t-SNAREs located on opposing membranes. A corollary to this hypothesis is that these SNARE proteins are prevented from spontaneous assembly by clamping proteins. Recent evidence suggests that Rab proteins (sec4p, rab8, rab10, and rab13) function in the initial membrane contact connecting the fusing membranes but are not involved in the fusion reaction itself. Interestingly, rab8 and rab13 are enriched on the plasma membrane at the apical junctional complex, which includes the TJs and the adherens junctions (Huber et al., 1993). Their restricted distribution depends on cell–cell contact (Weber et al., 1994; Zahraoui et al., 1994). Rab proteins by

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themselves cannot mark a site on the plasma membrane for vesicle docking since it is not restricted to sites of exocytosis on the plasma membrane before the arrival of transport vesicles (Mays and Nelson, 1992; Grindstaff et al., 1998). Proteins that constitute the targeting site become restricted after initiation of cell–cell or cell–ECM contact. Moreover, it should be restricted to sites of exocytosis before the arrival of transport vesicles. Recent work from Nelson’s laboratory indicates that a multiprotein complex consisting of mammalian homologues of the yeast Sec3, Sec5, Sec6, Sec8, Sec10, Sec15, and Exo70 gene products meets the abovementioned criteria (Grindstaff et al., 1998). In yeast, this protein complex is restricted to the plasma membrane sites of active exocytosis (TerBush and Novick, 1995). In yeast, neurons, and MDCK cells these homologous proteins are present in large complexes (Hsu et al., 1999) that appear diffusely in the cytosol in a single MDCK cell. Once cell–cell contacts are induced, Sec6/8 complex is recruited rapidly to the membrane, at sites of cell–cell contacts; moreover, it is codistributed with E-cadherin and ZO-1 along the length of each cell–cell contact, but does not extend beyond the boundary of these contacts. As the monolayer becomes polarized, the distribution of Sec6/8 becomes restricted to the apex of the lateral membrane and no longer extends along the lateral membrane. Nevertheless, disruption of E-cadherin-mediated cell–cell contacts results in dissociation of Sec6/8 complex from the plasma membrane indicating that the localization of the complex at the apical junctional complex is still dependent on Ca+2-dependent cell–cell adhesion. When permeabilized MDCK cells are treated with antibodies against Sec8, the basolateral delivery of vesicles carrying LDL-receptor is reduced significantly, while that of vesicles carrying the apical protein p75NTR is not affected. If the anti-Sec8 antibody interferes with the vesicle docking, the results mentioned above imply that docking and fusion of basolateral vesicles occur near the apical junctional complex. The mechanisms that regulate assembly of targeting patches for transport vesicles on different membrane domains are unknown. Nevertheless, evidence indicates that cellular components involved in sorting and targeting of membrane proteins to the basolateral membrane domain are present in cells before cell–cell adhesion (Grindstaff et al., 1998; Lapierre et al., 1999a). Furthermore, because direct targeting requires cadherin-mediated cell–cell adhesion, it is likely that components of the targeting patch, at least for vesicles containing basolateral proteins, are somehow associated with the cadherin adhesion complex. 8.6.3.2 Membrane Skeleton Specialized cytoskeletal networks assembled at different membrane domains in response to spatial cues may organize vesicle-targeting patches. In turn, these targeting patches specify polarized protein delivery from sorting compartments (Yeaman et al., 1999), thereby reinforcing and maintaining differences in cell surface protein distribution. Formation of the actin cytoskeleton precedes the assembly of a fodrin-based membrane skeleton at sites of cell adhesion (Nelson and Veshnock, 1986). Fodrin, a member of the spectrin family (Bennett, 1990a), is a long, rod-shaped protein that assembles with actin, protein 4.1, aduccin, and others to form a protein membrane

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skeleton. This skeleton is associated with the cadherin/catenin complex, probably through binding of either actin or fodrin to α-catenin. Fodrin also binds to ankyrin, which in turn binds with high affinity to integral membrane proteins, including Na+,K+-ATPase (Nelson and Veshnock, 1987) and Cl–/HCO3– exchanger (Bennett, 1990b). These interactions result in localized assembly of the fodrin-based skeleton and a selective stabilization of specific membrane proteins at sites of cell adhesion in epithelial cells. The native spectrin molecule consists of high-molecular-weight α and β subunits that assemble as elongated heterotetramers. In insect epithelial cells, spectrin associates with ankyrin, which in turn associates with the cell adhesion molecule neuroglian and presumably other integral plasma membrane proteins. The Drosophila Na+,K+-ATPase conserves an ankyrin-binding site (Zhang et al., 1998), and it codistributes with ankyrin and spectrin in polarized fly cells (Baumann et al., 1994; Dubreuil et al., 1997). Yet, despite these conserved features, Na+,K+-ATPase polarity was not altered in epithelial cells from spectrin-null mutants (Lee et al., 1993; 1997). These results led to the conclusion that basolateral accumulation of the Na+,K+ATPase in Drosophila epithelia did not require a stabilizing interaction with the spectrin membrane skeleton. Recently, Dubreuil et al. (2000) using mutations in the Drosophila β spectrin gene provided the first direct evidence that spectrin contributes to the polarized distribution of the Na+,K+-ATPase in epithelial cells and, unexpectedly, that the β subunit of spectrin carries out this role independently of spectrin. Therefore, assembly of a membrane skeleton may be one of the initial steps in propagating signals from extrinsic spatial cues to initiate both localized assembly of specialized membrane domains (targeting patches) and global changes in the organization of other cytoskeletal complexes and membrane compartments. It is well established that the actin cytoskeleton organizes the microvilli at the apical domain and that this organization is regulated by the actin-binding proteins fimbrin and villin (Friederich et al., 1990). Nevertheless, little is known about linkage of membrane proteins to this membrane skeleton, and about the role it may play in stabilizing apically polarized proteins.

8.7 POLARITY OF TIGHT JUNCTIONS Occludin targeting to the TJs seems to rely on a basolateral signal located in its C-terminal domain (Matter and Balda, 1998), on the clustering to the framework of cytosolic proteins (ZO-1, ZO-2, and ZO-3) (Mitic et al., 1999), and on the lateral association to another occludin molecule (Nusrat, 2000). Occludin–connexin chimeras reach the TJs when their clustering activity is preserved. Nevertheless, occludin–Fc or glycophorin chimeras, lacking lateral-clustering ability, are delivered to the basolateral membrane but do not reach the TJs (Matter and Balda, 1998; Fanning et al., 1999). Occludin itself hardly forms TJ strands when expressed in fibroblasts (Furuse et al., 1993; 1996). On the other hand, claudins develop extensive strands expressed under the same conditions (Furuse et al., 1998). Therefore, lateral clustering of claudins is sufficient to develop strands, even when the C-terminal domain, which associates to plaque proteins, is deleted (Furuse et al., 1999). It is worth mentioning that the expression of a fully functional TJ requires more than strands

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detected by freeze fracture. The TJ is formed by a dozen different molecules arranged in a complex manner, it has several functions that can be gauged through different parameters (transepithelial electrical resistance, permeability, fluorescent lipid probes), and its presence is acknowledged when its typical membrane strands are found in freeze-fracture replicas. Obviously, the presence of a single marker or the detection of one of these characteristics does not ensure that the other would necessarily be present (Cereijido et al., 2000). The clustering process is fundamental for the proper localization of TJ proteins (Fanning et al., 1999; Fanning and Anderson, 1999a). Clustering proteins may contain several different modules that specifically bind to segments in other regions of the same polypeptide, or to specific sites in a different molecule. As a result of these interactions, the shape and electronic profile adopted by proteins prompt them to bind specifically to other proteins of the same (multimerization) or different molecular species. In turn, this association induces a redistribution of charges that may enable one of the assembled proteins then to combine specifically with still other proteins that thereby become new members of the cluster. Usually, the affinity of one of the molecules for a given protein species is increased (or decreased) not only by the interaction with a third protein molecule, but also by phosphorylation, combination with Ca2+, K+, or Na+ changes in local pH, etc. Therefore, the assembly of adherens junctions, TJs, and focal adhesions and the link of several of its proteins in a scaffold that includes the cytoskeleton may result from a series of interactions/inductions (Cereijido and Rotunno, 1971) that were initiated by the external cues. Incorporation of occludin to the TJs is therefore a result of a selective targeting to the basolateral membrane through the specific delivery machinery (see below). Once in the basolateral membrane, occludin is sequestered in the framework of plaque proteins through direct association to ZO-1, ZO-2, and ZO-3, and recruited to the TJ. This process is regulated by serine and threonine phosphorylation since just the highly phosphorylated occludin reaches the TJs, whereas the poorly or nonphosphorylated remains in the basolateral membrane (Sakakibara et al., 1997). Exogenous occludin expressed in fibroblasts recruits ZO-1 to the contacting membranes (Van Itallie and Anderson, 1997). This fact demonstrates that the ZO-1 interaction with occludin, as well as the interaction between occludins of neighboring cells, is crucial for the localization of TJ proteins (Figure 8.4). Before contact formation, ZO-1, E-cadherin, and β-catenin are found to colocalize in intracellular compartments; occludin remains largely excluded from these complexes (Rajasekaran et al., 1996; Sakakibara et al., 1997; Grindstaff et al., 1998; Ando-Akatsuka et al., 1999). During TJ formation, ZO-1, E-cadherin, and Sec6/8, a component of the basolateral delivery machinery, are expressed coordinately on initial contacts (Sakakibara et al., 1997; Grindstaff et al., 1998). This marks the site where the protein complex named “Exocyst” is assembled. Basolateral membrane components will be inserted in the exocyst and then recruited to its final destination (Guo et al., 1997; Hsu et al., 1999). Furthermore, it has been demonstrated that the N-terminal half of ZO-1 associates with TJs, whereas the C-terminal half binds actin (Itoh et al., 1997; Fanning et al., 1998). Taken together, these observations suggest that ZO-1 is crucial for clustering TJ proteins, segregating them from other lateral junctions, and stabilizing them through anchorage to cortical actin cytoskeleton.

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OCCLUDIN

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ZO-1

FIGURE 8.4 COS-1 cells transfected with occludin-HA. Occludin (left) detected by immunofluorescence with antibodies against the HA tag. Endogenous ZO-1 (right) detected in the same field with a specific antibody. The filled arrow indicates contacting border between two cells that express transfected occludin. The empty arrow indicates a border between a transfected and a nontransfected cell.

8.8 POLARITY OF ADHERENS JUNCTION PROTEINS In polarized MDCK cells, E-cadherin is localized to the lateral membrane (Behrens et al., 1985). Newly synthesized E-cadherin is preferentially sorted to the basal-lateral membrane (Le Bivic et al., 1990), after the formation of a strong binding with the β-catenin (Hinck et al., 1994). The cytoplasmic domain of E-cadherin contains two putative basal-lateral, tyrosine-based sorting motifs. Mutation of these sites does not affect the fidelity of newly synthesized E-cadherin delivery to the basal-lateral membrane of MDCK cells (Chen et al., 1999). Nevertheless, these sorting signals target efficiently to the basolateral membrane a chimeric protein with the extracellular domain of an apical membrane protein (GP2), and the intracellular and transmembrane domains of E-cadherin. Thus, β-catenin binding to the whole cytoplasmic domain of E-cadherin correlates with efficient and targeted delivery of E-cadherin to the lateral plasma membrane. In this capacity, the authors suggest that β-catenin acts as a chauffeur, to facilitate transport of E-cadherin out of the endoplasmic reticulum and the plasma membrane. Another catenin molecule associated with adherens junctions is α-catenin. It seems that α-catenin joins the E-cadherin/β-catenin complex once it is inserted to the membrane (Hinck et al., 1994). In this respect, αcatenin is the connector of E-cadherin/β-catenin complex to the cytoskeleton. Hence, α-catenin recruitment to adherent junctions might be through a different route.

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8.9 DYNAMICS OF CELL MEMBRANE POLARITY Cell polarity does not constitute a static feature, but rather is a highly dynamic arrangement of molecules in the plasma membrane that remains sensitive to the nature of the ECM, contacts with neighboring cells, growth and differentiating factors, hormones, pharmacological agents, and even to cell cycle stages. Thus, MDCK cells express Na+,K+-ATPase and E-cadherin in the lateral membrane, provided these proteins are also expressed by the neighboring cell (see Figure 8.2). Similarly, Na+, K+-ATPase polarity in epithelia derived from the neuroepithelium (Rizzolo, 1999) and from honeybee receptors (Baumann and Takeyasu, 1993) seems to depend strictly on signals from the extracellular environment. Typically, this ion pump is expressed in basolateral membranes, but in the epithelium of the choroid plexus and the retinal pigment epithelium (RPE) is confined to the apical plasma membrane (Rizzolo, 1999). Cells from the RPE in situ, besides contacting each other, establish contacts with the retinal cells through their apical domains, where the submembranal cytoskeleton is located. In vitro, RPE cells (RPE-J cell line) express an endogenous cadherin and form adherens junctions and a tight monolayer, but Na+,K+-ATPase is localized to both apical and baso-lateral membranes (Marrs et al., 1995). Moreover, expression of E-cadherin in RPE-J cells results in restriction and accumulation of both Na+,K+-ATPase and the membrane cytoskeleton at the lateral membrane. Changes in the ECM revert the polarity of intercalated cells from collecting duct (see Section 8.6.1.2). There is now clear evidence of overlapping signals. Thus, the LDL and polyimmunoglobulin receptors are expressed on the basolateral milieu because of a dominant signal related to clathrin. However, once these receptors are exposed to the basolateral milieu and bind their respective ligands (LD, IgG, or IgM) they are readdressed and transcytosed to the apical domain (Casanova et al., 1990; Matter et al., 1994). Thy-1, a GPI-anchored protein addressed to the apical domain, is still delivered to this address upon removal of the GPI, implying again that it must have a second apical signal, or else that it is sent to this domain through a default mechanism (Powell et al., 1991).

8.10 RELATIONSHIP BETWEEN TIGHT JUNCTIONS AND ADHERENS JUNCTIONS 8.10.1 THE ESTABLISHMENT OF STRUCTURAL AND MOLECULAR ASYMMETRY AT THE CELL SURFACE REQUIRES AN EXTERNAL SIGNAL As mentioned above, extracellular Ca2+ triggers the process of epithelial polarization. It is also demonstrated that the extracellular domains of cadherins change conformation in response to calcium, engaging in homotypic interactions to specify cell–cell connections (Nose et al., 1990; Shapiro et al., 1995). This feature is thought to account fully for the requirement of calcium in intercellular adhesion. Nevertheless, recent findings of Vasioukhin et al. (2000) clearly demonstrate that Ca2+ is

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required to stimulate filopodia, which penetrate and embed into neighboring cells (Figure 8.5). Thus, extracellular calcium is required for initiation and maintenance of epithelial polarity both for homotypic cadherin association and for “zipper formation.” Nevertheless, it is becoming apparent that the driving force for adherens junction formation in epithelial cells is a direct actin polymerization and organization (Vasioukhin et al., 2000). It can be speculated that the initial calcium-dependent cell adhesions are likely to occur between the few E-cadherin molecules that are randomly distributed at the filopodia membrane, hence probably forming a weak adhesion (Imamura et al., 1999). Once the first cadherin–catenin complexes are immobilized within the cell–cell contact by associating to actin cytoskeleton, more freely diffusing cadherin is progressively recruited and clustered to form a stable and strong adherens junction structure (Adams and Nelson, 1998).

8.10.2 THE ROLE OF SEC6/8 COMPLEX OF CELL–CELL CONTACT

IN THE INITIAL

STAGE

Sec6/8 complex (or exocyst complex) is initially localized at the cell–cell contact tips of epithelial MDCK cells (Grindstaff et al., 1998). It is not yet clear which is the mechanism that recruits the exocyst complex to the primordial cell–cell contacting membrane. Early studies of polarity and TJ biogenesis demonstrated that a cascade of signal transduction pathways is involved in this initial step of Ca2+induced polarity and TJ assembly (Gonzalez-Mariscal et al., 1985; 1990; Balda et al., 1991; Contreras et al., 1992a, b). Therefore, one can speculate that the mechanism includes an interaction with a signal-transduction messenger. Nevertheless, other mechanisms such as post-translational modifications or an activation of a membrane receptor for the complex are not excluded. If the mechanism involves interaction of signal-transduction messenger with sec6/8 complex, it is anticipated that when MDCK cells expressing E-cadherin are mixed with Ma104 cells expressing N-cadherin, the monolayer would polarize and TJs will assemble in spite of the lack of adherent junctions at the heterologous border. The authors’ results, presented in Figure 8.6, show that although there is no E-cadherin and Na+,K+-ATPase in the mixed border (see Figures 8.3 and Contreras et al., 1995b), all other components of TJ and adherent junctions are well localized. This implies that a soluble message induced by homologous contacting membrane is sufficient for recruitment of junctional proteins to all lateral membrane. It would therefore be interesting to study the localization of sec6/8 in the heterologous border. In this respect, α-catenin in L-cells wt was sufficient to stimulate puncta formation in contacts with an L cell that lacks α-catenin. In this case, cytoskeleton anchorage on one side of an adhesion junction is essential for puncta formation and stabilization. Yonemura et al. (1995) have shown that at the initial stage of cell–cell contact, E-cadherin and ZO-1 appear to be simultaneously recruited to the primordial form of spotlike junctions at the tips of cellular processes. Moreover, Grindstaff et al. (1998) reveal that sec6/8 complex is recruited to the same domain and with similar timing. Therefore, an interesting question would be whether vesicles that carry the sec6/8 complex to sites of initial cell–cell contacts include E-cadherin and ZO-1.

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A

B

C

D

TJs P

E

AJs

AP

BL

FIGURE 8.5 Model of cell–cell contacts and cell polarity generation. (A) Nonpolarized precursors. The establishment of cell–substrate (B) or cell–cell contacts defines initial polarization.

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8.10.3 CELL JUNCTION PROTEINS ARE RECRUITED LATERAL MEMBRANE

185 TO INITIAL

Rajasekaran et al. (1996) reported a probable association of β-catenin with ZO-1 at the intracellular pool previous to Ca2+-dependent cell adhesion induction, and a colocalization of these two molecules at the tips of cell–cell contacts or “zipper.” On the other hand, Hinck et al. (1994) reported that shortly after E-cadherin is synthesized, it forms a strong interaction with β-catenin, which is essential for the incorporation of E-cadherin to the lateral plasma membrane (Wittchen et al., 1999). Balda et al. (1993) have found colocalization of ZO-1, ZO-2, and ZO-3 proteins in low Ca2+ conditions, suggesting that at least these three TJ plaque proteins are packed together in the same compartment prior to TJ assembly. Nevertheless, Sakakibara et al. (1997) found that occludin, the transmembrane protein of TJs, does not colocalize with ZO-1 in the cytosolic pool, prior to induction of cell–cell adhesion. Given that the initial lateral membrane is very limited in space, and that sec6/8 marks the site for docking and fusion of basolateral vesicles in this domain, it is difficult to assess whether junction molecules colocalize to the same domain because they were all packed together in the same cytosolic pool, for example, in basolateral targeting vesicles containing sec6/8 complex, or whether they get there from separate pools and happen to meet at a very restricted site. What is fairly clear is that as cellular polarization proceeded, ZO-1 is sorted to form spotlike junctions and occludin is gradually accumulated at the ZO-1-positive spots to form beltlike TJ. In a complementary manner, E-cadherin is sorted out from the ZO-1-positive spotlike junctions to form beltlike adherens junctions (AJ). The molecular mechanism of TJ/AJ formation during epithelial cellular polarization is not so obvious; however, it appears that ZO-1 plays a fundamental role in this mechanism thanks to its capability to interact with junction proteins via PDZ domains.

FIGURE 8.5 (continued) Model of cell–cell contacts and cell polarity generation. (A) Nonpolarized precursors. The establishment of cell–substrate (B) or cell–cell contacts defines initial polarization. A small quantity of E-cadherin (ovals) is in the membrane of noncontacting cells, and a pool of intracellular cadherin is associated with ZO-1 (small empty circle) and with members of the secretory machinery (sec6/8, triangle) in intracellular vesicles (big empty circle). Occludin (striped circle) is located in a different vesicle. Membrane E-cadherin forms occasionally weak contacts triggering a signal that is conveyed to the intracellular delivery machinery (C). Ca2+ increments the assembly of actin cytoskeleton filopodia from monomeric actin (black dots), that penetrate deep into neighboring cells and promotes the formation of strong contacts (D). The delivery machinery, containing ZO-1, Sec6/8, and E-cadherin (D), is incorporated in these puncta. Vesicles with occludin are incorporated later to the basolateral membrane. Phosphorylation is necessary to anchor occludin to the TJs (P in part E). TJs and AJs are segregated (E). The exocyst constitutes the site for delivery of basolateral membrane components and is identified by the Sec6/8 protein near the TJs. Maintaining of the polarized phenotype is carried out by selective targeting of apical (AP, gray circle) and basolateral (BL, dotted circle) membrane components and by retention in the well-formed microdomains (TJs and AJs).

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A

E

B

F

C

G

D

H

FIGURE 8.6 Fluorescence image of monolayers formed as indicated in Figure 8.3. (A and B) Occludin (B, C) is present in all the borders. (C and D) E-cadherin, stained with DECMA-1 antibody (green) is only observed in MDCK–MDCK borders (filled arrow) but not in MDCK borders in contact with Ma104 cells stained with CMTMR (E, F, G, H), as indicated by the empty arrows. Images obtained by confocal laser scanning microscopy from the X–Y (A, C, E, and G) and X–Z (B, D, F, and H) planes.

8.10.4 MEMBRANE SKELETON STABILIZES MEMBRANE PROTEINS A fodrin-based membrane skeleton is assembled in parallel to the organization of the AJ at the contacting membranes of adjacent cells. The role of this submembranal compartment is to reinforce the polarity already established by cell adhesion and actin cytoskeleton. Therefore, membrane proteins with ankyrin-binding domain, such as the Na,K-ATPase, anchor to the membrane skeleton and stabilize there. Nevertheless, fodrin/ankyrin membrane skeleton by itself is an asymmetric structure, and how this domain becomes polarized is not clear. Jefford and Dubreuil (2000), studying the expression of the L1 family cell adhesion molecule neuroglian in Drosophila, found that neuroglian adhesion generates a spatial cue for polarized assembly of ankyrin and the spectrin cytoskeleton. Again, an external stimulus of a membrane receptor apparently induces the polarization of a specific intracellular

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structure. It is also reminiscent of the case of PECAM-1 in endothelial cells (Sun et al., 2000), where a similar mechanism is apparent.

8.10.5 TJ IS A TARGETING PATCH Once the sec6/8 complex is recruited to the cell–cell contact, it restricts the subsequent localization of specific molecules synthesized de novo to the immediate vicinity of the complex. It seems that the sec6/8 complex in yeast, MDCK cells, and neurons is the membrane site for exocytosis (Hsu et al., 1999). Moreover, in polarized monolayer of MDCK cells it specifically constitutes the basolateral-targeting site (Grindstaff et al., 1998), which localizes at the TJs. Thus, Lapierre et al. (1999b) found that VAP-33, a protein implicated in vesicle docking/fusion, interacts with occludin in vitro and colocalizes with occludin at the TJs. Recent works of Bilder et al. (2000; Bilder and Perrimon, 2000) provide a related example from Drosophila epithelia. Mutations in Drosophila scribble (scrib), which encodes a multi-PDZ and leucine-rich-repeat protein, cause aberrant cell shapes and loss of the monolayer organization of embryonic epithelia. Scrib is localized to the epithelial septate junction, the analogue of the vertebrate TJ, at the boundary of the apical and basolateral cell surfaces. Loss of scrib function results in the misdistribution of apical proteins and AJs to the cell surface, but basolateral protein localization remains intact, thus suggesting that the lateral domain of epithelia, particularly the septate junction, functions in restricting apical membrane identity and correctly placing AJs. Moreover, recruitment of Lgl (which promotes fusion of vesicles with target membranes) into the proximity of membrane t-SNAREs requires proper localization of Scrib and Dlg (septate junction molecule homologous to ZO-1), thus potentially linking the transmembrane proteins that establish polarity to the protein-targeting system that preserves it. Since VACs and leucine amino peptidase were also seen to fuse at the vicinity of TJs (Louvard, 1980; Vega-Salas et al., 1987), a putative “apical exocytosis site” could exist adjacent to TJs, probably from its apical side. Recently, Nusrat et al. (2000) demonstrated that TJs are in fact microdomains that include lipid rafts and caveolin-1. Hence, TJs seem to restrict polarity not just by their fence function but also by including specific targeting patches in them.

8.10.6 SELECTIVE TARGETING AND STABILIZATION MAINTAIN EPITHELIAL CONTACTS AND POLARITY In mature epithelial monolayer, TJs and AJs function as specialized membrane microdomains. Newly synthesized components of these microdomains are delivered to the membrane through selective targeting mechanisms, and stabilized at their functional site through protein–protein clustering and association with cytoskeleton. Nevertheless, the relationship between TJs and AJs may be more complex than it appears today. Thus, Troxell et al. (2000) found typical junctional strands and some TJ markers, such as ZO-1 and occludin, in cells whose expression of endogenous E-cadherin had been severely reduced. In this respect, it is worth remembering that the TJ is formed by a dozen molecular species arranged in a complex manner and has several functions and that the presence of a marker or the detection of one of

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the TJ characteristics (measuring transepithelial electrical resistance, or TER, permeability, fluorescence lipid probe mobility) does not ensure that the other would be necessarily present.

8.11 CONCLUDING REMARKS The intimate relationship between cell–cell contacts and polarity has been observed for many years. It is known that E-cadherin expression is a prerequisite for TJ formation and polarity establishment, but a growing body of evidences, obtained in the last years, demonstrates that TJ components participate in AJ and polarity biogenesis. The initial development of cell–cell contacts and polarity are carried out by a complex structure, the filopodia, constituted by components of TJs, AJs, actin cytoskeleton, and the machinery of basolateral delivery. In mature epithelial cells, TJs are not just a cell–cell contact, but also a membrane functional microdomain that acts as a barrier or fence, as well as in the insertion site of membrane components to the basolateral domain. Protein phosphorylation and clustering emerge as key processes for TJs and polarity generation and maintenance.

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9

Optical Methods for the Study of Tight Junctions Olga N. Kovbasnjuk and Kenneth R. Spring

CONTENTS 9.1 9.2

Overview ......................................................................................................199 Previous Applications of Light Microscopy in the Study of Tight Junctions.......................................................................................................200 9.2.1 Immunofluorescence Microscopy ....................................................200 9.2.2 Cytoskeleton–Tight Junction Interactions .......................................202 9.2.3 Solute Permeability of the Tight Junction.......................................203 9.2.4 Water Flow across the Tight Junction .............................................205 9.3 Prospects for Light Microscopic Methods in the Study of the Tight Junction ........................................................................................................208 9.3.1 Protein–Protein Interactions Using FRET.......................................208 9.3.2 Tight Junction Dynamics Studied with GFP-Labeled Proteins ......209 9.3.3 Lipid Distribution.............................................................................209 References..............................................................................................................210

9.1 OVERVIEW Light microscopy provides many of the tools used to understand the three classical functions of the TJ — bridge, gate, and fence — described by Diamond (1977) in his Bowditch lecture. The multiple proteins that constitute the bridge connecting neighboring cells have largely been identified and localized by immunofluorescence microscopy. Because functional information cannot be directly ascertained from the immunofluorescence images, the role of the TJ as a transepithelial permeability barrier (the gate) has traditionally been determined from the transepithelial electrical resistance or from the flux of a marker substance such as a fluorescent dye or radiolabeled tracer. These methods, described in detail in other chapters, report the average properties of the entire epithelium and not of an individual TJ. In many cases, it would be of benefit to know the permeability properties of a defined region of the TJ free of concerns about the influence of the transcellular pathway. To this end, light microscopic methods for determining the water and solute permeabilities of individual TJs have recently been developed. Finally, the role of the TJ as a fence, a barrier to the intermixing of the lipids and proteins of the apical and basolateral 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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membranes, was clearly demonstrated by Dragsten et al. (1981) from microscope images of fluorescently labeled lipid and lectin probes. Epithelia, in general, and TJs, in particular, pose special technical problems for the light microscopist. Light scattering in the relatively tall, columnar cells of many epithelia limits the visualization of intracellular organelles or subcellular structures, such as the TJs. Optical sectioning together with digital deblurring or confocal fluorescence microscopy can provide some information about the three-dimensional structure and volume of epithelial cells, but the number of z-sections required may be very great and may hamper subsequent analyses. Study of native epithelial sheets, such as gallbladder and intestine, or cultured epithelial cells grown on permeable supports, necessitates the use of dual-sided perfusion chambers to control the composition of both bathing solutions. Because of the presence of a layer of solution between the cover glass and the epithelium, the optical microscopic properties of such chambers are less than ideal, further compromising resolution and image quality.

9.2 PREVIOUS APPLICATIONS OF LIGHT MICROSCOPY IN THE STUDY OF TIGHT JUNCTIONS 9.2.1 IMMUNOFLUORESCENCE MICROSCOPY One of the most important applications of fluorescence microscopy is immunofluorescence. It is based on an antigen–antibody reaction in which the antibody is labeled with a fluorophore. The direct immunofluorescence method utilizes a specific fluorescently labeled antibody that conjugates with the appropriate antigen in the specimen; visualization of the fluorescence emission enables determination of the distribution of the antigen in the specimen. Indirect immunofluorescence employs an unlabeled antibody that combines with the related antigen. A fluorophore attached to an anti-antibody is then introduced that will interact with the antibody–antigen complex enabling its detection by fluorescence microscopy. The specificity of antibodies and the sensitivity of fluorescence microscopy make immunofluorescence a powerful method for the study of protein and lipid distribution in cell culture and tissues. Determination of the colocalization of antibodies labeled with spectrally distinct fluorophores also permits the identification of possible molecular associations. Over the past 15 years, numerous associated and transmembrane proteins of TJs have been identified, and immunofluorescence microscopy has become an important tool in the localization of these proteins to the junctional complex and in identifying possible interactions. The first tight junctional protein found, termed ZO-1 (Stevenson et al., 1986), was detected by a monoclonal antiserum specific for an intercellular junction antibody. As assayed by immunofluorescence staining of cryostat sections of whole tissue, the ZO-1 antibody localized to the cytoplasmic side of the junctional complex of a number of epithelia, including colon, kidney, liver, and testis, Madin–Darby canine kidney (MDCK) cells, as well as to arterial endothelium. A second TJ-associated protein, ZO-2, was subsequently coimmunoprecipitated with ZO-1 (Gumbiner et al., 1991; Jesaitis and Goodenough, 1994). Another 130-kDa protein that coimmunoprecipitated with the TJ protein ZO-1 was purified from

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MDCK cells (Haskins et al., 1998). Immunofluorescent costaining showed that this new protein, named ZO-3, was localized to sites of cell–cell interaction, identical to the distribution of ZO-1 in MDCK cells, but with additional staining of the cytoplasm (Haskins et al., 1998). A component of cell–cell contacts, symplekin, associated with the cytoplasmic face of the TJ of polarized epithelial cells and of Sertoli cells of testis but absent from the junctions of vascular endothelia, was detected with a fluorescent monoclonal antibody (Keon et al., 1996). However, as was shown by immunofluorescence, this protein could also be detected in a wide range of cell types that do not form TJs or were even completely devoid of any stable cell contacts. Subsequent analyses revealed that the protein occurred in diverse cells in the nucleoplasm, and only in those cells forming TJs was it recruited, partially but specifically, to the zonula occludens. Symplekin represents a group of dual-residence proteins that occur and probably function in the nucleus as well as in the plaques formed as part of TJs, adherens junctions, or desmosomes (Keon et al., 1996). Occludin, an integral 65-kD membrane protein, was localized by immunofluorescence microscopy to the TJ of both epithelial and endothelial cells (Furuse et al., 1993). At the electron microscopic level, the labels were detected directly over the points of membrane contact in TJs. Later, it was shown that phosphorylation of occludin played an important role in its location and may be a key step in TJ assembly (Sakakibara et al., 1997; Wong, 1997). Fluorescent antibodies that recognized only the higher-molecular-weight phosphorylated isoform selectively stained the TJ of intestinal epithelial cells, whereas other antioccludin antibodies, which recognized the nonphosphorylated or less phosphorylated protein were detected on the basolateral membrane. Recently, the claudin family of proteins (Furuse et al., 1998a) was discovered; the family now includes 16 isoforms. All of them are small, about 20 to 22 kDa, integral membrane proteins with four transmembrane domains that are exclusively concentrated at TJs. Claudins function as a major structural component of TJ strands, whereas occludin serves as an accessory protein (Furuse et al., 1998b). Immunofluorescence confocal microscopy revealed colocalization of claudins 1 to 8 with occludin at the TJ in various tissues (Morita et al., 1996). Thus, a fluorescent antibody against claudin-3 revealed its presence in liver bile canaliculi, while claudins 4 and 8 were detected in distal tubules in kidney and in lesser amount in proximal tubules. Claudin-11 was shown to be expressed in the brain and testis (Morita et al., 1999). Imunofluorescence microscopy with anti-claudin-11 polyclonal antibody and an antineurofilament antibody revealed that, in the brain, claudin-11 formed interlamellar strands that spiral around neurofilament-positive axons. In testis, the well-developed tight junctional strands of Sertoli cells were specifically labeled with anticlaudin-11 antibody. Another important member of the family, paracellin-1 (claudin16), was found predominantly in the thick ascending loop of Henle, a tubule segment where the divalent cations magnesium and calcium are reabsorbed paracellularly from the lumen (Simon et al., 1999). Immunofluorescence microscopy not only has the potential to colocalize structural elements of the TJ, but also to detect some protein interactions and observe their influence on TJ assembly and function. Thus, immunofluorescence microscopy

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has shown that lack of afadin, an actin filament-binding protein that binds to nectin during embryogenesis, completely disrupts the distribution of ZO-1 in mouse ectoderm (Ikeda et al., 1999). In the calcium switch model of MDCK cells, immunofluorescence microscopy confirmed a close association of β-catenin (but not E-cadherin) and ZO-1 in the first 2 h after the calcium switch (Rajasekaran et al., 1996). At low calcium concentrations, ZO-1 was distributed intracellularly and colocalized with E-cadherin in granular clusters. In fully polarized monolayers of MDCK cells, the ZO-1–catenin complex was not detected, suggesting that catenin participated only in the mobilization of ZO-1 from the cytosol to the cell surface early in the development of the tight junction. Immunofluorescence microscopy revealed that RhoA and Rac1, small GTP-ases, regulate both the gate and fence functions of the TJ, and play a role in the spatial organization of TJ proteins (Wittchen et al., 1999). Binding interactions of ZO-2, ZO-3, and occludin were revealed by immunofluorescence, which showed that all three proteins colocalized with actin aggregates at the cell border in cytochalasin D–treated MDCK cells, and that ZO-2 bound directly to both ZO-1 and occludin (Jou et al., 1998). However, any apparent molecular association detected by immunofluorescence is limited by the 0.2-µm spatial resolution of light microscopy. The expected distances between colocalized proteins or lipids may be far less than the spatial resolution of the microscope as this is gigantic at the molecular scale. A high-resolution light microscopy method, fluorescence resonance energy transfer (FRET), is a useful tool for investigating molecular associations at a length scale of <10 nm, comparable to the size of the larger molecules themselves (Kenworthy and Edidin, 1997). FRET effectively increases the resolution of light microscopy to the molecular level and will be considered in more detail below.

9.2.2 CYTOSKELETON–TIGHT JUNCTION INTERACTIONS It has been proposed that the association of intestinal epithelial tight junctional complexes with cytoskeletal components, especially actin filaments, could mediate changes in the structure and permeability of the junctions (Madara et al., 1988). Manipulations of the actin cytoskeleton, such as polymerization and depolymerization of F-actin, have been widely used in investigations on the role of actin in a variety of epithelial functions. A useful tool for the study of actin in the cytoskeleton is phalloidin, a bicyclic toxic peptide isolated from the Amanita phalloides mushroom. Phalloidin specifically binds to polymerized actin filaments and shifts the monomer/polymer equilibrium toward the polymer, lowering the critical concentration for polymerization. Phalloidin is available with various attached fluorophores; most notable are new photostable fluorescent dyes, such as Oregon green 488 and 514, that result in brighter, longer-lasting fluorescence. Another useful toxin, cytochalasin D, disrupts F-actin shifting the equilibrium toward the monomeric form (Cooper, 1987). The thin optical sectioning through fluorescently labeled cells made possible by confocal microscopy has enabled visualization of the distribution of actin filaments stained with fluorescently labeled phalloidin. A perijunctional actin–myosin ring is a characteristic feature of epithelial cells (Nusrat et al., 1995). Quantitative analysis

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of filamentous actin from confocal fluorescence images of MDCK cells showed that an increased amount of F-actin in the perijunctional ring correlated with a higher sodium permeability of the TJ (Kovbasnjuk et al., 1998a). Other studies (Hecht et al., 1996; Gandhi et al., 1997) have shown that the paracellular permeability of MDCK cell epithelium is increased after stiffening of the cytoskeleton by expression of the catalytic domain of myosin light-chain kinase. Although these studies support early observations that the permeability of TJs can be regulated, they do not prove that alterations in the cytoskeleton are the mechanism by which the permeability change occurs. More-sophisticated studies, examining the dynamics of the cytoskeleton as elegantly demonstrated by Waterman-Storer and Salmon (1999) in their studies of tubulin, will be required to understand the relationship between structural changes in the perijunctional actin–myosin ring and the permeability properties of the adjacent TJs.

9.2.3 SOLUTE PERMEABILITY

OF THE

TIGHT JUNCTION

The composition of fluids on opposite sides of epithelial cell layers is often different, and the epithelium serves as a barrier to movement of fluid, electrolytes, nutrients, pathogens, and cells. Cellular membranes provide most of the barrier while the TJs seal the paracellular pathway. Understanding of the barrier function of the TJ arose, in part, from a comparison of the electrical resistance of “tight” epithelia, such as urinary bladder, to that of “leaky” epithelial such as the renal proximal tubule. The TJs of some epithelia were shown to be a selectively permeable, regulated gate to the paracellular movement of solutes. Delineation of the permeability properties of the epithelial TJs by electrical resistance or tracer methods is complicated by the effects of unstirred layers, concerns about monolayer integrity, and the effects of the narrow, tortuous lateral intercellular spaces (LIS) adjoining the TJs. Recent application of fluorescence microscopy to the study of the paracellular pathway has enabled direct, noninvasive determination of the permeability to ions and fluorescent solutes of defined regions of the junctional complex of the living cells (reviewed in Spring, 1998). Fluorescence microscopy was used to determine the flux of fluorescent tracers into salivary gland acini (Segawa, 1994) and the kinetics of sodium movement across the TJs of MDCK cells (Chatton and Spring, 1995; Kovbasnjuk et al., 1995). A review of the approach used in the authors’ laboratory illustrates the principle of the use of fluorescent indicator dyes to determine directly the ionic permeability of the TJ as well as to estimate the transcellular fluxes. The method has also been employed to determine the influence of several agents on the sodium permeability of the TJ (Kovbasnjuk et al., 1998a). As shown in Figure 9.1, measurement of the sodium fluxes requires, first, filling the LIS with a solution containing a sodiumsensitive fluorescent dye (Figure 9.1A), then using ratio imaging to measure the rate of change of LIS sodium concentration in response to changes in the composition of the perfusion solution (Figure 9.1B). When the perfusion medium with a high sodium concentration is rapidly switched to one in which most of the sodium is replaced by lithium (Figure 9.1C), the fluorescence emission intensity ratio reflects the decrease in the concentration of sodium in the LIS as a function of time because

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FIGURE 9.1 Steps involved in determination of the TJ sodium permeability by fluorescence microscopy. First, the indicator dye (SBFO or SBFO dextran) is microinjected into an adjacent dome or between the epithelium and the glass coverslip and allowed to diffuse into nearby LIS (panel A); second, the fluorescence from an undisturbed, dye-filled region of LIS is measured to determine the LIS sodium concentration (panel B); third, the sodium concentration of the perfusate is suddenly reduced from 142 to 24 mM with the sodium replaced by lithium (panel C); fourth, the sodium efflux from the LIS is measured from the rate of decrease of the sodium concentration indicated by the fluorescence (panel D).

the dye is insensitive to lithium. The rate of decline in LIS sodium concentration is a measure of the relative sodium permeability of the TJ, enabling comparison with measurements in the presence of modulators of TJ permeability (Figure 9.1D). Determination of the absolute magnitude of the sodium flux across the paracellular and transcellular pathways requires calibration of the sodium concentration within the LIS as was done previously by Chatton and Spring (1995). As shown in Figure 9.2, similar experiments performed when the perfusate sodium concentration is returned to control levels in the absence or presence of ouabain to inhibit the Na,K-ATPase allow estimation of the transcellular sodium flux. Kinetic analysis permitted calculation of the sodium back-flux from the LIS across the TJ into the apical bath as well as the sodium influx into the LIS across both the TJ and cellular pathways. These studies showed that about 43% of the sodium content of an MDCK cell is transported into the LIS every minute and that about 20% of that leaks back into the apical bath across the TJ driven by the elevated sodium chloride concentration within the LIS (Kovbasnjuk et al., 1995). Measurements of the chloride fluxes across MDCK cell TJs by a similar approach were in good agreement with the sodium results (Xia et al., 1995; Kovbasnjuk et al., 1995). A similar experimental approach based on fluorescence ratio imaging of LIS with other indicator dyes could be used to measure the paracellular fluxes in cultured epithelia of solutes such as potassium, magnesium, protons, as well as fluorescent anions. Although the permeability of the TJ to the solute of interest can be readily determined by this method, ascertainment of the magnitude of the transcellular flux

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FIGURE 9.2 Steps involved in determination of the transcellular and paracellular sodium fluxes by fluorescence microscopy. The preparation has been loaded with dye as described in Figure 9.1 and the perfusate sodium concentration has previously been reduced to 24 mM so that a steady-state sodium concentration has been achieved in both cell and LIS. Under this condition, the sodium pump is presumably inactive because of the low intracellular sodium concentration. When the perfusate sodium concentration is suddenly increased to control value (142 mM) as shown in panel A, sodium should enter both the cells and LIS as shown in panel B. The rate of increase in LIS sodium concentration is measured from the fluorescence of the indicator dye trapped in the LIS (panel B). When ouabain is added to inhibit the Na,KATPase, the efflux of sodium to the LIS from the cells is assumed to be completely blocked (panel C). Repetition of the sequence in panels A and B (i.e., the increase of perfusate sodium to control levels and measurement of the fluorescence of the sodium indicator dye in the LIS) then gives an estimate of the sodium influx into the LIS across the TJ. From the difference between the rates in panels B and D, it is possible to estimate both the transcellular and paracellular sodium fluxes as well as any backflux from LIS to apical bathing solution that occurs under control conditions.

requires the use of a specific inhibitor of the transcellular pathway. Imaging of solute fluxes across the TJ enables both the determination of the permeability of an individual TJ to that solute as well as a direct analysis of the regulation of that permeability.

9.2.4 WATER FLOW

ACROSS THE

TIGHT JUNCTION

Fluid absorption is a hallmark of epithelia such as small intestine, gallbladder, and renal proximal tubules. It is well known that fluid absorption across intestinal and renal epithelia occurs in the absence of any external osmotic or hydrostatic gradients and is secondary to active solute transport. Although the mechanism of isosmotic transepithelial fluid transport was attributed to local osmotic gradients within the tissues, the site of coupling between active solute transport and water flow across epithelia was not defined until the study of Whitlock and Wheeler, who suggested the lateral intercellular spaces between the cells (reviewed in Spring, 1998). In principle, there are two possible routes for transepithelial water flow: transcellular — via apical and basolateral membrane water channels and/or diffusion across the

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respective lipid bilayer — or paracellular — across the TJ. The discovery of substantial ionic permeability of the TJ led investigators to propose significant water flux as well through the paracellular pathway. Some previous estimates of the magnitude of paracellular fluid flow have been based on the difference between total fluid transport by a tissue before and after attempting to block the water permeability of the TJ or the cell membrane. However, to date, specific inhibitors of TJ permeability or cellular water transport have not been found, as all of them affected both paracellular and transcellular fluid flow (reviewed in Spring, 1998). As described in previous chapters, the transepithelial fluxes of radioactive or fluorescent nonelectrolyte tracers have also been used to estimate the magnitude of the paracellular water flow, as well as the pore size of the TJ. In this approach, the apical-to-basal flux of solutes of different molecular weight was measured as a function of molecular size and transepithelial volume flow. These experiments demonstrated substantial fluxes of relatively large solutes across the TJ. The TJ pore size determined should permit water flow through the same pathway. Because the calculated magnitude of the water flow across the TJ deduced from the nonelectrolyte fluxes was dependent on additional parameters of undetermined magnitude, such as solvent-drag effects, local concentration of solutes in unstirred layers both on extracellular surfaces and within the LIS, it was virtually impossible to assess the validity of these estimates (reviewed in Spring, 1998). Another approach to calculate the paracellular fluid flow was based on the comparison of cell membrane and transepithelial water permeabilities. Light microscopy provides a direct method for the determination of cell membrane water permeability by measurement of the epithelial cell volume change in response to an osmotic gradient (Strange and Spring, 1986; Alvarez-Leefmans et al., 1995; Raat et al., 1996). The rate of cell swelling or shrinkage in hypotonic or hypertonic bathing solutions, together with estimated cell surface area, allows calculation of water permeability for both apical and basolateral cell membranes as was done in Necturus gallbladder (Persson and Spring, 1982), rabbit renal cortical collecting duct (Strange and Spring, 1987), rabbit renal proximal tubule (Carpi-Medina and Whittembury, 1988), and MDCK cells (Timbs and Spring, 1996). However, transepithelial water permeability is far more difficult to measure accurately as it is always underestimated because of unstirred layer effects (Diamond, 1979), with a resultant overestimate of the calculated paracellular water permeability (reviewed in Spring, 1998). Recently, a confocal fluorescence microscopy method was used to measure directly the fluid flux across the TJ of MDCK cells (Kovbasnjuk et al., 1998b). It is based on the assumption that fluid flow within the LIS can be visualized by introducing a high-molecular-weight fluorescent dye into the LIS and observing its concentration profile along the LIS from the TJ to the basement membrane. Figure 9.3 illustrates the principle of the method. Within the small LIS dimensions, the concentration profile of the dye depends on two factors: (1) fluid convection, tending to build a dye concentration gradient by sweeping (solid arrows in Figure 9.3), and (2) diffusion, tending to dissipate the flow-induced concentration gradient (dashed, two-headed arrow in Figure 9.3). The observed dye profile is also a function of LIS geometry, and of the diffusion coefficient of the dye. Thus, small molecules with high diffusion coefficients will result in flat profile along the LIS as

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FIGURE 9.3 Illustration of the principle of determination of any water flux across the TJ. The epithelium was grown on a permeable support that has been treated to reduce its pore size. The LIS is depicted as filled with dots representing a high-molecular-weight fluoresceindextran marker that was microinjected into an adjacent region. The solid arrows show the pathway for water movement across the cells in the absence of any transjunctional water flow. The dashed, two-headed arrow depicts the effects of diffusion tending to negate the marker concentration gradient created by the sweeping effects of the water movement. The thin dashed arrow with the question mark shows the pathway for putative transjunctional water flow. The lower portion shows a graph of fluorescence (F) as a function of distance along the LIS from the TJ for the case of no water flow across the TJ (solid line) or for transjunctional flow (dashed line).

diffusion outpaces convection, whereas the profile of a high-molecular-weight dye will be influenced more strongly by the local rate of water flow across the epithelium and will, therefore, be sensitive to the water permeability of the TJs. The flow velocity at each point within the LIS can then be determined from knowledge of the local marker concentration, the diffusion coefficient of the marker in the LIS, and the cross-sectional area of the region (an example profile is illustrated at the bottom of Figure 9.3). If significant water flows occur across the TJ (thin dashed arrow in Figure 9.3), the marker concentration would be expected to fall precipitously in the region of the LIS immediately adjacent to the junction (dashed curve in the flow profile at the bottom of Figure 9.3). The small dimensions of the LIS in this region would result in a relatively high flow velocity and powerful convection. These experiments showed that the magnitude of fluid flow across the TJ of MDCK cells grown on permeable supports was indistinguishable from zero (Kovbasnjuk et al., 1998b). Even when junctional permeability was greatly increased by the stimulation of protein kinase A, the fraction of transepithelial flow that was paracellular was not of significant magnitude. Similar measurements of fluid flow across

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the TJs of native tissues where transjunctional flows have been postulated, such as gallbladder and small intestine, have yet to be accomplished. Support for the conclusion that transjunctional water flow is not a significant component of transepithelial absorption also comes from experiments on renal proximal tubules from aquaporin-1 gene knockout mice. Transepithelial water permeability was decreased by ~80% in the knockout mice compared with control animals (Schnermann et al., 1998), about what would be expected if the only remaining permeability was associated with the water diffusion across the lipid bilayer of the cell membrane. These results were incompatible with the large paracellular fraction of fluid transport across the proximal tubule previously estimated by indirect methods, and offered strong support for a wholly transcellular route for water, as in the case of cultured MDCK epithelium.

9.3 PROSPECTS FOR LIGHT MICROSCOPIC METHODS IN THE STUDY OF THE TIGHT JUNCTION 9.3.1 PROTEIN–PROTEIN INTERACTIONS USING FRET The limiting resolution of the light microscope in the visible spectrum is about 0.2 µm (200 nm), although an approximately twofold improvement can be achieved by the use of digital deconvolution methods. From the point of view of a protein molecule, 100 nm is still a relatively large distance. The only optical microscopic technique suited for the study of molecular interactions is FRET. FRET occurs when two fluorophores are within 10 nm of each other and when the fluorescence emission spectrum of one fluorophore (the “donor”) overlaps the absorption spectrum of the other fluorophore (the “acceptor”). This situation is sometimes referred to as “sensitized fluorescence” as the excitation of the acceptor is accomplished by virtue of its coupling to the donor molecule rather than by direct absorption of a photon. Under these conditions, the energy normally associated with the fluorescence emission by the donor molecule is coupled, nonradiatively, to the acceptor molecule. This means that the fluorescence emission by the donor diminishes concomitantly with an increase in that of the acceptor. The extent of FRET is inversely proportional to the sixth power of the distance separating the fluorophores and is, therefore, an extremely sensitive measure of the intermolecular separation. FRET may be measured in a number of ways, most commonly by determination of the intensity of the fluorescence of both donor and acceptor (Kenworthy and Edidin, 1997; Gordon et al., 1998). FRET suffers from some significant technical limitations that must be taken into account before determinations of the degree of transfer can be made (Gordon et al., 1998). The most obvious problem arises from direct excitation of the acceptor molecule by light intended for the donor. The result is fluorescence emission by the acceptor when the donor is absent, a situation for which a correction can be made provided that the acceptor molecule concentration is known and the donor/acceptor ratio is well controlled (Gordon et al., 1998). A less obvious determinant of the extent of FRET is the degree of alignment of the emission dipole of donor molecule

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with the absorption dipole of the acceptor. If neither fluorophore is free to rotate because they are rigidly attached to their respective proteins, the extent of FRET may differ only because of the relative spatial orientation of the two fluorophores. FRET may be altered then not by changes in intermolecular distance between donor and acceptor but by the orientation of their dipoles.

9.3.2 TIGHT JUNCTION DYNAMICS STUDIED GFP-LABELED PROTEINS

WITH

To date, limited use has been made of green fluorescent protein (GFP) labels in the study of the protein components of the tight junctional complex (Furuse et al., 1998a; Lapierre et al., 1999). The development of new GFP variants has allowed the analysis of the spatial relationship of multiple proteins as well as provide the possibility of FRET as a method for understanding intermolecular interactions. Although it has been suggested by several authors (Claude, 1978; Cereijido et al., 1989; Hill and Shachar-Hill, 1997) that the tight junctions are dynamic, periodically altering their seal to admit large solutes, no direct experimental evidence has been obtained in support of these proposals (Timbs and Spring, 1996). Kovbasnjuk et al. (1995) and others (Steward, 1982; Whittembury et al., 1988) have reported that solutes with molecular weights as high as 1000 are capable of crossing the TJs of epithelia exhibiting electrically highly selective TJs. To date, the site of penetration of these solutes across the paracellular pathway has not been identified, and GFP-labeled junctional proteins may provide the necessary information. The use of multiple GFP labels on the putative protein components of the TJ may lead to an increased understanding of the structure and regulation of TJ components and permeability.

9.3.3 LIPID DISTRIBUTION In their seminal fluorescence microscopy study, Dragsten et al. (1981) reported that only some lipid probes added to the apical solution appeared on the basolateral membrane. They speculated that these probes were not solely confined to the outer leaflet of the cell membrane and could “flip-flop” into the inner leaflet. Further, they suggested that upon reaching the inner leaflet, the probes could bypass the TJ and diffuse into the basolateral membrane. At the time, nothing was known about the structure of the TJ and, in fact, it had been suggested that they were entirely lipidic in nature (Kachar and Reese, 1982). Now that it is known that the proteins forming the junctional complex span the entire membrane, the question of how some lipid probes can move across this barrier needs to be reexamined. As a greater understanding of the organization of the lipids of cell membranes is gained (Brown and London, 2000), the mechanism(s) that maintain the different compositions of the apical and basolateral membranes may also be amenable to investigation. Clearly, light microscopy has been a very useful tool for the study of the three functions of the TJ. The prospects for the future use of the method are bright, indeed, and assure a greater understanding of this complex and essential component of epithelia in days to come.

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Ikeda, W., Nakanishi, H., Miyoshi, J., Mandai, K., Ishizaki, H., Tanaka, M., Togawa, A., Takahashi, K., Nishioka, H., Yoshida, H., Mizoguchi, A., Nishikawa, S., and Takai, Y. 1999. Afadin: a key molecule essential for structural organization of cell–cell junctions of polarized epithelia during embryogenesis. J. Cell Biol., 146:1117–1132. Jesaitis, L. A. and Goodenough, D. A. 1994. Molecular characterization and tissue distribution of ZO-2, a tight junction protein homologous to ZO-1 and the Drosophila discs-large tumor suppressor protein. J. Cell Biol., 124:949–961. Jou, T-S., Schneeberger, E. E., and Nelson, W. J. 1998. Structural and functional regulation of tight junctions by RhoA and Rac1 small GTPases. J. Cell Biol., 142:101–115. Kachar, B. and Reese, T. S. 1982. Evidence for the lipidic nature of tight junction strands. Nature, 296:464–466. Kenworthy, A. K. and Edidin, M. 1997. Imaging fluorescence resonance energy transfer as probe of membrane organization and molecular association of GPI-anchored proteins. Methods Mol. Biol., 116:37–49. Keon, B. H., Schafer, S., Kuhn, C., Grund, C., and Franke, W. W. 1996. Symplekin, a novel type of tight junction plaque protein. J. Cell Biol., 134:1003–1018. Kovbasnjuk, O., Chatton, J.-Y., Friauf, W. S., and Spring, K. R. 1995. Determination of the Na permeability of the tight junctions of MDCK cells by fluorescence microscopy. J. Membr. Biol., 148:223–232. Kovbasnjuk, O. N., Smulowicz, U., and Spring, K. R. 1998a. Regulation of the MDCK cell tight junction. J. Membr. Biol., 161:93–104. Kovbasnjuk, O. N., Leader, J. P., Weinstein, A. M., and Spring, K. R. 1998b. Water does not flow across the tight junctions of MDCK epithelial cells. Proc. Natl. Acad. Sci. U.S.A., 95:6526–6530. LaPierre, L. A., Tuma, P. L., Navarre, J., Goldenring, J. R., and Anderson, J. M. 1999. VAP33 localizes to both an intracellular vesicle population and with occludin at the tight junction. J. Cell Sci., 112:3723–3732. Madara, J. L., Stafford, J., Barenberg, D., and Carlson, S. 1988. Functional coupling of tight coupling of tight junctions and microfilaments in T84 monolayers. Am. J. Physiol., 254:G416–G423. Morita, K., Furuse, M., Fujimoto, K., and Tsukita, S. 1996. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc. Natl. Acad. Sci. U.S.A., 96:511–516. Morita, K., Sasaki, H., Fujimoto, K., Furuse, M., and Tsukita, S. 1999. Claudin-11/OSPbased tight junctions of myelin sheaths in brain and Sertoli cells in testis. J. Cell Biol., 145:579–588. Nusrat, A., Giry, M., Turner, J. R., Colgan, S. P., Parkos, C. A., Carnes, D., Lemichez, E., Boquet, P., and Madara, J. L. 1995. Rho protein regulates tight junctions and perijunctional actin organization in polarized epithelia. Proc. Natl. Acad. Sci. U.S.A., 92:10629–10633. Persson, B. E. and Spring, K. R. 1982. Gallbladder epithelial cell hydraulic water permeability and volume regulation. J. Gen. Physiol., 79:481–505. Raat, N. J. H., De Smet, P., Van Driessche, W., Bindels, R. J. M., and Van Os, C. H. 1996. Measuring volume perturbations of proximal tubular cells in primary culture with three different techniques. Am. J. Physiol., 271:C235–C241. Rajasekaran, A. K., Hojo, M., Huima, T., and Rodriguez-Boulan, E. 1996. Catenins and zonula occludens-1 form a complex during early stages in the assembly of tight junctions. J. Cell Biol., 132:451–463. Sakakibara, A., Furuse, M., Saitou, M., Ando-Akatsuka, Y., and Tsukita, S. 1997. Possible involvement of phosphorylation of occludin in tight junction formation. J. Cell Biol., 137:1393–1401.

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Schnermann, J., Chou, C.-L., Ma, T., Traynor, T., Knepper, M. A., and Verkman, A. S. 1998. Defective proximal tubular fluid reabsorption in transgenic aquaporin-1 null mice. Proc. Natl. Acad. Sci. U.S.A., 95:9660–9664. Segawa, A. 1994. Tight junctional permeability in living cells: dynamic changes directly visualized by confocal laser microscopy. J. Electron Microsc., 43:290–298. Simon, D. B., Lu, Y., Choate, K. A., Velazquez, H., Al-Sabban, E., Praga, M., Casari, G., Bettinelli, A., Colussi, G., Rodriguez-Soriano, J., McCredie, D., Milford, D., Sanjad, S., and Lifton, R. P. 1999. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science, 285:103–106. Spring, K. R. 1998. Routes and mechanism of fluid transport by epithelia. Annu. Rev. Physiol., 60:105–119. Stevenson, B. R., Siliciano, J. D., Mooseker, M. S., and Goodenough, D. A. 1986. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J. Cell Biol., 103:755–766. Steward, M. C. 1982. Paracellular nonelectrolyte permeation during fluid transport across rabbit gallbladder epithelium. J. Physiol., 322:419–439. Strange, K. and Spring, K. R. 1986. Methods for imaging renal tubule cells. Kidney Int., 30:192–200. Strange, K. and Spring, K. R. 1987. Cell membrane water permeability of rabbit cortical collecting duct. J. Membr. Biol., 96:27–43. Timbs, M. M. and Spring, K. R. 1996. Hydraulic properties of MDCK cell epithelium. J. Membr. Biol., 153:1–11. Waterman-Storer, C. M. and Salmon, E. D. 1999. Fluorescent speckle microscopy of microtubules: how low can you go? FASEB J., 13 (Suppl 2):S225–S230. Whittembury, G., Malnic, G., Mello-Aries, M., and Amorena, C. 1988. Solvent drag of sucrose during absorption indicates paracellular water flow in the rat kidney proximal tubule. Eur. J. Physiol., 412:541–547. Wittchen, E. S., Haskins, J., and Stevenson, B. R. 1999. Protein interactions at the tight junction. J. Biol. Chem., 274:35179–35185. Wong, V. 1997. Phosphorylation of occludin correlates with occludin localization and function at the tight junction. Am. J. Physiol., 273:C1859–C1867. Xia, P., Persson, B.-E., and Spring, K. R. 1995. The chloride concentration in the lateral intercellular spaces of MDCK cell monolayers. J. Membr. Biol., 144:21–20.

10

Occludin and Claudins: Transmembrane Proteins of the Tight Junction Laura L. Mitic and Christina M. Van Itallie

CONTENTS 10.1 Introduction .................................................................................................213 10.2 Occludin ......................................................................................................214 10.2.1 Identification of Occludin..............................................................214 10.2.2 Sequence of Occludin....................................................................214 10.2.3 Freeze-Fracture Characteristics of Occludin.................................215 10.2.4 Distribution of Occludin................................................................215 10.3 Claudins ......................................................................................................216 10.3.1 Identification of Claudins ..............................................................216 10.3.2 Sequence Comparisons of Claudins..............................................216 10.3.3 Freeze-Fracture Characteristics of Claudins .................................218 10.3.4 Distribution of Claudins ................................................................218 10.4 Functional Analysis of Occludin and Claudins..........................................220 10.4.1 Transepithelial Electrical Resistance.............................................220 10.4.1.1 Occludin and Transepithelial Electrical Resistance......221 10.4.1.2 Claudin and Transepithelial Electrical Resistance........222 10.4.2 Occludin and Claudins Are Cell–Cell Adhesion Molecules ........222 10.5 Interactions of Occludin and Claudins with Other Proteins......................224 10.6 Occludin Phosphorylation as a Possible Determinant of Fibril Organization ................................................................................................225 10.7 Future Directions ........................................................................................227 Acknowledgments..................................................................................................227 References..............................................................................................................227

10.1 INTRODUCTION The unique structural element of the tight junction (TJ) is the network of fibrils encircling epithelial cells that can be visualized by freeze-fracture electron microscopic analysis (see Chapter 2). The biochemical makeup of these fibrils and their associated proteins defines the physiological characteristics of TJs; namely, the 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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barrier both to the paracellular movement of solutes between epithelial cells and to the movement of membrane proteins and lipids between the apical and basolateral domains of the plasma membrane. To explain the nature of paracellular barriers formed by TJs, fibril constituents must be able to influence size and charge selectivity of the barrier in a tissue-specific and regulated fashion. In addition, because TJs are characterized by points of cell–cell membrane contact in transmission electron microscopic images, fibril components must be able to interact adhesively with complementary components on adjacent cells. Finally, the fibril components must be appropriately localized and be capable of lateral polymerization. To date, two proteins of the TJ fibrils have been identified; occludin (Furuse et al., 1993) and claudins (Furuse et al., 1998a). The latter are a protein family with at least 20 different members. Although an additional TJ integral membrane protein family, JAMs (junctional adhesion molecule), has been described (Martin-Padura et al., 1998; Aurrand-Lions et al., 2000), its identity as a constituent of the freeze-fracture fibrils has not yet been demonstrated. Since the discovery of claudins and occludin, a considerable number of studies has documented their distributions, functions, and binding partners. These recent advances in understanding the molecular structure of the TJ support older physiological models implying that the paracellular barrier had different characteristics in different tissues. Selective expression of claudins and occludin are likely to underlie variable and discriminating solute permeation. The goal of this chapter is to review this information in the context of how it contributes to an understanding of the structural and functional characteristics of the paracellular barrier.

10.2 OCCLUDIN 10.2.1 IDENTIFICATION

OF

OCCLUDIN

In 1993, Tsukita and co-workers (Furuse et al., 1993) used an isolated junctionenriched fraction from chick liver as an antigen to generate monoclonal antibodies. Some of the resulting antibodies identified a 65 kDa protein in rats that by both immunofluorescent and immunoelectron microscopic analysis specifically localized to TJs. Analysis of the sequence of the corresponding cDNA suggested that the predicted 504 amino acid polypeptide contained four transmembrane domains with internal N and C termini. An overall similarity in hydrophobicity plots with the gap junction protein connexin was recognized in this initial study, although the authors noted that the extracellular loops of occludin were larger than those of connexin. These larger extracellular loops were difficult to reconcile with the observation that distances between apposed membranes were smaller at TJs than at gap junctions, but these authors suggested that this might be related to the unusual chemistry of the extracellular loops. Both loops lack charged residues and are very rich in tyrosine; more than half (55%) of the first loop is composed of tyrosine plus glycine residues.

10.2.2 SEQUENCE

OF

OCCLUDIN

Subsequent studies by this group demonstrated the close identity among mammalian occludins (around 90%) and a lower level of sequence identity (around 50%) between

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chick and mammalian occludin (Ando-Akatsuka et al., 1996). Interestingly, even when primary sequence in the first extracellular loop was not strictly conserved, the tyrosine- and glycine-rich amino acid composition was maintained. Occludin appears to be encoded by a single gene (Saitou et al., 1997); however, expression of a longer splice variant of occludin was recently reported (Muresan et al., 2000). This occludin isoform, named occludin 1B, includes a novel 56 amino acid amino terminal insertion. When the subcellular distribution of occludin 1B was analyzed by immunofluorescent microscopic analysis, it appeared to be identical to that of the more prevalent smaller occludin isoform. Immunoblot analysis revealed some differences in the relative levels of the two isoforms, but in general the tissue distributions were similar. However, one possible basis for generation of diversity in the physiological properties of paracellular barriers is variability in the expression ratios of the two occludin isoforms. The small size difference between occludin 1B and occludin and the minor contribution of this splice variant to the total occludin protein are likely explanations for why this form of occludin was not previously recognized.

10.2.3 FREEZE-FRACTURE CHARACTERISTICS

OF

OCCLUDIN

Immunoelectron microscopic analysis of freeze-fracture replicas of chicken liver (Furuse et al., 1996) and rat kidney (Saitou et al., 1997) demonstrated convincingly that occludin was a component of the TJ fibrils. In addition, overexpression of occludin in Sf9 cells by recombinant baculovirus infection resulted in the formation of multilamellar bodies in the insect cells that were characterized by fused apposing membranes (Furuse et al., 1996). Freeze-fracture images of the occludin-enriched structures were characterized by intramembrane particles of approximately 10 nm, which occasionally coalesced to form short strands. This 10 nm particle size is approximately that seen in freeze-fracture replicas of TJ fibrils from unfixed or formaldehyde-fixed tissues. Differences in membrane lipid composition in the insect cells were invoked as a possible explanation for why overexpression of occludin did not result in true strand formation in Sf9 cells (Furuse et al., 1996). In type II MDCK (Madin–Darby canine kidney) cells, overexpression of wild-type occludin resulted in slightly increased fibril number (Balda et al., 1996; McCarthy et al., 1996) and increased the amount of apparent side-to-side fibril aggregation, resulting in some thicker fibrils (McCarthy et al., 1996; Medina et al., 2000). However, there was no consistent correlation between fibril number and functional changes in TJ resistance (Balda et al., 1996).

10.2.4 DISTRIBUTION

OF

OCCLUDIN

Examination of the expression levels and distribution of occludin revealed a correlation between the immunofluorescent staining intensity of occludin and the number of TJ strands in a variety of epithelial cells (Saitou et al., 1997). For example, occludin staining was intense in kidney distal tubules, which have elaborate TJs, while staining was weak in kidney proximal tubules, which have only one or two fibrils in their rudimentary TJs. However, closer examination revealed occasional discrepancies between the presence of fibrils and occludin expression. Occludin was

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present in the TJs of Sertoli cells in mouse and rat, but absent from those in guinea pig and human (Moroi et al., 1998). In addition, occludin was easily detectable in brain endothelial cell TJs, but not in peripheral endothelial cell junctions where there were demonstrated freeze-fracture fibrils (Hirase et al., 1997). Based on the lack of occludin in human testis, Moroi and coauthors (1998) speculated the existence of an additional integral membrane protein of TJs, revealed as claudins by this group later the same year (Furuse et al., 1998a).

10.3 CLAUDINS 10.3.1 IDENTIFICATION

OF

CLAUDINS

Although as indicated above, there were suggestions that occludin was not the only fibril protein of TJs, the most convincing evidence came from experiments in which both alleles of the occludin gene were disrupted by homologous recombination in mouse embryonic stem (ES) cells (Saitou et al., 1998). These occludin-deficient cells were still capable of developing apparently normal TJs, with normal-appearing freeze-fracture fibrils, and of generating a paracellular diffusion barrier (Saitou et al., 1998). This finding led to a renewed effort by Tsukita and co-workers (Furuse et al., 1998a) to identify other integral membrane proteins of the TJ, using the same chick liver fraction used to identify occludin. These authors used cofractionation with occludin on sucrose step gradients as a means to identify a single 22 kDa band as a candidate integral membrane TJ protein. Peptide sequencing revealed that this band contained at least two related proteins, which were cloned and named claudin1 and claudin-2.

10.3.2 SEQUENCE COMPARISONS

OF

CLAUDINS

Through a combination of database searching and both cDNA and genomic cloning, the claudin family has now been expanded to at least 20 genes. All claudins are predicted to encode 20 to 27 kDa proteins with two extracellular loops and short Cterminal intracellular tails. Although the four transmembrane domains are very similar, the extracellular loops have both conserved and divergent regions, and the C-terminal tails are strikingly divergent. This divergence could reflect differences in binding or regulatory interactions. The highly conserved exception in the cytoplasmic domain is the YV motif at the carboxy terminus, a sequence reminiscent of that seen in PDZ-binding proteins, except for claudins 11 (OSP), 12, 13, and 16. Because the extracellular loops are likely to form the paracellular barrier, they are of particular interest. The first loop is approximately 53 amino acids in length, whereas the second is much shorter, only around 24 amino acids. Two particular features of claudin extracellular loops invite speculation. The first is the variability in distribution and number of charged amino acid residues in the loops that might be expected to be important in influencing the passage of differently charged ions in the paracellular space (Table 10.1). This variability gives rise to isoelectric points for the first loop that range from 4.17 for claudin-16/paracellin to 10.49 for claudin-14 (Table 10.2), although placement of the charged residues may

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TABLE 10.1 Comparison of the First Extracellular Loop of Claudins 1, 2, 5, 11, and 16 + – – + – + 1 PQWRIYSYAGDNIVTAQAMY––EGLWMSCVSQ–STGQIQCKVFDSLLNLSS––LQATR + + – + – – 2 PNWRTSSYVGASIVTAVGFS––KGLWMECATH–STGITQCDIYSTLLGLPA–DIQAAQ –+ + + + – – + 5 PMWQVTAFLDHNIVTAQTTW––KGLWMSCVVQ–STGHMQCKVYDSVLALST–EVQAAR – ++ –– + – + + – + 11 NDWVVTCGYTIPTCRKLDELGSKGLWADCVM––ATGLYHCKPLVDILILPGY–VQACR – –– – + + – – + –– – –+ + + 16 ATW–TDCWMVNADDSLEVSTKCRGLWWECVTNAFDGIRTCDEYDSILAEHPLKLVVTR * *** ** ** * * * ** * D,E = acidic amino acids. K,R,H = basic amino acids. * = conserved 4/5.

TABLE 10.2 Comparison of Claudin Extracellular Loop pKIs

m. Claudin-1 m. Claudin-2 m. Claudin-3 m. Claudin-4 m. Claudin-5 m. Claudin-6 m. Claudin-7 m. Claudin-8 m. Claudin-9 m. Claudin-10 m. Claudin-11 h. Claudin-12 m. Claudin-13 m. Claudin-14 h. Claudin-15 h. Claudin-16 h. Claudin-17 h. Claudin-18 h. Claudin-19 h. Claudin-20 a

pKI Loop I 6.89 5.12 6.97 6.97 8.00 6.97 6.89 9.84 6.97 4.36 7.75 8.53 4.45 10.49 5.54 4.17 9.31 6.31 5.49 7.28

pKI Loop II 4.25 4.05 7.12 9.84 4.25 4.21 4.05 4.36 4.25 4.05 7.84 7.14 10.50 4.05 4.37 5.28 8.86 9.04 4.44 4.76

1st v. 2nd loop Acidic or Basica Neutral/acidic Acidic/acidic Neutral/neutral Neutral/basic Basic/acidic Neutral/acidic Neutral/acidic Basic/acidic Neutral/acidic Acidic/acidic Basic/basic Basic/neutral Acidic/basic Basic/acidic Acidic/acidic Acidic/acidic Basic/basic Acidic/basic Acidic/acidic Neutral/acidic

The pKIs of sequences predicted to encode the first and second loops were calculated using DNASTAR (Madison, WI).

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be as or more important than the number of charged amino acids. Claudin-16, a presumptive cation pore (Simon et al., 1999), is highly negatively charged in loop 2 as well, with a pKI of 5.28. Using claudin-16 as a model, one would predict that claudins 2, 10, 15, 16, and 19 are cation pores, while claudins 11 and 17 are anion pores. A second feature is the central (G)LW(M) common to all claudins in the center of the first loop, which could function in claudin-dependent adhesion. The first extracellular loop also contains highly conserved cysteine residues, which may stabilize intra- and interloop interactions via disulfide bridges, as is found in the extracellular domains of the connexins (Foote et al., 1998).

10.3.3 FREEZE-FRACTURE CHARACTERISTICS

OF

CLAUDINS

Transfection of claudin-1, -2, -3, -5, or -11 into L-cell fibroblasts all resulted in the novel appearance of continuous freeze-fracture fibrils. These results strongly suggest that claudins constitute the primary fibril-forming proteins (reviewed in Tsukita and Furuse, 1999) (Figure 10.1a). This is in contrast to occludin, which forms short fragments of strands when expressed in L cells (Figure 10.1b). Knockout of the claudin-11 gene in mice resulted in a complete loss of TJ fibrils in testicular Sertoli cells and central nervous system oligodendrocytes (Gow et al., 1999), demonstrating in vivo the importance of a single gene for fibril formation. In addition, the studies by Furuse and co-workers demonstrated that intrinsic properties of claudins might explain old freeze-fracture observations. For example, claudins 1 and 3 exogenously expressed in L cells produce continuous, smooth fibrils on the protoplasmic surface (P face) of replicas, while claudins 2 and 11 form discontinuous P-face fibrils (Furuse et al., 1998b; 1999; Morita et al., 1999a). The variable proportions of each claudin among tissues may now explain the variability in how TJs of different tissues fracture. The influence of cytoplasmic interactions on fibril formation and fracture face preference is cast into doubt by the observation that claudin-1, lacking the entire C-terminal tail and, therefore, unable to interact with ZOs or other plaque proteins, produces fibrils in L cells that look and fracture indistinguishably from full-length claudin-1 (Furuse et al., 1999).

10.3.4 DISTRIBUTION

OF

CLAUDINS

Individual claudins show different distribution patterns in different tissues (Gow et al., 1999; Simon et al., 1999; Rahner et al., 2001) supporting the idea that each claudin possesses distinctive functional properties. Within a single tissue, the distribution of different claudins can be quite variable. For example, immunofluorescent analysis of rat liver shows that both claudins 2 and 3 are concentrated at TJs, but claudin-2 shows a gradient in expression that increases from periportal to pericentral hepatocytes. On the other hand, claudin-3 is distributed in the same TJs but is expressed evenly across the liver lobule (Rahner et al., 2001). Claudin-5 was originally described as exclusively present at endothelial TJs (Morita et al., 1999b), but has more recently been found associated with TJs of pancreatic acinar cells (Rahner et al., 2001). Unexpectedly, the same claudins can be distributed along the lateral cell membranes in some tissues and be exclusively localized to the TJ in others.

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FIGURE 10.1 Freeze-fracture fibrils resulting from transfection of claudin-3 (a) or occludin (b) in L-cell fibroblasts. (a) Claudin-3-expressing L-cell transfectants reconstitute a highly developed network of parallel and branched fibrils. (From Sonoda, N. et al., J. Cell Biol., 147:195–204, 1999. With permission.) (b) Occludin-expressing L-cell fibroblasts produced short, strandlike structures (arrows), which were occasionally seen associated with gap junctions (asterisks). (From Furuse, M. et al., J. Cell Biol., 143:391–401, 1998. With permission.)

Immunofluorescent analysis of the distribution of claudin-4, for example, shows lateral membrane staining in a subset of colonic surface cells, but TJ staining in pancreatic duct epithelial cells and acini (Figure 10.2; Rahner et al., 2001). The observed tissue-specific physiological differences in TJs could be explained by differences in the composition of claudins within specific TJs. However, non-TJ localization of claudins is less easy to understand, since freeze-fracture fibrils are not normally observed on lateral cell membranes. One possibility is that claudins distributed on the lateral cell surface represent an available pool that can be recruited

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FIGURE 10.2 Localization of claudin-4 in rat pancreas (left) and colon (right). Claudin-4 in pancreatic duct epithelium (indicated with an asterisk) and acini (arrow) is restricted solely to the TJ. In contrast, claudin-4 is localized to a subset of surface colonic epithelial cells both at the TJ and along the lateral cell surface. (Courtesy of Dr. Christoph Rahner, Yale University.)

to form fibrils in response to various stimuli. An alternative explanation is that lateral claudins could have a second function independent of their role in TJs (Rahner et al., 2001).

10.4 FUNCTIONAL ANALYSIS OF OCCLUDIN AND CLAUDINS 10.4.1 TRANSEPITHELIAL ELECTRICAL RESISTANCE TJ-containing epithelial cells are characterized as having “tight” or “leaky” junctions based on their relative transepithelial electrical resistance, which roughly correlates with the number of TJ strands in the respective tissues (Diamond, 1974). Claude (1978) noted that the relationship between fibril number and resistance was not linear but, instead, that junctional resistance varied exponentially with strand number. She postulated that TJ fibrils contained aqueous pores that could vary between open and closed states (Claude, 1978). This model, as modifed by Cereijido et al. (1989) suggested that a solute traveling across the paracellular space would have to proceed in a stepwise fashion between adjacent TJ strands; the independent probability of encountering a compartmentalized open pore in each fibril would give rise to the observed relationship between strand number and resistance. Although this model fits many tissues, there are examples in which there is no correlation between fibril number and resistance. Type I and type II MDCK cells have identical fibril numbers but their transepithelial electrical resistance (TER) varies by nearly an order of magnitude, from less than 100 Ω·cm2 for low-resistance type II MDCK cells to about 800 Ω·cm2 for type I MDCK cells (Stevenson et al., 1988, Gonzalez-Mariscal et al., 1989). In addition, cooling MDCK cells from 37° to 3°C increases the transepithelial electrical resistance threefold, without affecting strand number or appearance. The

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basis for this and other discrepancies between fibril number and resistance are not explained by the Claude hypothesis. In view of the newly available information about the biochemical diversity of TJs, it is now clear that fibril number is only one possible determinant of paracellular permeability. The large number of members of the claudin family and their variable distribution and biochemical properties suggest that fibrils in any given tissue may have specific paracellular barrier properties depending on their claudin composition. For example, MDCK I cells might have fibrils containing claudins with intrinsically lower ion permeability than a different set of claudins in MDCK II cells. In addition, the large number of fibrils in a “tight” tissue might not only increase the physical barriers for ion transit across the junction, but, if these fibrils were associated with expression of a specific subset of claudins, more fibrils would also be expected to confer a higher level of selective permeability, analogous to increasing the height of an ion-exchange column. The identification of claudins and occludin as fibril constituents allows one to begin to ask how the biochemical characteristics of these proteins might function in generating particular paracellular barrier characteristics. 10.4.1.1 Occludin and Transepithelial Electrical Resistance Unfortunately, initial studies on the physiological role of occludin and claudin in the TJ have been difficult to interpret. Both Balda et al. (1996) and McCarthy et al. (1996) demonstrated that overexpression of chicken occludin in MDCK cells resulted in increases in TER. The effects of overexpression of occludin were somewhat variable, in that McCarthy et al. (1996), using an IPTG-inducible transgene expression system, reported an increase in TER that was about 30% above control values, along with a small increase in fibril number. In addition, these authors noted that there was an increase in the number of freeze-fracture strands that appeared as doublets (McCarthy et al., 1996). On the other hand, Balda and co-workers (1996), using multiple transfected MDCK cell clones as well as transgene induction with sodium butyrate, described a two- to fourfold increase in TER above control values with no significant effect on fibril number or appearance. To add to the confusion, another study demonstrated that overexpression of occludin in MDCK cells resulted in dramatic decreases in TER (Bamforth et al., 1999). Both McCarthy et al. (1996) and Balda et al. (1996) described an unexpected increase in flux of mannitol across the transfected cell monolayers. Although one criticism of these data had been the possibility for structural mismatch between exogenous chicken both groups used and endogenous canine occludin, this contradictory rise in both TER and flux was recently confirmed using canine occludin in MDCK cells (McCarthy et al., 2000). The increase in flux did not temporally parallel the increase in TER or the increase in occludin expression, suggesting that it was an effect secondary to the increased occludin expression, perhaps related to occludin mislocalization or induction of a second protein. One mechanism suggested by both groups was that occludin overexpression might regulate or increase the number of aqueous pores within the TJ strands. An increase in pore number might explain the discrepancy between the increased TER and flux, since TER is instantaneous measure of permeability, while flux is a measurement made over minutes to hours. Alternatively, McCarthy et al. (2000) also

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suggested that overexpression of occludin may titer available levels of scaffolding proteins such as ZOs, resulting in disrupted linkage to the contractile cytoskeletal network and altered pore regulation. Consistent with this idea was the demonstration that removal of the ZO-binding domain of occludin resulted in much larger increases in paracellular permeability in MDCK cells than was seen after transfection of fulllength occludin (Balda et al., 1996). In addition, although overexpressed occludin concentrated at TJs, it also localized to the lateral cell membranes (Medina et al., 2000), with an unknown contribution to physiological parameters. 10.4.1.2 Claudin and Transepithelial Electrical Resistance Although it now seems clear that claudins are the main structural element of the TJ fibrils, there are only two published reports to date on functional effects of claudin expression on TER and flux, and their results differ. Inai et al. (1999) demonstrated that expression of transfected claudin-1 in MDCK cells increases TER fourfold over untransfected MDCK cells. However, unlike the results seen with occludin transfection, there was reduced paracellular flux of both 4 and 40 kDa FITC-labeled dextrans (Inai et al., 1999). McCarthy et al. (2000) also obtained an increase in TER (fivefold) following overexpression of myc-epitope tagged claudin-1 in MDCK cells, but, in contrast to Inai et al. (1999), reported a rise in mannitol and 4 kDa FITC dextran flux. Interestingly, both untagged and C-terminally epitope-tagged claudin-1 increased TER, but only tagged claudin-1 affected flux (McCarthy et al., 2000). McCarthy et al. (2000) postulate that since C-terminally tagged claudin-1 is unlikely to interact with ZOs (presumably because the tag blocks the PDZ binding site), the paradoxical rise in TER and flux is likely a result of a perturbed and consequently unregulated claudin–cytoplasmic TJ protein linkage. In fact, these authors demonstrate the formation of aberrant fibrils, which lack associated ZO-1, along the lateral surface only in cells expressing tagged claudin-1. As noted above, the actual number of TJ strands is likely to be only one determinant of TER and may be less important than the combination of claudins, occludin, and potentially other proteins that compose specific cell-type fibrils. Overexpression of a single component of TJ fibrils might actually act to disorganize endogenous fibril proteins and thus confound interpretation of their functions. In addition, the interactions of the integral membrane proteins with ZO proteins and other cytoplasmic scaffolding proteins, and the importance of these interactions in regulating resistance, is unknown. However, it might be expected that overexpression of exogenous fibril proteins might also deform or overwhelm cytoplasmic scaffolding, resulting in undetermined effects on TER and flux. In summary, although both occludin and claudin are functional components of the TJ barrier, understanding their specific contributions will require more study and cautious interpretation of results.

10.4.2 OCCLUDIN AND CLAUDINS ARE CELL–CELL ADHESION MOLECULES Transmission electron microscopic images of TJs show intermittent areas of membrane apposition between adjacent cells, suggestive of adhesive contacts, which likely correspond to fibrils seen in freeze-fracture images. The ability of protein

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components of these fibrils to adhere to partners on adjacent cells is likely to be critical for formation of the physiological barrier. Both occludins and claudins have demonstrated adhesive activity; however, it seems likely that while occludin-dependent cell adhesion is weak and requires occludin localization to cadherin/ZO-1-based cell contacts, claudin-dependent adhesion is stronger and is independent of interactions with cytoplasmic proteins. Overexpression of occludin in insect cells by recombinant baculovirus infection resulted in the production of multilamellar bodies in which the luminal space was collapsed and membranes were fused, reminiscent of adherent sites in TJs (Furuse et al., 1996). This indirect demonstration of occludin adhesion was followed by studies in which administration of peptides corresponding to the second extracellular loop of occludin to Xenopus kidney epithelial cells could disrupt occludin localization and increase paracellular permeability (Wong and Gumbiner, 1997), suggesting that occludin localization and barrier function might be dependent on interactions of the extracellular loops. Similar peptides corresponding to the first extracellular loop of occludin delayed resealing of TJs disrupted by calcium removal (LacazVieira et al., 1999). Direct demonstration of occludin-dependent adhesion was shown in Rat-1 and NRK (but not mouse L-cell) fibroblasts that normally lack occludin. Expression of occludin in these cells resulted in increases in calcium-independent adhesiveness that could be blocked by peptides corresponding to the extracellular loops of occludin (Van Itallie and Anderson, 1997). Fibroblast expression studies also enabled the investigation of how claudins interact. With 20 members, the possibilities for combinatorial associations are many. However, initial studies of mixing cells expressing different claudins demonstrated that claudins may show defined partner preferences (Furuse et al., 1999). Similar to occludin studies, claudin-based calcium-independent adhesion was demonstrated by transfection of claudins 1, 2, and 3 into L-cell fibroblasts (Kubota et al., 1999). Transfection of any individual claudin into L-cells resulted in the formation of elaborate plaques between cells, which by freeze-fracture analysis contained welldefined fibrils. Adhesion was measured by an aggregation assay and aggregation was accompanied by the appearance of very close membrane appositions between transfected L-cells that were not normally seen in untransfected cells. Additional analysis of the interactions between these three claudins was carried out in a more recent study (Furuse et al., 1999). Coculture of claudin-1 and claudin-3 transfected cells resulted in the formation of plaques containing both proteins between adjacent cells. However, coculture of claudin-2 with either claudin-1 or claudin-3 expressing cells did not result in any apparent interactions. These authors suggested that although all three of these claudins could form plaques through homophilic interactions, only a subset was capable of heterophilic interactions that could lead to plaque formation. Since claudin distribution studies suggest that more than one type of claudin can exist at a single junction, specificity in claudin interactions would have very important implications in the generation of junction diversity. The adhesive interactions of occludin and claudins have differing dependencies on cytosolic interactions. The ability of occludin to form adhesive contacts is normally dependent on cytosolic interactions. Occludin was not adhesive in cells that lacked cadherin-based, ZO-containing cell contacts. In addition, occludin lacking

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the last 150 amino acids of its carboxy terminus, which contains the ZO-binding region, did not localize to sites of cell contact. This truncated occludin was not adhesive even in cells that had well-differentiated cadherin-based cell contacts (Van Itallie and Anderson, 1997). Unlike occludin, the ability of claudins to form adhesive plaques at cell contacts was independent of cytosolic interactions, since these proteins localized to adhesive plaques in L cells and formed apparently normal fibrils even when the cytosolic carboxy terminus was removed (Furuse et al., 1999). This suggested that occludin, which lacks the intrinsic ability to polymerize that is characteristic of at least some claudins (see Figure 10.1), might not be sufficiently concentrated to be adhesive in the absence of a localization signal such as ZO proteins. At least some of the claudins, on the other hand, aggregated both laterally and transcellularly, suggesting either that the lateral aggregation concentrated claudins sufficiently to promote transcellular adhesion or that the adhesion mediated by claudins was so much stronger than that by occludin that concentration of claudin by a targeting signal was not required. Along with this adhesive interaction demonstrated between cells, claudins and occludin interacted to form the linear fibrils characteristic of the TJs. Coexpression studies in L-cells revealed that claudin-1 could recruit occludin into continuous claudin-1 fibril networks (Furuse et al., 1998b). These results suggested claudin-1 may directly associate with occludin, or that the two proteins are coassembled indirectly by virtue of their joint interactions with cytoplasmic plaque proteins. If claudin and occludin can associate directly, it would be laterally in the membrane, as occludin and claudins do not associate transcellularly (Furuse et al., 1999). In fact, Furuse et al. (1999) demonstrated that claudins 1, 2, and 3 could all copolymerize within a single strand, suggesting that specificity of claudin interactions is more likely to be regulated in transcellular adhesion than lateral polymerization. Although the ability for various claudins and occludin to participate in lateral association provides insights into the overall composition of fibrils, what is missing is insight into the organization of the particles that make up the freeze-fracture strands.

10.5 INTERACTIONS OF OCCLUDIN AND CLAUDINS WITH OTHER PROTEINS In addition to their interactions with each other, both occludin and claudins associate with a number of other proteins. Occludin binds to itself (Chen et al., 1997), the ZO proteins, ZO-1, ZO-2, and ZO-3 (Furuse et al., 1994; Itoh et al., 1999b; Wittchen et al., 1999), actin (Wittchen et al., 1999), and VAP-33 (Lapierre et al., 1999). Using a bait peptide of 27 amino acids near the C-terminal end of occludin, Nusrat et al. (2000) demonstrated that this region of occludin could also bind to protein kinase C-ζ, the protein tyrosine kinase c-yes, the regulatory subunit of phosphatidylinositol 3-kinase, and the gap junction protein connexin-26; however, these interactions were not confirmed by coimmunoprecipitations of endogenous proteins (Nusrat et al., 2000). The claudins, as mentioned above, can associate with themselves, occludin, and in some circumstances with other claudins. In addition, most of the claudin tails end in YV, a sequence reminiscent of a PDZ-binding motif (Woods and Bryant, 1993; Anderson, 1996). Indeed, a direct interaction of the first PDZ domains of ZO-1,

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ZO-2, and ZO-3 with glutathione S-transferase (GST)-fusion proteins encoding the eight amino acid carboxy terminal tails of claudins 1 to 8 was recently demonstrated (Itoh et al., 1999a). The role of interactions with these cytoplasmic plaque proteins in the organization of the transmembrane proteins is unclear. In the case of occludin, the ZO-binding region can act as a signal for occludin localization. As mentioned above, in the absence of endogenous TJs, occludin lacking the ZO-binding region cannot localize to cell contact sites (Van Itallie and Anderson, 1997). Further, when the ZO-binding region of occludin is coupled to the extracellular and transmembrane domains of connexin-32, the resulting chimeric protein is incorporated into TJ fibrils (Mitic et al., 1999). In spite of the ability of the ZO-binding region to direct this localization, further results suggest that the ZO-binding region is neither necessary nor sufficient to direct occludin localization in MDCK cells (Matter and Balda, 1998; Medina et al., 2000). In the presence of endogenous occludin and claudin, occludin lacking the entire carboxyl terminus localized to TJs (Balda et al., 1996). In addition, truncation of the second extracellular loop of occludin results in a lateral but not TJ localization in MDCK cells, even in the presence of the ZO-binding domain (Medina et al., 2000). The simplest interpretation of both the chimeric and truncated occludin results is that an adhesive association mediated by the extracellular domains of occludin or connexin may be more important in maintaining TJ localization for occludin than its interactions with cytosolic plaque proteins. A primary dependence of localization on adhesive interactions rather than on interactions with cytoplasmic proteins may also be true for the claudins. Removal of the last three amino acids of claudin-1 comprising the PDZ-binding motif did not affect claudin localization in mouse L-cell fibroblasts, but did eliminate the ability of transfected claudins to cluster the ZO proteins at claudin-containing plaques (Furuse et al., 1999). However, the strongly adhesive nature of claudins may actually create de novo contact sites in fibroblasts, so that it is not clear how this can be interpreted in view of the very restricted distribution of fibrils in epithelial cells. Insight may be gained from claudin-11-null mice (Gow et al., 1999), where in the absence of claudin-based TJ fibrils in Sertoli cells there are still parallel depressions in the plasma membranes that show the same spacing as normal fibrils. These claudinnull mice still contain occludin and ZO-1, and it is not clear if these depressions are organized by interactions of occludin with cytoplasmic plaque proteins, by some modifications in the lipid organization by occludin or cytoplasmic plaque proteins, or by some other organizing principle. At any rate, the presence of the regular “fibril templates” strongly suggests that proteins as well as claudins organize the fibrils.

10.6 OCCLUDIN PHOSPHORYLATION AS A POSSIBLE DETERMINANT OF FIBRIL ORGANIZATION Claudins are not known to be subject to post-translational modification, but occludin is a variably serine- and threonine-phosphorylated protein, and the level of phosphorylation correlates with its localization in the junction. Sakakibara et al. (1997) took advantage of monoclonal antibodies that could distinguish hyperphosphorylated occludin from less-phosphorylated forms. They found hyperphosphorylated occludin

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FIGURE 10.3 Confocal immunofluorescent localization of occludin in frozen sections of chick intestine. An antibody that recognizes both hyperphosphorylated and less-phosphorylated forms of occludin labeled both junction-associated (arrow) and basolateral membraneassociated (arrowheads) pools of occludin. Further analysis using an antibody specific for hyperphosphorylated occludin revealed that it localized predominately at junctions and not basolateral membrane domains (not shown). (From Sakakibara, A. et al., J. Cell Biol., 137:1393–1401, 1997. With permission.)

was highly concentrated at the TJ; although less-phosphorylated occludin was junctional as well, but it was also distributed on the lateral surface of epithelial cells (Figure 10.3), a finding that led the authors to suggest that occludin phosphorylation was a key step in junction assembly (Sakakibara et al., 1997). Attempts to correlate changes in occludin phosphorylation state have been associated with changes in the state of TJ assembly or permeability, but the results have been somewhat difficult to interpret. Fashori and Kachar (1999) showed that dephosphorylation of occludin correlated with opening of TJs; while phosphorylation correlated with junction reformation (Farshori and Kachar, 1999). Similarly, Tsukamoto and Nigam (1999) showed occludin dephosphorylation after ATP depletion of MDCK cells that was associated with a dramatic fall in TER, followed during a recovery phase by increased phosphorylation and TER. However, Cordenonsi et al. (1997) found that occludin dephosphorylation was associated with de novo assembly of TJs in Xenopus embryos (Cordenonsi et al., 1997). Further, inhibition of tyrosine phosphatases results in occludin proteolysis and redistribution from the junction (Wachtel et al., 1999). In addition, vascular endothelial growth factor, which increases permeability of TJs, caused rapid increases in occludin phosphorylation (Antonetti et al., 1999). It appears that changes in occludin phosphorylation can be associated with changes in TJ function or assembly; however, the importance of this post-translational modification is unclear. Interestingly, occludin 1B is not phosphorylated, suggesting that the sequences occludin lacks but that occludin 1B contains may promote dephosphorylation (Muresan et al., 2000). The presence of a less-phosphorylated lateral membrane

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pool of occludin suggests that one potential mechanism for rapidly changing paracellular permeability might be recruitment of lateral occludin into fibrils. However, an alternative possibility is that the hyperphosphorylation of TJ-associated occludin is adventitious and unrelated to occludin function.

10.7 FUTURE DIRECTIONS The discovery of occludin and the members of the claudin family of proteins has allowed investigators to begin to ask fundamental questions about the nature of the long-recognized TJ fibrils. One critical question is that now that it is known that TJ fibrils are made up of a number of related but different proteins in varying combinations, how does this aid the understanding of tissue-specific resistance? If one imagines that a TJ fibril has both resistive and channel functions, are they both variable? Or is the resistive element simply the other side of a channel — that is, a closed channel? One pressing issue that remains is the nature of the particle that makes up freeze-fracture fibrils. It seems likely to be a claudin oligomer, but the number and arrangement of monomers into this structure is at present unknown. In addition, is it a homo-oligomer or a heteromeric structure composed of more than one claudin type? Does occludin form part of this structure or is it separately organized into the TJ fibrils? A second important issue that remains to be resolved is how the many members of the claudin family contribute to the variable tissuespecific paracellular transport properties observed. One obvious hypothesis is that the differing charged residues in the extracellular loops contribute to tissue-specific permeability properties, and that different claudins in a single fibril or particle might all contribute to the overall characteristics of different tissues. Again, the role of occludin in this process remains to be determined. Finally, the role of the cytoskeletal proteins in organizing and regulating the transmembrane proteins is imperfectly understood, and much available information needs to be reexamined once a better understanding of the organization of the TJ fibrils is achieved.

ACKNOWLEDGMENTS The authors are supported by grants from the National Institutes of Health (DK 45134, DK 38979).

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The Cytoplasmic Plaque Proteins of the Tight Junction Sandra Citi

CONTENTS 11.1 Introduction .................................................................................................232 11.2 TJ Plaque Proteins Containing PDZ Domains...........................................232 11.2.1 ZO-1...............................................................................................232 11.2.2 ZO-2...............................................................................................237 11.2.3 ZO-3...............................................................................................240 11.2.4 MAGI-1/BAP1 ...............................................................................240 11.2.5 AF-6 ...............................................................................................241 11.2.6 ASIP/PAR-3 and PAR-6 ................................................................242 11.3 Non-PDZ and Other TJ Plaque Proteins....................................................243 11.3.1 Cingulin .........................................................................................243 11.3.2 Symplekin ......................................................................................244 11.3.3 ZONAB..........................................................................................246 11.3.4 ASH1..............................................................................................247 11.3.5 Protein 4.1R ...................................................................................247 11.3.6 BG9.1 Antigen, 220 kDa Protein, 7H6, 19B1, Sec6/8 .................248 11.4 Cytoskeletal Proteins ..................................................................................248 11.4.1 Actin...............................................................................................248 11.4.2 Spectrin ..........................................................................................249 11.5 GTP-Binding Proteins and Protein Kinases...............................................249 11.5.1 RAB Proteins .................................................................................249 11.5.2 G Proteins ......................................................................................250 11.5.3 Protein Kinases ..............................................................................250 11.6 Phosphorylation of TJ Plaque Proteins ......................................................251 11.7 The Architecture of the Cytoplasmic Plaque of TJ ...................................251 11.8 The Role of TJ Cytoplasmic Plaque Proteins in TJ Function ...................252 11.9 Nuclear Localization of TJ Plaque Proteins: A Role in Regulation of Gene Expression? .......................................................................................252 11.10 TJ Plaque Proteins and Disease .................................................................253 11.11 Concluding Remarks...................................................................................254 Acknowledgments..................................................................................................254 References..............................................................................................................254 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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11.1 INTRODUCTION The cytoplasmic plaque of tight junctions (TJs) is the area of cytoplasm beneath the TJ membrane. Farquhar and Palade (1963) described the cytoplasmic plaque of TJs as a 0.2- to 0.5-µm-long “diffuse band of dense cytoplasmic material,” the development of which varies from one epithelium to another (Figure 11.1). The condensed material in the TJ plaque is less conspicuous and fibrillar than that of adherens-type junctions (zonula adhaerens and desmosome) (Farquhar and Palade, 1963). It is now clear that the cytoplasm beneath the TJ membrane is highly organized, and contains proteins whose presumed functions are to link TJ membrane proteins to the cytoskeleton, and to act as targets and effectors of signaling pathways involved in cell growth and differentiation and in modulation of TJ function. The aim of this chapter is to summarize the present state of knowledge about the structure, expression, and molecular interactions of TJ plaque proteins. The reader is referred to other chapters in this book for a more detailed description of TJ plaque proteins in development (Chapter 13 by Fleming et al.), signaling pathways (Chapter 17 by Balda and others), and cytoskeletal organization (Chapter 12 by Fanning). In this chapter, TJ plaque proteins have been subdivided arbitrarily into five groups (Figure 11.2): (1) TJ plaque proteins containing PDZ domains; (2) non-PDZ TJ plaque proteins; (3) other TJ plaque proteins; (4) cytoskeletal proteins; (5) signaling proteins (GTP-binding proteins and protein kinases).

11.2 TJ PLAQUE PROTEINS CONTAINING PDZ DOMAINS A significant group of TJ plaque proteins is characterized by the presence of a ~90residue-long region, called the PDZ domain (from PSD-95/SAP90, Discs-large, and ZO-1, the first proteins where such domains were identified). PDZ domains mediate protein–protein interactions with other PDZ-containing proteins, and with specific sequences at the carboxy-terminal ends of membrane protein, and may thus function by clustering protein complexes at the plasma membrane (Kennedy, 1995; Kim et al., 1995; Fanning and Anderson, 1996; Craven and Bredt, 1998). The TJ plaque proteins with PDZ domains include a subset (ZO-1, ZO-2, ZO-3, and MAGI-1/BAP1) belonging to the MAGUK (membrane-associated guanylate kinase) family of proteins (see Figure 11.2). MAGUK proteins contain a domain homologous to yeast guanylate kinase (GUK) in addition to a variable number (three to five) of PDZ domains. MAGUK proteins are localized on the cytoplasmic face of the plasma membrane in several cell types, where they are believed to have scaffolding and signaling functions (Anderson, 1996; Fanning and Anderson, 1999).

11.2.1 ZO-1 ZO-1 (zonula occludens-1) was identified as an Mr 225 kDa antigen recognized by a monoclonal antibody generated against a membrane preparation of mouse liver (Stevenson et al., 1986). By immunoelectron microscopy the anti-ZO-1 antibody labeled the cytoplasmic faces of TJ in bile canaliculus-enriched plasma membranes

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FIGURE 11.1 The ultrastructure of the cytoplasmic plaque of TJs. Transmission electron micrograph of the junctional complex of cells of the distal convoluted tubule of rat kidney. The TJ extends along the apicobasal axis between the arrows (~0.33 µm long). The dashed box indicates the area of cytoplasm extending to ~65 nm from the midline of the TJ membrane. (Modified from Farquhar, M.G. and Palade, G. E., J. Cell Biol., 17, 375–412, 1963. By copyright permission of the Rockefeller University Press.)

(Stevenson et al., 1986) (Table 11.1). ZO-1 was subsequently found to be localized at adherens-type junctions in fibroblasts and other cells devoid of TJ (Itoh et al., 1991; Howarth et al., 1992). Partial cDNA sequencing showed that ZO-1 exists in

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FIGURE 11.2 A simplified classification of TJ plaque proteins.

two isoforms (ZO-1α+ and ZO-1α–) differing by an internal 80-residue domain, termed domain α (Willott et al., 1992). Most epithelial cells express both isoforms, but exclusive expression of the α- isoform appears restricted to cells with a higher degree of junctional plasticity, such as endothelia, seminiferous tubules, and glomerular podocytes (Kurihara et al., 1992; Balda and Anderson, 1993) or to cells forming immature junctions during development (Sheth et al., 1997). Additional splicing regions have been identified in canine ZO-1 (Gonzalez-Mariscal et al., 1999). Mouse ZO-1 was independently identified as an Mr 220 kDa component of cadherin-based adherens junctions (Itoh et al., 1991). Cloning and sequencing of this protein showed that it was identical to ZO-1, based on the presence of the α domain sequence (Itoh et al., 1993). Analysis of human ZO-1 sequence revealed that the N-terminal half of ZO-1 contains three (PDZ) domains homologous to domains detected in the Drosophila melanogaster dlg tumor suppressor gene product, the rat brain PSD-95 protein, and the human erythrocyte p55 protein (Willott et al., 1993). In addition, ZO-1 contains one repeat homologous to the src oncogene homology region 3 (SH3) and one repeat homologous to yeast guanylate kinase (GUK). The

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TABLE 11.1 An Alphabetical List of Vertebrate TJ Plaque Proteins with Known GenBank Accession Numbers Protein

Mr (kDa)

AF-6

180–195

ASH1 ASIP

~330 140–180

Cingulin

MAGI-1

140 160 140 137–148

PAR-6

37

Rab13 Rab3B

24 25

Symplekin

110–150

ZO-1

210–225

ZO-2

ZO-3

ZONAB

160

130

47–55

Reference Prasad et al., 1993 Yamamoto et al., 1997 Nakamura et al., 2000 Izumi et al., 1998 Izumi et al., 1998 Joberty et al., 2000 Citi et al., 1988 Cordenonsi et al., 1999 Citi et al., 2000 Nagase et al., 2000 Dobrosotskaya et al., 1997 Shiratsuchi et al., 1998 Ide et al., 1999 Hung and Kemphues, 1999 Johansson et al., 2000 Choi et al., 2000 Zahraoui et al., 1994 Weber et al., 1994 Klengel et al., 1997 Keon et al., 1996 Ueki et al., 1997 Takagaki et al., 2000 Stevenson et al., 1986 Willott et al., 1993 Itoh et al. 1993 Gonzalez-Mariscal et al., 1999 Gumbiner et al., 1991 Jesaitis et al., 1994 Duclos et al., 1994 Collins and Rizzolo, 1998 Itoh et al., 1999b Balda et al., 1993 Haskins et al., 1998 Itoh et al., 1999a Itoh et al., 1999a Balda and Matter, 2000

Species/ Chromosome

GenBank

Human 6q27

U02478

Human 1q21 Rat Human 10p11.2 Human Chicken Xenopus Human 1q21 Human Mouse Human Rat Mouse Human Xenopus Human Human Rat Human Human 19q13.3

AF257305 AB005549

Notes TJ local.

Mouse Human 15q13 Mouse Canine Canine Canine Human 9q13-q21 Chicken Mouse Canine Canine Mouse Mouse Canine

AF252293

Chr. local. (Par3) Id./TJ local.

AF207901 AF263462 AB037740 AF027503/4/5 AB010894

Unidentif. (BAP1) TJ local.

AF070970 CAB85490 AF152346 X75593 M28214 Y14019 U49240 U88726 Id./TJ local. L14837 D14340 U55935 Id. L27152 L27476 AF085184 AF113005

(X104)

Id. (p130) AF023617 AF157006 NM_013769 AF171061/2

(Tjp3)

Note: The first column lists the protein names. The second column lists the molecular sizes (determined in most cases on the basis of mobility by SDS-PAGE). The third column lists key references for the identification and/or TJ localization of each protein, and/or the description of cDNA/protein sequences. The fourth column lists the species and human chromosomal localization (when applicable) related to each reference. The fifth column lists the GenBank accession numbers. The last column contains additional information for references in which no new sequence was reported, but in which the protein was identified and/or localized at TJs by immunoelectron microscopy. Id. = paper reporting identification of protein; Chr. local. = paper reporting human gene chromosomal localization; TJ local. = paper reporting TJ localization of protein; Unidentif. = sequence not identified as cingulin by Nagase et al. (2000). The names in parentheses in this column refer to alternative names by which the same protein is known in different species.

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C-terminal half of human ZO-1 comprises an acidic region, the α motif, and a large proline-rich region (Willott et al., 1993) (Figure 11.3). ZO-1 associates with a number of junctional and cytoskeletal proteins (Figure 11.3). Its PDZ domains interact with the TJ membrane protein claudin (PDZ1) (Itoh et al., 1999a), with the TJ plaque proteins ZO-2 (PDZ2) (Fanning et al., 1998; Itoh et al., 1999b), and ZO-3 (PDZ2) (Itoh et al., 1999a), and the gap junction protein connexin-43 (PDZ2) (Giepmans and Moolenaar, 1998) (see Figure 11.3). ZO-2 and ZO-3 are detected in ZO-1 immunoprecipitates (Gumbiner et al., 1991; Balda et al., 1993; Wittchen et al., 1999), indicating that a complex of ZO-1 with these proteins occurs in vivo. The SH3 domain is a ~60-residue motif detected in several cytoskeletal and signaling proteins that link protein tyrosine kinases to different signaling pathways (Musacchio et al., 1992; Schlessinger, 1994). The SH3 domain of ZO-1 interacts with a serine protein kinase (ZAK) (Balda et al., 1996) and with the nucleic acid–binding protein ZONAB (Balda and Matter, 2000) (see Figure 11.3). The activity of the GUK domain, whose structure is similar to that of small GTP-binding proteins, could elevate the GDP/GTP ratio in the cells and thus inactivate G proteins (Bryant and Woods, 1992). However, no GUK activity has been detected for ZO-1, and it is unlikely that such an activity exists, since key residues in the catalytic site of the ZO-1 GUK domain are missing (Willott et al., 1993). Although it remains to be determined whether ZO-1 interacts with G-protein-binding proteins, a 244-residue region comprising the GUK domain associates with occludin (Fanning et al., 1998) (see Figure 11.3). Several other protein interactions of ZO-1 have been reported. In low-salt extracts of isolated adherens junctions, ZO-1 immunoprecipitates with α-spectrin and appears to bind to a position ~10 to 20 nm from the midpoint of the spectrin tetramer (Itoh et al., 1991). An N-terminal fragment of ZO-1 interacts with α-catenin (Kd = ~0.5 nM) (Itoh et al., 1997) and with AF-6 (Yamamoto et al., 1997). An N-terminal fragment of cingulin interacts with full-length ZO-1 (Kd = ~5 nM) (Cordenonsi et al., 1999a). JAM coimmunoprecipitates with ZO-1 and the C-terminal cytoplasmic tail of JAM binds to ZO-1 in vitro (Bazzoni et al., 2000). The proline-rich region of ZO-1, which contains the differentially spliced motif α, interacts with F-actin (Kd = ~10 nM) (Itoh et al., 1997; Fanning et al., 1998). ZO-1 coimmunoprecipitates with cortactin (Katsube et al., 1998), an F-actin binding protein originally identified as a substrate for Src protein tyrosine kinase (Wu and Parsons, 1993). Interestingly, cortactin interacts with the proline-rich C-terminal region of Tamou, the D. melanogaster homologue of ZO-1 (Takahisa et al., 1996), suggesting that the same region of ZO-1 is involved in cortactin binding. The product of the D. melanogaster tamou gene (Takahisa et al., 1996) is more closely related to ZO-1 than the product of dlg gene. Mutations of tamou lead to a defective development of sensory organs, a phenotype similar to that generated by mutations of the extramacrochaetae (emc) gene, a repressor of helix-loop-helix transcription factors involved in the initiation of neural development. Since the tamou gene product (Tam) has been localized at cell–cell junctions, it has been postulated that Tam protein is implicated in the signaling pathway that activates emc expression

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(Takahisa et al., 1996). Recent experiments suggest that ZO-1 may also be implicated in gene expression (see Section 11.9). In addition to a potential role in regulation of gene expression and cell growth (Section 11.9), the protein interactions, expression, and subcellular localization of ZO-1 indicate that this protein has a scaffolding function and is implicated in junction biogenesis and architecture. The N-terminal half of ZO-1 may cluster transmembrane junctional proteins and other PDZ-containing proteins at sites of cell–cell contact, and create a submembrane network that is anchored to the cytoskeleton through the C-terminal half of ZO-1.

11.2.2 ZO-2 ZO-2 was identified as an Mr 160 kDa polypeptide that coimmunoprecipitates with ZO-1 but has a distinct antibody reactivity, peptide map, and turnover rate (Gumbiner et al., 1991) (see Table 11.1). Partial cDNA sequence showed that ZO-2 contains three PDZ domains, one SH3, and one GUK domain and is a member of the MAGUK family (Jesaitis and Goodenough, 1994) (see Figure 11.3). A human cDNA clone for ZO-2 was cloned while attempting to identify the Friedreich ataxia gene locus (Duclos et al., 1994). The complete ZO-2 protein has overall 51% sequence identity with ZO-1, contains a 36-residue alternatively spliced region (β) at the C-terminal end, and a C-terminal proline-rich region with only 25% amino acid identity with ZO-1 (Beatch et al., 1996) (see Figure 11.3). The protein-binding MAGUK domains are most conserved between ZO-2 from avian and mammalian species (Collins and Rizzolo, 1998) and between ZO-2 and other MAGUK proteins. ZO-2 has been localized to the TJ plaque by immunoelectron microscopy of MDCK liver membranes and is junctionally localized in liver, kidney, intestine, and testis epithelia (Jesaitis and Goodenough, 1994). Like ZO-1, ZO-2 interacts in vitro with occludin and α-catenin through an N-terminal fragment (Itoh et al., 1999b) (see Figure 11.3). A C-terminal fragment of ZO-2 associates with actin microfilaments in transfected fibroblasts, suggesting that the proline-rich region of ZO-2 associates with actin (Itoh et al., 1999b). Indeed, purified recombinant ZO-2 interacts with F-actin in vitro (Wittchen et al., 1999). Recruitment of ZO-2 to TJ in transfected cells requires the interaction of ZO-2 with claudin through the PDZ1 domain and/or with ZO-1 through the PDZ2 domain (Itoh et al., 1999a, b). Finally, an N-terminal fragment of cingulin interacts with full-length ZO-2 in vitro, and cingulin coimmunoprecipitates with ZO-2 (Cordenonsi et al., 1999a) (Figure 11.3). One interesting aspect of ZO-1 and ZO-2 is that both proteins have been detected in many types of nonepithelial cells. In these cells, ZO-1 and ZO-2 are found in areas of cell–cell contact, associated with cadherins (Itoh et al., 1991; 1993; 1997; 1999b; Howarth et al., 1992; Jesaitis and Goodenough, 1994; Yonemura et al., 1995). Studies on developing embryos and cultured cells indicate that epithelial cell polarization involves the redistribution of occludin into ZO-1-containing contact sites, whereas cadherin is sorted out from the same sites (Sheth et al., 1997; AndoAkatsuka et al., 1999) (see Chapter 13 by Fleming et al.). Why ZO-1 and ZO-2 are not associated with cadherins in TJ-bearing cells remains unclear, although a higher

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FIGURE 11.3 The domain organization and protein interactions of some TJ plaque proteins. h = human; c = canine; r = rat; m = mouse; x = Xenopus. The names of the protein domains are indicated above or below the schematic diagram of the protein (see text for details). NLS (nuclear localization signals) determined by PSORTII program are indicated by arrowheads

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FIGURE 11.3 (continued) below the diagram. Note that for h-ASH1 only bipartite NLS were indicated. Lines below the diagram indicate the region(s) involved in interactions with other proteins. Some of the protein interactions indicated have been established with protein from a species other than the one shown in the diagram.

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affinity of interaction with specific TJ proteins such as occludin and claudins remains a plausible hypothesis. In summary, ZO-2 appears similar to ZO-1 in its domain organization, protein interactions, tissue expression, and subcellular localization, and it is unclear whether ZO-1 and ZO-2 play redundant or distinct roles.

11.2.3 ZO-3 ZO-3 was identified in ZO-1 immunoprecipitates as a phosphorylated Mr 130 kDa polypeptide (p130) (Balda et al., 1993) (see Table 11.1). ZO-3 is homologous to ZO-1, ZO-2 (47% amino acid identity), and other MAGUK proteins. The sequence linking PDZ2 and PDZ3 domains of ZO-3 is unique, and is rich in proline residues (Haskins et al., 1998) (see Figure 11.3). When epitope-tagged ZO-3 is transfected into MDCK cells, it colocalizes with ZO-1 in TJ membranes by immunoelectron microscopy (Haskins et al., 1998). An in vitro interaction between ZO-3 and ZO-1, but not between ZO-3 and ZO-2, has been detected, suggesting that ZO-1 forms independent complexes with ZO-2 and ZO-3 (Haskins et al., 1998; Wittchen et al., 1999). Purified recombinant ZO-3 interacts with F-actin (Wittchen et al., 1999), and the first PDZ domain interacts with the cytoplasmic domain of claudin-1 in vitro (Itoh et al., 1999a) (see Figure 11.3). Some of the in vitro binding partners of ZO-1/ZO-2/ZO-3 may be relevant for their localization in vivo. Transfection experiments in L cells indicate that the interaction of claudin-1 with the PDZ1 domain of ZO-1, ZO-2, and ZO-3 and the interactions between the PDZ2 domains of the ZO proteins can recruit these proteins to claudin-based networks and may be responsible for TJ targeting in epithelial cells (Itoh et al., 1999a). Transfection of JAM into CHO cells results in an increased accumulation of junctional ZO-1 (Bazzoni et al., 2000), but anti-JAM antibodies do not prevent junctional assembly of ZO-1 and partial recovery of TJ barrier function in epithelial (T84) monolayers after calcium switch (Liu et al., 2000). These observations suggest that increased cell–cell adhesion favors ZO-1 accumulation at junctions, but JAM-mediated homophylic adhesion is not critical for ZO-1 localization. Occludin interaction with ZO-1, ZO-2, and ZO-3 is not important for the recruitment of these proteins to the TJ, since occludin-deficient cells and tissues show normal junctional localization of ZO-1, ZO-2, and ZO-3 (Saitou et al., 1998; Itoh et al., 1999a). Similarly, AF-6-deficient mouse embryonic stem cells show normal junctional localization of ZO-1, indicating that junctional targeting of ZO-1 does not require AF-6 (Zhadanov et al., 1999).

11.2.4 MAGI-1/BAP1 MAGI-1 (membrane associated guanylate kinase with an inverted arrangement of protein–protein interaction domains) was identified in a yeast two hybrid screen as a mouse protein interacting with the small GTP-binding protein K-RasB (Dobrosotskaya et al., 1997). BAP1 (BAI-associated Protein-1) is the human counterpart of MAGI-1, and was identified by its interaction with brain angiogenesis inhibitor (BAI) (Shiratsuchi et al., 1998) (Table 11.1). These proteins are characterized by the presence

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of an N-terminal GUK domain, two WW (tryptophan–tryptophan) domains, and four (MAGI-1) or five (BAP1) PDZ domains (see Figure 11.3). WW domains were first identified as two repeats in mouse YAP-65 (Yes-associated protein of 65 kDa) (Sudol et al., 1995) and interact with polyproline sequences conforming to the consensus PPXY (PY motif) and with other sequences (Chan et al., 1996). Thus, there is a protein-interaction overlap between WW domains and SH3 domains, raising the possibility that these domains may compete for interaction with the same ligand (Chan et al., 1996). The “inverted” arrangement of protein–protein interaction domains refers to the fact that, unlike DlgA and ZO-1, in MAGI-1/BAP-1 the GUK domain is in the N-terminal part of the molecule and the PDZ domains are in the C-terminal part of the molecule (Dobrosotskaya et al., 1997). Recently, MAGI-1/BAP1 was localized at the TJ cytoplasmic plaque by immunoelectron microscopy of rat intestinal cells and by colocalization with ZO-1 in MDCK cells (Ide et al., 1999). The antibodies used (Ide et al., 1999) were raised against the WW domain of S-SCAM, a neuronal protein that displays 50% sequence homology to MAGI-1/BAP1 (Hirao et al., 1998). The interactions of MAGI-1 with other TJ proteins and its possible role in TJ are not known.

11.2.5 AF-6 AF-6 is a protein essential for neuroepithelial development, and its localization at TJ is controversial. The AF-6 gene (from ALL-1 Fusion partner-6) was originally identified as the fusion partner of the ALL-1 gene in the t(6:11) translocation found in some acute myeloid leukemias (Prasad et al., 1993) (Table 11.1). AF-6 contains one PDZ domain (Prasad et al., 1993), one domain homologous to class V myosins (Prasad et al., 1993; Ponting, 1995), and one domain homologous to the Caenorhabditis elegans kinesin-like protein unc-104 (Ponting, 1995) (see Figure 11.3). Its domain organization is similar to Canoe, a D. melanogaster protein involved in signaling pathways regulating the number of eye cone cells (Miyamoto et al., 1995; Ponting, 1995). AF-6 was also identified as a Ras partner by 2-hybrid (Van Aelst et al., 1994) and GSTpulldown (Kuriyama et al., 1996) assays. In fact, two Ras-binding domains have been identified in the N-terminal region of AF-6 (Ponting and Benjamin, 1996) (see Figure 11.3). The idea that AF-6 may be a Ras effector is supported by the characterization of mutants of the C. elegans homologue Ce-AF-6 (Watari et al., 1998). AF-6 was localized by immunoelectron microscopy in the TJ of MDCK cells (Yamamoto et al., 1997). However, a splicing variant of AF-6 (l-afadin), which contains an additional F-actin-binding domain of ~200 residues at the C-terminal end, was localized by immunoelectron microscopy in cadherin-based adherens junctions, but not TJ of epithelial intestinal cells (Mandai et al., 1997). Thus, the TJ localization of AF-6 is controversial, and may depend on splicing variant and cell type. Interestingly, in cadherin-deficient L cells, AF-6 is still localized at cell–cell contact sites, whereas ZO-1 is localized at the tip area of cell processes (Sakisaka et al., 1999). In addition to interacting with Ras (Van Aelst et al., 1994; Kuriyama et al., 1996; Yamamoto et al., 1999), an N-terminal fragment of AF-6 associates in vitro with ZO-1 and occludin, but not E-cadherin or catenins (Yamamoto et al., 1997) (see Figure 11.3). The interaction between ZO-1 and AF-6 is disrupted by activated Ras

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(Yamamoto et al., 1997) suggesting that ZO-1- and Ras-binding sites are overlapping. Interestingly, the fly homologue of ZO-1 (Tam) interacts with Canoe, an AF-6 homologue (Takahashi et al., 1998). The PDZ domain of AF-6 interacts in vitro with Eph-related receptor tyrosine kinases, neurexin, and the Notch ligand Jagged (Hock et al., 1998; Buchert et al., 1999). It also interacts with nectin/PRR (poliovirus receptor–related protein) (Takahashi et al., 1999) and with the TJ membrane protein JAM (Ebnet et al., 2000) (see Figure 11.3), indicating that AF-6 may function as a scaffolding module for the subcellular targeting of transmembrane proteins containing the PDZ-binding C-terminal motif. The interaction between AF-6 and the receptor tyrosine kinase EphB3 occurs in vivo and depends on receptor kinase activity, and AF-6 is an in vivo substrate of EphB3 (Hock et al., 1998; Buchert et al., 1999). The AF-6 interaction with nectin, a homophylic cell–cell adhesion molecule of the immunoglobulin superfamily, is required for the recruitment of nectin to cadherin-based adherens junctions (Takahashi et al., 1999). Similarly, the interaction of the PDZ domain of AF-6 with JAM may serve to recruit JAM into TJ, and clustering JAM with other structural or signaling proteins of cell–cell junctions (Ebnet et al., 2000). The C-terminal region of AF-6 contains proline-rich sequences and interacts with the Fam deubiquinating enzyme (Taya et al., 1998). Fam (Wood et al., 1997) is the mammalian homologue of the Drosophila fat facets gene, which encodes a deubiquinating enzyme involved in cell fate determination (Fischer-Vize et al., 1992; Cadavid et al., 2000). Fam colocalizes with AF-6 during mouse eye development (Kanai-Azuma et al., 2000). Recently, null mutations in the murine AF-6 (Zhadanov et al., 1999) and l-afadin (Ikeda et al., 1999) loci were generated. In both cases, homozygous mutant embryos died at ~10 days postcoitum as a result of a disorganization of the embryonic ectoderm, with loss of epithelial polarity and cell–cell junctions. Somites and other structures derived from ectoderm and mesoderm failed to form in the mutant embryos. However, the structural integrity of epithelia of trophoectoderm and visceral endoderm was maintained, suggesting that AF-6/afadin plays an important role only in neuroepithelial cells, such as those of the primitive streak and neural fold/groove, but not in junctions of preimplantation embryos. It has been proposed (Zhadanov et al., 1999) that AF-6 plays a key role in modulating cell–cell junctions downstream of Ras during the physiological reorganization of the neuroepithelium, which has been shown to involve loss of occludin and TJ, but not ZO-1 (Aaku-Saraste et al., 1996).

11.2.6 ASIP/PAR-3

AND

PAR-6

ASIP (atypical-PKC-specific interacting protein) was identified by screening NIH3T3 cell libraries with [32P]-labeled atypical protein kinase C zeta (PKCζ) (Izumi et al., 1998). ASIP was shown to bind to and colocalize with atypical protein kinase C (aPKC) isoforms zeta and lambda (PKCζ and PKCλ) at sites of cell–cell contact (Izumi et al., 1998) (see Table 11.1). Immunofluorescence microscopy shows that ASIP colocalizes with ZO-1 in cell–cell contact areas of fibroblasts and epithelial cells, and immunoelectron microscopy shows TJ localization in intestinal epithelial cells (Izumi et al., 1998).

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Sequence analysis shows that ASIP is the rat homologue of the C. elegans PAR-3 protein (Izumi et al., 1998). The par (partitioning defective) mutations in the C. elegans zygote lead to defects in cytoplasmic organization, distribution of cortical microfilaments, and establishment of anterior/posterior polarity (Kirby et al., 1990). PAR-3 is asymmetrically distributed in the C. elegans zygote, is required for correct spindle orientation (Etemad-Moghadam et al., 1995), and associates with the atypical protein kinase C PKC-3 (Tabuse et al., 1998). PAR-3-related proteins have been identified in D. melanogaster (Muller and Wieschaus, 1996; Kuchinke et al., 1998), mouse (Lin et al., 1999), and in humans (Joberty et al., 2000). ASIP and PAR-3 proteins contain an N-terminal conserved region (CR1), three PDZ domains (conserved region 2-CR2), and a third conserved region (CR3) contained within the aPKC binding domain (Figure 11.3). A cDNA corresponding to the mouse homologue of PAR-3 (PHIP) was isolated by screening a mouse embryo cDNA expression library with a peptide probe based on the putative PDZ domain binding site of ephrin 3B (Lin et al., 1999), suggesting an interaction between ASIP/PAR-3 and B-type ephrins (Figure 11.3). B-type ephrins are transmembrane proteins that function as ligands for B class Eph receptor tyrosine kinases and may function as regulators of cell–cell repulsion and adhesion (Wilkinson, 2000). The N-terminal region of human PAR-3, comprising the first PDZ domain, binds to human PAR-6B (Joberty et al., 2000) (Figure 11.3). PAR-6, another member of the C. elegans “par” family of genes (Watts et al., 1996) contains only one PDZ domain and may function to link PAR-3 to atypical PKCs (Hung and Kemphues, 1999; Joberty et al., 2000) and Cdc42 (Joberty et al., 2000; Johansson et al., 2000). In MDCK cells, endogenous PAR-6 shows a junctional localization overlapping with ZO-1 staining, suggesting that it is a TJ plaque component (Johansson et al., 2000). Unlike PAR-3, PAR-6 is also localized in the nuclei (Johansson et al., 2000) (see Section 11.9). Transfection of PAR-6 in MDCK cells perturbs the localization of PAR-3 and ZO-1, but not the organization of adherens junctions (Joberty et al., 2000). These and other studies (Wodarz et al., 2000) indicate that the complex comprising PAR-3, PAR-6, and atypical protein kinases C and Cdc42 is part of an evolutionary conserved mechanism that controls the establishment of apicobasal polarity in epithelial and other cell types. Northern and Western blot analysis indicate that mammalian PAR-3 and PAR-6 exist in different forms, with distinct sizes and patterns of tissue expression (Izumi et al., 1998; Joberty et al., 2000; Johansson et al., 2000), suggesting isoform- and tissue-specific functions.

11.3 NON-PDZ AND OTHER TJ PLAQUE PROTEINS 11.3.1 CINGULIN Cingulin (from the Latin cingere = to form a belt around) was identified using monoclonal antibodies raised against a preparation of purified subfragment-1 (S1) of brush-border myosin that contained cingulin fragments as minor contaminants (Citi et al., 1988) (see Table 11.1). Cingulin was localized to the cytoplasmic plaque of epithelial TJ by immunoelectron microscopy, and has been detected in polarized and stratified epithelia of several vertebrate species, but not in nonepithelial cells,

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“leaky” endothelia, and invertebrate organisms (Citi et al., 1988; 1989; 1991; Cardellini et al., 1996). The apparent size of cingulin is ~140 kDa in all mammalian species studied so far, whereas in Xenopus laevis it is ~160 kDa. The amino acid sequences of X. laevis and human cingulins show an N-terminal globular region (“head”), a central coiled-coil region (“rod”), and a C-terminal globular region (“tail”) (Cordenonsi et al., 1999a; Citi et al., 2000) (see Figure 11.3). The sequence of the coiled-coil region appears more conserved than the head region when comparing human and frog sequences (47.3 vs. 33% identity) (Citi et al., 2000). The rod domain of cingulin contains four (human) or five (X. laevis) repeated subdomains, suggesting the occurrence of gene duplication events within the rod sequence (see Figure 11.3). Sequence analysis indicates that the coiled-coil region of cingulin forms parallel dimers (Cordenonsi et al., 1999a), suggesting a molecular organization as shown in Figure 11.4A. This model is supported by electron microscopy examination of purified chicken cingulin rod and by the observation that purified chicken cingulin rod migrates as a dimer in non-reducing SDS-PAGE (Citi et al., 1988; 1989). Selfassociation of cingulin appears to depend primarily on rod interactions, as shown by GST pulldown assays using bacterially expressed X. laevis cingulin fragments (Cordenonsi et al., 1999a). Cingulin interacts with several TJ and cytoskeletal proteins (Cordenonsi et al., 1999a) (Figure 11.3). An N-terminal fragment of cingulin, comprising most of the head, interacts in vitro with ZO-1 (Kd = ~5 nM), ZO-2, ZO-3, myosin, AF-6 (Cordenonsi et al., 1999a), and JAM (Bazzoni et al., 2000). The association with ZO-1 and ZO-2 probably occurs in vivo, since ZO-1 and ZO-2 immunoprecipitates contain cingulin (Cordenonsi et al., 1999a). A C-terminal fragment of cingulin interacts with myosin (Cordenonsi et al., 1999a), in agreement with the observation that cingulin and myosin copurify from brush-border cell lysates (Citi et al., 1989). Full-length cingulin interacts in vitro with the cytoplasmic domain of occludin (Cordenonsi et al., 1999b) and with F-actin (D’Atri and Citi, unpublished observations). The bacterially expressed N-terminal fragment of cingulin interacts with several TJ proteins in vitro (Cordenonsi et al., 1999a). However, when the same fragment is transiently transfected into epithelial cells, the expressed protein does not target to TJ, but is localized in the nucleus and cytoplasm, suggesting that the head requires rod domain sequences to be stably targeted to TJ in vivo (Cordenonsi et al., 1999a). The rod domain may stabilize TJ targeting by interacting with specific sorting machineries or by allowing the molecule to fold correctly and dimerize. Indeed, cingulin stable targeting to TJ requires specific ZO-1-binding head sequences and more than ~100 residues of coiled-coil rod sequence (D’Atri et al., unpublished results).

11.3.2 SYMPLEKIN Symplekin (from the Greek συµπλεκειν = to tie together) was identified by a monoclonal antibody recognizing an Mr ~150 kDa antigen localized in the TJ plaque of epithelial cells and in the nucleus of epithelial and nonepithelial cells (Keon et al., 1996) (see Table 11.1). A cDNA coding for a protein of 1142 amino acids, and a predicted size of Mr ~126 kDa, was isolated (Keon et al., 1996). It was first reported

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FIGURE 11.4 Cingulin and the architecture of the TJ plaque. (A) Schematic diagram of a cingulin molecule, consisting of two parallel subunits, each with a globular head and a globular tail, separated by a central coiled-coil domain. The length of the coiled-coil region is ~120 to 130 nm. (B) Histogram showing the distribution of cingulin immunogold particles as a function of the distance from the midline of the TJ membrane in chicken intestinal epithelial cells labeled as in Citi et al. (1989). (C) Speculative diagram of the architecture of the TJ plaque, based on the measured distances of the immunogold particles for each protein. The inner area of the TJ plaque contains symplekin and ZO-1, and presumably also ZO-2 and ZO-3, plus other proteins. The outer area of the TJ plaque contains cingulin, 7H6, and other proteins.

that symplekin could not be detected in the cadherin-based adhesion sites of nonepithelial cells such as fibroblasts (Keon et al., 1996). However, recent studies indicate that a novel type of adherens-type junction in the retina contains symplekin together with ZO-1, vinculin, β-catenin, and plakophilin-2 (Paffenholz et al., 1999). Symplekin was subsequently identified by its interaction with huntingtin, a 350 kDa protein that is mutated in Huntington’s disease (Faber et al., 1998), and with CstF-64, a component of the heterotrimer cleavage stimulation factor (CstF) involved in the first step of mRNA precursor polyadenylation (Takagaki and Manley, 2000). cDNAs coding for two different forms of symplekin, symplekin-I (1273 amino acids,

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size by SDS-PAGE ~135 kDa) and symplekin-II (1058 amino acids, size by SDSPAGE ~110 kDa), were isolated (Takagaki and Manley, 2000). These cDNAs are identical to each other up to amino acid residue 964, but diverge thereafter. They differ from the sequence reported by Keon et al. (1996) since they contain an extra 187 amino acid residues in the N-terminal region. The presence of additional sequences 5′ to the originally reported 3.7-kB symplekin cDNA is also suggested by characterization of symplekin exon sequences (Ueki et al., 1997). Different forms of symplekin could arise from alternative splicing, although Northern blot analysis apparently detects only a 4.2 kB signal in most human tissues (Ueki et al., 1997). A central region of symplekin displays homology with the known yeast protein PTA1 (Takagaki and Manley, 2000), an essential component of the yeast polyadenylation machinery (Preker et al., 1997; Zhao et al., 1999). Thus, it has been suggested that symplekin plays a role as an assembly/scaffolding factor in mRNA polyadenylation (Takagaki and Manley, 2000). The first ~900 amino acids of symplekin, which contain the PTA1 homology region, display significant homology to protein sequences retrieved from D. melanogaster and C. elegans databases, suggesting that symplekin may be conserved through evolution. The interactions of symplekin with TJ proteins and the sequences involved in symplekin interaction with huntingtin and CstF-64 have not been characterized.

11.3.3 ZONAB ZONAB (ZO-1-associated nucleic acid-binding protein) was identified by screening an MDCK expression library with a radiolabeled GST fusion protein containing the SH3 and PDZ3 domain of ZO-1 (Balda and Matter, 2000) (Table 11.1). This protein exists in two closely related isoforms, ZONAB-A (47 kDa) and ZONAB-B (55 kDa), differing by a 68-residue alternative domain (AD) (see Figure 11.3). The sequence of ZONAB indicates that it is a member of the Y-box transcription factor family (Matsumoto and Wolffe, 1998) and is probably the canine homologue of human and mouse DNA-binding protein A (dbpA) (Sakura et al., 1988) and rat Y-box binding protein-a (RYB-a) (Ito et al., 1994). All these proteins contain conserved cold-shock and arginine- and proline-rich domains. ZO-1 and ZONAB coimmunoprecipitate from MDCK cell lysates, and GST pulldown experiments show that the SH3 domain of ZO-1 is responsible for the interaction (Balda and Matter, 2000) (see Figure 11.3). By immunofluorescence, ZONAB and ZO-1 are colocalized in the junctional regions of MDCK cells, although in growing cells a large fraction of ZONAB is detected in the nucleus. ZONAB interacts with promoter sequences of genes coding for ErbB-2 and cell cycle regulatory proteins (Balda and Matter, 2000). In transfected cells, ZO-1 and ZONAB regulate endogenous ErbB-2 expression in a concerted fashion, suggesting a model where ZONAB functions as a repressor of ErbB-2 expression that is sequestered to TJ by ZO-1 binding (Balda and Matter, 2000). ZONAB expression and distribution correlates with cell density and growth and inversely correlates with ZO-1 expression. In growing cells, ZO-1 expression levels are low, and ZONAB expression levels are high, and ZONAB localizes mostly in the nucleus. In confluent cells, ZO-1 expression levels are high, ZONAB expression levels are low, and low amounts of ZONAB are

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detectable in the nucleus (Balda and Matter, 2000). The idea that ZONAB is a growthinducible gene agrees with the observation that expression of the putative rat homologue (RYB-A) is induced by stimulation of quiescent fibroblasts with serum (Ito et al., 1994). Although the pattern of tissue expression of ZONAB has not been characterized, mRNA levels for the putative rat homologue RYB-a are high in skeletal muscle, spleen, and fetal liver, but very low in newborn and adult livers, suggesting that its expression is under developmental regulation (Ito et al., 1994).

11.3.4 ASH1 Human ASH1 protein was recently shown to be localized at TJ of cultured epithelial cells by colocalization with ZO-1 and cingulin (Nakamura et al., 2000). Northern analysis indicates that ASH1 is expressed in epithelial and nonepithelial tissues, with highest levels in brain, heart, and kidney. In addition to the junctional localization, ASH1 is localized in the nucleoplasm. Human ASH1 protein is predicted to be a very large (Mr ~330 kDa), basic protein (see Table 11.1). HuASH1 is the human homologue of the D. melanogaster gene ash1 (absent, small, or homeotic 1) (Tripoulas et al., 1996), a member of the trithorax transcription factor family, that regulates transcription of homeotic and other genes. HuASH1 protein sequence shows motifs (AT hooks, SET, bromodomain, PHD finger) shared with D. melanogaster ash1 and with other proteins involved in transcriptional regulation through chromatin alterations (see Figure 11.3). The AT hook is a 22-residue DNA-binding protein motif that may function to tether DNA-binding proteins to the minor groove of DNA (Aravind and Landsman, 1998). The SET domain has been identified in at least three D. melanogaster gene products (SUVAR(3)9, E(Z)PCO, and TRX). SET domain-containing proteins have been implicated in integration of upstream signaling pathways to epigenetic regulation and growth control (Firestein et al., 2000). Drosophila melanogaster ash1 interacts directly with trithorax (trx), and the interaction is mediated by the SET domains of the proteins (Rozovskaia et al., 2000) (Figure 11.3). The bromodomain is a conserved structure that recognizes acetylated residues and may serve as a signaling domain (Haynes et al., 1992). The PHD finger (plant homeodomain-finger) is a DNA-binding motif first detected in plants (Schindler et al., 1993). In D. melanogaster, mutations in ash1 cause transformation of the arista to leg, first leg to second leg, and other transformations, suggesting that ASH1 serves to maintain the expression pattern of homeotic selector genes (Shearn, 1989). The role of HuASH1 in TJ and its interactions with TJ proteins are not known.

11.3.5 PROTEIN 4.1R Two nonerythroid isoforms of the red cell protein 4.1R, with apparent size of Mr 135 kDa and Mr 150 kDa, were recently found to bind to human ZO-2 in yeast 2-hybrid screens, and to associate with ZO-1, occludin, spectrin, and actin in a complex (Mattagajasingh et al., 2000). The protein was localized to the TJ plaque by immunoelectron microscopy with antibodies against the N-terminal extension of 4.1R (Mattagajasingh et al., 2000). In vitro binding experiments indicate that the 4.1R-binding region of ZO-2 is located in the proline-rich C-terminal domain of

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ZO-2 (see Figure 11.3). Other MAGUK proteins, such as human Dlg and LIN2, interact with 4.1R through a conserved lysine-rich sequence between the SH3 and GUK domains (Lue et al., 1994; Marfatia et al., 1995; Cohen et al., 1998), which is missing in ZO-2. ZO-1 and occludin appear to interact only weakly with 4.1R by yeast 2-hybrid assay, suggesting that 4.1R links TJ plaque and membrane protein to the actin and spectrin cytoskeleton via interaction with ZO-2. Since other members of the protein 4.1 superfamily (merlin, ezrin, radixin, moesin) are implicated in Rho signaling pathways (reviewed in Tsukita and Yonemura, 1999), 4.1R may have a role in the functional linkage between TJ and the actin cytoskeleton.

11.3.6 BG9.1 ANTIGEN, 220 KDA PROTEIN, 7H6, 19B1, SEC6/8 The proteins listed below have been localized to the TJ plaque, but their amino acid sequences and/or interactions with TJ proteins have not yet been characterized. The BG9.1 antigen is an Mr 192 kDa mouse protein recognized by a monoclonal antibody raised against teratocarcinoma cells, and was localized at the cytoplasmic plaque of hepatocyte TJ by immunoelectron microscopy (Chapman and Eddy, 1989). A 220 kDa polypeptide identified in brain clathrin preparations was localized to rat epithelial TJ by immunoelectron microscopy, and may correspond to ZO-1 (Enrich et al., 1989). The 7H6 antigen was identified as an Mr 155 kDa protein recognized by a monoclonal antibody (7H6) raised against a bile canaliculus-rich fraction from rat liver, and was localized at TJ by immunoelectron microscopy (Zhong et al., 1993). An altered localization and decreased phosphorylation of 7H6 was detected in MDCK cells after ATP depletion (Zhong et al., 1994a). A partial protein sequence of 7H6 was reported, which shows a coiled-coil structure and ~30% sequence identity to yeast and vertebrate chromosomal segregation proteins (Ezoe et al., 1995). The monoclonal antibody 19B1 was generated using SDS-denatured X. laevis lung membrane preparation as immunogen, and the antigen was shown to colocalize with ZO-1 in A6 X. laevis kidney epithelial cells by immunofluorescence and immunogold microscopy (Merzdorf and Goodenough, 1997). The extractability properties of the 19B1 antigen indicate that it is a peripheral membrane protein. Sec6 (86 kDa) and Sec8 (110 kDa) are components of a ~17S complex in MDCK cells and show immunofluorescent colocalization with ZO-1 (Grindstaff et al., 1998). Upon calcium-dependent assembly of junctions, Sec6/8 is recruited to the apical junctional complex with kinetics similar to ZO-1, and becomes tightly associated with the membrane (Grindstaff et al., 1998). Antibodies against Sec8 inhibit basallateral protein delivery, suggesting that Sec6/8 may be involved in delivering membrane vesicles at the apical junctional complex.

11.4 CYTOSKELETAL PROTEINS 11.4.1 ACTIN Pharmacological and transfection studies show that the actin cytoskeleton plays an important role in TJ organization (see Chapter 12 by Fanning and other chapters in this book). The presence of actin in the TJ plaque was demonstrated by electron

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microscopy. First, the subfragment-1 of myosin-II decorated the cytoplasmic face of the TJ membrane (Madara, 1987). Second, actin was detected along the entire lateral plasma membrane of the TJ by immunoelectron microscopy, with a label density similar to that of the zonula adherens (Drenckhahn and Dermietzel, 1988). Thus, F-actin filaments are concentrated in the fibrillar electron-dense material of the TJ plaque. F-actin interacts in vitro with a growing number of protein components of the TJ plaque, including ZO-1 (Itoh et al., 1997; Fanning et al., 1998), ZO-2, ZO-3, occludin (Wittchen et al., 1999), and cingulin (D’Atri and Citi, unpublished observations). Thus, it is likely that direct interactions between F-actin and one or more TJ proteins contribute to TJ regulation by the actin cytoskeleton. Contraction of the actin cytoskeleton requires the actin-activated MgATPase activity of myosin. In addition, cingulin interacts with nonmuscle myosin in vitro (Cordenonsi et al., 1999a), and was originally identified as a minor contaminant of brush-border myosin preparations (Citi et al., 1988). These observations suggest that nonmuscle myosin II may associate with cingulin near the TJ plaque. However, no significant accumulation of myosin immunogold labeling was detected immediately beneath the TJ membrane by Drenckhahn and Dermietzel (1988). The lack of labeling could be caused by a low concentration of myosin in the TJ plaque, since myosin is distributed at high concentrations in the terminal web, and at much lower concentrations throughout the lateral membrane region of intestinal cells (Citi and Kendrick-Jones, 1991).

11.4.2 SPECTRIN The family of spectrin/fodrin proteins contains several isoforms of large, acidic actinbinding proteins, which are usually associated with the plasma membrane, polymerize into heterodimers and heterotetramers, and link transmembrane proteins to the cytoskeleton (Bennett and Gilligan, 1993). The first evidence linking spectrin to TJ was the observation that ZO-1 is present in α-spectrin immunoprecipitates and binds to a spectrin affinity column (Itoh et al., 1991). Furthermore, in MDCK monolayers ATP depletion leads to the association of fodrin and ZO-1 in an insoluble fraction (Tsukamoto and Nigam, 1997). When connexin-43 and ZO-1 are transfected into cultured cells and immunoprecipitated with anti-tag antibodies, they form a complex that contains spectrin, suggesting that ZO-1 may serve to localize connexin-43 at the intercalated disks (Toyofuku et al., 1998). Spectrin was immunoprecipitated with ZO-1 and other TJ proteins from extracts of confluent MDCK cells, but not nonconfluent MDCK cells (Mattagajasingh et al., 2000) and other cell types (Howarth and Stevenson, 1995). These observations suggest that association of ZO-1 with spectrin/fodrin may require the expression or post-translational modification of specific proteins associated with TJs of confluent epithelia.

11.5 GTP-BINDING PROTEINS AND PROTEIN KINASES 11.5.1 RAB PROTEINS Rab proteins are small GTP-binding proteins homologous with the proto-oncogene ras, and are involved in regulation of membrane traffic (Chavrier and Goud, 1999).

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Rab13 is closely related to the yeast Sec4 gene product and was localized to the junctional complex of several epithelia, but not in cardiac intercalated disks (Zahraoui et al., 1994) (Table 11.1). Although Rab13 is also detected in a cytoplasmic localization in nonepithelial cells, it colocalizes with ZO-1 at epithelial TJ and is not recruited to junctions by E-cadherin, suggesting some specific role in TJ assembly, function, or vesicle docking (Zahraoui et al., 1994). Rab3B is an epithelial-specific isoform of rab3 and was localized at TJs of cultured epithelial cells by immunoelectron microscopy (Weber et al., 1981) (Table 11.1). Rab3B targets to the apical pole during cell polarization following junction assembly induced by calcium-switch, and may therefore be involved in recruiting junctional and/or apical proteins during epithelial polarization (Weber et al., 1981), or in secretory functions (Klengel et al., 1997). Another member of the Rab family (Rab8) shows partial colocalization with ZO-1 (Huber et al., 1993).

11.5.2 G PROTEINS In normal renal and intestinal cell lines, the alpha subunit of heterotrimeric G protein (Gαi-2) is localized, at least in part, in the lateral junctional region (Hamilton and Nathanson, 1997). Its junctional distribution overlaps in part with the distribution of ZO-1 (de Almeida et al., 1994; Denker et al., 1996; Dodane and Kachar, 1996). In addition, transfected Gαo subunit is coimmunoprecipitated with ZO-1 from lysates of MDCK cells transfected with a Gαo construct (Denker et al., 1996). This evidence suggests that ZO-1 and/or other MAGUK TJ proteins may directly or indirectly associate with G proteins.

11.5.3 PROTEIN KINASES Atypical protein kinases C (aPKC = PKCζ and/or PKCλ) colocalize with ZO-1 (Dodane and Kachar, 1996; Izumi et al., 1998) and ASIP (Izumi et al., 1998) at sites of cell–cell contact by immunofluorescence, indicating that they are specific components of TJs. In addition, aPKC interact in vitro with ASIP and PAR-3 (Izumi et al., 1998). Atypical protein kinases C are distinct from conventional protein kinases C because they are not directly regulated by Ca2+, phorbol esters, or diacylglycerols, but are stimulated by signaling pathways involving PI3-kinase (Akimoto et al., 1996; Ettinger et al., 1996; Standaert et al., 1997). PKCζ and PKCλ associate with the GTP-binding protein Cdc42 and have been implicated in rasmediated reorganization of the actin cytoskeleton (Uberall et al., 1999; Coghlan et al., 2000; Hellbert et al., 2000). Another protein kinase that may be associated with TJs is ZAK (ZO-1 associated kinase), which was identified in MDCK cell lysates by its interaction with a GST fusion protein containing the third PDZ domain and the SH3 domain of human ZO-1 (Balda et al., 1996). The SH3 domain is sufficient for associating in vitro with ZAK kinase activity, but does not contain the Ser phosphorylation site, which is located C terminal to the SH3 domain. A kinase activity that phosphorylates the human GST fusion protein containing PDZ3 and SH3 domains can also be immunoprecipitated from MDCK lysate using anti-ZO-1 antibodies (Balda et al., 1996). The identity of this kinase and its subcellular localization have not been determined.

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11.6 PHOSPHORYLATION OF TJ PLAQUE PROTEINS In MDCK cells, ZO-1, ZO-2, ZO-3 (Anderson et al., 1988; Balda et al., 1993), and cingulin (Citi and Denisenko, 1995) are phosphorylated. Phosphorylation occurs on Ser residues in ZO-1 (Anderson et al., 1988) and cingulin (Citi and Denisenko, 1995). In addition, ZO-1, ZO-2, and ZO-3 can be phosphorylated on Tyr residues in cultured cells where tyrosine kinases has been activated or tyrosine phosphatase has been inhibited by a variety of experimental treatments (Staddon et al., 1995; Takeda et al., 1995; Van Itallie et al., 1995; Tsukamoto and Nigam, 1999). ZO-1 is also Tyr-phosphorylated in vivo in kidney glomeruli after perfusion with protamine sulfate, which induces the assembly of the glomerular slit filter (Kurihara et al., 1995). In another study, increased tyrosine phosphorylation of ZO-1 was reported during TJ assembly induced by the MEK1 inhibitor PD98059 in Ras-transformed MDCK cells (Chen et al., 2000). Although these studies indicate that Ser and Tyr phosphorylation of TJ plaque proteins may have important roles in the physiological or pathological modulation of TJ, direct evidence on the identity of in vivo phosphorylation sites and their role in protein function is missing.

11.7 THE ARCHITECTURE OF THE CYTOPLASMIC PLAQUE OF TJ Analysis of immunogold label density of specific TJ proteins can be used to reconstruct the spatial organization of the TJ plaque, within a margin of error that depends on the size of the antibody complex, the shape of the antigen, the physical localization of the epitopes on the antigen, and the plane of sectioning. In heart (Itoh et al., 1991) and epithelial cells (Stevenson et al., 1989), ZO-1 labeling is confined to a narrow area 0 to 15 nm from the membrane. In epithelial cells, cingulin labeling is localized at a mean distance from the TJ membrane of ~40 nm (Citi et al., 1988; 1989) to ~65 nm (Stevenson et al., 1989). The distribution of cingulin immunogold label density in intestinal epithelial cells indicates that cingulin is present in one layer within the TJ plaque (Figure 11.4B). The mean distance of 7H6-associated gold particles from TJ membrane in rat liver bile canaliculus fractions is ~41 nm (Zhong et al., 1993). Symplekin appears to be located at the interface between the plaque structure and the membrane proper (Keon et al., 1996). On the basis of these measurements, the TJ plaque could comprise an inner layer, containing (among others) ZO-1, symplekin (and presumably also ZO-2 and ZO-3, since they bind to ZO-1), and an outer layer, containing (among others) 7H6 and cingulin (Figure 11.4C). Actin microfilaments and spectrin probably extend throughout the two layers of the plaque and may further cross-link TJ proteins, since actin and spectrin interact with ZO-1 and ZO-2, actin and myosin interact with cingulin, and complexes containing both TJ plaque and cytoskeletal proteins can be isolated (see Section 11.4). It is unclear whether changes in the architecture of the TJ plaque occur in the physiological or pathophysiological modulation of TJ function. In animal models, the decrease in TJ barrier function occurring during inflammation does not correlate with changes in TJ ultrastructure or distribution of TJ plaque proteins (Cui et al.,

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1996; Lora et al., 1997). Conversely, an altered expression and localization of ZO-1 was described in rat hepatocytes during cholestasis induced by bile duct ligation (Fallon et al., 1993). In addition, several pharmacological treatments that disrupt TJ barrier function induce redistribution of TJ plaque proteins in cultured cells (Siliciano and Goodenough, 1988; Citi, 1992; Stevenson and Begg, 1994; Chen et al., 2000).

11.8 THE ROLE OF TJ CYTOPLASMIC PLAQUE PROTEINS IN TJ FUNCTION Although some of the components of the cytoplasmic plaque of TJ have been known for over a decade, little information is available on the role of any TJ cytoplasmic plaque protein in the barrier and fence functions of TJs. The functional properties of TJs in mutant cells and embryos lacking AF-6 have not been characterized (Zhadanov et al., 1999; Ikeda et al., 1999). At the ultrastructural level, junctions of mutant neuroectoderm were reported to possess basally displaced, short, electron-dense regions (Zhadanov et al., 1999). However, it is not clear whether these junctions still provide a permeability barrier or maintain the polarized distribution of protein and lipid components of the plasma membrane. Evidence suggesting a role of ZO-1 in regulating TJ permeability was recently reported using cultured epithelial cells (Balda and Matter, 2000; Reichert et al., 2000). MDCK-I cells stably transfected with a ZO-1 mutant construct containing only the three PDZ domains showed no transepithelial resistance, indicating that TJs were disrupted (Reichert et al., 2000). Conversely, cells transfected with fulllength human ZO-1 or with a mutant containing the PDZ, SH3, and GUK domains showed about a threefold increase in transepithelial resistance, suggesting an increased barrier function. In another study, MDCK-II cell clones stably transfected with full-length ZO-1 showed no change in transepithelial resistance, but about a threefold increase in paracellular permeability to [3H]mannitol (Balda and Matter, 2000), suggesting a reduced barrier function. The discrepancy between the results obtained by Reichert (Reichert et al., 2000) and Balda (Balda and Matter, 2000) is unclear, but could be due to the different strain of MDCK cells used. The increase in paracellular permeability induced by ZO-1 was reversed by cotransfection of ZO-1 with ZONAB, suggesting that ZO-1 and ZONAB functionally interact in the regulation of paracellular permeability (Balda and Matter, 2000).

11.9 NUCLEAR LOCALIZATION OF TJ PLAQUE PROTEINS: A ROLE IN REGULATION OF GENE EXPRESSION? Virtually all TJ plaque proteins contain putative nuclear localization signals (see Figure 11.3), and several TJ plaque proteins have been localized by immunofluorescence in the nucleus: ZO-1 (Gottardi et al., 1996), symplekin (Keon et al., 1996), cingulin (Citi and Cordenonsi, 1999), ZONAB (Balda and Matter, 2000), ASH1 (Nakamura et al., 2000), PAR-6 (Johansson et al., 2000). Detection of TJ plaque proteins in the nucleus depends on cell type, fixation, permeabilization protocols used for immunofluorescence, and the type of antibody (Keon et al., 1996). In nonepithelial cells, symplekin appears to be localized exclusively in the nucleus

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(Keon et al., 1996). The nuclear staining of ZO-1 was detected in subconfluent, but not confluent, cell cultures, at sites of wounding of an epithelial monolayer, and in cells along the outer tip of intestinal villi, suggesting that nuclear accumulation of ZO-1 is stimulated by loss or reduction in cell–cell contact (Gottardi et al., 1996). On the other hand, symplekin nuclear staining does not depend on the proliferative state of epithelial cells (Keon et al., 1996). The nuclear localization of TJ plaque proteins suggests that they may be implicated in regulation of gene expression. Transcription factors whose nuclear concentration is critical for activity may be exported into the cytoplasm and be recruited at TJs by TJ proteins. This could provide a mechanism to sequester transcription factors and thus regulate their activity. Another scenario could be that TJ plaque proteins, similarly to adherens junction proteins (Heasman et al., 1994; Behrens et al., 1996), are directly involved in triggering signals from the cell–cell contact area by translocating into the nucleus and functionally interact with transcription factors. Recent studies indicate that overexpression of ZO-1 mutants influences gene expression and cell differentiation (Balda and Matter, 2000; Reichert et al., 2000; Ryeom et al., 2000). When MDCKI cells are transfected with constructs encoding the N-terminal half of human ZO-1 (lacking the GUK and proline-rich region), they undergo an epithelial–mesenchymal transition, display a fibroblast-like morphology, are more tumorigenic, and show a repression of epithelial marker genes and an induction of mesenchymal marker genes (Reichert et al., 2000). These changes are associated with an activation of the β-catenin/Tcf/Lef signaling pathway, and are reversed by ectopic expression of the adenomatous polyposis coli (APC) protein (Reichert et al., 2000), suggesting a functional interaction between signaling pathways associated with tight and adherens-type junctions. In corneal epithelial cells, long-term stable expression of mutants of ZO-1 containing only an N-terminal fragment result in the partial transformation from an epithelial morphology to a mesenchymal cell type, with downregulation of occludin and upregulation of vimentin and smooth muscle actin (Ryeom et al., 2000). There is evidence that TJ plaque proteins bind to transcription factors. PAR-6 was identified by yeast 2-hybrid assay as a protein binding to the HTLV-1 oncoprotein Tax (Rousset et al., 1998). Furthermore, ZO-1 interacts in vitro and functionally in vivo with the transcription factor ZONAB, resulting in the regulation of expression of the ErbB2 gene (Balda and Matter, 2000). Whether other TJ plaque proteins are involved in epithelial–mesenchymal transitions or interaction with transcription factors remains unclear.

11.10 TJ PLAQUE PROTEINS AND DISEASE A decreased permeability barrier function of epithelia has been implicated in a number of pathological processes (see chapters on disease-related aspects of TJs). However, there are at present no reports demonstrating a specific role of any TJ plaque proteins in the pathogenesis of human disease. The vast majority of cancers originates from epithelial cells, and malignancy is often correlated with loss of the polarized epithelial phenotype. Is expression of TJ plaque proteins altered in cancer? Cingulin expression levels in undifferentiated colon adenocarcinomas are high, and

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its localization is aberrant (Citi et al., 1991). 7H6 expression is decreased in rat liver tumors (Zhong et al., 1994b). ZO-1 staining is decreased in undifferentiated breast adenocarcinomas, and reduction in ZO-1 staining was correlated with reduced E-cadherin staining (Hoover et al., 1998). Pancreatic duct carcinoma cells do not express a splicing variant of ZO-2, which lacks a 23 amino acid motif at the N-terminal end, whereas the longer form is detected in both normal and cancer cells (Chlenski et al., 1999; 2000). These observations suggest that up- or downregulation of TJ plaque protein expression and specific isoform expression may be related to control of cell growth and cancer progression. Interestingly, null mutants of the dlg gene in D. melanogaster show loss of septate junctions and neoplastic growth of cells lining the imaginal disk (Woods and Bryant, 1991), suggesting that in this model system mutation of a MAGUK protein might play a causative role.

11.11 CONCLUDING REMARKS The cytoplasmic plaque of TJs comprises over a dozen proteins that have now been characterized at the molecular level, and several others that await further characterization. By interacting with each other, with TJ membrane proteins, and with cytoskeletal proteins, TJ plaque proteins assemble and maintain a highly organized structure at the interface between the membrane domain of TJs and the cytoplasm. Some TJ plaque proteins are homologous with invertebrate proteins involved in control of cell fate, and can be localized in the nucleus and interact with transcription factors. In addition, TJ plaque proteins appear to act as targets and effectors of different signaling pathways. Thus, is the TJ plaque a specific signaling center that clusters proteins involved in control of cell growth and differentiation? How are TJ plaque proteins involved in modulating TJ barrier function? The challenge is now to address these questions, and to clarify the molecular signals and hierarchies of interactions leading to the assembly of the TJ plaque.

ACKNOWLEDGMENTS The author is grateful to Marylin Farquhar for generously providing electron micrographs, Benjamin Geiger for data about cingulin immunogold label distribution, Nicholas Roggli for graphic assistance, Fabio D’Atri and David Shore for comments, and the Swiss National Fonds, State of Geneva, MURST, and CNR for support.

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12

Organization and Regulation of the Tight Junction by the Actin–Myosin Cytoskeleton Alan S. Fanning

CONTENTS 12.1 Introduction .................................................................................................266 12.2 Organization of F-Actin at the Tight Junction of Epithelial Cells ............266 12.3 The Cytoskeleton and Paracellular Permeability: Effects of Physiological, Pharmacological, and Pathogenic Agents ..........................267 12.3.1 Cytochalasins .................................................................................267 12.3.2 Pharmacological Agents ................................................................269 12.3.3 Bacterial Pathogens .......................................................................269 12.3.4 Small GTP-Binding Proteins.........................................................270 12.3.5 Physiological Agents: Glucose ......................................................270 12.4 Possible Mechanisms of Cytoskeletal Action on Tight Junction Organizaton and Function...........................................................................271 12.5 The Actin Cytoskeleton Interacts Directly with Molecular Components of the Tight Junction...................................................................................272 12.6 Functional Significance of Direct Interactions between the Cytoskeleton and TJ Proteins: De Novo Assembly, Reorganization, and Acute Regulation of Paracellular Permeability ...................................274 12.6.1 Regulation of Paracellular Permeability .......................................274 12.6.2 De Novo Assembly of Tight Junctions..........................................275 12.6.3 Clues from Invertebrate Model Systems.......................................277 12.7 Concluding Remarks...................................................................................277 Acknowledgments..................................................................................................279 References..............................................................................................................279

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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12.1 INTRODUCTION The cytoskeleton is the fundamental determinant of cell shape, cytosolic organization, and motility. The actin cytoskeleton in particular has been associated with organization of plasma membrane–associated structures. An extensive literature has accumulated that implicates the actin cytoskeleton in regulation of tight junctions (TJs) in response to pharmacological and pathological stimuli. The purpose of this chapter is less to review this literature, than to examine the possible molecular basis of these phenomena. It has become increasingly clear that many of the molecular components of TJs are intimately associated with F-actin or other actin-binding proteins. This chapter discusses the nature of these interactions, and what is currently known about the functional role of these interactions during assembly and regulation of the paracellular barrier.

12.2 ORGANIZATION OF F-ACTIN AT THE TIGHT JUNCTION OF EPITHELIAL CELLS The actin cytoskeleton of polarized epithelial cells is organized into distinct apical, lateral, and basal plasma membrane domains, which are functionally and structurally distinct (Mooseker, 1985). For example, the apical membrane includes microvilli containing bundled arrays of parallel microfilaments that extend into the cytoplasm and terminate in a fibrous array of F-actin known as the terminal web (Drenckhahn and Groschel-Stewart, 1980). The basal and lateral membranes are associated with a dense cortical cytoskeleton composed of short cross-linked actin filaments and associated proteins like spectrin, protein 4.1, and ankyrin, which cross-link actin filaments and anchor them to the plasma membrane (Morrow et al., 1997). In cultured cells the basal membrane is also associated with stress fibers — contractile bundles of actin and nonmuscle myosin which terminate at the plasma membrane at integrin-based adhesion sites known as focal contacts. Finally, the lateral surface is associated with several sites of cell–cell contact (Hirokawa and Tilney, 1982). The most prominent of these is the apical junctional complex, which includes both TJs and the adherens junctions. The TJ creates a seal within the paracellular space, and delineates the apical and basal-lateral domains by forming a barrier to the diffusion of transmembrane proteins and lipids (see Chapters 8 and 14). The adherens junction, which in vertebrates is positioned more basal relative to the TJ in the lateral membrane, forms an adhesive structure of cadherin-based cell–cell contacts. Electron microscopy studies of the apical junctional complex reveal that both TJs and adherens junctions are associated with microfilaments (Hirokawa and Tilney, 1982; Hirokawa et al., 1983; Drenckhahn and Dermietzel, 1988). Adjacent to the adherens junction is a highly ordered band of antiparallel actin filaments associated with the conventional myosin II, which is referred to as the perijunctional actomyosin ring (PAMR). This structure has been demonstrated to have contractile activity both in vitro and in vivo (Keller and Mooseker, 1982), and it has been proposed that the tensile strength generated by the contraction of this ring is required for the integrity of epithelial sheets. In contrast, the microfilaments associated with TJs are much fewer in number and less organized (Hirokawa and Tilney, 1982; Madara, 1987). However, analysis of detergent-extracted membranes decorated with myosin subfragment-1

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FIGURE 12.1 Electron micrograph of a detergent-extracted preparation of guinea pig ileum (left) and corresponding sketch (right). Actin filaments were identified by decoration with myosin subfragment-1, which gives them a barbed appearance. Note that the decorated filaments appear to terminate at dense plaques associated with membrane contact sites, or “kisses.” (From Madara, J. L., Am. J. Physiol., 253, C171, 1987. With permission.)

suggests that microfilaments extend from the terminal web and make direct contact with the dense plaque material at sites of membrane contacts, or “kisses,” within the TJ (Figure 12.1; Madara, 1987). These observations imply that the actin cytoskeleton makes direct contact with elements of the TJ.

12.3 THE CYTOSKELETON AND PARACELLULAR PERMEABILITY: EFFECTS OF PHYSIOLOGICAL, PHARMACOLOGICAL, AND PATHOGENIC AGENTS 12.3.1 CYTOCHALASINS Early investigators were quick to note the functional implication of direct interaction between the cortical cytoskeleton and the TJ — that the actin cytoskeleton might regulate some aspect of TJ assembly or permeability. Initial support for this hypothesis

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TABLE 12.1 The Effects of Different Agents on Paracellular Permeability and the Actin Cytoskeleton Treatment Cytochalasin B or cytochalasin D

Cellular Effects Regulate actin dynamics

Cytoskeletal Effects F-actin condensation filament reorganization ↑ MLC phosphorylation

Tight Junction Effects ↓TER/↑flux f.f. fibril disorganization Altered ZO-1 distribution

Phalloidin

Stabilize F-actin

↑ number actin filaments

↑TER f.f. fibril reorganization

Plant cytokinins (kinetin, zeatin)

Pleiotropic

F-actin condensation filament reorganization

↑TER f.f. fibril disorganization

D-Glucose

Activate Na+-glucose transporter (SGLT1)

F-actin condensation at PAMR MLC phosphorylation

↓TER/↑flux f.f. fibril reorganization

PMA/dIC8

PKC agonists

↑ F-actin at PAMR MLCK phosphorylation MLC dephosphorylation

↑/↓ TER Altered dynamics of junction assembly and disassembly

Antimycin A + 2-deoxyglucose

ATP depletion

F-actin disruption at PAMR

↓TER f.f. fibril disorganization

Constitutively active MLCK

Activate actomyosin contraction

No change F-actin distribution

↓TER/↑flux

TNF-α + IFN-γ

Pleiotropic

Reorganization of F-actin into punctate cell–cell contacts

↓TER Redistribution of ZO-1 into punctate cell–cell contacts

Ethanol

Pleiotropic

F-actin reorganization Stimulation of MLCK

↓TER/↑flux Retraction of ZO-1

Clostridium difficile toxin A/B

Glucosylation (inhibition) of Rho GTPase

↓F-actin at PAMR and/or F-actin condensation

↓TER/↑flux

C3 transferase

ADP ribosylation (inactivation) of Rho GTPase

Disruption F-actin at PAMR

↓TER/↑flux Displacement of ZO-1

Dominant active (DA) or negative (DN) Rac and Rho GTPases

Regulate F-actin dynamics

DA — ↑ accumulation of F-actin at PAMR DN — disruption F-actin

DA — altered ZO-1 and occludin distribution Altered f.f. fibril morphology DA/DN — ↓TER/↑flux

Abbreviations: f.f., freeze fracture; TER, transepithelial electrical resistance; F-actin, filamentous actin; PAMR, perijunctional actomyosin ring; MLCK, myosin light-chain kinase; MLC, myosin light chain. See text for references.

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came from an analysis of the effects of actin-disrupting agents on paracellular permeability (Table 12.1). Treatment of tissue explants or cultured cells with cytochalasin D or the plant cytokinin zeatin resulted in a marked decrease in transepithelial electrical resistance (TER) and a corresponding increase the flux of radiolabeled tracers through the paracellular space (Meza et al., 1980; Bentzel et al., 1980; Madara et al., 1986; Stevenson and Begg, 1994). This was accompanied by disruption of freeze-fracture fibrils and changes cell shape and the distribution of F-actin. Freeze-fracture fibrils became disorganized, with numerous gaps and strands that appeared disconnected from the fibril network (Bentzel et al., 1980; Madara et al., 1987). Actin microfilaments retracted from the plasma membrane (Meza et al., 1980; Madara et al., 1988) or accumulated in condensations characteristic of actin–myosin contraction (Madara et al., 1986; 1987), and the apical plasma membrane became rounded (Bentzel et al., 1980; Madara et al., 1986). In marked contrast to these observations, microtubule-disrupting agents had little effect on junction structure or paracellular permeability (Gonzalez-Mariscal et al., 1985).

12.3.2 PHARMACOLOGICAL AGENTS Subsequent investigations with more physiologically relevant agents have demonstrated a similar correlation between cytoskeletal organization and TJ properties (see Table 12.1). For example, several pharmacological agents that alter paracellular permeability also invoke changes in cytoskeletal organization. Agents that deplete intracellular ATP, such as antimycin A and 2-deoxyglucose, cause disruption of the PAMR, disorganization of freeze-fracture fibrils, and decreases in TER (Bacallao et al., 1994; Tsukamoto and Nigam, 1997). Protein kinase C (PKC) agonists can either increase or decrease paracellular permeability (Mullin and O’Brien, 1986; Hecht et al., 1994; Rosson et al., 1997; Turner et al., 1999; Clarke et al., 2000), depending on the cell line used (Ellis et al., 1992). These agents have also been demonstrated to promote accumulation of F-actin at the PAMR, disorganization of cells within the monolayer (Hecht et al., 1994), and altered organization of proteins within TJs (Clarke et al., 2000) of confluent monolayers. Perhaps more significantly, these agents can dramatically alter the dynamics of junction assembly and disassembly (Balda et al., 1991; 1993; Citi, 1998; Clarke et al., 2000). PKC agonists accelerate the development of TER and stimulate the reorganization of actin filaments, TJ proteins, and freeze-fracture fibrils into organized junctions in cultured cells following extracellular calcium depletion and readdition; the so-called calciumswitch experiment (Balda et al., 1991; 1993). Other factors have similar effects on junction assembly and TER. Dual application of tumor necrosis factor-α and interferon-γ to endothelial cells promotes decreases in TER and reorganization of F-actin and TJ proteins into punctate–cell–cell contacts (Blum et al., 1997).

12.3.3 BACTERIAL PATHOGENS Bacterial pathogens and their associated toxins can also have dramatic effects on both cytoskeletal organization and paracellular permeability (see Chapter 23 by Hecht). Reductions in TER and increased mannitol flux in cultured cells treated with Clostridium difficile toxins A or B are accompanied by reorganization of F-actin in the PAMR

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into discrete plaques (Hecht et al., 1988; 1992). The Vibrio cholerae zonula occludens toxin (ZOT) induces a marked reduction TER, reduced complexity of TJ fibrils (Fasano et al., 1991), and a redistribution of F-actin. A similar correlation between F-actin disruption and altered permeability has also been observed with the 20-kDa toxin of Bacteriodes fragilis (also known as BFT or fragilysin) (Koshy et al., 1996; Obiso et al., 1997), the Yersinia pseudotuberculosis outer membrane protein YopE (Tafazoli et al., 2000), and with infection of certain strains of enteropathogenic Escherichia coli (EPEC) (Spitz et al., 1995; Philpott et al., 1996; Peiffer et al., 2000). In these latter cases the increased paracellular permeability and F-actin rearrangements are also associated with redistribution of several TJ proteins, including ZO-1 and occludin.

12.3.4 SMALL GTP-BINDING PROTEINS More recently, investigators have identified several key regulators of cytoskeletal dynamics that can also regulate the assembly and permeability of TJs; the Rac and Rho GTPases. These proteins interact with cell signaling pathways downstream of several transmembrane receptors and regulate the assembly of F-actin at the plasma membrane. Overexpression of dominant active or dominant negative forms of Rac or Rho result in alterations in paracellular permeability that are accompanied by reorganization of F-actin at the PAMR, disorganization of freeze-fracture fibrils, and altered distribution of the TJ proteins ZO-1 and occludin (Takaishi et al., 1997; Jou et al., 1998). Inhibitors of Rho, such as the Clostridium botulinum toxin C3 transferase, have similar effects (Nusrat et al., 1995). In fact, there is evidence that several of the bacterial toxins that affect junction permeability work through the small GTPbinding protein Rho. For example, the Clostridium toxins glucosylate Rho, and prevent binding to effector proteins. The YopE protein of Yersinia acts as RhoGAP, stimulating GTPase activity of Rho, which results in its inactivation (Aktories et al., 2000). These results suggest that cellular signaling pathways that regulate F-actin assembly may also regulate the assembly and permeability of TJs. However, since both Rac and Rho also affect cell adhesion and general cytoskeletal organization, it is not yet clear how directly these proteins affect TJs.

12.3.5 PHYSIOLOGICAL AGENTS: GLUCOSE In several instances changes in paracellular permeability are also accompanied by changes in the cytoskeleton characteristic of actomyosin contraction of the PAMR, suggesting a more direct role of cytoskeleton in regulation paracellular permeability. Perhaps the most convincing evidence for regulation of TJ permeability by actomyosin contraction comes from examination of glucose uptake in the intestine, which is discussed in greater detail in Chapter 15 by Turner and Madara. Briefly, activation of the sodium glucose transporter in tissue explants (Madara and Pappenheimer, 1987; Atisook et al., 1990) or cultured cell systems (Turner et al., 1996; 1997) leads to increased flux of glucose analogues and other small solutes through the paracellular space. These changes are accompanied by condensation of F-actin within the PAMR, ultrastructural changes within the TJ, and changes in cell shape characteristic of contraction within the PAMR. Significantly, these changes are paralleled by phosphorylation of the myosin light chain (MLC), which is required to activate actomyosin

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contraction in nonmuscle cells (Turner et al., 1997). In addition, the changes in paracellular permeability and junction structure in response to glucose are inhibited by the MLC kinase (MLCK) inhibitors ML-7 and ML-9 (Turner et al., 1997), suggesting that there is a direct relationship between MLC phosphorylation by MLCK, actomyosin contraction, and changes in paracellular permeability. In support of this observation, Hecht et al. (1996) have found that overexpression of a constitutively activated MLCK in MDCK cells can reduce TER to less than 10% of normal values. Notably, a similar correlation between increased paracellular permeability, cytoskeletal reorganization, MLCK activation, and/or MLC phosphorylation has also been observed in cultured cells treated with ethanol (Ma et al., 1999), cytochalasin D (Ma et al., 2000), PKC agonists (Turner et al., 1999), or in cells infected with EPEC (Yuhan et al., 1997). Although the physiological relevance of passive glucose transport through the paracellular space in vivo is still a matter of some debate (Diamond 1995), these observations nevertheless suggest that actomyosin contraction within the PAMR may be a general mechanism that regulates paracellular permeability.

12.4 POSSIBLE MECHANISMS OF CYTOSKELETAL ACTION ON TIGHT JUNCTION ORGANIZATON AND FUNCTION The major drawback of the experimental manipulations described above is that they are relatively indirect, which makes mechanistic interpretations particularly difficult. However, several general interpretations of these observations have been offered. The correlation between F-actin rearrangement and disorganization of junction structure in the preceding examples implies that loss of cytoskeletal integrity causes disruption of TJ structure and function. This would suggest that the actin cytoskeleton has an organizational role in the TJ. One possibility is that the cytoskeleton is directly involved in the assembly and/or maintenance of junction structure, perhaps through interactions with junction-associated proteins (discussed below). Thus, the changes in junction structure and permeability in treated cells reflect a disruption of the normal cytoskeletal role in junction assembly. The other possibility is that effects on TJs are secondary to more global changes in cell–cell adhesion or overall cellular organization caused by cytoskeletal reorganization. Resolution of this issue awaits a better understanding of how the cytoskeleton interacts with molecular components of the TJ. In either case, the changes in paracellular permeability in this scenario are secondary to disruption of structure. However, the actin condensations, cell shape changes, and activation of MLC and MLCK observed in several of these reports suggest that actomyosin contraction within the PAMR might underlie the changes in paracellular permeability. This implies that the actin cytoskeleton has an active and much more direct role regulating the opening and closing of the paracellular space in epithelia. How actomyosin contraction might regulate TJ permeability is currently only a matter for speculation. Turner and Madara (Chapter 15) have proposed that contraction of the PAMR creates physical distortions of plasma membranes, leading to openings within the paracellular space. However, given the recent understanding of the extensive interaction between junctional and cytoskeletal proteins, it is also possible that the cytoskeleton

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may have more direct effects on the junctional proteins themselves, perhaps affecting the interaction between transmembrane components of the TJ. These possibilities are discussed further below.

12.5 THE ACTIN CYTOSKELETON INTERACTS DIRECTLY WITH MOLECULAR COMPONENTS OF THE TIGHT JUNCTION The last decade has seen a rapid advancement in knowledge of the molecular composition of the TJ (see Chapter 10, by Mitic and Van Itallie, Chapter 11, by Citi). One of the more significant revelations arising from these studies has been the recognition that many, if not all, of these proteins are intimately associated with the actin cytoskeleton. In fact, recent studies have demonstrated that many of the components of the TJ bind directly to F-actin in vitro (Table 12.2). These include members of the MAGUK family of membrane-associated scaffolding proteins, ZO1, ZO-2, and ZO-3 (Itoh et al., 1997; Fanning et al., 1998; Wittchen et al., 1999), as well as the 140- to 160-kDa phosphoprotein cingulin (Cordenonsi et al., 1999). Furthermore, at least one transmembrane component of the TJ, occludin, may also bind directly to F-actin (Wittchen et al., 1999). The Ras tyrosine kinase substrate l-afadin has also been demonstrated to bind to F-actin (Mandai et al., 1997). However, it is currently not clear whether l-afadin, like the shorter splice form AF-6/safadin (Yamamoto et al., 1997), is localized to TJs. Many of these proteins also have additional connections to the actin cytoskeleton through interactions with other F-actin binding and regulatory proteins (Table 12.2

TABLE 12.2 Molecular Interactions between Tight Junction Proteins and the Actin Cytoskeleton Indirect Cytoskeletal Interactions Via Protein ZO-1

Binds F-Actin Direct and indirect

ZO-2

Direct and indirect

ZO-3 Cingulin Occludin AF-6/s-afadin

Direct Direct and indirect Direct and indirect Directa and indirect

F-Actin-Binding Proteins α-Catenin Cortactin Fodrin Protein 4.1R α-Catenin Protein 4.1R Nonmuscle myosin-II Protein 4.1R Profilin

Tight Junction Proteins ZO-2/3 Cingulin AF-6 ZO-1 ZO-1 ZO-1/2/3 ZO-1/2/3 ZO-1

a Only the longer isoform, l-afadin, binds directly to F-actin. l-Afadin is a component of cadherinbased cell–cell contacts, and its presence in TJs is still a matter of debate.

See text for references.

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FIGURE 12.2 Hypothetical organization of cortical cytoskeleton at the TJ. The figure is meant to demonstrate the possible contacts between actin and proteins of the TJ, as well as those between different TJ proteins. The exact nature of the complexes formed by these proteins is currently unknown. (Adapted from Mitic et al., 2000. With permission.)

and Figure 12.2). This is particularly evident among the ZO MAGUK proteins. For example, both ZO-1 and ZO-2 bind to a novel isoform of the erythrocyte protein 4.1 that localizes to TJs (Mattagajasingh et al., 2000). In erythrocytes, protein 4.1R links transmembrane proteins like glycophorin-C or band 3 to cytoskeletal proteins like spectrin/fodrin (Morrow et al., 1997). Interestingly, ZO-1 has also been demonstrated to coimmunoprecipitate with fodrin, an actin cross-linking protein of the cortical cytoskeleton that is required for cell shape and plasma membrane stability (Itoh et al., 1993). Surprisingly, the transmembrane protein occludin has also been demonstrated to bind protein 4.1R (Mattagajasingh et al., 2000). Taken together, these observations reinforce ultrastructural evidence that ZO proteins are closely associated with the cortical cytoskeleton. The ZO proteins also bind to cytoskeletal proteins not normally associated with TJs. Both ZO-1 and ZO-2 bind to α-catenin, a component of cadherin-based cell–cell contacts (Itoh et al., 1997; 1999). In addition, ZO-1 binds cortactin (Katsube et al., 1998), a prominent tyrosine kinase substrate associated with lamellae and some cell–cell junctions (Wu and Parsons, 1993). α-Catenin appears to link components of adherens junction to the actin cytoskeleton, and is required for the formation of stable cell–cell adhesions (Provost and Rimm, 1999). In contrast, cortactin is associated with dynamic arrays of F-actin in cultured cells (Wu and Parsons, 1993), and

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is thought to be involved in regulating the de novo assembly of F-actin at the plasma membrane through interactions with proteins like the F-actin nucleating protein ARP 2/3 (Weed et al., 2000). Both α-catenin and cortactin can cross-link microfilaments in vitro (Rimm et al., 1995; Huang et al., 1997). The functional role of these protein interactions in TJ physiology is at present unclear. It is possible that they are involved early in TJ biogenesis, but it is also possible that they reflect roles for proteins like ZO-1 and ZO-2 outside of TJs. Several other TJ proteins bind to cytoskeletal proteins that have established roles regulating actin dynamics. For example, cingulin binds to nonmuscle myosin II, the predominant myosin mechanoenzyme of the PAMR and terminal web (Cordenonsi et al., 1999). AF-6/afadin interacts with profilin, a protein that promotes F-actin assembly at the plasma membrane by regulating the availability of actin monomers (Boettner et al., 2000). These interactions raise the intriguing possibility that junction proteins may actually regulate the assembly of F-actin at the plasma membrane in the vicinity of the TJ through interactions with proteins like cortactin or profilin. It is notable that many TJ proteins that bind F-actin also bind to each other (Figure 12.2). This is particularly evident among the ZO proteins. For example, ZO-1 dimerizes in vivo with either ZO-2 or ZO-3 (Fanning et al., 1998; Wittchen et al., 1999), and has also been demonstrated to bind directly to cingulin, occludin, and AF-6 (Furuse et al., 1994; Yamamoto et al., 1997; Cordenonsi et al., 1999). The other ZO proteins, ZO-2 and ZO-3, also bind cingulin and occludin (Itoh et al., 1999; Cordenonsi et al., 1999; Wittchen et al., 1999). This implies that TJ proteins form an intricate and highly interconnected network with multiple contacts to the actin cytoskeleton (see Figure 12.2).

12.6 FUNCTIONAL SIGNIFICANCE OF DIRECT INTERACTIONS BETWEEN THE CYTOSKELETON AND TJ PROTEINS: DE NOVO ASSEMBLY, REORGANIZATION, AND ACUTE REGULATION OF PARACELLULAR PERMEABILITY 12.6.1 REGULATION

OF

PARACELLULAR PERMEABILITY

The demonstration that many of the molecular components of TJs interact directly with F-actin has reinforced the hypothesis that the actin cytoskeleton is a major determinant of junction organization and regulation. However, although numerous contacts between the TJ and the cytoskeleton have been identified, the functional significance of these interactions is still far from clear. One hypothesis, an extension of those outlined above, is that the connections between TJ proteins and the cytoskeleton are required for acute regulation of paracellular permeability in response to physiological stimuli. Specifically, these connections may transduce tension generated by actomyosin contraction to the cortical cytoskeleton at TJs, resulting in transient opening of the paracellular space. Alternatively, they may transduce tension directly to the transmembrane sealing proteins claudin or occludin. At present, there is little evidence to support or deny this model. Experiments that have examined the

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effects of ectopic expression of TJ proteins that bind to F-actin have provided few clues. Although expression of a ZO-1 transgene was noted to have a subtle effect on the paracellular flux of macromolecules in one report (Balda and Matter, 2000), the mechanism underlying this change is unknown. In another study, overexpression of a ZO-1 construct lacking the actin-binding domain was demonstrated to have little effect on TER or cytoskeletal organization in MDCK cells (Fanning et al., 1998; and unpublished results). However, a major caveat to interpretation of experiments of this type is that these cells also express the endogenous ZO-1, as well as ZO-2 and ZO-3. These experiments point out the need for experimental systems, such as mouse knockouts, lacking endogenous ZO-1 to address appropriately questions about the physiological relevance of protein–protein interactions.

12.6.2 DE NOVO ASSEMBLY

OF

TIGHT JUNCTIONS

Another working hypothesis is that F-actin linkages are required for de novo assembly of proteins into the TJ. TJ proteins like ZO-1, occludin, and cingulin are intimately associated with F-actin during junction assembly in early mouse and Xenopus embryogenesis (Fleming et al., 1989; Fesenko et al., 2000; reviewed in Fleming et al., 2000). Similar associations have been documented during the normal assembly of junctions in cultured cells (Yonemura et al., 1995; Howarth and Stevenson, 1995; Ando-Akatsuka et al., 1999) and during junction assembly following a calcium switch (Silicano and Goodenough, 1988; Citi, 1998). Disruption of F-actin with cytochalasin D during early mouse embryogenesis prevents assembly of TJ proteins like ZO-1 into the apical junctional complex (Fleming et al., 1989). Similarly, treatment of cultured cells with cytochalasin D during calcium-triggered junction assembly (calcium switch) inhibits the development of TER (Meza et al., 1980) and the assembly of proteins into the TJ. Redistribution of ZO-1 in endothelial cells following TNF treatment is also sensitive to cytochalasin D (Blum et al., 1997). These observations support a role for the actin cytoskeleton in the recruitment or assembly of proteins into the TJ. However, it may be more likely that contacts between F-actin and TJ proteins maintain TJ structure. Actin may anchor TJ proteins at plasma membrane or provide a “glue” that holds the assembled complex together. Treatment of cultured cells with cytochalasin D results in retraction of F-actin and TJ proteins like ZO-1 and occludin into punctate aggregates (Takaishi et al., 1997), suggesting that association of TJ proteins with plasma membrane requires F-actin contacts. A similar retraction of F-actin and junction proteins has been observed following chelation of extracellular calcium (Meza et al., 1980; Silicano and Goodenough, 1988; Citi, 1998). Interestingly, cytochalasin D markedly inhibits the drop in TER and the redistribution of ZO-1 following calcium chelation (Takaishi et al., 1997), suggesting that the localization of ZO-1 within these cells is dependent on F-actin. However, this latter conclusion is probably too simplistic, since many of the TJ proteins examined (ZO-1, occludin, and AF-6) have been demonstrated to localize to the TJ even when actin-binding sites are deleted (Balda et al., 1996; Chen et al., 1997; Yamamoto et al., 1997; Fanning et al., 1998). This is perhaps best illustrated by ZO-1. Actin cosedimentation assays with ZO-1 transgenes have localized the

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FIGURE 12.3 Localization of myc-tagged ZO-1 transgenes in MDCK cells. Epitope-tagged fragments encoding the N-terminal half (z1-876), C-terminal half (del 67-1033), or full-length ZO-1 (Zo1myc) were introduced into cultured MDCK cells by calcium phosphate precipitation. Stable cell lines expressing these transgenes were fixed and stained with rhodamine phalloidin to visualize F-actin, and with antibodies against the c-myc epitope to localize the ZO-1 transgene. Note that a myc-tagged construct encoding the N-terminal half of ZO-1 (z1876), like the full-length protein ZO1myc, will still target to the apical junctional complex, even though it lacks the binding sites for actin, protein 4.1, and cortactin. The C-terminal half ZO-1 (del 67-1033) localizes to F-actin. (From Fanning, A. S. et al., J. Biol. Chem., 273, 29745, 1998. With permission.)

actin-binding site to the C-terminal half of ZO-1 (Itoh et al., 1997; Fanning et al., 1998). Interestingly, this region also contains the binding sites for cortactin (Katsube et al., 1998) and protein 4.1 (Mattagajasingh et al., 2000). Consistent with this observation, it was found that an epitope-tagged transgene encoding the C-terminal half of ZO-1 (del 67-1033) colocalizes with actin filaments in either epithelial and nonepithelial cells (Figure 12.3) (Itoh et al., 1997; Fanning et al., 1998). However, a construct encoding the N-terminal half of ZO-1 (z1-876), like the full-length protein ZO1myc, will still target to the apical junctional complex, even though it lacks the binding sites for actin, protein 4.1, and cortactin (Figure 12.3). These

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observations indicate that direct binding to F-actin is not required for assembly of ZO-1 into preformed junctions. However, because proteins like ZO-1 are assumed to exist in a large macromolecular complex with other TJ proteins, many of which bind F-actin, it is likely that interactions between ZO-1 and other TJ proteins can mediate localization in the absence of direct cytoskeletal contacts. It has also been demonstrated that changes in F-actin distribution and paracellular permeability in cultured cell monolayers do not always correlate with changes in the distribution of TJ proteins. For example, the F-actin-disrupting drug mycalolide B seems to have little effect on the distribution of ZO-1 or the organization of freezefracture fibrils, although both F-actin and paracellular permeability are markedly disrupted (Takakuwa et al., 2000). In a separate example, the changes in TER and actin organization associated with ATP depletion precede changes in ZO-1 localization by 60 min (Bacallao et al., 1994). Observations like these underscore the complexity of the mechanisms that regulate junction assembly.

12.6.3 CLUES

FROM INVERTEBRATE

MODEL SYSTEMS

The recent identification of the invertebrate homologues of ZO-1 and AF-6/afadin, known as polychaetoid and canoe, may provide some insight into another possible role for the cytoskeleton. These proteins, like their vertebrate counterparts, bind to each other and localize to the apical junctional complex of epithelial cells (Takahashi et al., 1998). Normally, in response to cellular signals that arise during embryogenesis, epithelial cells within the dorsal epidermis undergo rapid migration and dramatic shape changes that are associated with changes in actin cytoskeleton and cell–cell junctions. In hypomorphic double mutants of canoe and polychaetoid, both migration and cell shape changes are attenuated (Takahashi et al., 1998). These results were interpreted to suggest that the polychaetoid/canoe complex regulates the actin cytoskeleton during embryogenesis. The vertebrate homologue AF-6/afadin appears to have a similar role during mouse embryogenesis. Mouse embryos with a targeted disruption of the AF-6/afadin gene initially show normal junction assembly early in embryogenesis. It is not until gastrulation, when cell–cell junctions normally undergo a dramatic reorganization, that TJ structure begins to break down (Zhadanov et al., 1999; Ideda et al., 1999). Thus, AF-6/afadin does not appear to be required for de novo assembly of junctions. Instead, these observations suggest that interaction between cytoskeleton and proteins like AF-6 or ZO-1 are required for the dynamic reorganization at cell–cell junctions in response to cellular signaling cascades. It is interesting to speculate that interactions between the cytoskeleton and TJ may also be involved during other transient events that involve junction reorganization, such as leukocyte transmigration, wound repair, or apoptosis.

12.7 CONCLUDING REMARKS From the examples presented in this chapter, it is clear that cytoskeletal integrity is required for physiological regulation of TJs. Furthermore, it has also become apparent that the molecular components of the TJ are intimately associated with the cortical cytoskeleton through both direct and indirect contacts. However, it is still

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not clear how directly the cytoskeleton can influence junction assembly and paracellular permeability. Resolution of this issue will require a better understanding of how these proteins interact with F-actin. This includes the precise resolution of the actin-binding sites, as well as identification of the mechanisms that can regulate binding in vivo. Furthermore, it will require the development of better experimental systems that permit direct examination of the functional relevance of these interactions. One obvious, albeit ambitious, approach is to insert junction proteins lacking actin-binding motifs into cells in which the endogenous proteins have been eliminated. This could be accomplished through the use of cell systems derived from targeted gene deletion in mice, although to date only targeted disruption of occludin has been reported. Another approach is to utilize the more tractable invertebrate model systems. Homologues of the ZO proteins have now been identified in both Drosophila and Caenorhabditis elegans. These systems allow relatively easy genetic manipulation, and would also permit the introduction of modified junction proteins lacking actin-binding domains in otherwise null backgrounds. The drawback of invertebrate systems is that the permeability properties of invertebrate junctions are at present poorly understood, and it is not clear to what extent the functional roles of these junctions are analogous to those of vertebrate TJs. It is also important to point out that cytoskeletal regulation is likely not the only mechanism by which paracellular permeability is regulated, since in many cases factors that regulate paracellular permeability are not necessarily accompanied by visible changes in the distribution of cortical actin, cortical contraction, or changes in the distribution of other TJ proteins. For example, the addition of dexamethasone to cultured cells results in a dramatic change in TER without visible changes in the distribution of F-actin or ZO-1 (Zettl et al., 1992). Chelation of intracellular calcium slows the development of TER and assembly of ZO-1 following calcium switch, but it does not appear to affect the distribution of actin (Stuart et al., 1994). Bacallao et al. (1994) have found that the decrease in TER following ATP depletion precedes any observable structural changes in F-actin or redistribution of ZO-1. Finally, addition of sodium butyrate to MDCK cells can cause a two- to threefold change in TER, but has no discernible effect on cytoskeletal organization, freeze-fracture fibril morphology, or distribution of ZO-1 and occludin (A. S. Fanning, E. E. Schneeberger, and J. M. Anderson, unpublished observations). The nature of these other mechanisms is currently a matter of great interest. Recent attention has been focused on the role of cellular signaling pathways and post-translational modification of TJ proteins as one potential mechanism. Such modifications might regulate the ability of TJ proteins to assemble into a macromolecular complex, or perhaps directly alter the sealing properties of transmembrane proteins. However, post-translational modification could also regulate associations with the cytoskeleton. Although many tight junction proteins are post-translationally modified (see Chapter 17, by Balda et al.), it is currently not clear how this affects junctional permeability, assembly, or interaction with cytoskeleton. In some cases, however, the cytoskeleton appears to be required for post-translational modification of junction proteins (Van Itallie et al., 1995; Tsukamoto and Nigam, 1999). For example, it has been observed that the tyrosine phosphorylation of ZO-1 that occurs in epithelial cells treated with epidermal growth factor is eliminated if cells are

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treated with cytochalasin D (Van Itallie et al., 1995). Thus, it is likely that elaboration of the signaling pathways that act on TJs will provide further insight into the role of the cytoskeleton at the TJs. In conclusion, the identification of the molecular components of the TJ, and the identification of the molecular contacts between these proteins and F-actin, has now made it possible to test directly the functional role of the cytoskeleton during the assembly and regulation of TJs.

ACKNOWLEDGMENTS The author thanks the members of the Anderson Laboratory, past and present, for their thoughtful discussions and constant support.

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13

Developmental Assembly of the Tight Junction Tom P. Fleming, Bhavwanti Sheth, Fay Thomas, Irina Fesenko, and Judith Eckert

CONTENTS 13.1 Introduction .................................................................................................285 13.2 The Trophectoderm Model of Tight Junction Assembly ...........................286 13.2.1 Molecular Maturation of the Tight Junction during Trophectoderm Differentiation ......................................................287 13.2.2 Cell Adhesion and the Regulation of Tight Junction Assembly.....292 13.3 Regulation of Tight Junction Assembly During Early Xenopus Development ...............................................................................................293 13.4 Tight Junction Proteins as Regulators of Cell Differentiation During Development ...............................................................................................295 13.5 Conclusions .................................................................................................296 Acknowledgments..................................................................................................296 References..............................................................................................................296

13.1 INTRODUCTION During early development in most organisms, epithelia are generated to construct a compartmentalized body plan. These epithelia are a useful resource for the embryo, not just for their polarized transport function but also because they constitute a twodimensional tissue sheet that can be molded into three-dimensional shapes of increasing complexity. Shaping epithelia into the correct form at the correct time in development has been conserved throughout evolution to underpin those basic embryological transitions such as blastula formation (or blastocyst in mammals), gastrulation, and neurulation. A second important mechanism of developmental biology is to construct or disperse the epithelial phenotype in a dynamic way, according to the requirements of the emerging body plan. Thus, epithelial–mesenchymal interconversion of cells allows for regulated cell movement and changes in cellular organization and morphology. The formation or loss of epithelial tissues

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contributes to gastrulation, neurulation, and neural crest cell formation, and to later organogenesis, such as in the development of the kidney. The regulation of the epithelial phenotype, either in the shaping of tissue sheets during early development or in the formation or loss of epithelia during cell movement or organogenesis, is primarily coordinated by the activity of the intercellular junctions and associated cytoskeleton. The activity of the E-cadherin/catenin protein complex of the adherens junction in regulating epithelial tissue formation and developmental signaling is now well established (Takeichi, 1991; McCrea et al., 1993). Tight junctions (TJs), in contrast, have long been regarded as rather static membrane complexes that surround the apical border of epithelial cells and seal them to their neighbors, thereby preserving the integrity of the transepithelial permeability barrier and the apicobasal polarity inherent within each cell. However, examination of developmental epithelia has provided evidence of a more dynamic role for the TJ in coordinating epithelial function and, from this, contributing to developmental transitions. In particular, the characteristics of molecular interactions and the expression profile of individual proteins of the TJ complex can regulate the timing of TJ biogenesis and, indeed, the capacity of the epithelium to function in developmental processes. This chapter examines the mechanisms of TJ biogenesis during early development in vertebrates, with emphasis on mammalian blastocyst and Xenopus blastula formation. The relationship between TJ assembly and the functioning of the respective epithelia is emphasized. In addition, mechanisms that may facilitate the participation of TJ proteins in dynamic changes in epithelial phenotype that occur during development are considered.

13.2 THE TROPHECTODERM MODEL OF TIGHT JUNCTION ASSEMBLY The trophectoderm arises from cleavage of the fertilized mammalian egg. This epithelium forms the outer layer of cells of the blastocyst, a spherical cyst some 100 µm in diameter that also contains the blastocoelic cavity and another population of cells, the inner cell mass (ICM) situated underneath one region of trophectoderm and adjacent to the blastocoel (Figure 13.1A and B). During cleavage, cells enter either the trophectoderm or ICM cell lineage by a process of differentiative cell division whereby daughter cells allocated toward the embryo interior constitute the ICM (Johnson and Ziomek, 1981; Fleming, 1987). The blastocyst forms 3 to 4 days after fertilization when the embryo has around 32 cells. The blastocyst is the first stage where developmental pattern formation is evident, and has been most intensively studied in the mouse. The outer trophectoderm, when it completes its differentiation at the 32-cell stage, has an outward-facing apical membrane and initiates vectorial transport of fluid across from the outer environment (maternal uterine cavity) into the embryo interior to form the blastocoel. This tranport activity is driven by Na+,K+-ATPase, localized on the basolateral membrane domain (DiZio and Tasca, 1977; Vorbrodt et al., 1977; Watson and Kidder, 1988; MacPhee et al., 1994; 2000; Jones et al., 1997). At this stage, the ICM is therefore dependent upon the transport function of trophectoderm cells for influx and efflux of molecules

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FIGURE 13.1 Mouse blastocyst shown diagrammatically (A), in phase contrast (B), and following confocal microscopic analysis for the detection of TJ-associated protein ZO-1α– isoform (C). The outer trophectoderm (TE) and inner cell mass (ICM) lineages are apparent (unshaded and shaded, respectively, in A), together with the blastocoel (b) and surrounding zona pellucida (ZP in B). In blastocyst midplane, ZO-1α– protein is localized to the junctional site between trophectoderm cells (arrows in C).

and ions. Indeed, transport systems for amino acids, sugars, ions, and transcytosis of growth factors and other macromolecules have been identified between the maternal environment and the blastocoel cavity (DiZio and Tasca, 1977; Fleming and Pickering, 1985; Heyner et al., 1989; Manejwala et al., 1986; Dardik and Schultz, 1991; Wiley et al., 1991; Aghayan et al., 1992; Hewitson and Leese, 1993; Brison et al., 1993; Smith et al., 1993; Barr et al., 1998). The blastocoel acts, therefore, as a controlled microenvironment for the ICM and a reservoir of nutrients and other factors necessary for subsequent ICM development. After full expansion of the blastocoel, it is the trophectoderm that initiates implantation into the uterine wall and continues to differentiate into the extraembryonic lineages comprising the trophoblast, ectoplacental cone, and ultimately the chorio-allantoic placenta. The protected ICM gives rise to the entire fetus and the extraembryonic visceral and parietal endoderm lineages.

13.2.1 MOLECULAR MATURATION OF THE TIGHT JUNCTION TROPHECTODERM DIFFERENTIATION

DURING

TJs are present between trophectoderm cells and are essential for blastocoel cavity formation (Ducibella et al., 1975; Magnuson et al., 1977; Figure 13.1C). Together with cadherin-based adherens junctions and desmosomes, they also maintain the integrity of the epithelium and the entire blastocyst during blastocoel expansion. The maturation of TJs takes place during the period of trophectoderm differentiation. When the mouse embryo has only eight cells (about 24 h prior to the onset of blastocoel formation), the cells of the embryo activate the E-cadherin/catenin adhesion system located along cell contact sites resulting in the adherence of cells into a tight ball (now called a morula) in a process called compaction (Vestweber et al., 1987; Ohsugi et al., 1996). While compaction occurs, the cells become polarized and exhibit an outer apical membrane rich in microvilli and a nonmicrovillous

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TABLE 13.1 Phases in the Molecular Maturation of the Tight Junction during Mouse Trophectoderm Differentiation 8-Cell Stage E-cadherin/catenin adhesion and epithelial polarity initiates ZO-1α–, rab13, and JAM membrane assembly at apicolateral contact site Permeable junction 16-Cell Stage Cingulin assembly at apicolateral contact site Permeable junction 32-Cell Stage ZO-1α+ de novo expression and intracellular association with occludin ZO-1α+ and occludin assembly at apicolateral contact site Claudin-1 membrane assembly Segregation of TJ and E-cadherin/catenin adherens junction Impermeable junction Blastocoel cavity formation begins Note: See text for references.

basolateral membrane (Handyside, 1980; Ziomek and Johnson, 1980; Reeve and Ziomek, 1981). Within the cytoplasm, specific organelles and the cytoskeleton also realign along the protoepithelial apicobasal axis of polarity (Johnson and Maro, 1984; Fleming and Pickering, 1985; Houliston et al., 1987). It is the initiation of cadherin adhesion at compaction that catalyzes the onset of cell polarization and regulates its apicobasal orientation (Johnson et al., 1986). Similarly, cadherin adhesion acts as a positive regulator for membrane assembly of the TJ which begins within hours after compaction (Fleming et al., 1989). The assembly of TJs from the late eight-cell stage is characterized by a protracted maturation in their molecular composition with at least three phases of assembly being evident (Table 13.1). For information on the molecular composition of TJs, see the reviews by Mitic and Van Itallie (Chapter 10), Citi (Chapter 11), and Fanning (Chapter 12) in this book. The first phase, occurring during the eight-cell stage, is characterized by the assembly of the TJ cytoplasmic plaque protein, ZO-1 (Fleming et al., 1989). ZO-1 (Stevenson et al., 1986; Anderson et al., 1988; Itoh et al., 1993; Willott et al., 1993) is expressed in two alternatively spliced variants, either with or without an 80 amino acid C-terminal α domain (Willott et al., 1992; Balda and Anderson, 1993). More recent studies have shown that it is exclusively the isoform lacking the α domain (ZO-1α–) that assembles at the apicolateral contact site at the eight-cell stage (Sheth et al., 1997). This isoform is expressed as both a maternal transcript and as an early transcript from the embryonic genome that activates from the late one-cell/early two-cell stage in the mouse. It is therefore detectable throughout all stages of cleavage at mRNA (RT-PCR [reverse transcription-polymerase chain reaction], in situ hybridization) and protein (immunoblotting, immunoprecipitation) levels. Use of isoform-specific antiserum and confocal microscopy has shown

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ZO-1α– first assembles at the apicolateral contact site as punctate foci and only becomes a continuous beltlike ring of immunostaining several hours later, usually after further division to the 16-cell stage (Fleming et al., 1989; Sheth et al., 1997; Figure 13.2A and B). In recent studies, the authors have found that the newly identified TJ transmembrane protein, junction adhesion molecule (JAM; Martin-Padura et al., 1998), also assembles at the apicolateral junction site from the late eight-cell stage (F. Thomas et al., manuscript in preparation). In addition, it was found that ZO-1α– assembly is coordinated with that of the rab GTPase, rab13 (Sheth et al., 2000b). The rab proteins are a subfamily of the Ras superfamily of regulatory GTPases and are involved in the targeting of proteins to specific destinations during intracellular transport (Novick and Zerial, 1997; Chavrier and Goud, 1999). Rab13 contains a C-terminal CaaX motif (C, cys; a, mostly aliphatic; X, any amino acid) similar to those of Ras/Rho proteins and is isoprenylated in vivo and geranylgeranylated in vitro, which may facilitate its anchorage to the lipid bilayer during membrane attachment (Joberty et al., 1993). Regulation of membrane association and disassociation is achieved by rab13 specific binding to rod cGMP phosphodiesterase δ subunit (δ-PDE; Marzesco et al., 1998). Significantly, rab13 has been shown to associate with the TJ in mature epithelial cells, indicative of a specific role in targeting and docking of TJ proteins (Zahraoui et al., 1994). In the mouse embryo, rab13 is expressed throughout cleavage and assembles at the apicolateral junction site in precise colocalization with ZO-1α– after compaction at the eight-cell stage (Sheth et al., 2000b). This early assembly of rab13 during TJ biogenesis may suggest that it is involved not only in maintaining the integrity of the junction in mature cells but also in the specification of this domain during differentiation. Rab13 regulatory protein, δ-PDE, exhibits two putative C-terminal sequences necessary for interaction with PDZ motifs (PSD95, Dlg, ZO-1) (Marzesco et al., 1998) which, significantly, are present on ZO-1 and other ZO family TJ-associated proteins (Anderson, 1996; Dimitratos et al., 1999). Rab13/δ-PDE is therefore suitably equipped to initiate ZO-1 targeting to the putative TJ site. The second phase in TJ assembly occurs during the 16-cell stage, some 12 h after ZO-1α– and rab13 assembly. At this time, the cytoplasmic plaque protein cingulin (Citi et al., 1988; 1989; Cordenonsi et al., 1999a) colocalizes with ZO-1α– at the apicolateral contact site (Fleming et al., 1993). Cingulin, like ZO-1, is expressed by both maternal and embryonic genomes (Javed et al., 1993). Maternal cingulin in the unfertilized egg is localized in the egg membrane skeleton (Fleming et al., 1993), possibly as a result of cingulin binding sites with myosin (Cordenonsi et al., 1999a), which also occurs in the egg cortex. This pool of cingulin may play a role in the adherence of cumulus cells that surround the oocyte up to and immediately after fertilization. However, it is only embryonically expressed cingulin that assembles at the nascent TJ site, the maternal pool is internalized by endocytosis during cleavage and appears to undergo degradation (Fleming et al., 1993). The third and final phase in TJ assembly occurs some 12 h later again from phase two, and is restricted to the early 32-cell stage. This step is characterized by the coassembly of the transmembrane component occludin (Furuse et al., 1993; Ando-Akatsuka et al., 1996) and the ZO-1α+ isoform as a complex at the junction

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site (Sheth et al., 1997; 2000a; Figure 13.2C to F). Occludin is detectable as a transcript throughout cleavage and at least four separate bands in immunoblots of molecular mass ranging between 67 to 72 kDa (Figure 13.2G). Biochemical and quantitative densitometric analyses have revealed that the different forms of occludin increase or decrease their relative amount during cleavage and blastocyst formation and that they may represent post-translational modifications (Sheth et al., 2000a). Distinct post-translational forms of occludin have been detected in many cell types (Cordenonsi et al., 1997; Sakakibara et al., 1997; Wong, 1997). One form of occludin in the embryo (band 2, 65 to 67 kDa) increases in abundance during cleavage and appears to undergo phosphorylation close to the time of blastocyst formation and exclusively enters the detergent-insoluble pool (Figure 13.2G). It is this form of occludin that appears to assemble at the junction since only assembled occludin is detergent-insoluble (Sheth et al., 2000a). Occludin assembly at the junction site at the 32-cell stage is followed very rapidly by polarized fluid transport across the trophectoderm layer and the onset of blastocoel cavitation. Recent studies have shown that the TJ membrane protein claudin-1 (Furuse et al. 1998; Morita et al., 1999) also assembles during this late phase of TJ biogenesis in the embryo (F. Thomas et al., manuscript in preparation). It is only from the time of cavitation that the trophectoderm becomes impermeable to electron dense tracer and vital dyes (Magnuson et al., 1978; Sheth et al., 2000b). This suggests that the timing of occludin band 2 post-translational modification and assembly (and possibly also claudin-1 assembly) may act as a limiting factor in TJ sealing and thereby regulate the timing of blastocyst morphogenesis. How might this be achieved? The evidence indicates an important role for the ZO-1α+ isoform. ZO-1α+ is first transcribed in the embryo during the late 16-cell stage and detectable as a distinct protein band in immunoblots soon after. ZO-1α+, when first expressed, is localized to perinuclear Golgi sites before delivery to the membrane TJ during the early 32-cell stage (Sheth et al., 1997; Figure 13.2C and D). Double-label confocal microscopy has revealed that occludin and ZO-1α+ colocalize at the perinuclear sites before their membrane assembly as a complex. An example of occludin localization at cytoplasmic and junctional sites in late morulae is shown in Figure 13.2E and F. If membrane assembly of this complex is inhibited by transient brefeldin A treatment, blastocyst cavitation is likewise inhibited. The timing of FIGURE 13.2 Immunoconfocal (A to F) and immunoblotting (G) of TJ proteins in mouse embryos at different stages of cleavage. (A) Immediately following compaction in eight-cell embryos, ZO-1α– assembles at the apical border between blastomeres as a discontinuous series of spots (arrowheads; image is in tangential optical plane, grazing the surface of the embryo). (B) During the 16-cell stage, ZO-1α– becomes a continuous belt of staining around each outer cell (arrowheads). (C, D) During the early 32-cell stage, ZO-1α+ isoform is expressed and is first evident at perinuclear sites within outer (trophectoderm lineage) cells (C, arrowheads) before assembly at the TJ site (D, arrowheads). (E, F). During the 32-cell stage, occludin protein is first evident at perinuclear sites (E, arrowheads) where colocalization with ZO-1α+ occurs, before assembly at the membrane (F, arrowheads). (G) Immunoblot of occludin in eight-cell, blastocyst, and control lung tissues showing both total and detergentinsoluble fractions. Occludin is apparent within four main bands as indicated. It is only band 2 that enters the insoluble fraction in blastocysts.

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completion of TJ assembly in the early embryo, and the transition from morula to blastocyst, may therefore be regulated by the timing of ZO-1α+ transcription (Sheth et al., 1997; 2000a).

13.2.2 CELL ADHESION AND JUNCTION ASSEMBLY

THE

REGULATION

OF

TIGHT

The trophectoderm model for TJ assembly during development has provided strong evidence for E-cadherin adhesion at compaction as a permissive state for subsequent assembly of the TJ. Thus, if E-cadherin adhesion during the eight-cell stage is prevented by neutralizing antibody, ZO-1α– membrane assembly is delayed and occurs in a nonpolarized and irregular manner (Fleming et al., 1989). Moreover, in E-cadherin or α-catenin knock-out embryos, normal trophectoderm differentiation is inhibited (Larue et al., 1994; Riethmacher et al., 1995; Torres et al., 1997) and ZO-1 assembly is disturbed (Ohsugi et al., 1997). A similar phenotype results from E-cadherin antisense RNA treatment (Wianny and Zernicka-Goetz, 2000). In addition to regulating membrane assembly of TJ plaque proteins, activation of E-cadherin adhesion at compaction also enhances their stability, probably by cytoskeletal anchorage. Thus, the half-life of newly synthesized cingulin in pulse-chase immunoprecipitation analysis is significantly extended following initiation of compaction and is shortened if E-cadherin adhesion is reversed (Javed et al., 1993). Activation of E-cadherin/catenin adhesion in the embryo, unlike the completion of TJ formation, does not appear to be regulated by the timing of expression of the main protein constituents that are present on membranes throughout cleavage (Vestweber et al., 1987; Sefton et al., 1992; Ohsugi et al., 1996). Rather, post-translational modifications appear to be involved since activation of protein kinase C (PKC) with phorbol ester can induce premature compaction at the four-cell stage (Winkel et al., 1990) and PKCα isoform redistributes to the cell contact sites at the time of compaction (Pauken and Capco, 1999). E-cadherin becomes phosphorylated from the eight-cell stage (Sefton et al., 1992) but is unlikely to be a direct substrate of PKC since the timing of phosphorylation is not modified during phorbol ester–induced premature compaction (Sefton et al., 1996). In contrast, phosphorylation of β-catenin occurs at serine-threonine residues not only at the time of natural compaction but also during prematurely induced compaction (Pauken and Capco, 1999), indicating it may be one substrate for PKC-mediated control of E-cadherin adhesion. β-catenin is also heavily tyrosine phosphorylated prior to compaction and gradually becomes tyrosine dephosphorylated in the morula (Ohsugi et al., 1999) but inhibition of tyrosine kinase with genistein did not modulate the timing of compaction (Goval and Alexandre, 2000). These data indicate, therefore, that TJ formation during trophectoderm differentiation may be activated secondarily in response to PKC signaling acting directly to initiate E-cadherin/catenin adhesion. However, recent evidence has also indicated that activation of specific isoforms of PCK stimulates the membrane assembly of ZO-1 and other TJ proteins in the embryo independently of the onset of E-cadherin adhesion (J. Eckert et al., manuscript in preparation), as has been implicated for culture cells (Balda et al., 1991; Stuart and Nigam, 1995; see also Chapter 17, by Balda et al.).

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The trophectoderm model has also provided evidence that the E-cadherin/catenin adhesion system contributes to the assembly of the TJ not only in permissive but also in structural terms. The late membrane assembly of occludin and claudin-1 during the 32-cell stage raises the question of the binding site of plaque proteins during earlier stages of cleavage. In mature TJs, in vitro and in vivo evidence has indicated that plaque proteins associate with the cytoplasmic domain of occludin and claudins (Furuse et al., 1994; Itoh et al., 1999a; Cordenonsi et al., 1999b). However, ZO-1 and ZO-2 can also associate with catenins and the E-cadherin membrane complex (Rajasekaran et al., 1996; Itoh et al., 1997; 1999b). Interestingly, during formation of an epithelium in vitro after replating of culture cells, ZO-1 transiently interacts with the E-cadherin complex before binding to occludin (Rajasekaran et al., 1996; Ando-Akatsuka et al., 1999). Recently, the authors examined the localization of ZO-1 and catenins in doublelabeled embryos at distinct stages of cleavage and found that ZO-1 is indeed precisely colocalized with catenins at apicolateral contact sites until the time of occludin assembly when they segregate, with ZO-1 (both isoforms) colocalizing with occludin while catenins are positioned subjacent to this site, within the zonula adherens and along the lateral membrane (Sheth et al., 2000b). Thus, the TJ appears to emerge from the adherens junction as a distinct entity during the terminal phase of epithelial differentiation in the embryo. This is supported by ultrastructural evidence. In recently compacted eight-cell embryos, the apicolateral contact sites are first unspecialized but then exhibit adherens-like junction plaques (Ducibella and Anderson, 1975; Sheth et al., 2000b; Figure 13.3A and B) and the embryo is fully permeable to electrondense and vital tracers. It is only after occludin assembly at the 32-cell stage that a characteristic TJ is apparent apical to the adherens junction, coinciding with the sealing of apposed membranes (Magnuson et al., 1978; Sheth et al., 2000b; Figure 13.3C).

13.3 REGULATION OF TIGHT JUNCTION ASSEMBLY DURING EARLY XENOPUS DEVELOPMENT The large size and rapid rate of cellularization of the amphibian egg impose greater structural difficulties in generating an epithelial permeability seal during blastula formation than those encountered by the mammalian egg. TJ assembly is under maternal genomic control and initiates from the time of first cleavage when epithelial membrane polarity is established (Roberts et al., 1992; Muller and Hausen, 1995). The Xenopus egg membrane is relatively impermeable to ions and is devoid of cadherins at the time of fertilization; this membrane becomes the apical membrane of the primary epithelium, which forms during cleavage. The early origin of cell polarity in Xenopus contrasts with the later onset of epithelial differentiation in the mouse at compaction. E-cadherin is not expressed until gastrulation in Xenopus. Instead, two maternally encoded and closely related cadherins, EP- and XB/Ucadherin, are expressed and become localized to newly formed basolateral membranes (Angres et al., 1991; Ginsberg et al., 1991; Herzberg et al., 1991; Levi et al., 1991; Muller et al., 1994). A distinct boundary is therefore formed after first cleavage between apical and basolateral membranes (Gawantka et al., 1992).

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FIGURE 13.3 Ultrastructure of apicolateral regions of contact between cells at the time of compaction in the eight-cell embryo (A, B) and within the trophectoderm of the blastocyst (C). After compaction occurs, an adherens-like junction forms (B, arrowhead), whereas in the blastocyst, an apical tight junction (C, arrow) and subjacent adherens junction (C, arrowhead) are apparent.

In addition to the onset of epithelial polarity, the blastocoel also forms in early cleavage and can be detected from the two-cell stage. However, ultrastructural, freeze-fracture, and electrophysiological analyses have produced inconsistent results and have variously indicated that TJs form from the two-cell (Kalt, 1971; Slack and Warner, 1973), eight-cell (Sanders and DiCaprio, 1976; Cardellini and Rasotto, 1988), 32-cell (Muller and Hausen, 1995), or later stages (Palmer and Slack, 1970; Regen and Steinhardt, 1986). More recent functional and immunocytochemical analyses have provided strong support for TJs being established at the two-cell stage. First, use of a biotin permeability assay indicated TJ sealing and exclusion from the nascent blastocoel was operative from the two-cell stage (Merzdorf et al., 1998). Second, cingulin protein was found to be localized at the border between apical and basolateral membrane domains from the two-cell stage (Cardellini et al., 1996). Xenopus embryo occludin, like mouse embryo occludin (see above), is expressed in different post-translational forms, including different phosphorylated states (Cordenonsi et al., 1997). As in the mouse embryo, occludin forms change following fertilization but in Xenopus it appears that dephosphorylation rather than phosphorylation regulates the onset of membrane assembly (Cordenonsi et al., 1997). Occludin assembly takes place from the two-cell stage and, like in the mouse embryo, follows assembly of ZO-1 and cingulin and corresponds to the time a permeability seal is formed (Fesenko et al., 2000).

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Despite the rapid nature of TJ formation in the Xenopus embryo compared with the mouse embryo, in both models junctions are assembled after establishment of epithelial polarity and lead immediately to the accumulation of blastocoelic fluid. In both models, TJs form by sequential assembly of constituents (although this occurs over a protracted time period in the mouse) with a potentially important regulatory role for occludin post-translational change in the timing of barrier formation. However, the relationship between cadherin adhesion and TJ formation may be distinct. In the mouse embryo, a significant role for cadherin adhesion in TJ assembly has been identified (see above). In contrast, in the Xenopus embryo, extracellular calcium depletion or vitelline membrane removal from before first cleavage abolishes cadherin adhesion but does not significantly alter the timing or spatial patterning of TJ assembly at the apical-basolateral membrane border (Cardellini et al., 1996; Fesenko et al., 2000). This suggests that the site of assembly is defined by cell polarity independently of cell adhesion.

13.4 TIGHT JUNCTION PROTEINS AS REGULATORS OF CELL DIFFERENTIATION DURING DEVELOPMENT During early development, assembly of TJs can act in a regulatory way to control the timing of epithelial transport function and dramatically alter the state of morphogenesis with the formation of a blastocyst or blastula. Assembly and disassembly of TJs during later stages of development can also have morphogenetic consequences. Thus, after mammalian blastocyst expansion, the embryo must implant in the uterus wall for continued development. To accomplish blastocyst attachment, uterine epithelial cells appear to downregulate ZO-1 expression and TJ integrity such that E-cadherin/catenins become redistributed to apical surfaces to enhance adhesiveness (Thie et al., 1996). Moreover, as implantation progresses, ZO-1 and E-cadherin are expressed in the underlying stromal cells at the implantation site, suggesting that a new permeability barrier is established to regulate access of immunologically competent maternal cells (Paria et al., 1999). Later, during neurulation in chick and mouse embryos, the cells of the neural plate undergo a loss of occludin expression and TJ permeability seal as they fold to form the neural tube (Aaku-Saraste et al., 1996). How might TJ proteins themselves contribute to the dynamic nature of developmental transitions? The plaque constituents ZO-1, ZO-2, and ZO-3 all belong to the MAGUK protein family of signaling molecules (Anderson, 1996; Dimitratos et al., 1999; see also Chaper 11 by Citi) and ZO-1 has homology with the Drosophila tumor suppressor, discs-large A (Willott et al., 1993). There is now growing evidence that ZO and other TJ proteins, in addition to being structural components of epithelial cells, also contribute to signaling pathways regulating cell differentiation and proliferation. Thus, ZO-1 is downregulated at cell junctions during breast cancer progression (Hoover et al., 1998), transcription of the A form of ZO-2 is inactivated in pancreatic duct adenocarcinoma (Chlenski et al., 1999), and occludin downregulation and ZO-1 relocation occurs in response to Raf-1-induced epithelial oncogenesis (Li and Mrsny, 2000). ZO-1 can be found in the nucleus as well as at the TJ (Gottardi et al., 1996) and, recently, it has been shown that the SH3 domain of ZO-1 is a

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binding site for the transcription factor, ZONAB, which in turn binds to the promoter of the cell cycle regulator, Erb-B2, to control its expression (Balda and Matter, 2000). ZO-1, as has been shown for the adherens junction constituent, β-catenin (Behrens et al., 1996; Huber et al., 1996; Barker et al., 2000), may therefore contribute to dynamic developmental events by the nature of its molecular interactions that will direct its participation in structural (TJ) vs. signaling (cytoplasmic/nuclear) activities. Growth factor signaling during development may provide the cue to coordinate the participation of ZO-1 and other TJ proteins in structural vs. signaling functions. Such extracellular factors may utilize intracellular kinase pathways already known to modulate TJ organization (Balda et al., 1991; Stuart and Nigam, 1995; see also Chapter 17 by Balda and Matter). Thus, hepatocyte growth factor/scatter factor (HGF/SF), acting through the Met receptor, induces epithelial–mesenchymal transformation and motility of activated cells. In addition, ZO-1 appears to be a downstream target of HGF/SF signaling since, in treated MDCK cells, ZO-1 relocates from the TJ to the cytoplasm (Grisendi et al., 1998). Moreover, it has been found that expression of mutated forms of ZO-1 similarly activate epithelial to mesenchymal conversion of MDCK cells (Reichert et al., 2000) and corneal epithelial cells (Ryeom et al., 2000).

13.5 CONCLUSIONS Examination of the biogenesis of TJs during development has shown that these structures not only perform an essential role in epithelial transport function but also contribute to changes in developmental state. Once E-cadherin adhesion and epithelial cell polarity are established, the characteristics of TJ protein expression and membrane assembly can act as a mechanism to activate epithelial function at the completion of differentiation. This indicates a role for the TJ in morphogenesis and the formation of a compartmentalized body plan. The dynamic nature of TJ assembly and disassembly during development, together with the growing evidence of signaling properties of certain constitutive proteins, suggests that these membrane complexes perform a critical role in epithelial differentiation and phenotype transformation.

ACKNOWLEDGMENTS The financial support of the Medical Research Council, Wellcome Trust, and European Union for research in T.P.F.’s laboratory on tight junction assembly and trophectoderm differentiation during development is gratefully acknowledged.

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14

Tight Junctions and Cell Surface Lipid Polarity Nanette Kälin and Gerrit van Meer

CONTENTS 14.1 Lipids Are Heterogeneously Distributed over Cellular Membranes .........306 14.1.1 Major Lipid Classes in Mammalian Membranes .........................306 14.1.2 Lipids of Different Cellular Membranes.......................................306 14.1.3 Lipid Heterogeneity within a Single Membrane ..........................307 14.2 Surface Polarity of Lipids...........................................................................308 14.2.1 Epithelial Surface Domains...........................................................308 14.2.2 Apical Lipid Composition .............................................................308 14.2.3 Protective Functions of Glycosphingolipids .................................309 14.3 Tight Junctions — A Diffusion Barrier to Membrane Components .........310 14.3.1 Lateral Diffusion............................................................................310 14.3.2 Tight Junction Structure ................................................................310 14.3.3 How Do Tight Junctions Affect Lipid Diffusion? ........................312 14.4 Lateral Lipid Macrodomains That Are Not Stabilized by Cell–Cell Contacts: Putative Mechanisms ..................................................................314 14.4.1 Myelin and Neuronal Cells ...........................................................314 14.4.2 Spermatozoa...................................................................................315 14.4.3 Other Polarized Cells.....................................................................316 14.5 Generation of Cell Surface Lipid Polarity .................................................316 14.5.1 Lipid Synthesis ..............................................................................316 14.5.2 Lipid Transport and Sorting: General Principles ..........................317 14.5.3 Vesicular Traffic to the Apical and Basolateral Plasma Membrane Domains ......................................................................318 14.5.3.1 Sorting by Lateral Segregation and Coupling to Specific Membrane Coats .............................................318 14.5.3.2 Transport, Docking, and Fusion....................................321 14.5.4 Monomeric Lipid Transport and Cell Surface Lipid Polarity ......321 14.5.5 Transmembrane Lipid Asymmetry and Cell Surface Lipid Polarity...........................................................................................322 14.5.5.1 The Aminophospholipid Translocase............................322 14.5.5.2 ABC-Transporters..........................................................323 References..............................................................................................................323

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14.1 LIPIDS ARE HETEROGENEOUSLY DISTRIBUTED OVER CELLULAR MEMBRANES 14.1.1 MAJOR LIPID CLASSES

IN

MAMMALIAN MEMBRANES

The membranes of living organisms are composed of an oily film of lipids, the lipid bilayer. The bilayer “dissolves” hydrophobic polypeptide domains, and the transmembrane domains of membrane proteins and anchors proteins via their acyl, prenyl, or phospholipid modification. A wide variety of different lipids are found in biological membranes. The main lipid classes in eukaryotic cells are glycerophospholipids (GL), sphingolipids (SL), and cholesterol. Glycerophospholipid species can be distinguished by their head groups: the most common lipid species are phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), and inositolphospholipids (PI). The head groups of mammalian sphingolipids are either a phosphorylcholine (sphingomyelin, SM) or carbohydrate (glycosphingolipids), in the simplest case a single glucose as in glucosylceramide (GlcCer) or galactose as in galactosylceramide (GalCer). The head groups of glycosphingolipids contain mostly not more than 15 sugar residues but display an extreme variety of combinations. Hundreds of different connections are possible for a carbohydrate tree of only three different sugars, theoretically a biological variety that exceeds even that of proteins and DNA. Among the lipids, glycosphingolipids can thus be considered experts in structural diversity. The two big lipid classes, glycerolipids and sphingolipids, are characterized by their lipid backbone, which in the first case is a glycerol and in the second a sphingoid base, a condensation product of serine and a C16 or C18 fatty acid. This structure is similar to a glycerol with a typical fatty acid in the sn-1 position. The striking difference between the two classes, however, manifests itself in the sn-2 position. Here, the fatty acids of glycerolipids are mostly (poly)unsaturated (e.g., C18:1, C18:2, or C20:4), whereas in sphingolipids the amide-linked fatty acid is long and saturated (C22:0, C24:0, C24:1) and can be hydroxylated, as is found in a large fraction of glycosphingolipids. Thus, the hydrophobic part of sphingolipids is strongly asymmetric concerning the length of the two fatty chains, is narrower, and extends deeper into the lipid phase than glycerophospholipids. As a consequence it increases the thickness of the membrane.

14.1.2 LIPIDS

OF

DIFFERENT CELLULAR MEMBRANES

The membranes of a cell are composed of a wide variety of lipids. The same lipids are found in all membranes of a cell, but the ratio by which a lipid species contributes to the composition is characteristic. In mammalian cells, which are the focus in this chapter, exceptions are cardiolipin, which is exclusively found in the inner membrane of mitochondria (Daum, 1985) and may be a remembrance of its bacterial origin, and lysobisphosphatidate, which is a marker for endosomes/lysosomes (Kobayashi et al., 1998). The relative abundance of a lipid species within each membrane or of a distinct membrane in a cell is cell type dependent. Table 14.1 summarizes data on the relative abundance of the lipids in a mammalian fibroblastic cell line (BHK; Allan, 1996). There is general agreement that the plasma membrane is enriched in

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TABLE 14.1 Mass Distribution of Lipid Species in Total Cellular Membranes and in the Plasma Membrane, and the Percentage of Each Lipid Present in the Plasma Membrane SM PC PS PI PE CL LBPA GSL Chol. TAG Total Cell 6 39 5 5 20 3 1.5 3 14.5 3 100 Plasma membrane 15 17 11 2 18.5 — — 7 29.5 — 100 Percent of each 67 11 60 10 25 — — 67 55 — 27 lipid in plasma membrane Abbreviations: SM = sphingomyelin; PC = phosphatidylcholine; PS = phosphatidylserine; PI = phosphatidylinositol; PE = phosphatidylethanolamine; CL = cardiolipin; LBPA = lysobisphosphatidate; GSL = glycosphingolipids; Chol. = cholesterol; TAG = triacylglycerol. Source: Modified from Allan, D., Mol. Membr. Biol., 13, 81, 1996.

cholesterol, glycosphingolipids, sphingomyelin, and PS, whereas the intracellular membranes contain more PC and PI. Most striking discrepancies between studies concern the portion of cholesterol present in the plasma membrane (24 vs. 90%; van Meer, 1987; Lange et al., 1989) and the percentage of total membranes made up by the plasma membrane (1 to 2%, Blouin et al., 1977; 15%, Griffiths et al., 1989; 50%, Lange et al., 1989).

14.1.3 LIPID HETEROGENEITY

WITHIN A

SINGLE MEMBRANE

Heterogeneity in lipid composition is not restricted to different membrane entities but extends to the level of a single membrane where lateral domains with distinct lipid composition can be distinguished. The saturation of the long fatty acid chains of sphingolipids allows tighter packing of the molecules and stronger hydrophobic interactions. These, together with pronounced intermolecular hydrogen bonding at the interface of lipid and aqueous phase, constitute the clustering capacity and drive sphingolipids to form membrane domains (Brown, 1998). Depending on their size, domains can be specified as macrodomains, such as the apical and basolateral part of the plasma membrane (van Meer and Simons, 1988), or as microdomains, such as cholesterol/sphingolipid rafts that are characterized by their detergent insolubility at 4˚C, but not at 37˚C, and their fragility upon cholesterol extraction (Brown and London, 1998b). These domains can be visualized upon clustering with polyvalent agents and size estimations spread between 70 nm (Friedrichson and Kurzchalia, 1998; Varma and Mayor, 1998) and a few hundred nanometers (Sheets et al., 1997; Pralle et al., 1999). Membranes are not only characterized by lateral lipid heterogeneity, but also exhibit lipid asymmetry across the plane of the bilayer (Devaux, 1991). At the plasma membrane, sphingolipids and cholesterol are enriched at the cell surface, whereas the aminophospholipids PE and PS are predominantly located at the cytosolic side of the membrane. The transbilayer distribution of cholesterol is still unclear. Differences in lipid composition are associated with differences in fluidity. Apical plasma

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membrane domains have been found to be less fluid than basolateral domains (Brasitus and Schachter, 1980), and this difference has been assigned to the outer leaflet of the plasma membrane bilayer (van Meer, 1988). The higher fluidity of the inner leaflet of plasma membranes has been attributed to the greater degree of unsaturation of the lipid chains within this monolayer (Tanaka and Ohnishi, 1976; El Hage Chahine et al., 1993). One could envision that the intramolecular differences in fatty chain length of sphingolipids entails a packing defect of two opposing sphingolipid molecules in the bilayer that could be circumvented by interdigitation of the confronting molecules or interactions of sphingolipids with cholesterol. For the latter, a formation as bilayer spanning dimers has also been proposed. Such a connection of the membrane leaflets by sphingolipids and/or cholesterol depends on the distribution of these lipids within the cytoplasmic membrane leaflet (which so far has not been thoroughly investigated), and would provide a model to explain the specificity of sorting and signaling processes at the cytosolic surface in membrane domain structures.

14.2 SURFACE POLARITY OF LIPIDS 14.2.1 EPITHELIAL SURFACE DOMAINS The intramembrane lipid heterogeneity is most eye-catching in epithelia where tight junctions (TJs) weld adjacent cells together at the apical pole, separating an apical from a basolateral membrane domain that differs in protein and lipid composition. The apical surface of epithelial cells lines a milieu that is outside of the body where the environmental conditions are no longer constant but are dramatically changing and often aggressive, where surfaces can be colonized with bacteria and fungi, and where the weapons of the immune system do not spare the lining host tissue. A cell surface that is in contact with such an environment has to protect the cell from the outside milieu without isolating it. Typically, these cell surfaces are shielded by the glycocalyx, a structure analogous to the cell wall in plants and microorganisms, and is built by the carbohydrate moiety of the glycoproteins and glycolipids of the plasma membrane together with extracellular layers of proteoglycans and glycoproteins. The glycocalyx creates a buffered ion milieu that shields and connects the cell with its environment. In contrast, the basolateral surface faces the internal milieu. Along the lateral sides, epithelial cells are connected to the neighbor cells. Their basal surface interacts with the extracellular matrix, the basal lamina.

14.2.2 APICAL LIPID COMPOSITION The membrane lipid composition of the outward-directed cell surface reflects its protective function. The apical membrane of epithelial cells of the intestine (Forstner et al., 1968; Douglas et al., 1972; Kawai et al., 1974; Brasitus and Schachter, 1980; Hauser et al., 1980), of the urinary bladder (Stubbs et al., 1979), and of MDCK cells derived from the distal nephron (Simons and van Meer, 1988) is two- to fourfold enriched in glycosphingolipids at the expense of PC in comparison with the basolateral surface domain. The concentration of glycosphingolipids at the apical cell

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TABLE 14.2 Lipid Composition of the Apical and Basolateral Plasma Membrane Domain of Rat Intestinal Cells in Comparison with That of a Nonpolarized Cell Percentage (mol/mol) Lipid class Glycosphingolipids Phospholipid PC PE PS + PI + SM Cholesterol a b

Rat Intestinal Cellsa Apical Basolateral 33 8 33 66 8 34 16 18 9 14 33 26

BHK Fibroblastsb Plasma membrane 6 58 23 12 17 37

Kawai et al., 1974. Renkonen et al., 1972.

surface is remarkable. For the brush border of intestinal cells, glycosphingolipids, phospholipids, and cholesterol contribute equally to the molar composition (Forstner et al., 1968; Kawai et al., 1974; Brasitus and Schachter, 1980), whereas the basolateral membrane contains two to four times less glycosphingolipids. For nonepithelial cells, a molar ratio of glycosphingolipid/phospholipid/cholesterol was reported of 1:51:48 (Table 14.2; Cooper, 1978; Allan, 1996). When taking into account that glycosphingolipids are restricted to the outer plasma membrane leaflet (Thompson et al., 1986), and assuming that cholesterol is homogeneously distributed over the two leaflets, the apical cell surface of an epithelial cell can be considered a glycosphingolipid/cholesterol macrodomain.

14.2.3 PROTECTIVE FUNCTIONS

OF

GLYCOSPHINGOLIPIDS

This specialized plasma membrane composition protects the cell not only against physical and mechanical stress, a condition where apical membranes of kidney and intestine increase the number of sphingolipid hydroxyl groups probably to make the exposed surface more rigid and impermeable (Pascher, 1976), but it also provides a barrier against lipases and bile acids that are intrinsic to the digestive tract. Lipolytic enzymes, such as phospholipases C and D and sphingomyelinase, also originate from bacteria. The secreted enzymes attack glycerophospholipids and sphingomyelin at the epithelial cell surface and can increase the virulence of a bacterial infection. The effects range from minor alterations in cell membrane composition and function to lethality at low concentrations (Songer, 1997). The epithelial cell surface is an attachment site for different microorganisms, pathogens as well as commensals. Lactosylceramide was proposed to serve as a low-affinity receptor for enteric bacteria (Karlsson, 1989). These bacteria secrete glycosidases specific for lacto-series glycosphingolipids and thereby promote their own interaction with the host (Falk et al., 1990). So, the mucosal epithelial cell surface must permit colonization while protecting

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the cell from its detrimental effects. Less beneficial interactions have been reported between the glycosphingolipid GM1 and cholera toxin (Orlandi and Fishman, 1998), between Gb3 and Shiga toxin/verotoxin (Jacewicz et al., 1986), and between GalCer and HIV (Bhat et al., 1991). On the basolateral surface, glycolipids were found to be involved in cell–cell communication and cell–matrix interactions (Zinkl et al., 1996). In kidney cortex epithelial cells the level of glycosphingolipids is low (Spiegel et al., 1988), and a high level of SM has been found in their apical membrane instead (Chapelle and Gilles-Baillien, 1983; Schwertz et al., 1983; Carmel et al., 1985; Molitoris and Simon, 1985; Venien and Le Grimellec, 1988). This may suggest that these cells face an external environment that is less aggressive than the intestinal lumen and that they are normally not colonized by microorganisms, permitting lower glycosphingolipid levels and reduced surface shielding.

14.3 TIGHT JUNCTIONS — A DIFFUSION BARRIER TO MEMBRANE COMPONENTS 14.3.1 LATERAL DIFFUSION The original fluid mosaic model of membrane structure as presented by Singer and Nicholson (1972) envisaged proteins and lipids as freely diffusing within the oily film of a membrane. Lipid molecules move micrometers within seconds (Edidin, 1974; Bretscher, 1980) and typical lateral diffusion coefficients in cellular membranes are in the order of 10–8 cm2/s for lipids and 10–9 to 10–11 cm2/s for proteins (Angelides et al., 1988). Taken an average diameter of an animal cell of 10 to 20 µm, a lipid at the apical pole would reach the opposite basolateral side within a minute. In comparison, internalization of the cell surface via endocytosis occurs with a halftime of approximately an hour (Griffiths et al., 1989). Thus, in the absence of mechanisms that restrict diffusion, the lipid composition of the cell would be expected to be homogeneous. The formation of a barrier is the easiest way to separate domains and, indeed, epithelial cells use such a membrane “fence,” the TJs, to prevent intermixing of apical and basolateral membrane components. TJs are anastomosing intramembrane strands that surround the apical pole of an epithelial cell and thoroughly connect adjacent cells. They impose a paracellular barrier to the diffusion of water and ions and prevent the lateral passage of amphiphilic molecules within the cell surface membrane.

14.3.2 TIGHT JUNCTION STRUCTURE This barrier could be an intramembranous protein structure, e.g., consisting of transmembrane proteins. Alternatively, it could be lipidic in nature (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982; Verkleij, 1984). It has been proposed that the core of the TJ consists of a hexagonal II phase, which would be a cylindrical inverted lipid micelle inserted between the two membrane leaflets: the lipid head groups face the membrane interior, whereas the fatty acids are in contact with those of the bilayer lipids (Figure 14.1a). Arguments for this simple lipid model of TJs

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(b)

FIGURE 14.1 Structural models of the TJ. (a) A simple lipid model of TJs (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982; Verkleij, 1984). The core of TJs is formed by a cylinder of inverted lipids that surrounds the apical poles of epithelial cells and results in partial fusion of adjacent plasma membranes. This model explains the paracellular barrier between adjacent cells, separating the mucosal (light red) from the serosal environment (light blue) and the presence of an asymmetric barrier within the bilayer of the plasma membrane. At the cell surface, diffusion of polar membrane components is restricted within either the apical or basolateral (red and blue) leaflet, maintaining the differences in polar lipid composition. No such barrier is formed for polar components at the cytosolic surface (orange) or for apolar molecules within the hydrophobic core of the bilayer (yellow). However, this simple lipid model proposes lipid continuity between the cell surfaces of adjacent cells. This was found not to be the case. (b) A protein-based model of TJs (van Meer and Simons, 1986; Wong and Goodenough, 1999). Various transmembrane proteins of the adjacent cells have recently been shown to contribute to the formation of TJs. Upon lateral polymerization the resulting belts glue adjacent cells together and separate apical from basolateral compartments. Interaction of such a protein structure with the cell surface membrane interface abolishes the passage of polar components, not only between the aqueous environments (light red and light blue), but also between the apical and basolateral membrane interfaces (red and blue), preventing the passage of the charged or polar lipid head groups. There is no reason such a protein structure would impair the diffusion at the cytosolic leaflet or within the hydrophobic core of the plasma membrane. On the cytosolic side, the tight junctional protein complex is connected to other intracellular proteins that connect the complex with the cytoskeleton. In the paracellular pathway, the tight junctional protein complex forms selective pores that permit the exchange of specific compounds between the mucosal and serosal compartments (Wong and Goodenough, 1999).

were found in the perfect complementarity of P and E faces in freeze-fracture replicas that were considered to be consistent with an intralipid but not with an intraprotein structure (Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982) and which have been observed in pure lipid systems (Meyer, 1983). However, since then, the detergent stability of tight junctional strands visualized by electron microscopy (Stevenson and Goodenough, 1984) and the detection of several protein components,

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the transmembrane proteins occludin (Furuse et al., 1993), different isoforms of the claudin family (Furuse et al., 1998a,b), and JAM (Bazzoni et al., 2000) demonstrate that proteins are substantially involved in the tight junctional ultrastructure. On the cytosolic surface, the tight junctional protein complex interacts with cytosolic and peripheral membrane proteins that are thought to be involved in cross-linking TJs with the underlying actin-based cytoskeleton, in cellular signaling and in vesicle targeting (Figure 14.1b).

14.3.3 HOW DO TIGHT JUNCTIONS AFFECT LIPID DIFFUSION? Additional arguments against the lipid model have been presented by functional studies. In the lipid model of TJs, the exoplasmic bilayer leaflet of the apical membrane is continuous with that of the adjacent cells (see Figure 14.1a). To address this prediction, the diffusion of a cell-type-specific lipid was investigated in a polarized mixed monolayer of Madin–Darby canine kidney I (MDCK I) and MDCK II cells. MDCK II cells express Forssman antigen (an antigenic glycosphingolipid of the globo series) on their cell surface, whereas MDCK I cells do not. Immunofluorescence (Nichols et al., 1986; van Meer et al., 1986) and immunoelectronmicroscopy (Figure 14.2; van Genderen et al., 1991) demonstrated that the lipid antigen remained restricted to the host cell. No membrane continuity of the apical and basolateral cell surfaces was established upon generation of a polarized epithelial monolayer (Figure 14.2). This restriction was not a result of cell heterogeneity but was similarly seen when fluorescent-labeled lipids were incorporated into a subpopulation of cells in a homogenous MDCK monolayer (van Meer et al., 1986). Diffusion from one cell to the next has been observed for the short-chain lipid analogue C6-NBD-PC (Grebenkämper and Galla, 1994). However, the fact that this lipid rapidly exchanges in a monomeric form across the aqueous phase does not allow the conclusion of membrane continuity. The detected intercellular separation of apical cell surface membrane leaflets is not in accordance with a simple lipidic model of TJs (see Figure 14.1a). To study the effect of TJs on lipid diffusion between the apical and basolateral domain, fluorescent amphiphiles (Dragsten et al., 1981) and lipid analogues (Spiegel et al., 1985; van Meer and Simons, 1986; van Meer et al., 1987; Calderón et al., 1998) were incorporated apically or basolaterally, and their lateral diffusion into the opposite plasma membrane domain was recorded. In the first investigations fluorescent amphiphiles were added to the apical surface (Dragsten et al., 1981). Some probes remained on the apical surface while part diffused to the basolateral domain. The authors proposed that some probes moved across the lipid bilayer and concluded that lateral diffusion was inhibited in the outer but not the inner plasma membrane leaflet. However, the unknown transbilayer behavior and the partial water solubility of the probes did not allow unequivocal conclusions. An abrogation of lateral lipid diffusion at the TJs was also demonstrated for exogenously added fluorescent phospholipids and glycosphingolipids (Spiegel et al., 1985; Mandel et al., 1993; van Genderen and van Meer, 1995; Calderón et al., 1998), and for natural glycosphingolipid GM1 added to the outside of the membrane or generated by exogenous neuraminidase (Spiegel et al., 1985).

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FIGURE 14.2 Immunogold labeling of Forssman glycolipid in a mixed monolayer of cocultured MDCK I and MDCK II cells (van Genderen et al., 1991). Polarization of the cells is indicated by the formation of TJs at the apical cell contacts. The two cell types can be distinguished by the presence (MDCK II cell, right) or absence (MDCK I cells, left) of the lipid antigen. The glycosphingolipid is expressed in intracellular membranes and at the cell surface. The immunoelectronmicrograph demonstrates the restriction of the sphingolipid to MDCK II cells. The lipid does not diffuse to the apical or basolateral cell surface of the adjacent MDCK I cell, indicating the absence of lipid continuity between the outer membrane leaflets of adjacent cells. (From van Genderen, I. L. et al., J. Cell Biol., 115, 1009, 1991. With copyright permission of the Rockefeller University Press.)

Other investigators (van Meer and Simons, 1986; Knoll et al., 1988) inserted water-insoluble fluorescent (N-rhodamine-PE) or radioactive phospholipids ([3H]PC) into the apical plasma membrane by a fusion protocol and studied lipid diffusion to the basolateral compartment by fluorescent microscopy or lipid autoradiography. When fluorescent lipids were fused exclusively into the exoplasmic leaflet of the apical plasma membrane by using liposomes, where the fluorescent lipid was restricted to the vesicle outside, the lipid remained apical until the TJs were opened by calcium removal. However, when fused into both the exo- and cytoplasmic leaflets by using symmetrical liposomes, fluorescent lipid redistribution into the basolateral membrane compartment was observed. This led to the conclusion that TJs form a lipid diffusion barrier at the exoplasmic but not cytoplasmic leaflet of the membrane. Similar results were obtained after fusion of symmetrical liposomes containing [3H]PC with the apical membrane, rapid-freezing, freeze-substitution, and localization of the PC by autoradiography. Yet another technical strategy has taken advantage of the biosynthetic capacity of the cell. When cells were fed with NBD-ceramide, a lipid precursor that contains

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a fluorescent C6-fatty acid in the sn-2 position, this resulted in the synthesis of NBDsphingomyelin and NBD-glucosylceramide (Lipsky and Pagano, 1985). After 1 h at 37˚C, the products were found at the cell surface by fluorescence microscopy. In addition, the fluorescent lipids were completely extractable by BSA (van Meer et al., 1987) indicating that the analogues were present in the outer, exoplasmic leaflet. Performing these BSA back-exchange experiments at either the apical or basolateral surface completely abolished membrane fluorescence of that compartment without affecting the opposite membrane domain (van Meer et al., 1987). This implied an inhibition of lateral diffusion at the cell surface across the TJ area. These results were not restricted to kidney (van Meer et al., 1987) and intestinal cells (van‘t Hof and van Meer, 1990), but were also confirmed for hepatocytes (Zaal et al., 1994). All these experiments led to the conclusion that TJs act as a diffusion barrier in the exoplasmic membrane leaflet for amphiphilic lipids in addition to its well-known barrier function in the paracellular passage of water and ions. The fact that the barrier is limited to one leaflet of the lipid bilayer demonstrates that the lipids do not rapidly redistribute over the two membrane monolayers. A rapid transversal movement, as is expected for all apolar lipids, should abolish the effect of a barrier that is present in one leaflet. After flipping into the cytosolic leaflet, the lipids could rapidly diffuse through the continuous cytosolic leaflet. Indeed, cholesterol concentrations of apical and basolateral domains seem very similar (see Table 14.2; Kawai et al., 1974; van Meer, 1988). Still, it should be realized that this equal distribution may be coincidental, as according to freeze-fracture studies using filipin the organization of cholesterol in the two domains is different (Miller and Baldridge, 1985). Most likely, the disposition of cholesterol is governed solely by its affinity for the lipids in the two domains. The restriction of the barrier to the exoplasmic leaflet implies that the differences in lipid composition between apical and basolateral membranes reside in their exoplasmic leaflets. This makes it possible to calculate the distribution of the various lipid classes across the bilayer. Whereas the glycosphingolipids (and SM in proximal kidney cells) are enriched in the exoplasmic leaflet of the apical membrane, PC (and SM in intestinal cells and distal kidney cells) is enriched in the exoplasmic leaflet of the basolateral membrane. PE is enriched in the cytosolic leaflet (van Meer and Simons, 1986).

14.4 LATERAL LIPID MACRODOMAINS THAT ARE NOT STABILIZED BY CELL–CELL CONTACTS: PUTATIVE MECHANISMS 14.4.1 MYELIN

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NEURONAL CELLS

Myelin is a specialized domain of the plasma membrane of oligodendrocytes and Schwann cells. The glycosphingolipids GalCer and sulfatide constitute roughly 25 mol% of the myelin lipids (Morell et al., 1994). TJ structures (Tabira et al., 1978) and a tight junctional protein of the claudin family (Morita et al., 1999) have been localized between successive lamellae of the myelin sheath, and they turn out to be essential for myelin function (Gow et al., 1999). These structural features plus the finding of the typical apical protein MAL in myelin (Frank, 2000) support the idea

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that myelin forms a (series of) apical domain(s) that is separated from the plasma membrane of the cell body (Gumbiner and Louvard, 1985). The innermost lamella of the myelin sheath forms junctions with the axonal membrane (Gumbiner and Louvard, 1985). These contain proteins other than those in TJs. One protein component has been identified as a member of the neurexin family (Menegoz et al., 1997), members of which had been shown to be part of septate junctions (Baumgartner et al., 1996). Although less tight than TJs, septate junctions may well prevent the lateral diffusion of proteins and lipids (Aschenbrenner and Walz, 1998). Interestingly, even in nonmyelinated nerve cells evidence has been presented for a lateral diffusion barrier between the axonal and somatodendritic plasma membrane domains at the axon hillock. Long-chain fluorescent lipid and the natural glycolipid GD1a, after insertion into the axonal membrane by specific liposome fusion, essentially remained in the axon (Kobayashi et al., 1992). In contrast, the axon hillock did not impair lateral diffusion of the phospholipid rhodamine-PE (Angelides et al., 1988) or the amphiphile DiI (Winckler and Poo, 1996), and it has been argued that in the Kobayashi experiments the sensitivity of the fluorescence measurements was too low to detect free diffusion (Futerman et al., 1993). Controversial results were also reported concerning the surface distribution of the endogenous glycosphingolipid GM1. Whereas Ledesma et al. reported that GM1 became restricted to the axon upon maturation of the neurons (Ledesma et al., 1999), Futerman and colleagues were unable to detect a polarized distribution of GM1 under any condition (Sofer and Futerman, 1995; Shogomori et al., 1999). Still, an impediment of lateral diffusion was observed at the axon hillock when membrane proteins coupled to beads were dragged laterally along the axon by using “laser tweezers.” Interestingly, not only the diffusion of transmembrane proteins was affected but also that of Thy1, a protein anchored by a lipid (GPI-)tail. Further characterization of the diffusion barrier revealed an enrichment in ankyrin, a cytoskeletal protein also implicated in restricting proteins to the nodes of Ranvier (Davis et al., 1996), and the barrier required an intact submembranous cytoskeleton (Winckler et al., 1999). A high density of transmembrane proteins anchored to the cytoskeleton (Bennett and Gilligan, 1993) could impede diffusion of membrane proteins. By inducing a kind of intramembrane “traffic-jam,” membrane components that cannot interact with the cytoskeleton, such as GPI-anchored proteins or lipids, could also be prevented from traversing this region.

14.4.2 SPERMATOZOA The prime example of a cell possessing plasma membrane domains in the absence of TJs is the free-swimming sperm cell (Friend, 1989). The plasma membrane of sperm cells consists of two obvious macrodomains, head and tail, that are further divided into subdomains. The domains are defined according to their morphology, to protein antigens, and surprisingly to lipid composition and membrane fluidity. The lipid heterogeneity suggests the presence of barriers to lipid diffusion (Wolf and Voglmayr, 1984; Bearer and Friend, 1990; Gadella et al., 1994; Ladha et al., 1997). Although dense accretions have been identified by electron microscopy within

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and below the plasma membrane bilayer, the posterior ring and the annulus, a direct correlation between these structures and a function as a membrane barrier has not been established so far. In principle, the existence of lipid domains could also be due to the immiscibility of lipid phases (Wolf et al., 1988). If this is the case in the sperm plasma membrane, the challenge is to find out how the domains maintain their relative positions on the cell surface.

14.4.3 OTHER POLARIZED CELLS Several cell types display functionally differentiated surface domains that are not defined by the presence of TJs. One example is the cytotoxic T-cell, which attaches to a target cell and extends the contact into a membrane cleft, into which it secretes the pore-forming perforin and granzymes (Berke, 1994). The actual mechanism by which the T cell protects its own membrane against these harmful substances is unclear but most likely involves sealing the cleft to prevent leakage and the generation of a specialized resistant membrane domain lining the cleft. A similar mechanism may be used by the bone-resorbing osteoclast. The cell membrane of a resorbing osteoclast is divided into four different domains. The ruffled border is a secretory membrane, which contacts the bone and is bordered by the sealing zone, an area of close bone attachment. A “basolateral” and an “apical” membrane domain can be distinguished at the opposite side of the cell. The basolateral domain is distinguished from the apical domain, a central area at the top of the first, by the targeting of proteins that in epithelial cells are sorted to the respective domain and the characteristic distribution of certain carbohydrate structures. The mechanisms by which this lateral membrane heterogeneity is generated and maintained are unclear. The membrane organization is dynamic and reversible: domains appear and disappear various times during the lifetime of the cell depending on its physiological state (Salo et al., 1996).

14.5 GENERATION OF CELL SURFACE LIPID POLARITY 14.5.1 LIPID SYNTHESIS In intestinal cells and distal kidney cells glycosphingolipids are apically enriched at the expense of PC and SM, which as a consequence appear enriched at the basolateral membrane. In proximal kidney cells, SM is apically enriched whereas PC is enriched on the basolateral surface. Theoretically, this cell surface lipid polarity could be generated at the level of lipid biosynthesis. For this, lipids should be synthesized at the cell surface. This is not the case. In eukaryotic cells, essentially all lipid synthesis occurs within intracellular membranes. The glycerophospholipids (PC, PE, PS, PI), cholesterol, and ceramide are assembled at the cytoplasmic surface of the endoplasmic reticulum (ER; Dennis and Vance, 1992). Cholesterol synthesis presumably also takes place in peroxisomes (Krisans, 1992) and some PE is generated in mitochondria (Dennis and Kennedy, 1972). At the ER, distinct activities mediate base exchange among the glycerolipids PC, PE, and PS (Kanfer, 1980). Phospholipid base-exchange activities have also been detected at the plasma membrane (Siddiqui and Exton, 1992).

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The bulk of the sphingolipids is generated in the Golgi complex. After synthesis in the ER, ceramide is transported to the Golgi for conversion into SM, GlcCer, and higher glycosphingolipids. SM is generated within the lumenal leaflet of the cis-/medial-Golgi (Futerman et al., 1990; Jeckel et al., 1990) and at the plasma membrane, in epithelial cells on the basolateral surface (Kallen et al., 1993; van Helvoort et al., 1994). The majority of the higher glycosphingolipids originates from sequential modifications of GlcCer. GlcCer is generated at the cytosolic surface of the cis/medialGolgi by the UDP-glucose ceramide glucosyltransferase (Futerman and Pagano, 1991; Jeckel et al., 1992). In the lumen of the medial- and trans-Golgi (Lannert et al., 1994; Burger et al., 1996) GlcCer is galactosylated to LacCer, which is the basis of specific series of complex glycolipids such as the gangliosides. Whereas GlcCer and its derivatives are the major glycosphingolipids in murine intestine, human intestinal epithelia mainly contain GalCer (Falk et al., 1979). GalCer is generated within a lumenal membrane leaflet of the ER (Sprong et al., 1998). Also, GalCer can be further modified in the lumen of the Golgi. However, the addition of head group moieties is restricted to sulfation, further galactosylation, and/or sialylation. Clearly, the localization of lipid-synthesizing activities in the ER and the Golgi cannot explain the polarity of lipids on the epithelial cell surface. The only synthetic enzyme activity on the cell surface that is relevant in this respect is the phosphocholine exchange between PC and SM found on the basolateral surface (van Helvoort et al., 1994). If there would be a supply of diacylglycerol in the plasma membrane, SM would be converted to PC specifically on the basolateral surface, which would result in a basolateral enrichment of PC and a loss of basolateral SM. Indeed, a higher turnover was reported for the basolateral SM in proximal kidney epithelia. However, these cells did not contain a particularly low level of basolateral SM, but an exceptionally high concentration of SM in the apical membrane of 35 to 45% (Schwertz et al., 1983; Molitoris and Simon, 1985). This would rather favor domainmediated sorting. An alternative mechanism of generating lipid polarity would be the selective hydrolysis of lipids on the cell surface opposite to where they are enriched. There is no evidence for hydrolysis of glycosphingolipids on the basolateral surface, nor for a difference in PC turnover in apical and basolateral membranes (Molitoris and Simon, 1985).

14.5.2 LIPID TRANSPORT

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SORTING: GENERAL PRINCIPLES

Lipids are transported between the various cellular locations by a variety of mechanisms. With the exception of mitochondria and peroxisomes the cellular organelles are connected by vesicular traffic. A typical half-time for a vesicular transport step is 2 to 5 min (Griffiths et al., 1989). Transport of lipids within the membrane of a vesicle can occur in either of both leaflets, and the transbilayer distribution of the donor membrane should determine the localization of the lipids within the acceptor membrane. Lipids of the cytosolic leaflet are inserted into the cytosolic leaflet of the target membrane, and lumenal lipids end up in the external membrane leaflet. Lipids can also exchange between the cytosolic surface of cellular organelles as monomers. For most membrane lipids, this is a slow process with half-times in the order of hours (Brown, 1992). The rate is inversely correlated to the hydrophobicity

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of the lipid, and can be enhanced by having close membrane contacts or by the involvement of transfer proteins. Within an individual membrane, lipids diffuse laterally in the plane of the membrane and transversely from one leaflet into the other leaflet. Whereas lipids diffuse all over the cell surface in less than a minute, spontaneous translocation across a lipid bilayer is generally slow (hours) unless mediated by facilitators or stimulated by translocators. If there were no selectivity in the various transport processes the lipid composition of cellular membranes would be random. To generate and maintain the surface polarity of lipids, lipid traffic must be selective. Lipids must be sorted during transport. Vesicular traffic (Figure 14.3a) involves the budding of a membrane vesicle from a donor compartment and subsequent fusion with a target membrane. Two elements can be discriminated that convey specificity to this process. One, components that must be transported by different pathways emanating from a common compartment are laterally segregated. Two, the increase in surface density of a component is coupled to the cytoplasmic coat components that provide specificity for the next step, the targeting of the transport vesicle to the correct membrane. After the association between the membrane cargo and the proper cytosolic proteins, budding occurs, and the resulting transport vesicle contains all information needed for targeting, docking, and fusion with the correct membrane. The monomeric exchange of lipids (Figure 14.3b) between two membranes would be expected to result in equilibration of the lipid classes involved (Brown, 1992). However, exchange can lead to an enrichment of a lipid in one membrane if that membrane has a higher affinity for the lipid. For example, because of a higher affinity for cholesterol, sphingolipid-rich membranes can accumulate this lipid via monomeric exchange (Wattenberg and Silbert, 1983), and a loss of sphingolipids results in the subsequent loss of cholesterol (Slotte and Bierman, 1988). Alternatively, one of the two membranes could act as a sink for this lipid if it contained a lipid converting activity, as in the case of PS transport to the mitochondria by exchange followed by decarboxylation (Heikinheimo and Somerharju, 1998). Yet another possibility by which monomeric transport could be unidirectional is that the target membrane contains a translocator that eliminates the lipid from the cytosolic surface by pumping it into the noncytoplasmic leaflet. A nonphysiological example of the latter is the monomeric transfer of multiple short-chain lipids from the ER to the plasma membrane and translocation by multidrug transporters of the ABCtransporter family (van Helvoort et al., 1996; Raggers et al., 1999).

14.5.3 VESICULAR TRAFFIC TO THE APICAL AND BASOLATERAL PLASMA MEMBRANE DOMAINS (FIGURE 14.3A) 14.5.3.1 Sorting by Lateral Segregation and Coupling to Specific Membrane Coats After synthesis in the ER and the Golgi, proteins and lipids are transported to the two plasma membrane domains by the secretory pathway. The membrane units that are shipped to the apical and basolateral plasma membrane domain are not only small vesicles but also larger tubulovesicular structures undergoing continuous

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(b)

FIGURE 14.3 Generation of cell surface lipid polarity along the biosynthetic pathway. (a) Apical and basolateral lipids are sorted along the secretory pathway. After synthesis of most lipids in the ER and Golgi, they undergo sorting within the Golgi complex. Upon movement from the cis- to the trans-side the Golgi membrane matures (1), generating transversal and lateral membrane lipid asymmetry. Certain lipids become included in the forward movement of the Golgi stack, whereas others experience retrograde transport (2). This separation is driven by lateral lipid aggregation at the lumenal surface of the Golgi and leads to the formation of sphingolipid/cholesterol microdomains/rafts and in membrane domains that are enriched in glycerophospholipids. The inclusion of the different domains that are either enriched in glycosphingolipids or in PC in distinct vesicle populations that are destined to fuse either with the apical (3) or the basolateral (4) plasma membrane can generate cell surface lipid polarity (van Meer and Simons, 1986; 1988; van Meer et al., 1987; Simons and van Meer, 1988; Simons and Ikonen, 1997). (b) Lipids are also transported between organelles on the cytoplasmic surface of vesicles or by monomeric exchange between membranes where transfer proteins could facilitate the traversal of the membrane interspace. To reach the cell surface, these lipids must be transported across the plasma membrane bilayer. The restriction of the translocating activities to the apical (2) or basolateral membrane (3), respectively, could generate cell surface lipid polarity. The demonstration of natural GlcCer outward transport by MDR1 P-glycoprotein (see text), which in epithelial cells is restricted to the apical surface, renders such a model realistic. This lipid outward movement would then be balanced by an inward movement (1), e.g., of PE and PS by the aminophospholipid translocase.

fusion and fission along the way (Toomre et al., 1999). Since proteins destined for both domains are still found together in the trans-Golgi network, the sorting of apical and basolateral proteins by lateral segregation has been proposed to occur in this membrane (Fuller et al., 1985). It has been proposed that glycosphingolipids segregate from PC in the trans-Golgi network by a process of self-aggregation (van Meer and Simons, 1986; van Meer et al., 1987). Higher glycosphingolipids and SM are assembled on the lumenal, exoplasmic surface of the Golgi. In addition, the differences in lipid composition reside in the exoplasmic leaflet of the plasma membrane and translocation of these lipids to the cytosolic surface seems insignificant. Therefore, lipid segregation should occur in the lumenal leaflet of the Golgi membrane.

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Over the years, the “lipid microdomain” hypothesis has been extended into a “lipid raft” hypothesis to include the membrane proteins (Simons and van Meer, 1988; Simons and Ikonen, 1997). Membrane sorting in polarized cells is best understood for the proteins targeted to the basolateral membrane. Here, peptide motifs have been recognized in the cytoplasmic domains of membrane proteins that resemble clathrin-coated pit determinants specifying endocytosis. The amino acid-based basolateral targeting signals rely on critical tyrosine or dileucine motifs, which are frequently followed by a cluster of acidic residues. Clathrin-coated pit signals are recognized by adaptor complexes, heterotetramers of different adaptins, that are linked to the clathrin vesicle coat. Different types of adaptor complexes mediate specificity: AP-2 is involved in endocytosis, whereas at the trans-Golgi network and/or endosome AP-1 and AP-3 are thought to mediate lysosomal targeting. AP-4 is a new member of the growing family of adaptor complexes. It exhibits a ubiquitous expression pattern but specificity for a trafficking pathway is still unknown (Kirchhausen, 1999). Generally, µ adaptins interact with tyrosine-based sorting determinants and β subunits with dileucine signals. A specific adaptor complex has been implicated in basolateral sorting. Pig kidney-derived LLC-PK1 cells do not express the epithelial-specific AP-1 subunit µ1B and missort basolateral proteins with tyrosine-based sorting signals to the apical cell surface. Correct sorting was restored by expression of the specific subunit (Fölsch et al., 1999). Thus, the two steps of the sorting of basolateral proteins in the trans-Golgi network, aggregation and inclusion in a transport vesicle containing the coat machinery for basolateral targeting, are mediated by the same coat proteins. Apical protein targeting is less clearly defined. In addition to cytoplasmic domain determinants, a variety of signals have been proposed to be relevant such as the lumenal protein modifications, N- and O-linked carbohydrates, and membrane-linked determinants such as the transmembrane domain or lipidic protein anchors. In contrast to basolateral sorting, where lateral aggregation relies on the interactions between proteins and a specific vesicle coat, apical recognition has been proposed to involve the sphingolipid/cholesterol “rafts” (Simons and Ikonen, 1997). The formation of these rafts is based on the lateral mobility of membrane components and the mutual attraction between their constituents. By their aggregation sphingolipids and cholesterol exclude PC. Apical proteins partition into the apical precursor membrane domain by direct lipid–lipid interactions, like GPI-anchored proteins (Brown and London, 1998a), acylated proteins (Melkonian et al., 1999), or a proteolipid (Cheong et al., 1999; Puertollano and Alonso, 1999). Other proteins can then be dragged into the domain by protein–protein interactions. Basolateral proteins may be excluded from the rafts by lack of interaction and, as a consequence, expulsion. The importance of lipids for protein sorting has become apparent in studies where a decrease in sphingolipid (Mays et al., 1995) or cholesterol (Keller and Simons, 1998) content abolished sorting of apical proteins. It has been proposed that the proteolipid MAL is an integral part of the apical sorting machinery (Cheong et al., 1999; Puertollano et al., 1999). Although little is known concerning the coat machinery that recognizes the apical precursor domain in the TGN and is responsible for targeting towards the apical plasma membrane domain, apical exocytosis involves specific members of the typical SNAP/SNARE machinery (Gaisano et al., 1996;

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Low et al., 1996). This was also specifically demonstrated for the raft pathway (Lafont et al., 1999). The formation of sphingolipid/cholesterol rafts in the Golgi has been proposed to underlie the cisternal maturation from the cis- to the trans-side of the organelle (Holthuis et al., 2001). Sphingolipids and cholesterol become included in the forming domains, whereas unsaturated lipids, such as glycerophospholipids, do not fit properly into the lateral membrane scaffolds and are excluded from the maturing cisterna by retrograde transport. During its anterograde movement, the cisternal membrane becomes more rigid and thicker and displays more and more the structure of the plasma membrane. The morphological changes within the Golgi stack had already been documented in excellent electron micrographs in 1966 but at that time the appearance of thickened luminal membrane patches in the late Golgi cisternae could not be interpreted (Hicks, 1966). 14.5.3.2 Transport, Docking, and Fusion After the transport vesicle has budded from the donor membrane, its coat components contain all information for targeting. Microtubule motors are known to ferry the vesicles from the trans-Golgi network to the two plasma membrane domains (Lafont et al., 1994). Specificity of the subsequent docking and fusion is mediated by recognition between proteins on the vesicle and the plasma membrane. Candidates are SNAREs, cognate receptors that are found on vesicles (v-SNARE) and on the target membrane (t-SNARE). Since the receptor components have distinct polarized distributions in several epithelial cell types (Low et al., 1996; Weimbs et al., 1997), they could specify delivery of transport vesicles to the apical and basolateral membrane. However, distribution of the receptor isoforms were found different but not restricted to a single membrane domain (Fujita et al., 1998). Recent studies identified a different protein complex, Sec6/8, that upon cell polarization was recruited to the regions of cell–cell contact and established a cue for the fusion of basolateral vesicles with the plasma membrane (Grindstaff et al., 1998; Charron et al., 2000). Membrane fusion at the targeting patch could thus generate the basolateral membrane domain. The homologous complex in yeast localizes at the site of membrane expansion, the membrane area where the bud forms. Other factors, such as the small GTPases rab8 (Huber et al., 1993), rab13 (Zahraoui et al., 1994), and rab3B (Weber et al., 1994), were also shown to orient toward the cell–cell contact sites and could be involved in polarized vesicle trafficking. Interestingly, it has been reported that also an apical protein was inserted at the TJ of MDCK cells (Louvard, 1980). This raises the possibility that the TJ complex itself contains topological information that directs docking and fusion of apical and basolateral membrane vesicles on either side of the barrier.

14.5.4 MONOMERIC LIPID TRANSPORT POLARITY (FIGURE 14.3B)

AND

CELL SURFACE LIPID

Potentially, lipids can also desorb from the cytosolic leaflet of one membrane and integrate into the cytosolic leaflet of a different membrane. This is favored with increasing hydrophilicity of the desorbing membrane component and decreasing

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aqueous distance that has to be bridged. Membrane contact zones between the ER and the outer membrane of the mitochondrion appear to be involved in translocation of PS from the ER to the outer membrane of the mitochondrion (Daum and Vance, 1997). However, the lipids that display a polarized distribution are not very hydrophilic and the trans-Golgi network is not close to either plasma membrane domain. Importantly, the transport of lipid monomers can be facilitated by their binding to “lipid transfer proteins.” These have been found for PI (Li et al., 2000), PC (van Helvoort et al., 1999), and glycolipids (Lin et al., 2000). Indeed, both PC (Kaplan and Simoni, 1985) and GlcCer (Warnock et al., 1994) seem to be transported to the plasma membrane on a timescale and under conditions incompatible with vesicular traffic. However, it is unlikely that monomeric exchange by itself contributes to the enrichment of lipids on the apical or the basolateral surface for the following reasons. Lipids are transferred between cytosolic surfaces that are continuous between the apical and basolateral domains. This implies that lipids inserted from the cytosol into the apical membrane instantly mix with lipids inserted into the basolateral domain. In the case of the PC transfer protein, this protein always carries a PC molecule, and it is unclear how this protein could cause unidirectional transport. The latter is not the case for the glycolipid transfer protein, which may therefore be involved in net mass transfer (Sasaki and Demel, 1985; Mattjus et al., 2000).

14.5.5 TRANSMEMBRANE LIPID ASYMMETRY AND CELL SURFACE LIPID POLARITY (FIGURE 14.3B) 14.5.5.1 The Aminophospholipid Translocase PC is enriched on the basolateral cell surface. As described above, this is thought to be due to a lateral segregation between sphingolipids and PC in the lumenal leaflet of the trans-Golgi network and inclusion of PC into basolateral transport vesicles. Why PC and not PE or PS? Based on the present literature, two scenarios seem feasible. One possibility is that after synthesis in the cytosolic surface of the ER, PC but not PE or PS can equilibrate across the ER membrane, for example, via a PC-specific “flippase” in the ER membrane (Bishop and Bell, 1985; Backer and Dawidowicz, 1987; Menon et al., 2000). The second possibility is that all newly synthesized phospholipids can equilibrate across the ER membrane (Zilversmit and Hughes, 1977; Herrmann et al., 1990; Buton et al., 1996), and that a specific energyrequiring translocator is required to remove PS and PE from the exoplasmic leaflet. Indeed, an aminophospholipid translocase activity has been demonstrated by numerous investigators (Seigneuret and Devaux, 1984; Daleke and Lyles, 2000), and a candidate protein has been cloned. It belongs to the family of P-type ATPases (Tang et al., 1996). The activity has been found in both the apical and basolateral plasma membrane domain of human intestinal and dog kidney epithelial cells. Surprisingly, the activity in kidney-derived MDCK cells was not specific for PS and PE but also translocated PC (Pomorski et al., 1999). It is at present unclear at what stage of the secretory pathway the aminophospholipid translocase becomes active. One of the yeast homologues of the bovine P-type ATPase, Drs2p, has been localized in the Golgi (Chen et al., 1999).

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14.5.5.2 ABC-Transporters A third potential mechanism to enrich PC in the exoplasmic leaflet of the plasma membrane was discovered when studying PC secretion into bile. PC was found to be missing from the bile in mice carrying null-alleles for the ATP-binding cassette (ABC) transporter MDR3 P-glycoprotein (Smit et al., 1993). This led to the conclusion that the MDR3 P-glycoprotein acts as a PC pump or a PC translocator across the bile canalicular membrane, the apical plasma membrane of the hepatocyte (Smith et al., 1994). The protein is not involved in the enrichment of PC on the basolateral surface of epithelia, as it is virtually only expressed in the apical membrane of hepatocytes (Smith et al., 1994). The localization of complex glycosphingolipids and SM in the lumenal, exoplasmic leaflet of Golgi membranes results from the fact that they are synthesized on the lumenal side of the Golgi membrane and do not translocate across the Golgi membrane (Jeckel et al., 1992; Lannert et al., 1994; Burger et al., 1996). This is not the case for GlcCer, the simple glycolipid that constitutes a large fraction of the apical glycosphingolipids especially in rodents (Kawai et al., 1974). GlcCer is synthesized on the cytosolic surface of the Golgi. By an unknown mechanism, it can translocate to the lumenal leaflet of the Golgi membrane (Lannert et al., 1994; Burger et al., 1996), where it can be converted to higher glycosphingolipids and/or segregate into glycosphingolipid/cholesterol rafts. However, an alternative pathway by which GlcCer could reach the outer leaflet of the plasma membrane would be transport on the cytosolic surface of transport vesicles or by monomeric transport into the cytosolic surface of the plasma membrane followed by translocation across the lipid bilayer. If the translocator were exclusively present in the apical domain, this process would result in the selective transport of GlcCer to the apical surface. No selectivity would be required at the level of transport between the Golgi and the plasma membrane. This is no longer just a theoretical possibility as it has been found that MDR1 P-glycoprotein, a ubiquitous apical ABC-transporter known to pump amphiphilic molecules out of the cell, has been reported capable of translocating analogues of GlcCer specifically across the apical membrane (van Helvoort et al., 1996). It has now been found to be able to translocate natural, long-chain GlcCer (R. J. Raggers, D. J. Sillence, J. Wijnholds, P. Borst, J. M. F. G. Aerts, K. Sandhoff, G. van Meer, in preparation). It will be interesting to know what fraction of the apical GlcCer had crossed the Golgi membrane and reached the apical membrane via sorting in the TGN and what fraction translocated across the plasma membrane. The regulation of these processes may be an important parameter in generating the cell surface polarity of other molecules.

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15

Physiological Regulation of Tight Junction Permeability by Na+Nutrient Cotransport Jerrold R. Turner and James L. Madara

CONTENTS 15.1 Introduction .................................................................................................333 15.2 Dynamic Regulation of Tight Junction Permeability.................................334 15.2.1 Physiological Regulation of Tight Junction Permeability in Mammalian Small Intestine...........................................................334 15.2.2 In Vivo Evidence for Regulation of Paracellular Absorption by Na+-Glucose Cotransport..........................................................336 15.3 Proximal Signals Linking Na+-Glucose Cotransport to Myosin LightChain Phosphorylation................................................................................337 15.4 Distal Events in the Na+-Glucose Cotransport to Tight Junction Signaling Pathway ......................................................................................339 15.5 Cytoskeletal Regulation of the Tight Junction — A Common Final End Point?...................................................................................................341 15.6 A Unified Model of Na+-Nutrient Cotransport-Dependent Tight Junction Regulation ....................................................................................343 Acknowledgments..................................................................................................343 References..............................................................................................................344

15.1 INTRODUCTION In the mammalian gastrointestinal tract, a primary function of the tight junction (TJ) is to form the barrier that restricts the flux of molecules between the lumen and the interstitium along the paracellular pathway. In the small intestine, both transcellular and paracellular absorption of nutrients and water contribute to transport across the epithelial barrier (see Chapter 4 by Reuss). Although transcellular transport across cell membranes is accomplished by specific transporters, paracellular absorption occurs across the TJ. However, the small intestine is also charged with maintaining 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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a protective barrier separating the lumen, a potentially toxic milieu, from the interstitium. Thus, TJ permeability must be precisely tuned to permit water and nutrient absorption while preventing passage of toxic substances.

15.2 DYNAMIC REGULATION OF TIGHT JUNCTION PERMEABILITY Early studies presumed that TJ permeability was fixed. This assumption was modified after the demonstration of increases in Necturus gallbladder transmucosal resistance following exposure to cyclic AMP analogues (Duffey et al., 1981). Both charge selectivity and ultrastructure of TJs changed in parallel with resistance changes, suggesting that increases in resistance were due to decreased TJ permeability. Further studies showed that in Necturus gallbladder the number of TJ strands increased along with transmucosal resistance following treatment with Ca2+ ionophore (Palant et al., 1983). Thus, it became clear that TJ permeability was not static, but could be regulated by intracellular signaling molecules.

15.2.1 PHYSIOLOGICAL REGULATION OF TIGHT JUNCTION PERMEABILITY IN MAMMALIAN SMALL INTESTINE In 1987, Pappenheimer, Madara, and Reiss proposed a new theory of small intestinal nutrient absorption. They posited that the presence of Na+-cotransported nutrients, such as glucose and some amino acids, triggered the opening of TJs to allow mass transport of solutes through these paracellular channels (Madara and Pappenheimer, 1987; Pappenheimer, 1987; Pappenheimer and Reiss, 1987). This theory suggests that when luminal nutrient concentrations are high, such as after a meal, a primary purpose of active transcellular transport is to trigger increases in TJ permeability and to provide the osmotic force necessary to drive paracellular flow (Madara and Pappenheimer, 1987; Pappenheimer, 1987; Pappenheimer and Reiss, 1987). The latter occurs as a result of the deposition of the Na+-cotransported nutrient and Na+ ions in the subjunctional basolateral space following coordinated apical and basolateral transmembrane transporter. Consequently, solvent drag, the paracellular absorption of water and water soluble nutrients, becomes a major mechanism of mass transport of hydrophilic nutrients (Madara and Pappenheimer, 1987; Pappenheimer, 1987; Pappenheimer and Reiss, 1987) (Figure 15.1). The model proposed by Pappenheimer, Madara, and Reiss is also consistent with a variety of other observations. First, in vivo human data show that, at up to 500 mM, intestinal glucose absorption is nearly proportional to the luminal concentration (Cummins, 1952; Holdsworth and Dawson, 1964; Gisolfi et al., 1992) and that glucose absorption from perfused rat intestine is fourfold greater at 246 than at 28 mM (Fullerton and Parsons, 1956). These observations are inconsistent with the reported Km of 0.11 mM for the cloned intestinal Na+-glucose cotransporter SGLT1 (Hediger et al., 1987; Ikeda et al., 1989). Additional studies confirmed this discrepancy for humans and showed that rates of glucose ingestion and absorption by mice, rabbits, and rats also exceeded the maximal rates of active transmucosal transport by as much as fivefold (Fullerton and Parsons, 1956; Pappenheimer, 1990). Thus,

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FIGURE 15.1 Amplification of active transcellular by solvent drag-mediated paracellular nutrient absorption. As glucose is absorbed transcellularly, the paracellular pathway opens and becomes more permeable to small nutrients. The basolateral deposition of Na+ and nutrients establishes an osmotic gradient that provides the force for water flow. Small nutrients can then be carried across the paracellular pathway with the water by solvent drag. This paracellular transport is facilitated by opening of TJs following transcellular Na+-nutrient cotransport. Thus, total glucose absorption continues to increase well after transcellular absorption achieves Vmax. Drawn after Pappenheimer (1993) and Pappenheimer and Reiss (1987).

it is reasonable to suppose that a second paracellular mechanism of glucose absorption must amplify transcellular nutrient absorption. Experimentally, Pappenheimer and Reiss (1987) showed that addition of 25 mM glucose to the mucosal surface of rat small intestine mounted in Üssing chambers caused a 193% increase in the paracellular absorption of creatinine. Notably, this exceeds the 91% increase in fluid absorption, a result that is best explained by the apparent increase in pore radius of the TJ (Pappenheimer and Reiss, 1987). Parallel electrophysiological analyses showed that addition of glucose or SGLT1-transported glucose analogues resulted in two- to threefold decreases in transmucosal impedance (Pappenheimer, 1987). This was accompanied by simultaneous increases in capacitance and conductance (Pappenheimer, 1987). The latter indicate increases in membrane surface area and width of intercellular junctions, respectively (Pappenheimer, 1987). Consistent with this observation, recent mathematical modeling studies of the epithelial barrier have also demonstrated increased numbers of small paracellular pores during active Na+-glucose cotransports (Fihn et al., 2000). Similar effects on impedance, capacitance, and conductance were noted when other Na+-coupled amino acid transporters were activated (Pappenheimer, 1987). These data suggest that Na+ cotransport of several luminal nutrients can trigger increases in TJ channel pore size, i.e., permeability, and is consistent with increased paracellular absorption due to solvent drag. The necessity for Na+-nutrient cotransport in TJ regulation was confirmed by subsequent studies showing that regulation of transmucosal resistance did not occur in the absence of extracellular Na+ (despite the presence of apical glucose) and was prevented by the specific Na+-glucose cotransporter inhibitor phloridzin (Atisook et al., 1990). Decreases in transmucosal resistance were also induced by the Na+cotransported amino acid alanine, even in the presence of phloridzin (Atisook et al.,

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FIGURE 15.2 Na+-glucose cotransport induces intra-TJ dilatations with increased luminal access. (A) Initiation of Na+-glucose cotransport in rodent small intestine results in the development of small dilatations within the TJ region (arrow). (B) When the small hemeconjugated oligopeptide MP-11, which retains peroxidase activity, is added to the luminal (apical) surface, it is seen to gain access to the normally restricted intra-TJ region and accumulates within TJ dilatations (arrow).

1990). This effect of apical Na+-nutrient cotransport on TJ permeability, as assessed by transmucosal resistance, did not occur as a consequence of experimentally induced hyperglycemia (See and Bass, 1993), suggesting that the presence of luminal, rather than basolateral, Na+ and nutrients was necessary for this regulation to occur. Electron microscopic examination of TJ structure during Na+-glucose cotransport demonstrated striking dilatations within the TJ (Madara and Pappenheimer, 1987; Pappenheimer, 1987; Atisook et al., 1990; Atisook and Madara, 1991; Madara and Carlson, 1991; Pappenheimer and Volpp, 1992) (Figure 15.2). Consistent with the demonstrated alterations in TJ permeability, these dilatations are penetrated by an oligopeptide tracer applied to the apical (luminal) surface, thus confirming that the TJ is the site of increased intestinal permeability (Figure 15.2) (Atisook and Madara, 1991). These observations demonstrate a clear relationship between ultrastructural and functional responses of the TJ to Na+-nutrient cotransport. An immunoelectron microscopic study of the distribution of the TJ protein ZO-1 during glucose-induced regulation of paracellular permeability showed that a spatial dissociation between ZO-1 and the morphologically identified TJ occurs after Na+-glucose cotransport (Madara et al., 1993). This suggests that the biochemical association between ZO-1 and the junctional fibrils is modified during physiological regulation of TJ permeability (see Chapter 11 by Citi and Chapter 12 by Fanning).

15.2.2 IN VIVO EVIDENCE FOR REGULATION OF PARACELLULAR ABSORPTION BY NA+-GLUCOSE COTRANSPORT Although some controversy has surrounded the in vivo relevance of Na+-nutrient cotransport-dependent TJ regulation (Fine et al., 1993; 1994; Soergel, 1993; Madara, 1994; Schwartz et al., 1995; Turner and Madara, 1995; Uhing and Kimura, 1995a; Lane et al., 1999), the original reports of Na+-glucose cotransport-dependent regulation of small intestinal permeability included studies in unanesthetized rats (Pappenheimer

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and Reiss, 1987). In those studies unanesthetized rats were starved for 24 h and then given 50 to 100 mg of creatinine by gastric intubation (Pappenheimer and Reiss, 1987). The rats were then fed water or water with 20% glucose, respectively, ad libitum. Since creatinine is absorbed by the paracellular pathway, is confined to the extracellular space, is not metabolized, and is freely filtered by the glomerulus, measurements of urinary creatinine recovery serve as a surrogate for measurements of intestinal creatinine absorption. In rats with access to glucose, 53 ± 2.4% of the exogenous creatinine load was recovered within 15 h, whereas only 37 ± 2.3% of creatinine was recovered in rats receiving only water (Pappenheimer and Reiss, 1987). Thus, the data obtained from the in vitro studies of isolated mucosa also appear to be representative of in vivo events. The authors recently reported the results of a similar study assessing nutrientinduced augmentation of paracellular absorption in humans (Turner et al., 2000b). Essential criteria for the experimental approach included (1) complete glucose absorption (Madara, 1994), (2) the absence of surgical manipulation and anesthesia (Uhing and Kimura, 1995b), and (3) assay of a significant intestinal length (Madara, 1994). Molecular probes of appropriate size were employed to allow detection of changes in paracellular absorption. Both the radius (3.2 Å) and molecular weight (113.12 g) of creatinine are similar to those of glucose (3.7 Å, 180.16 g) and other small nutrients. It was therefore concluded that creatinine would be an appropriate probe to measure changes in intestinal paracellular permeability induced by luminal glucose in humans. Sufficient creatinine was administered to render endogenous creatinine insignificant. When creatinine was ingested with glucose, urinary creatinine recovery was 55 ± 4% of ingested creatinine. In contrast, urinary creatinine recovery was only 38 ± 9% of ingested creatinine when ingested without an Na+cotransported nutrient (p < 0.01). Thus, human intestinal paracellular absorption is increased by the presence of luminal glucose, consistent with in vivo regulation of intestinal permeability by Na+-glucose cotransport (Turner et al., 2000b).

15.3 PROXIMAL SIGNALS LINKING NA+-GLUCOSE COTRANSPORT TO MYOSIN LIGHT CHAIN PHOSPHORYLATION Further progress on the mechanisms by which TJ permeability is regulated following initiation of Na+-glucose cotransport were significantly limited by the technical challenges inherent in studies using isolated mammalian mucosa. To circumvent these problems, an in vitro model of physiological Na+-glucose cotransport-dependent TJ regulation was established using the human intestinal epithelial cell line Caco-2 (Turner et al., 1997). The Caco-2 cell line was selected because it differentiates with a small intestinal absorptive cell phenotype (Pinto et al., 1983) and a well-developed brush border (Peterson and Mooseker, 1992). The authors stably transfected Caco-2 cells to express the intestinal Na+-glucose cotransporter, SGLT1, and showed that SGLT1 expressed in these cells had a Km of 0.31 mM, a KNa 43 meq/l, and a Hill coefficient of 1.96 (Turner et al., 1996), comparable to that of native intestinal SGLT1. Moreover, in differentiated monolayers, SGLT1 protein was apically polarized and was able to mediate vectorial Na+ and glucose cotransport (Turner et al., 1996).

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FIGURE 15.3 Na+-glucose cotransport-dependent TJ regulation in cultured cell monolayers. SGLT1-transfected Caco-2 monolayers were cultured overnight with 0.5 mM phloridzin, a specific SGLT1 inhibitor. Glucose was added (and Na+-glucose cotransport activated) by transfer to HBSS with 25 mM glucose (filled circles) and resulted in progressive decreases in transepithelial resistance for the first 90 min, after which transepithelial resistance stabilized. In contrast, after transfer to HBSS with 5 mM glucose and 2 mM phloridzin (closed circles) transepithelial resistance remained high.

Monolayers of these Caco-2 cells exhibit reversible regulation of TJ permeability following activation or inhibition of SGLT1-dependent Na+-glucose cotransport (Turner et al., 1997). For example, activation of SGLT1 induces an approximately 25% decrease in transepithelial resistance (Figure 15.3). Moreover, Üssing chamber studies showed that decreases in transepithelial resistance after initiation of Na+glucose cotransport were accompanied by increases in transepithelial permeability to mannitol, but not to the larger tracer inulin (Turner et al., 1997), suggesting that the Na+-glucose cotransport-dependent increase in permeability is due to regulation of TJ pores. Furthermore, since mannitol but not inulin flux was affected, the change is size selective, consistent with increased permeability to small nutrient-sized molecules but not to larger substances (Turner et al., 1997). Thus, this transfected Caco-2 cell model appears to reflect accurately Na+-nutrient cotransport-dependent TJ regulation in vitro, in isolated native mucosa, as well as in vivo. This cultured cell model of Na+-glucose cotransport-dependent TJ regulation has been used to characterize early events in the signaling pathway linking these events (Turner et al., 2000a). Cell swelling is a well-recognized consequence of SGLT1mediated Na+-glucose cotransport (MacLeod and Hamilton, 1991). Moreover, the degree to which activation of various Na+-solute cotransporters results in altered TJ permeability correlates with variation in the ability of the cotransporters to induce cell swelling (i.e., with rates of internalization) (Pappenheimer and Volpp, 1992). Thus, it was suggested that cell swelling may be the trigger that links Na+ cotransport of differing nutrients and results in the common end point of increased TJ permeability (Pappenheimer and Volpp, 1992; Turner and Madara, 1995). Following cell swelling, a regulatory volume decrease response normalizes cell volume within minutes (MacLeod and Hamilton, 1991). Although the signaling events involved in

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regulatory volume decrease induced by various stimuli differ (MacLeod et al., 1992), it has been shown that regulatory volume decrease after hypotonic swelling of enterocytes requires cytoplasmic alkalinization (MacLeod and Hamilton, 1996). This alkalinization appears to be mediated by Na+-H+ exchange, since both alkalinization and regulatory volume decrease can be prevented by Na+-H+ exchange inhibitors (MacLeod and Hamilton, 1996). Thus, the regulation of Na+-H+ exchange by Na+glucose cotransport was explored. Initiation of Na+-glucose cotransport was found to cause a mild cytoplasmic alkalinization and this alkalinization required both extracellular Na+ and glucose (Turner and Black, 2000). Evaluation of the SGLT1-transfected Caco-2 cells used showed that these cells express all three intestinal Na+-H+ exchanger isoforms, NHE1, NHE2, and NHE3. NHE1 is a basolaterally localized Na+-H+ exchanger that is expressed ubiquitously. In contrast, NHE2 and NHE3 are localized to the apical brush border and are expressed in only a limited number of cell types. To determine which, if any, of these Na+-H+ exchanger isoforms was involved in cytoplasmic alkalinization after Na+-glucose cotransport, the ability of relatively isoform-specific Na+-H+ exchange inhibitors to prevent alkalinization was examined. It was found that the preferential NHE3 inhibitor S3226 prevented alkalinization at 1 µM, consistent with inhibition of NHE3 (Ki = 0.02 to 0.2 µM), but not NHE1 (Ki = 3.5 µM) or NHE2 (Ki = 80 µM). Moreover, HOE694, which preferentially inhibits NHE1 and NHE2 (Ki values of 0.16 and 5 µM, respectively) could not prevent cytoplasmic alkalinization at doses that did not significantly inhibit NHE3 (Ki = 650 µM). Thus, it was concluded that activation of NHE3 occurs following the initiation of Na+glucose cotransport. More recently, these observations have been extended to show that the osmotically responsive MAP kinase p38 is an essential intermediate in Na+glucose cotransport-dependent NHE3 activation (Turner and Black, 2000). Since NHE3 is activated following initiation of Na+-glucose cotransport, one could envision a signal transduction pathway in which NHE3 activation is an intermediate between Na+-glucose cotransport and TJ regulation. Evaluation of the effects of Na+-H+ exchange inhibitors on transepithelial resistance showed that inhibition of Na+-H+ exchange caused marked increases in transepithelial resistance (Turner et al., 2000a). Pharmacological evaluation of isoform-specific Na+-H+ exchange inhibitors, as above, showed that the brush border NHE3 isoform, but not NHE1 or NHE2, was critical for the effects of these inhibitors on transepithelial resistance (Turner et al., 2000a). Thus, it appears that NHE3 activation may be a critical component of the signaling pathway for Na+-glucose cotransport-dependent TJ regulation.

15.4 DISTAL EVENTS IN THE NA+-GLUCOSE COTRANSPORT — TIGHT JUNCTION SIGNALING PATHWAY Ultrastructural examination of intestinal mucosae provided additional clues to the mechanistic basis for Na+-nutrient cotransport-dependent changes in TJ permeability (Madara and Pappenheimer, 1987; Atisook et al., 1990; Atisook and Madara, 1991). These studies showed two primary changes in the TJ region of enterocytes with

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FIGURE 15.4 Na+-glucose cotransport induces morphological condensation of the perijunctional actomyosin ring. (A) In the absence of Na+-glucose cotransport, the perijunctional actomyosin ring (arrow) of rodent small intestinal enterocytes is indistinct (tight junction indicated by arrowhead). (B) When Na+-glucose cotransport is initiated, a striking condensation of the perijunctional actomyosin ring (arrow) adjacent to the TJ (arrowhead) and the adherens junction is evident.

Na+-glucose cotransport-induced increases in TJ permeability (Madara and Pappenheimer, 1987; Atisook et al., 1990; Atisook and Madara, 1991). First, dilatations were seen within the TJ (Madara and Pappenheimer, 1987) (Figure 15.2). These dilatations could be seen by transmission electron microscopy as lucent areas within the TJ and by freeze-fracture electron microscopy as disruptions of the TJ strands (Madara and Pappenheimer, 1987). Thus, these localized disruptions of TJ structure were considered to represent the anatomic correlate of increased TJ permeability and increased water flux (Madara and Pappenheimer, 1987). The second morphological change identified within the TJ region of glucosetreated mucosa was condensation of the perijunctional cytoskeleton (Madara and Pappenheimer, 1987; Atisook et al., 1990) (Figure 15.4). This ring of actin and myosin II encircles the apical pole of columnar epithelial cells at the level of the junctional complex. Thus, condensation of the perijunctional cytoskeleton is consistent with contraction and suggests a mechanistic linkage between the cytoskeleton and TJ permeability. Such a linkage between TJ permeability and the perijunctional actomyosin ring has also been suggested by the observations that pharmacological inhibitors of actin polymerization cause both perijunctional actomyosin ring disassembly and markedly increased TJ permeability (Bentzel et al., 1976; Madara et al., 1986) (see Chapter 12 by Fanning). The above-described cultured cell model of Na+-glucose cotransport-dependent TJ regulation has been used to evaluate a biochemical marker of actomyosin contraction, phosphorylation of the myosin II regulatory light chain. Myosin light chain phosphorylation at serine-19 by myosin light chain kinase, or, possibly, by other kinases, activates actomyosin contraction. Consistent with the hypothesis that contraction of the perijunctional actomyosin ring effects TJ regulation, it was shown that activation of SGLT1 leads to increased 32P incorporation into myosin light chain (Turner et al., 1997) and that this 32P incorporation occurs at ser-19 (Turner et al., 1999). To determine whether such increases in myosin light chain phosphorylation were mechanistically linked to changes in TJ permeability, the effects of the related

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FIGURE 15.5 Myosin II regulatory light chain phosphorylation is reduced by inhibition of Na+-glucose cotransport or myosin light chain kinase. The left three lanes show the 20-kDa region of an SDS-PAGE autoradiograph of myosin light chain from 32P-labeled Caco-2 cells. All cells were preloaded with 32P and then incubated with PO4-free media with 25 mM glucose, 25 mM glucose with 40 µM ML9, or 5 mM glucose, 20 mM mannitol, and 2 mM phloridzin (left to right, respectively). The arrow indicates the position of myosin light chain. The far right lane is an immunoblot of the phloridzin-treated monolayers (performed after the autoradiograph, as described; Turner et al., 1997).

myosin light chain kinase inhibitors ML-7 and ML-9 were evaluated (Turner et al., 1997). In monolayers of SGLT1-expressing Caco-2 cells, inhibition of myosin light chain kinase decreased both myosin light chain phosphorylation and TJ permeability despite ongoing Na+-glucose cotransport (Turner et al., 1997) (Figure 15.5). Similarly, in isolated native mucosa, inhibition of myosin light chain kinase prevented Na+-glucose cotransport-dependent TJ regulation (Turner et al., 1997). Notably, inhibition of myosin light chain kinase did not prevent the formation of intrajunctional dilatations. Thus, it is possible that the dilatations are the result of increased paracellular fluid movement (triggered by transcellular Na+ and glucose flux) and do not primarily reflect TJ regulation. Based on such observations, it appears that myosin light chain phosphorylation may be a critical intermediate in Na+-glucose cotransport-dependent regulation of intestinal epithelial TJ permeability. Moreover, it was demonstrated that inhibition of Na+-H+ exchange also reduced myosin light chain phosphorylation (Turner et al., 2000a), consistent with the hypothesis that Na+glucose cotransport-dependent activation of NHE3 is linked to increased myosin light chain phosphorylation.

15.5 CYTOSKELETAL REGULATION OF THE TIGHT JUNCTION — A COMMON FINAL END POINT? A role for myosin light chain phosphorylation in TJ regulation has also been suggested in other intestinal epithelial models. For example, in intestinal epithelial cells, myosin light chain phosphorylation occurs after colonization with enteropathogenic Escherichia coli (Yuhan et al., 1997). Reorganization of the apical cytoskeleton and increased TJ permeability occur in parallel with this myosin light chain phosphorylation and all three are prevented by inhibitors of myosin light chain kinase (Yuhan et al., 1997). However, this mechanism differs from physiological Na+-glucose cotransport-dependent TJ regulation in that the latter is more rapid, reversible, and size selective (Turner et al., 1997).

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The role of myosin light chain phosphorylation in TJ regulation has been characterized further in a study using a transfected Madin–Darby canine kidney (MDCK) cell line model. These cells were transfected with a truncated myosin light chain kinase gene construct (Hecht et al., 1996) that lacks the inhibitory domain necessary for calmodulin dependence (Itoh et al., 1989; Ito et al., 1991). Thus, the truncated myosin light chain kinase expressed in the transfected cells is constitutively active. When the transfected cells were grown as monolayers, they developed transepithelial resistance that was less than 10% of that developed in monolayers of control cells (Hecht et al., 1996). These studies may be the strongest data available to demonstrate a direct effect of myosin light chain phosphorylation on epithelial TJ permeability. However, since the truncated myosin light chain kinase was expressed continuously, this model does not allow further dissection of the effects of myosin light chain phosphorylation. For example, it is not possible to use this model to differentiate effects of myosin light chain phosphorylation on TJ assembly from effects of myosin light chain phosphorylation on permeability of the assembled TJ. A tetracycline-dependent regulated expression system has recently been developed in the Caco-2 intestinal epithelial cell line. This model was used to express a truncated myosin light chain kinase construct similar to that used in the studies of MDCK cells. Removal of doxycycline from the culture media induces expression of the transgene and a marked increase in myosin light chain kinase activity and myosin light chain phosphorylation within 18 h (Turner et al., 2000c). Coincident with these increases in myosin light chain phosphorylation, a 20 to 30% decrease in transepithelial resistance, and a comparable increase in transepithelial mannitol flux were observed (Turner et al., 2000c). These data confirm that myosin light chain phosphorylation may directly regulate the permeability of TJs in assembled monolayers. Further evidence for the specific role of myosin light chain phosphorylation in these events comes from the observation that myosin light chain kinase inhibitors rapidly increase the transepithelial resistance of monolayers expressing the truncated myosin light chain kinase to that of controls not expressing the transgene (Turner et al., 2000c). Although further details of the specific molecular interactions that link contraction of the perijunctional actomyosin ring to TJ regulation are not known, it is tempting to speculate that a multimolecular complex is involved. Likely members of this complex include ZO-1, which is both an actin-binding and cross-linking protein (Wittchen et al., 1999), as well as cingulin, which can interact with ZO-1, ZO-2, ZO-3, and enterocyte myosin heavy-chain (Cordenonsi et al., 1999). Thus, it is possible that ZO-1, ZO-2, and ZO-3 interactions with cingulin and actin, cingulin binding to myosin, and actomyosin interactions serve as bridges for the assembly of claudin- and occludin-containing complexes and mediate regulation of TJs by the perijunctional actomyosin ring (Figure 15.6). Consistent with this hypothesis, it was recently discovered that TJ proteins are present in specialized membrane microdomains with physical characteristics of detergent-insoluble glycolipid-rich membrane rafts (Nusrat et al., 2000). Moreover, regulation of TJ permeability by myosin light chain phosphorylation causes a subtle change in the physical characteristics of TJ membrane microdomains (Turner et al., 2000c). Thus, it is possible that myosin light chain phosphorylation causes both changes in TJ protein–protein interactions and reorganization of TJ membranes.

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FIGURE 15.6 Model of cytoskeletal regulation via the TJ. TJ protein complexes can bind to both actin filaments and myosin (Cordenonsi et al., 1999; Wittchen et al., 1999). When the perijunctional actomyosin ring is relaxed, i.e., when there is no active Na+-nutrient cotransport (left panel), junctional complexes on adjacent cells form closer associations that limit paracellular permeability. When perijunctional actomyosin ring contraction occurs, i.e., when active Na+-nutrient cotransport is initiated (right panel), both the interaction of the TJ protein complexes with actomyosin and the organization of TJ membrane microdomains are altered, resulting in increased TJ permeability.

15.6 A UNIFIED MODEL OF NA+-NUTRIENT COTRANSPORT-DEPENDENT TIGHT JUNCTION REGULATION Taken together, the studies described in this chapter provide strong evidence that the permeability of intestinal TJs can be regulated by activation of Na+-glucose cotransport. Additionally, a variety of recent studies have shed light on the intracellular mechanisms involved in this regulation (Figure 15.7). Activation of the brush-border Na+-H+ exchanger NHE3 appears to be an early event following initiation of Na+-glucose cotransport that, by as yet undetermined mechanisms, leads to increased phosphorylation of myosin light chain and contraction of the perijunctional actomyosin ring. In turn, this actomyosin contraction leads to an ill-defined reorganization of the TJ that results in increased permeability. As signaling pathways and structural events that cause TJ regulation are defined, rational approaches to diagnosis and treatment of intestinal diseases with altered mucosal permeability may become available.

ACKNOWLEDGMENTS This work was supported in part by National Institutes of Health Grants DK02503 and DK56121 (J.R.T.) and DK35932 and DK47662 (J.L.M.).

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FIGURE 15.7 Proposed signaling pathway of Na+-glucose cotransport-dependent TJ regulation. The diagram illustrates the authors’ working model of intracellular signaling events that occur following the initiation of Na+-glucose cotransport. The first events after initiation of Na+-glucose cotransport must include cell swelling. Possibly due to this cell swelling, p38 MAP kinase is activated. Further unknown intermediates may be involved or p38 MAP kinase may directly cause activation of the brush border Na+-H+ exchanger NHE3. In turn, NHE3 triggers increased activity of myosin light chain kinase (MLCK), resulting in increased phosphorylation of myosin regulatory light chain and contraction of the perijunctional actomyosin ring. This actomyosin contraction leads to modulation of TJ protein interactions and membrane microdomains, and, ultimately, causes increased TJ permeability.

REFERENCES Atisook, K., and Madara, J. L. 1991. An oligopeptide permeates intestinal tight junctions at glucose-elicited dilatations. Implications for oligopeptide absorption. Gastroenterology, 100, 719–724. Atisook, K., Carlson, S., and Madara, J. L. 1990. Effects of phlorizin and sodium on glucoseelicited alterations of cell junctions in intenstinal epithelia. Am. J. Physiol., 258, C77–C85. Bentzel, C. J., Hainau, B., Edelman, A., Anagnostopoulos, T., and Benedetti, E. L. 1976. Effect of plant cytokinins on microfilaments and tight junction permeability. Nature, 264, 666–668. Cordenonsi, M., D’Atri, F., Hammar, E., Parry, D. A., Kendrick-Jones, J., Shore, D., and Citi, S. 1999. Cingulin contains globular and coiled-coil domains and interacts with ZO-1, ZO-2, ZO-3, and myosin. J. Cell Biol., 147, 1569–1582. Cummins, A. J. 1952. Absorption of glucose and methionine from the human intestine: the influence of glucose concentrations in the blood and in the intestinal lumen. J. Clin. Invest., 31, 928–937. Duffey, M. E., Hainau, B., Ho, S., and Bentzel, C. J. 1981. Regulation of epithelial tight junction permeability by cyclic AMP. Nature, 294, 451–453.

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Fihn, B. M., Sjoqvist, A., and Jodal, M. 2000. Permeability of the rat small intestinal epithelium along the villus-crypt axis: effects of glucose transport. Gastroenterology, 119:1029–1036. Fine, K. D., Santa Ana, C. A., Porter, J. L., and Fordtran, J. S. 1993. Effect of D-glucose on intestinal permeability and its passive absorption in human small intestine in vivo. Gastroenterology, 105, 1117–1125. Fine, K. D., Santa Ana, C. A., Porter, J. L., and Fordtran, J. S. 1994. Mechanism by which glucose stimulates the passive absorption of small solutes by the human jejunum in vivo. Gastroenterology, 107, 389–395. Fullerton, P. M., and Parsons, D. S. 1956. Absorption of sugars and water from rat intestine in vivo. Q. J. Exp. Physiol., 41, 387–397. Gisolfi, C. V., Summers, R. W., Schedl, H. P., and Bleiler, T. L. 1992. Intestinal water absorption from select carbohydrate solutions in humans. J. Appl. Physiol., 73, 2142–2150. Hecht, G., Pestic, L., Nikcevic, G., Koutsouris, A., Tripuraneni, J., Lorimer, D. D., Nowak, G., Guerriero, V., Jr., Elson, E. L., and Lanerolle, P. D. 1996. Expression of the catalytic domain of myosin light chain kinase increases paracellular permeability. Am. J. Physiol., 271, C1678–1684. Hediger, M. A., Coady, M. J., Ikeda, T. S., and Wright, E. M. 1987. Expression cloning and cDNA sequencing of the Na+/glucose cotransporter. Nature, 330, 379–381. Holdsworth, C. D., and Dawson, A. M. 1964. The absoprtion of monosaccharides in man. Clin. Sci., 27, 371–379. Ikeda, T. S., Hwang, E.-S., Coady, M. J., Hirayama, B. A., Hediger, M. A., and Wright, E. M. 1989. Characterization of a Na+/glucose cotransporter cloned from rabbit small intestine. J. Membr. Biol., 110, 87–95. Ito, M., Guerriero, V., Jr., Chen, X. M., and Hartshorne, D. J. 1991. Definition of the inhibitory domain of smooth muscle myosin light chain kinase by site-directed mutagenesis. Biochemistry, 30, 3498–3503. Itoh, T., Ikebe, M., Kargacin, G. J., Hartshorne, D. J., Kemp, B. E., and Fay, F. S. 1989. Effects of modulators of myosin light-chain kinase activity in single smooth muscle cells. Nature, 338, 164–167. Lane, J. S., Whang, E. E., Rigberg, D. A., Hines, O. J., Kwan, D., Zinner, M. J., McFadden, D. W., Diamond, J., and Ashley, S. W. 1999. Paracellular glucose transport plays a minor role in the unanesthetized dog. Am. J. Physiol., 276, G789–794. MacLeod, R. J., and Hamilton, J. R. 1991. Volume regulation initiated by Na+-nutrient cotransport in isolated mammalian villus enterocytes. Am. J. Physiol., 260, G26–33. MacLeod, R. J., and Hamilton, J. R. 1996. Activation of Na+/H+ exchange is required for regulatory volume decrease after modest “physiological” volume increases in jejunal villus epithelial cells. J. Biol. Chem., 271, 23138–23145. MacLeod, R. J., Lembessis, P., and Hamilton, J. R. 1992. Differences in Ca2+-mediation of hypotonic and Na(+)-nutrient regulatory volume decrease in suspensions of jejunal enterocytes. J. Membr. Biol., 130, 23–31. Madara, J. L. 1994. Sodium-glucose cotransport and epithelial permeability. Gastroenterology, 107, 319–320. Madara, J. L., and Carlson, S. 1991. Supraphysiologic L-tryptophan elicits cytoskeletal and macromolecular permeability alterations in hamster small intestinal epithelium in vitro. J. Clin. Invest., 87, 454–462. Madara, J. L., and Pappenheimer, J. R. 1987. Structural basis for physiological regulation of paracellular pathways in intestinal epithelia. J. Membr. Biol., 100, 149–164.

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Madara, J. L., Barenberg, D., and Carlson, S. 1986. Effects of cytochalasin D on occluding junctions of intestinal absorptive cells: further evidence that the cytoskeleton may influence paracellular permeability and junctional charge selectivity. J. Cell Biol., 102, 2125–2136. Madara, J. L., Carlson, S., and Anderson, J. M. 1993. ZO-1 maintains its spatial distribution but dissociates from junctional fibrils during tight junction regulation. Am. J. Physiol., 264, C1096–1101. Nusrat, A., Parkos, C. A., Verkade, P., Foley, C. S., Liang, T. W., Innis-Whitehouse, W., Eastburn, K. K., and Madara, J. L. 2000. Tight junctions are membrane microdomains. J. Cell Sci., 113, 1771–1781. Palant, C. E., Duffey, M. E., Mookerjee, B. K., Ho, S., and Bentzel, C. J. 1983. Ca2+ regulation of tight-junction permeability and structure in Necturus gallbladder. Am. J. Physiol., 245, C203–212. Pappenheimer, J. R. 1987. Physiological regulation of transepithelial impedance in the intestinal mucosa of rats and hamsters. J. Membr. Biol., 100, 137–148. Pappenheimer, J. R. 1990. Paracellular intestinal absorption of glucose, creatinine, and mannitol in normal animals: relation to body size. Am. J. Physiol., 259, G290–299. Pappenheimer, J. R. 1993. On the coupling of membrane digestion with intestinal absorption of sugars and amino acids. Am. J. Physiol., 265, G409–417. Pappenheimer, J. R., and Reiss, K. Z. 1987. Contribution of solvent drag through intercellular junctions to absorption of nutrients by the small intestine of the rat. J. Membr. Biol., 100, 123–136. Pappenheimer, J. R., and Volpp, K. 1992. Transmucosal impedance of small intestine: correlation with transport of sugars and amino acids. Am. J. Physiol., 263, C480–493. Peterson, M. D., and Mooseker, M. S. 1992. Characterization of the enterocyte-like brush border cytoskeleton of the C2BBe clones of the human intestinal cell line, Caco-2. J. Cell Sci., 102, 581–600. Pinto, M., Robine-Leon, S., Appay, M. D., Kedinger, M., Triadou, N., Dussaulx, E., Lacroix, B., Simon-Assmann, P., Haffen, K., Fogh, J., and Zweibaum, A. 1983. Enterocytelike differentiation and polarization of the human colon carcinoma cell line Caco-2 in culture. Biol. Cell, 47, 323–330. Schwartz, R. M., Furner, J. K., and Levitt, M. D. 1995. Paracellular intestinal transport of six-carbon sugars is negligible in the rat. Gastroenterology, 109, 1206–1213. See, N. A., and Bass, P. 1993. Nutrient-induced changes in the permeability of the rat jejunal mucosa. J. Pharm. Sci., 82, 721–724. Soergel, K. H. 1993. Showdown at the tight junction. Gastroenterology, 105, 1247–1250. Turner, J. R., and Black, E. D. 2000. SGLT1-mediated Na+-glucose cotransport activates NHE3 via a p38 MAPK-dependent pathway. Gastroenterology, 118, A870 (abstract). Turner, J. R., and Madara, J. L. 1995. Physiological regulation of intestinal epithelial tight junctions as a consequence of Na+-coupled nutrient transport. Gastroenterology, 109, 1391–1396. Turner, J. R., Lencer, W. I., Carlson, S., and Madara, J. L. 1996. Carboxy-terminal vesicular stomatitis virus G protein-tagged intestinal Na+-dependent glucose cotransporter (SGLT1): maintenance of surface expression and global transport function with selective perturbation of transport kinetics and polarized expression. J. Biol. Chem., 271, 7738–7744. Turner, J. R., Rill, B. K., Carlson, S. L., Carnes, D., Kerner, R., Mrsny, R. J., and Madara, J. L. 1997. Physiological regulation of epithelial tight junctions is associated with myosin light chain phosphorylation. Am. J. Physiol., 273, C1378–1385.

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Turner, J. R., Angle, J. M., Black, E. D., Joyal, J. L., Sacks, D. B., and Madara, J. L. 1999. Protein kinase C-dependent regulation of transepithelial resistance: the roles of myosin light chain and myosin light chain kinase. Am. J. Physiol., 277, C554–562. Turner, J. R., Black, E. D., Ward, J., Tse, C.-M., Uchwat, F. A., Alli, H. A., Donowitz, M., Madara, J. L., and Angle, J. M. 2000a. Transepithelial resistance can be regulated by the intestinal brush border Na+-H+ exchanger NHE3. Am. J. Physiol., 279, 1918–1924. Turner, J. R., Cohen, D. E., Mrsny, R. J., and Madara, J. L. 2000b. Noninvasive in vivo analysis of human small intestinal paracellular absorption: regulation by Na+-glucose cotransport. Dig. Dis. Sci., 45, 2122–2126. Turner, J. R., Guerriero, V., Jr., Black, E. D., and Haelewyn, K. 2000c. Regulated expression of the myosin light chain kinase catalytic domain increases paracellular permeability and alters tight junction structure. Gastroenterology, 118, A432 (abstract). Uhing, M. R., and Kimura, R. E. 1995a. Active transport of 3-O-methyl glucose by the small intestine in chronically catheterized rats. J. Clin. Invest., 95, 2799–2805. Uhing, M. R., and Kimura, R. E. 1995b. The effect of surgical bowel manipulation and anesthesia on intestinal glucose absorption in rats. J. Clin. Invest., 95, 2790–2798. Wittchen, E. S., Haskins, J., and Stevenson, B. R. 1999. Protein interactions at the tight junction. Actin has multiple binding partners, and ZO-1 forms independent complexes with ZO-2 and ZO-3. J. Biol. Chem., 274, 35179–35185. Yuhan, R., Koutsouris, A., Savkovic, S. D., and Hecht, G. 1997. Enteropathogenic Escherichia coli-induced myosin light chain phosphorylation alters intestinal epithelial permeability. Gastroenterology, 113, 1873–1882.

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Extracellular Macromolecules Modulate Epithelial Permeability Simon A. Lewis

CONTENTS 16.1 Introduction .................................................................................................349 16.2 The Epithelium as a Barrier .......................................................................350 16.3 Alteration by Proteases...............................................................................353 16.3.1 Physiological Role of Proteases ....................................................353 16.3.2 Pathophysiological Role of Proteases ...........................................354 16.4 Xenobiotics .................................................................................................355 16.4.1 Nonbacterial...................................................................................355 16.4.2 Bacterial .........................................................................................356 16.5 Modulation by Leukocytes .........................................................................357 16.5.1 Eosinophil Proteins........................................................................358 16.5.2 Neutrophil Proteins........................................................................359 16.6 Modulation by Cellular Constituents .........................................................360 16.6.1 Protamine .......................................................................................360 16.6.2 Histones..........................................................................................361 16.7 Cationic Proteins.........................................................................................362 16.8 Summary .....................................................................................................363 Acknowledgments..................................................................................................364 References..............................................................................................................364

16.1 INTRODUCTION Epithelia perform two distinct functions that are required for the survival of an organism. First, they actively transport electrolytes and nonelectrolytes between the two compartments formed by the epithelium. Second, they selectively restrict the movement of substances between the two compartments. One of the compartments formed by the epithelium is in close contact with a blood supply and is the serosal 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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or plasma compartment. In many instances, the other compartment is in contact with the outside environment and is the lumen or the mucosal compartment. Net movement of substances from the lumen to the plasma compartment is absorption, while movement in the opposite direction is secretion. Net absorption or secretion can be passive, occurring down electrical or chemical gradients, or active, requiring metabolic energy. The structure of the epithelium allows it to perform these two important functions. Architecturally an epithelium is a sheet of cells resting on a basement membrane, with the cells joined together along one edge by a continuous band of proteins, i.e., the tight junction (TJ). The TJ separates the membrane of the cell into two domains: (1) the apical, or luminal, domain, which is in series with the (2) basolateral, or serosal, domain. Movement of substances across the epithelium then has two pathways, one through the cell (transcellular pathway) and the other between the cells (paracellular pathway). Absorption via the transcellular pathway requires the substance to enter the cell from the lumen across the apical membrane and exit the cell across the basolateral membrane into the serosal compartment. Movement of substances through the paracellular pathway requires that the substance diffuse across the TJs and the series lateral intercellular space. The lateral membranes of the individual epithelial cells bound the lateral intercellular space. Alteration of the properties of either of these pathways can change the direction and magnitude of the movement of substances between the two compartments. There are numerous studies investigating the cellular mechanisms for the regulation of electrolyte and nonelectrolyte transport. Transcellular movement of an electrolyte or nonelectrolyte requires that the two series membranes (apical and basolateral) have complementary but different transport proteins. Regulation of transcellular transport can occur by the activation/inactivation, insertion/withdrawal, or modulation of membrane transporters as well as an energy supply to the primary active transporters. The signal for an alteration in cellular transport can be either extracellular (hormones, neurotransmitters, etc.) or intracellular (calcium, pH, etc.). Although less is known about the regulation of the paracellular pathway, alteration of TJ permeability can have dramatic effects on the ability of epithelia to generate a transepithelial solute gradient. Loss of the ability of epithelia to generate solute gradients will have both osmotic and homeostatic consequences. The purpose of this chapter is to describe the role and mechanism of action of extracellular molecules on epithelial permeability and epithelial integrity. Five groups of molecules are considered: (1) proteases, (2) xenobiotics (bacterial and nonbacterial), (3) cellular constituents, (4) leukocyte secretions, and (5) cationic proteins.

16.2 THE EPITHELIUM AS A BARRIER All substances encounter a series of barriers when moving from the lumen of an organ system to the blood compartment (Figure 16.1A and B). Starting in the lumen, the first barrier is the unstirred water layer, then the glycocalyx (fuzzy coat that is attached to the epithelial cell membrane), followed by the parallel combination of the epithelial cells and paracellular pathway and, finally, the basement membrane, interstitium, and capillary wall. Which one of these barriers offers the greatest

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FIGURE 16.1 (A) Schematic representation of the barriers to solvent and solute movement across an epithelium and associated structures. Solute and solvent must cross a number of barriers arranged in series and/or in parallel. These barriers are the unstirred water layer, the glycosaminoglycan layer (GAG), the parallel combination of the cell and TJ, followed by the basement membrane, the intersitium, and finally the capillary wall. The cell is composed of the series arrangement of the apical and basolateral membrane. The relative efficacy of these barriers in reducing solute movement can be dependent on the physical properties of the solute.

impediment to movement of a substance from one compartment to another? The answer to this question must consider the nature of the substance moving from lumen to blood as well as the type of epithelium being studied. Although it is generally accepted that the epithelial cell membranes (either or both the apical and basolateral membrane) or paracellular pathway is the major barrier to movement of small electrolytes and nonelectrolytes, the glycocalyx or basement membrane of the epithelium might limit the movement of large molecules or microorganisms. Further, the glycocalyx might play a stimulatory or inhibitory role in transepithelial movement of microorganisms. If the bacterium does not adhere to the glycocalyx, then this structure acts as a significant barrier to the movement of the bacterium across the epithelium. Conversely, bacterial adhesion to the glycocalyx is considered a required step for invasion. In many instances, the movement of a bacterium across an epithelium is an active, energy-requiring process. Such an active movement of a macromolecule can cause the glycocalyx to be the limiting barrier in transepithelial movement.

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FIGURE 16.1 (B) Electrical-equivalent circuit model of the barriers to solute movement across an epithelium and associated extracellular structures. The zigzag line represents the conductance of the individual barriers.

The unstirred layer (the adhering layer of water on both the apical and basolateral sides) can represent a significant barrier to the movement of some substances. If the transepithelial permeability of a substance is high (as an example, alcohol), then one must correct for unstirred layer effects. Thus, when interpreting results concerning the movement of a substance across a tissue, one must consider the influence of structures outside the epithelial cell membrane and the paracellular pathway. The paracellular pathway is composed of the series arrangement of the TJs and the lateral intercellular space. It is important to make a distinction between alterations in the permeability of the paracellular pathway (as defined above) and the formation of an extracellular pathway that might be caused by cell damage, necrosis, or desquamation. Alteration of the paracellular pathway might suggest regulated changes in the permeability of the TJs or the dimensions of the lateral intercellular space. Alterations in the extracellular pathway might suggest more global dysfunction of the epithelium, such as bacterial invasion or exposure to toxins. A description of the experimental methods used to measure the individual membrane conductances and extracellular conductances (see Figure 16.1B) is

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beyond the scope of this chapter. Detailed descriptions of the electrical methods used to measure the individual conductances in Figure 16.1B have been previously published (Lewis 1996a, b; Lewis et al., 1996).

16.3 ALTERATION BY PROTEASES Proteases are known to perform a number of essential physiological functions. The best known is the nutritional role that proteases play in the hydrolysis of proteins into di- and tripeptides and free amino acids for absorption by the small intestine. This digestion begins in the stomach by pepsin (secreted by chief cells) and is completed by a host of proteases released by the pancreas (trypsin, chymotrypsin, carboxypeptidases, elastase, etc.) and present on the brush border of small-intestinal cells. Proteases are also present in the bloodstream where they perform important functions such as dissolving blood clots (the fibrinolytic system) and activation or inactivation of circulating proteins, e.g., plasma kallikrein converts kininogen to bradykinin, a vasodilator, and kininases cleave bradykinin so it is no longer active. Proteases have been used in the biological sciences for a number of purposes, including protein sequencing, membrane protein topology, cell dissociation, isolation, and purification. Proteases can alter or modulate epithelial permeability either by physiological or pathophysiological methods.

16.3.1 PHYSIOLOGICAL ROLE

OF

PROTEASES

There are a number of instances where proteases have been shown to alter the sodium transport properties of tight epithelia. Sodium absorption by the rabbit urinary bladder epithelium (a tight epithelium) occurs in the following manner (Lewis and Diamond, 1976). Sodium enters the cell across the apical membrane through sodium channels (ENaC) down a net electrochemical gradient. Once in the cell, the sodium exits across the basolateral membrane via the Na,K-ATPase, in which three sodium ions are extruded from the cell for two potassium ions taken into the cell. The potassium ions that enter the cell then recycle across the basolateral membrane through potassium channels. This outward movement of potassium across the basolateral membrane makes the cell interior electrically negative, which aids in the uptake of sodium across the apical membrane. In toad urinary bladder, Garty and Edelman (1983) demonstrated that luminal trypsin (1 mg/ml) reduced the sodium transport of this epithelium by about 50%, presumably by decreasing the apical membrane permeability, i.e., hydrolyzing the apical membrane ENaC. Of interest is that trypsin did not alter the permeability properties of the TJs. In rabbit urinary bladder epithelium, Lewis and Clausen (1991) also demonstrated that luminal trypsin irreversibly reduced apical membrane conductance without any effect on TJ ion permeability. The resistance of the urinary bladder TJs to trypsin hydrolysis is in agreement with results found for tissue-cultured epithelial cells. When epithelial cells are grown on plastic supports, they form a confluent cell monolayer with welldeveloped TJs. Addition of trypsin to the surface of these confluent cells does not affect the integrity of the epithelial monolayer. In contrast, if the TJs are opened

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using low-calcium solutions, and then trypsin is added, the cells rapidly lift off the plastic supports. Thus, in the presence of normal calcium, the TJs are impermeable to trypsin, and trypsin does not increase the permeability of the TJs. However, trypsin did alter the permeability of the cell membrane by hydrolyzing the sodium channels found in the apical membrane. Do endogenous proteases alter epithelial permeability? The following observations suggested that urinary bladder ENaC was modified by some urinary constituent. First, in addition to possessing sodium channels in the apical membrane, this membrane also contains both a nonselective cation channel and a nonselective cation channel that seems to partition into and out of the apical membrane. Second, the density of channels in the apical membrane was lower than predicted based on measurements of channel density of cytoplasmic vesicles (Lewis and de Moura, 1982). Urine is known to contain three serine proteases: urokinase, plasmin, and kallikrein. The first two are part of the fibrinolytic system, are secreted by distal nephron segements, and function to maintain patency of the urinary tract. The function of kallikrein is unknown, but it is proposed to be important in sodium homeostasis. Lewis and colleagues (Lewis and Alles, 1986; Lewis and Clausen, 1991) studied the effects of kallikrein on the epithelial sodium channel. These authors found that kallikrein decreased the density of sodium channels in the apical membrane of the rabbit urinary bladder epithelium. Of interest is that not only did kallikrein decrease the density of sodium channels, but it also changed the quantity of nonselective cation channels and increased the quantity of the nonselective cation channel that partitions into and out of the apical membrane. Thus, urinary kallikrein can explain previous observations on channel density and channel properties. Since aldosterone increases kallikrein secretion by distal segments of the nephron (and increases apical membrane conductance), it is possible that kallikrein acts as a negative-feedback system for regulating apical membrane sodium channel density (Lewis and Clausen, 1991). The previous section described how urinary protease could act as a downregulator of sodium absorption. Recently, it has been proposed that proteases can act as upregulators of sodium transport. Vallet and colleagues (1997) demonstrated that the serine protease inhibitor aprotinin caused a decrease in sodium absorption by the renal A6 epithelial cells. These authors then showed that low levels of trypsin (2 µg/ml) also increased the sodium transport rate across A6 monolayers. In a functional complementation assay, these authors demonstrated the presence of a 329 amino acid protein in A6 cells with sequence homology to serine proteases that could activate Xenopus ENaC. The activation of ENaC was inhibited by aprotinin (an inhibitor of some serine proteases). The authors named this protein CAP (channel-activating protease) and speculated that it either modified ENaC directly or modified an ENaC-associated protein.

16.3.2 PATHOPHYSIOLOGICAL ROLE

OF

PROTEASES

Are there instances of pathophysiological effects of protease? This question is more difficult to demonstrate. However, there are some scenarios where endogenous proteases might exacerbate a pathophysiological condition. As an example, during gastric ulceration, there is ready access of the gastric contents (low pH and the

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endopeptidase pepsin) to the surface epithelial cells and underlying basement membrane. The presence of low pH, proteolytic enzymes, and perhaps initiating factors such as ethanol, bile salts, or aspirin could cause rapid epithelial necrosis, leading to the loss of large areas of surface epithelium. In the absence of basement membrane, there is an inhibition of migration of epithelial cells to cover the denuded area, thus extending the time for wound repair. In this example, the increase in epithelial permeability is due to an increase in the extracellular permeability as opposed to an increase in cellular or paracellular permeability. Under physiological conditions, epithelial integrity seems not to be altered by proteases. This is desirable when one considers the high levels and types of proteases in the lumen of the small intestine during a meal. This protection might be due to resistant cell membranes, resistant TJs and the buffering capacity of food in the lumen. That epithelial TJs are inert to proteases from the lumen has been confirmed in rabbit and toad urinary bladder and in tissue-cultured epithelial cells grown on plastic supports (see above). It has also been demonstrated that elastase (porcine pancreas or human leukocyte) does not alter the TJ resistance in Madin–Darby canine kidney (MDCK) cells as measured by electrical resistance, nor is it toxic to MDCK cells as assessed using 51Cr release assay (Azghani et al., 1993). One might then think that TJs are resistant to all elastases. However, these authors demonstrated that the addition of a similar activity of elastase from Pseudomonas caused a decrease in transepithelial resistance over a 4-h period without an increase in 51Cr release. These results suggest a specific increase in paracellular permeability as opposed to an increase in extracellular permeability as a result of cell necrosis and desquamation. That the TJ was the site of action was further demonstrated by measuring a decrease in staining of the TJ associated protein ZO-1 as well as a decrease in F-actin staining. Of interest is that there was no measured alteration in the morphology of the epithelium, nor was there an alteration in the staining of E-cadherin. The mechanism of action of bacterial elastase was not determined. One might speculate that since ZO-1 and F-actin are intracellular proteins that the elastase is acting from the cytoplasmic side. However, one cannot rule out the possibility that the effect of bacterial elastase is more indirect, i.e., altering a protein that results in an alteration of the distribution of ZO-1 and F-actin.

16.4 XENOBIOTICS A xenobiotic is a natural or anthropogenic chemical that is present in a living system to which it is foreign. Therapeutic drugs, non-nutritive constituents of foods, and anthropogenic chemicals, as well as natural products, solvents, pesticides, and environmental pollutants, are all examples of xenobiotics. This section is not intended to be an exhaustive list of xenobiotics that alter epithelial permeability but to list several examples and (where available) the mechanism of action.

16.4.1 NONBACTERIAL An example of a nonbacterial xenobiotic is palytoxin, which has been described as the most potent toxin of nonbacterial origin (LD50 of 10 to 450 ng/kg). The mechanism

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of toxicity of palytoxin is very interesting. It has been proposed that palytoxin binds to the Na,K-ATPase and forms nonselective cation channels; thus, cells lose potassium, gain sodium, followed by cell swelling and ultimately cell lysis (Habermann, 1989). There is one report of the use of palytoxin in epithelia. Mullin and colleagues (1991) demonstrated that the addition of 1 × 10–9 M palytoxin to mucosal and serosal solution led to an irreversible increase in transepithelial conductance of the renal tissue culture cell line LLC-PK1. The increase in conductance was biphasic with an initial rapid increase which plateaued after 2 to 3 h, followed by a slow increase in conductance that started 5 h after the initial addition of palytoxin. The initial increase in conductance (at both apical and basolateral membrane) might be due to the change in membrane conductance as a result of interaction with the Na,K-ATPase, and the secondary change might be due to cell swelling and lysis. Thus, palytoxin over the long term increases the extracellular conductance of the epithelium. An interesting observation is that addition of palytoxin to only the mucosal solution increased only the apical membrane conductance. The magnitude of the increase in apical conductance decreased as a function of the time postseeding the LLC-PK1 cells on permeable supports. This suggests that there is either a decrease in the targeting of the Na,K-ATPase to the apical membrane or a decrease in the residency time of the pump in the apical membrane. Asbestos is an example of a natural product and environmental pollutant. After long-term exposure, asbestos can result in asbestosis, i.e., fibrosis of the lung. In animal models it has been demonstrated that an early effect of asbestos is an increase in epithelial permeability. To determine whether this increase was a direct effect of asbestos on the epithelium or due to a secondary inflammatory response, Peterson and colleagues (1993) studied the effect of asbestos on the permeability of HBE (human bronchial epithelia cell line) grown on permeable supports. In brief, these authors demonstrated that asbestos at a concentration as low as 10 µg/ml caused an increase in mannitol permeability after an 8-h exposure. The permeability of the monolayer to mannitol continued to increase with prolonged asbestos exposure. The asbestos-induced increase was not reversible if the monolayers were pretreated with asbestos for 24 h. Although it is not clear what caused the increase in permeability, a number of possible mechanisms were excluded, including cytotoxicity (as determined by LDH release), alteration in the actin filament of the cell, mechanical alteration due to particle size or surface-bound iron.

16.4.2 BACTERIAL There are a number of excellent examples of the effects of bacterial xenobiotics on epithelial permeability. An example of a bacterial xenobiotic already discussed is bacterial elastase (see section on proteases). Another example is the antibiotic polymyxin B, which is secreted by Bacillus polymyxa. This antibiotic is used clinically in antibacterial ointments, as a bactericidal agent during long-term bladder irrigation, in ophthalmic solutions, and on a limited basis as an inhalant in patients with cystic fibrosis with Pseudomonas aureginous airway infections. If administered systemically, this antibiotic is nephrotoxic and neurotoxic. Recent work (Berg et al., 1996;

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FIGURE 16.2 The time course of the effect of polymyxin B on the transepithelial conductance (Gt) and the current flow across the epithelium (I). Polymyxin B is added to the luminal solution (40 µM) of the bladder while the transepithelial voltage is held at –70 mV (serosal solution ground). At this holding voltage, the apical membrane voltage is 15 mV (cell positive) and there is no effect of the antibiotic on the transepithelial conductance or current. At the arrow, the transepithelial holding voltage is then changed to 0 mV (the apical membrane voltage is then –55 mV, cell negative). Note that there is an increase in both the transepithelial conductance and current. This increase in current suggests that polymyxin B is increasing the conductance of the cell membrane.

1998) has been directed to determining the mechanism of nephrotoxicity of polymyxin B. Polymyxin is a cationic cyclic decapeptide with a fatty acyl tail, and a molecular weight of 1400. Using the mammalian urinary bladder epithelium, these authors demonstrated that polymyxin B increased the transepithelial conductance of the urinary bladder epithelium in a voltage-dependent manner (Figure 16.2). The site of the short-term conductance increase was at the apical membrane. Although short exposure times were readily reversible, prolonged exposure resulted in an irreversible increase in transepithelial conductance. This loss of barrier function was due to an increase in the extracellular conductance caused by cell swelling and lysis.

16.5 MODULATION BY LEUKOCYTES Leukocytes release a number of factors, including cytokines and cytolytic proteins, that alter epithelial permeability. This section focuses on the effects of cytolytic proteins (released by leukocytes) on epithelial permeabiltiy. The mechanism of action of cytokines and the transmigration of leukocytes are discussed in separate chapters of this book.

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16.5.1 EOSINOPHIL PROTEINS Eosinophils are leukocytes involved in allergies and infection, and use proteins that alter membrane permeability to kill parasites and other pathogens. The following sequences of events are involved in eosinophil toxicity. The eosinophil adheres to an immunoglobulin G-coated pathogen, forming a small pocket composed of the membrane of the pathogen and the membrane of the eosinophil. The contents of the granules of the eosinophil are emptied into this pocket allowing an interaction between the proteins and membrane of the pathogen. The proteins are eosinophil cationic protein (ECP), eosinophil peroxidase (EPO), major basic protein (MBP), and eosinophil-derived neurotoxin (EDN). These proteins are cationic (contain a net positive charge) and cytotoxic to a wide range of cells including mammalian cells. It is generally accepted that eosinophil proteins play a major role in the pathogenesis of asthma. Three of the proteins (ECP, MBP, and EPO) have been shown to cause the rupture of tracheal epithelium (Motojima et al., 1989). In addition, lung tissue from people with asthma has been demonstrated to have high concentrations of these three proteins, as well as eosinophils. These results suggest that a component of the pathology of asthma is due to the effect of these proteins on the epithelial cell membrane. One proposed mechanism for the effect of eosinophil proteins is by directly increasing the membrane permeability of the epithelial cell membrane by ion channel formation. Of interest is that one of the proteins (ECP) has been shown to form ion channels in lipid bilayer membranes (Young et al., 1986). The channels were nonselective and not voltage dependent. Although similar studies have not been performed on MBP, this protein has been shown to interact with negatively charged lipid vesicles (Abu-Ghazaleh et al., 1992). This suggests that a protein–lipid interaction is an integral step in the MBP toxic effect on tracheal epithelium. The effect of MBP and EPO has been tested on the permeability properties of the rabbit urinary bladder epithelium (Kleine et al., 1998; 1999). The addition of micromolar quantities of either protein to the luminal solution resulted in a voltagedependent increase in apical membrane conductance. If the apical membrane voltage was held such that the cell interior was electrically positive compared with the luminal solution, the subsequent addition of either protein to the luminal solution did not result in a change in transepithelial conductance. If the apical membrane voltage was then changed to cell interior negative, there was a time-dependent increase in the transepithelial conductance (Kleine et al., 1998; 1999). The site of the increase in conductance was at the apical membrane and not the TJ. The induced conductance was nonselective; i.e., sodium potassium and chloride were all equally permeable through the induced conductance. This increase in apical membrane conductance could be reversed (at short exposure times) by changing the apical membrane voltage back to cell interior positive, by removing the protein from the bath, or by adding calcium to the luminal solution. The mechanism of the calcium effect is proposed to be threefold: (1) it competes with MBP or EPO for a negatively charged membrane-binding site (probably a phospholipid); (2) it acts as an open channel blocker; and (3) it increases the rate of loss of the conductance from the membrane when the protein is removed from the bath or the membrane voltage is changed to cell interior positive. After long exposure, removing the protein from the

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bath results in an incomplete return of the transepithelial conductance to control values. This long-term increase in conductance was not due to an increase in the apical membrane conductance, but rather to an increase in the paracellular or extracellular conductance. It is speculated that the conductance increase is not a change in the TJ structure, but rather is due to the loss of some of the epithelial cells, i.e., nonspecific extracellular pathway. The loss of these cells was probably due to the influx of sodium and chloride, and a concomitant increase in cell osmotic pressure leading to cell swelling and lysis. In addition to an increase in membrane permeability, MBP has been reported to increase prostaglandin synthesis and release in tracheal epithelium (White et al., 1993). Whether prostaglandin synthesis is a byproduct of an increase in cell calcium (the calcium ionophore A23187 also stimulates prostaglandin synthesis) or a more specific interaction is not known; neither is the effect of prostaglandin on epithelial integrity known.

16.5.2 NEUTROPHIL PROTEINS Neutrophils are felt to represent the body’s first line of defense during tissue injury or infection. After phagocytosing the target, the phagocytic vesicle fuses with azurophilic granules, which contain toxic proteins including cationic proteins — defensins, cathepsin G, azurocidin, or cationic antimicrobial protein (CAP-37) — and bacterial/permeability-increasing protein (BPI or CAP-57), elastase, lysozyme, and myeloperoxidase. Neutrophils use two general mechanisms for killing target organisms; one is oxygen dependent and the other is oxygen independent. The cationic proteins that are cytotoxic via an oxygen-independent mechanism are discussed here. There are only a few studies of the effects of defensins on epithelial integrity. Defensins are toxic to a lung adenocarcinoma cell line (A549, a cell line that has characteristics similar to alveolar epithelial cells). The mechanism of toxicity is not known, but it is speculated that it might involve endocytosis and intracellular modification (Okrent et al., 1990). In another study, defensins were shown to increase the mannitol permeability of MDCK monolayers as well as to increase the transepithelial conductance of the monolayers (Nygaard et al., 1993). Lencer and colleagues (1997) demonstrated that cryptdins 2 and 3 (defensins isolated from intestinal paneth cells) stimulated chloride secretion in an intestinal epithelial cell line T84. Of interest is that the chloride current was not due to cAMP- or cGMP-stimulated chloride channels and that the induced conductance was large enough to allow the movement of carboxyfluorescein into the cells. As shown in lipid bilayer studies (Kagan et al., 1990), this increase in chloride current is most likely due to the ability of defensins to form channels directly in the apical membrane. These bilayer studies demonstrated that defensins form a voltage-dependent conductance, which was weakly anion selective. Single-channel conductances range from 10 to 1000 pS, suggesting a barrel stave mechanism for pore formation, where the large conductance channel is composed of more barrel staves. Although defensins were originally discovered in neutrophils, there are a number of epithelial tissues that secrete defensins (or defensin-like molecules) into their lumen (see Schröder, 1999, for a review). The effect of cathepsin-G has been studied on cultured type II pneumocytes (Rochat et al., 1988). These authors demonstrated that cathepsin-G increased transepithelial

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mannitol permeability and transepithelial conductance. The increase in mannitol permeability suggests an increase in either the paracellular or extracellular pathway, while the increase in conductance might be due to an increase in transcellular and paracellular (extracellular) conductance. The effect of azurocidin, CAP-37, and CAP-57 has not been studied on epithelial permeability or integrity.

16.6 MODULATION BY CELLULAR CONSTITUENTS Although the above substances are released into the extracellular milieu, proteins that play a predominantly intracellular function can also alter transepithelial permeability. Protamine and histones are cationic proteins localized to sperm head (protamine and histones) or the nucleus (histones). When found in the extracellular environment as a result of pharmacological use or through the breakdown of cells, protamine and histones can have pathophysiological effects.

16.6.1 PROTAMINE Protamine is a DNA-binding protein found in sperm head. This protein is used clinically as a sequestering agent during surgical procedures that require heparin as an anticoagulant. There are a number of side effects of intravenous injection of protamine, which include neutropenia, thrombocytopenia, pulmonary hyper- or hypotension, and bradycardia. Protamine is an unusual molecule in that approximately 67% of the protein is arginine; thus, it is a highly positively charged protein. It is the high charge density that makes it ideal for compacting DNA in sperm head and binding heparin; however, it is also the high charge density that might result in the side effects. The effects of protamine on epithelial permeability have been extensively studied. Table 16.1 lists the effects of protamine on a number of different epithelia. The effect of luminal protamine on epithelial permeability fell into three categories. First, protamine decreased transepithelial conductance of Necturus gallbladder (Bentzel et al., 1987), ciliary epithelium (Straub and Wiederholt, 1991), and Henle’s loop (Koyama et al., 1991a); the decrease in conductance was due to a decrease in paracellular conductance. Second, protamine increased transepithelial conductance in Necturus gallbladder (Poler and Reuss, 1987), proximal tubule (Okada et al., 1975), and urinary bladder (Tzan et al., 1993); the increase was due to an increase in apical membrane conductance. Last, protamine increased transepithelial conductance in MDCK cells, and the increase was due to an increase in the paracellular conductance (Peterson and Gruenhaupt, 1990). That there might be a cellular site of action is discussed below. Fromm and colleagues (1990) proposed that the different response and site of protamine action on gallbladder was due in part to the pH of the luminal solution. A comparison of protamine action on MDCK cells to gallbladder, proximal tubule, and urinary bladder suggests that in MDCK cells protamine increases TJ conductance whereas in the other epithelia it increases cell conductance. A possible explanation is that the effects of protamine in gallbladder, proximal tubule, and urinary bladder were measured soon after adding protein to the bath (minutes), whereas for MDCK cells the increase in TJ conductance was measured 1 to 4 h after adding protein.

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TABLE 16.1 Site of Action of Protamine Is Dependent on the Type of Epithelium Tissue NGB Ciliary epithelium Descending limb, Henle’s loop Ascending thin limb, Henle’s loop NGB Proximal tubule Urinary bladder MDCK cells

Gt ↓ ↓ ↓ ↓ ↑ ↑ ↑ ↑ ↑ ↑

Gj ↓ ↓ ↓ ↓ ? ↑(CL) ↔ ↑(CL) ? ↑

Gc ↔ ↔ ↔ ↔ ↑ ↑ ↑ ↑ ↑ ?

Comment

L.T. S.T. L.T. S.T. L.T.

Ref. Bentzel et al., 1987 Straub and Wiederholt, 1991 Koyama et al., 1991b Koyama et al., 1991a Poler and Reuss, 1987 Okada et al., 1975 Tzan et al., 1993 Peterson and Gruenhaupt, 1990

Gt = transepithelial conductance; Gj = tight junction conductance; Gc = cellular conductance; ↑ = increase; ↓ = decrease; ↔ = no change; ? = unknown; CL = increase in junctional conductance due to cell lysis not TJ structure; L.T. = long term (hours); S.T. = short term (minutes); NGB = Necturus gallbladder; MDCK = Madin–Darby canine kidney.

Indeed, in urinary bladder, prolonged exposure to protamine results in an irreversible increase in extracellular conductance (Tzan et al., 1993). Interestingly, in MDCK cells there is evidence for an increase in cell membrane conductance over the first 5 to 10 min of exposure to protamine (Peterson and Gruenhaupt, 1990). The effect of protamine on epithelial structure has been reported for gallbladder and MDCK cells. In gallbladder, Quinton and Philpott (1973) reported that adjacent membranes of the gallbladder epithelium fused, microvilli lost rigidity, and there was an apparent increase in membrane permeability. Bentzel and colleagues (1987) reported variable changes in microvilli structure but an increase in the number of strands that comprise the TJ. Confocal microscopy of MDCK monolayers treated with polylysine (a molecule similar to protamine) demonstrated a loss of the TJ associated protein ZO-1 and inspection of F-actin staining showed breaks in the TJ structure (i.e., there was a patent paracellular pathway). These confocal images were acquired after a 10-min exposure to a high concentration of polylysine (5 µM of 10,000 mw polylysine) (McEwan et al., 1993).

16.6.2 HISTONES Histones are cationic DNA-binding proteins found in the nucleus of somatic cells of most organisms and in sperm head of humans. Histones with a short length of DNA wrapped around it form the nucleosome. The molecular weight of histones ranges from 11,000 Da for H4 to 24,000 Da for H1. The fraction of positively charged amino acids ranges from 22.5% for H4 to 34.7% for H5. The tertiary structure of histone is also variable; H1 and H5 have a central globular domain between two random coil tails. In contrast, H4 has an α helical component between two random coil tails. Histones have been shown to increase the permeability of mitochondria (Johnson et al., 1967) and to alter the architecture of gallbladder

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epithelium (Quinton and Philpott, 1973). Addition of a histone (H1, H4, or H5) to the luminal solution of the mammalian urinary bladder epithelium caused an increase in the transepithelial conductance. The site of action of histone (short time) was at the apical membrane (Kleine et al., 1995). The increase in the apical membrane conductance by histone required that the apical membrane voltage be cell interior negative. The induced conductance was nonselective, i.e., the conductance was permeable to sodium, potassium, and chloride. For short time periods, the conductance could be reversed at cell-negative voltages by removing the protein from the bath by washing, complexing the protein with DNA, or hydrolyzing the protein with trypsin. In the presence of bath histone, changing the apical membrane voltage to cell interior positive can reverse the induced conductance. Over long-term exposure to histone, there was an irreversible increase in the transepithelial conductance. This toxic effect of histone might be due to a mechanism similar to the eosinophil proteins, i.e., cell lysis due to swelling of the epithelial cells. The effect of protamine and histones on epithelial integrity is of particular interest when considering the results of Mendizabal and Naftalin (1992). These authors demonstrated that human semen is cytotoxic to the in vivo colon of the rat. Sperm breaks down in the colon in about 1 h and the toxicity of the semen to the colon occurred after about 3 to 4 h. Histology of the colon demonstrated that after 3 to 4 h there were no epithelial cells present where the semen had been placed. These authors (Mendizabal and Naftalin, 1992) proposed that cationic amines (particularly spermidine, spermine, and putricine) and collagenase were responsible for the loss (by lysis or desquamation) of the colonic epithelial cells. Histones and protamine, which are cationic and found in micromolar concentrations in sperm, might also contribute to the toxicity of semen. These proteins might then provide access for pathogenic viruses and other organisms across the colonic mucosal barrier.

16.7 CATIONIC PROTEINS With a few exceptions (Table 16.2), the short-term effect of cationic proteins is to increase membrane permeability, whereas the long-term effect of cationic proteins (independent of their origin) is to increase transepithelial conductance by increasing the permeability of an extracellular pathway. The cationic proteins discussed in this chapter seem to act in a similar manner. Table 16.2 summarizes some of the features of cationic protein-induced conductance. In all cases, the proteins increase the apical membrane conductance in a voltagedependent manner. The apical membrane potential must be cell interior negative before there is a measurable increase in membrane conductance. The relationship between the magnitude of the cell membrane voltage and the induced conductance is exponential. Thus, increasing the membrane voltage from –50 to –200 mV will increase the membrane conductance more than 200-fold. In all instances the induced conductance could be reversed if the exposure time were short. This reversal could be achieved by either removing the protein from the bath or changing the apical membrane voltage to a cell-positive value. The ion selectivity of the induced conductance did not discriminate between cations and anions. In the case of protamine,

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TABLE 16.2 Properties of Cationic Protein-Induced Conductance Protein Protamine Histone H4 MBP EPO Polymyxin B

Gt ↑ ↑ ↑ ↑ ↑

Site Apical Apical Apical Apical Apical

Voltage Yes, exp Yes, exp Yes, exp Yes, ? Yes, exp

Reversible (short term) Yes Yes Yes Yes Yes

Selectivity Nonselective Nonselective Nonselective Nonselective Nonselective

Dose Response Saturating Saturating Saturating ? Saturating

Gt = transepithelial conductance; ↑ = increase; site = the site of action of the protein on the epithelium; apical = apical membrane; voltage = protein requires a negative cell interior voltage to increase membrane conductance; exp = the relationship between magnitude of the voltage and magnitude of conductance increase is an exponential; reversible = the induced conductance can be reversed by voltage or removing the protein from the bath; selectivity = nonselective = the induced conductance does not discriminate between cations and anions; dose response = the relationship between the conductance change as a function of the concentration of protein added to bath. Saturating suggests a Michaelis–Menten curve.

the induced conductance was large enough to allow the movement of 12,000-mw dextran. This suggests a large aqueous pore. Last (not shown in Table 16.2), calcium reduced the ability of the cationic protein to induce a conductance. For the specific case of protamine, calcium was shown to act by three mechanisms. First, calcium and protamine competed for the same binding site; this binding site might be negatively charged phospholipid. Second, calcium blocks the protamine-induced conductance in a rapid and reversible manner; this was called “conductive block” (Tzan et al., 1994). Last, in the absence of bath calcium, the induced conductance could not be completely reversed by removing protamine from the bath. Thus, calcium is important for loss of conductance from the membrane. Possible mechanisms by which these proteins increase the extracellular/paracellular permeability are illustrated in Figure 16.3. These include osmotic disequilibrium caused by influx of sodium and chloride, alteration in the intracellular milieu, disruption of metabolism, altered enzyme activity, or interaction with the cell cytoskeleton.

16.8 SUMMARY This chapter has attempted to demonstrate that there are a number of mechanisms by which the barrier function of an epithelium can be altered. The compounds vary from proteases to leukocyte secretion, cell constituents, and xenobiotics of both nonbacterial and bacterial origin. Although the final result of a loss of barrier function is similar for these compounds, the path to the final result might be quite different. Elucidating and understanding these pathways will lead to therapeutic approaches to circumvent the loss of epithelial barrier function.

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FIGURE 16.3 This figure shows possible initial sites of cationic protein action, which ultimately results in an increase in extracellular permeability. (i) The cationic proteins increase the cell membrane conductance to small monovalent cations, resulting in an osmotic disequilibrium, cell swelling, and subsequent cell lysis. (ii) The increased membrane permeability results in the loss of essential cell constituents (e.g., ATP or proteins) that are important for the maintenance of the TJs. (iii) The increase in membrane permeability by the cationic proteins increases or decreases divalent cation concentration, which can increase TJ permeability. (iv) The cationic proteins enter the epithelial cells and disrupt mitochondrial metabolism. This results in a decrease in cell ATP levels. (v) Internalization of the protein might alter cell enzyme activity such as kinases or phosphatases, which regulate TJ assembly and disassembly. (vi) The increase in extracellular conductance might not depend on an alteration in membrane conductance; rather, it might be due to an association of the protein with the cell membrane, which alters the cytoskeleton leading to a conductance increase.

ACKNOWLEDGMENTS This work was supported in part by National Institutes of Health Grant DK51382. The author thanks Jamie R. Lewis for helpful comments on this manuscript.

REFERENCES Abu-Ghazaleh, R. I., Gleich, G. J., and Prendergast, F. G. 1992. Interaction of eosinophil granule major basic protein with synthetic lipid bilayers: a mechanism for toxicity. J. Membr. Biol., 128:153–164. Azghani, A. O., Gray, L. D., and Johnson, A. R. 1993. A bacterial protease perturbs the paracellular barrier function of transporting epithelial monolayers in culture. Infect. Immunol., 61:2681–2686. Bentzel, C. J., Fromm, M., Palant, C. E., and Hegel, U. 1987. Protamine alters structure and conductance of Necturus gallbladder tight junctions without major electrical effects on the apical cell membrane. J. Membr. Biol., 95:9–20. Berg, J. R., Spilker, C. M., and Lewis, S. A. 1996. Effects of polymyxin B on mammalian urinary bladder. J. Membr. Biol., 154:119–130.

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Berg, J. R., Spilker, C. M., and Lewis, S. A. 1998. Modulation of polymyxin B effects on mammalian urinary bladder. Am. J. Physiol., 275:F204–F215. Fromm, M., Tykocinski, M., Schulzke, J.-D., Hegel, U., and Bentzel, C. J. 1990. pH dependence of protamine action on apical membrane permeability in Necturus gallbladder epithelium. Biochim. Biophys. Acta, 1027:179–184. Garty, H. and Edelman, I. S. 1983. Amiloride-sensitive trypsination of apical sodium channels. J. Gen. Physiol., 81:785–803. Habermann, E. 1989. Palytoxin acts through Na+,K+-ATPase. Toxicon, 11:1171–1187. Johnson, C. L., Mauritzen, C. M., Starbuck, W. C., and Schwartz, A. 1967. Histones and mitochondrial ion transport. Biochemistry, 6:1121–1127. Kagan, B. L., Selsted, M. E., Ganz, T., and Lehrer, R. I. 1990. Antimicrobial defensin peptides form voltage-dependent ion-permeable channels in planar lipid bilayer membranes. Proc. Natl. Acad. Sci. U.S.A., 87:210–214. Kleine, T. J., Gladfelter, A., Lewis, P. N., and Lewis, S. A. 1995. Histone-induced damage of a mammalian epithelium: the conductive effect. Am. J. Physiol., 37:C1114–C1125. Kleine, T. J., Gleich, G. J., and Lewis, S. A. 1998. Eosinophil major basic protein increases membrane permeability in mammalian urinary bladder epithelium. Am. J. Physiol. Cell Physiol., 44:C93–C103. Kleine, T. J., Gleich, G. J., and Lewis, S. A. 1999. Eosinophil peroxidase (EPO) increases membrane permeability in mamalian urinary bladder epithelium. Am. J. Physiol., 276:C638–C647. Koyama, S., Yoshitomi, K., and Imai, M. 1991a. Effect of protamine on ion conductance of ascending thin limb of Henle’s loop from hamsters. Am. J. Physiol. Renal Fluid Electrolyte Physiol., 261:F593–F599. Koyama, S., Yoshitomi, K., and Imai, M. 1991b. Effect of protamine on ion conductance of upper portion of descending limb of long-looped nephron from hamsters. Am. J. Physiol. Renal Fluid Electrolyte Physiol., 260:F839–F847 Lencer, W. I., Cheung, G., Strohmeier, G. R., Currie, M. G., Ouellette, A. J., Selsted, M. E., and Madara, J. L. 1997. Induction of epithelial chloride secretion by channel-forming cryptdins 2 and 3. Proc. Natl. Acad. Sci. U.S.A., 94:8585–8589. Lewis, S. A. 1996a. Epithelial electrophysiology, in Epithelial Transport: A Guide to Methods and Experimental Analysis. N. K. Wills, L. Reuss, and S. A. Lewis, Eds., Chapman & Hall, London, 93–117. Lewis, S. A. 1996b. Epithelial structure and function, in Epithelial Transport: A Guide to Methods and Experimental Analysis. Vol. 1, N. K. Wills, L. Reuss, and S. A. Lewis, Eds., Chapman & Hall, London, 1–20. Lewis, S. A. and Alles, W. P. 1986. Urinary kallikrein: a physiological regulator of epithelial Na absorption. Proc. Natl. Acad. Sci. U.S.A., 83:5345–5348. Lewis, S. A. and Clausen, C. 1991. Urinary proteases degrade epithelial sodium channels. J. Membr. Biol., 122:77–88. Lewis, S. A. and de Moura, J. L. C. 1982. Incorporation of cytoplasmic vesicles into apical membrane of mammalian urinary bladder epithelium. Nature, 297:685–688. Lewis, S. A. and Diamond, J. M. 1976. Na+ transport by rabbit urinary bladder, a tight epithelium. J. Membr. Biol., 28:1–40. Lewis, S. A., Clausen, C., and Wills, N. K. 1996. Impedance analysis of epithelia, in Epithelial Transport: A Guide to Methods and Experimental Analysis. Vol. 1, N. K. Wills, L. Reuss, and S. A. Lewis, Eds., Chapman & Hall, London, 118–145. McEwan, G. T. A., Jepson, M. A., Hirst, B. H., and Simmons, N. L. 1993. Polycation-induced enhancement of epithelial paracellular permeability is independent of tight junctional characteristics. Biochim. Biophys. Acta Biol. Membr., 1148:51–60.

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Mendizabal, M. V. and Naftalin, R. J. 1992. In vivo exposure of rat colonic mucosa to human semen induces mucosal cytolysis, abolishes fluid absorption and raises paracellular permeability. J. Physiol., 446:411P Motojima, S., Frigas, E., Loegering, D. A., and Gleich, G. J. 1989. Toxicity of eosinophil cationic proteins for guinea pig tracheal epithelium in vitro. Am. Rev. Respir. Dis., 139:801–805. Mullin, J. M., Snock, K. V., and McGinn, M. T. 1991. Effects of apical vs. basolateral palytoxin on LLC-PK1 renal epithelia. Am. J. Physiol. Cell Physiol., 260:C1201–C1211. Nygaard, S. D., Ganz, T., and Peterson, M. W. 1993. Defensins reduce the barrier integrity of a cultured epithelial monolayer without cytotoxicity. Am. J. Respir. Crit. Care Med., 8:193–200. Okada, Y., Sato, T., and Inouye, A. 1975. Effects of potassium ions and sodium ions on membrane potential of epithelial cells in rat duodenum. Biochim. Biophys. Acta, 413:104–115. Okrent, D. G., Lichtenstein, A. K., and Ganz, T. 1990. Direct cytotoxicity of polymorphonuclear leukocyte granule proteins to human lung-derived cells and endothelial cells. Am. Rev. Respir. Dis., 141:179–185. Peterson, M. W. and Gruenhaupt, D. 1990. Protamine increases the permeability of cultured epithelial monolayers. J. Appl. Physiol., 68:220–227. Peterson, M. W., Walter, M. E., and Gross, T. J. 1993. Asbestos directly increases lung epithelial permeablity. Am. J. Physiol., 265:L308–L317. Poler, S. M. and Reuss, L. 1987. Protamine alters apical membrane K+ and Cl– permeability in gallbladder epithelium. Am. J. Physiol., 253:C662–C671. Quinton, P. M. and Philpott, C. W. 1973. A role for anionic sites in epithelial architecture. J. Cell Biol., 56:787–796. Rochat, T., Casale, J., Hunninghake, G. W., and Peterson, M. W. 1988. Neutrophil cathepsin G increases permeability of cultured type II pneumocytes. Am. J. Physiol., 255:C603–C611 Schröder, J. M. 1999. Epithelial peptide antibiotics. Biochem. Pharmacol., 57:121–134. Straub, O. and Wiederholt, M. 1991. Transepithelial resistance of ciliary epithelial cells in culture: functional modification by protamine and extracellular calcium. Comp. Biochem. Physiol., 100A:987–993. Tzan, C. J., Berg, J. R., and Lewis, S. A. 1993. Effect of protamine sulfate on the permeability properties of the mammalian urinary bladder. J. Membr. Biol., 133:227–242. Tzan, C. J., Berg, J. R., and Lewis, S. A. 1994. Mammalian urinary bladder permeability is altered by cationic proteins: modulation by divalent cations. Am. J. Physiol. Cell Physiol., 267:C1013–C1026 Vallet, V., Chraïbi, A., Gaeggeler, H. P., Horisberger, J.-D., and Rossier, B. C. 1997. An epithelial serine protease activates the amiloride-sensitive sodium channel. Nature, 389:607–610. White, S. R., Sigrist, K. S., and Spaethe, S. M. 1993. Prostaglandin secretion by guinea pig tracheal epithelial cells caused by eosinophil major basic protein. Am. J. Physiol. Lung Cell. Mol. Physiol., 265:L234–L242. Young, J. D. E., Peterson, C. G. B., Venge, P., and Cohn, Z. A. 1986. Mechanism of membrane damage mediated by human eosinophil cationic protein. Nature, 321:613–616.

17

Intracellular Signaling in Classical and New Tight Junction Functions Gaëlle Benais-Pont, Karl Matter, and Maria S. Balda

CONTENTS 17.1 Tight Junction Functions ............................................................................367 17.2 Experimental Systems.................................................................................368 17.3 The GTP-Binding Protein Pathways ..........................................................369 17.3.1 Heterotrimeric G Proteins .............................................................369 17.3.2 Small GTPases...............................................................................372 17.4 The Protein Kinases C Pathway .................................................................373 17.5 The Protein Kinase A Pathway ..................................................................374 17.6 Phosphorylation of Tight Junction Proteins ...............................................375 17.6.1 Serine/Threonine Phosphorylation ................................................375 17.6.2 Tyrosine Phosphorylation ..............................................................377 17.7 Regulation of Paracellular Permeability by Multiple Signaling Pathways .....................................................................................................379 17.8 Cell–Cell Junctions in Gene Expression and Differentiation ....................380 17.8.1 Ras-Mediated Transformation and the Junctional Complex ........380 17.8.2 Cell–Cell Junctions and Transcription Factors .............................383 Note Added in Proof..............................................................................................385 Acknowledgments..................................................................................................385 References..............................................................................................................385

17.1 TIGHT JUNCTION FUNCTIONS Multicellular organisms are formed by compartments of different composition. To maintain this compartmentalization, they possess specialized cells, epithelia and endothelia, that form selective barriers. These cells are polarized in an apical and a basolateral cell surface domain and establish an intercellular junctional complex. In epithelial cells, the most apical structure of the junctional complex is the zonula occludens or tight junction (TJ) (see Chapter 1 by Anderson and Cereijido and Chapter 2 by Schneeberger and Lyunch; Farquhar and Palade, 1963). TJs act as a 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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fence between the apical and basolateral cell surface domains and regulate the diffusion of molecules through the paracellular space. Different types of extracellular stimuli were found to modify transepithelial electrical resistance (TER) and paracellular permeability (see Chapter 15 by Turner and Madara, Chapter 16 by Lewis, and Chapter 18 by Nguyen et al.), the two experimental parameters that are commonly used to characterize the junctional paracellular barrier (see Chapter 4 by Reuss). Moreover, recent evidence shows that these two TJ functions can be dissociated (i.e., modified independently) from each other (Balda et al., 1996a; McCarthy et al., 1996; Calderon et al., 1998; Hasegawa et al., 1999), and that TJ proteins also participate in signaling processes that govern gene expression and differentiation (Balda and Matter, 2000a; Li and Mrsny, 2000; Nakamura et al., 2000; Reichert et al., 2000; Ryeom et al., 2000). TJs are not simple barriers but are semipermeable since they permit selective diffusion of tracers with particular characteristics (see Chapter 4 by Reuss; Lindemann and Solomon, 1962; van Os et al., 1974). The molecular mechanism, however, that permits selective paracellular diffusion has not yet been identified (Balda and Matter, 2000b). Because of the existence of such a selective diffusion pathway, paracellular permeability can increase for two reasons: increased selective permeability of TJs or disruption of intercellular junctions. The latter possibility results in a nonselective increase in paracellular permeability due to, for example, activation of a transforming signaling pathway. Although this is not always possible, this chapter will try to differentiate between these two types of regulation. Multiple questions must be answered to understand the signaling processes that occur at TJs. Are there general or TJ-specific receptors? How are signals transmitted to TJs and how are TJ proteins modified in response to such signals? How do TJ proteins transduce signals in response to stimulation? What are the transducing signaling molecules at the TJ? Is there a signaling complex at the TJ that receives and transmits information from junctional proteins within the cell and between neighboring cells? Although the knowledge on TJs has significantly increased since the last edition of this book, only partial answers can be given to these fascinating questions. In this chapter, the observations on TJ signaling are summarized and an attempt is made to integrate them in a working model. The chapter begins with observations focusing on the two most-studied types of signal transduction molecules, GTPbinding proteins and protein kinases, continues with observations on the phosphorylation of TJ proteins, and ends with a discussion of the involvement of TJs in the regulation of gene expression and differentiation.

17.2 EXPERIMENTAL SYSTEMS Different experimental model systems and strategies have been developed to try to simplify the analysis of different signaling pathways involved in TJ regulation. These experimental protocols include the Ca-switch model (see Chapter 8 by Shoshani and Contreras; González-Mariscal et al., 1985), ATP depletion (see Chapter 25 by Marrs and Molitoris), induction of TJs in brain endothelial cells (Rubin and Staddon, 1999; Bauer and Bauer, 2000; Kniesel and Wolburg, 2000), detachment from the substrate

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or transformation (Contreras et al., 1999; Chen et al., 2000; Li and Mrsny, 2000), and leukocyte migration through brain endothelial or epithelial cells (see Chapter 29 by Burns et al.; Parkos, 1997; Hofman et al., 1998; Huber et al., 1998; Adamson et al., 1999).

17.3 THE GTP-BINDING PROTEIN PATHWAYS GTP-binding proteins are important switches that cycle between an active (GTPbound) and an inactive (GDP-bound) state. There are two types of GTP-binding proteins, and several subfamilies are known in both cases. The heterotrimeric G proteins, or simply G proteins, are membrane-signaling systems that contribute to transduce a receptor-mediated signal from one side of a membrane to the other. G proteins are formed by three subunits, α, β, and γ. Gα subunits consist of domains for binding and hydrolysis of GTP, and for association with activated receptors, effectors, and dimers composed of β and γ subunits. Over 20 different Gα subunits are expressed in mammals; some of them are additionally expressed as alternatively spliced isoforms. They are grouped into four classes: Gα-s, Gα-i, Gα-q, and Gα-12 (Morris and Malbon, 1999). There is evidence that suggests an involvement of Gα-s, Gα-i, and Gα-q in TJ functions (Balda et al., 1991; Denker et al., 1996; Dodane and Kachar, 1996; see Table 17.1). Members of the Gα-s (for stimulatory) class activate adenyl cyclase and are substrates for cholera toxin. Members of the Gα-i (for inhibitory) class are involved in the regulation of ion channel activity, adenyl cyclase, and phosphodiesterase. G proteins of this class are substrates for pertussis toxin. Gα-q class members are substrates for neither cholera nor pertussis toxin and are regulators of specific phospholipase C isoforms, which are enzymes that are involved in the regulation of PKC (protein kinase C). The other type of GTP-binding proteins are monomeric G proteins, or small G proteins. They have an intrinsic GTPase activity that is regulated by other proteins. They associate with a GEF (guanine nucleotide exchange factor) that increases their affinity for GTP to become activated or with a GAP (GTPase-activating protein) to hydrolyze the bound GTP to become inactivated. Based on the yeast genome, there are at least five subfamilies of small GTPases: Ras, Rho, Ran, Rab, and ARF GTPases (Garcia-Ranea and Valencia, 1998). Ras proteins contribute to the regulation of cell growth–stimulating signaling pathways (Campbell et al., 1998). Rho-family GTPases (e.g., RhoA, Rac1, and Cdc42) regulate several cellular processes, such as actin dynamics, cell cycle progression, cell adhesion, and gene expression (Bishop and Hall, 2000). Ran GTPases regulate nuclear import and export (Mattaj and Englmeier, 1998). Rabs and ARFs are involved in vesicular traffic (Moss and Vaughan, 1998; Schimmoller et al., 1998).

17.3.1 HETEROTRIMERIC G PROTEINS Early pharmacological studies suggested the participation of G proteins in the regulation of TJ functions. The use of a nonhydrolizable GTP analogue and AlF4– that activates GTP-binding proteins was shown to inhibit assembly of ZO-1 (Zonula occluden 1) and TER development induced by calcium, respectively (Balda et al.,

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TABLE 17.1 Signaling Pathways Regulating Tight Junction Signaling Molecule PTX AlF4– GTP-γ-S Gα− i0 transfection Gα− i2 transfection

C3 transferase

RhoA or Rac1 transfection RhoA transfection PG EP3β receptor transfection RhoA transfection RhoA transfection Rac1 transfection RhoA transfection Rho-kinase

MLC kinase transfection MLC phosphorylation State changes MLCK inhibitors

PMA diC8 PKC inhibitors

Effect on Tight Junction

Ref.

Heterotrimeric G protein Accelerated TJ assembly Balda et al., 1991 Increased TER Inhibition TJ assembly Balda et al., 1991 Increased TER Saha et al., 1998 Inhibition of ZO-1 assembly Balda et al., 1991 Accelerated TJ assembly Denker et al., 1996 Increased TER Saha et al., 1998 Small GTPases Decreased TER Inhibitiion of TJ assembly Inhibition of lymphocye migration Decreased TER Loss of intramembrane lipid diffusion barrier prevention of TJ disassembly by ATP depletion Increased TER Increased paracellular permeability Decreased TER Loss of intramembrane lipid diffusion barrier Prevention of dissasembly by ATP depletion Increased TER Increased paracellular permeability

Nusrat et al., 1995 Takaishi et al., 1997; Zhong et al., 1997 Adamson et al., 1999 Jou et al., 1998

Gopalakrishnan et al., 1998

Hasegawa et al., 1999 Jou et al., 1998 Gopalakrishnan et al., 1998 Fujita et al., 2000 Essler et al., 1998

Myosin light chain pathway Decreased TER Hecht et al., 1996 Decreased permeability Turner et al., 1999 Increased permeability Essler et al., 1998 Prevention of increase in Ma et al., 1999; Turner et al., 1997 permeability Philpott et al., 1998 Prevention of TJ disassembly Ma et al., 2000 Protein Kinases C TJ dissasembly and transformation Accelerated TJ assembly Induction of TJ strands Inhibition of TJ assembly Inhibition of TJ dissasembly by EDTA

Fey and Penman, 1984; Ojakian, 1981 Ben-Ze'ev, 1986; Clarke et al., 2000 Balda et al., 1991 Balda et al., 1993 Balda et al., 1991; Nigan et al., 1991 Citi, 1992

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TABLE 17.1 (continued) Signaling Pathways Regulating Tight Junction Signaling Molecule

cPKCβ1 transfection nPKCδ transfection

Forskolin cAMP

Effect on Tight Junction Inhibition of lymphocyte transmigration Inhibition of TJ disassembly by ZOT Prevention of NO or vasopressin effects Increased paracellular permeability Increased permeability and transformation Protein Kinase A Collapse of the paracellular space Inhibition of TJ disassembly Increased TER Decreased TER No change of paracellular permeability No change of TER Inhibition of TJ assembly Inhibition of TJ dissasembly Redistribution of 7H6 Prevetion of RhoAV14 effects Induction of TJ strands

PKA inhibitors cAMP+ astrocytes media

Inhibition of TJ disassembly Increased TER

Ref. Etienne-Manneville et al., 2000 Fasano et al., 1995 Burgstahler and Nathanson, 1995; Nathanson et al., 1992 Vuong et al., 1998 Mullin et al., 1998

Kottra and Fromter, 1993 Nilsson et al., 1996 Duffey et al., 1981; Ladino et al., 1991 Janecki et al., 1991; Satoh et al., 1996 Ellis et al., 1992; Janecki et al., 1991 Le Varlet B, 1995 Lowe et al., 1988 Balda et al., 1991 Balda et al., 1991 Behrens et al., 1985 Satoh et al., 1996 Adamson et al., 1998; Duffey et al., 1981 Klingler et al., 2000 Wolburg et al., 1994

1991). The use of pertussis toxin, which ADP-ribosylates Gα-i proteins, in MDCK (Madin–Darby canine kidney) cells, resulted in increased TER after TJ assembly induced by calcium as well as after treatment of established monolayers, indicating an involvement of heterotrimeric G proteins (Balda et al., 1991). Indeed, Gα-i2 was found at cell–cell contacts, where it colocalizes with ZO-1 (de Almeida et al., 1994; Denker et al., 1996; Dodane and Kachar, 1996). A function of Gα-i2 in the regulation of TJ assembly was further demonstrated by the overexpression of wild-type and constitutively activated Gα-i2 in MDCK cells, which was found to accelerate TJ assembly induced by calcium and to increase TER in confluent cells (Saha et al., 1998). Furthermore, a transfected Gα-i protein that is not endogenously expressed by MDCK cells, Gα-i0, is targeted to cell–cell junctions, and, if a constitutively activated form is expressed, also accelerates TJ assembly induced by calcium (Denker et al., 1996). Interestingly, this protein coimmunoprecipitates with ZO-1.

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The reason for the contradiction between the pharmacological and the transfection experiments is not clear, but it may be caused by the inhibition of several different Gα-i proteins by pertussis toxin or, alternatively, by a stoichiometric disequilibrium caused by the overexpression of a single α subunit. No information is available about the receptors that regulate the activity of these G proteins in TJ assembly and regulation, and how they transduce their signals to specific TJ components.

17.3.2 SMALL GTPASES Small GTPases are not directly coupled with cell surface receptors but are intermediate switches in different signaling pathways. Ras, Rho, and Rabs were directly or indirectly implicated in the regulation of TJs. The small GTPases Rab13, Rab8, and Rab 3B were shown to localize at the TJ by immunoelectron microscopy or to colocalize with TJ proteins by immunofluorescence (Huber et al., 1993; Weber et al., 1994; Zahraoui et al., 1994; see Table 17.1). The role of these Rabs in TJ functions is not known. However, since Rabs also regulate membrane traffic, they may play a role either in TJ assembly or in a function of cell–cell junctions in membrane traffic (see Chapter 7 by Rodriguez-Boulan et al. and Chapter 8 by Shoshani and Contreras; Grindstaff et al., 1998; Yeaman et al., 1999; Zahraoui et al., 2000). In contrast to Rabs, Rho-family GTPases were early-on shown to modify the actin cytoskeleton (Hall, 1998) and may regulate adherens and TJ functions through this interaction. C3 transferase, a toxin that mainly inactivates Rho family GTPases, inhibits assembly of adherens and TJs (Nusrat et al., 1995; Takaishi et al., 1997; Zhong et al., 1997). Moreover, inducible overexpression of constitutively activated forms of RhoA and Rac1 were reported to perturb paracellular permeability and the intramembrane diffusion barrier (Jou et al., 1998). In contrast, an increase in TER paralleled by an increase in paracellular permeability was obtained when RhoA was activated through a prostaglandin EP3β receptor (Hasegawa et al., 1999). It is not clear whether this means that prostaglandin has other effects apart from RhoA activation or whether this experimental discrepancy is caused by different levels of activation. RhoA has also been implicated in disassembly of TJs in two different experimental models. Activation of RhoA signaling by transient expression of RhoA-V14 can prevent TJ disassembly in response to ATP depletion in MDCK cells (Gopalakrishnan et al., 1998). In contrast, inactivation of Rho family GTPases by C3 transferase inhibits lymphocyte migration through brain endothelial cells (Adamson et al., 1999). Therefore, Rho family GTPase activities appear to be carefully balanced, and neither high nor low levels of Rho activity might be optimal for TJ integrity. Alternatively, the regulation of TJs by Rho GTPases may vary depending on cell type and/or ligand that induces the response. Effector proteins that are activated by Rho-type GTPases and that regulate TJs have not yet been studied in detail. However, the RhoA-dependent increase of TER involves Rho-associated kinase (Fujita et al., 2000). Regulation of myosin light chain (MLC) phosphorylation by Rho signaling may be a common pathway. For example, Pasteurella multocida toxin increases endothelial permeability via Rho kinase and produces an increase in MLC phosphorylation and cell retraction (Essler et al.,

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1998). Moreover, expression of the catalytic domain of MLC kinase was shown to increase paracellular permeability in MDCK cells (Hecht et al., 1996), and MLC kinase inhibitors prevent the increase of TJ permeability induced by different experimental conditions (Turner et al., 1997; Philpott et al., 1998; Ma et al., 1999; 2000). Therefore, the stimulation of Rho-associated kinase by RhoA modulates paracellular permeability by increasing MLC phosphorylation. However, strong stimulation of Rho function may result in transformation and thus result in a disassembly of the junctional apparatus, providing a possible explanation for the contradictory results reported for the regulation of TER by RhoA. Ras has also been reported to affect TJs. Ras signaling is discussed in Section 17.8, cell–cell junctions in gene expression and differentiation.

17.4 THE PROTEIN KINASES C PATHWAY This type of serine/threonine kinases, which exists as several isotypes, is the most extensively studied kinase in the TJ field. The PKC family is divided into three subfamilies: classical (cPKC), novel (nPKC), and atypical (aPKC). The cPKC (α, β, γ) and nPKC isotypes (δ, ε, η, and θ) are activated by DAG (diacylglycerol) and phosphoserine, and are targets of phorbol esters (phorbol 12-myristate 13-acetate, or PMA), which activate the enzyme in the absence of DAG. cPKCs additionally require calcium whereas nPKCs do not. aPKCs (ζ, λ, and PRK, for PKC-related kinases) are calcium insensitive and in general do not respond to PMA (Mellor and Parker, 1998). The effect of PKC activators and inhibitors was analyzed in confluent cells and in some models of TJ assembly (see Table 17.1). The phorbol ester PMA was shown to produce disassembly of TJs in many different cell lines (Ojakian, 1981; Fey and Penman, 1984; Ben-Ze’ev, 1986; Mullin and McGinn, 1988; Clarke et al., 2000a). The data obtained with PMA, however, are not easy to reconcile with those obtained with other PKC activators (e.g., the DAG analogue diC8) or with inhibitors of PKCs. For example, inhibition of PKCs blocks TJ assembly (Balda et al., 1991; Nigan et al., 1991; Citi et al., 1994), and activation by diC8 accelerates TJ formation (Balda et al., 1993). It was also shown that inhibition of PKCs blocks TJ disassembly produced by calcium removal (Citi, 1992). This contradiction between the effects of PMA, diC8, and PKC inhibitors could be because PMA can produce activation or downregulation of two PKC isotypes or because of binding of the drugs to targets other than PKC (Mellor and Parker, 1998). Additionally, the effect of PMA varies widely depending on the concentration, the time of incubation, and the type of cell (Oliver, 1990; Ellis et al., 1992; Mullin et al., 1992b). Since several PKC isotypes are known, an important question is which of them affect TJs. Some PKC isotypes were shown to localize at the lateral membrane, such as cPKCα and aPKCζ, suggesting that they may play a role in the formation of cell–cell junctions (Dong et al., 1993; Stuart and Nigan, 1995; Dodane and Kachar, 1996). Moreover, overexpression or downregulation of cPKCβ1 seems to correlate with changes in permeability of endothelial cells (Vuong et al., 1998), and overexpression of nPKCδ was shown to induce transformation of LLC-PK1 cells (porcine kidney epithelial cell line) and, consequently, increased TJ permeability (Mullin

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et al., 1998). More recently, aPKCλ was shown to interact directly with the epithelial TJ protein ASIP, a mammalian homologue of the Caenorhabditis elegans polarity protein PAR-3, making it a prime candidate for a regulator of TJ assembly and disassembly (Izumi et al., 1998). Different PKC inhibitors were used to block the increase in TJ permeability produced by different manipulations. For example, the effect of a toxin that opens TJs, ZOT (Zonula occludens toxin), was blocked by CGP41251 (Fasano et al., 1995), whereas the increase of TJ permeability produced by nitric oxide (NO) or vasopressin in hepatocytes can be blocked by the PKC inhibitor H-7 (Nathanson et al., 1992; Burgstahler and Nathanson, 1995; Roma et al., 1998) and transendothelial lymphocyte migration by bisindolylmaleimide PKC inhibitors (Etienne-Manneville et al., 2000). It will be interesting to test whether the same isotype of PKC is responsible for these different PKC effects or whether different PKC isotypes are involved. To test this type of question, it will be necessary to develop approaches to modulate specifically PKC isotypes, such as the use of antisense oligonucleotides and inhibitory peptides, or expression of dominant negative mutants (Way et al., 2000; see Note added in proof).

17.5 THE PROTEIN KINASE A PATHWAY The influence of protein kinase A (PKA) on TJs is controversial since some studies reported increased TER upon PKA stimulation, whereas others observed decreases or no significant effects (Duffey et al., 1981; Lowe et al., 1988; Balda et al., 1991; Ladino et al., 1991; Ellis et al., 1992; Schneeberger and Lynch, 1992; Wolburg et al., 1994; Le Varlet et al., 1995; Kovbasnjuk et al., 1998) (see Table 17.1). These differences could be due to the use of different concentrations of activators since high concentrations may result in nonspecific effects, to the presence of certain growth factors that may modulate PKA signaling, or to the cell types analyzed. In Sertoli cells, for example, the concentration of the activator is critical since high concentrations of dibutyryl cyclic AMP inhibited and low concentrations stimulated TER development (Janecki et al., 1991). Additionally, stimulation of PKA can also increase TER by mechanisms that do not involve TJs: in Necturus gallbladder epithelium, using forskolin instead of cAMP, the increase in TER was shown to correlate with the collapse of the lateral space (Kottra and Fromter, 1993). PKA activation was also shown to inhibit TJ assembly and disassembly. Cyclic AMP or forskolin pretreatment can inhibit TJ assembly induced by calcium (Balda et al., 1991) but also blocks TJ disassembly triggered by removal of calcium (Nilsson et al., 1996) or addition of anti-E-cadherin antibodies (Behrens et al., 1985). PKA inhibition can also prevent junction disassembly induced by removal of calcium (Klingler et al., 2000). Hence, PKA may regulate the dynamics of TJ components. Indeed, stimulation of PKA can induce a redistribution of the TJ protein 7H6 at the cell borders (Satoh et al., 1996) and, in certain experimental systems, increase the number of TJ strands (Duffey et al., 1981; Adamson et al., 1998). However, it is not clear whether PKA directly phosphorylates TJ components or whether the observed pharmacological effects are due to indirect effects.

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17.6 PHOSPHORYLATION OF TIGHT JUNCTION PROTEINS 17.6.1 SERINE/THREONINE PHOSPHORYLATION In terms of phosphorylation, ZO-1 is one of the most extensively studied TJ proteins (see Table 17.2). ZO-1 is a cytoplasmic TJ protein that is expressed in epithelial and nonepithelial cells (see Chapter 11 by Citi; Stevenson et al., 1986; Balda and Anderson, 1993; Itoh et al., 1993; Howarth and Stevenson, 1995). Phosphorylation of ZO-1 has been studied in a variety of conditions. In most studies, no correlation between ZO-1 phosphorylation and TJ assembly or functioning was found. For example, in two strains of MDCK cells with different TER, the phosphorylation state of ZO-1 is the same (Stevenson et al., 1989). Similarly, neither ZO-1, ZO-2, nor p130/ZO-3 was found to change its phosphorylation content by addition of a PKC agonist or

TABLE 17.2 Phosphorylation of Tight Junction Proteins A. Ser/Thr phosphorylation ZO-1 Similar in MDCK clone I and II cells Not affected during calcium induced assembly Phosphorylated in vitro by brain PKC Phosphorylated in vitro by ZAK Reduced after 48 h calcium depletion Reduced by PKC inhibition ZO-2 Not affected during calcium induced TJ assembly p130/ZO-3 Not affected during calcium induced TJ assembly occludin Increased during TJ assembly in MDCK Decreased during TJ assembly in Xenopus Phosphorylated in vitro by CK II Dephosphorylation by PMA

cingulin

Dephosphorylation by calcium removal Phosphorylated in vitro by OAK Not affected during calcium induced TJ assembly

B. Tyrosine phosphorylation ZO-1 Increased by tyrosine phosphatase inhibition Increased by protamine Increased by EGF Increased by VEGF Increased by v-src transformation Increased when MEK-1 inhibitor reversed transformation ZO-2 increased by EGF Increased by v-src transformation Occludin Increased when MEK-1 inhibitor reversed transformation Increased by VEGF

Stevenson et al., 1989 Balda et al., 1993 Stuart and Nigan, 1995 Balda et al., 1996b Howarth et al., 1994 Stuart and Nigan, 1995 Balda et al., 1993 Balda et al., 1993 Sakakibara et al., 1997; Wong, 1997 Cordenonsi et al., 1997 Cordenonsi et al., 1999 Clarke et al., 2000; Farshori and Kachar, 1999 Farshori and Kachar, 1999 (In this chapter) Citi and Denisenko, 1995

Staddon et al., 1995 Kurihara et al., 1995 Van Itallie et al., 1995 Antonetti et al., 1999 Takeda and Tsukita, 1995 Chen et al., 2000 Van Itallie et al., 1995 Takeda and Tsukita, 1995 Chen et al., 2000 Antonetti et al., 1999

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during a Ca2+-switch experiment (Balda et al., 1993). However, inhibition of TJ assembly by a PKC inhibitor correlates with reduced phosphorylation of ZO-1 (Stuart and Nigan, 1995). Because more than 48 h of calcium depletion is required to reduce phosphorylation of ZO-1, dephosphorylation is unlikely to be a direct consequence of calcium depletion and TJ disassembly (Howarth et al., 1994). The phosphorylation content of another TJ protein, cingulin, was also found to remain constant during the assembly induced by calcium (Citi and Denisenko, 1995). PKC activation appears to be an intermediate step between E-cadherin activation and TJ assembly (Balda et al., 1993). Therefore, some adherens junction proteins were tested as targets of PKC. Vinculin, but not α-actinin, seems to be phosphorylated by PKC during junction assembly induced by calcium (Perez-Moreno et al., 1998). PKC also phosphorylates MLC kinase, which in turn affects MLC phosphorylation and cytoskeleton organization (Turner et al., 1999). Although PKC can phosphorylate ZO-1 in vitro (Stuart and Nigan, 1995), it is conceivable that PKC only indirectly affects TJ assembly upon activation of adherens junction formation. ZO-1 coimmunoprecipates with an associated serine/threonine kinase activity, which is called ZAK (ZO-1-associated protein kinase) and binds specifically to the SH3 domain of ZO-1 (Balda et al., 1996b). In vitro, ZAK phosphorylates one or two serines C terminal to the SH3 domain. Based on its immunological and biochemical properties, ZAK does not resemble known kinases that are related to TJ functions (e.g., aPKCζ, PKA, or MLC kinase; M. S. Balda and K. Matter, unpublished results). However, the molecular identity of this kinase is not yet known. Phosphorylation of occludin, a transmembrane protein of TJ, has been investigated by several groups (see Table 17.2). Immunoblots of occludin from MDCK cells show several bands that can be reduced to one of a lower apparent molecular weight by alkaline phosphatase treatment. This is due to removal of phosphate groups on serine and threonine residues (Sakakibara et al., 1997). Alkaline phosphatasesensitive bands are more detergent insoluble than the lower-molecular-weight forms and accumulate during junctional assembly (Sakakibara et al., 1997; Wong, 1997). These results suggest that a high phosphocontent of occludin correlates with junctional localization. However, a decrease in the phosphocontent of occludin was observed during TJ assembly at early stages of embryogenesis in Xenopus (Cordenonsi et al., 1997). It is not known whether these differences are due to different phosphorylation sites or to a different regulation of TJ assembly in different species and tissues. Dephosphorylation of occludin is induced by PMA or prolonged calcium depletion (Farshori and Kachar, 1999; Clarke et al., 2000b). Although the C-terminal domain of occludin is an in vitro substrate for different kinases (Cordenonsi et al., 1999), the kinases that phosphorylate occludin in vivo are not known. Occludin has four transmembrane domains and three cytosolic domains. To test which of the cytoplasmic domains is phosphorylated in vivo, MDCK cells were metabolically labeled with 32P-orthophosphate. Wild-type and transfected MDCK cells stably expressing the following constructs were studied: full-length wild-type occludin with an N-terminal epitope tag, HA-occludinCT3 (which lacks the entire C-terminal domain but possesses the N-terminal cytosolic domain and the small intracellular loop), and a chimeric protein containing only the C-terminal domain of occludin in fusion with the transmembrane and extracellular part of a mouse

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Fc receptor (Figure 17.1D). Then, the different constructs were immunoprecipitated and the incorporation of 32P-orthophosphate analyzed by gel electrophoresis and autoradiography. Figure 17.1A shows that in the absence of the C-terminal domain, occludin is only poorly phosphorylated in comparison with the wild-type protein (a low signal can be observed after longer times of exposure), demonstrating that, in vivo, most of the phosphorylation of occludin occurs in the C-terminal domain. Nevertheless, occludin possesses phosphorylation sites outside the C-terminal cytosolic domain. Furthermore, the chimeric protein containing only the C-terminal domain of occludin shows a strong phosphorylation. Hence, the main phosphorylation sites are in the C-terminal domain. Interestingly, this chimeric protein is not associated with TJs but accumulates in the lateral membrane (Matter and Balda, 1998), indicating that phosphorylation of the C-terminal cytosolic domain is not sufficient for integration into TJs. Because the C-terminal domain of occludin is involved in the regulation of paracellular permeability (Balda et al., 1996a), it was next tested whether phosphorylation of this domain could be involved in this regulatory process. Therefore, a fusion protein containing the C-terminal domain of occludin fused to His6 was produced and it was speculated that the cytoplasmic domain of occludin may bind a protein kinase that also uses this domain as a substrate. This fusion protein was incubated coupled to beads with MDCK cell extract, was washed extensively, and an in vitro protein kinase assay was performed. Figure 17.1B shows that the C-terminal domain of occludin binds to a protein kinase that can phosphorylate this domain. Negative controls did not pull down this kinase activity. As with occludin in vivo (Cordenonsi et al., 1997; Sakakibara et al., 1997), the in vitro phosphorylated occludin fusion protein was phosphorylated on serine and threonine residues (Figure 17.1C). Moreover, similar phosphopeptides were detected in the in vitro phosphorylated fusion protein, as in in vivo phosphorylated occludin (not shown). Since casein kinase II (CK II) can phosphorylate a fusion protein containing the C-terminal domain of occludin in vitro (Cordenonsi et al., 1997), it was next tested whether the occludin-associated kinase is CK II. However, CK II could not be detected with specific antibodies in fusion protein pull-downs nor do various biochemical properties of CK II match with those of the occludin-associated kinase (not shown). Hence, the identified kinase activity does not correspond to CK II. Thus, it is tempting to call this kinase OAK, for occludin-associated kinase. Although the functional analysis of OAK will require its molecular identification, OAK is a good candidate for a direct regulator of occludin function.

17.6.2 TYROSINE PHOSPHORYLATION Tyrosine phosphorylation of ZO-1 was observed in different conditions (see Table 17.2); some of them correlated with junction disruption and others with formation. The use of general tyrosine phosphatase inhibitors produces junctional disruption with increased tyrosine phosphorylation of cell–cell junctional proteins (Volberg et al., 1991), including ZO-1 (Staddon et al., 1995). Overexpression of v-src results in transformation and is paralleled by tyrosine phosphorylation of ZO-1 and ZO-2 (Takeda and Tsukita, 1995). In contrast, upon induction of TJ-like structures

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FIGURE 17.1 Occludin phosphorylation and enzymatic identification of OAK. (A) Wildtype (wt MDCK) and transfected MDCK cells expressing occludin with an N-terminal epitope tag (HAoccludin), epitope-tagged occludin lacking the C-terminal cytosolic domain (HAoccludinCT3; results from two different clones, c4 and c12, are shown), or a chimeric protein consisting of the mouse Fc receptor ecto- and transmembrane domains and the C-terminal cytosolic domain of occludin were grown to confluence and then metabolically labeled with [32P]-orthophosphate for 15 h. The cells were extracted and endogenous occludin (lane 1) or the transfected proteins (lanes 2 to 5) were immunprecipitated. The precipitates were analyzed by SDS-PAGE and autoradiography. Note the intense labeling of the chimeric protein containing the C-terminal domain of occludin. The arrow marks the position of HAoccludinCT3, which, although only weakly, was also labeled, suggesting that the N-terminal cytosolic domain and/or the cytosolic loop between the second and the third transmembrane domain can also be phosphorylated. (B) Probond Ni-resin beads without (background) or with bound fusion protein consisting of a His-tag and the C-terminal cytosolic domain of occludin (His-occludinCTD) were incubated with MDCK cell extract. After extensive washing, the recombinant fusion protein was added to the background sample, and both samples were incubated with kinase buffer and [32P]-ATP (Balda et al., 1996b). The samples were then analyzed by SDS-PAGE and autoradiography. Note the intensive labeling of the fusion protein when the beads were incubated with cell extracts in the presence (lane 2) but not in the absence (lane 3) of the His-tagged C-terminal domain of occludin. Since the kinase activity

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between glomerular foot process produced by protamine, tyrosine phosphorylation of ZO-1 was also observed (Kurihara et al., 1995). Similarly, upon treatment of A431 cells with EGF, tyrosine phosphorylation of ZO-1 correlated with cell–cell junction formation (Van Itallie et al., 1995). A role for tyrosine phosphorylation in TJ assembly was also suggested by the observation that the tyrosine kinase inhibitor genistein blocks reassembly of TJ and rephosphorylation of occludin, ZO-2, and ZO-3 after junction disruption by ATP depletion of MDCK cells (Tsukamoto and Nigam, 1999). Moreover, MEK1 inhibition reverses Ras transformation in MDCK cells, allowing the development of TJs; during this TJ assembly process, tyrosine phosphorylation of occludin and ZO-1 has been observed (Chen et al., 2000). On the other hand, tyrosine phosphorylation also appears to play a role in the regulation of junctional permeability. Tyrosine phosphorylation of ZO-1 and occludin is increased by vascular endothelial growth factor (VEGF), which produces increased paracellular permeability of retinal endothelial cells (Antonetti et al., 1999), and the increases in paracellular permeability produced by certain inflammatory mediators, such as tumor necrosis factor, can be blocked with tyrosine kinase inhibitors (Mullin et al., 1992a; Schmitz et al., 1999). The protein tyrosine kinases that phosphorylate TJ proteins have not been identified. It is also unclear whether the different responses to tyrosine phosphorylation are due to phosphorylation of different sites or to differences in other parameters that determine whether tyrosine phosphorylation of TJ protein results in junction formation, disassembly, or modulation of paracellular permeability.

17.7 REGULATION OF PARACELLULAR PERMEABILITY BY MULTIPLE SIGNALING PATHWAYS One can imagine two basic mechanisms by which intracellular signals regulate selective paracellular permeability. A stimulus may directly act on the TJ components that mediate selective paracellular permeability or, alternatively, a stimulus may result in increased selective paracellular permeability via a modification of the actinbased cytoskeleton that causes, for example, a redistribution of the components that mediate paracellular permeability. There is evidence for both regulatory mechanisms. Although the molecular mechanism that underlies selective paracellular permeability is still unknown, two types of TJ-associated transmembrane proteins, occludin and claudins, have been experimentally connected to selective paracellular permeability FIGURE 17.1 (continued) was also found to coprecipitate with occludin, the kinase was named OAK for occludin-associated kinase. (C) The amino acid specificity of OAK was determined by a phosphoamino acid analysis after a total hydrolysis using two-dimensional thin-layer chromatography (Balda et al., 1996b). The spots representing phosphoserine (P-S) and phosphothreonine (P-T) are indicated. Phosphotyrosine (P-Y) was not detected. (D) The schematic structures of the occludin constructs analyzed in A are illustrated. HA indicates the N-terminal epitope attached to some constructs. In the FcOCD chimera, the ecto- and transmembrane domain of a mouse Fc receptor (indicated in black) were fused to the C-terminal cytosolic domains of occludin (indicated in gray).

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(Balda et al., 2000; Balda and Matter, 2000b; Tsukita and Furuse, 2000). Although the mechanisms used by claudins are not yet clear, occludin may be of central importance in the translation of intracellular stimuli to increased selective paracellular permeability. The C-terminal cytosolic domain of occludin, which is heavily phosphorylated and binds the protein kinase OAK (see Figure 17.1A), is also critical for the regulation of selective paracellular permeability in transfected MDCK cells and Xenopus (Balda et al., 1996a; Chen et al., 1997); hence, protein kinases like OAK or CK II may regulate selective paracellular permeability by phosphorylating occludin (Figure 17.2). Alternatively, increases in selective paracellular permeability could also involve changes in ZO-1 since increased paracellular permeability was observed when ZO-1 was overexpressed in MDCK cells (Balda and Matter, 2000). Activation of protein kinases that phosphorylate TJ proteins and/or cytoskeletal proteins may be stimulated by signaling mechanisms involving, for example, G proteins and PKC, PKA, and/or GEF-dependent pathways. Changes in the actin-based cytoskeleton were early-on suggested to be involved in TJ assembly and sealing (see Chapter 12 by Fanning; Bentzel et al., 1976; Meza et al., 1982). Since actin can be seen at the TJ (Hirokawa and Tilney, 1982; Madara, 1987) and several TJ components can directly bind to actin (Itoh et al., 1997; Fanning and Anderson, 1998; Wittchen et al., 1999), it could be that the actin cytoskeleton regulates permeability by contraction of the actin ring or by transmission of signals directly to TJ proteins. Essentially all of the above-described signaling proteins can modify the actin cytoskeleton. The problem now is how to arrange the different signaling pathways described above in an integrated manner. The Rho pathway appears to be of central importance. Several trimeric G proteins — including Gα-q, Gα-12, and Gα-i2 — can activate Rho GTPases in a manner that depends on the receptor and the cell type. This activation could be mediated via the PLC-PKC or PKA pathway, or via a GEF (Hasegawa et al., 1999; Essler et al., 2000; Somlyo and Somlyo, 2000). Further downstream, activated Rho activates Rho kinase, which inactivates MLC phosphatase and phosphorylates MLC, resulting in changes in MLC phosphorylation; this was shown to correlate with changes in paracellular permeability (Hecht et al., 1996; Turner et al., 1999). Thus, some types of stimulation of Rho may result in only modest and local changes in the actin cytoskeleton and hence only changed permeability properties of TJ, whereas stimulation of several pathways that converge to activate Rho may cause a radical reorganization of the actin cytoskeleton and, hence, dissociation of the junctions (Ridley et al., 1999).

17.8 CELL–CELL JUNCTIONS IN GENE EXPRESSION AND DIFFERENTIATION 17.8.1 RAS-MEDIATED TRANSFORMATION

AND THE JUNCTIONAL

COMPLEX

Differentiation of epithelial cells is regulated by several stimuli and environmental parameters, such as hormones, growth factors, extracellular matrix components, and cell adhesion (Birchmeier et al., 1997; Adams and Nelson, 1998; Boudreau and Bissell, 1998; Soriano et al., 1998; Gumbiner, 2000). Loss of differentiation of

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FIGURE 17.2 Regulation of paracellular permeability by intracellular signaling pathways. The major identified signaling proteins (shown as ovals) and pathways (given by the name) that have been implicated in the regulation of paracellular permeability are schematically indicated. The homologous ZO-1, ZO-2, and ZO-3 proteins are indicated as ZOs; however, only ZO-1 has thus far been shown to function in the regulation of paracellular permeability. Y and X represent possible additional regulators such as, for example, other known junctional proteins that interact with the components shown here. Regulation of the actin cytoskeleton by changing the activities of MLC phosphatase (MLC-P) and MLC kinase (MLCK) appears to be a major mechanism of regulation and may be important for all known signaling pathways that regulate paracellular permeability. Alternatively, permeability may also be controlled by, for example, Rho-family GTPases and their function in the regulation of actin polymerization and depolymerization. Additionally, the indicated regulatory mechanisms may directly affect the activities of junctional components. For example, it is conceivable that phosphorylation of the occludin C-terminal domain (by OAK, CK-II, or another protein kinase) regulates its interaction with the submembrane cytoskeleton and thereby regulates the function of occludin in selective paracellular permeability. Nevertheless, the molecular mechanisms that are involved in the regulation of TJ components by the major signaling pathways are largely unknown.

epithelial cells has been correlated in many cancers with mutations in Ras. The Ras pathways not only were shown to be involved in growth factor and extracellular matrix signaling but also in disruption of cell adhesion (Behrens et al., 1989; Ridley et al., 1995; Potempa and Ridley, 1998; Cary et al., 1999; Tan and Kim, 1999). There are three human ras genes, H-ras, K-ras, and N-ras. Only knockouts of K-ras are lethal in mice (Johnson et al., 1997). Ras proteins interact with multiple effectors and thus can stimulate signaling along different pathways (Campbell et al.,

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1998; Marshall, 1999; Shields et al., 2000). For example, via the Ral GEF, Ras can regulate signaling based on Rho-family GTPases. Ras can also activate Raf and thus stimulate signaling along the MEK/ERK pathway, resulting in activation of the transcription factor Elk1. Additionally, Ras can regulate transcription via MEKK and JNK, leading to activation of Jun, or stimulate PI3K (phosphatidyl inositol 3-kinase), resulting in activation of the Akt/PKB pathway. Other Ras effectors are known as well, such as the junction-associated protein AF-6 (Yamamoto et al., 1997), but their signaling activities are not known. As the multitude of signaling pathways suggests, the response to Ras activation depends on the cell type analyzed (Shields et al., 2000). For reasons of simplicity, this chapter therefore considers only the most relevant data obtained with epithelial (mainly MDCK) cells. Overexpression of K-Ras or H-Ras transforms MDCK cells in different ways, suggesting that different Ras isotypes activate different signaling pathways in MDCK cells. In the case of viral K-Ras, this induces multilayers without a significant redistribution of ZO-1 (Schoenenberger et al., 1991). If viral H-Ras is expressed, the transformed MDCK cells produce malignant tumors in nude mice with reduced expression of E-cadherin (Behrens et al., 1989; Mareel et al., 1991). Interestingly, H-Ras-stimulated cell spreading is paralleled by a loss of E-cadherin and β-catenin from the adherens junctions, but desmosomes and TJs appear to remain intact (Potempa and Ridley, 1998). The Raf-mediated pathway is a well-characterized Ras-stimulated pathway. There are three Raf kinases: Raf-1, A-Raf, and B-Raf (Hagemann and Rapp, 1999). In epithelia, Raf-1 has been most studied, but whether or not expression of activated Raf-1 causes transformation depends on the analyzed epithelial cell type (Oldham et al., 1996; Li and Mrsny, 2000). The downstream effectors of Raf are the kinases MEK and ERK, and the transcription factor Elk1. Expression of constitutively active MEK1 results in transformation of MDCK cells (Schramek et al., 1997). Consequently, treatment of H-Ras-transformed MDCK cells with the selective MEK1 inhibitor PD98059 was shown to result in increased expression of adherens junction proteins and assembly of functional adherens and TJs (Lu et al., 1998; Chen et al., 2000). An important function of Raf pathway components in the arrangement of the cell–cell junctional complex is also suggested by the fact that inhibition of MEK1/MEK2 allows reorganization of cell junctions in MDCK cells that detach because of inhibition of Na+,K+-ATPase (Contreras et al., 1999). Interestingly, TJs appear to be intimately connected to Raf-1 signaling since overexpression of occludin is able to reverse Raf-1-mediated transformation of a salivary gland cell line (Li and Mrsny, 2000). It will be interesting to analyze whether occludin reverses MEKmediated transformation and whether Raf-1 transforms MDCK cells. Cell–cell junctions also harbor upstream and downstream components of Rassignaling pathways. For example, a Ras GAP, IQGAP, accumulates at cell–cell junctions and may constitute a link between Cdc42/Rac1 and Ras signaling (Kuroda et al., 1996). However, loss of IQGAP1 does not affect tumor development or tumor progression, but adult mutant mice deficient in IQGAP1 exhibit gastric hyperplasia (Li et al., 2000). Moreover, the Ras effector AF-6 colocalizes and interacts with ZO-1, and activation of Ras inhibits the interaction (Yamamoto et al., 1997). Although the mechanism by which AF-6 mediates Ras signaling is not known, AF-6

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is critical for the establishment of cell–cell junctions during mouse development (Zhadanov et al., 1999). As mentioned above in the section on small GTPases, RhoA and Rac1 are involved in TJ functions. However, the activities of RhoA and Rac1 are also regulated by Ras. For example, in Ras-transformed MDCK cells activation of Rac restores E-cadherin-mediated adhesion (Hordijk et al., 1997), and oncogenic Ras downregulates Rac activity, resulting in increased Rho activity and epithelial–mesenchymal transition (Zondag et al., 2000). Hence, it is conceivable that Ras also functions in the physiological regulation of paracellular permeability. However, as for signaling via Rho-family GTPases, it is at the moment difficult to differentiate between physiological regulation of permeability and transformation-induced complete or partial dissociation of cell–cell junctions.

17.8.2 CELL–CELL JUNCTIONS

AND

TRANSCRIPTION FACTORS

Cell adhesion is an important determinant of cell growth and differentiation. Some of the signaling pathways that transmit information from sites of cell–cell adhesion to the nucleus involve direct modulation of transcription factor activities. This was first described for adherens junctions. The transmembrane protein E-cadherin was shown early-on to be involved in tumor suppression (Takeichi, 1993; Kirkpatrick and Peifer, 1995; Birchmeier et al., 1996; Gumbiner, 1997). The different steps involved in this pathway were more recently characterized and extensively reviewed (Behrens, 1999; Roose and Clevers, 1999; Peifer and Polakis, 2000). The central component in this system is β-catenin, a coactivator of transcription factors such as TCF (tall factor) and LEF (lymphoid enhancer factor). The pool of β-catenin that can associate with a transcription factor is determined by the level of E-cadherin, which stabilizes β-catenin at adherens junctions, and by other proteins (e.g., GSK-3β (glycogen synthetase kinase), axin, and APC (adenomatous polyposis coli gene product)), which control β-catenin activation and degradation. Active β-catenin acts as a transcriptional activator by recruiting elements of the transcription machinery, such as p300/CBP acetyltransferase (Hecht et al., 2000). Although two TJ proteins, symplekin and ZO-1, were observed in the nucleus (Gottardi et al., 1996; Keon et al., 1996), a direct pathway linking TJ to the control of gene expression was discovered only recently and involves ZO-1. ZO-1 belongs to a family of proteins, the MAGUKs (membrane associated guanylate kinases). These proteins are often associated with specific plasma membrane domains such as epithelial junctions or neuronal synapses (Dimitratos et al., 1999; Fanning and Anderson, 1999; Garner et al., 2000). One of the founding members of this protein family is DlgA, a Drosophila tumor suppressor associated with septate junctions. Interestingly, all signaling functions of DlgA depend on two domains that are conserved in all MAGUKs, the SH3 and the guanylate kinase homology domain (Woods et al., 1996). Because of the critical involvement of the SH3 domain in DlgA signaling, the SH3 domain of ZO-1 was used to search for a possible signaling molecule of this MAGUK member. This resulted in the identification of ZONAB (ZO-1-associated nucleic acid binding), a Y-box transcription factor (Balda and Matter, 2000a).

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FIGURE 17.3 Cell density-dependent regulation of gene expression by ZO-1 and ZONAB. In low-density MDCK epithelial cells that are still in a growth phase, only a little ZO-1 is expressed and it is primarily associated with TJs. ZONAB is expressed at high levels, however, and most of it is in the nucleus and only little is associated with ZO-1. In cells grown to a high density, ZO-1 is expressed at high levels and recruits ZONAB, which is present at low levels only, to TJs. Since ZONAB represses the expression of the transmembrane tyrosine kinase ErbB-2, the expression profile of this proto-oncogene develops in the same way as that of ZO-1: ErbB-2 expression levels are low in growing cells and high in mature monolayers (Balda and Matter, 2000a). Moreover, the same mechanism appears to regulate the cell densitydependent regulation of cell growth (Balda and Matter, in preparation).

ZONAB contains a cold-shock domain and, in vitro, binds to sequences of different promoters containing an inverted CCAAT box (ATTGG box). ZONAB localizes at the TJ and in the nucleus. The relative distribution of ZONAB is determined by the cell density. In cells grown to a high density, which contain a great deal of ZO-1 and little ZONAB, ZONAB localizes primarily to TJs, whereas in cells at low density, it is also in the nucleus. The interaction between ZO-1 and ZONAB is of functional relevance since the two proteins interact in the regulation of paracellular permeability and expression of the proto-oncogene ErbB-2 (Balda and Matter, 2000a). There is also evidence indicating that ZO-1 and ZONAB functionally interact to regulate G1 to S-phase transition during the cell cycle (M. S. Balda and K. Matter, in preparation). Therefore, a working model is proposed in which ZO-1 and ZONAB are part of a cell density–dependent signaling mechanism at the TJ that regulates expression of specific genes that are important for epithelial cell growth and differentiation (Figure 17.3). The principle of controlling the activity of a transcription factor by interaction with a cell–cell junction or adhesion–receptor associated protein becomes more and more common. Another example is the MAGUK family member and human homologue of C. elegans lin-2, CASK, which binds to the cytoplasmic domain of syndecan (Cohen et al., 1998). CASK binds to the T-box transcription factor Tbr-1, and the

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two proteins coactivate transcription of reelin (Hsueh et al., 2000). Both reelin and Tbr-1 are important for brain development (Lambert de Rouvroit and Goffinet, 1998). Moreover, a second TJ-associated potential transcription factor has recently been described. HuASH1 had been cloned because of its homology to Drosophila ASH1, a transcription factor involved in homeotic gene expression during development, and was then found to colocalize with TJ proteins (Nakamura et al., 2000). Although the analysis of TJ-associated transcription factors has already given some initial interesting results, this is clearly one of the topics that should be studied in detail in the future. It will be important to determine the types of genes that are regulated by TJs, the types of transcription factors that are controlled by TJ association, the types of proteins and signaling pathways that control these transcription factors, and the types of structural and functional interactions between different TJassociated transcription factors. Perhaps one of the most challenging undertakings will be to understand the cross talk between pathways originating at TJs, adherens junctions, sites of cell–extracellular matrix adhesion, and growth factor receptors and to integrate these signaling pathways into a unifying model of growth control.

NOTE ADDED IN PROOF Overexpression of a dominant-negative mutant of aPKC in MDCK II cells affects the biogenesis of tight junctions as reported by Suzuki, A., Yamanaka, T., Hirose, T., Manabe, N., Mizuno, K., Shimizu, M., Akimoto, K., Izumi, Y., Ohnishi, T., and Ohno, S. Atypical protein kinase C is involved in the evolutionary consrved PAR protein complex and plays a critical role in establishing epithelia-specific junctional structures. J. Cell. Biol. 2001. 152(6):1183–1196.

ACKNOWLEDGMENTS The authors thank Dr. John Greenwood for critical reading of the manuscript. The research in the authors’ laboratories is supported by the Swiss National Science Foundation, The Cancer Research Campaign [CRC] (SP2562/0101), and The Wellcome Trust (063661).

REFERENCES Adams, C. L., and W. J. Nelson. 1998. Cytomechanics of cadherin-mediated cell–cell adhesion. Curr. Opin. Cell Biol., 10:572–577. Adamson, P., S. Etienne, P. O. Couraud, V. Calder, and J. Greenwood. 1999. Lymphocyte migration through brain endothelial cell monolayers involves signaling through endothelial ICAM-1 via a rho-dependent pathway. J. Immunol., 162:2964–2973. Adamson, R. H., B. Liu, G. N. Fry, L. L. Rubin, and F. E. Curry. 1998. Microvascular permeability and number of tight junctions are modulated by cAMP. Am. J. Physiol., 274:H1885–1894. Antonetti, D. A., A. J. Barber, L. A. Hollinger, E. B. Wolpert, and T. W. Gardner. 1999. Vascular endothelial growth factor induces rapid phosphorylation of tight junction proteins occludin and zonula occluden 1. A potential mechanism for vascular permeability in diabetic retinopathy and tumors. J. Biol. Chem., 274:23463–23467.

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18

Regulation of Tight Junction Permeability in the Mammary Gland Duy-Ai D. Nguyen, Neal Beeman, and Margaret C. Neville

CONTENTS 18.1 Introduction .................................................................................................395 18.2 Tight Junction Permeability in Mammary Glands of Pregnant and Lactating Animals .......................................................................................397 18.3 The Transition from Pregnancy to Lactation .............................................399 18.3.1 Hormonal Regulation.....................................................................399 18.3.2 Mechanism of Tight Junction Closure ..........................................402 18.4 In Vivo Regulation of Mammary Tight Junctions by Milk Stasis, Mastitis, and Oxytocin................................................................................405 18.4.1 Milk Stasis .....................................................................................405 18.4.2 Mastitis...........................................................................................406 18.4.3 Oxytocin.........................................................................................407 18.5 Hormonal Regulation of Tight Junction Permeability in In Vitro Mammary Systems .....................................................................................408 18.5.1 Glucocorticoids ..............................................................................408 18.5.2 Prolactin .........................................................................................409 18.6 A Role for TGF-β? .....................................................................................410 18.7 Perspective ..................................................................................................410 Acknowledgments..................................................................................................411 References..............................................................................................................411

18.1 INTRODUCTION During lactation, milk is secreted by a continuous monolayer of epithelial cells that line the lumina of the alveoli and ducts of the mammary gland (Figure 18.1). These cells are joined at their apical borders by highly impermeable tight junctions (TJs) that prevent backflux of all milk components from the alveolar lumen into the interstitial space. Mammary epithelial TJs are dynamic: although they are highly

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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FIGURE 18.1 Junctional complexes in the lactating mammary epithelium. The mammary alveolus consists of a single continuous layer of secretory mammary cells surrounding alveoli (a) and ducts (b). These cells secrete milk by a number of complex processes depicted in the cartoon of the mammary alveolar cell and are joined near their apical borders by TJs, adherens junctions, and gap junctions.

impermeable during lactation, the epithelium becomes very leaky under a number of conditions. In the pregnant animal, the mammary epithelium allows passage of large macromolecules; at this stage plasma constituents such as sodium, chloride, and albumin enter directly into the mammary lumen, and milk constituents such as lactose and α-lactalbumin leak into the interstitial space and plasma. As shall be seen the transition between this highly permeable state in pregnancy and the impermeable state of lactation is governed by the hormonal status of the animal. Mammary epithelial permeability is also increased by milk stasis, high doses of oxytocin, and mastitis. The mechanisms are poorly understood, although local factors, such as intramammary pressure, prostaglandins, and (TGF-β), and systemic factors, such as prolactin, progesterone, and glucocorticoids, all have been implicated. The permeability of the mammary TJs appears to be closely linked to the rate of milk secretion suggesting a functional connection between TJ status and cellular activity. This chapter first describes methods for measuring the permeability of mammary TJs in vivo, describing the findings in pregnant and lactating glands using these techniques. The chapter then discusses the transition between pregnancy and lactation considering both the hormones that bring about this transition and the mechanism, in so far as is known, by which it occurs. Then other circumstances are considered under

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which, in vivo, the mammary epithelium becomes permeable to large and small molecules. Finally, findings from in vitro model systems that shed some light on the molecular mechanisms involved in mammary TJ regulation are considered. The perspective section concludes by considering some of the questions that remain for future research.

18.2 TIGHT JUNCTION PERMEABILITY IN MAMMARY GLANDS OF PREGNANT AND LACTATING ANIMALS A variety of techniques have been used to monitor the permeability of the mammary epithelium in vivo. These include passage of nonmetabolizable radioisotopes such as [14C]sucrose from milk space to blood, or vice versa; measurements of the electrical potential difference across the epithelium; monitoring of milk composition; and injection of fluorescent dyes into the lumen and interstitial space of the mammary gland followed by fluorescence microscopy of sections of the gland. The earliest experiments on the permeability of the mammary TJs were carried out by Linzell and Peaker in the 1970s (Linzell and Peaker, 1971; 1974). These investigators injected [14C]sucrose through the teat canal into the lumen of the mammary gland of the pregnant goat and observed that the tracer appeared immediately in the blood. Only a fraction of the injected tracer could be recovered in the milk. In contrast, during lactation no tracer could be observed in the blood, and all injected tracer remained within the mammary gland until removed by milking. Additionally, they found that [14C]sucrose moved from the bloodstream into the milk during pregnancy but not during lactation. Higher concentrations of the isotope were observed in milk fractions derived from the alveoli than from the ducts, suggesting that most of the leaky TJs are in the alveolar epithelium. The geometry of the mammary epithelium precludes measurement of the transepithelial resistance in vivo. However, the electrical potential across the mammary epithelium can be measured by placing electrodes in the bloodstream and milk space. Leaky TJs short-circuit any potential difference that might arise as the result of different membrane potentials across the apical and basolateral membranes of the mammary epithelial cell; correspondingly, no potential could be measured across the mammary epithelium of the pregnant goat (Peaker, 1977). During lactation, Peaker (1977) measured a blood–milk potential of about 20 mV in goats; Berga (1984) found a blood–milk potential of 37 mV in the lactating mouse. The existence of this high potential difference across the mammary epithelium indicates that the TJs are highly impermeable during lactation. Because the direct passage of interstitial molecules such as sodium and chloride through the paracellular pathway into the milk space is possible during pregnancy, higher concentrations of these ions are expected to be present in the prepartum mammary secretion product. TJ closure around parturition is expected to result in a decrease in the concentrations of these ions as well as an increase in the concentration of milk components such as lactose and potassium. Such changes in milk composition have been fully documented during the onset of lactation in both goats (Linzell and Peaker, 1974) and humans (Neville et al., 1991; Neville, 1995) and can serve as an indicator of TJ status under conditions where direct tracer measurement of the permeability of the mammary epithelium is not possible (Figure 18.2).

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Sodium

Chloride

Volume

800

180

600 120 400 60 200

0

Milk Volume (ml/day)

[Lactose], [Na] or [Cl] (mM/L)

Lactose

0

0

2

4

6

8

Days Postpartum FIGURE 18.2 Changes in milk composition denoting TJ closure after parturition in women. A fall in the sodium and chloride concentration and a rise in the lactose concentration denoting TJ closure occurs 24 h prior to the milk volume increase during the onset of lactation. (Data replotted from Neville et al., 1991.)

Macromolecules labeled with fluorescent tracers can be injected into the lumen of the mouse mammary gland through a glass pipette placed in the teat canal, as described in the legend to Figure 18.3 (Nguyen et al., 2000). These molecules pass immediately into the interstitial space of the gland during pregnancy and are retained in the lumen during lactation. The permeability of the mammary TJs can also be assessed visually by overlaying the exposed mouse mammary gland with a solution containing a fluorescently labeled marker and incubating for 1 to 2 h prior to removal of the gland and sectioning for observation under the microscope. In sections from the pregnant gland, the dye can be observed to pass through the TJs; it is stopped at the TJ during lactation. This experiment shows that the permeable TJs observed during pregnancy are not the result of damage to the epithelium brought about by the pressure of intraductal injection. To determine the size of molecules to which the paracellular pathway is permeable, fluorescent tracers of increasing size, FITC-bovine serum albumin (mol wt 60 kDa), FITC-rabbit IgG (mol wt 150 kDa), and FITC-dextran (2000 kDa), were injected into the lumen of the mammary gland of pregnancy (Nguyen and Neville, 1998). Both bovine serum albumin and rabbit IgG quickly leaked across the mammary epithelium. Even such a very large macromolecule as 2000-kDa FITC-dextran was observed to pass in a limited way through the highly permeable TJs of pregnancy. Together these findings from isotope tracer, blood–milk potential, milk composition, and fluorescent tracer experiments indicate that the mammary TJs, particularly in the alveoli, are leaky during pregnancy and close around parturition to form a tight barrier that prevents paracellular movement of molecules across the mammary epithelium. The next section considers the mechanism for TJ closure during the transition from pregnancy to lactation, focusing on the hormonal regulation of this transition and the possible mechanisms involved.

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FIGURE 18.3 FITC-labeled IgG passage across the mammary epithelium. A glass pipette containing 10 µl of 3 mg/ml of FITC-labeled IgG in PBS was injected intraductally into the lumen of the fourth mammary gland of anesthetized mice, followed immediately by an intraductal injection of 25 µl of 10% paraformaldehyde to immobilize the tracer as quickly as possible. After dissection and embedding, sections of the gland were viewed by fluorescence microscopy. (A) gland from a pregnant animal; (B) gland from a lactating animal; AT, adipose tissue; F, fat droplet in milk or mammary cell; arrows point at interstitial space filled with tracer in A and empty of tracer in B.

Considerable evidence indicates that TJ permeability increases during involution and with mastitis. Much less research has been devoted to these stages, during which it is not even known whether changes in permeability result from loss of necrotic or apoptotic cells.

18.3 THE TRANSITION FROM PREGNANCY TO LACTATION The authors were able to model the transition from pregnancy to lactation by removing the ovaries in a 17-day pregnant mouse. The time course of TJ closure after this procedure was followed using repeated intraductal injections of [14C]sucrose (Figure 18.4). After about a 10-h delay the height of the sucrose peak began to decrease and by 20 h passage of the isotope into the bloodstream was barely detectable. The process was highly reproducible with the mid-point of the curve occurring at 13.6 ± 1.5 h after the surgery (Nguyen et al., 2001). This procedure was used to examine both the hormones responsible for junction closure and the mechanism by which it occurred.

18.3.1 HORMONAL REGULATION A high level of progesterone maintains pregnancy, and a drop in the level of this hormone leads to parturition and initiates lactation (Kuhn, 1969; Deis and Delouis,

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FIGURE 18.4 Use of intraductal [14C]sucrose injection to assess TJ permeability in the mouse mammary gland. The isotope (2 × 106 cpm) was injected intraductally 5, 11, 23, 17, and 21 h after ovariectomy of a midpregnant mouse. To 11 h postinjection the peak amount of [14C]sucrose in the blood was similar to that observed prior to surgery. By 13 h the peak began to decrease, until at 21 h no sucrose could be observed in the bloodstream. (Inset) The blood sucrose is plotted as a function of the time postovariectomy. The peaks are fitted with an empirical curve y = (1 – A)/(1 + exp(a(t – t1/2))), where A is the minimum value approached by the sigmoidal curve and reflects sucrose permeability after TJ closure, a expresses the steepness of the transition, and t1/2 is the time at which the sigmoidal curve falls to half the maximum value. The t1/2 is defined as the TJ closure time. (Replotted from Nguyen et al., 2001, with permission of the Society for Endocrinology.)

1983; Nishikawa et al., 1994). To determine whether progesterone withdrawal brought about by ovariectomy was responsible for the change in TJ permeability exogenous progesterone was injected 1.5 h after ovariectomy; closure was delayed. In addition, injection of the progesterone antagonist RU486 into late pregnant mice mimicked progesterone withdrawal. Both types of experiment provide strong evidence that loss of progesterone is the specific trigger for TJ closure. As expected, progesterone withdrawal also triggered parturition; however, parturition itself did not appear to be the direct cause of TJ closure, since the timing of parturition was erratic, usually occurred after closure, and showed no correlation to the relatively consistent time course of TJ closure. A large body of evidence demonstrates a relation between glucocorticoids and prolactin and the onset of lactation (Neville, 1983). Additionally, glucocorticoids have been implicated in the regulation of mammary TJ permeability in a number of studies both in vivo and in vitro. Thompson (1996) showed that local injection of cortisol into the mammary gland of late pregnant goats decreased TJ permeability.

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Stelwagen et al. (1995) found a decrease in the permeability of the mammary gland of lactating cows when the level of cortisol was increased by ACTH treatment. As described in more detail below, Firestone and colleagues (Zettl et al., 1992; Buse et al., 1995) demonstrated that glucocorticoid decreased the transepithelial resistance and paracellular flux in 31EG4 nontransformed mouse mammary epithelial cells and Con 8 rat tumor cells grown on permeable supports. This laboratory (Nguyen et al., 2001) found that low levels of glucocorticoid were essential if TJs were to close in response to progesterone withdrawal. In addition, when dexamethasone was given in conjunction with ovariectomy, the permeability of the junctions was lower 20 h after ovariectomy than observed in response to progesterone withdrawal alone. Both findings suggest that glucocorticoids are essential for TJ closure around parturition. Prolactin has also been associated with the regulation of TJs in the mammary epithelium. Linzell and colleagues (1975) found that milk composition gradually changed during late lactation in rabbits. Milk lactose and potassium decreased, and milk sodium increased, consistent with an increase in the permeability of the mammary epithelium. Prolactin treatment returned milk composition to that of established lactation. Furthermore, when the source of prolactin in the lactating rabbit was removed by hypophysectomy, milk composition gradually came to resemble that of late lactation (Linzell et al., 1975). Inhibition of prolactin secretion by bromocryptine produced a similar change in the composition of mouse milk (Flint and Gardner, 1994). The authors’ study (Nguyen et al., 2001) found that ovariectomy-induced TJ closure can proceed when prolactin secretion is inhibited by bromocryptine or when the source of placental lactogen, a hormone that also interacts with prolactin receptors, was removed by hysterectomy, and dexamethasone was given to replace fetal glucocorticoid. However, when the sources of both prolactin and placental lactogen were eliminated by both bromocryptine and hysterectomy, junction closure did not occur, suggesting that activation of prolactin receptors is necessary. The effect of prolactin on TJ permeability may be secondary to its effect on the maintenance of the mammary epithelium. In the study by Linzell et al. (1975) conducted in rabbits, prolactin had an effect on TJ permeability only during late lactation, when cell loss was occurring at a higher rate, and not during midlactation. Flint and Gardner (1994) found that the inhibition of prolactin secretion not only increased mammary epithelial permeability, but also increased cell loss, as shown by a decrease in the DNA content of rat mammary gland. In addition, Sheffield and Kotolski (1992) showed that prolactin treatment inhibited apoptosis during involution. These results suggest that prolactin might maintain the integrity of the mammary epithelium by preventing apoptosis, with secondary effects on TJ permeability. Nevertheless, a direct effect of prolactin on mammary epithelial TJ permeability cannot at present be ruled out. An unknown milk factor may also be implicated in TJ closure around parturition (Peaker et al., 1997). Peaker and colleagues observed that milk removal from one gland in late pregnancy led to changes in the composition of the secretion consistent with junction closure in that gland whereas the composition of the secretion from the unmilked gland did not change. They suggested that a substance that prevents

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Local Factors

Ovary RU486 Progesterone

_

Pregnancy OPEN

Lactation CLOSED

+

Placental Prolactin

Lactogen

Bromocryptine

Corticosterone

Fetus and Placenta

Adnenal Cortex

Anterior Pituitary

FIGURE 18.5 Model of regulation of TJ closure at parturition. Ovarian progesterone inhibits TJ closure possibly assisted by a local factor. Progesterone withdrawal can be induced by ovariectomy or the inhibitor RU486. Corticosterone and a lactogen, either placental lactogen or prolactin, are required for closure. The fetal adrenal cortex can supply sufficient glucocorticoid for closure by itself; the glucocorticoid can be replaced by dexamethasone (not shown). Either placental lactogen from the placenta or bromocryptine from the maternal pituitary is sufficient to allow closure to occur upon progesterone withdrawal. (Modified from Nguyen et al., 2001.)

junction closure is produced by the mammary gland during pregnancy. Near parturition, as hormonal changes occur, production ceases and any substance remaining in the gland is catabolized so that TJ closure occurs. Consistent with this hypothesis, when secretion product from periparturient goats was injected into the udder of lactating goats, milk composition changes consistent with opening of TJs were observed (Blatchford et al., 1985). The nature of the substance is unknown. Maule Walker and Peaker (1980) presented evidence that prostaglandin F2α could be involved, but this notion has not been further tested. A diagram depicting the current understanding of the hormonal regulation of mammary TJs at parturition is shown in Figure 18.5.

18.3.2 MECHANISM

OF

TIGHT JUNCTION CLOSURE

With the very high permeability of the mammary epithelium of pregnancy, one might question the presence of TJs in the epithelium. However, Pitelka and colleagues (1973), using freeze-fracture electron microscopy, found that the TJ network of the mammary epithelium was present in the mouse mammary gland in late pregnancy, although it was more disorganized, and had fewer interconnections, more loose ends, and fewer TJ strands between the lumen and the basolateral space than the TJ network of lactation (Figure 18.6). This finding has been confirmed in this laboratory (Nguyen and Neville, 1999). Further, it was demonstrated using immunocytochemistry that

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FIGURE 18.6 Freeze-fracture images of the TJs of the mammary epithelium from the pregnant (upper) and lactating (lower) mouse. Circled arrows indicate the orientation of the junctional strands with respect to the apical pole of the cell. Both inner (A) and outer (B) leaflet are present. LU, lumen; closed arrows point to loose ends and open arrows to gap junctions. Bars represent 0.5 µM. (From Pitelka, D. R., J. Cell Biol., 56, 797, 1973. With copyright permission of the Rockefeller University Press.)

all TJ components were present and appropriately arranged at the apical borders of the mammary epithelial cells in the mammary gland from the pregnant animal (Figure 18.7). All these findings suggest that the process of closure involves, not assembly of TJs, but a change in the interaction of these components. A previous section described the finding that the leaky TJs of pregnancy are permeable to molecules as large as 2000 kDa. The radius of a 2000-kDa FITCdextran is approximately 200 Å (Burns-Bellhorn et al., 1978; Bellhorn, 1981; Armstrong et al., 1987), whereas the width of a TJ strand is 80 Å. The large size of the opening places a limit on the mechanisms that could be responsible for the highly permeable junctions of pregnancy. Assuming that the TJ strand is formed from the polymerization of subunits with a diameter of 80 Å, then it is unlikely that the opening is a pore within a single subunit or formed by a few subunits. A large opening that would allow passage of 2000-kDa dextran could form from a break in the TJ strand, or from detachment of a strand end from an interconnection point. An opening formed by these mechanisms could potentially accommodate molecules larger than 2000 kDa. Analysis of TJ strand number showed that a molecule crossing the TJ barrier during lactation would have to cross an average of 11.3 ± 0.4 strands, whereas it would only have to cross an average of 5.5 ± 0.4 strands during pregnancy (Nguyen and Neville, unpublished). Further, during pregnancy 5% of the epithelial barrier consisted of single strands and a distributed analysis of flux density through the

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FIGURE 18.7 ZO-1 staining in the mammary gland from a pregnant (A) and a lactating mouse (B). The staining completely surrounds the apical border of each cell. Similar views can be obtained by staining for actin, ZO-1, occludin, β-catenin, α-catenin, and E-cadherin. (From Nguyen, D.-A. D. and Neville, M. C., J. Mammary Gland Biol. Neoplasia, 3, 233, 1998. Used by permission of Kluwer Academic/Plenum Press.)

single-stranded regions indicated that they could account for one half the flux through the epithelium of pregnancy. No regions containing fewer than five strands were found during lactation. This analysis provides additional evidence that the mechanism of TJ closure involves strand reorganization into a multistrand barrier. However, another possibility is that short sections of the TJ strand could lose their interaction with the corresponding strand in the membrane of the neighboring cell, forming an opening between cells. The size of such openings could be large and variable, depending on the number of subunits in such a section. There is currently no evidence that would allow evaluation of this proposition. Unlike TJs in most other epithelia, the TJs of the mammary epithelium, especially during lactation, exhibit a high degree of double-strandedness (Figure 18.8B). This characteristic may enhance the tensile strength of the junctions and allow them

FIGURE 18.8 Side-to-side interactions of TJ strands 18 h after ovariectomy (A) and in the lactating mammary epithelium (B). Arrowheads = regions of double-strandedness; arrows = regions of double-strandedness forming during TJ closure after ovariectomy.

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to withstand the stress generated as the lumen expands and contracts with milk secretion and expulsion, respectively. In addition, the side-to-side interactions that give rise to double-strandedness may also play a role in the organization of the TJ network. For example, they could help to recruit loose strands into the network or remove areas with low strand numbers from the network. Examination of the TJ network 18 h after ovariectomy revealed numerous sites where side-to-side interactions act to pinch off an area with low strand number (Figure 18.8A). This analysis of the freeze-fracture evidence suggests that a change in strand-tostrand interaction within the membrane of a single cell is the basis of the change in permeability of the mammary TJ between pregnancy and lactation. As yet, it has not been possible to identify any TJ components that are missing in one stage or another, although the authors have detected an increase in phosphorylated occludin following ovariectomy of the late pregnant mouse (Nguyen and Neville, unpublished).

18.4 IN VIVO REGULATION OF MAMMARY TIGHT JUNCTIONS BY MILK STASIS, MASTITIS, AND OXYTOCIN 18.4.1 MILK STASIS In the earliest experiments on the effects of milk stasis Fleet and Peaker (1978), using lactating goats, found that 3 days after cessation of milking sodium and chloride increased and potassium and lactose decreased in a manner consistent with a “loss of integrity” of the mammary epithelium. More recently, Stelwagen and colleagues (1994; 1997) investigated the effects of once-daily milking of goats (1994) and cows (1997) on TJ opening using the appearance of lactose in the blood as a measure of the status of the TJs. In both cases, a reduction in milk yield and an increase in blood lactose were observed 20 h after the last milking. In the experiments in cows, the lactose concentration in the plasma decreased rapidly once milking was reinitiated. These observations have led to the dual concepts that milk stasis leads to junction opening and that milk secretion and TJ status are coupled. Consistent with the second idea, infusion of EGTA up the teat of goats led both to opening of the junctional complexes between the cells and to a reduction in the rate of milk secretion (Neville and Peaker, 1981; Stelwagen et al., 1995). The relation between milk stasis and TJ opening raises the following questions: When do TJs open in response to milk stasis and what is the mechanism? A number of hypotheses have been proposed, ranging from a signal generated by cell damage (Stelwagen and Lacy-Hulbert, 1996), to simple stretch of the mammary epithelial cells (Sudlow and Burgoyne, 1997), to the presence of a feedback inhibitor in milk (Wilde et al., 1995). Cell damage when cows went unmilked for up to 24 h was ruled out by the observation that there was no increase in the concentration of N-acetyl-β-D-glucosaminidase, an indicator of damaged cells, in milk at the time when lactose appeared in the bloodstream (Stelwagen and Lacy-Hulbert, 1996). Simple stretch does not seem to be the mechanism since large changes in milk sodium and chloride were not observed until 3 days after cessation of milking in the early study by Fleet and Peaker (1978). The feedback inhibitor isolated by Wilde

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and colleagues does not alter milk composition (Peaker et al., 1997), suggesting that at least this particular chemical agent is not involved in TJ opening. Recently, Wilde and colleagues analyzed changes in the mammary epithelium associated with oncedaily milking in goats (Li et al., 1999) and found that a portion of the mammary alveoli was always undergoing involution, suggesting that changes in the permeability of the mammary epithelium may be associated with apoptosis of mammary epithelial cells. Consistent with this notion, the authors have found that TJs open about 48 h after the onset of milk stasis in the mouse (Beeman and Neville, unpublished), a time when massive apoptosis of the epithelium is initiated.

18.4.2 MASTITIS Mastitis is inflammation of the mammary gland and is often caused by pathogenic microorganisms. Linzell and Peaker (1972) described increased sodium and chloride in the milk and a decrease in the blood–milk potential from 20 mV in the normal lactating gland to 0 mV during mastitis. Both changes are consistent with decreased TJ permeability or with loss of cells from the epithelium. Other researchers (Symons and Wright, 1974; Frost et al., 1984) observed an increase in permeability, as indicated by increased sodium and plasma protein level in milk, with mastitis induced by injections of bacterial endotoxins and exotoxins into the udder. Although it is generally understood that the mammary epithelium is more permeable during mastitis than during healthy lactation, the exact role the TJ plays in the increased permeability of mastitis is not well understood. Most likely, there are several components to this permeability change. An obfuscating factor is the tendency of the immune system to increase TJ permeability during the inflammatory response. Products of the inflammatory reaction such as histamine, tumor necrosis factor (TNF), interferon-γ (IFNγ) (Leach and Firth, 2000; Madsen et al., 1997), and the newly discovered zonulin (Fasano et al., 2000) have been shown to increase permeability across the endothelial and epithelial layers. Zonulin is a recently discovered endogenous protein that is highly related to the Vibrio cholerae–derived Zonula occludens toxin (Zot). Zot has been shown to induce TJ disassembly in intestinal epithelia and a subsequent increase in intestinal permeability. Zonulin expression was raised in intestinal tissues during celiac disease, a clinical condition in which TJs are opened and permeability is increased. Both Zot and zonulin proteins were shown to bind a 45-kDa zonulin/Zot receptor homologous to 14-kDa MRP-8, a member of the S-100 family of calcium-binding proteins expressed by endothelial cells forming the blood–brain barrier (Fasano et al., 2000; Lu et al., 2000). The Gram-negative bacterial cell wall component lipopolysaccharide (LPS) has been shown to induce mastitis when injected intraductally into the udders of cows (Frost et al., 1984) or into the mammary glands of mice (Beeman and Neville, unpublished). The same toxin was shown to decrease transepithelial electrical resistance, increase mannitol flux, and cause the disappearance of the 7H6 antigen from the TJ of the IEC-6 intestinal epithelial cell line (Kimura et al., 1997). Interestingly, LPS had no effect upon the cultured intestinal cells when applied apically, unlike the situation in the mammary glands. These results imply a mammary epithelial cell surface receptor for LPS.

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The Clostridium perfringens enterotoxin (CPE) was shown to bind directly to the TJ structural proteins claudins 3 and 4. CPE was shown to be cytotoxic to cultured fibroblasts expressing claudins 3 or 4 but not to the closely related claudins 1 or 2. The COOH terminal 136 amino acids of CPE when expressed alone were shown to reduce transepithelial electrical resistance (TER) and to increase dextran flux without cytotoxicity across monolayers of Madin–Darby canine kidney (MDCK) cells expressing claudin-4 but not claudin-1. Claudin-4 localization to the TJ was shown to decrease upon CPE treatment. Freeze-fracture study of claudin-4 MDCK cells treated with the CPE COOH terminus demonstrated a decrease in mean strand number and a decrease in complexity concurrent with an increase in free end number (Katahira et al., 1997; Sonoda et al., 1999). The structural and physiological changes in the TJ of claudin-4 expressing MDCK cells brought about by exposure to the CPE terminus appear to be the inverse of changes in TJ structure in the mammary gland during the switch from pregnancy to lactation (see Figure 18.6). Thus, bacterial infection of the mammary gland may be expected to affect TJ permeability via at least three major avenues: (1) direct action upon TJ proteins as in the case of CPE, (2) interaction with an epithelial cell surface receptor and subsequent downstream modification of TJ permeability in the same cell as in the cases of V. cholera toxin and most probably LPS, and (3) through the effects of immune modulators released as higher-order responses to bacterial inflammation.

18.4.3 OXYTOCIN Oxytocin is a peptide hormone secreted by the posterior pituitary gland that causes the myoepithelial cells surrounding the mammary alveoli to contract, leading to ejection of milk. Oxytocin is generally used in the laboratory as an aid to milk removal. However, treatment with oxytocin, especially in large doses, can produce changes in milk composition consistent with an increase in TJ permeability. This effect of oxytocin has been observed in rabbits, mice, goats, and cows (Linzell et al., 1975; Peaker, 1977; Berga and Neville, 1985; Allen, 1990). Linzell et al. (1975) examined the effect of oxytocin on the composition of milk in rabbits, finding that a single dose of 100 mU of oxytocin, comparable to the amount of oxytocin released during a single nursing bout with a whole litter, was not sufficient to cause a change in milk composition. A single injection of 1 U, ten times the physiological dose, or a continuous infusion of 50 mU of oxytocin/min did increase milk sodium and chloride levels and decrease milk potassium and lactose levels. Tracer experiments using [14C]sucrose infused into the bloodstream showed that the higher doses of oxytocin increased the leakage of [14C]sucrose into milk. These results suggest that oxytocin administered at doses above the physiological range can cause mammary TJs to open. Large doses of oxytocin can decrease milk yield in addition to causing TJ leakage. Allen (1990) found that milk sodium and plasma lactose, both indicators of TJ leakage, increased in a dose-dependent manner with oxytocin treatment in dairy cows. A concomitant decrease in milk yield was observed. From recovery of lactose in the urine Allen was able to determine that the decreased milk yield was due to a decrease in the rate of milk secretion and not to leakage through TJs.

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The mechanism by which oxytocin increases TJ permeability and decreases milk secretion is not known at this time. Oxytocin receptors are found on the myoepithelial cells of the mammary epithelium, but not on the epithelial cells, which are the site of TJs, suggesting that oxytocin must act indirectly. Possibly the shape change in alveolar epithelial cells brought about by myoepithelial cell hypercontraction triggers mechanotransduction signaling (Sudlow and Burgoyne, 1997; Millar et al., 1997). It has also been proposed that TJ leakage could decrease milk secretion by increasing the Na:K ratio of milk (Allen, 1990). Because the apical plasma membrane of the mammary epithelial cell is highly permeable to sodium and potassium (Blatchford and Peaker, 1988), an increase in the intracellular Na:K ratio could follow. Such changes in intracellular ions have been shown to affect a wide range of cellular activities (Ledbetter and Lubin, 1977; Falconer et al., 1978). What is currently clear is that high doses of oxytocin alter TJ permeability and the milk secretion rate in a coordinated fashion. The mechanisms are entirely unknown.

18.5 HORMONAL REGULATION OF TIGHT JUNCTION PERMEABILITY IN IN VITRO MAMMARY SYSTEMS To this point experiments dealing with TJ permeability in the in vivo mammary gland have been discussed. This discussion is appropriate because it reveals the extent and physiological relevance of TJ regulation in the animal. However, for studies of the molecular mechanisms of TJ regulation it is usually more convenient to study in vitro systems. Some mammary culture systems that respond to hormones with changes in junctional permeability exist. This section summarizes data from such systems. The most important thing to note is that most model systems do not reproduce the change in junction permeability observed during the transition from pregnancy to lactation, a change that appears to involve, not assembly of the TJs, but reorganization of disorganized junctional strands into an orderly array that blocks flux across the epithelial monolayer.

18.5.1 GLUCOCORTICOIDS Firestone and colleagues developed in vitro culture systems for studying TJ regulation using 31EG4 nontransformed mouse mammary epithelial cells (Zettl et al., 1992) or Con8 rat mammary tumor cells (Buse et al., 1995) grown on permeable supports. They found that dexamethasone, a powerful glucocorticoid, increased the transepithelial resistance from 0 to greater than 1000 Ω·cm2 and reduced the diffusion of the tracers [14C]mannitol and [3H]inulin across the monolayer by more than tenfold after 96 h. The dexamethasone effect was dose dependent and the half maximal effective concentration, 8 nM, was comparable to the dissociation constant of the glucocorticoid receptor. Indirect immunofluorescence microscopy and Western blotting with an antibody against ZO-1, a cytoplasmic component of the TJ complex, showed that the decrease in TJ permeability in the 31EG4 cells was accompanied by a 2.3-fold increase in ZO-1 protein, but no gross structural changes in TJ structure. ZO-1 stained in a narrow, continuous band surrounding the periphery of each cell, as is typical for

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normal TJs. Cycloheximide, a protein synthesis inhibitor, and okadaic acid, a protein kinase C (PKC) inhibitor, inhibited both the change in transepithelial resistance and the increase in ZO-1 (Singer et al., 1994). These observations suggested that both assembly of TJ components like ZO-1 and phosphorylation of some TJ component(s) are important in the effect of glucocorticoids. Using Con8 cells, Firestone and colleagues (Woo et al., 1999) found that Rasdependent signals are required for TJ sealing but not for assembly of TJ components. In this landmark study, glucocorticoids stimulated the translocation of TJ components such as ZO-1 to the apical border of the cells in the monolayer and increased the transepithelial resistance to about 800 Ω·cm2. A number of inhibitors of the Rassignaling pathway, including dominant negative N17 Ras, the MEK inhibitor of MAPK signaling, PD09805, and the PI-3 kinase inhibitors Wortmannin and LY294002, decreased transepithelial resistance without affecting ZO-1 localization. When both the MAPK pathway and the PI-3 kinase pathway were inhibited, transepithelial resistance fell nearly to 0 but the TJ structure, at least at the light microscope level, was unchanged. These results firmly implicate Ras signaling in TJ sealing. Further, this set of experiments appears to represent the first time that TJ assembly and sealing have been dissociated in an in vitro mammary system. It remains to be seen whether Ras inhibition or inactivity is involved in the leaky TJ of pregnancy or whether the permeability characteristics and freeze-fracture morphology of the Ras-inhibited junction in Con8 cells resemble those of the in vivo junction. If so, the Con8 system may be an appropriate model for detailed studies of the mechanism by which the in vivo junction is regulated during the transition from pregnancy to lactation. More recently, Firestone and colleagues (Woo et al., 2000) have identified one of the signal transduction molecules that appear to be associated with the glucocorticoid response in Con8 cells. They found that dexamethasone treatment stimulated the level of the Id-1 protein, a serum-inducible helix-loop-helix transcriptional repressor. That this protein plays a critical role in the glucocorticoid effect is suggested by the observation that overexpression of Id-1 in a cell line that responds poorly to dexamethasone enhanced the TJ formation and sealing on treatment with dexamethasone. Antisense Id-1 prevented this effect. These results implicate Id-1 as a critical regulator of the glucocorticoid induction of TJs in mammary epithelial cell–cell interactions.

18.5.2 PROLACTIN In contrast to glucocorticoid, little attention has been paid to prolactin in tissue culture models of the mammary epithelium, although a number of prolactin responsive cell lines do exist. Stelwagen et al. (1999) have recently reported that prolactin stimulates TJ formation in the Comma 1D normal mouse mammary cell line and its derivative, HC11 cells. The authors have confirmed this finding using a derivative of the Comma 1D, the CIT3 line, derived on the basis of its ability to form TJs. This line forms TJs in the absence of either prolactin or glucocorticoid over a period of days. Both prolactin and glucocorticoid independently increase the rate of TJ formation, and together cause TJs to form quite rapidly. These findings suggest that prolactin has an

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effect on TJs independent of its inhibitory effects on apoptosis discussed above, but its mechanism of action is currently unknown. The existence of a prolactin-responsive mammary cell line may facilitate dissection of its mechanism of action.

18.6 A ROLE FOR TGF-? TGF-β isoforms are expressed with morphologically and developmentally specific patterns within the mammary gland. The isoforms TGF-β2 and TGF-β3 are strongly expressed in developing alveoli and their associated ducts during pregnancy, but only weakly during lactation (Silberstein et al., 1996; Daniel et al., 1996). This pattern of expression suggests that they may play a role in the differentiation of mammary epithelial cells that leads to lactation. In vitro, Firestone and colleagues (Woo et al., 1996) found that TGF-β treatment prevented TJ closure in dexamethasone-treated 31EG4 cells by a mechanism that resulted in disorganization and dispersal of ZO-1 staining. However, TGF-β had no effect on the amount of ZO-1 present in the cells or the activity of the glucocorticoid receptor, suggesting that TGF-β may act downstream of glucocorticoid effects on junctional assembly. In addition, the effect of TGF-β on the junctions was separate from its mitogenic effect on 31EG4 cells. Thus, hydroxyurea, a DNA synthesis inhibitor, inhibited mitosis, but not the increase in TJ permeability observed after TGF-β treatment. TJ permeability is only one facet of the antagonism between TGF-β and glucocorticoids in the mammary gland. In addition to closing TJs, dexamethasone suppresses cell division and, with the addition of prolactin and insulin, can induce 31EG4 and Comma 1D cells to secrete some milk components (Parry et al., 1987; Strange et al., 1991). TGF-β has the opposite effect, disrupting TJs and stimulating cell proliferation. These effects of TGF-β and glucocorticoid, the expression pattern of TGF-β, especially of TGF-β2 and TGF-β3 (Robinson et al., 1993), and the glucocorticoid requirement for TJ closure after ovariectomy in the mouse together suggest that the antagonism between glucocorticoid and TGF-β could play an important role in the stimulation of TJ closure. However, this hypothesis has not been directly tested experimentally.

18.7 PERSPECTIVE Mammary TJs are particularly interesting because of their high degree of physiological regulation. The most thoroughly studied regulatory event is the closure of TJs accompanying parturition. During pregnancy, the junctions are permeable to very large macromolecules, but they become completely impermeable to even the smallest ions during lactation. Closure around the time of parturition appears to be independent of the assembly of TJ components and to involve a reorganization of TJ strands. Uncoupling of TJ assembly from permeability is rare in most tissueculture model systems so far studied. However, the Firestone laboratory has demonstrated that inhibitors of the RAS-signaling pathway inhibit sealing but not assembly. It will be of interest to determine whether the inhibited cells show TJ disorganization

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on freeze-fracture analysis similar to that observed in the mammary gland from the pregnant mouse. It will also be interesting to determine whether inhibitors of RAS signaling open the junctions of the lactating gland. There is a large amount of literature that suggests that TJ permeability and milk secretion are coupled. Signaling from the adherens junction has been extensively studied. It will be of interest to determine whether the mammary epithelial cell, or any other cell, for that matter, in some way tests the integrity of its TJs before commencing the vectorial secretion of milk, a process that is compromised if the junctions are open.

ACKNOWLEDGMENTS Preparation of this chapter was supported in part by National Institutes of Health Grant R37-HD19547.

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Unique Aspects of the Blood–Brain Barrier Steven D. Wilt and Lawrence J. Rizzolo

CONTENTS 19.1 Introduction .................................................................................................415 19.2 Structure of Different Regions ...................................................................416 19.2.1 Endothelia ......................................................................................416 19.2.2 Epithelia .........................................................................................417 19.3 Protein Composition ...................................................................................419 19.4 Development ...............................................................................................420 19.4.1 Endothelia ......................................................................................420 19.4.1.1 Environmental Interactions during Development .........421 19.4.1.2 Culture Models of the Endothelial Blood–Brain Barrier............................................................................422 19.4.1.2.1 Identity of the factors that regulate endothelial junctions..................................424 19.4.1.2.2 Effects of the factors that regulate endothelial junctions..................................424 19.4.2 Epithelia .........................................................................................425 19.4.2.1 Choroid Plexus ..............................................................425 19.4.2.2 Development of the RPE ..............................................426 19.4.2.2.1 Culture model of chick RPE .....................431 19.4.2.2.2 Identity of the factors that regulate RPE junctions.....................................................432 19.4.2.2.3 Effects of the factors that regulate RPE junctions.....................................................432 19.5 Summary .....................................................................................................436 Acknowledgments..................................................................................................437 References..............................................................................................................437

19.1 INTRODUCTION This chapter examines how the environment regulates the permeability of tight junctions (TJs) in the central nervous system (CNS). The CNS requires a specialized environment that is created and maintained by the blood–brain barrier (BBB). The barrier is formed by endothelia and epithelia with unique properties. The endothelia 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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are much less permeable than those of the systemic circulation. A measure of permeability is the transendothelial or transepithelial electrical resistance (TER). The TER of CNS capillaries is approximately 1000 to 2000 Ω·cm2 vs. 1 to 10 Ω·cm2 for systemic capillaries. This requires a relatively impermeable seal to block diffusion across the paracellular spaces that lie between neighboring endothelial cells. This seal, the TJ, is present in all endothelia and epithelia, but its permeability varies widely among various tissues. The epithelial regions include the epithelium of the choroid plexus and the retinal pigment epithelium (RPE). Here the capillaries are fenestrated and the associated epithelial layer forms the barrier. Depending upon the species, these regions have an intermediate TER that ranges from 135 to 600 Ω·cm2. The concept of a BBB (Bradbury, 1979) arose from the early studies of Erhlich and his colleague Goldman who injected dye into the vasculature or the cerebral ventricles. Most tissues were labeled by intravenous injection except brain, testes, and placenta, suggesting they possessed a unique blood–tissue barrier (Goldman, 1909). As a control, brain could be labeled by intraventricular injection (Goldman, 1913). The concept originally concerned the transbarrier movement of injected proteins or dyes that bound serum protein. The mechanisms that blocked movement from blood into parenchyma involved TJs and reduced transcytosis. The concept grew to include the regulation of transcellular transport of all solutes by various mechanisms, including active transport, facilitated diffusion, and metabolic and catabolic pathways. Although this chapter focuses on the regulation of TJs, one must be mindful of these other mechanisms, as illustrated by experiments with the Necturus gallbladder (Kottra et al., 1993). Paracellular permeability decreased in response to an increase in intracellular cAMP, but the mechanism did not involve TJs. Instead, an increase in chloride conductance across the apical membrane lowered the intracellular concentration of chloride. In turn, this caused an influx of chloride and water across the lateral membrane that collapsed the paracellular space. This narrowing of the paracellular space decreased the paracellular permeability (Claude, 1978). The mechanisms that regulate the BBB develop progressively during embryogenesis (Dermietzel and Krause, 1991; Rizzolo, 1997; 1999; Kniesel and Wolburg, 2000). Different transporters, channels, barrier-specific antigens, and junctional proteins appear at different stages. The polarized distributions of various apical and basolateral membrane proteins change progressively. Finally, there are changes in the composition, structure, and permeability of the TJs. By this gradual development, nature has dissected the function of the TJs into component parts. Accordingly, culture models of development reveal a diversity of mechanisms that regulate TJs. This chapter first discusses the morphology of various regions of the barrier and the protein composition of the intracellular junctional complexes. Then insights gained by examining development in vivo and the culture models that have been devised to explore those insights are discussed.

19.2 STRUCTURE OF DIFFERENT REGIONS 19.2.1 ENDOTHELIA The endothelia come from mesodermal precursors that invade the CNS. When endothelial precursors invade a tissue, they recruit pericytes from the local mesenchyme,

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FIGURE 19.1 Structure of the endothelial regions. Intraendothelial cell junctional complexes are intermixed gap, adherens, and tight junctions. Endothelial-pericyte junctions are formed by gap and adhesion junctions that penetrate the basal lamina (light gray). The end feet of astrocytes envelop the complex.

which may account for tissue variability in the properties of pericytes (Hirschi and D’Amore, 1996; 1997). Pericytes form multiple contacts with endothelial cells that include gap and adherens junctions (Fujimoto, 1995; Liebner et al., 2000). In the CNS, pericytes are thought to function as phagocytic microglia (van Deurs, 1976), antigen-presenting cells (Hickey and Kimura, 1988), and regulators of capillary flow in the retina (Wallow and Burnside, 1980). Absent pericytes, the brain microvasculature forms, but is subject to edema, hemorrhage, and microaneurysms (Lindahl et al., 1997). The ratio of pericytes to endothelial cells is high in the brain and retina (estimates range from 1:1 to 1:5) as compared to muscle (1:100). In the CNS, microvessels and their associated pericytes are enveloped by the end-feet of astrocytes (Figure 19.1). Compared with systemic capillaries, CNS capillary walls are thinner, pinocytotic vesicles are fewer, and transcytosis is lower (Kandel et al., 2000). Unlike some capillaries, CNS capillary endothelia have a continuous network of TJs. In endothelial cells, the TJs are intermixed with adherens and gap junctions (Hüttner et al., 1973; Dermietzel, 1975; Schulze and Firth, 1993).

19.2.2 EPITHELIA The epithelia that contribute to the BBB derive from the neuroepithelium that forms the neural tube. A diverticulum of the tube forms the optic vesicle. The optic vesicle invaginates to form a two-layered optic cup with the apical surfaces of the two layers apposed (Figure 19.2A). The neural tube thickens as cells proliferate and differentiate into neurons and glia. However, the lumen is lined by a monolayer that retains epitheliod properties including the circumferential belt of apical junctional complexes. The lumen becomes the ventricular system and the subretinal (or interphotoreceptor) space, while the apical junctional complexes are remodeled according

FIGURE 19.2 Development of the epithelial regions. (A) The basal aspect of the neuroepithelium (thick line) faces the mesoderm, and the apical aspect with apical junctional complexes (thin line) faces a central lumen. The optic vesicle is a diverticulum of this tube that invaginates to form a two-layered optic cup with the apical surfaces apposed. The potential space between the layers is the subretinal space. The dark gray regions become simple epithelial monolayers with fenestrated capillary beds on their basal sides, whereas the white and light gray regions form neural tissue. (B) The linings of the ventricular system and subretinal space. Although adherens junctions encircle ependymal cells, the width of the lateral spaces is exaggerated to emphasize the lack of TJs. Not shown are the similar junctions that form the outer limiting membrane, which link photoreceptors and Müller cells at the apical surface of the neural retina. Dashed lines represent fenestrated capillaries. The box is enlarged in Figure 19.3. The iris and retinal capillary beds have been omitted for clarity. (Modified from Rizzolo, 1997; 1999.)

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to their position along the neurotube. The region lining the ventricles becomes the ependyma, a cuboidal cell layer that retains a circumferential band of adherens junctions, but loses the TJs (Brightman and Reese, 1969). A variation of this theme occurs in the neural retina, where the ventricular surface is lined by photoreceptors and Müller cells. Like the ependyma, these cells are joined by circumferential adherens junctions that collectively form the outer (external) limiting membrane (Bunt-Milam et al., 1985). There are exceptions to this general pattern. Tight junctions are retained by those ependymal cells (tanocytes) associated with various circumventricular organs and by the RPE (Brightman and Reese, 1969; Rizzolo, 1997). Here, adjacent capillaries are fenestrated and the BBB is formed by the epithelium (Figure 19.2B). The apical junctional complexes of the epithelia are unusual. Typically, the TJ lies just apical to a circumferential belt of adherens junctions. By contrast, the RPE and epithelium of the choroid plexus resemble endothelial cells with tight, adherens, and gap junctions intermixed (Hudspeth and Yee, 1973; Dermietzel et al., 1977; Møllgård et al., 1979). For endothelial cells, this intermixing might result from the very short lateral membranes of these squamous cells. But with the long lateral membrane of the epithelia, restriction of junctions to a narrow region suggests a functional relationship that differs from other epithelia.

19.3 PROTEIN COMPOSITION Endothelial and epithelial TJs have similar protein compositions (Mitic and Anderson, 1998; Rubin and Staddon, 1999; Tsukita et al., 1999). This section discusses members of the complex that have been studied in the BBB. Two transmembrane proteins, occludin and claudin, are specific for TJs and form the strands observed by freeze fracture. Occludin appears to be a regulatory protein that is itself regulated by phosphorylation. Claudins are an ever-growing protein family of 20 members that may form pores in the strands and thereby regulate selectivity (Tsukita and Furuse, 2000). Different claudins are specific for different tissues or regions. For example, claudin-16, also known as paracellin, regulates Mg2+ permeability in the kidney. Claudin-5 was reported to be specific for some endothelial cells, but the authors have also found it in RPE (Kojima and Rizzolo, 2000). Claudins and occludin bind a cytoplasmic complex of potential regulatory proteins. The scaffolding for this complex appears to be the zonula occludens proteins ZO-1, ZO-2, and ZO-3, which bind each other to form a network. For latter discussion, note that ZO-1 and ZO-2 also function in some adherens junctions. Each protein contains a series of globular protein-binding domains: PDZ1, PDZ2, PDZ3, SH3, and GuK. For ZO-2, these domains are highly conserved and distinct from the homologous domains of ZO-1 and ZO-3 (Collins and Rizzolo, 1998). For example, the sequence identity between chick and mammals was greater than 90%, compared with approximately 70% for any pairwise combination of the nine PDZ domains of ZO-1, ZO-2, and ZO-3. The preservation of sequence differences suggests the nine PDZ domains may have different binding affinities and specificities, and that a network of the three proteins has the capacity to assemble a diverse complex of regulatory proteins. The need for such a complex is discussed with culture models of RPE.

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TJs and zonula adherens junctions appear to function together to regulate permeability and cell growth. Their interrelationship appears even more intimate in the BBB, because the junctions are spatially intermixed. Further, the adherens junctions have unique protein compositions. Unlike most epithelia, the transmembrane protein E-cadherin is absent from the complex. Endothelia express cadherin-5 (VE-cadherin), while RPE expresses multiple cadherins (Marrs et al., 1993; Grunwald, 1996; Burke et al., 1999; Dejana et al., 1999). The cytoplasmic domains of cadherins bind phosphatases and catenins. N-cadherin and β-catenin are expressed transiently when endothelia first differentiate, but they appear to function in the initial interactions with pericytes. Unlike systemic endothelia, plakoglobin (γ-catenin) is preferred over β-catenin in interendothelial cells of the BBB (Liebner et al., 2000). In the epithelia of the CNS, B-cadherin (P-cadherin), R-cadherin, and N-cadherin are prominent, depending upon the species and stage of development (Marrs et al., 1993; Grunwald, 1996; Burke et al., 1999). Heterogeneous expression of E-cadherin in some cells of the RPE monolayer has been described, but the significance of this remains obscure (Burke et al., 1999). TJs and adherens junctions link to a circumferential band of actin filaments that may regulate junction permeability (Madara et al., 1987). The molecular details of this interaction are emerging from studies of model cell lines. The C-terminal domains of ZO-1 and ZO-2 bind actin, while the N-terminal domains bind α-catenin and the E-cadherin/catenin complex (Itoh et al., 1997; 1999; Fanning et al., 1998). Early in the formation of epithelial TJs, ZO-1 appears to be more intimately associated with the adherens junction (Rajasekaran et al., 1996). Intriguingly, overexpression of ZO-1 subdomains can result in an epithelial–mesenchymal transformation that includes stimulation of the β-catenin-signaling pathway (Reichert et al., 2000). The interrelationship between TJs and adherens junctions in the BBB is an important area for future study.

19.4 DEVELOPMENT 19.4.1 ENDOTHELIA Dermietzel and Krause (1991) described early, intermediate, and late phases of development of the BBB. In the early phase, ZO-1 and continuous strands of TJs are evident, but the junctions are leaky. During development, various barrier-related enzymes and antigens appear, and the TJs become less permeable. The details of this conversion are controversial. Although CNS endothelia appear leaky to injected horseradish peroxidase (HRP) until the intermediate phase, these data are subject to artifacts (Vinores, 1995; Saunders et al., 2000). HRP, especially type II, can cause vascular leakage mediated by histamine and serotonin. Further, the high volumes and protein concentrations used in some studies might damage the fragile vessels of early development. Less controversial is the decrease in permeability to small solutes. Butt et al. (1990) examined the TER of pial vessels of the rat. On embryonic days 17 to 20 (E17–20), endothelia blocked the diffusion of injected lanthanum and the TER was 300 Ω·cm2. At this stage a barrier to proteins was likely present, because epithelia with a TER of 150 Ω·cm2 block the diffusion of HRP (Madara, 1998). At birth (E21), the TER rose to 1100 Ω·cm2. By postnatal day 28 (PN28), the TER rose

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to 1500 Ω·cm2. Thus, a sharp decrease from intermediate to low levels of permeability occurred around birth, followed by further decreases in the postnatal period. Morphological studies of rat suggest that the decreased permeability resulted from changes in the TJs, rather than decreased fenestration, decreased transcytosis, shortened length of the paracellular path, or decreased length of the junctions along the wall of the capillary (as results from a decrease in cell density) (Claude, 1978). These alternatives are inconsistent with the observations of rat cerebrum and cerebellum (Stewart and Hayakawa, 1994). Fenestrations disappeared by E13. Further, cell density and the number of intracellular vesicles actually increased between E18 and PN1. Although the cells became thinner, the length of the paracellular path remained nearly constant. By contrast, the jump in TER around E21 correlated with structural changes in the TJs (Schulze and Firth, 1992; Stewart and Hayakawa, 1994; Kniesel et al., 1996). The most-compelling studies examined junctional strands in rat cerebral cortex by freeze fracture, and considered complexity (number of strands and branch points), density of particles, and association of particles with the E- or P-fracture face (Kniesel et al., 1996). Although particles associate with the P face of epithelia, they also associate with the E face of systemic endothelia. The grooves and ridges typical of tight junctional strands were present on E13, but they were smooth. Particles were evident on E15 and E18, and mostly associated with the E face. Between E18 and PN1, there were increases in complexity, particle density, and particle association with the P face. These structural changes are not yet correlated with changes in protein expression. ZO-1 and claudin-5 are present throughout development, but changes in their level of expression or post-translational modification were not carefully examined (Dermietzel and Krause, 1991; Morita et al., 1999). The expression of occludin does increase during development. In rat brain, it was barely detectable on PN9, but readily detectable on PN70 (Hirase et al., 1997). Further, CNS endothelia express more occludin than aortic endothelia. However, the reported increase in expression occurred after the sharp increase in TER that occurs around birth. Conceivably, the barrier develops later in the parenchyma of the brain than it does in the surface, pial vessels. Although TER was only measured in the pia, the time course for the development of vasculature morphology was similar in the pia and parenchyma (Stewart and Hayakawa, 1994). The relationship of protein expression to junction tightness will be reconsidered when culture studies are discussed below. 19.4.1.1 Environmental Interactions during Development The special properties of BBB endothelia are induced and maintained by the neural environment. The seminal finding was obtained using chick/quail chimeras (Stewart and Wiley, 1981). The transplanted quail tissues can be distinguished from the host chick tissues by morphology. Brain vessels that invaded and vascularized mesodermal tissues failed to express histochemical markers for the BBB. By contrast, these markers were induced in mesodermal vessels that invaded CNS tissues. Diffusible factors may be involved, because the pia is separated from the neuropil by a wide extracellular space and develops barrier properties simultaneously with the parenchyma (Stewart and Hayakawa, 1994).

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Astrocytes have received the most attention as the source of factors that induce or maintain the BBB. Other candidates include neurons and pericytes. These potential sources could affect endothelia directly or use each other as intermediaries. For example, noradrenaline affects astrocytic and vascular function and could be delivered by the nonsynaptic nerve terminals near astrocytes and vessels (Cohen et al., 1997). Pericytes differentiate when vessels form and precede the expression of many barrier-specific properties (e.g., loss of fenestrations, formation of continuous TJs, and expression of certain antigens) (Bauer et al., 1993; Balabanov and Dore-Duffy, 1998). These events precede the association of astrocytes with the vessels. Further, when astrocytes were destroyed by intraperitoneal injections of the gliotoxin, 6-aminonicotinamide, no effect was observed on the permeation of HRP or serum albumin (Krum and Rosenstein, 1993). Although this study did not address the permeation of small solutes, it demonstrates that junctions of intermediate strength can be maintained despite diminished contributions by astrocytes. As noted earlier, the ratio of pericytes to endothelial cells is much higher in the CNS than in other tissues, and they communicate with endothelia by adherens and gap junctions. Although these correlations advance the role of pericytes, there is direct evidence that astrocytes secrete factors that affect TJs (discussed below). In rat, astrocytes are present for the sharp decrease in permeability that occurs between E18 and PN1. These descriptive studies indicate a progression of events that gradually decrease the permeability of CNS endothelia. It is controversial whether astrocytes can induce barrier properties de novo, or whether they require a cell that is primed to be receptive to astrocytic signals. It is clearer that astrocytes produce a subset of the factors that regulate endothelial TJs. The data supporting this conclusion are presented in the next section. Later sections demonstrate that these factors differ from those that regulate the epithelial regions of the BBB. 19.4.1.2 Culture Models of the Endothelial Blood–Brain Barrier The transplantation studies described above were refined by using in situ cultures. Secondary cultures of astrocytes from cerebral cortex were implanted into the anterior chamber of the eye or onto the chorioallantoic membrane, where the cell aggregates were invaded by the leaky vessels of the iris or chorioallantoic membrane (Janzer and Raff, 1987; Holash et al., 1993; Tout et al., 1993). The most-convincing study was performed by Tout et al. (1993). Electron microscopy demonstrated that intravenous HRP decorated the luminal surface of the invading vessels, but not the lateral and ablumenal spaces. Thus, astrocytes induced a level of barrier function de novo, but, as noted earlier, HRP diffusion is not a stringent test. An intriguing difference between the deep and superficial vascular beds of the retina is that astrocytes are absent from the deep bed. Tout et al. (1993) tested the hypothesis that Müller cells replace the function of astrocytes. The observation that Müller cells also induced barrier function indicates that several glial cell types can supply the needed factors. The hypothesis that glial-derived factors modulate TJs was tested using in vitro cultures (reviewed in Reinhardt and Gloor, 1997; Rubin and Staddon, 1999). Cultures

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of brain-derived endothelial cells have a low TER of 10 to 120 Ω·cm2. The TJs form continuous strands of variable number and low complexity (Krause et al., 1991; Wolburg et al., 1994). The TER was doubled by culturing the cells with astrocyteconditioned medium or with astrocytes that did not contact the endothelial cells. A greater effect was obtained by increasing intracellular cAMP. The combination of factors was synergistic and increased the TER as high as 600 Ω·cm2 (Rubin et al., 1991; Raub, 1996). This still falls short of the high resistance observed in vivo. Commonly, the C6 glial tumor cell line was the source of conditioned medium, but the vasculature of tumors is relatively leaky. Nonetheless, similar results were obtained with purified astrocytes. Better results were obtained when endothelia and astrocytes were cultured on opposite sides of the same filter (Dehouck et al., 1990; Isobe et al., 1996; Hayashi et al., 1997). In this model, the expression of BBBspecific proteins was induced even in non-brain-derived endothelial cells. There are several interpretations. Astrocyte processes grew through the filter to contact the endothelial cells. The effect may be mediated by either cell–cell contact or by a dilute or labile paracrine effector. Brain-derived endothelial may not require close proximity to astrocytes for several reasons. They may be primed to respond and require lower concentrations of the factor, or they may no longer require one factor to respond to additional factors secreted by astrocytes. This issue was partially addressed in studies of an umbilical cord–derived cell line, ECV 304 (Kuchler-Bopp et al., 1999). Media conditioned by primary astrocyte cultures increased the TER from 19 to 150 Ω·cm2 and decreased the permeation of sucrose. Only fresh conditioned medium was effective, which implies a critical component was very labile. Part of the decreased permeability with glial cell contact may be an artifact. Permeability could be decreased by astrocyte processes that plug the filter and astrocyte cell bodies that coat one filter side. Some controls were reported, such as the barrier formed by astrocytes alone or substitution by COS cells. However, it was not determined whether coculture induced the formation of processes or increased the density of the astrocyte coating the filters. Further, various laboratories have used different assays to assess permeability, which makes direct comparisons more difficult. Other environmental factors may be important. Shear-stress from flow through the capillaries affects the endothelia. Endothelial cells were cultured along the lumen of a hollow fiber culture apparatus (Stanness et al., 1997). The combination of flow through the cultures and astrocytes cultured on the outside of the porous fibers led to very high TER. Unfortunately, it is difficult to compare directly TER measured by different culture models, because the geometry of the electrodes affects the result. Nonetheless, there was a tenfold difference between coculture and the sum of the TER from individual cultures. Again, the effect of chemoattraction on processes invading the filter was not addressed. Another factor is the dual role of astrocytes during vascular development and maintenance. In contrast to promoting vascular stability and low permeability in normoxia, retinal astrocytes secrete vascular endothelial growth factor (VEGF) to promote angiogenesis and vascular permeability in hypoxia (Stone and Maslin, 1997; Zhang et al., 1999). This emphasizes the role of the in vivo or culture environment on the behavior of astrocytes and cells that collaborate with astrocytes to regulate the behavior of endothelial cells.

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Despite the inability to recreate a BBB in vitro, the partial successes lend insight into how to convert a high-permeability junction to a low-permeability form. Because the TER is much lower than the transcellular resistance, these changes in TER are most likely due to effects on TJs. The TER is insensitive to effects on membrane pumps and channels until it approaches the transmembrane resistance, which is greater than 1000 Ω·cm2 (see Chapter 4 by Reuss). Clearly, astrocytes are a player, but what elevates cAMP or decreases permeability before astrocytes appear? Studies have addressed two questions: what is the nature of the astrocyte-derived factor and what are its effects, and that of cAMP, on TJ proteins. 19.4.1.2.1 Identity of the factors that regulate endothelial junctions Little progress has been made in identifying the active factor(s) in astrocyte-conditioned medium owing to limited quantities and the lability of a critical component. Using the potential-candidate approach, Igarashi et al. (1999) examined glial-derived neurotrophic factor (GDNF). Only CNS capillaries expressed the receptor for GDNF. Alone, GDNF had no effect on endothelia isolated from porcine brain, but it enhanced the effects of elevated cAMP. This effect happened within 8 h of exposure compared with 24 h for astrocyte-conditioned medium. Therefore, at least three factors affect endothelial TJs: GDNF, a distinct factor secreted by astrocytes, and a factor that stimulates an increase of intracellular cAMP. 19.4.1.2.2 Effects of the factors that regulate endothelial junctions The effects of astrocyte-conditioned medium are slow and may require changes in gene expression. By contrast, the effects of cAMP are rapid and reversible, which implies the relevant signaling pathway is already present. The two treatments had different effects on the morphology of TJs. Elevated cAMP increased the complexity of junctional strands, whereas astrocyte-conditioned medium and cAMP increased the P-face association of junctional particles (Wolburg et al., 1994; Adamson et al., 1998). The morphological changes were unaccompanied by obvious changes in the expression or distribution of TJ and adherens junction proteins (Rubin et al., 1991; Hirase et al., 1997). The proteins examined included ZO-1α+, ZO-1α–, occludin, β-catenin, VE-cadherin, and PECAM, but not claudin. The lack of a correlation between expression level and junction tightness was manifest a second way. The in vivo studies discussed earlier suggested that high levels of occludin were responsible for the different permeabilities of CNS and systemic endothelia. Despite the high levels of occludin expression in culture, the TER of untreated cells was low (Hirase et al., 1997). A similar conclusion can be drawn for ZO-1. The two splice variants of ZO-1 differ by an 80 amino acid “alpha” sequence (Willott et al., 1992). All endothelia express ZO-1α–, but cultured brain endothelia also express ZO-1α+ (Balda and Anderson, 1993; Hirase et al., 1997). Despite the presence of ZO-1α+ in brain endothelia, its permeability was similar to aortic endothelia that expressed only ZO-1α–. These data suggest that high levels of occludin or ZO-1 expression are insufficient and that astrocyte-conditioned medium and cAMP regulate the function of preexisting pools of junctional proteins.

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Conceivably, cAMP and astrocyte-conditioned medium might modulate protein phosphorylation. Although this has not been examined directly, the affect of phosphorylation has been examined by using growth factors and phosphatase inhibitors. VEGF and hepatic growth factor increased the permeability of endothelia, and demonstrated increased phosphorylation of occludin (Antonetti et al., 1999; Jiang et al., 1999). This result contradicts studies of epithelia where increased phosphorylation correlated with a decrease in permeability (Sakakibara et al., 1997; Farshori and Kachar, 1999). The problem is that growth factors have many effects. Besides occludin, VEGF phosphorylated ZO-1 on tyrosine. Further, it is unknown whether phosphorylation is an early or late event in lowering the permeability of endothelial junctions. Finally, VEGF increases permeability by promoting transcytosis, and the formation of fenestra and vesiculovacuolar organelles, which further complicates interpretation of the data (Roberts and Palade, 1995; Esser et al., 1998; Feng et al., 1999a, b). In epithelial and endothelial cells, phosphatase inhibitors increased tyrosine phosphorylation and disrupted TJs (Staddon et al., 1995). Besides ZO-1 and ZO-2, it may be more significant that the cadherins and catenins of adherens junctions were also phosphorylated (Rubin and Staddon, 1999). These shotgun approaches are too crude to dissect the various pathways that might modulate the function of TJs. In summary, neural tissue induces the BBB in endothelia and is required for its maintenance. In the early stage of development, it is unclear how pericytes, glia, and neurons contribute to the formation of endothelial TJs. These early junctions form a barrier to proteins and a moderately low permeability barrier to small solutes. Regardless of whether or not endothelia are primed by these early events, they can respond to astrocyte-conditioned medium or agents that elevate cAMP to decrease their permeability. In vivo, these latter interactions decrease permeability from the intermediate levels induced by early development to the low levels of the adult. GDNF together with undefined, labile factors from astrocytes participate in this late phase. The late phase includes an increase in the complexity of junctional strands and in the degree of P-face association of junctional particles. Many of these effects occur without overt changes in the expression or distribution of TJ or adherens junction proteins.

19.4.2 EPITHELIA The epithelial regions are easier to study, because many culture models exhibit TERs close to those in vivo (135 to 425 Ω·cm2, depending upon the species) (Wright, 1972; Gallemore et al., 1997). Although used to study epithelial polarity and transepithelial transport, most models have not been used to study TJs directly. It is unclear how the specialized culture media that are employed replace in vivo environmental interactions. A promising model that addresses these issues uses RPE from chicken embryos. The exciting observation is that the properties of the TJs reflect the embryonic age of the RPE at the time it was isolated, and the properties can be modified by factors produced by neighboring tissues. 19.4.2.1 Choroid Plexus Although the choroid plexus and the production of cerebral spinal fluid have been studied extensively in vivo (Dziegielewska et al., 2000; Saunders et al., 2000), there

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are few culture studies of the epithelium. The in vivo studies are complicated by the many factors that affect the composition of cerebral spinal fluid. In addition to permeability and transport properties of the epithelium, composition depends on the mass of the CNS, and the rate of fluid secretion, and flow through the ventricular system. Each parameter changes during development. TJs that block the diffusion of proteins are evident at early stages, but the permeability to small molecules decreases during development (Dermietzel et al., 1977; Møllgård et al., 1979; Wakai and Hirokawa, 1981). Nonetheless, the fine structure revealed by freeze fracture does not change during development (Møllgård et al., 1979). The in vitro tissue and primary culture models of the epithelium have been used to investigate transport mechanisms, but not the function of TJs. The cultures are polarized and form TJs by immunocytochemistry, freeze fracture, and transmission electron microscopy (Dermietzel et al., 1977; Marrs et al., 1993; Gath et al., 1997; Villalobos et al., 1997). The TER of porcine cultures ranged from 90 to 170 Ω·cm2, which is about half the 250 Ω·cm2 determined for bullfrog choroid plexus in vivo (Wright, 1972; Gath et al., 1997). 19.4.2.2 Development of the RPE The development of the RPE is coordinated with the development of the choroid on its basal side and the neural retina on its apical side (Figures 19.2 and 19.3). Coordination among these tissues was first documented for the neural retina and choroid. For all vertebrates studied, regardless of the length of gestation, or when birth occurs relative to ocular development, the developmental landmarks of photoreceptors correlate with those of Bruch’s membrane (Figure 19.4). Bruch’s mem-

FIGURE 19.3 Structure of the retinal pigment epithelium. At the apical pole, microvilli (MV) extend into the interphotoreceptor space and interdigitate with the outer segments of photoreceptors (OS). At the basal pole, numerous infoldings appose Bruch’s membrane, and the underlying choriocapillaris is fenestrated. The apical junctional complex forms a band that completely encircles each cell in the epithelium. The complexes are unusual in RPE and epithelium of the choroid plexus, because TJs, adherens junctions (AJ), and gap junctions intermix. (Modified from Rizzolo, 1997.)

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brane is the five layers of extracellular matrix that separates the RPE from the fenestrated capillaries (Greiner and Weidman, 1991). Further, the appearance of photoreceptor outer segments marks the formation of fenestrae in choroidal vessels. The maturation of photoreceptors also marks developmental events of the RPE. Three developmental phases can be defined that roughly correspond to the three phases described for the endothelial BBB (Dermietzel and Krause, 1991; Rizzolo, 1997). The boundary between the early and intermediate phases is when the inner segments of photoreceptors protrude through the outer limiting membrane (E8 in chick; E18 in rat). The boundary between the intermediate and late phases is when outer segments begin to form (E15 in chick; PN9 in rat). The development of cell morphology and polarity were recently reviewed (Rizzolo, 1997; 1999). The following discussion focuses on the apical junctional complex. In chick, the optic cup forms by E3 and the neuroepithelium begins its differentiation into neural retina and RPE. Adherens and gap junctions were present in RPE, but the adherens junction displayed morphological changes throughout embryogenesis (Fujisawa et al., 1976; Sandig and Kalnins, 1990). Membrane “kisses” were evident by transmission electron microscopy, but not freeze fracture. This indicates that rudimentary TJs may be present. Tracer studies did not address RPE directly, but suggested that a barrier to protein diffusion exists as early as E6 (Latker and Beebe, 1984). That study examined tissue after a 5-min exposure to HRP. To avoid the artifacts noted earlier with respect to HRP, the RPE was studied more rigorously in organ culture (Williams and Rizzolo, 1997). During a 45-min incubation, HRP penetrated the TJs of RPE until E12. This indicates a change in junction permeability occurred during the intermediate phase. Detailed freeze-fracture studies examined the period from E10 to hatching (Kniesel and Wolburg, 1993). Unlike endothelia, junctional particles principally associated with the P face. Between E10 and E15, strand number or complexity remained constant, but in the late phase of development these parameters increased. These changes in morphology and function correlate with changes in protein expression. On E3, the junctional complexes of the neuroepithelium contain ZO-1, N-cadherin, and occludin (Grunwald, 1996; Williams and Rizzolo, 1997). In chick, two putative isoforms of ZO-1 were detected whose expression was temporally and spatially regulated. They resembled mammalian ZO-1α+ and ZO-1α–, except that one of the chick isoforms failed to cross-react with the monoclonal antibody R40.76. Accordingly, the lower-molecular-mass isoform was termed the ZO-1-like protein or ZO-1LP (Williams and Rizzolo, 1997). The epitope for R40.76 lies near the α-region and a slight variation in the splice site might explain this anomaly. The data that suggest ZO-1LP is a ZO-1 isoform are that it cross-reacts with antibodies to ZO-1 peptides, localizes to the apical junctional complex, and binds to ZO-2 (Williams and Rizzolo, 1997; Collins and Rizzolo, 1998). Additionally, ZO-1LP binds to the cytoplasmic domain of occludin (Figure 19.5). Only ZO-1LP was found in the neuroepithelium and its expression continued in the neural retina as part of the outer limiting membrane. The outer limiting membrane is the network of zonula adherens junctions that bind photoreceptors and Müller cells and that lacks TJs. ZO-1 has been found in the outer limiting membrane of other species and in the adherens junctions of ependymal cells that lack TJs (Dermietzel and Krause, 1991;

Morphology 2,3

RPE monolayer forms

Early

Layer 1: RPE basal lamina is evident

Rudimentary Junctions

Na.K-ATPase becomes apically polarized in central region

Microvilli begin to elongate

Late Early

Late

Microvilli further elongate

Complexity of Morphological tight junctions maturation of adherens increases junctions is complete

Basal pool forms for: Spectrin, Ankyrin 5A11. REMP

Basal infoldings form

Junctions become impermeable to HRP

6 1 integrin becomes basally polarized

Outer segments begin to form

Layers 2-4: Elastin layer subdivides collagenous layer to complete Bruch's membrane Choriocapillaris is fenestrated

Intermediate

Layer 5: Choriocapillaris basal lamina is evident

Inner segments of photoreceptors protrude from outer limiting membrane

FIGURE 19.4 Time line for RPE development. Changes in the environment are indicated above the time line; changes in RPE are below the line. References: 1. Greiner and Weidman, 1991; 2. Braekevelt and Hollenberg, 1970; 3. Rizzolo, 1997; 4. Philp et al., 1995; 5. Huotari et al., 1995; 6. Fujisawa et al., 1976; 7. Williams and Rizzolo, 1997; 8. Kniesel and Wolburg, 1993; 9. Sandig and Kalnins, 1990; 10. Grunwald, 1996; 11. Liu et al., 1997.

Junctions 6,7,8,9

Microvilli Polarity/Expression 3,4,5 Apical -tubulin & centrosome Basal 1 3 integrin

RPE

Bruch's Membrane (5 layered structure)1,2,3

Photoreceptors

ENVIRONMENT

428 Tight Junctions

ZO-1LP, ZO-2

R

B

Relative Occludin Levels

FIGURE 19.4 (continued.)

TJ Proteins7

Relative Cadherins10,11 Levels

N

ZO-1

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FIGURE 19.5 ZO-1LP binds to the cytoplasmic domain of occludin. A chimera of glutathione-S-transferase (GST) and the cytoplasmic domain of occludin was bound to a sepharose-glutathione column. Cell lysates were passed over the column and the bound fraction resolved by sodium dodecylsulfate-polyacrylamide gel electrophoresis. Immunoblots were probed with antibodies that bind ZO-1 and ZO-1LP.

FIGURE 19.6 Steady-state level of ZO-2 decreases with development. Equal amounts of protein isolated from the indicated age and immunoblotted with antisera to ZO-2. Actin was blotted to verify equal protein loads.

Saitou et al., 1997). In the RPE, the level of total ZO-1 decreased during the intermediate phase of development and ZO-1LP was replaced by ZO-1 (which corresponds to ZO-1α+, see Figure 19.4). A parallel decrease in the expression of ZO-2 was also observed (Figure 19.6). These decreases will be revisited when remodeling of the adherens junction is discussed.

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Occludin was observed in the RPE and outer limiting membrane in the early phase of development, even when TJs were undetected by freeze fracture (Fujisawa et al., 1976; Williams and Rizzolo, 1997). During the intermediate phase, occludin fell below detectable limits in the outer limiting membrane (Williams and Rizzolo, 1997). The level of expression was constant in the RPE through the intermediate phase, but occludin shifted into a Triton X-100-insoluble pool. A corresponding shift in mobility on gel electrophoresis suggests phosphorylation, but this remains to be investigated directly. In contrast to occludin, preliminary data indicate that the expression of claudin-5 increases during the intermediate phase (Kojima and Rizzolo, 2000). Changes in protein expression also correlate with the morphologic maturation of adherens junctions (see Figure 19.4) (Grunwald, 1996; Liu et al., 1997). N-cadherin is widely expressed in the developing retina, but becomes confined to the RPE and outer limiting membrane. In the RPE, the level of expression is initially high, but it decreases throughout development. In the early phase, the expression of R-cadherin increases, but it decreases in the late phase. Only B-cadherin is relatively invariant. In several ways, the expression of ZO-1LP and ZO-2 mirrors N-cadherin. Both remain in the outer limiting membrane, and they gradually decrease with developmental age in the RPE. ZO-1 gradually increases until it exceeds ZO-1LP; then both isoforms decrease to basal levels. Because ZO-1 is known to function in adherens junctions of various types, it may be that ZO-1LP, ZO-2, and N-cadherin all function in adherens junctions early in development and later in the outer limiting membrane. In the RPE, these proteins are replaced as adherens junctions remodel. The basal levels of ZO-1 and ZO-2 would function mainly in TJs, as in most epithelia. 19.4.2.2.1 Culture model of chick RPE The most intriguing aspect of these observations is that during the intermediate phase the complexity of TJs does not change; yet there are changes in protein composition and permeability. The authors devised a culture model to explore these observations. Chick RPE was isolated from E7, E10, and E14, and cultured on semipermeable filters. Provided the filters were coated with laminin, the cells retained many properties exhibited by the RPE in vivo (Rizzolo, 1991; Ban and Rizzolo, 1997). The permeability of the cultures decreased with increased embryonic age. Permeability decreased further if cultures were incubated with medium conditioned by E14 neural retinas in organ culture. Notably, retinal-conditioned medium also compensated for the decreased diffusion of glucose through the paracellular pathway by increasing transport through the glucose transporter (Ban and Rizzolo, 2000b). Unlike endothelia and regardless of the presence of retinal-conditioned medium, permeability was insensitive to increased intracellular concentrations of cAMP (Rizzolo and Li, 1993; Ban et al., 2000). Like astrocyte-conditioned medium, retinal-conditioned medium required days to exert an effect and could involve changes in gene expression. Notably, RPE is only competent to respond to conditioned medium if the medium is presented in the first week of culture (Rizzolo and Li, 1993). Older cultures or passaged cells fail to respond. Immature RPE from many species fails to differentiate further in culture, but instead partially dedifferentiates (Zhao et al., 1997). Retinal-conditioned medium appears to maintain some of the differentiated phenotype.

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19.4.2.2.2 Identity of the factors that regulate RPE junctions The active factors in E14 retinal-conditioned medium were partially identified, but their source remains unresolved (Ban et al., 2000). The chick retina is avascular and lacks astrocytes. Müller cells are present on E14, but their differentiation is incomplete until E15. There was evidence for at least two active factors. Although conditioned medium doubles the TER, it does so by different mechanisms for E7 and E14 cultures. Like astrocyte conditioned medium, the factors that affect E7 cultures are labile. They are less than 10 kDa, insensitive to proteases, heat labile, and cannot be stored for more than 2 months at –80°C. Paracrine effectors of the retina include serotonin, dopamine, and adrenergic compounds, but antagonists of these had little effect. Heat-inactivated conditioned medium continued to exert a full effect on E14 cultures. The active factor in this case was trypsin sensitive. Gel filtration of conditioned medium revealed radiochemical amounts of a peptide with a mass of 49 kDa that the authors named RCM49. Although it may play a role, GDNF differs from the factors identified in retinal-conditioned medium. These results demonstrate fundamental differences between the development and regulation of the endothelial and epithelial TJs. Decreases in the permeability of endothelial junctions involve an increase in strand complexity and an increase in the association of particles with the P face. These properties are regulated by a combination of cAMP and astrocyte-conditioned medium. By contrast, these properties do not change in the epithelium of the choroid plexus or in intermediate-phase RPE. Although the active factors of astrocyte-conditioned medium have not been characterized, one factor may be GDNF. Different factors are active in retinalconditioned medium. These act sequentially at different phases of development. Unlike endothelia, there is no effect of cAMP or noradrenaline. 19.4.2.2.3 Effects of the factors that regulate RPE junctions The characterization of retinal-conditioned medium revealed that the function of TJs is multifaceted and that different properties of the junction can be regulated independently. Retinal-conditioned medium can be used as a reagent to manipulate different mechansims, because its specificity depends upon the embryonic age of the RPE used to establish the cultures. Other aspects of junctional function can be studied by comparing cultures derived from different ages. The TER is a gross measure that averages these functions with respect to the permeation of small ions, but obscures other functions. This was first apparent in studies of epithelial cells in which occludin was overexpressed (Balda et al., 1996; McCarthy et al., 1996). Although the TER increased, the permeability of mannitol also increased. This suggests the permeability of ions and small nonionic solutes might be regulated differently. A similar finding was obtained in RPE. The permeation of tracers of different size, monosaccharides of different charge, and different cations were measured; and differences with embryonic age and in response to retinal-conditioned medium were observed. The TER of E7 RPE was only 20 Ω·cm2 (Figure 19.7). There was a decrease in permeability between the E7 and E10 cultures that affected HRP (hydrodynamic radius of 30 Å), and not inulin (10 to 14 Å) or mannitol (4 Å). Despite the lack of an effect on mannitol, there was a small but significant increase in the TER. A

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FIGURE 19.7 Permeability of RPE cultures decreases with embryonic age and exposure to E14 retinal-conditioned medium. Chick RPE was isolated from embryos of the indicated age and cultured in serum-free growth medium (SF2) or conditioned medium (CM). The permeability (µl/cm2/min) of tracers added to the apical medium chamber and TER of the same cultures (Ω·cm2) were measured after 9 days in culture. * = p < 0.001; # = p < 0.05. (From Ban, Y. and Rizzolo, L. J., Mol. Vis., 3:18, 1997. With permission.)

different result was observed between E10 and E14. The permeation of HRP was unaffected, but there was a decrease in the permeation of inulin and a modest decrease in the permeation of mannitol. By contrast, the increase in TER was greater than might be predicted from the small decrease in mannitol permeation. This is notable, because in previous studies of leaky epithelia, mannitol was more sensitive to changes in TJs than TER (Madara, 1998). Retinal-conditioned medium decreased the permeability of each culture, but again the proportion of the effect depended upon the assay and the age of the cells used to establish the culture. With conditioned medium, each culture exhibited the same permeability to HRP. Similarly, there was no difference among cultures in the permeability to inulin or mannitol. By contrast, the age-related differences were maintained with respect to TER. Note that conditioned medium was able to increase the TER of E7 cultures to the basal level of E14 cultures. Since all activity was recovered in a <10-kDa fraction of conditioned medium, E7 cultures are incompetent

434

Tight Junctions

TABLE 19.1 The Ratio of Monosaccharide/Mannitol Permeability of Cultured RPE 3-O-Methylglucose Glucosamine N-Acetylneuraminic acid

SF2 CM SF2 CM SF2 CM

1.03 1.01 1.04 1.07 0.74 0.69

E7 ± 0.04 ± 0.03 ± 0.02 ± 0.04 ± 0.01 ± 0.01

E10 — — — — 0.72 ± 0.01 0.67 ± 0.01

E14 1.03 ± 0.03 1.06 ± 0.08 1.02 ± 0.02 1.00 ± 0.07 0.65 ± 0.02 0.65 ± 0.02

Note: Errors represent the SE of six experiments. For N-acetylneuraminic acid, significant differences were found between SF2 and CM in E7 and E10 cultures, and between E10 and E14 cultures in SF2 (p < 0.05). Source: From Ban, Y. and Rizzolo, L. J., Am. J. Physiol., 279:C744–750, 2000. With permission.

to respond to RCM49 (Rizzolo and Li, 1993). The <10-kDa fraction was not needed to maintain the basal TER of E14 cultures, but they could respond to retinalconditioned medium to raise the TER (Ban et al., 2000). The final TER was still lower than that of adult chick RPE (138 Ω·cm2), but the TER should rise further in the late stage when the complexity of the junctional strands increases (Kniesel and Wolburg, 1993; Gallemore et al., 1997). Control experiments indicated these were effects on TJs and not on transcytosis. Mannitol is larger than ions. To better assess the effect of charge, Ban and Rizzolo (2000a) measured the permeation of similar-sized, charged and uncharged monosaccharides (hydrodynamic radius 4 to 5 Å). In double-label experiments, the permeation of these tracers was measured relative to mannitol (Table 19.1). In each culture, regardless of the presence of conditioned medium, the ratio of 3-O-methylglucose or glucosamine to mannitol was 1.0. Only N-acetylneuraminic acid was selectively retarded. The ratio of N-acetylneuraminic acid to mannitol varied among cultures. The ratio decreased with age, but decreased with conditioned medium only in E7 and E10 cultures. Although conditioned medium reduced the permeability of E14 cultures, it had no effect on this ratio. Low-calcium medium could reduce the TER to <10 Ω·cm2 without affecting the close apposition of the lateral membranes. This increased the ratio to 0.76, which supports the interpretation that the difference in ratio is an effect of TJs. A common model for TJs is a network of anastomosing strands that contain pores whose open–closed state can be regulated. The model presented in Figure 19.8 was developed, because models of uniform pores were unable to explain the data from the RPE cultures. The following discussion was modified from Ban and Rizzolo (2000a). Pores are illustrated that vary in size and charge. These may be viewed as the total number of pores, or the number of pores in their open state. Because this model is concerned with the intermediate stage of development, the number of strands and their complexity are held constant.

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FIGURE 19.8 Conceptual model for regulating TJs. Three nonexclusive mechanisms indicate how junctional pores might be regulated. Anastomosing junctional strands are indicated as black lines. Pores in their open state are indicated as boxes of different size or charge. See text for details. (From Ban, Y. and Rizzolo, L. J., Am. J. Physiol., 279:C744–750, 2000. With permission.)

Mechanism A illustrates a shift in size distribution. The loss of large pores is compensated for by an increase in small pores. The total cross-sectional area of pores remains unchanged. With this mechanism, larger solutes would be excluded, but the permeation of smaller solutes would be unaffected. Mechanism A could account for the differences between E7 and E10 RPE that were cultured in SF2. Permeation might be affected by charge on the walls of the pores, if the predominant pores were close in size to the solute. For example, between E10 and E14 there is decrease in the permeation of inulin and N-acetylneuraminic acid, but there is less of an effect on mannitol. Mechanism B illustrates a decrease in the number of pores (or time in the open state) without affecting the size distribution. This would cause a decrease in the permeability of all solutes, without an effect on charge selectivity. This mechanism could explain the effects of retinal-conditioned medium on the E14 cultures. A variation on this mechanism is to decrease preferentially the number of large pores. In this scenario, the total cross-sectional area decreases, and the effects on charge discrimination by smaller pores becomes more evident. This might explain the effect of retinal-conditioned medium on the E7 and E10 cultures. Conditioned medium reduced the permeability of all solutes, but N-acetylneuraminic acid was affected more than mannitol. Pores smaller than 4 to 5 Å, but large enough for ions, would be invisible to the solutes mentioned above. A decrease in the number of such pores would explain an increase in TER that was out of proportion with a decrease in mannitol permeability. This was observed among cultures of different age that were maintained in either SF2 or retinal-conditioned medium. According to studies on other epithelia, the relationship between mannitol permeation and TER is nonlinear (Madara, 1998).

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Tight Junctions

When the TER is below 200 Ω·cm2, a decrease in junction permeability affects mannitol more than ions. In contrast, the percent increase in TER that was observed was always greater than, or equal to, the percent decrease in permeability to mannitol. Mechanism C illustrates a change in the charge density of the pores. This would be a nonexclusive, alternative explanation for the decreased N-acetylneuraminic acidto-mannitol ratio. However, it would not explain the concomitant changes in mannitol permeation or size discrimination. The authors looked for other evidence of this mechanism by determining ion selectivity. Among the various epithelia, the most selective is the gallbladder. Depending upon the species, ion selectivity conforms to the Eisenman sequence II or IV (Moreno and Diamond, 1975; Powell, 1981; Eisenman and Horn, 1983). These sequences correspond to channels of low or moderate “field strength.” The ion selectivity of RPE junctions always conformed to sequence I, which corresponds to channels of lowest field strength. Either the selectivity of the pores is low or the selectivity of small pores is masked by the flux through large, nonselective pores. The latter explanation, combined with variation in the distribution of pore size, could account for the variation in ion selectivity among various epithelia. A variation of mechanism C would be a change in the charge of the lateral membranes. For example, charge density would increase if the content of sialic acid increased in the membrane glycoproteins near the apical junctional complex. The diversity of tight junctional proteins might allow cells to regulate these various mechanisms independently. By itself, the level of ZO-1 expression is uninformative. Besides ZO-1LP, ZO-1 expression is turned on in E7 cultures without increasing TER. The expression of ZO-1 was unaffected by retinal-conditioned medium. Finally, as in vivo, the expression of ZO-1 was lower in E14 cultures than in E7 cultures (Ban and Rizzolo, 1997). The more important consideration may be the ligands of ZO-1, ZO-2, and ZO-3. As discussed earlier in Section 19.3, a complex of these three proteins potentially assembles many structural and regulatory proteins. By incorporating different mixtures of claudins and occludin and other regulatory proteins, pores of varying size and charge could be assembled. A flexible, regulatable scaffold could contribute to diverse pore structures. The authors are testing this hypothesis by overexpressing different protein-binding domains that will compete for scarce regulatory factors. Preliminary studies demonstrated dominant negative effects that depended upon embryonic age and the presence of retinal-conditioned medium (Wilt and Rizzolo, 2000).

19.5 SUMMARY The development of the BBB affords the opportunity to observe the natural conversion of a leaky junction to a tight phenotype. It highlights the differences between endothelial and epithelial junctions. In endothelial cells, changes in permeability correlated with changes in the fine structure of junctional strands. These included an increase in complexity (number of strands and number of anastomotic branch points) and changes in the distribution of junctional particles. These properties were influenced by glial factors and cAMP. In epithelial cells, changes in permeability were observed when the complexity of strands did not change. These changes

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correlated with changes in protein expression. Culture models have been devised to study these properties. Although endothelial cultures have been informative, an in vivo-like culture model has been difficult to develop. The TER tends to be low and modulators have modest effects. Further, the lability of the relevant glial factors has slowed progress. By contrast, epithelial cultures show great promise. They emphasize the differences with endothelial cultures, because they respond to different modulators of junction permeability. It is easy to isolate primary cultures of RPE, and the permeability of the cultures depends upon their developmental stage at the time the cells were isolated. Their permeability can be altered by their natural effectors, the secretions of the neural retina. These studies indicate there are multiple mechanisms that regulate different aspects of junctional permeability. The combination of exogenous gene expression with new assays of function will lead to a greater understanding of the mechanisms that regulate TJs.

ACKNOWLEDGMENTS Work in the authors’ laboratory was supported by National Institutes of Health Grants EY08694 (L.J.R.) and EY07031 (S.D.W.). The authors thank Dr. Laura Mitic for critically reviewing the manuscript.

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Teleost Chloride Cell Tight Junctions: Environmental Salinity and Dynamic Structural Changes Karl J. Karnaky, Jr.

CONTENTS 20.1 The Aquatic Environment: Ionic and Osmotic Challenges .......................445 20.2 Patterns of Osmoregulation in Seawater and in Fresh Water ....................446 20.2.1 Seawater Environment ...................................................................446 20.2.2 Freshwater Environment................................................................446 20.3 The Chloride Cell in Seawater-Adapted Teleosts ......................................447 20.4 Model for Sodium Chloride Secretion by the Seawater-Adapted Teleost Chloride Cell ..................................................................................449 20.5 Ion Absorption Mechanism in the Teleost Gill..........................................450 20.6 The Chloride Cell in Freshwater-Adapted Teleosts ...................................451 20.7 Insights From Isolated Epithelia Containing Chloride Cells.....................452 20.7.1 The Dynamic Nature of the Chloride Cell–Chloride Cell (Accessory Cell) Tight Junction in Seawater-Adapted Teleosts...452 20.7.2 Studies on Isolated Teleost Epithelia from Freshwater-Adapted Animals ........................................................452 20.8 New Observations from Gill Tissue in Culture .........................................453 20.8.1 Cultures with Chloride Cells.........................................................453 20.8.2 Cultures without Chloride Cells....................................................453 20.9 Chloride Cells in Nonteleost Species.........................................................454 20.10 Concluding Remarks...................................................................................454 Acknowledgments..................................................................................................455 References..............................................................................................................456

20.1 THE AQUATIC ENVIRONMENT: IONIC AND OSMOTIC CHALLENGES The fact that virtually all aquatic vertebrates have cell and interstitial ionic and osmotic characteristics different from that of their external environments is reflected 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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in the very special mechanisms evolved to maintain these differences. Whereas water and small nonelectrolytes can move through the plasma membrane at the external/internal interface, critical ions such as sodium, potassium, and chloride will not pass easily across biological membranes without the help of integral membrane transport proteins that span the lipid bilayer. It is obvious that the animal’s exposure to the external world includes not only the cell membrane of the surface cells but also the tight junctions (TJs) between these surface cells. The major focus of this chapter is the way the Na+ ion moves in a passive manner in and out of teleosts adapted to seawater, presumably in large part through the paracellular pathway composed of TJs (Karnaky, 1998). These paracellular pathway mechanisms join with other active and passive transmembrane transport mechanisms to achieve the net gain or loss of water and ions, and thus help solve osmotic problems. Because of space limitations, wherever possible, the latest reviews are referenced rather than original contributions.

20.2 PATTERNS OF OSMOREGULATION IN SEAWATER AND IN FRESH WATER The range of aquatic environments inhabited by teleosts extends from fresh water (<0.1 mOsmol/kg) to seawater (approximately 1000 mOsmol/kg), including environments several times the concentration of seawater. Despite this unusually wide range, teleosts maintain their plasma ion concentrations far below that of seawater and their internal osmotic concentration between 250 and 500 mOsmol/kg (Karnaky, 1980; 1998). Two terms are used to denote the adaptive ability of fish: stenohaline fish are limited to either freshwater or seawater environments, but not both; the euryhaline teleosts can adapt to a relatively wide range of salinities.

20.2.1 SEAWATER ENVIRONMENT The large osmotic gradients experienced by teleosts in seawater (600 to 800 mOsmol/kg) withdraw water from the animal, predominantly across the gills. Seawater teleosts drink to compensate for this exosmosis and drinking rates in seawater are three to ten times higher than in freshwater-adapted fishes (Karnaky, 1980; 1998). Most of the water and NaCl uptake occurs in the small intestine, with water absorption following osmotically the transport of ions. The low flow rates of only hypotonic or isotonic urine in teleosts do not permit the kidney to eliminate excess salt, so this required salt secretion is achieved by the gills (Evans, 1993). Thus, the gill is the focus of most research on NaCl extrusion processes in the teleost.

20.2.2 FRESHWATER ENVIRONMENT Teleosts in fresh water (plasma NaCl is approximately 150 mM) face the smallest osmotic gradients (~300 mOsmol/kg or less) (Karnaky, 1980; 1998). Because of the direction of the osmotic gradient, water flows into the hyperosmotic internal environment of the teleost, presumably mainly across the gills. Teleosts in fresh water drink little and produce large quantities of dilute urine (Karnaky, 1980; 1998) to

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compensate for this endosmosis. In addition, certain ions must be absorbed from the external environment, particularly Na+ and Cl–.

20.3 THE CHLORIDE CELL IN SEAWATER-ADAPTED TELEOSTS The first undeniable in vitro evidence for ion transport across any epithelium was provided by Keys’s historical description of a doubly perfused heart–gill preparation from the eel (Keys, 1931). This historic contribution was followed by another classic study, a morphological report in which Keys and Willmer (1932) described a large, acidophilic cell, the chloride-secreting cell (or chloride cell), in the gills of freshwater and seawater fish (reviewed in Marshall, 1995; Zadunaisky, 1996; Karnaky, 1998). Chloride cells are now thought to be present in the gills of all teleost species (Karnaky, 1998). The seawater chloride cell is approximately 8 µm wide and 20 µm long. Chloride cells from this environment exhibit an extremely rich population of mitochondria. The basal and lateral cell surface, but not the apical membrane surface, is tremendously amplified by means of branching invaginations of the plasma membrane. These take the form of tubules approximately 60 to 80 nm in diameter, and are collectively called the tubular system. For this chapter one major morphological feature should be the major focus: chloride cells in seawater teleosts do not exist as singlets, but always in multicellular complexes, effectively forming glands with a minimum of two cells (Karnaky, 1980; 1998). In these complexes, chloride cells may be continuous with other chloride cells or with cells termed accessory cells (Figures 20.1 and 20.2). The accessory cell is

FIGURE 20.1 To obtain a three-dimensional visualization of the multicellular complex architecture of seawater teleost chloride cell, several apical crypts were thin-sectioned completely through. Electron micrographs of each of these sections were digitized and combined using computer software. This figure represents the digitized image of a single thin section and clearly shows two chloride cells (CC1 and CC2) sharing an apical crypt (AC). Note that the chloride cell (or accessory cell) labeled CC2 has only a small apical exposure to the external environment. Two pavement cells (PC1 and PC2) overlap the chloride cells, leaving an opening for the exposure of the chloride cell apical membrane to the external environment. Cells marked SC are supporting, or sustentacular, cells. SW = seawater.

FIGURE 20.2 These two figures are computer reconstructions of two apical crypts composed of digital images such as that shown in Figure 20.1. These figures demonstrate the complexity of chloride cell–chloride cell and chloride cell–accessory cell interactions. For simplicity, only the chloride cell outlines have been included in these figures; the pavement cell profiles have been deleted.

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considered to be a maturing chloride cell (Wendelaar-Bonga and Jan der Meij; 1989) or a totally different type of cell with its own origin (Pisam and Rambourg; 1991). This issue has not been fully resolved. The accessory cell has fewer mitochondria and a less well developed tubular system. The adjacent chloride cells of these multicellular complexes share a relatively large invagination of their apical membranes, termed the apical crypt. The apical crypt is approximately 3 µm in diameter at its widest point, and usually has a very narrow opening to the external environment. The formation of multicellular complexes of chloride cells permits their special interaction through their shared TJs (Sardet et al., 1979; Ernst et al., 1980; Karnaky, 1998). Freeze-fracture studies have revealed the structure of TJs between chloride cells and pavement cells, the squamous cell type that covers much of the epithelial surface of the head region of the teleost. The TJs between chloride cells and pavement cells are deep (300 to 500 nm) and elaborate, as are those between adjacent pavement cells (Sardet et al., 1979; Ernst et al., 1980; 1981). In striking contrast, the TJs between contiguous chloride cells of seawater-adapted fish are shallow (25 nm) and simple (Sardet et al., 1979; Ernst et al., 1980; 1981). The multicellular complex/shallow chloride cell–chloride cell TJ is the hallmark of chloride cell morphology in seawater-adapted teleosts. The complexity of chloride cell interactions is emphasized in Figures 20.1 and 20.2.

20.4 MODEL FOR SODIUM CHLORIDE SECRETION BY THE SEAWATER-ADAPTED TELEOST CHLORIDE CELL The current model for chloride cell secretion of NaCl in teleosts adapted to seawater is shown in Figure 20.3. The basic mechanism is the subject of several recent reviews (Karnaky, 1998; Marshall and Bryson, 1998; Evans et al., 1999). The major underpinning for this model has been derived from studies of isolated jaw and opercular epithelia from the head region of several teleost species. These epithelia are flat sheets and are amenable to rigorous electrophysiological methods such as the shortcircuit current technique (reviewed in Karnaky, 1998). Studies from these epithelia showed that the Cl– ion is actively transported (Karnaky et al., 1977; Marshall and Bern, 1980; Foskett et al., 1981) and that the Na+ ion moves passively (Degnan et al., 1977; Marshall and Bern, 1980; Foskett et al., 1981). Further special analyses of isolated opercular epithelium of seawater-adapted Fundulus heteroclitus (Degnan and Zadunaisky, 1979; 1980) and the isolated skin of Gillichthys (Marshall, 1981) under open-circuit conditions have shown that the Na+ is passively distributed across these epithelia, and appears to move through a single barrier, presumably the paracellular conductive pathway. Finally, vibrating probe studies of teleost chloride cells in the tilapia opercular membrane demonstrated that chloride cells, and not pavement cells, provide the significant conductive elements in the epithelium (Foskett and Scheffey, 1982). In this model (Figure 20.3), tubular system Na+, K+-ATPase provides the primary driving force for chloride secretion by creating a steep Na+ gradient across the basolateral membrane. This Na+ gradient drives a Na+, K+, 2Cl– cotransporter, which transports these three types of ions into the cell. The Cl– ion diffuses to the apical

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FIGURE 20.3 The current model of major transport mechanisms in the chloride cell of seawater-adapted teleosts (panel at right). A chloride cell and a second chloride cell, the accessory cell, share a single apical crypt. Na+, K+, and Cl– ions enter the cell via a basolateral cotransporter, driven by the Na+ gradient generated by the action of the Na+,K+-ATPase. A basolateral K+ leak is also present in the basolateral membrane. The Cl– ion exits into the seawater via one or more Cl– channels at the apical membrane. At least one of these channels is the CFTR. Finally, the shallow TJ between adjacent chloride cells provides a paracellular pathway for the exit of Na+. Pavement cells and chloride cells are joined by deep TJs. In rapidly euryhaline teleosts adapted to fresh water, chloride cell morphology and interaction are dramatically different: chloride cells are joined by deep TJs (panel at lower left). Detailed mechanisms of freshwater ion transport are not as well understood as those for seawater NaCl secretion. In stenohaline teleosts in fresh water, chloride cells are generally found singly, and not in multicellular complexes (panel at upper right).

membrane and exits by electrical forces into the seawater environment through a Cl– channel or channels, one of which is undoubtedly the cystic fibrosis transmembrane conductance regulator (CFTR). This chloride movement establishes a transepithelial potential gradient of sufficient magnitude and sign (the seawater side of the epithelium is electronegative to the blood) to drive the Na+ ion out through the shallow, leaky TJs between chloride cells. The killifish, F. heteroclitus, gill CFTR gene sequence has been reported (Singer et al., 1998). Immunocytochemical studies reveal that CFTR is located in the apical crypt region (Wilson et al., 2000b) in 50% seawater-adapted mudskipper. Marshall et al. (1995) have reported electrophysiological evidence in killifish gill chloride cells for an anion channel sharing many similarities to CFTR.

20.5 ION ABSORPTION MECHANISM IN THE TELEOST GILL Models for ion uptake in fish adapted to fresh water have been derived mainly from studies of whole fish or two in vitro models: perfused gills or perfused head preparations. An original model derived from these studies suggested that chloride cells

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exhibited both a Na+/H+(NH4+ ) exchanger and a Cl–/HCO3– exchanger at their apical membrane (reviewed in Karnaky, 1998). More recent studies propose that a vacuolartype proton pump (H+-ATPase or V-ATPase) is located in the apical membrane of gill epithelial cells (Lin and Randall, 1995; Claiborne, 1998). This transporter generates a negative inside electrical potential by driving H+ from the cell into the external environment. This negative potential ultimately drives the Na+ ion into the gill cell via a conductive channel (reviewed in Claiborne, 1998; Karnaky, 1998; Wilson et al., 2000a). What is especially interesting about this model is that the two ions, Na+ and Cl–, may enter the freshwater-adapted fish via two different cell types. In a recent immunocytochemical study of freshwater tilapia (Oreochromis mossambicus) and rainbow trout (Oncorhynchus mykiss), the Na+ conductive channel and V-ATPase immunoreactivity were colocalized to pavement cells, the Cl–/HCO3– exchanger was localized to chloride cells, and the Na+/H+ exchanger was localized to pavement cells and accessory cells in tilapia gills (Wilson et al., 2000a). In this study, the rainbow trout Na+ channel was localized to both the chloride cell and the pavement cell. What is not clear in any of these freshwater transport models is the role of the TJ in any ion movement across the fish’s external epithelia. In these models, the major ions transported across the apical membranes of these cells move through integral membrane transporters via a transcellular, and not a paracellular, route. There is no need to invoke a paracellular pathway for the Na+ ion, as is the case for this ion in the NaCl secretory model for the seawater-adapted teleost chloride cell.

20.6 THE CHLORIDE CELL IN FRESHWATERADAPTED TELEOSTS Early freeze-fracture studies of freshwater chloride cells show that the chloride cell exists in a unicellular form, sharing only deep junctions with adjacent pavement cells (Sardet et al., 1979). Laurent and Dunel (1980) reached similar conclusions from electron microscopic study of thin sections. The chloride cell-rich opercular epithelium of F. heteroclitus has been studied after adaptation to 1% (Karnaky and Garretson, 1984; Karnaky and Kelly, 1987; Karnaky, 1992) or 8.6% seawater (Lacy, 1983), both essentially freshwater environments requiring ion absorption. In all these studies the chloride cells of the opercular epithelium remain in multicellular complexes, with deep junctions, approximately 600 nm deep, as observed in transmission electron micrographs of sectioned material (Karnaky, 1992). This deeper TJ between chloride cells and accessory cells has been described in other species. Hwang (1988) has described chloride cells in multicellular complexes in the ayu, carp, and tilapia. Specifically, the seawater-adapted tilapia chloride cell–chloride cell TJ is 20 to 40 nm deep, where the same junction is 70 to 300 nm deep (Hwang, 1987). In the rainbow trout (Salmo gairdneri), chloride cell–accessory cell TJs are 100 to 200 nm deep, whereas this same junction is 20 to 50 nm deep in seawater-adapted fish. Although this chapter has repeatedly mentioned chloride cells and accessory cells, the actual freshwater–seawater chloride cell story is apparently more complex. Pisam et al. (1987) have described two types of chloride cells in freshwater-adapted guppy, Lebistes reticulatus. Each of these is in a unicellular configuration. One of these chloride cell types disappears during adaptation to seawater, whereas the other

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chloride cell develops multicellular complexes with accessory cells. The TJ was not studied in great detail in this publication.

20.7 INSIGHTS FROM ISOLATED EPITHELIA CONTAINING CHLORIDE CELLS As mentioned above, isolated epithelia containing chloride cells have helped provide key data for the NaCl secretory model for seawater teleosts. For teleosts in this environment, electrophysiological data, combined with morphological observations, have also revealed the strikingly dynamic character of the chloride cell–chloride cell TJ.

20.7.1 THE DYNAMIC NATURE OF THE CHLORIDE CELL–CHLORIDE CELL (ACCESSORY CELL) TIGHT JUNCTION IN SEAWATER-ADAPTED TELEOSTS Stenohaline teleosts remain in fresh water or in seawater and exhibit the appropriate ion absorption or ion secretion mechanism necessary to maintain internal ionic and osmotic balance. In striking contrast, some rapidly euryhaline species, such as the killifish, F. heteroclitus, must alternate between these two sets of ion-transport mechanisms several times per day as the tides move in and out of estuarine environments. As noted above, in low-salinity environments, killifish chloride cells are arranged in multicellular complexes and the chloride cell–chloride cell TJ is deep. However, a surprising discovery in early studies was that the opercular epithelia of 1% seawater-adapted killifish develop a short-circuit current with a magnitude and time course resembling that of opercular epithelia from 100% seawater-adapted killifish (Degnan et al., 1977). Morphological studies of the opercular epithelium from 1% seawater-adapted killifish that were removed from Ussing chambers after establishment of a steady state of chloride secretion (45 min) revealed that chloride cells were still in multicellular complexes (Karnaky, 1992). However, the TJ between these chloride cells had disassembled (“unzipped”) to the shallow configuration normally seen in the chloride cell–chloride cell junction of seawater-adapted killifish. Although passive sodium fluxes were not measured during the 45-min period necessary to reach a steady-state chloride secretory current, it is likely that the TJ was being unzipped within minutes of initial exposure to physiological saline in the Ussing chamber. Clearly, the chloride cell TJ is dynamically regulated, at least when the fish is moving from the freshwater environment to the seawater environment. The killifish opercular epithelium appears to be a near-ideal model system to explore fully the morphological, biochemical, and transport properties of a dynamic TJ structure.

20.7.2 STUDIES ON ISOLATED TELEOST EPITHELIA ADAPTED ANIMALS

FROM

FRESHWATER-

As noted above, isolated, short-circuited opercular epithelia from teleosts adapted to 1% seawater exhibit a robust chloride secretion in physiological saline in Ussing chambers (Karnaky, 1986; 1992; 1998). Thus, it was initially concluded that the

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killifish opercular epithelium, studied in Ussing chambers under these conditions, would provide no insights into freshwater transport mechanisms. However, recent work has demonstrated that the killifish opercular epithelium adapted to extremely low salinity environments (1.0 mM NaCl and 0.1 mM Ca2+ for 10 days) shows active Cl– absorption (Wood and Marshall, 1994; Marshall, 1995; Marshall et al., 1997). Under these conditions, the transepithelial conductance is lower than that observed in seawater-adapted killifish opercular epithelium. Also, chloride cells are in multicellular complexes, with deep TJs. This preparation should prove to be extremely valuable in explorations of teleost ion absorption mechanisms. In summary, freshwater teleosts exhibit a low branchial ionic permeability. This low permeability may in part be because the TJs between pavement cells and between pavement cells and chloride cells are deep. In euryhaline fish adapted to fresh water the chloride cell–chloride cell junction is also deeper than it is in seawater-adapted fish chloride cells.

20.8 NEW OBSERVATIONS FROM GILL TISSUE IN CULTURE It has long been the goal of fish physiologists to generate tissue cultures of gill cells involved in osmoregulatory ion transport. Although early attempts were not fruitful, the last 6 years have produced some striking success, and some surprising findings, in this endeavor.

20.8.1 CULTURES

WITH

CHLORIDE CELLS

Recently, it has been reported that pavement cells and chloride cells from freshwater rainbow trout can be tissue-cultured together in a flat sheet suitable for studying transepithelial fluxes (Fletcher et al., 2000). In these double-seeded (gill cells from a second fish are added days after the initial culture is started), chloride cells comprised approximately 16% of the total cell numbers. The chloride cells formed deep TJs with pavement cells, and no accessory cells or shallow-type junctions typical of seawater chloride cells were observed. These electrically tight cultures exhibited extremely high transepithelial resistances, up to 34 kΩ·cm2, compared with the 50 to 100 Ω·cm2, which characterized opercular epithelia from seawateradapted killifish (Karnaky et al., 1977). Under symmetrical conditions, there was no active Na+ or Cl– transport. Active Ca2+ uptake was observed under symmetrical conditions. This is a very promising development in chloride cell physiology and should yield important insights into freshwater uptake mechanisms.

20.8.2 CULTURES

WITHOUT

CHLORIDE CELLS

Primary cultures of gills from a freshwater rainbow trout (Oncorhynchus mykiss) were first introduced by Pärt et al. (1993; Wood and Pärt, 1997). These cultures were composed exclusively of respiratory epithelial cells and did not contain chloride cells. Primary cultures of gill cells from sea bass (Dicentrarchus labrax) have been studied in the Ussing chamber (Avella and Ehrenfeld, 1997). These cultures do not

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contain chloride cells, but are composed almost exclusively of respiratory cells such as those that cover the respiratory lamellae of the teleost gill. Surprisingly, these cultures generate a short-circuit current and chloride secretion, although with a much lower magnitude than that produced by chloride cell-rich opercular epithelium of the killifish (Karnaky et al., 1977). This current is stimulated and inhibited (Avella et al., 1997; 1999) in a manner similar to that of chloride cell-containing opercular epithelia (Marshall, 1995; Marshall et al., 1997; Karnaky, 1998). Furthermore, patchclamp studies of the sea bass primary cultures revealed a low-conductance chloride channel (Duranton et al., 1997). The primary cultures in this study exhibited a very high resistance (5000 to 12,000 Ω·cm2) so that presumably freeze-fracture studies would reveal deep TJs between the respiratory cells. The way the Na+ ion would traverse this epithelium via a paracellular pathway, as described in the NaCl secretory model detailed above, remains unknown. Recall also that the vibrating probe study suggested that virtually all of the Na+ and Cl– current was carried by chloride cell elements (Foskett and Scheffey, 1982). In summary, work on sea bass primary cultures requires a reconsideration of the unique role in NaCl secretion ascribed to chloride cells for the last 70 years.

20.9 CHLORIDE CELLS IN NONTELEOST SPECIES Interestingly, chloride cells are present not only in teleosts, but also in another aquatic fish group, the lampreys. Lampreys are a subgroup of the class Agnatha, considered to be the most primitive of all living vertebrates. Lampreys live in both fresh water and seawater and are often euryhaline. These animals have osmotic and ionic concentrations approximately one third or less than that of seawater. Their osmotic and ionic regulation patterns are similar to those of teleosts (Karnaky, 1998). Not surprisingly, their gills have mitochondria-rich cells, with the extensive tubular system characteristic of the teleost chloride cell. Most importantly, freeze-fracture studies of seawater-adapted lamprey, Geotria australis, show that chloride cell–chloride cell TJs are shallow and the linear distance of these junctions is maximized by interdigitations (Bartels and Potter, 1991; Bartels et al., 1996). This morphology parallels the morphology described above for chloride cell–chloride cell TJs of seawateradapted teleost species.

20.10 CONCLUDING REMARKS The teleost chloride cell may be in a special class of epithelial cells, exhibiting one type of junction (deep) with other cell types and two types of dynamically modulated TJs (deep and shallow) with its own cell type. The type of tight junctional morphology exhibited depends on at least two factors: (1) the aquatic environment exposure (fresh water vs. seawater), and (2) the ability of the animal to move back and forth between these two environments. In summary, in freshwater-adapted teleosts limited to this environment, chloride cells are not found in multicellular complexes, and the TJ between pavement cells and chloride cells is deep. In euryhaline teleosts in freshwater environments, chloride cells (or chloride cells and accessory cells) are

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found in multicellular complexes, and chloride cells share a deep TJ with adjacent pavement cells and a deep junction with contiguous chloride cells. In euryhaline teleosts in seawater environments, chloride cells (or chloride cells and accessory cells) are found in multicellular complexes, and chloride cells share a deep TJ with adjacent pavement cells and a shallow junction with contiguous chloride cells. Finally, the deep chloride cell–chloride cell junction of freshwater-adapted teleosts can “unzip” in a matter of minutes as the fish moves from fresh water to seawater. Recent discoveries concerning the paracellular pathway for the Mg2+ ion in the thick ascending limb of Henle (TAL) offer a new perspective on the paracellular pathway provided by the shallow chloride cell–chloride cell TJ. In the TAL, renal Mg2+ resorption occurs predominantly through a paracellular conductance. Positional cloning was used to identify a human gene, paracellin-1 (PCLN-1), mutations in which cause renal Mg2+ wasting (Simon et al., 1999). PCLN-1 is related to the claudin family of TJ proteins and is located in TJs of the TAL. One could hypothesize that certain teleost chloride cells possess a special Na+-permeable, claudin-like molecule in the distalmost junctional strand, the strand closest to the seawater world. On the other hand, the deeper, more proximal strands limit Na+ permeability. In this view, this Na+-permeable strand would be present in chloride cells of euryhaline teleosts, whether they were adapted to fresh water or seawater. This special strand would also be present in stenohaline seawater teleosts, and a homologue of this Na+permeable strand would be present in chloride cells of the lamprey. This special strand would not be present in stenohaline freshwater teleosts. A recent review has noted a number of molecular advances in the broader field of fish osmoregulation, namely, the cloning and sequencing of at least nine genes for fish transport-related proteins including transporters, an important hormone, and receptors (Karnaky, 1998). Since the teleost chloride cell TJ was specifically reviewed approximately 10 years ago (Karnaky, 1992), progress has not been nearly as rapid in understanding transport mechanisms in fish as it has been in mammals. However, this author is confident that molecular strategies will be successfully applied to a number of fish transport mechanisms, including those at the important paracellular pathway. Exploration of the proteins involved in the dynamic chloride cell TJ should prove especially insightful. For example, with an increasing knowledge of the claudin family, it should be possible to elucidate the protein(s) responsible for Na+ permeability and those that are not. By combining gene cloning, bioinformatics, protein chemistry, and immunocytochemistry with appropriate antibodies, it should be possible to determine where these junctional proteins are present and how they change as fish move back and forth between the two environmental extremes. Many of these answers should be known before another 10 years pass.

ACKNOWLEDGMENTS The author thanks Martina Sedmerova for assistance in preparing the artwork. This work was supported by National Institutes of Health (GM24766, GM29099) and National Science Foundation (DCB-8409165) grants, an established investigatorship from the American Heart Association, and grants from the American Heart Association, South Carolina Affiliate, Ciba-Geigy Corporation, Cystic Fibrosis Foundation,

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the U.S. Department of Agriculture, and the Center for Membrane Toxicology Studies at the Mt. Desert Island Biological Laboratory (NIEHS Grant P30 E503828). The author was a senior fellow of the Salisbury Cove Research Fund, under a New Investigator Award, at the Mt. Desert Island Biological Laboratory, Salisbury Cove, ME during part of this work. Publication 176 of the Grice Marine Biology Laboratory of the University of Charleston.

REFERENCES Avella, M. and Ehrenfeld, J. 1997. Fish gill respiratory cells in culture: a new model for Cl–secreting epithelia, J. Membr. Biol., 156, 87. Avella, M., Part, P., and Ehrenfeld, J. 1999. Regulation of Cl– secretion in seawater fish (Dicentrarchus labrax) gill respiratory cells in primary culture, J. Physiol (London), 516, 353. Bartels, H. and Potter, I. C. 1991. Structural changes in the zonulae occludentes of the chloride cell of young adult lampreys following acclimation to seawater, Cell Tissue Res., 265, 447. Bartels, H., Moldenhauer, A., and Potter, I. C. 1996. Changes in the apical surface of chloride cells following acclimation of lampreys to seawater, Am. J. Physiol., 270, R125. Claiborne, J. B. 1998. Acid-base regulation, in The Physiology of Fishes, Evans, D. H., Ed., CRC Press, Boca Raton, FL. Degnan, K. J. and Zadunaisky, J. A. 1979. Open-circuit sodium and chloride fluxes across isolated opercular epithelia from the teleost Fundulus heteroclitus, J. Physiol., 294, 483. Degnan, K. J. and Zadunaisky, J. A. 1980. Passive sodium movement across the opercular epithelium: the paracellular shunt pathway and ionic conductance, J. Membr. Biol., 55, 175. Degnan, K. J., Karnaky, K. J., Jr., and Zadunaisky, J. A. 1977. Active chloride transport in the in vitro opercular skin of a teleost (Fundulus heteroclitus), a gill-like epithelium rich in chloride cells, J. Physiol. (London), 271, 155. Duranton, C., Tauc, M., Avella, M., and Poujeol, P. 1997. Chloride channels in primary cultures of seawater fish (Dicentrarchus labrax) gill, Am. J. Physiol., 273, C874. Ernst, S. A., Dodson, W. C., and Karnaky, K. J., Jr. 1980. Structural diversity of occluding junctions in the low-resistance chloride-secreting opercular epithelium of seawateradapted killifish (Fundulus heteroclitus), J. Cell Biol., 87, 488. Ernst, S. A., Hootman, S. R., Schreiber, J. H., and Riddle, C. V. 1981. Freeze-fracture and morphometric analysis of occluding junctions in rectal glands of elasmobranch fish, J. Membr. Biol., 58, 101. Evans, D. H. 1993. Osmotic and ionic regulation, in The Physiology of Fishes, Evans, D. H., Ed., CRC Press, Boca Raton, FL, 315. Evans, D. H., Piermarini, P. M., and Potts, W. T. W. 1999. Ionic transport in the fish gill epithelium, J. Exp. Zool., 283, 641. Fletcher, M., Kelly, S. P., Part, P., O’Donnell, M. J., and Wood, C. M. 2000. Transport properties of cultured branchial epithelia from freshwater rainbow trout: a novel preparation with mitochondria-rich cells, J. Exp. Biol., 203, 1523. Foskett, J. K. and Scheffey, C. 1982. The chloride cell: definitive identification as the saltsecretory cell in teleosts, Science, 215, 164. Foskett, J. K., Logsdon, C. D., Turner, T., Machen, T. E., and Bern, H. A. 1981. Differentiation of the chloride extrusion mechanism during sea water adaptation of a teleost fish, the cichlid Sarotherodon mossambicus, J. Exp. Biol., 93, 209.

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Hwang, P. P. 1987. Tolerance and ultrastructural response of the branchial chloride cells on salinity changes in euryhaline teleost, Oreochromis mossambicus, Mar. Biol., 94, 643. Hwang, P. P. 1988. Multicellular complex of chloride cells in the gills of freshwater teleosts, J. Morphol., 196, 15. Karnaky, K. J., Jr. 1980. Ion-secreting epithelia: chloride cells in the head region of Fundulus heteroclitus, Am. J. Physiol., 238, R185. Karnaky, K. J., Jr. 1986. Structure and function of the chloride cell of Fundulus heteroclitus and other teleosts, Am. Zool., 26, 209. Karnaky, K. J., Jr. 1992. Teleost osmoregulation: changes in the tight junction in response to the salinity of the environment, in Tight Junctions, Cereijido, M., Ed., CRC Press, Boca Raton, FL, 175. Karnaky, K. J., Jr. 1998. Osmotic and ionic regulation, in The Physiology of Fishes, Evans, D., Ed., CRC Press, Boca Raton, FL, 157. Karnaky, K. J., Jr. and Garretson, L. T. 1984. Rapid disassembly of chloride cell zonulae occludentes is synchronous with onset of chloride secretion, J. Cell Biol., 99, 294a. Karnaky, K. J., Jr. and Kelly, J. 1987. Control of chloride secretion in the teleost chloride opercular skin, Ped. Pulmon., Suppl. 1. Karnaky, K. J., Jr., Degnan, K. J., and Zadunaisky, J. A. 1977. Chloride transport across isolated opercular epithelium of killifish: a membrane rich in chloride cells, Science, 195, 203. Keys, A. B. 1931. Chloride and water secretion and absorption by the gills of the eel, Z. Vergl. Physiol., 15, 364. Keys, A. B. and Willmer, E. N. 1932. “Chloride-secreting cells” in the gills of fishes with special reference to the common eel, J. Physiol. (London), 76, 368. Lacy, E. R. 1983. Histochemical and biochemical studies of carbonic anhydrase activity in the opercular epithelium, Am. J. Anat., 166, 19. Laurent, P. and Dunel, S. 1980. Morphology of gill epithelia in fish, Am. J. Physiol., 238, R147. Lin, H. and Randall, D. 1995. Proton pumps in fish gill, in Cellular Approaches to Fish Ionic Regulation, Wood, C. M. and Shuttleworth, T. J., Eds., Academic Press, San Diego, 229. Marshall, W. S. 1981. Active transport of Rb+ across skin of the teleost Gillichthys mirabilis, Am. J. Physiol., 241, F482. Marshall, W. S. 1995. Transport processes in isolated teleost epithelia: opercular epithelium and urinary bladder, in Cellular Approaches to Fish Ionic Regulation, Wood, C. M. and Shuttleworth, T. J., Eds., Academic Press, San Diego, 1. Marshall, W. S. and Bern, H. A. 1980. Ion transport across the isolated skin of the teleost, Gillichthys mirabilis, in Epithelial Transport in the Lower Vertebrates, Lahlou, B., Ed., Cambridge University Press, Cambridge, 337. Marshall, W. S. and Bryson, S. E. 1998. Transport mechanisms of seawater teleost chloride cells: an inclusive model of a multifunctional cell, Comp. Biochem. Physiol., 119A, 97. Marshall, W. S., Bryson, S. E., Midelfart, A., and Hamilton, W. F. 1995. Low-conductance anion channel activated by cAMP in teleost Cl–-secreting cells, Am. J. Physiol., 268, R963. Marshall, W. S., Bryson, S. C., Darling, P., Whitten, C., Patrick, M., Wilkie, M., Wood, C. M., and Buckland-Nicks, J. 1997. NaCl transport and ultrastructure of opercular epithelium from a freshwater-adapted euryhaline teleost, Fundulus heteroclitus, J. Exp. Zool., 277, 23. Pärt, P., Norrgren, L., Bergstrom, E., and Sjoberg, P. 1993. Primary cultures of epithelial cells from rainbow trout gills, J. Exp. Biol., 175, 219.

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Pisam, M. and Rambourg, A. 1991. Mitochondria-rich cells in the gill epithelium of teleost fishes: an ultrastructural approach, Int. Rev. Cytol., 130, 191. Pisam, M., Caroff, A., and Rambourg, A. 1987. Two types of chloride cells in the gill epithelium of a freshwater-adapted euryhaline fish: Lebistes reticulatus; their modifications during adaptation to saltwater, Am. J. Anat., 179, 40. Sardet, C., Pisam, M., and Maetz, J. 1979. The surface epithelium of teleostean fish gills. Cellular and junctional adaptations of the chloride cell in relation to salt adaptation, J. Cell Biol., 80, 96. Simon, D. B., Lu, Y., Choate, K. A., Velazquez, H., Al-Sabban, E., Praga, M., Casari, G., Bettinelli, A., Colussi, G., Rodriguez-Soriano, J., McCredie, D., Milford, D., Sanjad, S., and Lifton, R. P. 1999. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption, Science, 285, 103. Singer, T. D., Tucker, S. J., Marshall, W. S., and Higgins, C. F. 1998. A divergent CFTR homologue: highly regarded salt transport in the euryhaline teleost F. heteroclitus, Am. J. Physiol., 274, C715. Wendelaar-Bonga, S. E. and van der Meij, C. J. M. 1989. Degeneration and death, by apoptosis and necrosis, of the pavement and chlordie cells in the gills of the teleost, Oreochromis mossambicus, Cell Tissue Res., 255, 235. Wilson, J. M., Laurent, P., Tufts, B. L., Benos, D. J., Donowitz, M., Vogl, A. W., and Randall, D. J. 2000a. NaCl uptake by the branchial epithelium in freshwater teleost fish: an immunological approach to ion-transport protein localization, J. Exp. Biol., 203, 2279. Wilson, J. M., Randall, D. J., Donowitz, M., Vogl, A. W., and Ip, A. K. Y. 2000b. Immunolocalization of ion-transport proteins to branchial epithelium mitochondria-rich cells in the mudskipper (Periophthalmodon schlosseri), J. Exp. Biol., 203, 2297. Wood, C. M. and Marshall, W. S. 1994. Ion balance, acid-base regulation, and chloride cell function in the common killifish, Fundulus heteroclitus. A euryhaline estuarine teleost, Estuaries, 17. Wood, C. M. and Pärt, P. 1997. Cultured branchial epithelia from freshwater fish gills, J. Exp. Biol., 200, 1047. Zadunaisky, J. A. 1996. Chloride cells and osmoregulation, Kidney Int., 49, 1563.

21

Tight Junctions and Proteases Yehuda Ben-Shaul and Ilana Ophir

CONTENTS 21.1 Introduction .................................................................................................459 21.2 Effects of Proteases.....................................................................................461 21.3 Tight Junctions in Human Colon Adenocarcinoma ...................................461 21.3.1 Primary Tumors and Derived Cell Lines ......................................461 21.3.2 Effects of Trypsin on Formation of Tight Junctions in HT29 Cells ...............................................................................................462 21.3.3 Separation of Induction and Assembly of Tight Junctions...........464 21.3.4 Effect of Temperature ....................................................................464 21.3.5 Calcium Ions..................................................................................466 21.3.6 Energy Requirement ......................................................................467 21.3.7 Protein Synthesis ...........................................................................468 21.3.8 Effects of Various Proteases on Tight Junction Formation ..........468 21.3.9 Protease Inhibitors .........................................................................469 21.4 Degradation of Tight Junctions ..................................................................473 21.5 Tight Junction Formation and Cell Differentiation....................................475 References..............................................................................................................478

21.1 INTRODUCTION Tight junctions (TJs) are specialized domains found at regions of contact between membranes of adjacent cells in epithelia and endothelia. They form a seal between neighboring cells and thus control the permeability of the paracellular pathway. They also maintain the apical and basolateral compartments of cell surface membranes, essential for the physiological activities of these tissues. TJs are best observed in freeze-fractured replicas of cell membranes, using electron microscopy, where they appear as anastomosing fibrils on the protoplasmic face (PF) of one cell and as corresponding furrows on the exoplasmic face (EF) of the neigboring cell (Farquhar and Palade, 1963; Staehelin, 1974; Gumbiner, 1987; Cereijido et al., 1998). Knowledge of the molecular nature of TJs has rapidly expanded in recent years. Since Stevenson et al. (1986) characterized the first (peripheral) protein ZO-1, localized at the tight-junctional area, nine peripheral tight-junctional proteins were identified, 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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as well as two types of TJ-specific integral membrane proteins: occludin and the family of claudins (Mitic and Anderson, 1998; Stevenson and Keon, 1998; Goodenough, 1999; Tsukita and Furuse, 1999). Functional TJs form a complex belt of sealing elements, named zonula occludens, at the apical position of lateral membranes of adjacent cells. Claude and Goodenough (1973) demonstrated a correlation between the complexity of zonula occludens structure as seen in freeze-fracture preparations and the electrical resistance of several epithelia. In epithelia with high transepithelial resistance, the zonula occludens consisted of many strands. In epithelia with low transepithelial resistance, TJs were much reduced and were sometimes composed of only one fibril. Today, exceptions to this rule are known. It was found that the barrier function of TJs is not only controlled by the complexity of their structure but is also dynamically regulated by modulations of the specific junction-associated proteins and by the actin cytoskeleton (Balda and Matter, 1998; Madara, 1998; Stevenson and Keon, 1998). Even the simple TJs exhibit strands on the PF face of one cell and corresponding grooves on the EF face of the neighboring cell, suggesting that their structure is similar to that of complex TJs. Sometimes, fibrils of TJs are not positioned at the apical area of lateral cell membranes but are rather scattered over the entire cell membrane. This type of TJ was named fascia occludens (Farquhar and Palade, 1963). Fujimoto (1995) developed a method of freeze-fracture replica immunolabeling and demonstrated that the TJ-specific protein occludin colocalizes with simple strands of TJs as well as with anastomosing fibrils. Fascia occludens were described in many biological systems. In mammalian embryos, TJs were observed at compaction (six to eight cell stage) and their formation was found to be a gradual process at the morphological level (from fascia to zonula) as well as at the molecular and physiological level (Ducibella et al., 1975; Dale et al., 1991; Fleming et al., 1992). Using newly generated tight-junctional markers, Mezdorf et al. (1998) found recently that at early developmental stages of Xenopus embryos, TJs were not at their normal apical positions and only later in development were apical/lateral boundaries and permeable fences formed. Fascia occludens were observed during morphogenesis of various tissues (Schneeberger et al., 1978; Polak-Charcon et al., 1980; Meyer and Overton, 1983; Yamamoto and Katoaka, 1988). They were also found in several invasive carcinomas, where a correlation between loss of cell polarity and presence of fascia occludens rather than zonula occludens was reported (Weinstein et al., 1976; Pitelka et al., 1980; PolakCharcon et al., 1980; Cochand-Priollet et al., 1998). It is widely accepted that TJs are dynamic structures subject to rapid modulations (Cereijido et al., 1998; Madara, 1998). The dynamic nature could be observed by transferring aggregates of epithelial cells into collagen gels. The collagen induced reversal of cell polarity and a rapid relocation of TJs from the periphery of the aggregate toward the central lumen (Kirkland, 1988; Wang et al., 1990; Ophir et al., 1995). Various experimental conditions have been reported to result in de novo formation of TJs in tissues and cultured cells. Rapid proliferation of TJs was observed in excised prostate tissue incubated in buffer (Kachar and Pinto da Silva, 1981), the osmotic gradient of mannitol induced formation of TJs in C1-1D cells (Azarnia

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et al., 1981) and in the intestinal mucosa of the rat (Madara, 1983), TJs were formed by vitamin A in embryonic skin (Elias and Friend, 1976), and rapid formation of TJs was reported in HT29 cells by hypertonic salt solutions (Faff et al., 1988).

21.2 EFFECTS OF PROTEASES Proteases were found to induce formation of TJs in a variety of experimental systems. Formation of tight-junctional strands on the lateral cell surfaces of excised pancreas tissue were observed after treatment with papain (Orci et al., 1973). Trypsin digestion induced proliferation of tight-junctional fibrils between the corneum and granular layer of rat oral mucosa (Shimono and Clementi, 1977). Application of trypsin (in the presence of Ca2+) to the basolateral surface of confluent Madin–Darby canine kidney (MDCK) cell monolayers with formed TJs induced formation of basolaterally oriented aberrant tight-junctional strands (Lynch et al., 1995). This was accompanied by an increase in transepithelial electrical resistance, as great as 90%. Under similar conditions, application of trypsin to the apical surface had almost no effect on either transepithelial resistance or the number of aberrant tight-junctional strands. ZO-1 extended below the TJs along the basolateral surface following a brief exposure to trypsin. Tight-junctional strands were not formed by trypsin in MDCK cells maintained in low-Ca2+ medium, suggesting that under these conditions tight-junctional precursors and/or trypsin-sensitive proteins, regulating tight-junctional strand assembly, are sequestered in a vesicular compartment that is inaccessible to exogenous trypsin. Different results were obtained when MDCK cells were exposed to rat mast cell protease, RMCP-II (Scudamore et al., 1998). Basolateral, but not apical, exposure of polarized MDCK monolayers on porous supports to RMCP-II led to an increase in electrical conductance and permeability to macromolecules via the paracellular route, accompanied by a decrease in the immunostaining of TJ-associated proteins occludin and ZO-1. These data were in accordance with studies that established a link between the release of mast cell granule chymases in inflammatory processes and increased mucosal permeability. It should be noted that most experimental systems that were used for the study of TJ formation express TJs permanently. In contrast, HT29 cells, a human colon adenocarcinoma cell line, grow in culture virtually without TJs and form extensively TJs following protease treatment (Polak-Charcon et al., 1978; Cohen et al., 1985). This cell system was used for an in-depth study of TJ formation that will be described in detail.

21.3 TIGHT JUNCTIONS IN HUMAN COLON ADENOCARCINOMA 21.3.1 PRIMARY TUMORS

AND

DERIVED CELL LINES

Primary tumors of human colon adenocarcinoma exhibit cells with morphological alterations in comparison to the normal colon mucosa. Various degrees of loss of cell polarity and changes in tissue architecture can be observed, usually in correlation with the pathological stage of the tumor. When tumor cells are freeze-fractured and

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replicas are observed by electron microscopy, many cells exhibit a fascia occludens type of TJ rather than zonula occludens, or even a complete loss of TJs. A correlation between loss of cell polarity and loss of the normal structure of TJs was found (Polak-Charcon et al., 1980). Many cell lines have been derived from human colon cancers. They vary in their differentiation characteristics, i.e., in their ability to polarize, to form TJs, apical brush border, and typical enterocytic enzymes, to secrete mucus, etc. (Zweibaum et al., 1991). These cell lines can serve not only as in vitro models for studies of human colon cancers, but also as cell models for studies of epithelial cell differentiation and polarization. Three of these cell lines are extensively used as in vitro models for polarization: Caco-2 and T84 that differentiate spontaneously at confluence, and HT29 that can be modulated to differentiate by changing the external conditions. To study cell differentiation, HT29 can serve as an important tool, since it is a pluripotent intestinal cell line and thus resembles the intestinal stem cell. This cell line was established in culture in 1964 by Fogh (Fogh and Trempe, 1975). HT29 cells have a very high consumption of glucose and therefore require a high concentration of glucose in the culture medium. When grown in Dulbecco Modified Eagle’s Medium (DMEM) at high glucose concentration and in the presence of 10% fetal calf serum (standard conditions), the cells are unpolarized and they grow as a multilayer. However, modifications in the culture medium can cause patterns of intestinal differentiation. Growth in DMEM containing galactose or inosine instead of glucose (Pinto et al., 1982; Trugnan et al., 1987; Le Bivic et al., 1988; Zweibaum et al., 1991), growth in RPMI medium (Polak-Charcon, 1989), and growth in the presence of suramin (Fantini et al., 1990) or in the presence of butyrate (Augeron and Laboisse, 1984) were found to cause differentiation and polarization of the cells. About 30 days of adaptation were required for differentiation of the cell culture. Many cell divisions occur during this period and it seems that a selection of polarizing cells that can survive and proliferate under the new conditions may be involved. HT29 cells can also be modulated to form rapidly the fascia occludens type of TJ by short treatment with proteases or by hypertonic salt solutions (Polak-Charcon et al., 1978; Cohen et al., 1985; Faff et al., 1988).

21.3.2 EFFECTS OF TRYPSIN IN HT29 CELLS

ON

FORMATION

OF

TIGHT JUNCTIONS

The effect of proteases on HT29 cells was extensively studied. It was found that when the cells were grown in culture dishes under standard conditions they were flat but not polar and were virtually without TJs. When incubated in a solution of 2.5 mg/ml trypsin, they became round and within 10 to 15 min most of them detached from the culture dish and formed small cell aggregates. By using the freeze-fracture technique it was found that TJs of the fascia occludens type were formed on the cell surface. This process was fast and reached its maximum after 15 min (Polak-Charcon et al., 1978). In contrast to zonula occludens that can be observed in freeze-fractured replicas of polarized epithelia in a definite location, underneath the brush border, fascia occludens formed in HT29 were not limited to a particular location on the

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FIGURE 21.1 Freeze-fractured membranes of HT29. (a) Untreated. (b) Treated with trypsin (1 mg/ml in DMEM, 15 min, 37°C). Bar = 1 µm. (From Ophir, I. et al., Eur. J. Cell Biol., 49: 116, 1989. With permission.)

cell surface membrane. To quantitize the effect of trypsin on TJ formation a statistical approach had to be developed. Replicas of freeze-fractured cells were systematically screened and cell membranes with or without TJs were counted. Every membrane fragment that could be unequivocally identified as part of the cell membrane (by topological aspects or presence of microvilli) was accepted as one unit, irrespective of size. This method of evaluation has a tendency to underestimate the percentage of cells with TJs, because the total surface of any given cell cannot be observed, and an exposed membrane without junctions may belong to a cell with TJs on other parts of the surface that were not exposed by freeze fracturing. A minimum of 60 membrane fragments were evaluated per each data point, yielding highly reproducible results (within 5% error limit) in repeated experiments. By using this method it was found that treatment with 2.5 mg/ml trypsin over a period of 15 min at 37°C induced the formation of junctions in about 60 to 70% of observed cell membrane surfaces, as compared with less than 1% in untreated cells (Figure 21.1) (Cohen et al., 1985). Table 21.1 summarizes the effect of trypsin concentration on TJ formation. It can be concluded that while very low concentrations of trypsin cannot induce TJ formation, 0.05 mg/ml is sufficient to induce the maximal effect at 37°C.

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TABLE 21.1 Induction of TJ in HT29 Cells by Trypsin Trypsin (µg/ml) 0 2.5 5 25 50 250 2500

Membranes Totala 523 87 48 240 72 57 154

With TJb 5 0 1 73 45 25 87

% TJc 0.9 0 2 30 62 43 56

Cells were treated for 15 min with solutions of trypsin in DMEM at 37°C. a b c

No. of membranes observed. No. of membranes with TJ. % of membranes with TJ.

Source: Cohen, E. et al., Exp. Cell Res., 156, 103, 1985. With permission.

21.3.3 SEPARATION OF INDUCTION OF TIGHT JUNCTIONS

AND

ASSEMBLY

Low temperature inhibited TJ formation by trypsin. Only a slight increase in TJs was observed, even after incubation of 3 h in 2.5 mg/ml trypsin at 0°C, and the few junctions that were formed were short and poorly developed. However, when cells were treated in the cold with trypsin, and trypsin was then removed by repeated washings with soybean trypsin inhibitor (STI) and the cells were incubated in DMEM (in the presence of STI) at 37°C for 15 min, an abundant assembly of TJs was observed. The number and complexity of the TJs were even higher than in the one-step induction. This two-step protocol, which separates the induction and assembly of TJs, enables a study of the factors required to operate in each respective phase, such as membrane fluidity, cytoskeletal involvement, etc. (Talmon et al., 1984; Cohen et al., 1985).

21.3.4 EFFECT

OF

TEMPERATURE

The temperature dependence of TJ formation was studied in detail (Cohen et al., 1990). It was found that when HT29 cells were treated with trypsin (0.05 or 1 mg/ml) for 15 min at various temperatures, or with trypsin at 4°C for 15 min and then further incubated for 15 min without trypsin at various temperatures (two-step protocol), no TJs were found at temperatures below 10°C and the fraction of cell membranes with TJs increased sharply to over 50% in a very narrow temperature range between 14 and 16°C, reaching maximal values of more than 60% at 20°C. Morphometric analysis of the length of tight-junctional strands formed per membrane area also

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FIGURE 21.2 Temperature dependence of induced TJ formation. (●) Cells treated with trypsin (1 mg/ml) for 15 min at the temperatures indicated; (, ) cells treated with trypsin (1 mg/ml or 50 µg/ml) at 4°C for 15 min and subsequently incubated with STI for 15 min at temperature indicated; (▫, ) cells treated with trypsin (1 mg/ml) in the presence of 0.32 M NaCl or CsSO4 for 30 min at the temperature indicated; (A) fraction of membrane with TJs; (B) length of TJ strands per membrane area (µm/µm2). (From Cohen, E. et al., J. Cell Sci, 97: 119, 1990. With permission.)

yielded a sigmoid curve very similar to the numerical evaluation (Figure 21.2). The possibility that the observed sigmoidal temperature curve could be associated with a sharp change in the lateral mobility of lipid molecules in the plasma was studied using the fluorescence photobleaching recovery technique with the fluorescent phospholipid N-NBD-PE (Axelrod et al., 1976). No measurable changes in the organization of the lipids in HT29 plasma membrane were found at the temperature range examined (6 to 35°C). The sigmoidal temperature characteristic of TJ assembly is reminiscent of folding/unfolding processes in macromolecules such as proteins or

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DNA and seems to indicate that likewise the assembly of TJ strand is a highly cooperative process.

21.3.5 CALCIUM IONS The influence of Ca2+ depletion on the permeability of tight-junctional seal was investigated in epithelia and in epithelial-like cell sheets (such as confluent MDCK cells) in culture. It was documented and recently reviewed (Cereijido et al., 1998) that Ca2+ is involved in synthesis of TJs and in maintenance of apical/basolateral polarity and is needed primarily on the extracellular side of the cell membrane, probably to activate E-cadherins. In contrast to these data, the absence of Ca2+ and Mg2+ had no effect on the formation and maintenance of the fascia occludens type of TJs induced by trypsin in HT29 cells (Faff et al., 1987). Cells incubated in Ca2+and Mg2+-free buffer in the presence of 3 mM EGTA or EDTA, and then treated with trypsin in the same buffer, formed equal numbers of TJs on cell membranes as did cells that were treated with trypsin in the presence of 1.2 mM Ca2+ and 0.8 mM Mg2+. In both cases, abundant TJs were observed on 57 to 70% of the exposed membranes (Figure 21.3). Transitions of fracture plane from PF to EF in the area covered by TJs, occurred on 65 to 77% of membranes with TJs. The PF/EF transition indicates that the membranes exposed belong to two closely apposed cells that were intimately connected by the TJ structure. No evidence for the occurrence of “split” TJs was found. It can be concluded that the structural integrity and formation of TJ

FIGURE 21.3 Freeze-fractured membrane of HT29 cells treated with trypsin (1.5 mg/ml) for 30 min at 37°C in the presence of (a) 1.2 mM Ca2+ + 0.8 mM Mg2+, (B) 3 mM EGTA, (C) 3 mM EDTA; wedges, PF/EF transitions; ×38,000. (From Faff, O. et al., Biochem. Biophys. Acta, 905: 48–56, 1987. With permission.)

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fibrils does not require extracellular divalent ions in this experimental system. Therefore, the focal TJs of HT29 cells (induced by trypsin) may be different from TJs in other epithelial systems where Ca2+ appears to be necessary for zonula occludens stability. Alternatively, the effect of chelators on cell sheets and epithelia, which have been usually documented by measurement of electric resistance, may be caused, not by actual splitting of tight-junctional strands, but rather by dynamic local modulations of junction-associated proteins.

21.3.6 ENERGY REQUIREMENT Energy requirement for the induction of TJ formation by trypsin in HT29 cells was evaluated using the metabolic inhibitors dinitrophenol (DNP) and deoxyglucose (DG) (Muckter et al., 1987). Levels of ATP in the cells were reduced by DG treatment to 20% of control values. However, no effect on the amount and complexity of induced TJ fibrils was found. Treatment of HT29 cells with 5 mM DNP (in glucose-free medium) gave a drastic reduction of ATP to 1 to 2% of normal value within 1 h. This reduction was fully reversible within 2 h when DNP was replaced by glucose. The low ATP level was accompanied by a marked reduction of trypsin-mediated TJ formation (Figure 21.4). TJs were observed only on 20% of the cell membranes (compared with 60% in controls) and were much reduced in length and complexity. Short, single-stranded segments of parallel strands with few anastomoses were a characteristic feature of energy-depleted cells (Figure 21.5). The reduction in length and in complexity of tight-junctional strands was confirmed by morphometry: the length of induced tight-junctional strands in DNP-treated cells was less than 10% as compared to controls, and anastomoses were almost absent in ATP-depleted cells.

FIGURE 21.4 TJ formation during depletion and recovery of ATP in HT29 cells treated with DNP. Cells were incubated in glucose-free DMEM containing 5 mM DNP at 37°C. At the times indicated, cells were treated with trypsin in DMEM containing 5.0 mM DNP for 15 min at 37°C. (●) TJ; () ATP content. Arrow indicates removal of DNP to allow for recovery of the ATP level. (From Muckter, H. et al., Eur. J. Cell Biol., 44: 258, 1987. With permission.)

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FIGURE 21.5 Transmission electron micrographs of freeze-fractured membrane fragments of HT29 cells. Parallel fibrils of TJ induced by trypsin (1 mg/ml) after pretreatment with 5 mM DNP and postincubation in 5 mM DNP for 45 min in the absence of glucose. ×40,000. (From Muckter, H. et al., Eur. J. Cell Biol., 44: 258, 1987. With permission.)

21.3.7 PROTEIN SYNTHESIS Involvement of protein synthesis in TJ formation by trypsin was studied using the protein synthesis inhibitors cycloheximide and puromycin (Ophir et al., 1989). Cells were incubated with the inhibitors (at concentrations of 20 µg/ml) for 1 to 12 h. Under these conditions protein synthesis was reduced to about 10% of controls. However, addition of trypsin to the cells induced high rates of TJ formation on cell surfaces, similar in amount and complexity to controls (Figure 21.6). Even extremely high concentrations of cycloheximide (2 mg/ml; 2 h) did not inhibit TJ formation, and TJs were found on 80 to 90% of cell surfaces. Cycloheximidine was found in another cell system (MDCK cell sheets) to increase the electrical resistance. It was suggested there, that the metabolic inhibitor might suppress the synthesis of a labile inhibitor of TJ assembly (Griepp et al., 1983; Gonzalez-Mariscal et al., 1985).

21.3.8 EFFECTS OF VARIOUS PROTEASES JUNCTION FORMATION

ON

TIGHT

HT29 cells are suitable for a comparative study on the effect of various proteases on TJ formation (Cohen et al., 1985). Table 21.2 summarizes experiments in which TJs were induced by a variety of proteases of different origins. Eight endopeptidases analyzed could induce TJ formation in HT29 cells, irrespective of the origin of the

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FIGURE 21.6 Effect of protein synthesis inhibitors on TJ formation in HT29 cells. Cells were incubated with the respective inhibitor (20 µg/ml) for the time indicated. They were then treated with trypsin (1 mg/ml in DMEM, 15 min, 37°C). (From Ophir, I. et al., Eur. J. Cell Biol., 49, 116, 1989. With permission.)

enzyme, from vertebrates, plants, or microorganisms. The structure and complexity of these junctions were similar to those formed by trypsin. Pronase, a bacterial protease mixture, showed a marked optimum at enzyme concentration of 10 to 100 µg/ml. Higher concentrations yielded progressively fewer TJs. Thrombin was the only endopeptidase that yielded low values of TJ formation. In contrast to endopeptidases, each of the three exopeptidases analyzed (aminopeptidase M, aminopeptidase C, and carboxypeptidase, all of mammalian origin) did not induce the formation of TJs. The data suggested that endoproteolytic modification of membrane proteins could induce the formation of TJs, irrespective of the specificity of the enzyme.

21.3.9 PROTEASE INHIBITORS Is a proteolytic cleavage of a cellular protein precursor a prerequisite step in TJ assembly? This question was addressed using protease inhibitors in three protocols of TJ formation: (1) induction of TJs by trypsin in HT29 cells; (2) induction of TJs by salt solutions in HT29 and Caco-2 cells; (3) spontaneous assembly of TJs in Caco-2 cell culture (Bacher et al., 1992). 1. The two-step protocol of separating induction of TJs by trypsin at 0°C and assembly without trypsin at 37°C was used to evaluate the importance of proteolytic modification of membrane proteins for the formation of TJs in HT29 cells. Trypsin was found to cleave artificial substrates rather efficiently even at low temperatures (Walsh, 1970). Although no TJs were assembled in HT29 by trypsin in the cold, the catalytic activity of the enzyme was absolutely required for the induction phase. Virtually no TJs were formed in the assembly phase when trypsin had been pretreated with protease inhibitors such STI or PMSF (phenylmethyl sulfonyl fluoride) (Cohen et al., 1985). 2. Involvement of a proteolytic modification in the formation of TJs was also studied in HT29 treated with salt solutions. Brief treatments of HT29

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TABLE 21.2 Induction of TJ in HT29 Cells by Various Proteases Protease Aminopeptidase C Aminopeptidase M Carboxypeptidase α-Chymotrypsin

Collagenase Elastase

Papain

Plasmin

Pronase E

Thrombin

Trypsin

Conc. (g/ml) 400 500 250 1250 125 1250 125 12.5 1.25 1250 125 1250 125 12.5 1.25 1250 500 125 12.5 1250 125 12.5 1250 125 12.5 1.25 0.125 5000 1250 500 50 5 0.5 1250 125 12.5 1.25

Membranes Total 54 53 52 81 45 55 53 50 76 201 53 52 63 46 72 71 50 45 73 53 63 49 327 246 125 118 71 60 62 203 146 90 110 72 54 49 85

a

With TJb 1 1 1 2 0 34 28 15 1 75 0 28 29 6 3 41 13 5 0 16 5 0 14 81 65 46 4 0 2 7 7 5 3 40 26 16 10

% TJc 1.8 1.8 2.0 2.4 0 62 53 30 1.3 37 0 54 46 13 4.2 58 26 11 0 30 7.9 0 4.3 33 52 39 5.6 0 3.2 3.4 4.8 5.5 2.7 56 48 33 12

Cells were incubated for 30 min at 37°C with a solution of the respective protease in DMEM. The specific activities of the proteases used were as follows: 0.1 U/mg aminopeptidase C; 7.6 U/mg aminopeptidase M; 24 U/mg carboxypeptidase; 34 U/mg chymotrypsin; 0.22 U/mg collagenase; 40 U/mg elastase; 20 U/mg papain; 4 U/mg plasmin; 8 U/mg pronase; 320 U/mg thrombin; 96 U/mg trypsin. a-c See Table 21.1. Source: Cohen, E. et al., Exp. Cell Res., 156, 103, 1985. With permission.

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60 50 40 30 20 10 0 0

250

500

750

1000

FIGURE 21.7 Formation of TJ in HT29 cells. Cells were treated for 30 min at 37°C with 320 mM cesium sulfate in HBSS containing leupeptin (●), dihydroleupeptin (), or antipain (). (From Bacher, A. et al., Exp. Cell Res., 200: 97, 1992. With permission.)

cells with appropriate salt solutions were found to induce TJs on the cell membranes (Faff et al., 1988). The amount and complexity of the TJs were similar to those found after induction by trypsin. A comparative study using a variety of inorganic and organic salts showed that all alkali sulfates induced TJs although with different yield. Both calcium and magnesium chloride were potent inducers, whereas sodium chloride did not induce TJs. The data suggested tentatively that antichaotropic ions that stabilize hydrophobic interactions between macromolecules (Washabaugh and Collins, 1986) had the potential to trigger the formation of TJs in HT29 cells. It was proposed that the induction of TJ formation by salt solutions could proceed via activation of endogenous protease. This hypothesis was tested by treatment of HT29 cells with cesium sulfate (a potent salt inducer) in the presence of the protease inhibitors leupeptin and antipain. Both inhibitors were found to suppress the formation of TJs (Figure 21.7). This effect was reversible and when the inhibitor was removed, cesium sulfate could form a large quantity of TJs. 3. The effect of protease inhibitors was also studied in the polar human colon adenocarcinoma cell line Caco-2 known to develop the zonula occludens type of TJs spontaneously. When grown on permeable support, the cell layer develops moderate transepithelial resistance of about 170 Ω·cm2. However, when these cells were cultured in the presence of 400 µM leupeptin (nontoxic concentration), the occurrence of the zonula type of TJ was reduced by a factor of about eight (morphometric study) and the cell layers did not develop transepithelial resistance even after growth for several days (Figure 21.8). The removal of the inhibitor was followed by a rapid development of transepithelial resistance and tight-junctional elements of the zonula occludens type. Treatment of Caco-2 cells with cesium

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FIGURE 21.8 Transepithelial electrical resistane of Caco-2 cell monolayers. Cells were grown on carbon-coated dialysis membranes, , cells grown in DMEM/F12 medium; ▫, cells grown in DMEM/F12 medum containing 400 µM leupeotin; , cells grown in DMEM/F12 containing 400 µM dihydroleupeptin. (From Bacher, A. et al., Exp. Cell Res., 200, 97, 1992. With permission.)

sulfate induced the fascia type of TJs (in addition to the zonula occludens) on lateral surfaces of the cells, similar to those induced in HT29 cells. Leupeptin inhibited the formation of this type of TJs as well. When the protease inhibitor was removed, cesium sulfate could induce new fascia occludens rapidly. The rapid restoration of the potential to form TJs in HT29 and Caco-2 cells after removal of the inhibitor, suggests that the protease involved in TJ formation is located on the outer surface of the cell membrane. If the protease were located in the cytoplasm, a lag phase to allow for efflux of the inhibitor from the cell was expected. On the basis of these data it was suggested that 1. TJ fibrils are assembled from protein precursors in the cell membrane. 2. Limited proteolysis is required to transform these proteins into functional elements of the TJs. 3. Correct proteolytic cleavage is normally executed by endogenous protease which can be inhibited by leupeptin. This protease may be located on the cell membrane. 4. The endogenous protease is nonfunctional in HT29 cells under standard culture conditions. 5. The endogenous protease activity may be substituted by an exogenous protease such as trypsin. Alternatively, a cellular protease can be recruited or activated by treatment with appropriate salt solutions.

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Recently, the presence and expression of trypsinogen mRNA was determined in undifferentiated and differentiated adenocarcinoma cell lines (Bernard-Perrone et al., 1998). It was found that Caco-2 cells and HT29 cells grown in the absence of glucose, both exhibiting enterocytic differentiation, exhibit also variations in trypsinogen I expression. In both cell lines a high and transient peak was observed during the first steps of differentiation. In contrast, undifferentiated HT29, grown in high concentration of glucose, did not exhibit similar variations in trypsinogen I expression.

21.4 DEGRADATION OF TIGHT JUNCTIONS HT29 cells provide an attractive model for qualitative and quantitative studies not only for TJ formation, but also for degradation of TJs. TJs formed in these cells by brief treatment with proteases or by hypertonic salt solutions degraded completely during a period of 2 to 3 h in the absence of the inducer. The removal of TJ fibrils from the cell surface involved a topological reorganization with formation of characteristic patches of roughly circular appearance, where the TJ strands were frequently characterized by parallel or concentric arrangements. Open networks and single strands were also observed and strand interruptions occurred frequently (Figure 21.9). Many vesicles with remnants of tight-junctional strands were found in the cytoplasm in the process of TJ degradation (Polak-Charcon and Ben-Shaul, 1979). Loss of complexity of zonula occludens and fragmentation of TJs were also observed during amphibian and avian neurulation and in early chick embryo. These observations prompted the hypothesis that TJs disappear by disintegration of the structural components followed by their redistribution in the plasma membrane (Revel et al., 1973; Decker, 1981). On the other hand, evidence for the removal of TJ fibrils by endocytosis-related processes came from numerous observations of cytoplasmic vesicles bearing remnants of tight-junctional structures and of an association of membrane fragment with lysosomal vesicles in different tissues and cell systems. These vesicles were shown in situ in the rat intestinal epithelium (Staehelin, 1974; Madara, 1990) in the human fetal hindgut (Polak-Charcon et al., 1980) in normal and dystrophic rat retina (Caldwell et al., 1984), in chick embryonic lung tissue after trypsin treatment (Talmon and Ben-Shaul, 1979), and in COLO 316 cells spontaneously released into the medium from confluent monolayers. The relationship of filipin–sterol complexes to the TJ complex on the plasma membrane and to the vesicle membrane with TJ remnants in COLO 316 suggested the degradation by lysosomes (Risinger and Larsen, 1983). Quantitative freeze-fracture electron microscopy was used to monitor the process of TJ degradation in HT29 cells. Replicas were screened and membranes with and without TJ strands were counted. In some experiments the length of TJ strands and the exposed membrane area were measured in sets of 30 membranes randomly collected. The majority of the cells removed TJ strands from the surface during a period of 2 h, and this process was not affected by the protein synthesis inhibitors puromycin and cycloheximide. The removal of TJ elements proceeded at the same velocity as in nontreated cells. However, it seems that the digestion of cytoplasmic vesicles was slower in cells treated with the inhibitors, as they contained large amounts of vesicles with TJ remnants in their cytoplasm (Ophir et al., 1989).

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FIGURE 21.9 Freeze-fractured plasma membrane of HT29 cells treated with trypsin for 15 min and then transferred to fresh medium for 1 h. Small groups of anastomosing ridges on PF faces. Final magnification ×31,000. (From Polak-Charcon, S. and Ben-Shaul, Y., J. Cell Sci., 35, 393, 1979. With permission.)

The removal of TJ strands from the cell surface was found to be an energydependent process (Keller et al., 1992). Deoxyglucose (DG), which inhibits glycolysis, reduced the level of ATP to about 20% of normal values in the absence of glucose, and it strongly inhibited the degradation process (60% of membranes with TJs after trypsin induction in the presence of the inhibitor and 53% after 13 h). The requirement of ATP for degradation was in contrast to findings on the assembly of TJs that was inhibited only by extremely low levels of ATP (Muckter et al., 1987). It was proposed that the assembly mechanism of TJ strands could represent a selforganizing system requiring no metabolic energy, whereas the removal of TJ strands from the cell surface requires energy in the form of ATP. The protonophor dinitrophenol (DNP), which acts as an efficient uncoupler of oxidative phosphorylation, reduced ATP to very low levels in the absence of glucose and completely blocked degradation in concentrations of 0.5 to 5 mM. DNP does not reduce ATP levels in the presence of glucose. However, at high concentrations (4 to 5 mM) it inhibited degradation of TJs even in the presence of glucose, possibly by interfering with the proton gradient of the cell membrane, which is required for removal of TJs from the cell surface by endocytosis.

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Although degradation was inhibited by the metabolic inhibitors, after long incubation in the presence of the inhibitors, some remodeling of TJs did occur. Thus, small patches of TJ strands were observed in the cell membranes and parallel or concentric arrangements of TJ strands were formed. A relatively high number of intracellular vesicular structures with TJ remnants were apparent several hours after the induction. This indicated that some degradation did occur even in the presence of the inhibitor. It seems that the intracellular processing of endocytosed TJ remnants proceeded with reduced velocity at low levels of ATP, thus leading to the accumulation of partially degraded TJ remnants. It should be noted that these features occurred in HT29 cells only after the induced formation of TJ during the process of their degradation, thus suggesting that endocytosis is at least one, if not the dominant mechanism involved in TJ degradation in HT29 cells.

21.5 TIGHT JUNCTION FORMATION AND CELL DIFFERENTIATION It was clearly demonstrated that proteases induce TJ formation in HT29 cells and that in the absence of the inducers the TJs degrade from the membranes in an energydependent process. The TJs formed were of the fascia occludens type and their formation was not followed by cell polarization nor by formation of the zonula occludens type of TJs. Forskolin and cholera toxin, known to activate adenylate cyclase and therefore to increase the intracellular level of cyclic AMP in intestinal cells, were found to induce the formation of a brush border on apical membranes of HT29 cells and TJs of the zonula occludens type between the cells (Ophir et al., 1995). Although only about 20 to 30% of the cells grown for 7 days (early confluence) responded to forskolin, almost all the cells at late confluence (80 to 90%) formed a brush border on their apical surface when incubated for 20 h with forskolin (Figure 21.10) (Cohen et al., 1999). Formation of both types of TJs, i.e., fascia occludens by proteases and zonula occludens by forskolin, was inhibited by cytochalasin D, in agreement with the growing knowledge of the involvement of perijunctional actin in the structure and function of TJs (Mitic and Anderson, 1998; Stevenson and Keon, 1998). In contrast, agents known to interfere with microtubules did not prevent TJ formation either by trypsin or by forskolin. Moreover, colchicine, nocodazole, and taxol were found to induce differentiation and apoptosis in HT29 cells (Cohen et al., 1999). Differentiation was characterized by flattening of the cells, formation of a brush border on the apical surfaces, and TJs (zonula occludens) between adjacent cells (Figure 21.11). Apoptosis was characterized by detachment of round cells from the cell layer (Figure 21.12), condensation of nuclear DNA, and annexin V binding to cell surfaces. The effect of cytoskeletal interfering agents and of forskolin on brush-border formation in HT29 cells is summarized in Figure 21.13. The effect of colchicine was rapid and incubation of 6 h in the presence of the inhibitor induced brush border on more than 60% of the cells. The effect of microtubule-interfering agents on cell death was also rapid, and after 24 h less than 50% of exponentially growing cells

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FIGURE 21.10 Scanning electron micrographs of HT29 cultured for 21 days and then treated with forskolin (15 µM, 20 h). (A) Epithelium-like cells. ×300. (B) High magnifications. ×1500. (From Cohen, E. et al., J. Cell Sci., 112, 2657. 1999. With permission.)

FIGURE 21.11 Effect of colchicine (10 µM, 20 h) on cells cultured for 21 days. Freezefractured membranes of adjacent cells. ×28,000. Ef, exoplasmic face, pf, protoplasmic face. Bar, 0.5 µm. (From Cohen, E. et al., J. Cell Sci., 112, 2657. 1999. With permission.)

were viable. Undifferentiated HT29 cells were found to possess E-cadherin. Most of it was found to be located at the cytoplasm, using immunofluorescence methods. However, after treatment with either forskolin or colchicine, E-cadherin was expressed mainly in the cell–cell contact areas.

FIGURE 21.12 Scanning electron micrographs of apoptotic cells. Cells were cultured for 21 days and treated with colchicine. Note brush borders in (A) and blebs (B) (10 µM, 20 h). ×1500. (From Cohen, E. et al., J. Cell Sci., 112, 2657. 1999. With permission.)

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FIGURE 21.13 Effect of cytoskeleton-interfering agents and forskolin on brush-border formation. Cells were cultured for 21 days and then treated for 20 h with forskolin (FS), cytochalasin D (CD), a combination of cytochalasin D and forskolin, colchicine (COL), colchicine and forskolin, nocodazole (NOC), and taxol (TAX). Apical surfaces were observed and counted by scanning electron microscopy. C, untreated cells. Concentrations of reagents are indicated in µM. (From Cohen, E. et al., J. Cell Sci., 112, 2657. 1999. With permission.)

All these data show that most HT29 cells have the potential to differentiate in a rapid process. Many molecules were reported to be involved in intracellular signaling pathways that affect cell growth and differentiation and also control assembly and disassembly of cell junctions. These include, among others, tyrosine kinases, protein kinase C, G proteins, phospholipase C, and also the second messenger, cyclic AMP (Balda and Matter, 1998; Mitic and Anderson, 1998). Signaling pathways may also be involved in the effect of microtubule-interfering agents on HT29 cells. Although these agents are known to arrest the cell cycle, it seems that this effect is not sufficient and additional phosphoregulating pathways may be required. Recently, it was reported that microtubule-interfering agents (colchicine, nocodazole, taxol, and others) activated signaling pathways like c-Jun, N-terminal kinase, Ras, and apoptosis signal-regulating kinase (ASK1) in a variety of human cells (Wang et al., 1998). Activation of signaling pathways and various kinases may mediate differentiation and apoptosis also in HT29 cells.

REFERENCES Augeron, C. and Laboisse, C. L. 1984. Emergence of permanently differentiated cell clones in a human colonic cancer cell line in culture after treatment with sodium butyrate, Cancer Res., 44, 3961. Axelrod, D., Koppel, D. E., Schlessinger, J., Elson, E. L., and Webb, W. W. 1976. Mobility measurement by analysis of fluorescence photobleaching recovery kinetics, Biophys. J., 16, 1055.

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Azarnia, R., Dahl, G., and Lowenstein, W. R. 1981. Cell junction and cyclic AMP. III. Promotion of junctional membrane permeability and junctional membrane particles in a junction-deficient cell type, J. Membr. Biol., 63, 133. Bacher, A., Griebl., K., Mackamul, S., Mitreiter, R., Mückter, H., and Ben-Shaul, Y. 1992. Protease inhibitors suppress the formation of tight junctions in gastrointestinal cell lines, Exp. Cell Res., 200, 97. Balda, M. S. and Matter, K. 1998. Tight junctions, J. Cell Sci., 111, 541. Bernard-Perrone, F., Carrere, J., Renaud, W., Moriscot, C., Thoreux, K., Bernard, P., Servin, A., Balas, D., and Senegas-Balas, F. 1998. Pancreatic trypsinogen I expression during cell growth and differentiation of two human colon carcinoma cells, Am. J. Physiol., 274, G1077. Caldwell, R. B., Wade, L. A., and McLaughlin, B. J. A. 1984. A quantitative study of intermembrane changes during cell junctional breakdown in the dystrophic rat retinal pigment epithelium, Exp. Cell Res., 150, 104. Cereijido, M., Valdes, J., Shoshani, L., and Contreras, R. G. 1998. Role of tight junctions in establishing and maintaining cell polarity, Annu. Rev. Physiol., 60, 161. Claude, P. and Goodenough, D. A. 1973. Fracture faces of zonulae occludentes from “tight” and “leaky” epithelia, J. Cell Biol., 58, 390. Cochand-Priollet, B., Raison, D., Molinie, V., Guillausseau, P. J., Wassef, M., and Bouchaud, C. 1998. Altered gap and tight junctions in human thyroid oncocytic tumors: a study of 8 cases by freeze-fracture, Ultrastruct. Pathol., 22, 413. Cohen, E., Talmon, A., Faff, O., Bacher, A., and Ben-Shaul, Y. 1985. Formation of tight junctions in epithelial cells. 1. Induction by proteases in human colon carcinoma cell line, Exp. Cell Res., 156, 103. Cohen, E., Ophir, I., Henis, Y. I., Bacher, A., and Ben-Shaul, Y. 1990. Effect of temperature on assembly of tight junctions and on the mobility of lipids in membranes of HT29 cells, J. Cell Sci., 97, 119. Cohen, E., Ophir, I., and Ben-Shaul, Y. 1999. Induced differentiation in HT29, a human colon adenocarcinoma cell line, J. Cell Sci., 112, 2657. Dale, B., Gualtieri, R., Talevi, R., Tosti, E., Santella, L., and Elder, K. 1991. Intercellular communication in early human embryo, Mol. Reprod. Dev., 29, 22. Decker, R. S. 1981. Disassembly of the zonula occludens during amphibian neurulation, Dev. Biol., 81, 12. Ducibella, T., Albertini, D. F., Anderson, E., and Biggers, J. D. 1975. The preimplantation mammalian embryo: characterization of intercellular junctions and their appearance during development, Dev. Biol., 45, 231. Elias, P. M. and Friend, D. S. 1976. Vitamin-A-induced mucous metaplasia. An in vitro system for modulating tight and gap junction differentiation, J. Cell Biol., 68, 188. Faff, O., Cohen, E., Bacher, A., and Ben-Shaul., Y. 1987. Protease-induced formation of focal tight junctions in HT29 adenocarcinoma cells does not require extracellular calcium, Biochem. Biophys. Acta, 905, 48. Faff, O., Mitreiter, R., Muckter, H., Ben-Shaul, Y., and Bacher, A. 1988. Rapid formation of tight junctions in HT29 human adenocarcinoma cells by hypertonic salt solutions, Exp. Cell Res., 177, 60. Fantini, J., Rognoni, J. B., Verrier, B., Lehmann, M., Roccabianca, M., Meauchamp, J., and Marvaldi, J. 1990. Suramin-treated HT29-D4 cells grown in the presence of glucose in permeable culture chambers form electrically active epithelial monolayers. A comparative study with HT29-D4 cells grown in the absence of glucose, Eur. J. Cell Biol., 51, 110. Farquhar, M. G. and Palade, G. E. 1963. Junctional complexes in various epithelia, J. Cell Biol., 17, 375.

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Fleming, T. P., Javed, Q., and Hay, M. 1992. Epithelial differentiation and intercellular junction formation in the mouse early embryo, Development, Suppl., 105. Fogh, J. and Trempe, G. 1975. New human tumor cell lines, in Human Tumor Cells “In vitro”, Fogh, J., Ed., Plenum Press, New York, 115. Fujimoto, K. 1995. Freeze-fracture replica electron microscopy combined with SDS digestion for cytochemical labeling of integral membrane proteins. Application to the immunogold labeling of intercellular junctional complexes, J. Cell Sci., 108, 3443. Gonzalez-Mariscal, L., Chavez De Ramirez, B., and Cereijido, M. 1985. Effect of temperature on occluding junctions of monolayers of epithelial cells (MDCK), J. Membr. Biol., 79, 175. Goodenough, D. A. 1999. Plugging the leaks, Proc. Natl. Acad. Sci. U.S.A., 96, 319. Griepp, E. B., Dolan, W. J., Robbins, E. S., and Sabatini, D. D. 1983. Participation of plasma membrane proteins in the formation of tight junctions by cultured epithelial cells, J. Cell Biol., 96, 693. Gumbiner, B. 1987. The structure, biochemistry and assembly of epithelial tight junctions, Am. J. Physiol., 253, C749. Kachar, B. and Pinto da Silva, P. 1981. Rapid massive assembly of tight junction strands, Science Wash., 213, 541. Keller, G., Ben-Shaul, Y., and Bacher, A. 1992. Influence of metabolic inhibitors on the degradation of tight junctions in HT29 cells, Exp. Cell Res., 200, 16. Kirkland, S. C. 1988. Polarity and differentiation of human rectal adenocarcinoma cells in suspension and collagen gel cultures, J. Cell Sci., 91, 615. Le Bivic, A., Hirn, M., and Reggio, H. 1988. HT-29 cells are an in vitro model for the generation of cell polarity in epithelia during embryonic differentiation, Proc. Natl. Acad. Sci. U.S.A., 85, 136. Lynch, R. D., Tkachuk-Ross, L. J., McCormack, J. M., McCarthy, K. M., Rogers, R. A., and Schneeberger, E. E. 1995. Basolateral but not apical application of protease results in a rapid use of transepithelial electric resistance and formation of aberrant tight junctional strands in MDCK cells, Eur. J. Cell Biol., 66, 257. Madara, J. L. 1983. Increases in guinea pig small intestine transepithelial resistance induced by osmotic loads are accompanied by rapid alterations in absorptive-cell tight-junction structure, J. Cell Biol., 97, 125. Madara, J. L. 1990. Maintenance of macromolecular barrier at cell extrusion sites in intestinal epithelium: physiological rearrangement of tight junctions. J. Membr. Biol., 116, 177. Madara, J. L. 1998. Regulation of the movement of solutes across tight junctions, Annu. Rev. Physiol., 60, 143. Merzdorf, C. S., Chen, Y. H., and Goodenough, D. A. 1998. Formation of functional tight junctions in Xenopus embryos, Dev. Biol., 195, 187. Meyer, R. A. and Overton, J. 1983. Changes in intercellular junctions. I. Embryonic chick liver development, Dev. Biol., 99, 172. Mitic, L. L. and Anderson, J. M. 1998. Molecular architecture of tight junctions. Annu. Rev. Physiol., 60, 121. Muckter, H., Ben-Shaul., Y., and Bacher, A. 1987. ATP requirement for induced tight junction formation in HT29 adenocarcinoma cells, Eur. J. Cell Biol., 44, 258. Ophir, I., Cohen, E., Bacher, A., and Ben-Shaul, Y. 1989. Effect of protein synthesis inhibitors on formation and degradation of tight junctions in HT29 adenocarcinoma cells, Eur. J. Cell Biol., 49, 116. Ophir, I., Cohen, E., and Ben-Shaul, Y. 1995. Apical polarity in human colon carcinoma cell lines, Tissue Cell, 27, 659.

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Orci, L., Amberdt, M., Henquin, J. C., Lambert, A. E., Unger, R. H., and Reinhold, A. E. 1973. Pronase effect on pancreatic beta cells secretion and morphology, Science, 180, 647. Pinto, M., Appay, M. D., Simon-Assmann, P., Chevalier, G., Dracopoli, N., Fogh, J., and Zweibaum, A. 1982. Enterocytic differentiation of cultured human colon cancer cells by replacement of glucose by galactose in the medium, Biol. Cell, 44, 193. Pitelka, D. R., Hamamoto, S. T., and Taggart, B. N. 1980. Epithelial cell junctions in primary and metastatic mammary tumors of mice, Cancer Res., 40, 1588. Polak-Charcon, S. and Ben-Shaul, Y. 1979. Degradation of tight junctions in HT29, a human colon adenocarcinoma cell line, J. Cell Sci., 35, 393. Polak-Charcon, S., Shoham, J., and Ben-Shaul, Y. 1978. Junction formation in trypsinized cells of human adenocarcinoma cell line, Exp. Cell Res., 116, 1. Polak-Charcon, S., Shoham, J., and Ben-Shaul, Y. 1980. Tight junctions in epithelial cells of human embryonic hindgut, normal colon and colon adenocarcinoma, J. Natl. Cancer Inst., 65, 53. Polak-Charcon, S., Hekmati, M., and Ben-Shaul, Y. 1989. The effect of modifying the culture medium on cell polarity in a human colon carcinoma cell line, Cell Differ. Dev., 26, 119. Revel, J. P., Yip, P., and Chang, L. L. 1973. Cell junctions in the early chick embryo. A freezeetch study, Dev. Biol., 35, 302. Risinger, M. A. and Larsen, W. J. 1983. Interaction of filipin with junctional membrane at different stages of the junction’s life history, Tissue Cell, 15, 1. Schneeberger, E. E., Walters, D. V., and Olver, R. E. 1978. Development of intercellular junctions in the pulmonary epithelium of the foetal lamb, J. Cell Sci., 32, 307. Scudamore, C. L., Jepson, M. A., Hirst, B. H., and Miller, H. D. 1998. The rat mucosal mast cell chymase, RMCP-II, alters epithelial cell monolayer permeability in association with altered distribution of the tight junction proteins ZO-1 and occludin, Eur. J. Cell Biol., 75, 321. Shimono, M. and Clementi, F. 1977. Intercellular junctions of oral epithelium. II. Ultrastructural changes in rat buccal epithelium induced by trypsin digestion, J. Ultrastruct. Res., 59, 101. Staehelin, L. A. 1974. Structure and function of intercellular junctions, Int. Rev. Cytol., 39, 191. Stevenson, B. R. and Keon, B. H. 1998. The tight junction: morphology to molecules, Annu. Rev. Dev. Biol., 14, 89. Stevenson, B. R., Siliciano, J. D., Mooseker, M. S., and Goodenough, D. A. 1986. Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia, J. Cell Biol., 103, 755. Talmon, A. and Ben-Shaul, Y. 1979. Tight junctions of dissociated and reaggregated embryonic lung cells, Cell Differ., 8, 437. Talmon, A., Cohen, E., Bacher, A., and Ben-Shaul, Y. 1984. Separation of induction and expression of tight junction formation mediated by proteinases, Biochem. Biophys. Acta, 769, 505. Trugnan, G., Rousset, M., Chantret, I., Barbat, A., and Zweibaum, A. 1987. The posttranslational processing of sucrase-isomaltase in HT-29 cells is a function of their state of enterocytic differentiation, J. Cell Biol., 104, 1199. Tsukita, S. and Furuse, M. 1999. Occludin and claudins in tight-junction strands: leading or supporting players? Trends Cell Biol., 9, 268. Walsh, K. A. 1970. Trypsinogens and trypsin in various species, in Methods in Enzymology, Vol. 19, Colowick, S. P. and Kaplan, N. O., Eds., In chief, Perlman, G. E., and Lorand, L., Eds., Academic Press, New York, 41.

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Wang, A. Z., Ojakian, G. K., and Nelson, W. J. 1990. Steps in the morphogenesis of a polarized epithelium. II. Disassembly and assembly of plasma membrane domains during reversal of epithelial cell polarity in multicellular epithelial (MDCK) cysts, J. Cell Sci., 95, 153. Wang, T. H., Wang, H. S., Ichijo, H., Giannakakou, P., Foster, J. S., Fojo, T., and Wimalasena, J. 1998. Microtubule-interfering agents activate c-Jun N-terminal kinase/stress-activated protein kinase through both ras and apoptosis signal-regulating kinase pathways, J. Biol. Chem., 273, 4928. Washabaugh, M. W. and Collins, K. D. 1986. The systematic characterization by aqueous column chromatography of solutes which affect protein stability, J. Biol. Chem., 261, 12477. Weinstein, R. S., Merk, F. B., and Alroy, J. 1976. The structure and function of intercellular junctions in cancer, Adv. Cancer Res., 23, 23. Yamamoto, M. and Katoaka, K. 1988. An electron microscope study of the development of the exocrine and endocrine pancreas with special reference to intercellular junctions, Arch. Histol. Cytol., 51, 315. Zweibaum, A., Laburthe, M., Grasset, E., and Louvard, D. 1991. Use of cultured cell lines in studies of intestinal cell differentiation and function, in Handbook of Physiology, Vol. 4, Field, A. and Frizzell, R. A., Eds., American Physiology Society, Bethesda, MD, 223.

22

Claudins Mediate Specific Paracellular Fluxes in Vivo: Paracellin-1 Is Required for Paracellular Mg2+ Flux Keith A. Choate, Yin Lu, and Richard P. Lifton

CONTENTS 22.1 Introduction .................................................................................................483 22.2 Normal Mg2+ Homeostasis .........................................................................484 22.3 Recessive Renal Hypomagnesemia with Hypercalciuria and Nephrocalcinosis .........................................................................................486 22.4 Positional Cloning of the Gene Causing Recessive Renal Hypomagnesemia........................................................................................487 22.5 Paracellin-1 and Paracellular “Channels” ..................................................489 22.6 Conclusion ..................................................................................................491 References..............................................................................................................491

22.1 INTRODUCTION Epithelial layers constitute barriers that maintain distinct compositions of the fluids on their apical and basolateral aspects. Nonetheless, an important role of epithelia in vivo is the selective and regulated flux of electrolytes and solutes from one side to the other. Such functions underlie physiological processes ranging from absorption of electrolytes, nutrients, and water from the gut, to the regulated passage of inflammatory cells from the vasculature into surrounding tissues. There are two pathways for such fluxes across an epithelial layer: the transcellular pathway through cells and the paracellular pathway between them. Although many ion channels, pumps, and exchangers have been cloned and their functions in transcellular flux pathways determined, specific mediators of paracellular conductance have remained unknown. The reasons for this are evident. Although there are many tools and model systems for studying mediators of transcellular flux, these have been lacking for the paracellular pathway. Thus, whereas transcellular conductance mediators can be cloned and their functions measured by in vitro expression in single cells, paracellular conductances can only be reconstituted in the context of an epithelial layer, greatly complicating the development of assays for determinants of paracellular permeability. 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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All evidence points to the tight junction (TJ) as the determinant of paracellular permeability. The TJ is known to constitute the major barrier to paracellular flux. Although the TJ was initially conceived of as an absolute seal, preventing all paracellular flux, careful physiological studies subsequently demonstrated that this is not the case, that different epithelia have greater or lesser paracellular resistance. Moreover, the flux through the paracellular pathway can be highly selective, suggesting that the junction may contain selective pores. Nonetheless, the nature of the molecules that mediate paracellular flux has remained elusive. The kidney is the major arbiter of electrolyte homeostasis and is one organ in which paracellular flux plays a key role. The basic filtering unit of the kidney is the nephron, and in a normal day, the nearly 1 × 106 nephrons in an adult filter over 170 L of plasma (Merlet-Bénichou et al., 1999). From this filtrate, roughly 85 to 99% of the electrolytes and water are reabsorbed. Remarkably, however, the fractional reabsorption of individual electrolytes and water can be independently and precisely increased or reduced depending upon environmental and dietary variation, thereby maintaining nearly constant levels of each electrolyte in plasma. Identification of the in vivo roles of specific mediators of electrolyte reabsorption has been greatly aided by application of genetic approaches. In particular, the investigation of Mendelian traits in which mutation in a single gene has effects on electrolyte homeostasis has provided substantial insight into integrated physiology, identifying gene products that are indispensable for normal homeostasis and whose altered function cannot be compensated for by other gene products. These disorders include those affecting reabsorption and/or secretion of Na+, K+, H+, Ca2+, HCO 3–, Cl–, and water. All of these determinants affect the transcellular pathway. Recent genetic studies of defects in Mg2+ homeostasis have identified a specific gene whose product is required for normal renal paracellular Mg2+ flux, thereby identifying the claudins as molecules that are required for specific paracellular fluxes.

22.2 NORMAL Mg2+ HOMEOSTASIS Mg2+ is the fourth most abundant intracellular cation, playing a key role as cofactor in a wide variety of enzymatic reactions, as well as a structural role in bone. Only 1% of total-body Mg2+ is in the plasma, with levels that normally vary little from 2 mg/dl. Typical dietary intake is 360 mg/day and about 80% of blood Mg2+ is freely filtered by the kidney (Kelepouris and Agus, 1998). Consequently, the maintenance of Mg2+ balance requires the renal reabsorption of about 90% of the filtered load. Unlike the other major biological cations such as sodium, calcium, and potassium, for which there are hormones that directly regulate their levels in the body, there are no such magnesium-regulating hormones known. Instead, magnesium reabsorption appears to be regulated directly at the level of the nephron, predominantly by serum magnesium levels (Sharhegi and Agus, 1982). The major site of renal Mg2+ reabsorption is the thick ascending limb of Henle’s loop (TAL), where 60% of the filtered load of Mg2+ is reabsorbed. The proximal and distal tubules each reabsorb about 15 and 10%, respectively. Importantly, nearly all the Mg2+ reabsorption in the TAL occurs via a paracellular pathway (Quamme, 1997), which is tightly regulated by serum magnesium levels. Renal magnesium

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+

Ca 2+ Mg 2+ Na +

485

Cl-

Na+ 2Cl K+ K+

Na+ ATP

K+

Ca 2+ Mg2+ Na+

FIGURE 22.1 Paracellular reabsorption of Mg2+ and Ca2+ in the renal TAL of Henle. A cell of the TAL is depicted with portions of adjacent cells. A lumen-positive potential is established by the Na+–K+–2Cl– cotransporter, which brings these ions into the cell as chloride leaves basolaterally via CLCNKB and potassium is recycled into the tubular lumen via ROMK. This results in the net reabsorption of two chloride ions for each sodium ion and yields a net lumen-positive potential, which drives paracellular reabsorption of Ca2+, Mg2+, and Na+.

excretion varies from 0.5 to 80% of the filtered load with low or high serum magnesium concentrations, respectively (Steele et al., 1968). The evidence for the paracellular nature of this pathway is twofold. First, magnesium reabsorption requires the generation of a lumen-positive transepithelial potential by active salt transport (Figure 22.1). Inhibition of this active transport in the TAL by pharmacological agents blocks magnesium reabsorption. Second, when active transport is blocked in isolated, perfused tubules, the direction of magnesium flux can be reversed from reabsorption to secretion by changing the orientation of an applied transepithelial potential (Sharhegi and Agus, 1982). Such a flux reversal is the hallmark of a paracellular pathway. Other electrolytes, including nearly 35% of the filtered load of Ca2+, are also reabsorbed in the TAL via a paracellular pathway (DiStefano et al., 1993). Although this paracellular pathway shows high conductance for cations, it is highly impermeable to water (Greger, 1981), revealing specificity and selectivity of this TJ. Factors that increase salt transport in the TAL produce an increased transepithelial potential, thereby increasing Mg2+ reabsorption. Thus, antidiuretic hormone (ADH), glucagon, parathyroid hormone (PTH), calcitonin, and insulin each can increase paracellular magnesium and calcium reabsorption (De Rouffignac and Quamme, 1994). PTH-induced upregulation of calcium and magnesium transport in isolated, perfused cTAL tubules has been shown to increase transepithelial potential by increasing activity of the Na–K–2Cl cotransporter. This effect is seen even when active transport is blocked by furosemide (Wittner et al., 1993), suggesting that PTH can also directly modify paracellular permeability.

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Physiological studies have characterized the regulation of magnesium and calcium reabsorption in the cTAL, by examining the effects of lumenal and serosal/serum concentrations. In vivo microperfusion studies in parathyroidectomized rats (Quamme and Dirks, 1980) demonstrate that with a constant delivered magnesium load in the tubule, increasing serum magnesium levels proportionally and significantly inhibits magnesium reabsorption, increasing fractional excretion from nearly 30% to greater than 90%. Hypermagnesemia also inhibits calcium reabsorption, although to a lesser extent, increasing fractional excretion from 20% to greater than 60%. Hypercalcemia similarly inhibits magnesium reabsorption, increasing its fractional excretion from 22 to 65%, and increasing fractional excretion of calcium from 14 to 42%. Interestingly, hypermagnesemia and hypercalcemia depress magnesium reabsorption more than calcium reabsorption. These studies suggested that magnesium and calcium may share a common paracellular reabsorption pathway in the cTAL. Within the physiological range of lumenal magnesium concentrations, it appears that magnesium and calcium do not compete for access to this pathway, but when concentrations greatly exceeding those seen in hypermagnesemia are delivered, reabsorption of calcium, magnesium, and sodium is inhibited (Quamme, 1982). With superphysiological lumenal concentrations of calcium or magnesium, transepithelial resistance has been demonstrated to rise in isolated, perfused cTAL segments, inhibiting reabsorption of calcium, magnesium, and sodium, suggesting that this inhibition of reabsorption may be due to a decrease in the overall permeability of the paracellular pathway (DiStefano et al., 1988). In physiological states, however, paracellular permeability to magnesium and calcium in the cTAL is regulated by their serum concentrations alone. These findings establish the paracellular nature of Mg2+ reabsorption in the TAL, begging the question of the molecular determinants of the paracellular permeability.

22.3 RECESSIVE RENAL HYPOMAGNESEMIA WITH HYPERCALCIURIA AND NEPHROCALCINOSIS Given the recognition of the important role of paracellular Mg2+ reabsorption in the TAL, an autosomal recessive human disease, renal hypomagnesemia with hypercalciuria and nephrocalcinosis is an interesting entity. Affected individuals have profound renal magnesium wasting, which results in severe hypomagnesemia uncorrected by oral or intravenous supplementation (Benigo et al., 2000). Concurrent hypercalciuria with normocalcemia maintained by high parathyroid hormone levels results in a progressive medullary nephrocalcinosis, a calcification of the renal parenchyma, which culminates in renal failure in most cases. Other clinical features include polydipsia, polyuria, kidney stones, hyperuricemia, and urinary tract infections. Significantly, renal transplantation cures the disease, suggesting that its cause is cellular, rather than hormonal, with the defect lying at the level of the renal tubule. Although the three hallmarks of this disease (renal hypomagnesemia, hypercalciuria, and nephrocalcinosis) are seen in all patients, the age of presentation, disease severity, and presence of other clinical features vary among affected individuals. The age at diagnosis ranges from 6 months to 25 years, and clinical presentations range from mild nephrocalcinosis identified by screening siblings of more severely affected individuals to advanced nephrocalcinosis and hypomagnesemia-induced convulsions

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at an early age (Praga et al., 1995). The magnesium and calcium wasting seen in affected individuals is consistent with a defect in the paracellular pathway of the TAL.

22.4 POSITIONAL CLONING OF THE GENE CAUSING RECESSIVE RENAL HYPOMAGNESEMIA A genetic approach using positional cloning has been employed to identify the molecular basis of recessive renal hypomagnesemia. Patients with the disease and their unaffected siblings were recruited, and all affected subjects had hypomagnesemia due to renal magnesium wasting, hypercalciuria with nephrocalcinosis, and progressive renal insufficiency. None had evidence of salt wasting, ruling out hypomagnesemia resulting from mutation in the sodium chloride cotransporter, the cause of Gitelman’s syndrome, and all patients had elevated parathyroid hormone levels with normal serum calcium before development of renal failure. Consistent with earlier reports of recessive renal hypomagnesemia, all patients had elevated serum uric acid levels. Of 12 index cases, 8 had a history of urinary tract infections, 5 had kidney stones, 3 had polyuria/polydipsia, and 2 had a congenital defect in eye development called coloboma. Autosomal recessive transmission was expected because parents were unaffected in all kindreds, and in ten kindreds affected subjects were the offspring of consanguineous union. Linkage analysis, comparing the inheritance of genetic markers distributed across all the autosomes with the inheritance of the disease in consanguineous kindreds demonstrated that the disease gene lay in a segment of chromosome 3q. Fine mapping of the gene was facilitated by the investigation of a large kindred from Saudi Arabia that included 13 distantly related affected members. This permitted mapping of the disease gene to a 1-cM interval in the human genome (Simon et al., 1999). Physical mapping of the interval with bacterial artificial chromosomes, followed by exon trapping on the spanning clones, revealed the presence of a novel gene in the interval, called paracellin-1. Exon trapping directly isolates transcribed segments from genomic DNA using a vector that identifies gene segments capable of participating in the splicing reaction required for exon joining (Buckler et al., 1991). Northern analysis demonstrated that paracellin-1 is highly expressed in the kidney, but not in other tissues, and nephron-segment-specific RT-PCR further localized its expression to cells of the thick ascending limb of Henle and the distal convoluted tubule. The genomic structure of the gene was determined, permitting resequencing of the gene in affected individuals. This analysis reveals a wide range of mutations in paracellin-1 that are predicted to impair its synthesis or function. For example, a nonsense mutation (R79TERM) causes a premature stop codon, truncating the protein in the first extracellular loop, and a missense mutation (G121R) inserts a charged reside in the third transmembrane domain. The finding of independent mutations that drastically alter the encoded protein, which cosegregate with the disease in families and which are not found in the general population, constitutes proof that mutations in paracellin-1 cause renal hypomagnesemia with hypercalciuria. Computational analysis revealed that paracellin-1 shares sequence and structural homology with the claudin family of TJ proteins (Morita et al., 1999) (Figure 22.2). The 20 members of the claudin family identified thus far have a similar structure

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FIGURE 22.2 Schematic of the paracellin-1 protein. Paracellin-1 shares sequence and structural homology with the claudin family of transmembrane domains. Like the claudins, it has four transmembrane domains, two extracellular loops, and intracellular amino and carboxy termini. It has a unique accumulation of negatively charged residues (black) in the first extracellular loop.

with four transmembrane domains and intracellular amino and carboxy termini. Paracellin-1 displays 14 to 21% amino acid identity with individual members, and phylogenetic analysis shows that paracellin-1 is one of the most distantly related members of the claudin family, with the greatest degree of homology seen in the transmembrane domains and significant divergence seen in the cytoplasmic carboxy terminus. Polyclonal antisera to a unique portion of the carboxy terminus of paracellin-1 used in immunohistochemistry demonstrated that paracellin-1 is localized to the TJs of cells within a subset of renal tubules. Costaining of human kidney sections with anti-Tamm Horsfall Protein (THP), a maker for the TAL, and a paracellin-1 antibody revealed that paracellin-1 expression is restricted specifically to the TAL. Confirmation that paracellin-1 resides in the TJ was obtained by costaining with an antibody to occludin, a TJ protein highly expressed in the TAL (Figure 22.3). Assembly of a series of images collected by confocal microscopy through the TJ demonstrated precise colocalization of paracellin and occludin throughout the junctional complex. These studies demonstrated that paracellin-1 is located within the TJ of cells in the thick ascending limb of Henle, the nephron segment responsible for the majority of renal magnesium handling. Interestingly, deletion of the first four exons of paracellin-1 in Japanese Black cattle has been reported to result in bovine chronic interstitial nephritis with fibrosis (CINF), an autosomal recessive disorder characterized by growth retardation and lethal renal failure before puberty (Hirano et al., 2000). Affected cattle have elongated hooves with a high degree of renal parenchymal inflammatory cell infiltration and fibrosis (Kobayashi et al., 2000). Although the clinical course of renal failure in bovine chronic interstitial nephritis is well documented in the literature, calcium and magnesium values are not, leaving open the question of whether CINF and human recessive renal hypomagnesemia are variants of the same disorder.

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FIGURE 22.3 Immunolocalization of paracellin-1. Frozen human kidney sections were fixed and stained with an affinity-purified rabbit polyclonal antisera to the unique carboxy terminus of paracellin-1 and an antibody to THP, a maker for the TAL of Henle. Confocal microscopy demonstrated that paracellin-1 stains the intercellular junctions of cells in a subset of renal tubules (A), which are identified as TAL of Henle segments by THP labeling (B) and are colocalized in C. This demonstrates that paracellin is localized to the TJ and that it is present in the physiologically relevant nephron segment for magnesium and calcium reabsorption.

22.5 PARACELLIN-1 AND PARACELLULAR “CHANNELS” The pathophysiology of recessive renal hypomagnesemia and the mutations seen in paracellin-1 suggested that the invocation of the “junctional pore” theory in the 1970s (Claude, 1978) to explain the physiology of the paracellular pathway might have a molecular correlate, with claudins providing the building blocks for specific paracellular “channels” within the TJ. When paracellin-1 was identified, the claudin family had just been discovered, with only 8 of the currently reported 20 family members identified. Although it had been established that claudins are components of the TJ and are necessary and sufficient to constitute junctional fibrils (Furuse et al., 1998), demonstration of their role in mediating specific paracellular conductances had not been made. The finding that mutations in paracellin-1 cause selective renal divalent cation wasting demonstrated for the first time the specific role of claudin family members in selective paracellular flux. Knowledge that other claudins engage in homotypic or heterotypic interactions across the TJ (Furuse et al., 1999), with the extracellular loops providing adhesion between cells, affords the strong suggestion that these molecules form intercellular pores in the TJ, allowing selective passage of electrolytes and solutes. The diversity of claudins suggests the potential diversity in paracellular permeabilities that might be produced by these molecules via their homotypic or heterotypic interactions. The finding that mutations in paracellin-1 result in recessive renal hypomagnesemia provided the first evidence that a member of the claudin family can generate a selective paracellular ion conductance. Although paracellin-1 shares sequence and structural homology with individual claudin members, examination of its unique features may offer insight into its function. Whereas the majority of claudins have a neutral or small net negative charge in their first extracellular loop (Table 22.1), paracellin-1 has ten negatively charged residues in its first extracellular loop, conferring

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TABLE 22.1 Charge of Extracellular Loops in Claudin Family Members Extracellular Loop 1 Claudin PCLN-1 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 17 18 19 20

(–) residues 10 3 3 3 3 3 3 3 5 3 5 6 6 9 2 3 3 5 4 3

(+) residues 5 3 2 3 3 3 3 3 5 3 4 4 8 6 6 2 7 4 3 3

Extracellular Loop 2 Net –5 0 –1 0 0 0 0 0 0 0 –1 –2 2 –3 4 –1 4 –1 –1 0

(–) residues 2 3 4 3 2 3 3 2 3 3 4 1 2 0 2 3 2 0 2 4

(+) residues 2 2 2 3 2 2 3 1 3 2 2 1 2 2 1 2 3 1 1 2

Net 0 –1 –2 0 0 –1 0 –1 0 –1 –2 0 0 2 –1 –1 1 1 –1 –2

Note: Examination of the amino acid composition of the extracellular loops of the claudin family members reveals that some have an accumulation of charged residues. Paracellin-1 has ten negatively charged residues in its first extracellular loop with a net charge of –5. Eight claudins have a net negative charge in their first extracellular loop, and only three have a net positive charge in the first loop: claudins 12, 14, and 17. The majority of claudins have neutral or negatively charged second extracellular loops. Charged loop residues may be important in generating claudin ion permeability and specificity.

a net charge of –5. These negatively charged residues could form a solvated pore either within the extracellular domain or between paracellin-1 molecules in the intercellular space; this charge barrier could contribute to the selectivity for cations. One other family member, claudin-13, has a similarly high density of negative charges, raising the question of whether this claudin might also mediate a paracellular cation flux. Conversely, claudins with net positive loop charge (claudins 12, 14, 17) could allow paracellular permeability to anions. Claudin distribution has been demonstrated to be tissue specific, often limited to a subset of functional epithelial cells (Mitic et al., 2000). Given that claudins can homotypically and heterotypically dimerize, multiple individual claudin pairings within epithelia are possible, each with a potentially different charge selectivity and/or permeability. An important question in this regard remains whether the properties of paracellular pores are determined only by individual claudin molecules or, alternatively, whether they are determined by higher-order interactions.

Claudins Mediate Specific Paracellular Fluxes in Vivo

491

22.6 CONCLUSION The finding that paracellin-1, a member of the claudin family of TJ proteins, is a necessary component of the paracellular pathway mediating Mg2+ reabsorption in the kidney provides the first identification of a molecular determinant of a specific paracellular flux in vivo. This finding has implications for the determinants of other paracellular fluxes. The large number of different claudins that show distinct tissue distributions implies that these different members will be involved in determination of the varied permeabilities of TJs in different tissues. These findings have substantial implications for normal and disease biology. For example, normal or abnormal paracellular permeability is believed to play a role in a wide range of disease states, including paracellular chloride flux in cystic fibrosis, increased epithelial and endothelial permeability in acute respiratory distress syndrome and septic shock, and the passage of inflammatory cells out of the vasculature in a wide range of disease states. The ability to determine the role of specific claudins in vivo by targeted disruption of their corresponding genes in animal models, as well as ability to reconstitute epithelia with TJs expressing specific claudins in vitro, will provide new insight into the role of individual claudins in the mediation of specific paracellular fluxes. In addition to providing new insight into this important area of biology, these studies may identify new targets for therapeutic intervention to treat disease states mediated by abnormal paracellular fluxes as well as opportunities for selective drug delivery.

REFERENCES Benigno, V., Canonica, C.S., Bettinelli, A., von Vigier, R.O., Truttmann, A.C., and Bianchetti MG. 2000. Hypomagnesaemia-hypercalciuria-nephrocalcinosis: a report of nine cases and a review. Nephrol. Dialysis Transplant., 15(5), 605–610. Buckler, A.J., Chang, D.D., Graw, S.L., Brook, J.D., Haber, D.A., Sharp, P.A., and Housman, D.E. 1991. Exon amplification: a strategy to isolate mammaliangenes based on RNA splicing. Proc. Natl. Acad. Sci. U.S.A., 88, 4005. Claude, P. 1978. Morphological factors influencing transepithelial permeability: a model for the resistance of the zonula occludens. J. Membr. Biol., 39, 219–232. De Rouffignac, C. and Quamme, G. 1994. Renal magnesium handling and its hormonal control. Physiol. Rev., 74, 305–322. DiStefano, A., Wittner, M., Gebler, B., and Greger, R. 1988. Increased Ca2+ or Mg2+ concentration rescues relative tight junction permeability to Na+ in the cortical thick ascending limb of Henle’s loop of rabbit kidney. Renal Physiol. Biochem., 11, 70–79. DiStefano, A., Roinel, N., deRouffignac, C., and Wittner, M. 1993. Transepithelial Ca2+ and Mg2+ transport in the cortical thick ascending limb of Henle’s loop of the mouse is a voltage-dependent process. Renal Physiol. Biochem., 16, 157–166. Furuse, M., Sasaki, H., Fujimoto, K., and Tsukita, S. 1998. A single gene product, claudin1 or -2, reconstitutes tight junction strands and recruits occludin in fibroblasts. J. Cell Biol., 143(2), 391–401. Furuse, M., Sasaki, H., and Tsukita, S. 1999. Manner of interaction of heterogenous claudin species within and between tight junction strands. J. Cell Biol., 147(4), 891–903. Greger, R. 1981. Cation selectivity of the isolated perfused cortical thick ascending limb of Henle’s loop of rabbit kidney. Pflügers Arch., 390, 30–37.

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Hirano, T., Kobayashi, N., Itoh, T., Takasuga, A., Nakamaru, T., Hirotsune, S., and Sugimoto, Y. 2000. Null mutation of PCLN-1/claudin-16 results in bovine chronic interstitial nephritis. Genome Res., 10(5), 659–63. Kelepouris, E. and Agus, Z.S. 1998. Hypomagnesemia: renal magnesium handling. Semin. Nephrol., 18(1), 58–73. Kobayashi, N., Hirano, T., Maruyama, S., Matsuno, H., Mukoujima, K., Morimoto, H., Noike, H., Tomimatsu, H., Hara, K., Itoh, T., Imakawa, K., Nakayama, H., Nakamaru, T., and Sugimoto, Y. 2000 Genetic mapping of a locus associated with bovine chronic interstitial nephritis to chromosome 1. Anim. Genet., 31(2), 91–95. Merlet-Bénichou, C., Gilbert, T., Vilar, J., Moreau, E., Freund, N., Lelièvre-Pégorier, M. 1999. Nephron number: variability is the rule: causes and consequences. Lab Invest., 79(5), 515. Mitic, L.L., Van Itallie, C.M., and Anderson, J.M. 2000. Molecular physiology and pathophysiology of tight junctions I. Tight junction structure and function: lessons from mutant animals and proteins. Am. J. Physiol. Gastrointest. Liver Physiol., 279(2), G250–254. Morita, K., Furuse, M., Fujimoto, K., and Tsukita, S. 1999. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc. Natl. Acad. Sci. U.S.A., 96, 511–516. Praga, M., Vara, J., Gonzalez-Parra, E., Andres, A., Alamo, C., Araque, A., Ortiz, A., and Rodicio, J.L. 1995. Familial hypomagnesemia with hypercalciuria and nephrocalcinosis. Kidney Int., 47(5), 1419–1425. Quamme, G.A. 1982. Effect of hypercalcemia on renal tubular handling of calcium and magnesium. Can. J. Physiol. Pharmacol., 60, 1275–1280. Quamme, G.A. 1989. Control of magnesium transport in the thick ascending limb. Am. J. Physiol., 256(2), F197–210. Quamme, G.A. 1997. Renal magnesium handling: new insights in understanding old problems. Kidney Int., 52, 1180–1195. Quamme, G.A. and Dirks, J.H. 1980. Magnesium transport in the nephron. Am. J. Physiol., 239(5), F393–401. Shareghi, R. and Agus, Z.S. 1982. Magnesium transport in the cortical thick ascending limb of Henle’s loop of the rabbit. J. Clin. Invest., 69, 759–769. Simon, D.B., Lu, Y., Choate, K.A., Velazquez, H., Al-Sabban, E., Praga, M., Casari, G., Bettinelli, A., Colussi, G., Rodriguez-Soriano, J., McCredie, D., Milford, D., Sanjad, S., and Lifton, R.P. 1999. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science, 285, 103–106. Steele, T. H., Wen, S. F., Evenson, M. A., and Rieselbach, R. E. 1968. The contribution of the chronically diseased kidney to magnesium homeostasis in man. J. Lab. Clin. Med., 71(3), 455–463. Wittner, M., Mandon, B., Roinel, N., deRouffignac, C., and DiStefano, A. 1993. Hormonal stimulation of Ca2+ and Mg2+ transport in the cortical thick ascending limb of Henle’s loop of the mouse: evidence for a change in the paracellular pathway permeability. Pflügers Arch., 423, 387–396.

23

Microbial Pathogens That Affect Tight Junctions Gail Hecht

CONTENTS 23.1 Introduction .................................................................................................493 23.2 Microbial Disruption of Tight Junctions via Targeting of the Cytoskeleton................................................................................................495 23.2.1 Clostridium difficile Toxins ...........................................................495 23.2.2 Clostridium botulinim Toxin .........................................................496 23.2.3 Bacteroides fragilis........................................................................496 23.2.4 Entamoeba histolytica ...................................................................498 23.3 Disruption of Tight Junctions by Bacterial Proteases................................498 23.3.1 Pseudomonas aeruginosa ..............................................................498 23.3.2 Porphyromonas gingivalis .............................................................499 23.3.3 Vibrio cholera ................................................................................501 23.4 Disruption of the Tight Junction Barrier by Pathogen-Stimulated Signaling Events .........................................................................................503 23.4.1 Vibrio cholera ................................................................................503 23.4.2 Pathogenic Escherichia coli ..........................................................504 23.4.3 Helicobacter pylori........................................................................507 23.4.4 Rotavirus ........................................................................................508 23.5 Access to Basolateral Ligands by Pathogen-Induced Alterations in Tight Junctions............................................................................................509 23.5.1 Yersinia pseudotuberculosis...........................................................509 References..............................................................................................................510

23.1 INTRODUCTION The simultaneous explosion of information regarding both host–pathogen interactions and tight junction (TJ) structure–function dynamics has resulted in the emergence of a new literature concerning the effects of microbes on these physiologically important epithelial structures. Initial investigations in this area were limited by the paucity of information regarding the composition of TJs. Thus, the level of exploration was restricted to correlating functional end points, such as transepithelial electrical resistance (TER) and paracellular solute flux, with morphological changes 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

493

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FIGURE 23.1 Regulation of the TJ barrier. There are two general regulatory arms that control TJ permeability: the cytoskeleton and TJ proteins. Most likely, direct and indirect interactions occur between these two arms. Signaling events may affect either regulatory pathway while proteases that initially attack specific TJ proteins may subsequently trigger changes in the cytoskeleton. The various levels of interaction have yet to be defined.

as determined by electron microscopy or freeze fracture. As the TJ has become better defined, however, studies concerning the effects of microbial pathogens on specific TJ proteins have, in some cases, begun to identify potentially new pathogenic mechanisms. Although the regulation of the TJ barrier is far from being completely understood, there is evidence to support two general arms of regulation (Figure 23.1). Cytoskeletal-mediated events define one arm and alterations of specific TJ proteins the other. Theoretically, cytoskeletal regulation may occur through direct or indirect processes. Direct regulation of the TJ barrier by the cytoskeleton is best represented by contraction of the perijunctional actomyosin ring, which immediately underlies the TJ (discussed in Chapters 12 and 15). This event appears to be controlled by phosphorylation of the 20-kDa myosin light-chain (MLC20) (Hecht et al., 1996; Turner et al., 1997). Contraction of this ring exerts tension on the cell membrane, which is transmitted upward to the TJ thus attenuating barrier function. Indirect cytoskeletal-mediated regulation of the TJ could theoretically occur as a result of the numerous interactions between specific TJ proteins, such as ZO-1, ZO-2, ZO-3, cingulin, occludin, and claudins with actin (Fanning et al., 1998; Haskins et al., 1998; Itoh et al., 1999; Wittchen et al., 1999) and/or myosin (Cordenonsi et al., 1999). Alternatively, direct effects on TJ-associated proteins may also alter the barrier (Balda et al., 1996; McCarthy et al., 1996; Wong and Gumbiner 1997; Bamforth et al., 1999; Sonoda et al., 1999). Because of the recent conception of this field, the mechanisms by which pathogens perturb the TJ barrier are ill-defined. For this reason, discussions will be divided into broad categories including cytoskeletal mediation of the TJ barrier by microbes; effects of bacterial proteases on TJs; pathogen-induced signaling pathways that alter TJ permeability; and exploitation of altered polarity by microbial disruption of TJs.

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FIGURE 23.2 Cytoskeletal-mediated disruption of TJs by microbial pathogens. The enteric pathogens C. difficile and C. botulinum produce enterotoxins, toxins A and B, and C3 transferase, respectively, that by inactivating Rho GTPases disrupt the actin cytoskeleton. As a result TJ proteins and permeability are altered.

23.2 MICROBIAL DISRUPTION OF TIGHT JUNCTIONS VIA TARGETING OF THE CYTOSKELETON 23.2.1 CLOSTRIDIUM

DIFFICILE

TOXINS

The anaerobic bacterium responsible for antibiotic-associated pseudomembranous colitis, Clostridium difficile, produces two exotoxins, toxin A and toxin B. The target of these toxins is the actin cytoskeleton through the disruption of Rho GTP-binding proteins including Rho, Rac, and Cdc42 (Just et al., 1994; 1995; Dillon et al., 1995). Toxins A and B both glucosylate Rho GTPases at threonine 37/35 using UDP-glucose as a cosubstrate, thus inactivating this enzyme (Schmidt and Aktories, 1998) and depolymerizing actin filaments. Both toxins have been shown to enhance the paracellular permeability of model host intestinal epithelia in a dose- and time-dependent manner, an event that correlates with disruption of the actomyosin perijunctional ring (Hecht et al., 1988; 1992) as depicted in Figure 23.2. Specifically, exposure of T84 intestinal epithelial monolayers to toxin A (Hecht et al., 1988) or toxin B (Hecht et al., 1992) was shown rapidly to decrease the transepithelial electrical resistance and enhance the flux of paracellular markers including mannitol and raffinose. This early study (Hecht et al., 1988) was the first demonstration that enteric pathogens and/or their toxins could influence the TJ barrier. Although these initial findings were made using an in vitro model, subsequent confirmation was reported in mammalian, including human, intestinal tissues (Moore et al., 1990; Riegler et al., 1995). Whether C. difficile exerts its effects on TJ permeability solely through the disruption

496

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of actin is not known, as the effects of these toxins on specific TJ proteins have not been examined. It is quite possible, however, that such alterations could occur in view of the many interactions between the actin cytoskeleton and TJ proteins (Fanning et al., 1998; Haskins et al., 1998; Itoh et al., 1999; Wittchen et al., 1999; Cordenonsi et al., 1999). With the recent identification of the TJ transmembrane proteins occludin (Furuse et al., 1993) and claudins (Furuse et al., 1998), further investigations into this area will more thoroughly define the effects of toxins A and B on TJs.

23.2.2 CLOSTRIDIUM

BOTULINIM

TOXIN

Another clostridial toxin, Clostridium botulinim toxin C3 transferase, which also targets the actin cytoskeleton through its effects on Rho (Schmidt and Aktories, 1998), has been shown to perturb the TJ barrier (Figure 23.2) (Nusrat et al., 1995). These experiments were cleverly executed by employing a chimeric toxin in which the C3 gene was fused with that encoding the cell-binding B domain, but not the active toxin domain, of diptheria toxin (DC3B). In this way, efficient delivery of C3 exoenzyme into cells was ensured. C3 transferase ADP-ribosylates Rho GTPases, but not Rac and Cdc42, thereby inhibiting its biological activity. As a result, actin filaments are disassembled as occurs in response to C. difficile toxins. When delivered into polarized intestinal epithelial T84 cells, actin comprising the apically situated perijunctional ring, but not the basal stress fibers, was disassembled. Correlating with this morphological alteration was a dose- and time-dependent perturbation of the TJ barrier, as reflected by a decrease in TER and a corresponding increase in the flux of 10-kDa dextran. While the adherens junction protein E-cadherin was unaffected by C3 transferase, the TJ protein ZO-1 became dissociated from its resident structure. This sentinel study (Nusrat et al., 1995) was the first to demonstrate that a toxin whose primary target is the actin cytoskeleton ultimately exerts effects on a TJ-associated protein, presumably as a result of the numerous interactions between actin and TJ proteins. Undoubtedly, additional investigations will establish this as a common paradigm. Conversely, other enteric pathogens, such as rotavirus (discussed below), appear independently to alter the actin cytoskeleton and TJ proteins (Obert et al., 2000).

23.2.3 BACTEROIDES

FRAGILIS

The production of a 20-kDa enterotoxin by Bacteroides fragilis (BFT), which causes diarrhea, is now recognized to be a zinc-dependent metalloprotease (Moncrief et al., 1995). BFT has been shown to have several cellular effects, including actin rearrangement, stimulation of chloride secretion, and diminution in barrier function when presented to the basolateral aspect of intestinal epithelial monolayers (Chambers et al., 1997). Since BFT is not internalized and requires access to the basolateral aspect of host epithelia to induce these physiological consequences, it was deduced that the substrate of BFT must reside on the basolateral membrane of the cell and possess an extracellular domain. Elegant studies from the laboratory of C. Sears and colleagues have begun to elucidate the mechanisms whereby BFT induces these physiological alterations (Wu et al., 1998). Three key cellular proteins fitting the

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FIGURE 23.3 BFT cleaves E-cadherin leading to downstream effects on TJs. BFT, a zinc metalloprotease, cleaves E-cadherin at an extracellular site near its transmembrane domain. In an energy-dependent fashion, cellular proteases then degrade intracellular E-cadherin, disrupting the interactions between catenins and F-actin. The alterations in the actin cytoskeleton are believed to be responsible for the subsequent redistribution of the TJ proteins ZO-1 and occludin. As such, TJ barrier function is perturbed. (Modified from Wu, S. et al., Proc. Natl. Acad. Sci. U.S.A., 95:14979–14984, 1998. With permission.)

above criteria were strategically selected for investigation: β1-integrin, E-cadherin, and occludin. Exposure of human intestinal epithelial HT29/C1 cells to BFT induced the cleavage of E-cadherin, but not β1-integrin or occludin. Specifically, degradation of the 120-kDa intact E-cadherin to 28- and 33-kDa fragments was seen. Cleavage is believed to occur near the plasma membrane (Figure 23.3), similar to that induced by Porphyromonas gingivalis (Katz et al., 2000) discussed below, since the molecular weights of the extracellular and transmembrane domains combined approximate 30 kDa. Although this initial cleavage step is ATP independent, subsequent degradation of the 28- to 33-kDa E-cadherin fragments was found to be dependent upon ATP and cellular proteases. Associated with the intracellular cleave of E-cadherin was the redistribution of ZO-1 and occludin. Following a 1-h exposure to 5 nM BFT, ZO-1 and occludin assumed a punctate distribution, as compared with uniform staining seen in control monolayers. Consistent with its intracellular location, no evidence of ZO-1 proteolysis could be demonstrated. It is speculated, therefore, that degradation of the intracellular domain of E-cadherin disrupts interactions with β-catenin, leading ultimately to changes in actin. Perturbation of the actin cytoskeleton may then be responsible for the altered cell morphology and disruption of the TJ barrier (see Figure 23.3). The possibility that initial effects of BFT on the adherens junction may stimulate signaling pathways that subsequently influence TJ permeability, however, cannot be ruled out.

498

23.2.4 ENTAMOEBA

Tight Junctions HISTOLYTICA

Although the characteristic effect of Entamoeba histolytica infection is host cell lysis (hence the name), previous studies have reported that, prior to this effect, trophozoites of E. histolytica rapidly diminish the TER of intestinal epithelial monolayers suggesting disruption of TJ complexes (Li et al., 1994; Leroy et al., 1997). Only recently, however, have the effects of this parasite on TJ proteins per se been explored (Leroy et al., 2000). Only apical infection with live trophozoites decreased TER in an inoculum- and time-dependent fashion. An inoculum of 2.5 × 104 organisms significantly diminished TER by 1 h postinfection, although lesser changes were detected as early as 15 to 30 min. This physiological alteration preceded the lysis of cell monolayers, as determined by morphological analysis and the release of 51Cr. Instead, alterations in specific TJ-associated proteins were observed. ZO-1/ZO-2 interactions were disrupted as determined by coimmunoprecipation; ZO-1 was degraded and less so ZO-2 (32%); and ZO-2 was dephosphorylated. Interestingly, no evidence of occludin or E-cadherin degradation could be detected and immunofluorescent staining for ZO-1, ZO-2, occludin, and cingulin showed no redistribution. The intensity of ZO-1 staining, however, appeared less intense. Live organisms were required to induce this effect as sonicated trophozoites did not alter TER suggesting that proteases were not responsible. In fact, a battery of protease inhibitors had no protective influence. Instead, two alternative mechanisms were offered. First, attachment of trophozoites to the membrane of host cells disrupts the apical actin cytoskeleton (Li et al., 1994), which may subsequently influence the TJ barrier. Second, signal transduction events may be responsible, although inhibitors of tyrosine kinases, phosphotidylinositol-3 kinase, or trimeric G proteins did not reverse the effect. Alternatively, E. histolytica trophozoites transfer a 170-kDa subunit of Gas/GalNAc-specific lectin to host cells at cell–cell borders, specifically at the area of the zonula adherens rather than the TJ. It is suggested that insertion of this lectin into the membrane may alter the distribution of proteins in sphingolipid–cholesterol rafts. In fact, Nusrat et al. (2000) have recently demonstrated that perturbation of the TJ barrier by chelating calcium with EGTA not only obliterates the TJ barrier but also shifts the distribution of TJ proteins into different membrane fractions. Certainly, this newly recognized paradigm will be investigated in the future with respect to host–pathogen interactions and TJs.

23.3 DISRUPTION OF TIGHT JUNCTIONS BY BACTERIAL PROTEASES Since the identification of the transmembrane, TJ-associated proteins occludin and claudins, recent efforts have focused on the effects of specific microbial pathogens on these molecules. One theme that has emerged as a result of these studies is disruption of TJs as a result of protein cleavage by bacterial proteases.

23.3.1 PSEUDOMONAS

AERUGINOSA

One of the first bacterial proteases reported to cleave TJ proteins was a product of Pseudomonas aeruginosa (Azghani, 1996). Pseudomonas aeruginosa is a frequent cause of pneumonia in the immunocompromised host, in particular those with cystic

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fibrosis. This microbial pathogen has devised ways to breach the epithelial lining of airways and destroy the underlying parenchyma. Using both animal (Azghani et al., 1990) and in vitro (Azghani et al., 1993) models, Azghani provided evidence that, in part, this occurs via the disruption of TJs. Here, P. aeruginosa protease was found to degrade ZO-1, an event that correlated with an increase in the permeability of the lung to albumin. A more recent study (Azghani, 1996) identified two virulence factors, (1) elastase, a zinc metalloprotease, and (2) exotoxin A, which are responsible for this physiological perturbation. Here, either primary isolates of type II pneumocytes or cultured Madin–Darby canine kidney (MDCK) cells were exposed to Pseudomonas elastase (PE) or exotoxin A (ExoA). Interestingly, PE rapidly (90 min) decreased TER and increased the flux of mannitol in a dose-dependent, nonpolar fashion. A shift of ZO-1 from the peripheral membrane to the cytoplasm was also observed with immunoblot analysis showing a dose-dependent decrease in ZO-1 and ZO-2. ExoA, on the other hand, disrupted paracellular permeability only when presented to the basolateral aspect of cultured epithelial cells and after prolonged (16-h) exposure. ZO-1 remained membrane-associated, but the intensity of the staining was diminished correlating with a decreased, but not ablated, signal for ZO-1 and ZO-2 by immunoblot analysis. Although the specific effects of PE and ExoA on ZO-1 and ZO-2 have not been defined, the physiological consequences can be extrapolated to host outcome. To exert its full impact on the host, P. aeruginosa must breach an intact epithelial barrier. Exposure of monolayers to either PE or ExoA enhanced the epithelial translocation of this pathogen, an event that correlated with the degree of TER reduction, suggesting that some movement through the paracellular space occurs. Protease inactivation by specific antibodies or heat attenuated bacterial penetration. The specificity of this response was revealed by the lack of similar effects in response to a nonbacterial protease, neutrophil elastase. The theoretical impact of such pathogen-induced alterations on the host (depicted in Figure 23.4) is that brief exposure of airway epithelia to PE opens the paracellular pathway, allowing bacteria to access the basal aspect of those cells and underlying tissues. Basal contact with ExoA further disrupts paracellular permeability and inhibits protein synthesis (Iglewski and Sadoff, 1979), thus diminishing the capacity of the host to restore barrier function and enhancing the opportunity for the establishment of infection. Although not demonstrated in this particular study, Reinecker and colleagues (Kinugasa et al., 2000) have recently shown that the expression of claudin-1 and claudin-2 is upregulated by interleukin-17 (IL-17), a finding that supports the premise that pathogen-induced cytokines may trigger host responses that would ultimately restore TJ integrity.

23.3.2 PORPHYROMONAS

GINGIVALIS

Another organism that appears to disrupt TJs to penetrate a host epithelial lining is Porphyromonas gingivalis. This pathogen, a Gram-negative bacterium, has acquired mechanisms by which it can penetrate the protective epithelial lining of the oral cavity, gain access to underlying tissues, and thus cause adult periodontal diseases. This mechanism, which is similar to that described for P. aeruginosa as discussed

500

Tight Junctions

P. aeruginosa

redistributes and degrades ZO-1/ZO-2

Pseudomonas elastase

Exotoxin A

P. gingivalis

increases TJ permeability

occludin ZO-1 ZO-2 ZO-1

ZO-2

ZO-1 ZO-2

E-cadherin

invasion and translocation

§1 integrin basolateral Exotoxin A diminishes ZO-1/ZO-2 release of protease degradation of proteins

FIGURE 23.4 Impact of bacterial proteases on the tight junction barrier. Two bacterial pathogens Pseudomonas aeruginosa and Phorphyromonas gingivalis, which elaborate proteases that degrade TJ proteins, are represented here. Pseudomonas aeruginosa elaborates Pseudomonas elastase, which degrades ZO-1 and ZO-2 when presented to the apical aspect of cells, thus opening the paracellular space. Intact organisms and a second proteolytic product, exotoxin A, can now access the basolateral domain of the cell where exotoxin A exerts its effects. Phorphyromonas gingivalis and its products also require basal positioning to exert effects on host tissue TJs. This is achieved by invasion of and translocation across the epithelial layer. P. gingivalis products, presumably proteases, then degrade not only the TJ protein occludin, but also β1-integrin and E-cadherin. With the subsequent loss of the paracellular barrier, organisms can translocate across this space as well, thus enhancing access to the basal domain of the cells.

above, was elucidated by examining the effect of P. gingivalis on epithelial MDCK TJs (Katz et al., 2000). In contrast to P. aeruginosa PE, however, only basolateral exposure of intact monolayers to P. gingivalis decreased TER and degraded several transmembrane proteins including β1-integrin, occludin, and the zonula adherens protein E-cadherin. Based on the membrane topology of E-cadherin, the specificity of the antibody for recognizing the cytoplasmic domain, and the size of the proteolytic fragments, it was predicted that cleavage occurs within the extracellular domain of the molecule adjacent to the plasma membrane, as has been described for BFT (see above). A secreted bacterial protease is believed to be responsible, but has not been definitively demonstrated, since exposure of purified protein to culture supernatants induced a similar response. Porphyromonas gingivalis likely accesses the basolateral aspect of host cells by invading and translocating across epithelial cells. In fact, the most efficient translocation was found to occur in the apical-basolateral direction. Once the basolateral

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surface is accessed, undefined products, most likely extracellular proteases, are released and degrade cell-adhesion proteins. The resulting disruption of the intercellular barrier then allows paracellular translocation of possibly whole organisms and secreted products (see Figure 23.4). Interestingly, the initial interactions between P. gingivalis and the apical surface of the host membrane evoke a protective increase in barrier function that is overcome once bacterial translocation and basolaterally mediated effects on key host proteins ensue. The specific enzymes responsible for these events and mechanisms underlying the host responses have yet to be defined.

23.3.3 VIBRIO

CHOLERA

Probably the best-defined direct effects of a bacterial protease on TJ proteins are those of the hemagglutinin protease (HA/P) of Vibrio cholera. HA/P is a member of a large family of bacterial metalloproteases that act on several physiological substrates including fibronectin, lactoferrin, and mucin (Hase and Finkelstein, 1993). In addition, it is this protease that activates cholera toxin (CT) by cleaving the A subunit into A1 and A2 (Booth et al., 1984). Interestingly, HA/P also inactivates the CTXΦ, which houses the V. cholera enterotoxins CT, ZOT, and ACE (Kimsey and Waldor, 1998). Initial studies by Wu et al. (1996) demonstrated that the application of HA/P to MDCK monolayers decreased TER and disrupted ZO-1 and the actin cytoskeleton in a dose-dependent manner. Later, these investigations both showed that the apical application of HA/P to epithelial monolayers degrades the transmembrane TJ protein occludin in a dose- and time-dependent manner (Wu et al., 2000). Specifically, MDCK monolayers were treated with culture supernatants from strain CVD110, a mutant of El Tor biotype that expresses active HA/P but not CTA, ZOT, Ace, or hemolysin. The higher-molecular-weight bands of occludin (particularly the 85-kDa form) were degraded to fragments of ∼50 and 35 kDa. Inhibition of bacterial metalloproteases with Zincov prevented the degradation. Immunofluorescent staining using antibody raised against the intracellular E domain of occludin (Figure 23.5) showed that the cytoplasmic portion of this protein remained localized to the cell periphery, suggesting that association of intracellular occludin with the TJ does not depend on intact extracellular domains. This is in agreement with the previous demonstration that the E domain is sufficient for targeting occludin to the TJ (Matter and Balda, 1998). Based on the sizes of the degradation products, the hydrophilicity plot of occludin, and the specificity of the antibody used, it was predicted that the 35-kDa degradation product represents the intracellular E domain plus the membrane-spanning segment M4 (see Figure 23.5). The 50-kDa product is predicted to represent a fragment consisting of M2, C, M3, D, M4, and E (see Figure 23.5). An unrelated protease, trypsin, failed to induce this change. Although ZO-1 is redistributed by HA/P, this morphological alteration could not be attributed to degradation. This phenomenon, however, was also prevented by bacterial metalloprotease inhibition, supporting the possibility that the primary effects of HA/P on occludin may perturb its interaction with ZO-1 via conformational changes that lead to the redistribution of ZO-1. Another tenable scenario is that the HA/P-induced cleavage of occludin perturbs the ZO-1/ZO-2/ZO-3 complex, thus triggering downstream signaling events that ultimately reorganize the actin cytoskeleton.

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FIGURE 23.5 Defined domains of occludin and predicted sites of cleavage by V. cholera HA/P. Occludin has a cytosolic N-terminus tail (A), four membrane-spanning domains (M1 to M4), two extracellular loops (B and D) with an intervening short cytosolic loop (C), and a long C-terminal tail. Evidence suggests the HA/P cleaves occludin at two extracellular sites adjacent to the membrane. Note that the domains are not drawn to scale.

Vilorio cholera has been previously reported to produce another toxin that specifically affects TJs, zonula occludens toxin (ZOT) (Fasano et al., 1991). How the effects of HA/P on TJ proteins relate to the previously described effects of ZOT on TJ structure and function (discussed below) is currently unknown. As predicted on the basis of the above data, however, HA/P was recently reported to disrupt not only TJ structure, but also function (Mel et al., 2000). Here, the impact of select V. cholera mutant strains on the TER of model human intestinal epithelial T84 monolayers was assessed. TER decreased by ∼80% following 3 h of exposure to supernatants from both zot+ and zot– strains, indicating that a factor other than ZOT was responsible. In fact, a strong correlation between HA/P production and decreased TER was demonstrated by creating a hap A deletion mutant. Culture supernatants from the deletion mutant strain had little protease activity and only modestly decreased TER (25%), whereas the activity of supernatants from a hap A complemented strain was indistinguishable from that of wild-type V. cholera. That other proteases were not responsible was evidenced by the lack of activity of 50-foldconcentrated supernatants from hap/A, the deletion mutant strain. Many pathogenic bacteria produce zinc metalloproteases that appear to have specific eukaryotic cell surface targets, for example, occludin, in the case of V. cholera HA/P, and E-cadherin, in the case of B. fragilis enterotoxin (Wu et al., 1998). Such findings suggest a common, but previously unrecognized, mechanism of bacterial pathogenesis. The speculation that perturbation of TJ permeability may

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be a means by which these organisms access nutrients from their host has been offered. In addition, the effects of HA/P on the TJ barrier may account for the residual diarrhea associated with infection by attenuated V. cholera vaccine strains that do not produce CT, ZOT, or ACE. In fact, CT and HA/P utilize the same secretory apparatus and deletion of CT may actually enhance the export of HA/P (Sandkvist et al., 1997). Additionally, strains with core region deletions, including CtxAB, ace, and zot, produce higher levels of HA/P than wild-type strains (Kimsey and Waldor, 1998).

23.4 DISRUPTION OF THE TIGHT JUNCTION BARRIER BY PATHOGEN-STIMULATED SIGNALING EVENTS 23.4.1 VIBRIO

CHOLERA

Prior to the identification of the effects of V. cholera HA/P on TJ proteins and permeability, a second V. cholera enterotoxin, which alters intestinal epithelial TJs, was described. Initial studies showed that the exposure of stripped rabbit ileal mucosa to culture supernatants of wild-type V. cholera strain 395 or CVD101, in which the gene encoding the A1 peptide of CT was deleted, increased conductance (Gt ; Fasano et al., 1991). The penetration of the paracellular space by wheat germ agglutinin–horseradish peroxidase, normally restricted by the TJ, was also seen following exposure to these culture supernatants. Freeze-fracture evaluation of TJs revealed a loss of TJ strands, primarily those oriented perpendicularly to the long axis of the TJ, hence the name zonula occludens toxin, or ZOT. Interestingly, the effects of ZOT appear to be restricted to the small intestine (Fasano et al., 1991; 1997) as colonocytes, neither cultured nor native, were affected. This may reflect the selective distribution of ZOT receptors. Although the mechanisms by which ZOT alters TJ permeability are undefined, protein kinase C (PKC)-induced effects on the actin cytoskeleton appear to be involved (Fasano et al., 1995). Exposure of enterocytes to ZOT enhances actin polymerization and redistributes it to the subcortical area of the cell. Both events correlate temporally with disruption of the TJ barrier and are reversible following the removal of supernatants. Inhibition of calcium-dependent PKC with staurosporine, or a more specific derivative, prevented the effects of ZOT on actin redistribution and attenuated the permeability response, suggesting that the involvement of PKC is proximal to both of these events. While the contribution of all PKC isoforms was not investigated, the activity of membrane-bound PKCα peaked at 1.7-fold following 3 min of ZOT exposure. Although the steps linking PKC activation and actin cytoskeletal rearrangements are not known, it is speculated that myristoylated alanine-rich C kinase substrate (MARCKS) may serve as this intermediate. Nonphosphorylated MARCKS is membrane associated and interacts with submembranous actin (Hartwig et al., 1992). Phosphorylation of MARCKS by PKC causes its dissociation from the membrane, thereby inducing actin rearrangement (Thelen et al., 1991) and possibly increasing TJ permeability. Whether or not there is a relationship between the activities of ZOT and HA/P on TJs has yet to be reconciled. Certainly there are regulatory interactions at the gene level (Kimsey and Waldor 1998). The fact that HA/P alters the permeability

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of T84 monolayers in a ZOT-independent manner is potentially explained by a lack of ZOT receptor expression in this cell line. On the other hand, the HA/P-independent effects of ZOT on native small intestinal tissues have yet to be explored.

23.4.2 PATHOGENIC ESCHERICHIA

COLI

Enteropathogenic Escherichia coli (EPEC) is a most interesting pathogen because despite its noninvasive/nontoxigenic nature it possesses the means to induce dramatic effects on host intestinal epithelial physiology. Certainly, its effects on TJs are the best characterized. Initial studies defining the model of infection of the host target tissue of this pathogen, intestinal epithelial cells, showed that TER was decreased in a time-dependent manner (Canil et al., 1993; Spitz et al., 1995; Philpott et al., 1996). This change was not attributable to cytotoxicity and correlated with the enhanced paracellular movement of mannitol (Spitz et al., 1995). Several earlier publications had demonstrated the impact of EPEC on the phosphorylation of host proteins. Interestingly, in the many different cell systems used, the most prominently phosphorylated band was MLC20 (Manjarrez-Hernandez et al., 1991; 1996). Using a genetic approach, the author and colleagues had previously shown that MLC20 phosphorylation was one mechanism by which TJ permeability could be regulated (Hecht et al., 1996). It has now been convincingly demonstrated that MLC20 phosphorylation provides a physiological means of TJ regulation (Madara and Pappenheimer, 1987; Turner et al., 1997; discussed in Chapter 15). It appears that EPEC exploits this pathway as inhibitors of MLC kinase (MLCK) attenuated the EPECinduced perturbation of the TJ barrier, whereas neither PKC nor tyrosine kinase inhibitors afforded protection against this response. Similarly, inhibition of MLCK has been shown to prevent partially the decrease in TER following infection with enterohemorrhagic E. coli (Philpott et al., 1998). In contrast to data concerning EPEC, however, PKC inhibitors also partially blocked the EHEC-induced resistance response. These findings are particularly interesting in view of the fact that the genes encoding the type III secretory system of EPEC and EHEC, which reside within a pathogenicity island called the locus of enterocyte effacement (LEE), share 98 to 100% (Perna et al., 1998) homology, suggesting that other virulence factors may be involved. The recent identification of TJ transmembrane proteins, such as occludin, that participate in the formation of the barrier (Farshori and Kachar, 1999) has allowed for investigation concerning the direct effect of EPEC on these proteins. As such, the author’s group has recently published data showing that EPEC infection of intestinal epithelial T84 cells causes the dissociation of occludin from TJs apparently by inducing dephosphorylation (Simonovic et al., 2000). The cytoplasmic tail of occludin has several threonine and serine sites available for phosphorylation (Sakakibara et al., 1997). Many investigators have demonstrated that the phosphorylation of occludin appears to be required for its association with the TJ (Sakakibara et al., 1997; Wong, 1997; Farshori and Kachar, 1999). The author’s group found that infection of T84 monolayers caused the redistribution of occludin from its normal localization at the peripheral membrane to a cytoplasmic location following 3 h of infection (Figure 23.6). When analyzed by immunoblot, occludin resolves as several

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FIGURE 23.6 Infection of intestinal epithelial cells with EPEC results in the redistribution of occludin without causing changes in perijunctional actin. Cells were infected with EPEC for the indicated periods of time and then stained for occludin or actin using fluorescent probes. (A) In uninfected control cells, occludin staining is uniform and is primarily localized to the periphery of the cell, indicating association with TJs. (B) After 1 h of infection with EPEC, occludin, although remaining primarily TJ associated, takes on a punctate pattern of staining. (C) At 3 h after EPEC infection, occludin appears to have dissociated from the membrane and relocalized to the cytoplasm. (D and E) In contrast to the dramatic changes seen in occludin localization in response to EPEC infection, filamentous actin within the perijunctional ring in control (D) and EPEC-infected (E) cells is essentially the same. (From Simonovic, I. et al., Cell. Microbiol., 2:305–315, 2000. With permission of Blackwell Science Ltd.)

bands ranging from 80 to 62 kDa depending on the level of phosphorylation. In T84 cells, occludin separates primarily as two bands, a higher-molecular-weight band to which the majority of occludin resolves and a lower-molecular-weight band. Following infection with EPEC, however, there is a progressive shift of occludin to the lower-molecular-weight form (Figure 23.7), suggesting dephosphorylation, which correlates temporally with the decrease in TER. In that EPEC is for the most part noninvasive, monolayers can be cleared of infection by treatment with antibiotics. Gentamicin treatment, followed by overnight recovery in fresh medium, resulted in the normalization of occludin distribution, a shift of occludin to the higher-molecular-weight form, suggesting an increased state of phosphorylation, and the return of TER to near baseline values. Incubation with the serine/threonine phosphatase inhibitor, calyculin A, prevented the redistribution of occludin as well as the fall in resistance in a dose-dependent fashion, suggesting that a phosphatase is responsible. Although the source of phosphatase is not known, it is possible that one of the EPEC-secreted proteins possesses this activity. Such enzyme activity by a bacterial

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B 80 70 60 50 40 30 20

FIGURE 23.7 EPEC infection of intestinal epithelial T84 cells induces a shift of occludin from a higher-molecular-weight band to a lower-molecular-weight form suggesting dephosphorylation. (A) Separation of occludin from control uninfected cells (first lane, labeled U) by SDS-PAGE and identification by immunoblotting results in the appearance of two bands, the higher band representing a hyperphosphorylated form and the lower band representing lesser or nonphosphorylated occludin. EPEC infection induces a progressive decrease in the density of the upper band and a corresponding increase in the lower-molecular-weight form, shown by the subsequent lanes representing occludin extracted from cells infected with EPEC for 30 min, 1 h, and 3 h, respectively. (B) Immunoblots from three separate experiments were quantified by densitometry, and the relative densities of the upper and lower bands are expressed in graphic form. Note that there is a temporal progression to a reciprocal relationship between these two bands after infection with EPEC; data represent mean SEM = 8 for time 0 and is <5 for all data points. (C) Time course of EPEC-induced decrease in TER correlates with the progressive shift of occludin to lower-molecular-weight forms. (From Simonovic, I. et al., Cell. Microbiol., 2:305–315, 2000. With permission of Blackwell Science Ltd.)

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FIGURE 23.8 Representative confocal laser scanning micrographs (Z-series, overlay) of T84 cell monolayers immunostained for ZO-1 with antibodies to this protein followed by secondary antibodies conjugated to fluorescein-isothiocyanate. (A) Uninfected T84 cells show a normal distribution of this TJ-associated protein outlining the perimeter of the cells. (B) T84 cells infected with EHEC CL56 for 15 h show that the intensity of the ZO-1 staining is decreased compared with that observed for uninfected cells. In addition, areas of ZO-1 disruption are present (arrows). Images were collected in 1 µm increments beginning at the apical aspect of the monolayers and optically sectioning to the basolateral membrane. Original magnification ×2400. (From Philpott, D. J. et al., Infect. Immun., 66:1680–1687, 1998. With permission.)

protein is not unprecedented, as YopH, secreted by type III secretion of Yersinia, is a tyrosine phosphatase that targets eukaryotic proteins (Guan and Dixon, 1990). Alternatively, it is possible that infection by EPEC activates a host cell phosphatase. Type III secretion is required for the full expression of effects. Whether infection by EHEC induces similar changes in occludin is not known; however, corresponding with the decrease in TER that has been reported is the redistribution of ZO-1 (Philpott et al., 1998). Specifically, in uninfected monolayers ZO-1 localizes, much like occludin, to the apical membrane, consistent with its TJ association. Following infection with EHEC, ZO-1 retains its membrane-associated localization; however, the intensity of staining is diminished and disruptions appear (Figure 23.8). Whether or not inhibition of PKC and/or MLCK also blocks the redistribution of ZO-1 has not been examined. The demonstration that pathogens, such as EPEC and EHEC, perturb TJs by inducing effects on both the cytoskeleton and TJ proteins raises the important question regarding the relationship between these two regulatory arms (see Figure 23.1). In that numerous interactions between the cytoskeleton, specifically actin and myosin, and TJ-associated proteins have been identified, it is predicted that initial alterations in the cytoskeleton would have secondary effects on TJ proteins, and vice versa. Additional studies will be required to dissect the functional aspects of these interactions.

23.4.3 HELICOBACTER

PYLORI

Published data regarding the effect of Helicobacter pylori on the TJ barrier conflict. This, however, may be attributable to the variation in model systems employed. An initial report demonstrating the dependence of H. pylori adhesion to epithelial cells

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on pH failed to show a significant impact of infection on TER (Corthesy-Thuelaz et al., 1996). In contrast to this report, which employed whole organisms, subsequent studies using either H. pylori sonicates (Terres et al., 1998) or purified H. pylori vacuolating toxin (VacA) (Papini et al., 1998) showed a decrease in TER. The active component of H. pylori sonicates responsible for the disruption of barrier function was not CagA or VacA, but rather was found to reside in a water-soluble fraction from the bacterial surface (Terres et al., 1998). Although activation of PKC by phorbol ester prevented this response, the precise signaling pathways have not been identified. It can be concluded from these studies, however, that direct interactions between H. pylori and host cells are not required to alter barrier function. Whether there are multiple H. pylori products that induce alterations in barrier function or if contamination of samples used in these studies explains the conflicting results is not known. Nevertheless, the enhanced diffusion of molecules suggests that usable substrates, such as glucose and amino acids, would potentially be made available to the organisms by this route. Conversely, diffusion of the slightly larger neutrophil chemoattractant f-MLP did not increase (Papini et al., 1998), thereby possibly limiting the inflammatory response. Also, of potential relevance to H. pylori survival was the enhanced basolateral-to-apical movement of critical ions, such as Fe3+ and Ni2+, following exposure of monolayers to VacA (Papini et al., 1998). Consistent with the reported lack of change in E-cadherin in response to H. pylori sonicates (Terres et al., 1998), VacA also failed to alter the distribution or concentration of E-cadherin (Papini et al., 1998). Similarly, no change in ZO-1, occludin, or cingulin could be demonstrated following exposure to VacA (Papini et al., 1998). Therefore, aside from the finding that activation of PKC seemed to protect against the disruption of barrier function, the mechanisms underlying this H. pylori-induced change remain unclear. One potential caveat of these experiments is that target gastric epithelial cells were not employed. Helicobacter pylori reportedly does not attach to intestinal cells in vivo. The fact that it does adhere to the transformed human intestinal epithelial cell line T84 may be due to the aberrant expression of specific receptor molecules. Even though studies of this nature provide useful data, additional investigations with gastric epithelial lines are needed. To date, however, none of the cultured human gastric epithelial lines forms confluent monolayers, thus preventing the development of TER. Encouragingly, a similar increase in permeability was observed following exposure of rat gastric mucosa to H. pylori sonicates (Terres et al., 1998). Further studies are needed to define the mechanisms by which this important human pathogen alters TJ integrity.

23.4.4 ROTAVIRUS Rotavirus, a double-stranded RNA, nonenveloped virus, is a major cause of gastroenteritis and diarrhea, especially in infants. Although destruction of intestinal villus enterocytes may in part explain the severe symptomatology, oftentimes significant diarrhea occurs in the face of mild histopathological changes. Recent studies have, therefore, focused on the impact of rotavirus on the TJ barrier. Using a model polarized human intestinal epithelial cell line, Caco-2, rotavirus infection was found to enhance TJ permeability as measured by the flux paracellular markers (Obert

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et al., 2000). Altered permeability was size selective allowing molecules such as mannitol (182 Da) and fluorescein-5 and -6 sulfonic acid (478 Da), but not 3H-PEG (900 and 4000 Da), to cross. These physiological changes were observed as early as 12 h prior to the detection of cell lysis. Correlating with enhanced permeability was the dissociation of occludin from the membrane. In contrast, E-cadherin retained its assigned location. Rotavirus infection of host cells has also been shown to disrupt the apical cytoskeleton (Jourdan et al., 1998). In contrast to the link between C. botulinum C3 transferase-induced alterations in actin and ZO-1, actin rearrangements in response to rotavirus infection appear not to be responsible for the redistribution of occludin. The independent nature of these events was demonstrated by stabilizing actin filaments with jaskosplikonide, a monocyclic peptide from the sea sponge Jaspis johnstoni. While jaskosplikonide prevented rotavirus-induced actin rearrangements, it had no impact on paracellular permeability, as measured by 3Hmannitol flux, or on occludin relocalization. The mechanism by which rotavirus alters permeability, therefore, seems to be linked to alterations in TJ-associated proteins and not the actin cytoskeleton. The mechanisms underlying these observations, however, have yet to be elucidated.

23.5 ACCESS TO BASOLATERAL LIGANDS BY PATHOGENINDUCED ALTERATIONS IN TIGHT JUNCTIONS 23.5.1 YERSINIA

PSEUDOTUBERCULOSIS

In addition to restricting passage through the paracellular space, TJs also serve to maintain polarity of the apical and basolateral membranes. Interestingly, some intestinal pathogens, Shigella flexneri (Mournier et al., 1991; Perdomo et al., 1994) and Yersinia pseudotuberculosis (Isberg et al., 1990), preferentially invade cells by accessing basolaterally residing molecules. In the case of Y. pseudotuberculosis, access to β1-integrin, which is recognized by the Yersinia outer membrane protein invasin (Isberg et al., 1987), is required. This seemingly untenable situation suggests that pathogens requiring access to the basolateral domain of host cells must have devised ways of reaching these molecules. Such a scenario has been described for Y. pseudotuberculosis, which involves disruption of TJs. McCormick et al. (1997), using an elegant in vitro approach, were the first to address the question of how this luminal pathogen might gain access to basolaterally situated β1-integrin. A very early host response to infection by virtually all enteric pathogens is the transepithelial migration of neutrophils. These acute inflammatory response cells reach the site of infection, the intestinal lumen, by traversing TJs (Nash et al., 1987). As a consequence of this physical disruption, TJ functions are interrupted, both barrier and maintenance of polarity. It was surmised, therefore, that neutrophil transmigration would potentially allow for the redistribution of membrane-associated proteins beyond their normal domains. In this particular study, neutrophil transmigration across confluent intestinal epithelial T84 monolayers was induced using the chemoattractant f-MLP. The distribution of β1-integrin was assessed prior to and following transepithelial migration. In unperturbed monolayers, as in native intestinal epithelia, β1-integrin localizes to the basolateral aspect of cells. Subsequent to neutrophil

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transmigration, however, β1-integrin was demonstrated to have relocalized to the apical membrane using both immunofluorescence localization and surface biotinylation approaches. As predicted, Y. pseudotuberculosis was found to associate with host cells only at areas where neutrophils had disrupted TJs and β1 was available for invasin binding. As a result, monolayers became more susceptible to invasion by this pathogen but not by a mutant strain in which the inv gene, which encodes invasin, had been deleted, thus demonstrating the specificity of this response. Redistribution of β1 polarity was not restricted to that induced by neutrophil transmigration as chelation of calcium with EGTA, known to disrupt intercellular junctions and cell polarity, also enhanced invasin-dependent, Yersinia internalization. This scenario may in fact not require the transmigration of neutrophils to open TJs as, more recently, infection of epithelial cells with Y. pseudotuberculosis was demonstrated to increase paracellular permeability by redistributing F-actin, ZO-1, and occludin (Tafazoli et al., 2000). Specifically, apically positioned F-actin was diminished and the TJ proteins assumed a punctate pattern of staining in infected cells as compared with the uniform distribution at the peripheral membrane seen in uninfected monolayers. Both TER and the paracellular permeation by dextrans were perturbed following 3 h of infection. Interestingly, in the cell lines used, MDCK and Caco-2, wild-type Y. pseudotuberculosis, but not inv deletion strains, was noted to associate with cells solely at junctional areas potentially indicating some level of β1 exposure on the apical surface. After 3 to 4 h of infection, bacteria were found to colocalize with β1-integrin within the paracellular space suggesting that the opening of TJs by bacteria enhances the access to β1-integrin. One specfic Yersinia outer membrane protein (Yop E), which is translocated into host cells via type III secretion and disrupts actin microfilaments (Rosqvist et al., 1991), was required for the alterations in TER as well as the redistribution of ZO-1 and occludin. Although both of these studies employed in vitro models, one prior publication reported a similar impact of Y. enterocolitica infection on the TJ barrier in the murine intestine (Gogarten et al., 1994), suggesting that these findings may be extrapolated to the intact host. Although these studies highlight different mechanisms by which Yersinia may gain access to its host cell ligand, they are not mutually exclusive events. In fact, they are complementary. One could envision that initial contact between host cell and pathogen would trigger two events: initiation of the inflammatory cascade that ultimately results in the transepithelial migration of acute inflammatory cells and a series of as-yet-undefined events that disrupt actin and key TJ proteins. Each of these culminates in a common end point, disruption of the TJ barrier and movement of β1-integrin to the apical membrane.

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Azghani, A. O., Gray, L. D., and Johnson, A. R. 1993. A bacterial protease perturbs the paracellular barrier function of transporting epithelial monolayers in culture. Infect. Immun., 61, 2681. Balda, M., Whitney, A., Flores, C., Gonzalez, S., Cereijido, M., and Matter, K. 1996. Functional dissociation of paracellular permeability and transepithelial electrical resistance and disruption of the apical-basolateral intramembrane diffusion barrier by expression of a mutant tight junction membrane protein. J. Cell Biol., 134, 1031. Bamforth, S. D., Kniesel, U., Wolberg, H., Engelhardt, B., and Risau, W. 1999. A dominant mutant of occludin disrupts tight junction structure and function. J. Cell Sci., 112, 1879. Booth, B. A., Boesman-Finkelstein, M., and Finkelstein, R. A. 1984. Vibrio cholerae hemagglutinin/protease nicks cholera enterotoxin. Infect. Immun., 45, 558. Canil, C., Rosenshine, S., Ruschkowski, S., Donnenberg, M. S., Kaper, J. B., and Finlay, B. B. 1993. Enteropathogenic E. coli decreases the transepithelial electrical resistance of polarized epithelial monolayers. Infect. Immun., 61, 2755. Chambers, F., Koshy, S., Saidi, R., Clark, D., Moore, R., and Sears, C. 1997. Bacteroides fragilis toxin exhibits polar activity on monolayers of human intestinal epithelial cells (T84 cells) in vitro. Infect. Immun., 65, 3561. Cordenonsi, M., D’Atri, F., Hammar, E., Parry, D. A. D., Kendrick-Jones, J., Shore, D., and Citi, S. 1999. Cingulin contains globular and coiled-coil domains and interacts with ZO-1, ZO-2, ZO-3, and myosin. J. Cell Biol., 147, 1569. Corthesy-Thuelaz, Porta, N., Pringault, E., Racine, L., Bogdanova, A., Kraehenbuhl, J.-P., Blum, A. L., and Michetti, P. 1996. Adhesion of Helicobacter pylori to polarized T84 human intestinal cell monolayers is pH dependent. Infect. Immun., 64, 3827. Dillon, S., Rubin, E., Yakubovich, M., Pothoulakis, C., LaMont, J. T., Feig, L., and Gilbert, R. 1995. Involvement of Ras-related Rho proteins in the mechanisms of action of Clostridium difficile toxin A and toxin B. Infect. Immun., 63, 1421. Fanning, A., Jameson, B. J., Jesaitis, L., and Anderson, J. M. 1998. The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J. Biol Chem, 273, 29745. Farshori, P. and Kachar, B. 1999. Redistribution and phosphorylation of occludin during opening and resealing of tight junctions in cultured epithelial cells. J. Membr. Biol., 170, 147. Fasano, A., Baudry, B., Pumplin, D., Wasserman, S., Tall, B., Ketley, J., and Kaper, J. 1991. Vibrio cholera produces a second enterotoxin, which affects intestinal tight junctions. Proc. Natl. Acad. Sci. U.S.A., 88, 5242. Fasano, A., Fiorentini, C., Donelli, G., Uzzau, S., Kaper, J., Margaretten, K., Ding, X., Guandalini, S., Comstock, L., and Goldblum, S. 1995. Zonula occludens toxin modulates toxin junctions through protein kinase C-dependent actin reorganization, in vitro. J. Clin. Invest., 96, 710. Fasano, A., Uzzau, S., Fiore, C., and Margaretten, K. 1997. The enterotoxic effect of zonula occludens toxin on rabbit small intestine involves the paracellular pathway. Gastroenterology, 112, 839. Furuse, M., Hirase, T., Itoh, M., Nagafuchi, A., Yonemura, S., Tsukita, S., and Tsukita, S. 1993. Occludin: a novel integral membrane protein localizing at tight junctions. J. Cell Biol., 123, 1777. Furuse, M., Fujita, K., Hiiragi, T., Fujimoto, K., and Tsukita, S. 1998. Claudin-1 and claudin-2: novel integral membrane proteins localizing at tight junctions with no sequence similarity to occludin. J. Cell Biol., 141, 1539. Gogarten, W., Kockerling, A., Fromm, M., Riecken, E. O., and Schulzke, J. D. 1994. Effect of acute Yersinia enterocolitica infection on intestinal barrier function in the mouse. Scand. J. Gastroenterol., 29, 814.

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Guan, K. and Dixon, J. E. 1990. Protein tyrosine phosphatase activity of an essential virulence determinant in Yersinia. Science, 249, 553. Hartwig, J. H., Thelen, M., Rosen, A., Janmey, P. A., Nairn, A. C., and Aderem, A. 1992. MARCKS is an actin filament cross linking protein regulated by protein kinase C and calcium-calmodulin. Nature (London), 356, 618. Hase, C. C. and Finkelstein, R. A. 1993. Bacterial extracellular zinc-containing metalloproteases. Microbiol. Rev., 57, 823. Haskins, J., Gu, L., Wittchen, E., Hibbard, J., and Stevenson, B. 1998. ZO-3, a novel member of the MAGUK protein family found at the tight junction, interacts with ZO-1 and occludin. J. Cell Biol., 141, 199. Hecht, G., Pothoulakis, C., LaMont, J. T., and Madara, J. L. 1988. Clostridium difficile toxin A perturbs cytoskeletal structure and tight junction permeability of cultured human intestinal epithelial monolayers. J. Clin. Invest., 82, 1516. Hecht, G., Koutsouris, A., Pothoulakis, C., LaMont, J. T., and Madara, J. 1992. Clostridium difficile toxin B disrupts the barrier function of T84 monolayers. Gastroenterology, 102, 416. Hecht, G., Pestic, L., Nikcevic, G., Koutsouris, A., Tripuraneni, D., Lorimer, D., Nowak, G., Guerriero, J. V., Elson, E. L., and de Lanerolle, P. 1996. Expression of the catalytic domain of myosin light chain kinase increases paracellular permeability. Am. J. Physiol., 271, C1678. Iglewski, B. H. and Sadoff, J. C. 1979. Toxin inhibitors of protein synthesis: production, purification, and assay of Pseudomonas aeruginosa toxin A. Methods Enzymol., 60, 780. Isberg, R. R., Voorhis, D. L., and Falkow, S. 1987. Identification of invasin: a protein that allows enteric bacteria to penetrate cultured mammalian cells. Cell, 50, 769. Isberg, R. R., Voorhis, D. L., and Leong, M. 1990. Multiple β1 chain integrins are receptors for invasin, a protein that promotes bacterial penetration into mammalian cells. Cell, 60, 861. Itoh, M., Furuse, M., Morita, K., Kubota, K., Saitou, M., and Tsukita, S. 1999. Direct binding of three tight junction-associated MAGUKs, ZO-1, ZO-2, and ZO-3, with the COOH termini of claudins. J. Cell Biol., 147, 1351. Jourdan, N., Brunet, J.-P., Sapin, C., Blais, A., Cotte-Laffitte, J., Forestier, F., Quero, A.-M., Trugnan, G., and Servin, A. 1998. Rotavirus infection reduces sucrase-isomaltase expression in human intestinal epithelial cells by perturbing protein targeting and organization of microvillar cytoskeleton. J. Virol., 72, 7228. Just, I., Fritz, G., Aktories, K., Giry, M., Popoff, M., Boquet, P., Hegenberth, S., and von Eichel-Streiber, C. 1994. Clostridium difficile toxin B acts on the GTP-binding protein Rho. J. Biol. Chem., 269, 10706. Just, I., Selzer, J., von Eichel-Streiber, C., and Aktories, K. 1995. The low molecular mass GTP-binding protein Rho is affected by toxin A from Clostridium difficile. J. Clin. Invest., 95, 1026. Katz, J., Sambandam, V., Wu, J., Michalek, S., and Balkovetz, D. 2000. Characterization of Phorphyromonas gingivalis-induced degradation of epithelial cell junctional complexes. Infect. Immun., 68, 1441. Kimsey, H. H. and Waldor, M. D. 1998. Vibrio cholerae hemagglutin/protease inactivates CTXΦ. Infect. Immun., 66, 4025. Kinugasa, T., Sakaguchi, T., Gu, X., and Reinecker, H. 2000. Claudins regulate the intestinal barrier in response to immune mediators. Gastroenterology, 118, 1001. Leroy, A., De Bruyne, G., Verspeelt, A., Lauwaet, T., Nelis, H., and Mareel, M. 1997. Bacterium-assisted invasion of Entamoeba histolytica through human enteric epithelia in two-compartment chambers. Invasion Metastasis, 17, 138.

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Leroy, A., Lauwaet, T., De Bruyne, G., Cornelissen, M., and Mareel, M. 2000. Entamoeba histolytica disturbs the tight junction complex in human enteric T84 cell layers. FASEB J., 14, 1139. Li, E., Stenson, W. F., Kunz-Jenkins, C., Swanson, C., Duncan, P. E., and Stanley, S. L., Jr. 1994. Entamoeba histolytica interactions with polarized human intestinal Caco-2 epithelial cells. Infect. Immun., 62, 5112. Madara, J. L. and Pappenheimer, J. R. 1987. Structural basis for physiological regulation of paracellular pathways in intestinal epithelia. J. Membr. Biol., 100, 149. Manjarrez-Hernandez, H. A., Amess, B., Sellers, L., Baldwin, T. J., Knutton, S., Williams, P. H., and Aitken, A. 1991. Purification of a 20 kDA phosphoprotein from epithelial cells and identification as myosin light chain. FEBS Lett., 292, 121. Manjarrez-Hernandez, H. A., Baldwin, T. J., Williams, P. H., Haigh, R., Knutton, S., and Aitken, A. 1996. Phosphorylation of the myosin light chain at distinct sites and its association with the cytoskeleton during enteropathogenic Escherichia coli infection. Infect. Immun., 64, 2368. Matter, K. and Balda, M. S. 1998. Biogenesis of tight junctions: the C-terminal domain of occludin mediates basolateral targeting. J. Cell Sci., 111, 511. McCarthy, K., Skare, L., Stankewich, M., Furuse, M., Tsukita, S., Rogers, R., Lynch, R., and Schneeberger, E. 1996. Occludin is a functional component of the tight junction. J. Cell Sci., 109, 2287. McCormick, B., Nusrat, A., Parkos, C., D’Andrea, L., Hofman, P., Carnes, D., Liang, T., and Madara, J. 1997. Unmasking of intestinal epithelial lateral membrane β1-integrin consequent to transepithelial neutrophil migration in vitro facilitates inv-mediated invasion by Yersinia pseudotuberculosis. Infect. Immun., 65, 1414. Mel, S., Fullner, K. J., Wimer-Mackin, S., Lencer, W., and Mekalanos, J. 2000. Association of protease activity in Vibrio cholerae vaccine strains with decreases in transcellular epithelial resistance of polarized T84 epithelial cells. Infect. Immun., 68:6487–6492. Moncrief, J. S., Obiso, R., Barroso, L. A., Kling, J. J., Wright, R. L., Van Tassell, R. L., Lyerly, D. M., and Wilkins, T. D. 1995. The enterotoxin of Bacteroides fragilis is a metallaprotease. Infect. Immun., 63. Moore, R., Pothoulakis, C., LaMont, J. T., Carlson, S., and Madara, J. 1990. C. difficile toxin A increases intestinal permeability and induces Cl– secretion. Am. J. Physiol., 90, G165. Mournier, J., Vasselon, T., Hellio, R., LeSourd, M., and Sansonetti, P. J. 1991. Shigella flexneri enters human Caco-2 epithelial cells through the basolateral pole. Infect. Immun., 60, 237. Nash, S., Stafford, J., and Madara, J. L. 1987. Effects of polymorphonuclear leukocyte transmigration on the barrier function of cultured intestinal epithelial monolayers. J. Clin. Invest., 80, 1104. Nusrat, A., Giry, M., Turner, J. R., Colgan, S. P., Parkos, C. A., Carnes, D., Lemichez, E., Boquet, P., and Madara, J. 1995. Rho protein regulates tight junctions and perijunctional actin organization in polarized epithelia. Proc. Natl. Acad. Sci. U.S.A., 92, 10629. Nusrat, A., Parkos, A., Verkade, P., Foley, C. S., Liang, T. W., Innis-Whitehouse, W., Eastburn, K. K., and Madara, J. L. 2000. Tight junctions are membrane microdomains. J. Cell Sci., 113, 1771. Obert, G., Peiffer, I., and Servin, A. 2000. Rotavirus-induced structural and functional alterations in tight junctions of polarized intestinal Caco-2 cell monolayers. J. Virol., 74, 4645. Papini, E., Satin, B., Norals, N., de Bernard, M., Telford, J., Rappuoli, R., and Montecucco, C. 1998. Selective increase of the permeability of polarized epithelial cell monolayers by Helicobacter pylori vacuolating toxin. J. Clin. Invest., 102, 813.

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Perdomo, J. J., Gounon, P., and Sansonetti, P. J. 1994. Polymorphonuclear leukocyte transmigration promotes invasion of colonic epithelial monolayers by Shigella flexneri. J. Clin. Invest., 93, 633. Perna, N. T., Mayhew, G. F., Posfai, G., Elliot, S. J., Donnenberg, M. S., Kaper, J. B., and Blattner, F. R. 1998. Molecular evolution of a pathogenicity island from enterohemorrhagic Escherichia coli O157:H7. Infect. Immun., 66, 3810. Philpott, D. J., McKay, D. M., Sherman, P. M., and Perdue, M. H. 1996. Infection of T84 cells with enteropathogenic Escherichia coli alters barrier and transport. Am. J. Physiol., 270, G634. Philpott, D., McKay, D., Mak, W., Perdue, M., and Sherman, P. 1998. Signal transduction pathways involved in enterohemorrhagic Escherichia coli-induced alterations in T84 epithelial permeability. Infect. Immun., 66, 1680. Riegler, M., Sedivy, R., Pothoulakis, C., Hamilton, G., Zacheri, J., Bischof, G., Consentini, E., Feil, W., Schiessel, R., LaMont, J. T., and Wenzly, E. 1995. E. Clostridium difficile toxin B is more potent than toxin A in damaging human colonic epithelim in vitro. J. Clin. Invest., 95, 2004. Rosqvist, R., Forsberg, A., and Wolf-Watz, H. 1991. Intracellular targeting of the Yersinia YopE cytotoxin in mammalian cells induces actin microfilament disruption. Infect. Immun., 59, 4562. Sakakibara, A., Furuse, M., Saitou, M., Ando-Akatsuka, Y., and Tsukita, S. 1997. Possible involvement of phosphorylation of occludin in tight junction formation. J. Cell Biol., 137, 1393. Sandkvist, M., Michel, L. O., Hough, L. P., Morales, V. M., Bagdasarian, M., Koomey, M., DiRita, V. J., and Bagdasarian, M. 1997. General secretion pathway (eps) genes required for toxin secretion and outer membrane biogensesis in Vibrio cholerai. J. Bacteriol., 179, 6994. Schmidt, G. and Aktories, K. 1998. Bacterial cytotoxins target Rho GTPases. Nature, 85, 253. Simonovic, I., Rosenberg, J., Koutsouris, A., and Hecht, G. 2000. Enteropathogenic E. coli dephosphorylates and dissociates occludin from intestinal epithelial tight junctions. Cell. Microbiol., 2, 305. Sonoda, N., Furuse, M., Sasaki, H., Yonemura, S., Katahira, J., Horiguchi, Y., and Tsukita, S. 1999. Clostridium perfringens enterotoxin fragment removes specific claudins from tight junction strands: evidence for direct involvement of claudins in tight junction barrier. J. Cell Biol., 147, 195. Spitz, J., Yuhan, R., Koutsouris, A., Blatt, C., Alverdy, J., and Hecht, G. 1995. Enteropathogenic Escherichia coli adherence to intestinal epithelial monolayers diminishes barrier function. Am. J. Physiol., 268, G374. Tafazoli, F., Holmstrom, A., Forsberg, A., and Magnusson, K.-E. 2000. Apically exposed, tight junction-associated β1-integrins allow binding and YopE-mediated perturbation of epithelial barriers by wild-type Yersinia bacteria. Infect. Immun., 68, 5335. Terres, A. M., Pajares, J. M., Hopkins, A. M., Murphy, A., Moran, A., Baird, A. W., and Kelleher, D. 1998. Helicobacter pylori disrupts epithelial barrier function in a process inhibited by protein kinase C activators. Infect. Immun., 66, 2943. Thelen, M., Rosen, A., Naim, A. C., and Aderem, A. 1991. Regulation by phosphorylation of reversible association of a myristoylated protein kinase C substrate with the plasma membrane. Nature (London), 351, 320. Turner, J. R., Rill, B. K., Carlson, S. L., Carnes, D., Kerner, R., Mrsny, R. J., and Madara, J. L. 1997. Physiological regulation of epithelial tight junctions is associated with myosin light-chain phosphorylation. Am. J. Physiol., 273, C1378.

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Wittchen, E., Haskins, J., and Stevenson, B. 1999. Protein interactions at the tight junction. J. Biol. Chem., 274, 35179. Wong, V. 1997. Phosphorylation of occludin correlates with occludin localization and function at the tight junction. Am. J. Physiol., 273, C1859. Wong, V. and Gumbiner, B. M. 1997. A synthetic peptide corresponding to the extracellular domain of occludin perturbs the tight junction permeabilty barrier. J. Cell Biol., 136, 399. Wu, S., Lim, K., Huang, J., Saidi, R., and Sears, C. 1998. Bacteroides fragilis enterotoxin cleaves the zonula adherens protein, E-cadherin. Proc. Natl. Acad. Sci. U.S.A., 95, 14979. Wu, Z., Milton, D., Nybom, P., Sjo, A., and Magnusson, K. E. 1996. Vibrio cholerae hemagglutinin/protease (HA/protease) causes morphological changes in cultured epithelial cells and perturbs their paracellular barrier function. Microb. Pathog., 21, 111. Wu, Z., Nybom, P., and Magnusson, K. 2000. Distinct effects of Vibrio cholerae haemagglutinin/protease on the structure and localization of the tight junction-associated proteins occludin and ZO-1. Cell Microb., 2, 11.

24

Interactions between Clostridium perfringens Enterotoxin and Tight Junction Proteins Bruce A. McClane and Usha Singh

CONTENTS 24.1 Introduction .................................................................................................517 24.2 The Role of CPE in Human Gastrointestinal Disease...............................518 24.3 CPE Mechanism of Action .........................................................................519 24.3.1 CPE Effects on the Gastrointestinal Tract ....................................519 24.3.2 CPE Effects on Mammalian Cells ................................................521 24.4 CPE Interactions with Tight Junction Proteins ..........................................522 24.4.1 Early Studies of CPE Receptors ...................................................522 24.4.2 Identification of Claudins 3 and 4 as Functional CPE Receptors........................................................................................523 24.4.3 CPE Also Interacts with Occludin, Another Important Structural Protein Component of Tight Junctions ........................524 24.5 Consequences of CPE: Tight Junction Protein Interactions for Tight Junction Structure .......................................................................................525 24.6 The Importance of CPE-Induced Changes in Tight Junction Structure for CPE-Associated Gastrointestinal Disease ............................................526 24.7 A Current Model for CPE Action ..............................................................527 24.8 Concluding Comments ...............................................................................529 Acknowledgments..................................................................................................530 References..............................................................................................................530

24.1 INTRODUCTION Clostridium perfringens is a Gram-positive, spore-forming, anaerobic bacterium that causes both histotoxic and enteric infections in humans and domestic animals (McDonel, 1986; Rood and Cole, 1991). The virulence of this important pathogen is largely attributable to its ability to produce a vast arsenal of potent protein 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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FIGURE 24.1 Functional regions of the CPE protein. CPE regions required for receptor binding, intermediate and large complex formation, and cytotoxicity are indicated. This map is compiled from Granum (1991; 1981), Hanna (1991; 1992), and Kokai-Kun (1997; 1999).

exotoxins. At least 15 different C. perfringens toxins have now been reported in the literature (McDonel, 1986; Rood and Cole, 1991; Gibert et al., 1997). However, an individual C. perfringens isolate can express only a subset of this total toxin repertoire, which provides the basis for a classification scheme that assigns individual C. perfringens isolates to one of five types (A through E), depending upon their ability to produce alpha, beta, epsilon, and iota toxins (McDonel, 1986). About 5% of all C. perfringens isolates, mostly belonging to type A, can produce C. perfringens enterotoxin (CPE) (Kokai-Kun et al., 1994; Daube et al., 1996; Songer and Meer, 1996). The CPE protein is a single 319 amino acid polypeptide that has a unique primary sequence, except for some limited homology (of unknown significance) with a non-neurotoxic protein produced by C. botulinum (Hauser et al., 1994). As depicted in Figure 24.1, recent structure–function mapping studies indicate that, like most bacterial toxins, CPE segregates its binding and activity regions. Receptor binding activity localizes to the C-terminal 30 amino acids of the enterotoxin protein (Hanna et al., 1991; Kokai-Kun and McClane, 1997; Kokai-Kun et al., 1999), while amino acid residues 45 to 116 in the N-terminal region of CPE are important for biologic activity (see Sections 24.3 and 24.4). Interestingly, removal of amino acid residues 1 to ~43 from the extreme N terminus of native CPE activates the biologic activity of the toxin about two- to threefold (Granum et al., 1981; Hanna et al., 1992; Kokai-Kun and McClane, 1997). Since both trypsin and chymotrypsin treatment can specifically remove these same extreme N-terminal CPE amino acid residues (Granum et al., 1981; Granum and Richardson, 1991), CPE may be proteolytically activated in the intestinal lumen during CPE-associated gastrointestinal disease (see below).

24.2 THE ROLE OF CPE IN HUMAN GASTROINTESTINAL DISEASE Enterotoxin-producing C. perfringens type A strains first became recognized as a cause of human gastrointestinal disease over 30 years ago, when they were associated with C. perfringens type A food poisoning (McClane, 1997). Clostridium perfringens type A food poisoning, the second most commonly reported foodborne disease in the United States and most other industrialized countries (Bean et al., 1996), develops when large numbers of cpe-positive C. perfringens type A vegetative cells are

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ingested in a contaminated food, typically a meat or poultry item (McClane, 1997; McClane et al., 2000). Once present in the small intestinal lumen, these bacteria multiply and then sporulate. It is during this in vivo sporulation process that the enterotoxin protein is produced; i.e., C. perfringens type A food poisoning is a true gastrointestinal infection, rather than a foodborne intoxication resulting from ingestion of a preformed toxin already present in foods. After expression, CPE is not secreted; instead, the enterotoxin accumulates in the cytoplasm of the sporulating C. perfringens cell and is only released into the intestinal lumen when the sporulating C. perfringens cell lyses to free its mature endospore. Symptoms of C. perfringens type A food poisoning typically include diarrhea and abdominal cramps, which develop about 12 to 16 h after ingestion of a contaminated food (McClane, 1997; McClane et al., 2000). Those gastrointestinal symptoms usually persist for 12 to 24 h, after which most victims of C. perfringens type A food poisoning recover without lasting effects. However, fatalities sometimes occur when elderly or debilitated individuals are sickened with this food poisoning. More recently, CPE-producing C. perfringens type A isolates have also become associated with several non-foodborne human gastrointestinal diseases, including both antibiotic-associated diarrhea and sporadic diarrhea (Carman, 1997; McClane et al., 2000). Epidemiological studies suggest that CPE-producing C. perfringens type A strains may be responsible for up to 5 to 20% of all cases of non-foodborne human gastrointestinal diseases (Carman, 1997; McClane et al., 2000). Interestingly, the diarrheic and cramping symptoms of CPE-associated non-foodborne gastrointestinal diseases tend to be more severe and longer lasting than those of C. perfringens type A food poisoning. These symptomology differences may reflect recently identified genotypic and phenotypic differences between CPE-positive non-foodborne gastrointestinal disease isolates vs. CPE-positive food poisoning isolates (Collie et al., 1998; Collie and McClane, 1998; Sarker et al., 2000). A wealth of epidemiological and experimental data now offers direct support for the importance of CPE in the pathogenesis of both C. perfringens type A food poisoning and CPE-associated non-foodborne human gastrointestinal diseases. For example, feeding experiments with human volunteers demonstrated that ingestion of highly purified CPE is sufficient to induce the diarrheic and cramping symptoms of CPE-associated gastrointestinal diseases (Skjelkvale and Uemura, 1977). Equally persuasive support for the importance of CPE expression in the gastrointestinal virulence of both CPE-associated food poisoning and non-foodborne human gastrointestinal diseases was recently provided by studies showing that specific inactivation of the cpe gene in both human foodborne and non-foodborne gastrointestinal disease isolates completely eliminates the ability of those isolates to induce gastrointestinal effects (see below) in rabbit ileal loops (Sarker et al., 1999).

24.3 CPE MECHANISM OF ACTION 24.3.1 CPE EFFECTS

ON THE

GASTROINTESTINAL TRACT

Results from animal model studies indicate that CPE can affect all segments of the small intestine, with the ileum being particularly sensitive (McDonel and Duncan,

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1977). Interestingly, the rabbit colon is reportedly insensitive to this enterotoxin, despite the presence of high numbers of CPE receptors (McDonel and Demers, 1982). Whether the human colon is similarly unaffected by CPE remains unknown. Addition of CPE to rabbit ileal loops rapidly induces fluid/electrolyte transport alterations (McDonel and Duncan, 1975; McDonel et al., 1978; Sherman et al., 1994). Those CPE-induced transport alterations initially start as an inhibition of fluid/electrolyte absorption, but develop into frank fluid/electrolyte secretion with longer CPE treatment periods. The intestinal action of CPE differs in several ways from that of the classical bacterial enterotoxin cholera toxin (McDonel, 1986). First, CPE (unlike cholera toxin) inhibits intestinal glucose uptake. Second, CPE does not increase intestinal cAMP levels, as has been observed in the cholera toxin-treated small intestine. Finally, CPE (but not cholera toxin) rapidly induces intestinal histopathological damage (Figure 24.2), which is characterized by both villus shortening and

FIGURE 24.2 CPE-induced histopathological damage. (A) The histology of normal rabbit ileum; (B) the effect of overnight CPE treatment on rabbit ileum. Note the severe epithelial desquamation and villi shortening present in the CPE-treated rabbit ileum (panel B). Both specimens are shown at 250× original magnification.

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severe desquamation of the intestinal epithelium, particularly at villi tips (McDonel and Duncan, 1975; 1977; McDonel et al., 1978; Sherman et al., 1994). Several observations suggest this histopathological damage initiates the fluid/electrolyte transport changes observed in the CPE-treated small intestine. For example, this histopathological damage develops concurrently with fluid/electrolyte transport changes in the CPE-treated rabbit ileum; i.e., in rabbit ileum treated with high doses of CPE, both histological damage and fluid/electrolyte transport alterations become detectable within 15 to 30 min (Sherman et al., 1994). Furthermore, other studies have established that only those CPE doses that induce histopathological damage are capable of causing fluid transport changes in rabbit ileal loops (McDonel and Duncan, 1975).

24.3.2 CPE EFFECTS

ON

MAMMALIAN CELLS

Available data indicate that the CPE-induced histopathological damage described in Section 24.3.1 results from the ability of the enterotoxin to kill mammalian cells rapidly. For example, studies have shown that CPE-induced cytotoxicity and histopathological damage develop concurrently (McClane and McDonel, 1979; Sherman et al., 1994). An early insight into how CPE kills mammalian cells was provided by studies showing that CPE affects plasma membranes prior to its effects on intracellular organelles. Additional studies showed that the enterotoxin remains associated with the plasma membrane throughout its action; i.e., CPE is not internalized into the cytoplasm of enterocytes or other mammalian cells (McClane, 1994). A plasma membrane site of action for CPE was confirmed when it was shown that this toxin disrupts the normal permeability properties of the mammalian plasma membrane (McClane, 1994); i.e., CPE-treated mammalian cells rapidly become more permeable to small molecules of <200 Da, including both ions and organic molecules (such as amino acids). Those CPE-induced small molecule permeability alterations collapse the colloid-osmotic equilibrium of the mammalian cell, leading to cell death from either lysis or metabolic effects, such as a shutdown of macromolecular synthesis (McClane, 1994). At 37°C, CPE interacts with eucaryotic proteins to form a series of complexes in the mammalian plasma membrane. These complexes can be differentiated on the basis of their size and sensitivity to dissociation by SDS. SDS-PAGE analyses using 6% acrylamide gels (no sample boiling) initially indicated that mammalian cells treated with CPE at 37°C contain only a single, SDS-resistant complex of ~160 kDa (Wnek and McClane, 1989). However, recent SDS-PAGE studies (Singh et al., 2000) using 4% acrylamide gels have now resolved (see Figure 24.2) three separate CPEcontaining species of SDS-resistant, high Mr material in CaCo-2 human intestinal cells treated with enterotoxin at 37°C, including complexes of ~135, ~155, and ~200 kDa. Kinetic studies suggest that the ~135 kDa complex may represent an intermediate precursor for formation of the ~155 and ~200 kDa large complexes in those CPE-treated CaCo-2 cells (Singh et al., 2000). Several experimental observations strongly suggest that formation of one or more of these three SDS-resistant, CPE-containing complexes is responsible for the

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small molecule membrane permeabilty alterations that kill CPE-treated mammalian cells. For example: 1. CPE binds to Vero cells at 4°C, yet fails to induce membrane permeability alterations at that temperature or to associate into the SDS-resistant complexes (McClane and Wnek, 1990). 2. If those Vero cells treated with CPE at 4°C are washed (to remove unbound CPE) and quickly shifted to 37°C, the SDS-resistant complexes form concurrently with the development of membrane permeability alterations (McClane and Wnek, 1990). 3. CPE fragments comprising residues 45 to 319 of the native enterotoxin can induce greater membrane permeability changes than does the native CPE; relative to native CPE, those hypertoxic CPE fragments also form about two- to threefold more of the SDS-resistant complexes (Kokai-Kun and McClane, 1997). 4. Random mutagenesis studies have identified several noncytoxic CPE point mutants that can bind similarly to native CPE, but are unable to form the ~135, ~155, or ~200 kDa complexes (Kokai-Kun et al., 1999). How might formation of the ~135, ~155, and/or ~200 kDa SDS-resistant CPE complexes alter mammalian plasma membrane permeability properties? Recent electrophysiology experiments strongly suggest that CaCo-2 cells develop pores in their apical membranes when treated with CPE at 37°C (Hardy et al., 1999), a condition where the SDS-resistant complexes can form. Furthermore, Pronase challenge studies indicate that, when CPE is localized in SDS-resistant complexes, it becomes closely associated with plasma membranes (Wieckowski et al., 1998). Collectively, these findings suggest that CPE becomes inserted into the membrane as part of a pore structure when it is present in one or more of the SDS-resistant complexes.

24.4 CPE INTERACTS WITH TIGHT JUNCTION PROTEINS 24.4.1 EARLY STUDIES

OF

CPE RECEPTORS

Since CPE can bind but does not form the ~135, ~155, or ~200 kDa complexes or induce cytotoxic effects when added to mammalian cells at 4°C (McClane and Wnek, 1990), one or more steps in CPE action obviously must precede formation of the SDS-resistant, high-Mr complexes. That conclusion has been experimentally proven by studies demonstrating that addition of CPE to Vero cells or CaCo-2 cells at 4°C results in the localization of CPE in an ~90 kDa, SDS-sensitive complex (Wieckowski et al., 1994). By using appropriate assays, that CPE-containing 90 kDa complex, now referred to as small complex, can also be detected in mammalian cells treated with toxin at 37°C (Wieckowski et al., 1994); i.e., under physiological conditions, CPE-treated CaCo-2 cells may contain at least four different CPE species, which include the ~90 kDa small complex, the ~135 kDa intermediate complex, and the ~155 and ~200 kDa large complexes.

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Kinetic analyses demonstrated that formation of the small CPE-containing complex closely coincides with CPE binding and that formation of this small complex is a precursor for formation of the larger, SDS-resistant, CPE-containing complexes (Wieckowski et al., 1994). Immunoprecipitation studies have also shown that an ~45 to 50 kDa eucaryotic protein is associated with the small CPE complex (Wieckowski et al., 1994). Based upon those findings, it was initially thought that small complex formation reflects the binding of CPE to its ~45 to 50 kDa protein receptor.

24.4.2 IDENTIFICATION OF CLAUDINS 3 CPE RECEPTORS

AND

4

AS

FUNCTIONAL

Just 3 years after the identification of an ~45 to 50 kDa protein as a putative CPE receptor (see Section 24.4.1), Katahira et al. (1997a) published elegant expression cloning studies that conclusively demonstrated that an ~22 kDa Vero cell protein can serve as a functional CPE receptor. Those expression cloning studies showed that Mouse L cells, which are not naturally affected by CPE because they are unable to bind this toxin, become very sensitive to CPE when transfected with a cDNA encoding an ~22 kDa Vero cell protein. Using Western overlay ligand blots and immunoprecipitation techniques, Katahira et al. demonstrated direct binding of CPE to that ~22 kDa protein. Evidence was also presented in that study suggesting that CPE treatment promotes formation of high-Mr, SDS-resistant, CPE-containing complexes in these L cell transfectants, although the electrophoresis conditions used were insufficient to resolve the size of those CPE-containing, high-Mr complexes. Finally, that initial study also presented data suggesting the ~22 kDa functional CPE receptor, which Katahira et al. originally named CPE-R, becomes localized in this SDS-resistant complex(es). Katahira et al. then published a second study revealing that human cells can express two different ~22 kDa proteins with homology with CPE-R (Katahira et al., 1997b). The first of those two human CPE-R homologues, which shares ~99% homology with CPE-R, was named hCPE-R. The second human CPE-R homologue, which shares ~65% homology with CPE-R, was named hRVP-1. In their second 1997 study, Katahira et al. also demonstrated that both hCPE-R and hRVP-1 can serve as functional CPE receptors; i.e., stable expression of either hCPE-R or hRVP1 in mouse L cell renders those transfectants sensitive to CPE-induced cytotoxicity. Finally, that study presented Northern blot analyses showing mouse homologues of hCPE-R and hRVP-1 are abundantly expressed in the mouse small intestine, as well as in several other mouse organs (e.g., lungs, liver, kidney). More recently, it was determined (Morita et al., 1999) that hCPE-R and hRVP-1 are members of the claudin family. Claudins are four-transmembrane-domain proteins that are now recognized as important structural components of epithelial tight junctions (TJs) (see Chapter 10). Consequently, hRVP-1 and hCPE-R (and their homologues in other animal species) were renamed claudins 3 and 4, respectively. Based upon differences in their C-terminal regions, at least 20 different claudins have now been identified. Interestingly, it appears that only certain claudins are able to function as CPE receptors, e.g., claudins 3, 4, 6, 7, 8, and 14, but not claudins 1,

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2, 5, and 10, can reportedly bind CPE (Fujita et al., 2000). Recent studies suggest that CPE appears to interact with the second extracellular loop of claudin-3 (Fujita et al., 2000).

24.4.3 CPE ALSO INTERACTS WITH OCCLUDIN, ANOTHER IMPORTANT STRUCTURAL PROTEIN COMPONENT OF TIGHT JUNCTIONS The identification of claudins 3 and 4 as functional CPE receptors raised questions regarding whether CPE might interact with other TJ-associated proteins. The possibility of CPE interactions with occludin, which is another major structural protein component of TJs (see Chapter 10), was particularly intriguing considering, 1. Claudins can sometimes associate with occludin in control (non-CPEtreated) mammalian cells (Morita et al., 1999), and 2. Early affinity chromatography and preparative electrophoresis studies had suggested that a protein of ~65 kDa, which matches the size of occludin, may associate with CPE in cells containing the SDS-resistant, high-Mr complexes (Wnek and McClane, 1989). By using preparative electrophoresis and immunoprecipitation approaches, a recent study confirmed the ability of CPE to interact with occludin in CaCo-2 cells (Singh et al., 2000). Specifically, that study demonstrated (Figure 24.3) the presence of occludin in the ~200 kDa CPE complex. However, occludin was found to be absent from the ~155 kDa large complex, the ~135 kDa intermediate complex, and the ~90 kDa small complex. Those findings imply that CPE–occludin interactions occur relatively late in the action of the enterotoxin. That conclusion received further support from another experiment presented in the Singh et al. study, which showed that Rat-1/R12 fibroblast transfectants expressing occludin (but no claudin proteins) fail to bind CPE and are insensitive to this toxin (Singh et al., 2000).

FIGURE 24.3 Presence of multiple SDS-resistant, CPE-containing complexes in CaCo-2 cells treated with CPE at 37°C. CaCo-2 cells were incubated at 37°C with CPE for desired time periods (as indicated) before the cells were extracted with SDS and the extracts were electrophoresed on 4% acrylamide gels containing SDS. Those gels were then immunoblotted using CPE or occludin antibodies, as indicated. The migration of myosin (212 kDa) and β-galactosidase (~122 kDa) are shown as size markers. The double, open, and closed arrows indicate the location of the ~200, ~155, and ~135 kDa complexes respectively. (From Singh, U. et al., J. Biol. Chem., 275, 18407, 2000. With permission.)

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The study by Singh et al. (2000) also demonstrated that several nontoxic CPE mutants carrying single point mutations in their N-terminal sequences are capable of binding and forming the SDS-sensitive, small-CPE complex, but are unable to promote formation of any of the larger, SDS-resistant CPE complexes. Those results indicate that the enterotoxin plays an active role in the association of occludin with the ~200 kDa large complex; i.e., the presence of occludin in the ~200 kDa large complex is not simply attributable to previously reported claudin–occludin interactions (in which case, occludin should have been present in all three intermediate/large CPE complexes, assuming that certain claudins represent the major CPE receptor). The inability of those nontoxic CPE point mutants to form any of the SDSresistant, high-Mr CPE complexes is also consistent with the ~135 kDa complex being an intermediate for formation of the ~155- and ~200 kDa complex. That result would also be consistent with all three SDS-resistant complexes being important for CPE-induced cytotoxicity. However, more recent data from Singh et al. (unpublished) indicates that confluent Transwell cultures of both CaCo-2 and Vero cells, which are highly sensitive to CPE-induced cytotoxicity, form detectable amounts of the ~155 kDa large complex, but not of the ~200 kDa large complex. Therefore, formation of the ~200 kDa large complex does not appear necessary for obtaining a CPE-induced cystotoxic response.

24.5 CONSEQUENCES OF CPE: TIGHT JUNCTION PROTEIN INTERACTIONS FOR TIGHT JUNCTION STRUCTURE Given the ability of CPE to interact with certain claudins and occludin (see Section 24.4), it became logical to ask whether this enterotoxin affects the structure of epithelial tight junctions. Results from two independent studies have now confirmed that, under certain experimental conditions, CPE (or CPE fragments) can affect TJ structure. A study by Sonoda et al. (1999) has demonstrated that application of C-CPE, a noncytotoxic CPE fragment comprising approximately the C-terminal half of the native enterotoxin protein, to the basolateral surface of Madin–Darby canine kidney (MDCK) cells causes the specific removal of claudin-4 from TJs of those cells. This effect results in a gradual disintegration of TJ fibrils in the CPE-treated MDCK cells (Sonoda et al., 1999). Independently, another study (Rahner et al., 1999) concluded that native CPE can also affect TJs, when it was found that application of CPE to the basolateral surface of hepatocytes in perfused rat liver rapidly induces dramatic fragmentation of TJ fibrils (Figure 24.4). Rahner et al. also reported that a nontoxic C-terminal CPE fragment, which was very similar to the C-CPE fragment used by Sonoda et al., fails to induce any TJ effects within a similar time frame in their perfused rat liver model system. It is not yet clear why native CPE induces TJ structural damage more quickly than C-terminal CPE fragments, at least in the rat liver model. Based upon Singh et al.’s recent results indicating that N-terminal CPE sequences are important for CPE associations with occludin (Singh et al., 2000), one possible explanation could be that TJ damage develops more quickly when rat liver is perfused with native CPE

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FIGURE 24.4 Effects of CPE treatment on TJs. (A) Freeze-fracture electron microscopy of perfused rat liver treated with a noncytotoxic C-terminal CPE fragment comprising amino acids 171 to 319 of native enterotoxin (Hanna et al., 1991; Kokai-Kun and McClane, 1997; Kokai-Kun et al., 1999). This specimen shows normal hepatocellular TJ strand morphology, with three to five anastomosing fibrils, which run parallel to the canalicular axis, forming the sealing elements. The canicular microvilli also appeared normal after treatment with this CPE fragment. Original primary magnification 39,000×. (B) Freeze-fracture electron microscopy of perfused rat liver after treatment with native CPE. This specimen shows dramatic fragmentation of TJ fibrils within minutes of toxin treatment. In addition to these TJ alterations, CPE treatment also resulted in a complete loss of canalicular microvilli paralleled by a flattened plasma membrane surface. Original primary magnification 39,000×. (This figure, and its interpretation, courtesy of Drs. Christoph Rahner and James Anderson, Yale University School of Medicine.)

vs. C-terminal CPE fragments because the native enterotoxin (but not C-terminal CPE fragments) can react with both claudins and occludin, two major structural components of the TJ.

24.6 THE IMPORTANCE OF CPE-INDUCED CHANGES IN TIGHT JUNCTION STRUCTURE FOR CPE-ASSOCIATED GASTROINTESTINAL DISEASE Sonoda et al. (1999) have also demonstrated that their noncytotoxic C-CPE fragment, which alters TJ morphology, increases paracellular permeability in polarized monolayers of MDCK cells. If similar paracellular permeability increases occurred in CPE-treated intestinal epithelium, they could certainly contribute to the diarrheic symptoms of CPE-associated human gastrointestinal diseases. However, several observations argue that TJ damage-mediated increases in paracellular permeability probably do not represent the initial effect of CPE on the

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intestines. The first of these observations has already been mentioned; i.e., CPEinduced histopathological damage, which apparently results from the cytotoxic action of this enterotoxin, appears to be necessary for initiating fluid/electrolyte losses in rabbit ileal loops. Second, Sonoda et al. (1999) observed TJ damage, and consequent paracellular permeability effects, only when their C-CPE fragment was added to the basolateral side of polarized MDCK monolayers. Similarly, Rahner et al. (1999) only observed TJ damage when native CPE was added to the basolateral side of rat liver cells. This ability of CPE (or C-CPE) to induce TJ structural effects and paracellular permeability changes only when applied to the basolateral surface of an epithelium provides a compelling argument that such effects are not likely to represent the primary, initiating action of the enterotoxin, if the following are considered: 1. C. perfringens cells are not known to invade the intestinal epithelium during CPE-associated gastrointestinal disease; instead those bacteria release CPE into the intestinal lumen, where the toxin first encounters the apical surface of enterocytes; and 2. CPE is not internalized inside mammalian cells (McClane, 1994). Therefore, available experimental evidence strongly suggests that the primary intestinal effect of CPE is to kill enterocytes by inducing membrane permeability alterations. The consequence of that CPE-induced enterocyte death is the histopathlogical damage that apparently initiates fluid/electrolyte transport alterations. In this regard, it is notable that CPE has recently been shown to induce the apparent formation of pores in apical membranes of CaCo-2 cells (Hardy et al., 1999). CPE-induced paracellular permeability changes still might conceivably contribute to diarrhea, particularly during the later stages of CPE-associated gastrointestinal disease after intestinal histopathological damage has developed. It is possible that this histopathological damage provides the enterotoxin access to the basolateral surface of some enterocytes, a location from which CPE might affect TJs and alter paracellular permeability.

24.7 A CURRENT MODEL FOR CPE ACTION Studies of CPE action are currently advancing so rapidly that proposing a model for CPE action that is capable of standing the “test of time” becomes highly problematic. Nonetheless, the model for CPE action depicted in Figure 24.5 is compatible with the experimental data available as this chapter is being written. This model proposes: 1. CPE action initiates when the enterotoxin binds to a receptor(s). Recent studies (Katahira et al., 1997a, b; Fujita et al., 2000) have convincingly shown that several claudins can serve as functional CPE receptors capable of conveying a cytotoxic response. However, it remains unclear if these claudins represent the only, or even the major, receptors used during the intestinal action of CPE. The possibility of additional CPE receptors, or a CPE coreceptor, receives support from biochemical studies (Wnek and

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FIGURE 24.5 Model for CPE action. Step 1: CPE binds to a protein receptor(s), which may involve CPE binding to claudin-3 or claudin-4 alone (left), to either a claudin receptor or an independent ~45 to 50 kDa eucaryotic protein receptor (middle), or a simultaneous binding of CPE to both a claudin and the ~45 to 50 kDa eucaryotic protein coreceptor. Step 2: After binding, CPE localizes on the plasma membrane surface in an SDS-sensitive small complex of ~90 kDa. Step 3: At 37°C, this small complex interacts with other protein(s) to form an SDS-resistant, intermediate complex of ~135 kDa. Step 4: Interaction of the intermediate complex with occludin results in formation of an ~200 kDa complex; this interaction may alter TJ structure and function, possibly including paracellular permeability. Step 5: Interaction of the intermediate complex with other protein(s) results in formation of an ~155 kDa complex, which alters the permeability properties of the plasma membrane, killing the enterocyte. This cytolethal effect results in histopathological damage, which initiates the fluid/electrolyte transport alterations that occur during CPE-associated gastrointestinal disease.

McClane, 1983; Sugii and Horiguchi, 1988; Wieckowski et al., 1994) demonstrating that an ~45 to 50 kDa eucaryotic protein quickly associates with membrane-bound enterotoxin. Whether that ~45 to 50 kDa protein interacts with CPE during binding or, instead, interacts with a claudin–CPE complex soon after binding remains to be clarified. 2. Following binding, CPE quickly becomes localized in a small complex of ~90 kDa. That small complex contains CPE and the ~45 to 50 kDa

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eucaryotic protein referred to in step 1 of this model (Wieckowski et al., 1994); since claudin receptors have been associated with high Mr material forming after small complex (Katahira et al., 1997), it is presumed that a claudin receptor(s) is also present in small complexes. 3. At 4°C, CPE action does not proceed beyond formation of this small complex and the mammalian cell remains viable (McClane and Wnek, 1990). However, at physiological temperatures (where diffusion of membrane proteins is facilitated), the small complex interacts with unidentified protein(s) to form the ~135 kDa CPE complex (Singh et al., 2000). 4. Kinetic studies suggest this ~135 kDa CPE complex is an intermediate precursor for formation of the ~155 kDa and/or ~200 kDa CPE complexes (Singh et al., 2000). When the ~135 kDa intermediate complex and other (still unidentified) protein(s) interact, the ~155 kDa complex forms (Singh et al., 2000). This ~155 kDa complex appears to be sufficient for mediating the cytotoxic activity of the enterotoxin since Transwell cultures of CaCo-2 cells, which are highly sensitive to the lethal effects of CPE, form the ~155 kDa complex but do not form an appreciable amount of the ~200 kDa complex (Singh and McClane, unpublished data). Formation of the ~155 kDa complex, which may represent a porelike structure, results in massive membrane permeability alterations (McClane, 1994). Those permeability alterations are lethal for the mammalian cell (McClane, 1994), and consequently produce the histopathological damage that apparently initiates CPE-induced fluid and electrolyte losses from the intestines (Sherman et al., 1994). 5. Interaction of the ~135 kDa intermediate complex with occludin (which in vivo may occur after CPE-induced histopathological damage provides the enterotoxin with access to the basolateral surface of enterocytes), results in formation of the ~200 kDa large complex species (Singh et al., 2000). Since the ~200 kDa large complex contains occludin, and presumably a claudin receptor, formation of this CPE species may account for the rapid TJ disintegration observed in rat liver perfused on the basolateral surface with native CPE. Formation of this ~200 kDa large complex species might also result in paracellular permeability alterations, which could be a secondary contributor to the diarrheic symptoms of CPEassociated gastrointestinal diseases.

24.8 CONCLUDING COMMENTS Despite the exciting recent progress described in this chapter, many important questions remain unanswered regarding the CPE mechanism of intestinal action. For example, what is the ~45 to 50 kDa protein that associates with CPE in small complex (and also presumably in larger complexes)? Is that ~45 to 50 kDa protein a receptor or coreceptor for CPE? Do CPE-induced TJ changes and paracellular permeability alterations occur in the intestines during CPE-associated gastrointestinal disease? If so, how important are those effects for disease pathology?

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While the questions posed above represent only a few of the many challenges remaining for CPE researchers, it is already quite clear that this enterotoxin (and its derivative fragments) represents powerful new tools for probing TJ structure and function. Therefore, TJ researchers may wish to keep abreast of future developments in the field of CPE research.

ACKNOWLEDGMENTS Preparation of this chapter was supported, in part, by Public Health Service Grant AI-19844-17 from the National Institute of Allergy and Infectious Diseases. The authors thank Dr. Mahfuzur Sarker for assistance with graphics.

REFERENCES Bean, N. H., Goulding, J. S., Lao, C., and Angulo, F. J. 1996. Surveillance for foodbornedisease outbreaks — United States, 1988–1992, Morbid. Mortal. Weekly Rep., 45, 1. Carman, R. J. 1997. Clostridium perfringens in spontaneous and antibiotic-associated diarrhoea of man and other animals, Rev. Med. Microbiol., 8 (Suppl. 1), S43. Collie, R. E. and McClane, B. A. 1998. Evidence that the enterotoxin gene can be episomal in Clostridium perfringens isolates associated with non-foodborne human gastrointestinal diseases, J. Clin. Microbiol., 36, 30. Collie, R. E., Kokai-Kun, J. F., and McClane, B. A. 1998. Phenotypic characterization of enterotoxigenic Clostridium perfringens isolates from non-foodborne human gastrointestinal diseases, Anaerobe, 4, 69. Daube, G., Simon, P., Limbourg, B., Manteca, C., Mainil, J., and Kaeckenbeeck, A. 1996. Hybridization of 2,659 Clostridium perfringens isolates with gene probes for seven toxins (α, β, ε, ι, θ, µ and enterotoxin) and for sialidase, Am. J. Vet. Res., 57, 496. Fujita, K., Katahira, J., Horiguchi, Y., Sonoda, N., Furuse, M., and Tskuita, S. 2000. Clostridium perfringens enterotoxin binds to the second extracellular loop of claudin-3, a tight junction membrane protein, FEBS Lett., 476, 258. Gibert, M., Jolivet-Reynaud, C., and Popoff, M. R. 1997. Beta2 toxin, a novel toxin produced by Clostridium perfringens, Gene, 203, 65. Granum, P. E. and Richardson, M. 1991. Chymotrypsin treatment increases the activity of Clostridium perfringens enterotoxin, Toxicon, 29, 445. Granum, P. E., Whitaker, J. R., and Skjelkvale, R. 1981. Trypsin activation of enterotoxin from Clostridium perfringens type A. Biochim. Biophys. Acta, 668, 325. Hanna, P. C., Mietzner, T. A., Schoolnik, G. K., and McClane, B. A. 1991. Localization of the receptor-binding region of Clostridium perfringens enterotoxin utilizing cloned toxin fragments and synthetic peptides. The 30 C-terminal amino acids define a functional binding region, J. Biol. Chem., 266, 11037. Hanna, P. C., Wieckowski, E. U., Mietzner, T. A., and McClane, B. A. 1992. Mapping functional regions of Clostridium perfringens type A enterotoxin, Infect. Immun., 60, 2110. Hardy, S. P., Denmead, M., Parekh, N., and Granum, P. E. 1999. Cationic currects induced by Clostridium perfringens type A enterotoxin in human intestinal Caco-2 cells, J. Med. Microbiol., 48, 235. Hauser, D., Eklund, M. W., Boquet, P., and Popoff, M. R. 1994. Organization of the botulinum neurotoxin C1 gene and its non-toxic protein genes in Clostridium botulinum C 468, Mol. Gen. Genet., 243, 631.

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Katahira, J., Inoue, N., Horiguchi, Y., Matsuda, M., and Sugimoto, N. 1997. Molecular cloning and functional characterization of the receptor for Clostridium perfringens enterotoxin, J. Cell Biol., 136, 1239. Katahira, J., Sugiyama, H., Inoue, N., Horiguchi, Y., Matsuda, M., and Sugimoto, N. 1997. Clostridium perfringens enterotoxin utilizes two structurally related membrane proteins as functional receptors in vivo, J. Biol. Chem., 272, 26652. Kokai-Kun, J. F. and McClane, B. A. 1997. Deletion analysis of the Clostridium perfringens enterotoxin, Infect. Immun., 65, 1014. Kokai-Kun, J. F., Songer, J. G., Czeczulin, J. R., Chen, F., and McClane, B. A. 1994. Comparison of Western immunoblots and gene detection assays for identification of potentially enterotoxigenic isolates of Clostridium perfringens, J. Clin. Microbiol., 32, 2533. Kokai-Kun, J. F., Benton, K., Wieckowski, E. U., and McClane, B. A. 1997. Identification of a Clostridium perfringens enterotoxin region required for large complex formation and cytotoxicity by random mutagenesis, Infect. Immun., 67, 6534. McClane, B. A. 1994. Clostridium perfringens enterotoxin acts by producing small molecule permeability alterations in plasma membranes, Toxicology, 87, 43. McClane, B. A. 1997. Clostridium perfringens, in Food Microbiology: Fundamentals and Frontiers, Doyle, M. P., Beuchat, L. R., and Montville, T. J., Eds., ASM Press, Washington, D.C., 307. McClane, B. A. and McDonel, J. L. 1979. The effects of Clostridium perfringens enterotoxin on morphology, viability and macromolecular synthesis, J. Cell Physiol., 99, 191. McClane, B. A. and Wnek, A. P. 1990. Studies of Clostridium perfringens enterotoxin action at different temperatures demonstrate a correlation between complex formation and cytotoxicity, Infect. Immun., 58, 3109. McClane, B. A., Lyerly, D. M., Moncrief, J. S., and Wilkins, T. D. 2000. Enterotoxic clostridia: Clostridium perfringens type A and Clostridium difficile, in Gram-Positive Pathogens, Fischetti, V. A., Novick, R. P., Ferretti, J. J., Portnoy, D. A., and Rood, J. I., Eds., ASM Press, Washington, D.C., 551. McDonel, J. L. 1986. Toxins of Clostridium perfringens types A, B, C, D, and E, in Pharmacology of Bacterial Toxins, Dorner, F. and Drews, H., Eds., Pergamon Press, Oxford, 477. McDonel, J. L. and Demers, G. W. 1982. In vivo effects of enterotoxin from Clostridium perfringens type A in rabbit colon: binding vs. biologic activity, J. Infect. Dis., 145, 490. McDonel, J. L. and Duncan, C. L. 1975. Histopathological effect of Clostridium perfringens enterotoxin in the rabbit ileum, Infect. Immun., 12, 1214. McDonel, J. L. and Duncan, C. L. 1977. Regional localization of activity of Clostridium perfringens type A enterotoxin in the rabbit ileum, jejunum and duodenum, J. Infect. Dis., 136, 661. McDonel, J. L., Chang, L. W., Pounds, J. L., and Duncan, C. L. 1978. The effects of Clostridium perfringens enterotoxin on rat and rabbit ileum: an electron microscopy study, Lab. Invest., 39, 210. Morita, K., Furuse, M., Fujimoto, K., and Tsukimoto, S. 1999. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands, Proc. Natl. Acad. Sci. U.S.A., 96, 511. Rahner, C., Mitic, L. L., McClane, B. A., and Anderson, J. M. 1999. Clostridium perfringens enterotoxin impairs bile flow in the isolated perfused rat liver and induces fragmentation of tight junction fibrils, Hepatology, 30, 326A. Rood, J. and Cole, S. T. 1991. Molecular genetics and pathogenesis of Clostridium perfringens, Microbiol. Rev., 55, 621.

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Sarker, M. R., Carman, R. J., and McClane, B. A. 1999. Inactivation of the gene (cpe) encoding Clostridium perfringens enterotoxin eliminates the ability of two cpe-positive C. perfringens type A human gastrointestinal disease isolates to affect rabbit ileal loops, Mol. Microbiol., 33, 946. Sarker, M. R., Shivers, R. P., Sparks, S. G., Juneja, V. K., and McClane, B. A. 2000. Comparative experiments to examine the effects of heating on vegetative cells and spores of Clostridium perfringens isolates carrying plasmid versus chromosomal enterotoxin genes, Appl. Environ. Microbiol., 66, 3234. Sherman, S., Klein, E., and McClane, B. A. 1994. Clostridium perfringens type A enterotoxin induces concurrent development of tissue damage and fluid accumulation in the rabbit ileum, J. Diarrheal Dis. Res., 12, 200. Singh, U., Van Itallie, C. M., Mitic, L. L., Anderson, J. M., and McClane, B. A. 2000. CaCo-2 cells treated with Clostridium perfringens enterotoxin form multiple large complex species, one of which contains the tight junction protein occludin, J. Biol. Chem., 275, 18407. Skjelkvale, R. and Uemura, T. 1977. Experimental diarrhea in human volunteers following oral administration of Clostridium perfringens enterotoxin, J. Appl. Bacteriol., 46, 281. Songer, J. G. and Meer, R. M. 1996. Genotyping of Clostridium perfringens by polymerase chain reaction is a useful adjunct to diagnosis of clostridial enteric disease in animals, Anaerobe, 2, 197. Sonoda, N., Furuse, M., Sasaki, H., Yonemura, S., Katahira, J., Horiguchi, Y., and Tsukita, S. 1999. Clostridium perfringens enterotoxin fragments remove specific claudins from tight junction strands: evidence for direct involvement of claudins in tight junction barrier, J. Cell Biol., 147, 195. Sugii, S. and Horiguchi, Y. 1988. Identification and isolation of the binding substance for Clostridium perfringens enterotoxin on Vero cells, FEMS Microbiol. Lett., 52, 85. Wieckowski, E. U., Wnek, A. P., and McClane, B. A. 1994. Evidence that an ~50kDa mammalian plasma membrane protein with receptor-like properties mediates the amphiphilicity of specifically-bound Clostridium perfringens enterotoxin, J. Biol. Chem., 269, 10838. Wieckowski, E., Kokai-Kun, J. F., and McClane, B. A. 1998. Characterization of membraneassociated Clostridium perfringens enterotoxin following Pronase treatment, Infect. Immun., 66, 5897. Wnek, A. P. and McClane, B. A. 1983. Identification of a 50,000 Mr protein from rabbit brush boarder membranes that binds Clostridium perfringens enterotoxin, Biochem. Biophys. Res. Commun., 112, 1099. Wnek, A. P. and McClane, B. A. 1989. Preliminary evidence that Clostridium perfringens type A enterotoxin is present in a 160,000-Mr complex in mammalian membranes, Infect. Immun., 57, 574.

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Ischemia-Induced Tight Junction Dysfunction in the Kidney James A. Marrs and Bruce A. Molitoris

CONTENTS 25.1 Introduction .................................................................................................533 25.1.1 The Kidney ....................................................................................533 25.1.2 Renal Ischemia ..............................................................................535 25.2 Renal Ischemia Disrupts Tight Junction Function .....................................536 25.3 Tight Junction Regulation and Ischemia ....................................................538 25.3.1 Actin Cytoskeleton Dysfunction and Tight Junction Regulation ......................................................................................539 25.3.2 Protein Kinases and Tight Junction Dysfunction .........................539 25.3.3 GTPase Regulation and Tight Junction Dysfunction....................540 25.3.4 Other Regulatory Pathways That May Be Disrupted by Ischemia .........................................................................................541 25.4 Summary ......................................................................................................545 Acknowledgments..................................................................................................546 References..............................................................................................................546

25.1 INTRODUCTION 25.1.1 THE KIDNEY The primary functions of the kidney, maintaining body volume and electrolyte balance, while removing nitrogenous and other waste products from the blood, are dependent upon size-selective filtration across the glomerulus and specific reabsorption of water, ions, and macromolecules by tubular epithelial cells. Regulation of these interrelated processes at the cellular level requires structural and biochemical polarization of the surface membrane of participating epithelial cells. For example, with normal renal function, approximately 150 liters of glomerular filtrate are produced every 24 h. About 99% of this volume is reabsorbed, and between 70 and 80% of this reabsorption occurs across proximal tubule cells. Therefore, small

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changes in reabsorptive characteristics across these cells can have profound effects on organ function. The nephron is composed of several distinct epithelial cell types, each having its own set of specialized functions and surface membrane characteristics. Tight junction (TJ) permeability varies over the length of the nephron becoming progressively tighter throughout the length of the nephron. Thus, proximal tubule cells have a low transepithelial electrical resistance (TER), whereas distal tubule cells, such as cortical collecting tubule cells, have high resistances (Brown and Stow, 1996). Proximal tubule cells possess a distinctive apical membrane, comprising microvilli, separated from a highly invaginated basolateral membrane domain by cellular junctional complexes, as shown in Figure 25.1A (Maunsbach and Christensen, 1992).

FIGURE 25.1 Ischemia opens TJs between proximal tubule but not between distal tubule cells. Selective ruthenium red staining of the apical plasma membrane domains of tubular epithelial cells and rapid gluteraldehyde fixation were accomplished using microperfusion techniques. Ruthenium red staining did not penetrate the intact TJs of proximal tubule cells of control kidneys in vivo (A), as shown by the white arrows. With as little as 5 min of ischemia, ruthenium red penetrated some (dark arrows) but not all TJs (B). After 15 (C) and 30 min (D) of ischemia 50 and 62% of proximal tubules TJs had ruthenium red penetration, respectively (Molitoris et al., 1989b). Distal tubule cell TJs were not disrupted by 30 min of ischemia (E, F). (Reprinted with permission of the American Society for Clinical Investigation.)

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These characteristics serve to enhance greatly the surface area of both membrane domains. As in other epithelial cells, TJs form a physical barrier separating biological compartments (urinary lumen from the blood). They also maintain unique membrane domain compositions for phospholipids and proteins such as ion channels, transport proteins, and specific enzymes (Brown and Stow, 1996). Thus, the integrity of these polarized plasma membrane domains and junctional complexes are critical for normal kidney function.

25.1.2 RENAL ISCHEMIA Maintenance of cellular integrity requires a continuous and adequate supply of substrate and oxygen for cellular energy production. Tissue ischemia results when blood flow is inadequate to maintain cellular ATP concentrations above a level required for homeostasis. Decreases in cellular ATP below 15% of physiological conditions are associated with cell death by necrosis, whereas ATP levels in the 25% range, for prolonged periods, are more likely associated with cell death by apoptosis (Lieberthal et al., 1998). Clinically, ischemic acute renal failure results when proximal tubule cell injury leads to acute organ dysfunction manifested primarily by a severe reduction in the glomerular filtration rate (GFR). In moderate and severe acute renal failure, dialysis is often required to replace renal function until cellular repair occurs and renal function has returned. Ischemia is the major cause of acute renal failure in hospitalized patients. Renal ischemia results primarily from events leading to severe reductions in renal blood flow such as cross-clamping the aorta during vascular surgery, excessive bleeding, bacterial sepsis, dehydration, or the use of therapeutic agents known to induce renal vasoconstriction such as radiocontrast dye. To mimic these clinical situations of ischemic acute renal failure, and to allow for further understanding at the cellular and molecular level, several complementary animal and cell culture models have been developed. In rats, cross-clamping the renal artery is used to induce variable durations, and therefore differing severity, of ischemic injury. In cultured cells, various mitochondrial inhibitors of oxidative phosphorylation, such as antimycin A or cyanide, are used in conjunction with either substrate depletion or inhibitors of glucose utilization, such as 2-deoxyglucose, to induce ATP depletion (Lieberthal and Nigam, 2000). Both animal models and tissue-culture cell ATP-depletion studies (chemical ischemia) recapitulate the pathological events known to occur during clinical renal ischemia (Lieberthal and Nigam, 2000). Therefore, these models have been used extensively to unravel the functional consequences and mechanisms of ischemia-induced TJ alterations. Ischemia-induced opening of proximal tubule cell TJs results in important clinical effects. These include a reduction in ion transport, especially Na+. This results in high distal delivery of Na+, K+, and Cl– and high distal flow rates. These factors cause afferent arteriole vasoconstriction and its attendant reduction in GFR, via a mechanism termed tubular glomerular feedback (Braam et al., 1993). Ischemiainduced TJ opening also results in increased and unregulated paracellular movement of ions and water, termed backleak (Donohoe et al., 1978). Both backleak and afferent arteriole vasoconstriction are important determinants in ischemia-induced reductions

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in GFR. Backleak also results in excessive accumulation of extracellular fluid resulting in tissue edema. Therefore, understanding the mechanisms of ischemia-induced TJ dysfunction may have important clinical and therapeutic implications. This chapter focuses on how ischemia affects TJ structure and function in proximal tubule cells. The potential role of actin cytoskeletal alterations in the overall process of junctional complex disruption will also be discussed.

25.2 RENAL ISCHEMIA DISRUPTS TIGHT JUNCTION FUNCTION Observations in the rat cross-clamp model revealed rapidly occurring, durationdependent opening of proximal tubule cell TJs (Molitoris et al., 1989b). In vivo ruthenium red microperfusion of renal tubules studies were performed. This procedure labels only the apical plasma membrane of perfused tubules with a functional TJ, and the label was prevented from accessing the basal-lateral plasma membrane domain. After the ischemic insult, ruthenium red staining permeated the TJ and was detected on the basal-lateral plasma membrane, indicating that an early consequence of ischemia was TJ barrier breakdown. As little as 5 min of ischemia was sufficient to induce the breakdown of the TJ barrier to ruthenium red (see Figure 25.1). Longer periods of ischemia caused greater proportions of proximal tubule TJs to become permeable. After only 15 min of ischemia, half the proximal tubule cells showed paracellular leakage through the TJ, with corresponding labeling of the basal-lateral plasma membrane domain (Molitoris et al., 1989b). These alterations in paracellular permeability were also accompanied by defects in surface membrane polarity (Fish and Molitoris, 1994). Lipid and Na+,K+-ATPase polarity was altered after short periods of ischemia (5 to 10 min) (Molitoris et al., 1985; 1988; 1989a, b), suggesting that lateral diffusion of membrane proteins and lipids between the apical and basallateral plasma membrane domains (through the TJ diffusion barrier) was responsible for the loss in surface membrane polarity. The TJs of distal tubule cells were unaffected by this duration of ischemia (see Figure 25.1E and F) (Molitoris et al., 1989b). This later result may be explained by the observation that distal tubule cells are more resistant to the cellular defects caused by similar levels of ATP depletion (Sheridan et al., 1993). Effects of ischemia on TJs were studied in human patients who had received renal allografts. Brief periods of warm ischemia during removal of kidneys from donors and extended periods (several hours) of cold ischemia result from the cold storage of the donor organ. Advances in storage perfusion solutions have reduced damage from cold ischemia (Southard and Belzer, 1995), but ischemic injury in renal allografts remains a significant factor important in transplant success (Scandling and Myers, 1997). Myers and colleagues studied numerous clinical measurements and cell biological features in post-transplant patients (Alejandro et al., 1995a, b; Kwon et al., 1998; 1999), including the examination of TJ dysfunction (Kwon et al., 1998). By using the renal allograft model, there was a measured ischemic event at the time of transplant that allowed the investigators to correlate the time interval of cold and warm ischemia with clinical and cell biological findings.

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The renal allograft model also permits the study of ischemic injury in the absence of ongoing renal disease. In addition, this powerful experimental approach showed that the ischemic injury to epithelial cell TJs in human patients was recapitulated by the rat cross-clamp model, further supporting the validity of this animal model. Patients in these studies were classified as having either sustained or recovering acute renal failure according to clinical features 3 to 7 days following transplantation, including glomerular filtration rate and urine output (Kwon et al., 1998). Patients with sustained acute renal failure showed poor recovery of function, and dialysis was required in most cases. Patients with recovering acute renal failure had prompt recovery of function post-transplant, and these patients did not require dialysis. To examine paracellular barrier dysfunction in the kidney, a series of measurements were made of the clearance of molecules from the blood to the urine (Kwon et al., 1998). Differential clearance of neutral dextran molecules of various sizes vs. the clearance of inulin, a small, uniformly sized molecule, was used to infer the amount of backleak of small molecules. Large dextran molecules generally do not pass the paracellular barrier, even if it is dysfunctional. Therefore, backleak can be calculated from the ratio of dextran clearance to inulin clearance. There was no difference in this ratio between control patients and patients with recovering acute renal failure, but the patients with sustained acute renal failure showed increased backleak in the allograft (Kwon et al., 1998). This study directly correlated the severity of acute renal failure in humans with TJ dysfunction. Myers and colleagues followed these observations by examining the distribution of TJ and adherens junction components by immunofluorescence in kidney biopsy tissue taken at the time of transplant and 7 days post-transplantation (Kwon et al., 1998). In control experiments, normal TJ and adherens junction component distributions were found in tubular epithelial cells from prenephrectomy biopsies of living kidney donors. Distributions of TJ and adherens junction components were not severely disrupted in patients categorized as recovering acute renal failure by clinical findings. In contrast, TJ and adherens junction components showed significantly altered distributions in patients with sustained acute renal failure. Comparing distributions of TJ and adherens junction components on the day of transplantation and 7 days post-transplantation showed that the altered distributions persisted in those patients with delayed renal allograft function, requiring dialysis (Kwon et al., 1998). In addition, Myers and colleagues studied effects on cell-to-substrate junctional complex protein (integrin and laminin) distributions (Kwon et al., 1998). The substrate adhesion receptor proteins α6-integrin and laminin showed no significantly altered distributions in biopsies from patients with sustained acute renal failure relative to biopsies from patients with recovering acute renal failure. Matlin and colleagues (Zuk et al., 1998) also studied the effects of ischemia on the substrate adhesion system using the rat cross-clamp model. These investigators showed that reperfusion injury, not the initial ischemia, leads to redistribution of integrins from a basal membrane distribution to one that includes lateral and apical plasma membrane domains. Matlin and colleagues (Zuk et al., 1998) also showed that distributions of basement membrane components (laminin and collagen IV) were not altered by ischemia/reperfusion injury. However, fibronectin accumulated in the lumen of renal tubules during reperfusion injury. This material may have been derived from

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the renal interstitium or from plasma sources due to denudation of the basement membrane (tubulorrhexis) and breakdown of the paracellular barrier (Zuk et al., 1998). In addition to producing the paracellular barrier, TJs also restrict the lateral diffusion of integral membrane proteins from mixing between the apical and basallateral plasma membrane domains. Dysfunction of the TJ could lead to changes in cell polarity. After short and moderate ischemia induced in the rat cross-clamp model, Na+,K+-ATPase and lipid polarity were disrupted (Molitoris et al., 1985; 1988; 1989a, b). Na+,K+-ATPase polarity effects were confirmed in tissue-culture cell models of ischemia, suggesting that ATP depletion directly leads to polarity defects (Canfield et al., 1991; Molitoris et al., 1991; Mandel et al., 1994). In addition, altered surface membrane polarity correlated with rearrangements in the actin cytoskeleton (Kellerman et al., 1990). Apically localized Na+,K+-ATPase remains functional and transports sodium across the apical membrane (Molitoris, 1993). As a working hypothesis, it was suggested that disrupting normal actin dynamics allows membrane proteins greater diffusion in the plasma membrane, and disrupting TJs allows membrane proteins to diffuse between the basal-lateral and apical plasma membrane domains (Molitoris, 1991). However, a detailed analysis in the Madin–Darby canine kidney (MDCK) cell model showed that the paracellular barrier was rapidly (10 min) compromised during ATP depletion, but the lateral diffusion barrier to lipids in the plasma membrane was not disrupted after 30 min of ATP depletion (Mandel et al., 1993). Different experimental models were used in various studies, and there may be differences in the amount of lateral diffusion that is permitted by proximal and distal cell types following ischemic injury. So, lateral diffusion of membrane proteins and lipids across the TJ barrier may play a part in the pathophysiological processes that lead to loss of normal cell polarity. In cultured cell models and in biopsy tissues from human transplant patients, ischemic injury leads to internalization of Na+,K+-ATPase from the basal-lateral plasma membrane (Mandel et al., 1994; Alejandro et al., 1995b; Kwon et al., 1999). Na+,K+-ATPase that was endocytosed could be misrouted to the apical plasma membrane. Another explanation for the origin of apical Na+,K+-ATPase could be that ischemic injury produces defects in polarized sorting mechanisms in the biosynthetic pathway. This effect could also explain altered polarity of other membrane proteins, for example, integrins (Zuk et al., 1998). The physiological consequences of open TJs and mislocalized Na+,K+-ATPase in renal tubular epithelial cells are devastating in acute renal failure. Understanding specific cellular processes that are altered by ischemia, leading to tubular cell dysfunction, is of utmost importance to designing better treatment strategies for acute renal failure.

25.3 TIGHT JUNCTION REGULATION AND ISCHEMIA Many factors regulate TJ assembly and function. These factors were reviewed in earlier chapters of this volume (see Chapters 15 through 17). This chapter discusses factors disrupted during ischemic injury to renal epithelial cells that regulate TJs and speculates on other TJ regulatory pathways that may be disrupted by ischemia. It is necessary to state that understanding of TJ regulation is still rudimentary, and,

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TABLE 25.1 Tight Junction Regulatory Mechanisms Potentially Affected by Ischemia GTPase inhibition Protein kinase signaling Actin cytoskeleton

Demonstrated Effects GTP-depletiona and Rho inhibitionb General inhibition of protein kinasesc Disorganization of actin cytoskeletond

Suspected Effects Inhibition of other GTPases, including Rac, Cdc42, Ras, and heterotrimeric G proteins Specific inhibition of protein kinases that regulate TJs (protein kinase C and MAP kinase) Myosin II dysfunction; specific regulatory mechanisms disrupted as a consequence of protein kinase inhibition and Rho-family GTPases

a

Dagher (2000). Hallet and Atkinson (in preparation). c Kobryn and Mandel (1994). d Reviewed in Molitoris (1997). b

thus, the list of known TJ regulatory pathways that are disrupted during ischemia will grow. Known TJ regulatory factors that are negatively affected by ATP depletion and ischemia include the stabilizing force of the actin cytoskeleton, protein kinase regulatory pathways, and Rho GTPase factors (Table 25.1).

25.3.1 ACTIN CYTOSKELETON DYSFUNCTION JUNCTION REGULATION

AND

TIGHT

The cortical actin ring that associates with the apical junctional complex (Hirokawa and Tilney, 1982) regulates TJ assembly and function (Madara, 1998). During ischemia, actin cytoskeleton structures are massively rearranged (Molitoris, 1997; Sutton and Molitoris, 1998; Wagner and Molitoris, 1999). Breakdown of apical microvillar cytoskeleton is the most obvious early effect of ischemic injury (Ashworth and Molitoris, 1999). TJ dysfunction is also a very early consequence of ischemia and ATP depletion (Molitoris et al., 1989b; Mandel et al., 1993). However, cortical actin cytoskeleton rearrangements and changes in TJ ultrastructure (freezefracture strands) are not altered until several minutes following the opening of the paracellular barrier (Mandel et al., 1993; Bacallao et al., 1994). Thus, TJ dysfunction is an early effect that precedes detectable morphological changes in the TJ and cortical actin cytoskeleton rearrangements. These and other observations suggest that the effect of ischemia and ATP depletion causing TJ dysfunction may be independent of effects on the cortical actin cytoskeleton, perhaps affecting TJ components directly through intracellular signaling pathways.

25.3.2 PROTEIN KINASES

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TIGHT JUNCTION DYSFUNCTION

Protein phosphorylation intracellular signal transduction pathways regulate TJ permeability and TJ component assembly (Madara, 1998). Both tyrosine kinase and serine/threonine kinase regulation of TJs have been documented (Ojakian, 1981; Mullin and O’Brien, 1986; Madara, 1988; Balda et al., 1993; Saxon et al., 1994;

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Citi and Denisenko, 1995; Staddon et al., 1995; Stuart and Nigam, 1995; Takeda and Tsukita, 1995; Van Itallie et al., 1995; Dodane and Kachar, 1996; Izumi et al., 1998; Kovbasnjuk et al., 1998; Tsukamoto and Nigam, 1999; Chen et al., 2000). During ischemia, ATP levels drop rapidly. ATP concentrations fall to levels that are below the Km for most protein kinases (Kobryn and Mandel, 1994). However, protein phosphatases remain active (Kobryn and Mandel, 1994). The consequence of ATP depletion for most phosphoproteins is rapid reduction in their phosphoamino acid content. Thus, ischemia is expected to disrupt most protein kinase–signaling pathways. ATP depletion of MDCK cells caused a rapid decline in phosphoamino acid content in ZO-1, ZO-2, and occludin (Gopalakrishnan et al., 1998). TJ regulation and assembly mechanisms that are dependent on protein kinase signaling will be severely compromised.

25.3.3 GTPASE REGULATION

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The authors have hypothesized that Rho GTPase family members, signaling molecules that regulate actin cytoskeleton and junctional complex assembly, are inactivated during ATP depletion. Many of the changes observed during ischemia, such as TJ dysfunction, breakdown of actin structures, and loss of cell–substrate adhesion (Hall, 1998), are similar to effects of inhibiting Rho GTPase family member signaling. Indeed, Dagher (2000) showed that ATP depletion is accompanied by a concomitant reduction in cellular GTP levels. Hallett and Atkinson (in preparation; personal communication) have examined the consequence of ATP depletion on Rho activity. They showed directly that Rho is very rapidly inactivated during ATP depletion (reduced Rho activity was detected after 5 min of ATP depletion), and Rho activity is restored during ATP repletion. The authors have also shown in experiments using constitutive active and dominant negative mutant Rho protein expression that Rho signaling protects TJs from disassembly during ATP depletion (Gopalakrishnan et al., 1998). Inhibiting Rho by expressing a dominant negative mutant form in MDCK cells caused more extensive loss of TJ components from cell contact sites during ATP depletion than control cells not expressing mutant Rho (Figure 25.2). MDCK cells expressing constitutively activated Rho mutant maintained TJs during ATP depletion significantly better than control cells (Figure 25.2). Thus, activating Rho signaling protects TJs from damage during ATP depletion (Gopalakrishnan et al., 1998) (Figure 25.2). In contrast, inhibiting or activating Rho in MDCK cells did not alter adherens junction disassembly during ATP depletion relative to control cells not expressing Rho mutants, showing that Rho signaling protective effects were specific for the TJ (Gopalakrishnan and Marrs, submitted). It may be that Rac or Cdc42 provide a protective signal for the adherens junction during injury. These results also indicate that the protection of TJs during ATP-depletion injury was not an indirect consequence of protecting adherens junctions. In addition to the protection of TJs by Rho signaling, cortical actin structures and stress fibers were also protected from disassembly during ATP depletion (Raman and Atkinson, 1999). Using microinjection experiments, Raman and Atkinson (1999) showed that the effect of Rho signaling was not required prior to or during ATP

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depletion. Cells injected with constitutive active Rho protein (Rho-V14) after ATP depletion recovered cortical actin cytoskeleton and stress fibers more rapidly than control, uninjected cells. In contrast, the microvillar actin cytoskeleton was not protected by Rho signaling. It is possible that Rac or Cdc42 protect the microvillar structure. These experiments illustrate the potential that Rho activation may be a therapeutic target for protecting kidney epithelial cells from ischemic injury and facilitating recovery from injury.

25.3.4 OTHER REGULATORY PATHWAYS THAT MAY BE DISRUPTED BY ISCHEMIA The observed effects of ischemia and ATP depletion on TJ regulatory mechanisms raise distinct possibilities that other regulatory pathways will be affected by ischemia and ATP depletion. The authors speculate that these pathways include the myosin contractility associated with the cortical actin ring, specific protein kinase regulatory pathways, and other GTP-binding proteins. These effects of ischemia and ATP depletion are only hypothetical, but they provide a framework for future investigations. Ischemia may affect contractility of the cortical actin ring by influencing myosin function. Myosin contractility appears to be a significant regulator of TJ function (Bentzel et al., 1980; Madara et al., 1987; Hecht et al., 1996; Turner et al., 1997; 1999; Kovbasnjuk et al., 1998), but mechanistic details of how myosin contraction affects TJ permeability are still unclear. It may be that tension produced by contraction of the cortical actin ring physically regulates TJ permeability (Figure 25.2). ATP depletion would tend to cause a rigor state of myosin, destroying the contractile regulation of the TJ. Effects of ischemia and ATP depletion on myosin II functional regulation have not been studied. However, myosin I distribution and association with the actin cytoskeleton is rearranged during ischemia and ATP depletion (Wagner and Molitoris, 1997), consistent with the idea that ATP depletion will lead to a rigor state for myosin. Future studies should examine the effect of ischemia and ATP depletion on myosin contractility in cortical actin ring tension. Myosin contraction is regulated by myosin light-chain phosphorylation (Somlyo and Somlyo, 1994), and myosin light-chain phosphorylation state is regulated by numerous factors, including protein kinase C and Rho signaling (Hecht et al., 1994; Amano et al., 1996; Kimura et al., 1996; Fukata et al., 1999b; Turner et al., 1999; Totsukawa et al., 2000), which also regulate TJs. These regulatory pathways may be profoundly altered by ischemia and ATP depletion (see below). Therefore, direct effects on myosin activity due to ATP depletion and indirect effects on myosin activity due to disruption of intracellular signaling pathways could act together and disrupt normal myosin regulation of TJs during ischemia and ATP depletion (Figure 25.3). There is a coordinate reduction in cellular GTP levels that accompanies ATP depletion (Dagher, 2000). GTP-binding proteins may be inactivated as a consequence of this GTP depletion, which would influence numerous GTP-binding protein signaling pathways. Trimeric G proteins are localized to the TJ and regulate its function (Denker et al., 1996; Dodane and Kachar, 1996; Saha et al., 1998). In addition, small GTP-binding proteins of the Ras superfamily regulate TJ assembly and function

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FIGURE 25.2 Effects of Rho signaling on ATP-depletion-dependent redistribution of occludin. MDCK cells were transiently transfected with (A) myc-tagged dominant negative Rho (Rho-N19) or (B) myc-tagged dominant active Rho (Rho-V14). Cells expressing Rho mutant proteins were ATP-depleted for 60 min (60 Minutes ATP-Depletion) or left untreated (Control). (A and B) Double-label, indirect immunofluorescence was performed using occludin antibodies (Occludin) and anti-myc antibodies (Myc) to detect mutant Rho GTPase expressing cells. (C and D) Mean fluorescence intensity was calculated for junctions between pairs of mutant Rho GTPase expressing cells (transfected; arrows) and between pairs of untransfected cells from the same experiment. A dashed line at 1 shows the mean fluorescence intensity ratio in untransfected junctions for reference. (E) Schematic, graphical interpretation of the role for Rho signaling in TJ protection during ATP depletion. ATP depletion induces disassembly of TJs (normal ischemia). Disassembly during ATP depletion was more extensive in cells in which Rho signaling was inhibited (decreased Rho signaling), and increasing Rho signaling reduced disassembly during ATP depletion (increased Rho signaling). Maintenance of TJ integrity during renal epithelial cell injury may help maintain normal cell polarity and transport functions and facilitate recovery from ischemic injury. (Reprinted with permission of the American Society for Clinical Investigation.)

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FIGURE 25.2 (continued.)

FIGURE 25.3 Illustration of cortical actin ring regulation of TJ permeability. Appropriate cortical actin ring tension can be maintained by myosin regulation, allowing the TJ to prevent paracellular transport. With increased myosin tension (myosin contraction), the cortical actin ring would contract, which could physically regulate the TJ seal and open the paracellular permeability barrier. Reduced tension may also open the paracellular permeability barrier. Disruption of the actin cytoskeleton during ischemia would uncouple the cortical actin ring from the TJ. Tension from the actin ring could not be produced. Also, myosin function may be inhibited by ATP depletion (see text). Together, the consequence of ischemia on the actin cytoskeleton may cause the paracellular permeability barrier to open.

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(Schoenenberger et al., 1991; Nusrat et al., 1995; Takaishi et al., 1997; Yamamoto et al., 1997; Gopalakrishnan et al., 1998; Jou et al., 1998; Woo et al., 1999; Hopkins et al., 2000; Li and Mrsny, 2000). In particular, Ras, Rho, and Rac were shown to regulate TJs. Ras transformation of epithelial cells was analyzed in several studies, and results showed that transforming Ras protein mutants (that activate Ras signaling) expressed in epithelial cells disrupted the normal epithelial phenotype (Hay and Zuk, 1995). Ras transformation caused an epithelial-to-mesenchymal transformation in differentiation and disassembly of adherens junctions and TJs. Ras signaling regulates the cadherin/catenin complex that can indirectly control TJ assembly (Schoenenberger et al., 1991; Kinch et al., 1995). Disrupting cadherin adhesion can lead to the disruption of TJ assembly. Preventing cadherin function using function-blocking antibodies or genetically, for example, in cells lacking α-catenin expression, also prevents TJ assembly (Gumbiner and Simons, 1986; Gumbiner et al., 1988; Watabe et al., 1994). Based on these findings, it is possible that inhibiting cadherin function through Ras transformation may cause TJ disassembly. However, expression of extracellular domain–deleted mutant cadherin in transfected epithelial cell lines had paradoxical effects on TJ assembly. Overexpressing this mutant cadherin molecule caused post-translational downregulation of the endogenous cadherin molecules (Nieman et al., 1999; Troxell et al., 1999). The rate of TJ assembly was inhibited in mutant cadherin-expressing cells (Troxell et al., 1999), but despite severe impairment of the cadherin adhesion system, the final extent of TJ assembly was increased in cells overexpressing this mutant cadherin molecule (Troxell et al., 2000). Therefore, regulation of TJ assembly by cadherin adhesion molecules is a complex process and requires additional investigation to clarify regulatory pathways and mechanisms. Ras signaling targets include the molecule AF-6 that is also called afadin (which has two alternatively spliced variants, the longer form, l-afadin, and the shorter form, s-afadin) (Mandai et al., 1997). Afadin binds actin filaments and has one PDZ domain (Mandai et al., 1997) that binds an immunoglobulin superfamily, homophilic cell adhesion molecule, nectin (Takahashi et al., 1999). Afadin also binds ponsin, a SH3 domain-containing protein (Mandai et al., 1999). Afadin was originally localized to TJs, and it forms a complex with ZO-1 (Yamamoto et al., 1997). The afadin/nectin complex recruits the cadherin/catenin complex to sites of cell–cell contact, which may be an early step in adherens junction assembly (Tachibana et al., 2000). Afadin knockout mice fail in development because they fail to assemble normal adherens junctions and TJs in the early embryonic ectodermal epithelium (Ikeda et al., 1999). Therefore, this molecule is an important regulator of both adherens junction and TJ assembly. Because Ras signaling may regulate the alfadin/nectin complex (Yamamoto et al., 1997), disruption of Ras signaling during ischemia could cause dysfunction of the alfadin/nectin complex, and thus dysfunction of junctional complex assembly. Ras signal transduction pathways control several cellular processes. Ras signaling through the mitogen-activated protein (MAP) kinase pathway regulates TJ assembly (Chen et al., 2000; Li and Mrsny, 2000). MAP kinase signaling appears to target selectively occludin protein expression (Li and Mrsny, 2000). A salivary gland cell line was transfected with an oncogenic Raf-1 (MAP kinase) construct,

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which disrupted normal epithelial phenotype, and occludin expression was suppressed. Forcing the expression of occludin by transfecting occludin cDNA into the Raf-1 transformed cells reestablished a more normal epithelial phenotype and reestablished the paracellular permeability barrier (Li and Mrsny, 2000). Effects of inhibiting MAP kinase in normal cells on TJ assembly have not been investigated. In ischemia, protein kinases are generally inactivated (Kobryn and Mandel, 1994), and this general effect on protein kinases may include effects on TJ assembly caused by MAP kinase inhibition. Rho, Rac, and Cdc42 signaling pathways have effects on junctional complex assembly in epithelial cells (Nusrat et al., 1995; Braga et al., 1997; Hordijk et al., 1997; Kuroda et al., 1997; 1998; Takaishi et al., 1997; Gopalakrishnan et al., 1998; Jou et al., 1998; Sander et al., 1998; 1999; Fukata et al., 1999a; Kaibuchi et al., 1999). Rho and Rac show dramatic effects on adherens junction and TJ assembly. Activating Rho and Rac increases adherens junction and TJ assembly, and inhibiting these GTPases decrease both adherens junction and TJ assembly. Signal transduction pathways used by Rho GTPase family members that regulate junctional complex assembly are still unclear. Rho activates Rho kinase, which leads to increased myosin activity by inhibiting myosin phosphatases activity (Amano et al., 1996; Kimura et al., 1996; Fukata et al., 1999b), regulating myosin activity and, thus, paracellular permeability (see above). General protein kinase inhibition during ischemia is likely to inhibit this Rho kinase pathway. The effects of ischemia on TJ regulation are still poorly understood. Clear effects on the actin cytoskeleton, protein kinase signaling, and Rho GTPase signaling are likely to represent only a fraction of negative consequences on these regulatory mechanisms. Ischemic reperfusion injury has a multitude of negative consequences on normal cellular physiology, and detailed studies of these numerous potential TJ regulators will be needed to dissect those pathways that play important roles or can be remedied clinically. However, our preliminary understanding of ischemic injury to TJ permeability points to clear pathways that can be examined experimentally in the future.

25.4 SUMMARY Clear data from various experimental models indicate ischemia (through cellular ATP depletion) results in TJ disruption and its attendant cell and organ physiology consequences. Significant advances have been made in understanding of TJ regulatory pathways that are affected by ischemia. However, considerable challenges remain for investigators to integrate effects on TJ regulatory pathways within the context of acute renal failure pathophysiology. For example, ischemic reperfusion injury has dramatic consequences on renal vasculature, particularly in the outer medulla (Mason et al., 1984), where prolonged ischemic injury occurs during reperfusion. This results from reduced blood flow caused by constriction of venous flow (Mason et al., 1984), which is a consequence of edema and red cell trapping in the interstitium (Olof et al., 1991). Both edema and red cell extrusion into the interstitium could result from ischemia induced dysfunction of endothelial TJs. Finally, the effects of ischemia on TJ permeability that have been described for renal epithelial cells may be a more general effect that has pathophysiological consequences in other

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organ systems. Future studies of the endothelial barrier in stroke (Schulze et al., 1997; Bolton et al., 1998; Rubin and Staddon, 1999) and cholangiocyte dysfunction in liver ischemia (Doctor et al., 1999) could benefit from the studies performed in the kidney.

ACKNOWLEDGMENTS The authors acknowledge support from the National Institutes of Health/NIDDK (DK54518 and DK53465). Dr. Osun Kwon is thanked for her critical reading of the manuscript.

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26

Tight Junctions in Intestinal Inflammation Jörg-Dieter Schulzke and Michael Fromm

CONTENTS 26.1 Introduction .................................................................................................554 26.2 Tight Junction Changes in Ulcerative Colitis ............................................554 26.2.1 Epithelial Permeability in Ulcerative Colitis ................................554 26.2.2 Freeze-Fracture Electron Microscopy ...........................................555 26.2.3 Tight Junction Molecules in Ulcerative Colitis ............................555 26.2.4 Mechanisms of Tight Junction Alteration in Ulcerative Colitis.............................................................................................557 26.3 Tight Junction Changes in Crohn’s Disease ..............................................558 26.3.1 Intestinal Permeability in Crohn’s Disease ...................................558 26.3.2 Tight Junction Morphology in Crohn’s Disease ...........................558 26.3.3 Tight Junction Molecules in Crohn’s Disease ..............................559 26.4 Tight Junction Changes in Celiac Sprue ....................................................559 26.4.1 Intestinal Permeability ...................................................................560 26.4.2 Tight Junction Structure in Celiac Sprue......................................560 26.4.3 Tight Junction Modulators in Celiac Sprue ..................................561 26.4.4 Diarrhea in Celiac Sprue ...............................................................562 26.5 Tight Junction Changes in Blind Loop Syndrome ....................................562 26.6 Lack of Tight Junction Changes in Short Bowel Syndrome .....................563 26.7 Infectious Intestinal Diseases and Epithelial Barrier Function .................564 26.7.1 Infectious Diarrhea ........................................................................564 26.7.2 HIV Enteropathy............................................................................564 26.8 Tight Junction Downregulation by Proinflammatory Cytokines ...............565 26.8.1 TNFα in HT-29/B6 Cells ..............................................................565 26.8.2 TNFα in CaCo-2, T84, and HT-29cl.19A Cells ...........................566 26.8.3 Interferon-γ in T84 Cells ...............................................................567 28.8.4 Occludin Promoter Activity in Response to Proinflammatory Cytokines .......................................................................................568 26.9 Tight Junctions and Apoptosis....................................................................570 References..............................................................................................................571

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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26.1 INTRODUCTION Most types of intestinal inflammation are associated with altered barrier function. In the intestine, an impairment of the mucosal barrier has two main consequences. First, small solutes and water may flow into the lumen and can cause leak flux diarrhea. In general, all types of diarrhea are driven by osmotic forces; i.e., fluid follows an impaired solute transport. This includes (1) motility-driven diarrhea due to insufficient contact time; (2) malabsorptive diarrhea due to reduced absorptive area, to defective transporters, or to enzymatic maldigestion of luminal macromolecules; (3) secretory diarrhea due to an excessive active (an)ion secretion; (4) and finally leak flux diarrhea due to an impaired epithelial barrier and a passive back leak of ions and water into the intestinal lumen. The second consequence concerns larger molecules, which under normal conditions are almost perfectly prevented from being taken up. If the barrier is disturbed, uptake of even small amounts of antigens may aggravate or even initiate inflammation. Although these changes are mainly due to altered tight junction (TJ) properties, other structural changes such as epithelial cell apoptosis and erosion/ulcer type lesions also play a role. This chapter describes the functional role and the structure of the epithelial TJ in the inflamed or morphologically transformed intestine as well as regulatory mechanisms involved in these changes.

26.2 TIGHT JUNCTION CHANGES IN ULCERATIVE COLITIS Ulcerative colitis (UC) is an autoimmune disease of unknown origin with inflammation of the colonic mucosa followed by profuse diarrhea. Possible mechanisms are reduced absorption and a defect in epithelial barrier function. Evidence for impaired ion transport was obtained in in vivo perfusion studies (Rask-Madsen and Brix Jensen, 1973). Using an in vitro technique, Sandle and co-workers (1986) found no evidence for the activation of active anion secretion but instead for decreased net sodium absorption, which improved after application of steroids. This was corroborated in a subsequent paper in which electrogenic sodium absorptive capacity decreased (Sandle et al., 1990). In that study, sodium pump impairment was assumed, but recent studies seem also to indicate lower cellular expression of epithelial sodium channels (ENaC) subunits (Greig and Sandle, 2000).

26.2.1 EPITHELIAL PERMEABILITY

IN

ULCERATIVE COLITIS

Barrier function and TJ structure was recently studied in inflamed sigmoid colon from patients with UC undergoing colectomy (Schmitz et al., 1999a). Specimens were chosen that exhibited only mild to moderate histological disease activity (i.e., no erosions and ulcer-type lesions). Histologically, the epithelial cell lining was intact and the properties of this diseased tissue refer to a mucosa that endoscopically appears hyperemic, edematous, granular, and vulnerable. Instead of measuring total tissue resistance (Rt) with conventional Ussing technique, alternating current impedance analysis was performed to determine also the epithelial (Re) and subepithelial (Rsub) components of Rt. In vivo Rsub is no part of

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the barrier, because the majority of the subepithelial tissues is situated below the microvasculature. The discrimination between Re and Rsub is crucial if both figures change in opposing directions. Although total wall resistance (Rt) was reduced in UC by only 50%, AC impedance analysis uncovered a much more pronounced barrier defect (Table 26.1). Epithelial resistance (Re) decreased by 79%, which in conventional Ussing technique would have been masked by the increase of the subepithelial resistance (Rsub) due to inflammatory cell infiltration and submucosal edema. Thus, the inflamed colonic mucosa in ulcerative colitis is characterized by a very pronounced loss of barrier function.

26.2.2 FREEZE-FRACTURE ELECTRON MICROSCOPY In her classical study, Claude (1978) has shown for several epithelia that large differences in resistance are associated with only small differences in the number of horizontally oriented strands (strand count). This was proved true also in UCaltered tissues where the pronounced decrease in epithelial resistance was paralleled by a rather small reduction in epithelial cell TJ complexity (Figure 26.1A/B and Table 26.1). This decrease in strand count was accompanied by a reduced depth of the main tight junctional meshwork in colitis epithelia (Table 26.1). This reduction reached statistical significance only in surface TJs, possibly because of enhanced cell proliferation in the crypts with increased TJ assembly. As a further structural feature, aberrant strands appeared below the main tight junctional meshwork of surface and crypt TJs in UC, which have also been correlated to the increase in cell proliferation in UC, since similar TJ alterations have been described during cell proliferation in other tissues (Tice et al., 1979). From the distribution of the strand counts (Figure 26.1C) it can be seen that TJs of UC tissues often had only one or two stands, whereas control tissues had at least five strands, although strand number retained its Gaussian distribution in UC, suggesting an orderly interference with the assembly/disassembly process of TJ formation. The appearance of TJ sites with very low strand count may be functionally even more important than mean strand number during inflammation and — together with other phenomena influencing mucosal barrier function such as epithelial apoptotic rate — could explain Claude’s observation. Strand discontinuities were not more frequent in UC than in controls. On a first view, this may argue against a contribution of strand discontinuities to the barrier defect. In reality, strand discontinuities may become functionally important at sites with only one or two strands (see Figure 26.1C). At these sites, strand discontinuities may enable bacterial and food antigens to pass the epithelial barrier.

26.2.3 TIGHT JUNCTION MOLECULES

IN

ULCERATIVE COLITIS

Very recently, first data were presented on the TJ proteins occludin and claudin-1 in UC. Occludin and the TJ-associated proteins ZO-1 and ZO-2 were downregulated, whereas claudin-1 was unaltered (Ooi et al., 2000). In Northern blots, mRNA expression was reduced only for ZO-1 but not for occludin.

Subepithelium 14 ± 1 (10) 36 ± 3 (11)*** 54 ± 4 (7)##

Crypt 7.3 ± 0.3 (9) 5.5 ± 0.4 (9)** —

TJ Strand Count Surface 6.9 ± 0.3 (9) 4.8 ± 0.5 (9)*** —

Crypt 330 ± 37 (9) 308 ± 31 (9) —

TJ Mesh Depth (nm) Surface 307 ± 22 (9) 244 ± 19 (9)* —

Note: In sigmoid colon of controls and patients with UC, epithelial and subepithelial resistance were measured by AC impedance analysis and number of horizontally oriented strands in the main compact meshwork (TJ strand count) and depth of the main tight junctional meshwork (TJ mesh depth) of the TJ were evaluated by freeze-fracture electron microscopy. All values are means ± s.e.m.; number of patients is given in parentheses. * = P < 0.05 vs. control. ** = P < 0.01 vs. control. *** = P < 0.001 vs. control. ## = P < 0.01 vs. mild–moderate UC.

Control Mild/moderate UC Severe UC

Epithelium 95 ± 5 (10) 20 ± 3 (11)*** 22 ± 4 (7)

Electrical Resistance (Ω·cm2)

TABLE 26.1 Electrical Resistance and Tight Junction Morphometry in Ulcerative Colitis

556 Tight Junctions

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FIGURE 26.1 TJs in ulcerative colitis. Freeze-fracture electron micrographs from TJs of colonic enterocytes. (A) Control. (B) UC; mv = microvilli, tj = tight junction strands (bar represents 200 nm). (C) Distribution of the number of horizontally oriented strands along the TJ under control conditions and in UC. Values represent the percentage of grid lines with the respective strand count. (From Schmitz, H. et al., Gastroenterology, 116, 301, 1999a. With permission.)

26.2.4 MECHANISMS OF TIGHT JUNCTION ALTERATION IN ULCERATIVE COLITIS The inherent mechanisms of the TJ alteration observed in UC may involve the action of proinflammatory cytokines upregulated in inflammatory bowel disease (Andus et al., 1991) and effects of impaired barrier related to upregulated apoptosis (Straeter et al., 1997; Gitter et al., 2000b). This is described in more detail below.

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Tight Junctions

26.3 TIGHT JUNCTION CHANGES IN CROHN’S DISEASE Quite similar to UC, Crohn’s disease is an autoimmune disease of unknown origin with macroscopic inflammation of intestinal segments with preference of the ileum but often also involving colonic segments. Recent pathogenetic concepts prefer a switch from immune tolerance to autoimmunity characterized by a Th1 profile with elevated levels of proinflammatory cytokines such as interleukin-1β (IL-1β), interferon-γ, and tumor necrosis factor-α (TNFα). The epithelial barrier function is disturbed in Crohn’s disease. Several groups have favored the hypothesis of a “primary barrier defect,” which means that the barrier defect precedes the inflammation and thus contributes to the onset of the disease (Hollander et al., 1986). Much effort has been made using in vivo permeability tests to further elucidate this question (Katz et al., 1989; Ruttenberg et al., 1992; Teahon et al., 1992; Wyatt et al., 1993). However, only little information exists on the cellular mechanisms that could be responsible for this barrier defect.

26.3.1 INTESTINAL PERMEABILITY

IN

CROHN’S DISEASE

As already mentioned above, a large body of evidence exists from in vivo permeability studies, where urinary excretion of orally administered test substances such as polyethylene glycol (PEG), lactulose/mannitol, lactulose/rhamnose, and 51Cr-EDTA were measured. These studies have clearly demonstrated a significant barrier disturbance in Crohn’s disease, although other factors such as altered intestinal blood flow through the villus core may have partially influenced this result. More importantly, however, this type of measurement cannot resolve regional differences in permeability, which makes it difficult to distinguish primary and secondary barrier defects. Intestinal permeability of distinct intestinal segments can be obtained in vitro (Schulzke et al., 1995b). In biopsies obtained endoscopically from noninflamed distal duodenum of patients with Crohn’s disease, epithelial resistance (Re) as determined by impedance analysis in miniaturized Ussing chambers was 15 ± 2 Ω·cm2 in control and unchanged in Crohn’s disease. Concomitantly, 3H-lactulose permeability was characterized by unidirectional mucosal-to-serosal tracer flux measurements. In control, Jlactulose was 0.20 ± 0.03 µmol·h–1 ·cm–2 and was not significantly different in Crohn’s disease. In a further set of experiments, inflamed distal small intestine obtained from surgical resection was studied in conventional Ussing chambers (Table 26.2). In contrast to the noninflamed intestinal segments above, Re was decreased from 28 ± 4 Ω·cm2 in control to 15 ± 2 Ω·cm2 in ileitis Crohn (p < 0.05). Taken together, no evidence was obtained for a primary barrier defect in the noninflamed small intestine of patients with Crohn’s disease, while epithelial barrier function was clearly affected in inflamed small intestinal segments. Therefore, barrier disturbances detected in in vivo permeability studies are most likely caused by inflamed intestinal segments.

26.3.2 TIGHT JUNCTION MORPHOLOGY

IN

CROHN’S DISEASE

One freeze-fracture electron microscopy analysis has been performed on inflamed ileum specimens of patients with Crohn’s disease (Marin et al., 1983). Although no

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TABLE 26.2 Epithelial Barrier Function in Crohn’s Disease and HIV Enteropathy

Crohn’s disease Control ileum Ileitis Crohn p HIV-enteropathy Control duodenum HIV-infection P

Epithelial Resistance

Subepithelial Resistance

28 ± 5 Ω·cm2 15 ± 2 Ω·cm2 <0.05

24 ± 2 Ω·cm2 48 ± 4 Ω·cm2 <0.001

8 8

21 ± 2 Ω·cm2 13 ± 1 Ω·cm2 <0.01

29 ± 2 Ω·cm2 25 ± 2 Ω·cm2 n.s.

10 8

n

Note: Epithelial and subepithelial resistances were measured by AC impedance analysis in inflamed ileal specimens from patients with Crohn’s disease undergoing resection and in duodenal biopsies of HIV-infected patients with diarrhea (AIDS with CD4 cell counts below 100/µl). All values are means ± s.e.m. n.s. = not significant.

morphometry was performed, a semiqualitative evaluation has yielded convincing evidence for disturbed epithelial TJ structure in inflamed regions with a decrease in strand complexity. Taking into account that similar TJ alterations have been described in a morphometrical analysis of inflamed colon in UC, it seems reasonable to conclude that downregulation also exists in inflamed intestinal segments of patients with Crohn’s disease. Finally, it should be mentioned that a tight junctional change may not be the only source of a barrier disturbance, as nothing is known, for example, about the quantitative importance of apoptotic events or changes of the transcellular route (Schuermann et al., 1999).

26.3.3 TIGHT JUNCTION MOLECULES

IN

CROHN’S DISEASE

Similar to findings in UC, downregulation of occludin was obtained in immunoblots of biopsies also from patients with Crohn’s disease (Ooi et al., 2000). Although less pronounced than in involved tissues, these changes were observed in uninvolved colonic tissues as well. This may suggest a primary barrier defect in Crohn’s colitis but could also represent minimal inflammatory alterations in the neighborhood of inflammatorily affected colonic regions.

26.4 TIGHT JUNCTION CHANGES IN CELIAC SPRUE Celiac sprue is a genetically influenced (HLA-DQ2) (auto)immune disease resulting in a malabsorption syndrome. Sensitivity to gluten, a component of most cereals, causes formation of gliadin-transglutaminase complexes and increases expression

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Tight Junctions

TABLE 26.3 Electrical Resistance and Tight Junction Morphometry in Celiac Sprue

Control Acute sprue Sprue gluten-free

Epithelial Resistance (Ω·cm2) 20 ± 2 (9) 9 ± 1 (9)*** 15 ± 1 (9) *

TJ Strand Count Villus/Surface 5.3 ± 0.1 (112) 3.0 ± 0.1 (172)*** 5.1 ± 0.1 (77) ns

Upper Crypt 4.6 ± 0.1 (202) 4.0 ± 0.1 (182)*** 4.3 ± 0.1 (121) **

Lower Crypt 4.3 ± 0.1 (107) 3.7 ± 0.1 (196)*** 3.8 ± 0.1 (52) **

Note: Epithelial resistance was measured by AC impedance analysis and the number of horizontally oriented strands in the main compact meshwork of the TJ (TJ strand count) was evaluated by freeze-fracture electron microscopy in jejunal biopsies of controls, children with acute celiac sprue, and children under a glutenfree diet. All values are means ± s.e.m., n values are given in parentheses, and in electron microscopy refer to the number of grid lines analyzed; n.s. = not significantly different from control; * = P < 0.05, ** = P < 0.01, *** = P < 0.001 vs. control.

of proinflammatory cytokines including TNFα and interferon-γ. The small intestinal mucosa exhibits “hyperregenerative transformation” resulting in villus atrophy. In active disease, diarrhea is frequent. Diminished villus area with reduced transport capacity for Na+–glucose cotransport and other substrate cotransporters plays an important role. Epithelial barrier function is also disturbed, as indicated by both in vivo and in vitro permeability tests. This increased paracellular permeability not only contributes to diarrhea by a leak flux mechanism, but may also perpetuate active disease state by permitting macromolecules, including gluten, to penetrate through the epithelium into the intestinal mucosa.

26.4.1 INTESTINAL PERMEABILITY Intestinal transport and barrier function in celiac sprue were characterized using a miniaturized Ussing device for measurements on jejunal biopsies (Schulzke et al., 1995a). Epithelial resistance (Re) as determined by AC impedance analysis was reduced from 20 ± 2 Ω·cm2 in control jejunum to 9 ± 1 Ω·cm2 in acute sprue (Table 26.3). In gluten-free-nourished patients with celiac sprue, Re only partly recovered to 15 ± 1 Ω·cm2. Unidirectional Na+ and Cl– fluxes were increased in both directions in acute sprue as a consequence of the decrease in resistance. However, ISC as well as Na+ and Cl– net fluxes were not significantly different from control, which argues against activation of anion secretion in acute celiac sprue. Although not present under basic conditions, a Cl–-dependent increase in ISC was obtained after stimulation with theophylline and PGE1. Vmax of this electrogenic Cl– secretion was not significantly different between sprue and control jejunum.

26.4.2 TIGHT JUNCTION STRUCTURE

IN

CELIAC SPRUE

Structural analysis of jejunal epithelial TJs using freeze-fracture electron microscopy had been performed in adult patients with sprue (Madara and Trier, 1980). Changes

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FIGURE 26.2 TJs in celiac sprue. Freeze-fracture electron micrographs from TJs of jejunal enterocytes of the lower villus (surface). (A) Control; (B) acute celiac sprue. MV = microvillus. (From Schulzke, J. D. et al., Pediatr. Res., 43, 435, 1998. With permission.)

compatible with a loss of barrier function were detected with a modest 20% decrease in strand count from 5.0 to 4.0 at the surface and no change in the crypt. In children with acute celiac sprue (Schulzke et al., 1998), strand number was reduced in all regions along the surface–crypt axis, from 5.5 ± 0.2 to 3.4 ± 0.3 at the surface and from 4.7 ± 0.2 to 3.6 ± 0.1 in the crypt (Figure 26.2 and Table 26.3). Meshwork depth was also reduced in all regions along the surface–crypt axis, and aberrant strands appeared below the main meshwork of crypt TJs in acute sprue. Strand discontinuities were more frequent in acute sprue than in control (see Figure 26.2B). In asymptomatic children treated with gluten-free diet, jejunal tight junctional structure only partially recovered. Strand number was restored to normal at the surface, but was still decreased in the crypts, from 4.7 ± 0.2 to 3.9 ± 0.3. The 20% decrease in adults compared with the 40% decrease in children probably reflects the more active disease state in the children’s group with complete “flat mucosa” and diarrhea.

26.4.3 TIGHT JUNCTION MODULATORS

IN

CELIAC SPRUE

Factors that could be responsible for the change in TJ structure are elevated levels of TNFα and interferon-γ as well as the endogenous ligand of the zonula occludens toxin (ZOT) called zonulin (Fasano et al., 2000).

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Tight Junctions

26.4.4 DIARRHEA

IN

CELIAC SPRUE

Active Cl– secretion does not contribute to diarrhea in celiac sprue, although this transporter is present in the diseased tissue. Besides by the established malabsorptive mechanisms, sprue diarrhea is caused by impaired epithelial barrier function due to a decreased number of TJ horizontal strands and the appearance of strand discontinuities. In addition, other factors have been identified that contribute to the epithelial barrier function in acute celiac disease such as induction of epithelial apoptosis.

26.5 TIGHT JUNCTION CHANGES IN BLIND LOOP SYNDROME TJ changes occur together with alterations in ion transport in the surgically created self-filling blind loop of rat jejunum, a model for the blind loop syndrome in humans (Schulzke et al., 1990). Stasis of ingesta in the blind loop leads to bacterial overgrowth with bacterial bile acid deconjugation. The stasis per se, bacterial proteases, and free bile acids are responsible for mucosal transformation. In this state, the hyperregeneratively transformed intestinal mucosa with increased crypt length due to enhanced cell proliferation and increased villus height is characterized by decreased active ion transport and an increased transepithelial resistance (Schulzke et al., 1987). When epithelial cell TJ structure was analyzed morphometrically along the crypt–villus axis (Table 26.4), the number of strands and tight junctional depth, including meshwork depth, decreased from crypt to villus tip in control jejunum. In

TABLE 26.4 Tight Junction Morphometry in Blind Loop Syndrome Upper Villus

Lower Villus Upper Crypt Number of Horizontal Stands 5.7 ± 0.1 (108) 5.8 ± 0.1 (181) 5.7 ± 0.2 (37) n.s. 5.6 ± 0.1 (146) n.s.

Lower Crypt

Control Blind loop

4.8 ± 0.2 (56) 5.2 ± 0.1 (47)*

6.3 ± 0.2 (79) 5.9 ± 0.1 (300) n.s.

Control Blind loop

Tight Junctional Meshwork Depth (nm) 189 ± 5 (56) 234 ± 4 (108) 243 ± 6 (181) 217 ± 5 (146) ** 232 ± 7 (47)*** 236 ± 8 (37) n.s.

250 ± 8 (79) 223 ± 5 (300) **

Control Blind loop

Depth of Total Tight Junction (nm) 201 ± 7 (56) 245 ± 5 (108) 257 ± 6 (181) 248 ± 10 (47)*** 279 ± 12 (37) ** 286 ± 11 (146) *

278 ± 12 (79) 373 ± 11 (300) ***

Note: Number of horizontally oriented strands in the main compact meshwork, depth of the main tight junctional meshwork, and depth of the total TJ including aberrant strands below the main tight junctional meshwork were measured by freeze-fracture electron microscopy in the jejunum of control rats and in surgically created self-filling blind loops. All values are means ± s.e.m., n values are given in parenthesis as are the number of grid lines analyzed; n.s. = not significantly different from control; * = P < 0.05, ** = P < 0.01, *** = P < 0.001 vs. control.

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563

the blind loop, aberrant strands appeared below the meshwork, particularly in crypt cells. Consequently, total junctional depth was greater than in controls. Furthermore, strand number and junctional meshwork depth were increased in blind loops at the villus tip, which is that site along the crypt–villus axis that showed the shallowest junction in control jejunum. It appears that strands are inserted into the junctional meshwork at a higher rate in blind loops than in controls, leading to development of a more complex junction in villus tip cells. This structural change is paralleled by a threefold increased epithelial resistance measured by AC impedance analysis and by a pronounced decrease in short-circuit current in blind loop epithelia (Schulzke et al., 1987). Relative Na+ over Cl– permeability obtained from dilution potential measurements was 1.50:1 in control and not significantly different in blind loops (Schulzke et al., 1990). Thus, the resistance increase in blind loops cannot be attributed to a collapse of the lateral intercellular space, but is due to TJ permeability changes, since otherwise a significant shift to free solution values (0.67:1) would have been observed (Krasny et al., 1982). The TJ alterations in the blind loop syndrome represent a functionally meaningful feedback between tight junctional conductance and intrinsic transport properties in an epithelium. Since intrinsic transport rates are diminished in epithelial cells of the blind loop mucosa, the decrease in transjunctional permeability serves to reduce back-leakage of transported ions from the interspaces to the mucosal compartment to increase transport efficiency. The stimulus for this adaptation may have come from the relatively low pressure/volume status of the cellular interspace in the blind loop epithelium.

26.6 LACK OF TIGHT JUNCTION CHANGES IN SHORT BOWEL SYNDROME Adaptational changes of epithelial barrier and transport functions occur in the short bowel syndrome. This was studied experimentally in ileal remnants 8 weeks after 70% proximal small intestinal resection in the rat (Schulzke et al., 1992). Epithelial resistance measured by AC impedance analysis decreased from 27 ± 1 to 21 ± 1 Ω·cm2 and PEG 4000 fluxes increased from 2.5 ± 0.3 to 3.6 ± 0.3 nmol·h–1·cm–2. Since active transport parameters were unchanged, it was suggested that this was due to TJ alterations. A morphometrical analysis of TJ structure by freeze-fracture electron microscopy at four positions along the crypt–villus axis yielded no changes in the mean number of horizontal strands in the short bowel (Table 26.5). To explain a resistance decrease in spite of no change in strand number, the change in mucosal surface area should be considered. Resistance is usually related to the serosal area as defined by the opening of the Ussing chamber. However, epithelial resistance and transport depend on the mucosal surface area. Under the light microscope, mucosal surface area was increased as a result of increased villus height by 30% during adaptation in the short bowel. Thus, the decrease in epithelial resistance is due to mucosal hyperplasia, leading to an increase in tight junctional area per square centimeter of serosal area exposed in the Ussing chamber.

564

Tight Junctions

TABLE 26.5 Electrical Resistance and Tight Junctions in Short Bowel Syndrome Epithelial Tight Junctional Strand Count Resistance, Ω·cm2 Upper Villus Lower Villus Upper Crypt Lower Crypt Control ileum 27 ± 1 (16) 5.8 ± 0.1 (141) 5.9 ± 0.1 (100) 6.0 ± 0.2 (55) 6.6 ± 0.2 (50) Short bowel 21 ± 1 (19) n.s. 5.9 ± 0.2 (46) n.s. 6.3 ± 0.2 (75) n.s. 6.2 ± 0.1 (193) n.s. 6.9 ± 0.1 (157) n.s. Note: Epithelial resistance was measured by AC impedance analysis and the number of horizontally oriented strands in the main compact meshwork of the tight junction was evaluated by freeze-fracture electron microscopy in control ileum and in the ileal remnants after 70% proximal small intestinal resection. All values are means ± s.e.m., n values are given in parentheses and in electron microscopy refer to the number of grid lines analyzed; n.s. = not significantly different from control.

Finally, transport capacity of sodium–glucose cotransport in short bowel was found to be increased to 250% of control, which is partly due to increased villus and microvillus surface area but predominantly to increased expression of this cotransport system in the short bowel mucosa (Schulzke et al., 1992). This is an example where both transport and TJ structure/function adapted together to an increased workload.

26.7 INFECTIOUS INTESTINAL DISEASES AND EPITHELIAL BARRIER FUNCTION 26.7.1 INFECTIOUS DIARRHEA For a few bacterial infections of the intestine, exact information is available on TJ regulation as, for example, for pseudomembranous colitis (Clostridium difficile toxin A and B) and for Vibrio cholerae (ZOT = zonula occludens toxin), both of which are described in detail in other chapters of this volume. Other infectious intestinal diseases without specialized toxin production may share pathomechanisms of TJ downregulation with those described in the UC section of this chapter, namely, the regulatory influence of proinflammatory cytokines. Although not experimentally studied, this can be hypothesized by analogy to the mechanisms of cytokine-dependent TJ regulation (see below).

26.7.2 HIV ENTEROPATHY The gastrointestinal tract is a major target organ of the HIV infection, and diarrhea is a common symptom with major importance for mortality. Frequently, a cause for diarrhea, for example, enteric pathogens, can be found. However, in 15 to 40%, HIV itself has been postulated to be the relevant cause (HIV enteropathy). The pathomechanisms include a malabsorptive mechanism due to mucosal atrophy and an intestinal barrier defect.

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In HIV enteropathy, measurements with a miniaturized Ussing chamber on forceps biopsy specimens for detection of active ion secretion and barrier dysfunction were performed in the duodenum of three groups of patients, namely, patients with AIDS with diarrhea, asymptomatic HIV-positive patients, and controls (Stockmann et al., 1998). As a result, active ion secretion was found to be activated in an intermediate stage of HIV infection (CD4+ helper cell count 100 to 250/µl) and epithelial barrier function was found to be disturbed in intermediate as well as in late stages of HIV infection (CD4+ helper count below 100/µl) (Table 26.2). In the latter, active ion secretion was not any longer detectable. These data are direct evidence for a disturbed epithelial barrier in the duodenum of HIV-infected patients. It may be mediated by a release of TNFα and/or interferon-γ. Further evidence for such a role of cytokines in HIV enteropathy was obtained from experiments with supernatants of HIV-infected HT-29/B6 cells (Schmitz et al., 1998). Monocyte-derived macrophages were infected with HIV strains SF162 or HTLV-IIIb. After 7 days, the macrophages were cocultured with homologous peripheral blood leucocytes (PBL) for 24 h. Increased cytokine levels in the coculture supernatants were detected for TNFα, IL-1β, interferon-α, and interferon-γ. Unspecific cytotoxic effects of supernatants were excluded by LDH release assay. Supernatants of the SF162-infected cocultures reduced Rt of confluent HT-29/B6 monolayers to 53 ± 8% of control values after 24 h. Stimulation with supernatants of the HTLV-IIIb group resulted in an even more pronounced decrease in Rt to 20 ± 2% of control. This indicates that the amount and pattern of cytokines produced by HIVinfected human immune cells is sufficient to impair epithelial barrier function.

26.8 TIGHT JUNCTION DOWNREGULATION BY PROINFLAMMATORY CYTOKINES Many intestinal inflammatory diseases are associated with enhanced subepithelial cytokine release. In has been shown in several cell lines (Table 26.6) and cytokine knockout models that proinflammatory cytokines, especially TNFα and interferon-γ, impair the epithelial barrier.

26.8.1 TNF

IN

HT-29/B6 CELLS

TNFα is produced by mononuclear cells and TNFα expression is upregulated in intestinal diseases, including inflammatory bowel disease, celiac sprue, and HIV enteropathy. Serosal addition of TNFα (100 ng/ml) decreased Rt by 81% function in the colonic epithelial cell line HT-29/B6 (see Table 26.6). This effect was dose dependent, not reversible as long as TNFα was present, and could be mimicked by serosal addition of antibodies against the p55 TNFα receptor (Schmitz et al., 1998). No cytotoxic effects were observed in LDH assays. Immunofluorescence localization with anti-ZO-1 antibodies revealed no evidence for disruption of the monolayer after TNFα treatment. In freeze-fracture electron microscopy (Figure 26.3A/B), TJ complexity was decreased by TNFα as indicated by a decrease in the number of strands from 4.7 to 3.4 with appearance of TJ regions with only one or two strands (Figure 26.3C). The tyrosine kinase blocker genistein and the protein kinase A

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Tight Junctions

TABLE 26.6 Cytokine Effects on Epithelial Barrier and Tight Junctions Duration, Concentration h HT-29/B6 Cells (Schmitz et al., 1999b) Control (no cytokine) 24 IFNγ 1 U/ml 24 10 U/ml 24 100 U/ml 24 1000 U/ml 24 TNFα 5 ng/ml 24 100 ng/ml 24 TNFα + IFNγ 5 ng/ml + 10 U/ml 24 Cytokine

Resistance in % of Initial Value 98 98 86 83 79 69 18 23

± ± ± ± ± ± ± ±

5 1 5 7 5 9 1 4

n.s. n.s. n.s.

** ** ** ***

IL-1β IL-8 PGE2

HT-29/B6 Cells (Bode et al., 1998) 10 ng/ml 24 160 ng/ml 24 1 µM 24

TNFα IFNγ

T84 Cells (Madara and Stafford, 1989) 100 U/ml 72 100 U/ml 72

99 ± 2 n.s. 26 ± 6 ***

IFNγ

T84 Cells (Youakim and Ahdieh, 1999) 10 ng/ml 24

60

TNFα

CaCo-2 cells (Marano et al., 1998) 100 ng/ml 24

71

89 ± 3 102 ± 5 102 ± 2

n.s. n.s. n.s.

Note: n.s. = not significantly different from control, ** p < 0.01, *** p < 0.001.

inhibitor H-8 reduced the effect of TNFα. A combination of TNFα with interferonγ acted synergistically on the epithelial barrier.

26.8.2 TNF

IN

CACO-2, T84,

AND

HT-29CL.19A CELLS

Also in CaCo-2 BBE cells, TNFα is a modulator of epithelial permeability and changes in tight junction structure could be detected in freeze-fracture electron microscopy analysis (see Table 26.6; Marano et al., 1998). Amount and time course of the TNFα effects, however, differ in respect to cell line model and cytokine concentrations. In the T84 cell line model no effect of TNFα was observed (see Table 26.6; Madara and Stafford, 1989), which may be due to the low TNFα concentration applied or to the absence of TNFα receptors. In the intestinal epithelial cell line HT-29cl.19A as well, even the high concentration of 100 ng/ml of TNFα had only a very small effect on Rt after 48 h (Heyman et al., 1994). In a further study by this group, the combination of TNFα with a small dose

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FIGURE 26.3 Cytokine effects on TJs. Freeze-fracture electron micrographs from TJs of HT-29/B6 enterocytes. (A) Control; (B) 8 h after TNFα treatment (100 ng/ml). mv = microvilli, arrow = TJ strand meshwork. (C) Percentage of grid lines with the respective strand count along the TJ of HT-29/B6 cells under control conditions and after incubation with 100 ng/ml TNFα for 24 h. (From Schmitz, H. et al., J. Cell Sci., 112, 137, 1999b. With permission.)

of interferon-γ led to a significant reduction of Rt with a concomitant change in tight junctional complexity in freeze-fracture analysis (Rodriguez et al., 1995), which could be due to upregulation of TNF receptors, as argued by the authors. However, it should be mentioned that interferon-γ may have also other, for example, metabolic, effects in this cell line, which could enhance TNFα susceptibility. Aggarwal and coworkers have described such a phenomenon. Induction of TNF receptors by interferon-γ was not the major mechanism of synergism, since not only interferon-γ but also other interferons acted synergistically with TNFα but only interferon-γ induced TNF receptors (Aggarwal and Eessalu, 1987).

26.8.3 INTERFERON-

IN

T84 CELLS

In contrast to TNFα, interferon-γ modulates intestinal barrier in T84 cell monolayers after 72 h (see Table 26.6; Madara and Stafford, 1989), which was accompanied by a change in ZO-1 expression and by a perturbation of apical actin organization leading to a disorganization of the TJ and an increase in paracellular permeability

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Tight Junctions

FIGURE 26.3 (continued.)

(Youakim and Ahdieh, 1999). As in HT-29/B6 cells, there is also a synergistic effect of TNFα and interferon-γ in T84 cells (Fish et al., 1999). Interferon-γ effects are also observed in other intestinal cell lines such as HT29/B6. In long-term experiments, this includes additional cytotoxic effects as indicated by LDH release (Schmitz et al., 1999b).

26.8.4 OCCLUDIN PROMOTER ACTIVITY IN RESPONSE TO PROINFLAMMATORY CYTOKINES The 65-kDa protein occludin is a membrane-spanning part of the epithelial TJ. The function of occludin as part of TJs is still poorly understood, and even less is known about the regulatory mechanisms that influence occludin gene expression. By using genome walking cloning of occludin-specific human genomic DNA sequences, a 1853-bp DNA fragment containing the transcription start point of occludin cDNA sequences was amplified and sequenced (Mankertz et al., 2000). In close proximity to the transcription start point for occludin mRNA, a number of transcription factor-binding sites, e.g., for nuclear factor interleukin-6 (NF-IL6) were identified. Subcloning of this fragment in front of the luciferase reporter-gene revealed strong expression of enzymatic activity after transfection of the human intestinal cell line HT-29/B6. With subsequent deletions of parts of the promoter fragment, its size was reduced to 208 bp, which is necessary and sufficient to mediate promoter activity. The decrease in resistance in HT-29/B6 cells induced by TNFα and interferon-γ was preceded by a decrease in occludin mRNA expression (Figure 26.4A). TNFα and interferon-γ diminished occludin promoter activity alone and even synergistically, suggesting genomic regulation (Figure 26.4B). Transcription factor NF-IL6 was suggested to be involved in TNFα-mediated gene expression

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FIGURE 26.4 Effect of cytokines on occludin mRNA expression and promoter activity. (A) Effect of TNFα on occludin mRNA expression. RNA from HT-29/B6 cells incubated in the absence (–) or presence (+) of TNFα for 6 or 24 h was subjected to Northern hybridization with occludin (OCLN)- and glyceraldehyde-3-phosphate dehydrogenase (GAPDH)-specific DNA probes. (B) Effect of TNFα and interferon-γ on occludin promoter activity; 10 or 100 ng/ml TNFα and 10 or 100 U/ml interferon-γ were added for 21 h to HT-29/B6 cells transiently transfected with pOCLNproluc7. Occludin promoter-mediated luciferase expression was monitored 24 h after transfection in a chemoluminiscent assay (relative light units).

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(Yamamoto et al., 1995) as well as in the regulation of TNFα gene expression itself (Zagariya et al., 1998). The intracellular pathways that mediate TNFα-dependent transcription of genes for TJ proteins remain to be elucidated. However, altered gene expression of the TJ protein occludin may lead to alterations in TJ strand formation and subsequently may influence barrier function.

26.9 TIGHT JUNCTIONS AND APOPTOSIS TJ alterations are not the only structural defect in UC. By means of the conductance scanning technique (Köckerling et al., 1993; Gitter et al., 1997; 2000a) for resolving the spatial distribution of epithelial conductance the authors have observed two different types of barrier defects in UC (unpublished data). First, a discontinuously distributed type of high conductance (“hot spots of conductance”) and, secondly, a homogeneously distributed barrier impairment throughout the epithelium, which could indeed be due to TJ alterations. Possible candidates for hot spots of conductance are apoptotic foci. Apoptosis of epithelial cells has recently been shown to be upregulated in UC as a result of which small spots of the epithelial area may lose their integrity (Straeter et al., 1997). Although apoptosis is a demanding task for TJ plasticity, from results of morphological studies it has been concluded so far that epithelial permeability is not affected by apoptosis (Madara, 1990; Mayhew et al., 1999). Very recently, local conductance measurements of single apoptoses have become possible in HT-29/B6 cells using the conductance scanning technique (Gitter et al., 2000b). It has turned out that under control conditions spontaneous apoptoses already exhibit leaks of up to 280 nS (mean 48 ± 19 nS). The results disprove the dogma that isolated cell apoptosis occurs without affecting the epithelial cell permeability barrier. After induction by TNFα, the apoptotic leaks were dramatically enhanced (Figure 26.5). Its frequency increased threefold and the mean conductance of single apoptoses increased 12-fold (597 ± 98 nS). Thus, apoptosis accounted for about half (56%) of the TNFα-induced permeability increase, and the other half is caused by alterations of TJs in nonapoptotic areas. The mechanisms of increasing conductance in TNFα-induced apoptotic events needs to be characterized but may include common intracellular signal transduction pathways that can induce both apoptosis and downregulation of TJs.

FIGURE 26.4 (continued from page 569.) Values are normalized for protein content and expression of cotransfected Renilla luciferase reportergene. Vertical bars represent standard error of the mean (n = 3). (From Mankertz, J. et al, J. Cell Sci., 113: 2085–2090, 2000. With permission.)

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FIGURE 26.5 Conductivity of apoptoses and of nonapoptotic tissue areas. Contribution of apoptoses and nonapoptotic areas to total tissue conductivity was obtained by conductance scanning in HT-29/B6 monolayers. Conductivity is expressed per square centimeter gross tissue area. Under control conditions, apoptosis contributed 5.5% to total epithelial conductivity. With TNFα (100 ng/ml), basic epithelial conductivity of nonapoptotic areas increased 2.6-fold vs. control, whereas that of apoptoses increased by a factor of 37. Hence, with TNFα, apoptosis contributed 45% to the total epithelial conductivity. (From Gitter, A. et al., FASBE J., 14, 1753, 2000. With permission.)

REFERENCES Aggarwal, B. B. and Eessalu, T. E. 1987. Induction of receptors for tumor necrosis factor-α by interferons is not a major mechanism for their synergistic cytotoxic response. J. Biol. Chem., 262: 10000–10007. Andus, T., Gross, V., Casar, I., Krumm, D., Hosp, J., David, M., and Schölmerich, J. 1991. Activation of monocytes during inflammatory bowel disease. Pathobiology, 59: 166–170. Bode, H., Schmitz, H., Fromm, M., Riecken, E. O., and Schulzke, J. D. 1998. Negative cytokines. Z. Gastroenterol., Suppl. 1: 83–87. Claude, P. 1978. Morphological factors influencing transepithelial permeability: a model for the resistance of the zonula occludens. J. Membr. Biol., 39: 219–232. Fasano, A., Not, T., Wang, W., Berti, I., Tommasini, A., and Goldblum, S. E. 2000. Zonulin, a new discovered modulator of intestinal permeability, and its expression in coeliac disease. Lancet, 355: 1518–1519. Fish, S. M., Proujansky, R., and Reenstra, W. W. 1999. Synergistic effects of IFNγ and TNFα on T84 cell function. Gut, 45: 191–198. Gitter, A. H., Bertog, M., Schulzke, J. D., and Fromm, M. 1997. Measurement of paracellular epithelial conductivity by conductance scanning. Pflugers Arch., 434: 830–840. Gitter, A. H., Bendfeldt, K., Schulzke, J. D., and Fromm, M. 2000a. Trans-/paracellular, surface/crypt, and epithelial/subepithelial resistances of mammalian colonic epithelia. Pflugers Arch., 439: 477–482. Gitter, A. H., Bendfeldt, K., Schulzke, J. D., and Fromm, M. 2000b. Leaks in the epithelial barrier caused by spontaneous and TNFa-induced single-cell apoptosis. FASEB J., 14: 1749–1753. Greig, E. and Sandle, G. I. 2000. Diarrhea in ulcerative colitis: the role of altered colonic sodium transport. Ann. N.Y. Acad. Sci., 915: 327–332.

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Heyman, M., Darmon, N., Dupont, C., Dugas, B., Hirribaren, A., Blaton, M. A., and Desjeux, J. F. 1994. Mononuclear cells from infants allergic to cow’s milk secrete tumor necrosis factor α, altering intestinal function. Gastroenterology, 106: 1514–1523. Hollander, D., Vadheim, C. M., Brettholz, E., Petersen, G. M., Delahunty, T., and Rotter, J. I. 1986. Increased intestinal permeability in patients with Crohn’s disease and their relatives. A possible etiologic factor. Ann. Intern. Med., 105: 883–885. Katz, K. D., Hollander, D., Vadheim, C. M., McElree, C., Delahunty, T., Dadufalza, V. D., Krugliak, P., and Rotter, J. I. 1989. Intestinal permeability in patients with Crohn’s disease and their healthy relatives. Gastroenterology, 97: 927–931. Köckerling, A., Sorgenfrei, D., and Fromm, M. 1993. Electrogenic Na+ absorption of rat distal colon is confined to surface epithelium. A voltage scanning study. Am. J. Physiol., 264: C1285–1293. Krasny, E. J., DiBona, A. I., and Frizzell, R. A. 1982. Regulation of paracellular permselectivity in flounder intestine. Bull. Mt. Desert Isl. Biol. Lab., 22: 83–85. Madara, J. L. 1990. Maintenance of the macromolecular barrier at cell extrusion sites in intestinal epithelium: physiological rearrangement of tight junctions. J. Membr. Biol., 116: 177–184. Madara, J. L. and Stafford, J. 1989. Interferon-γ directly affects barrier function of cultured intestinal epithelial monolayers. J. Clin. Invest., 83: 724–727. Madara, J. L. and Trier, J. S. 1980. Structural abnormalities of jejunal epithelial cell membranes in celiac sprue. Lab. Invest., 43: 254–261. Mankertz, J., Tavalali, S., Schmitz, H., Mankertz, A., Riecken, E. O., Fromm, M., and Schulzke, J. D. 2000. Expression from the human occludin promoter is affected by tumor necrosis factor alpha and interferon gamma. J. Cell Sci., 113: 2085–2090. Marano, C. W., Lewis, S. A., Garulacan, L. A., Soler, A. P., and Mullin, J. M. 1998. Tumor necrosis factor-alpha increases sodium and chloride conductance across the tight junction of CaCo-2 BBE, a human intestinal epithelial cell line. J. Membr. Biol., 161: 263–274. Marin, M. L., Greenstein, A. J., Geller, S. A., Gordon, R. E., and Aufses, A. H., Jr. 1983. A freeze fracture study of Crohn’s disease of the terminal ileum: changes in epithelial tight junction organization. Am. J. Gastroenterol., 78: 537–547. Mayhew, T. M., Myklebust, R., Whybrow, A., and Jenkins, R. 1999. Epithelial integrity, cell death and cell loss in mammalian small intestine. Histol. Histopathol., 14: 257–267. Ooi, C. J., Rosenberg, I. M., Reinecker, H. C., and Podolsky, D. K. 2000. Regulation of tight junction proteins in human subjects with inflammatory bowel disease. Gastroenterology, 118: A795. Rask-Madsen, J. and Brix Jensen, P. 1973. Electrolyte transport capacity and electrical potentials of the normal and the inflamed human rectum in vivo. Scand. J. Gastroenterol., 8: 169–175. Rodriguez, P., Heyman, M., Candalh, C., Blaton, M. A., and Bouchaud, C. 1995. Tumour necrosis factor-α induces morphological and functional alterations of intestinal HT29 cl.19A cell monolayers. Cytokine, 7: 441–448. Ruttenberg, D., Young, G. O., Wright, J. P., and Isaacs, S. 1992. PEG-400 excretion in patients with Crohn’s disease, their first degree relatives, and healthy volunteers. Dig. Dis. Sci., 37: 705–708. Sandle, G. I., Hayslett, J. P., and Binder, H. J. 1986. Effect of glucocorticoids on rectal transport in normal subjects and patients with ulcerative colitis. Gut, 27: 309–316. Sandle, G. I., Higgs, N., Crowe, P., Marsh, M. N., Venkatesan, S., and Peters, T. J. 1990. Cellular basis for defective electrolyte transport in inflamed human colon. Gastroenterology, 99: 97–105.

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Schmitz, H., Rokos, K., Fromm, M., Scholz, P., Bode, H., Riecken, E. O., Pauli, G., and Schulzke, J. D. 1998. Zellkulturüberstände von HIV-infizierten Immunzellen beeinträchtigen die Barrierefunktion von Colonepithelzellen (HT-29/B6). Z. Gastroenterol., Suppl. 1: 6–9. Schmitz, H., Barmeyer, C., Fromm, M., Runkel, N., Foss, H. D., Bentzel, C. J., Riecken, E. O., and Schulzke, J. D. 1999a. A decrease in tight junction complexity contributes to the severely impaired epithelial barrier function in ulcerative colitis. Gastroenterology, 116: 301–309. Schmitz, H., Fromm, M., Bentzel, C. J., Scholz, P., Bode, H., Epple, H. J., Riecken, E. O., and Schulzke, J. D. 1999b. Tumor necrosis factor-alpha (TNFα) regulates the epithelial barrier in the human intestinal cell line HT-29/B6. J. Cell Sci., 112: 137–146. Schuermann, G., Bruewer, M., Klotz, A., Schmidt, K. W., Senninger, N., and Zimmer, K. P. 1999. Transepithelial transport processes at the intestinal mucosa in inflammatory bowel disease. Int. J. Colorect. Dis., 14: 41–46. Schulzke, J. D., Fromm, M., Menge, H., and Riecken, E. O. 1987. Impaired intestinal sodium and chloride transport in the blind loop syndrome of the rat. Gastroenterology, 92: 693–698. Schulzke, J. D., Fromm, M., Zeitz, M., Menge, H., Riecken, E. O., and Bentzel, C. 1990. Regulation of the tight junction during impaired ion transport in the experimental blind loop syndrome of the rat. Res. Exp. Med., 190: 59–68. Schulzke, J. D., Fromm, M., Bentzel, C. J., Zeitz, M., Menge, H., and Riecken, E. O. 1992. Epithelial ion transport in the experimental short bowel syndrome of the rat. Gastroenterology, 102: 497–504. Schulzke, J. D., Schulzke, I., Fromm, M., and Riecken, E. O. 1995a. Epithelial barrier and ion transport in coeliac sprue: electrical measurements on intestinal aspiration biopsy specimens. Gut, 37: 777–782. Schulzke, J. D., Fromm, M., Bredenfeld, H., Herzog, A., and Riecken, E. O. 1995b. Direct permeability measurements on small intestinal biopsies in vitro point against a primary barrier defect in Crohn’s disease. Gastroenterology, 108: A914. Schulzke, J. D., Bentzel, C. J., Schulzke, I., Riecken, E. O., and Fromm, M. 1998. Epithelial tight junction structure in the jejunum of children with acute and treated celiac sprue. Pediatr. Res., 43: 435–441. Stockmann, M., Fromm, M., Schmitz, H., Schmidt, W., Riecken, E. O., and Schulzke, J. D. 1998. Duodenal biopsies of HIV infected patients with diarrhea show epithelial barrier defects but no secretion. AIDS, 12: 43–51. Straeter, J., Wellisch, I., Riedl, S., Walczak, H., Koretz, K., Tandara, A., Krammer, P. H., and Möller, P. 1997. CD95 (APO-1/Fas)-mediated apoptosis in colon epithelial cells: a possible role in ulcerative colitis. Gastroenterology, 113: 160–167. Teahon, K., Smethurst, P., Levi, A. J., Menzies, I. S., and Bjarnason, I. 1992. Intestinal permeability in patients with Crohn’s disease and their first degree relatives. Gut, 33: 320–323. Tice, L. W., Carter, R. L., and Cahill, M. B. 1979. Changes in tight junctions of rat intestinal crypt cells associated with changes in their mitotic activity. Tissue Cell, 11: 293–316. Wyatt, J., Vogelsang, H., Hubl, W., Waldhoer, T., and Lochs, H. 1993. Intestinal permeability and the prediction of relapse in Crohn’s disease. Lancet, 341: 1437–1439. Yamamoto, K., Arakawa, T., Ueda, N., and Yamamoto, S. 1995. Transcriptional roles of nuclear factor kappa B and nuclear factor-interleukin-6 in the tumor necrosis factor alpha-dependent induction of cyclooxygenase-2 in MC3T3-E1 cells. J. Biol. Chem., 270: 31315–31320.

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Youakim, A. and Ahdieh, M. 1999. Interferon-gamma decreases barrier function in T84 cells by reducing ZO-1 levels and disrupting apical actin. Am. J. Physiol., 276: G1279–1288. Zagariya, A., Mungre, S., Lovis, R., Birrer, M., Ness, S., Thimmapaya, B., and Pope, R. 1998. Tumor necrosis factor alpha gene regulation: enhancement of C/EBPbeta-induced activation by c-Jun. Mol. Cell Biol., 18: 2815–2824.

27

Tight Junctions in Liver Disease Lukas Landmann and Bruno Stieger

CONTENTS 27.1 The Role of Tight Juntions in Bile Secretion ............................................576 27.2 Hepatocellular Tight Junctions ...................................................................578 27.2.1 Morphology....................................................................................578 27.2.2 Probing of Tight Junctions in the Liver ........................................578 27.3 Tight Junctions in Liver Disease ................................................................580 27.3.1 Cholestasis .....................................................................................580 27.3.1.1 Extrahepatic Obstructive Cholestasis............................580 27.3.1.1.1 Structural alterations..................................580 27.3.1.1.2 Functional alterations ................................582 27.3.1.1.3 Effects on cell polarity ..............................585 27.3.1.1.4 Reversibility ...............................................585 27.3.1.1.5 Conclusion .................................................585 27.3.1.2 Ethinylestradiol..............................................................586 27.3.1.2.1 Structural alterations..................................586 27.3.1.2.2 Functional alterations ................................587 27.3.1.2.3 Effects on cell polarity ..............................587 27.3.1.2.4 Conclusion .................................................587 27.3.1.3 Bile Acid-Induced Cholestasis: Taurolithocholate........587 27.3.1.4 Drug- and Toxin-Induced Cholestasis...........................587 27.3.1.4.1 Cyclosporine A ..........................................587 27.3.1.4.2 ANIT ..........................................................588 27.3.1.4.3 Carmustin...................................................588 27.3.1.4.4 BSP ............................................................588 27.3.1.4.5 Phalloidin ...................................................588 27.3.1.4.6 Colchicine ..................................................589 27.3.2 Oxidative Stress .............................................................................589 27.3.3 Inflammatory Diseases ..................................................................589 27.3.4 Regeneration and Cell Proliferation..............................................589 27.4 Concluding Remarks...................................................................................590 References..............................................................................................................591

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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27.1 THE ROLE OF TIGHT JUNTIONS IN BILE SECRETION Three times the length of the Mississippi River, or 20,000 km, is the extent of hepatocellular tight junctions (TJs) within a human liver if one extrapolates data obtained from rats (Weibel, 1979; Stammler et al., 1990; Hornstein et al., 1992; Rahner et al., 1996). This seal is formed by 200 billion hepatocytes and confines as little as 3.5 cm3 of biliary space organized in an anastomosing network of tiny, 1-µm canaliculi. The secretory surface of the canalicular wall that is formed by the apical membrane domain of hepatocytes totals 115 m2, approximately the size of a fiveroom apartment. It is obvious that stowing these dimensions in a 1500-g organ necessitates an elaborate organization (Figure 27.1A). Consequently, hepatocytes are organized in one-cell-thick, irregular, and interconnected plates that radiate from the central vein and are intertwined with a threedimensional anastomosing network of capillaries, the sinusoids. These plates are lined by the sinusoidal endothelium, the cells of which are perforated by fenestrae. Therefore, noncellular blood components are in continuous exchange with the space of Disse localized between endothelial and liver cells and in direct contact with the basolateral plasma membrane of hepatocytes. Within the plates, the polygonal hepatocytes interact with each other by their lateral plasma membranes while the apical domains of adjacent cells delineate a chicken-wire-like network of bile canaliculi. Thus, the apical domain surrounds the hepatocyte periphery like a belt, whereas in simple surface epithelia it is confined to a single apex. In analogy to simple epithelia the apical domain is separated from the basolateral plasma membrane by TJs that form the only singular barrier between portal blood and biliary compartments. Bile formation is a major function of the liver. Bile acids, the main constituents of bile, are concentrated by a factor up to 1000 in bile as compared with portal blood, a process that depends on active transport. Since bile formation is an isoosmotic process, bile acids, actively transported across hepatocytes, are followed by trans- and paracellular water flow (Erlinger, 1994). Bile acids are taken up at the basolateral plasma membrane of hepatocytes, which is in direct contact with the blood plasma by Na-dependent and Na-independent transport proteins (KullakUblick et al., 2000). After transcellular movement (Agellon and Torchia, 2000), they are secreted across the apical or canalicular membrane by an ATP-dependent transporter (Kullak-Ublick et al., 2000). Other cholephilic compounds are translocated similarly by transport systems that in their majority are different from those used for bile acid transport (Kullak-Ublick et al., 2000). This vectorial transport is reflected by an asymmetric or polar expression of the involved transport systems, as well as of other plasma membrane proteins in the hepatocellular membrane domains. Secreted cholephilic compounds, because of their concentration in the canaliculus, exert an electrical transepithelial potential and osmotic gradient that are responsible for generation of bile secretion (Boyer et al., 1992). TJs partcipate in this process by allowing selective movement of water and small solutes (e.g., cations) and by preventing regurgitation of bile constituents through the paracellular pathway. Therefore, the barrier function of TJs is a dual one (Powell, 1981): they maintain a steep electro-osmotic gradient between two compartments (“gate” function) as well

B

FIGURE 27.1 (A) Hepatocytes (H) in a hepatic plate (HP). The plates are lined on both sides by liver sinusoids (LS) with endothelial cells (End) containing sieve plates (SP) and holes (Ho). Thus, cellular (E, erythrocyte) but not serum components of blood are prevented from entering the space of Disse (DS), which extends between endothelial and liver cells. Hepatocytes (H) face the liver sinusoids (LS) with microvilli (Mv) on their sinusoidal domain, while the smooth lateral membrane is in contact with an adjacent cell. The apical domain forms a longitudinal groove that together with a complementary groove of the adjacent cell closes to a bile canaliculus (BC). Canaliculi are sealed by TJs (ZO), which in turn are reinforced by zonulae adherentes (ZA). (From Krstic, R. V., Human Microscopic Anatomy, Springer-Verlag, Berlin, 1991. With permission.) (B) Freeze-fracture replica of normal rat liver showing a bile canaliculus (BC) with microvilli (Mv). The canaliculus is delineated on both sides by the TJs, which form a network of interconnected strands.

A

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as the functional and biochemical integrity of two membrane domains resulting in cell surface polarity (“fence” function). Both functions are crucial for bile formation, and their impairment is associated with elevated levels of bile constituents in blood.

27.2 HEPATOCELLULAR TIGHT JUNCTIONS 27.2.1 MORPHOLOGY In thin-section electron microscopy, TJs obliterate the intercellular space by the apparent fusion of the membranes of two adjacent cells. At high resolution, this region of membrane contact is discontinuous, displaying a varying number of close contact points. Freeze-fracture replication techniques of aldehyde-fixed material show a network of anastomosing strands and complementary grooves forming a continuous belt at the edges of the canaliculi (Figures 27.1B and 27.3A). These strands are reflected as contact points in thin sections. Junctional depth measured perpendicularly to the canalicular axis is approximately 0.2 µm and displays three to five strands, a structural organization considered to reflect a quite leaky type of TJs (Anderson, 1993). The molecular equivalents of these structures are dealt with in detail elsewhere in this book. Although it has become clear in recent years that claudins and occludin are components of TJ strands, very little is known to date about the properties of these molecules establishing and modulating barrier function. Occludin and claudins have two extracellular loops thought to interdigitate in a zipperlike way to create the paracellular seal (Tsukita et al., 1999; Tsukita and Furuse, 2000). In addition, plaque proteins such as ZO-1 mediate the mutual interaction of junctional integral membrane proteins and the actin cytoskeleton (Mitic and Anderson, 1998), which plays an important role in maintenance and regulation of TJ structure and function (see Chapter 12). Which of these proteins participate in separation of membrane domains is unknown to date. Therefore, characterization at the molecular level of TJ gate and fence function is not yet possible.

27.2.2 PROBING

OF

TIGHT JUNCTIONS

IN THE

LIVER

The characteristics of the TJ barrier conventionally are defined by its size and charge selectivity, both measured as transepithelial electrical resistance (Powell, 1981) (see Chapter 4). In liver, unfortunately, TJs in situ are inaccessible to direct electrophysiological examination since bile canaliculi are only about 1 µm in diameter. Therefore, understanding of their physiological properties is largely indirect. At the organ level (in situ or in the isolated perfused liver) studies are performed by applying markers to the vascular system and by sampling bile from the common bile duct. Metabolically inert, lipid-insoluble compounds of different size and charge that in hepatocytes are neither transported by a carrier nor bound to a receptor are used as probes for canalicular permeability (Erlinger, 1994). Monitoring of their biliary clearance by measuring their blood-to-bile ratio led to the conclusion that the paracellular pathway is the major route of entry into the canalicular bile compartment and that permeability is controlled by a barrier restricting selectively movement of

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FIGURE 27.2 Appearance of HRP in bile from an isolated rat liver perfused with nonrecirculating Krebs–Ringer bicarbonate buffer. HRP was injected into the perfusate at zero time and bile was collected every 12 s. Absorbance at 450 nm is a measure of HRP concentration in bile. (A) Data from a control experiment with an early peak at 2.5 min and a late peak at 12 min. (B) Data from an experiment in which 10–8 M vasopressin was infused 1 min before and 5 min after injection of HRP. This treatment greatly increases the early peak, an effect reflecting impairment of the functional integrity of TJs. (From Lowe, P. J. et al., Am. J. Physiol., 255, G454, 1988. With permission.)

anions and large molecules such as higher-Mr serum proteins (Forker, 1970; Bradley and Herz, 1978). Bolus injection or continuous infusion of horseradish peroxidase (HRP) results in a biphasic appearance in bile with an early peak after 2 to 5 min and a late peak at 15 to 30 min after injection (Lowe et al., 1985; Stieger et al., 1994; Rahner et al., 1996) (Figure 27.2A). The evidence linking these peaks with paracellular and transcellular routes includes coappearance of HRP with [3H]inulin and [14C]sucrose at the early peak and with pIgA at the late peak (Lowe et al., 1985; Coleman, 1987); both compounds are classical markers for the paracellular and transcytotic route, respectively. Detailed examination of the early peak using anionic and cationic HRP iosoenzymes corroberated the charge selectivity of the paracellular pathway (Hardison et al., 1989), whereas its size discrimination was confirmed by administration of dextrans of various molecular weights (Rahner et al., 1996). Since the TJ is the only physical barrier in the paracellular pathway, it is considered to control the characteristics of the early peak. The isolated rat hepatocyte couplets (IRHC) model (Boyer et al., 1988; Gautam et al., 1989) consists of adhering pairs of hepatocytes that are prepared by modified hepatocyte isolation procedures (Gautam et al., 1987; Wilton et al., 1991). This model eliminates potentially distorting contributions of the vascular system and biliary epithelium. IRHCs in primary culture reorganize their canalicular membrane domain including TJs and form a closed canalicular space (Graf et al., 1984; Roma et al., 1997) that increases with ongoing bile secretion (Gautam et al., 1989). In contrast to the intact organ, this space has been punctured with micropipettes (Graf et al., 1987), and TJ resistance was measured as 25 MΩ, corresponding to a conductance of

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37 µS/cm. This value is close to estimates based on structural parameters of TJs (Rahner et al., 1996), an approach that models the TJ as an electrical circuit consisting of individual resistors arranged in parallel (Claude, 1978; Marcial et al., 1984; Madara and Dharmsathaphorn, 1985) (see Chapter 4). The calculated specific TJ resistance of 48 kΩ·cm (Rahner et al., 1996) ranges with renal proximal tubule or gallbladder (Claude, 1978; Powell, 1981) among the leaky junctions and, if computed per singular hepatocyte (or couplet, 4 MΩ), is less than one order of magnitude off the 25 MΩ determined electrophysiologically in IRHC. All these indirect approaches demonstrated that permeability of TJs is modulated by a variety of factors (Nathanson and Boyer, 1991) (see Chapters 15 through 17), including protein kinase C (Corasanti et al., 1989), intracellular Ca2+ (Nathanson and Burgstahler, 1992), and hormones modulating its intracellular concentration such as vasopressin (Lowe et al., 1988) (Figure 27.2B).

27.3 TIGHT JUNCTIONS IN LIVER DISEASE 27.3.1 CHOLESTASIS Cholestasis is a pathophysiological condition originally defined by the presence of “bile plugs” in hepatocytes, canaliculi, and intrahepatic bile ducts (Popper and Szanto, 1956). This pathological definition was extended by a physiological one requiring decreased bile flow as sole criterion (Javitt and Arias, 1967). The many forms of cholestasis involve a variety of biochemical, physiological, structural, and clinical alterations (Desmet, 1992; Reichen and Simon, 1994). However, some features, including elevated serum levels of cholephilic compounds (e.g., bilirubin and bile acids) or activities of liver-associated enzymes (alkaline phosphatase, γ-glutamyltranspeptidase) as well as increased paracellular permeability associated with impaired TJ integrity, are common to all. Defective barrier function results in dissipation of electrochemical gradients between bile and blood, leads to regurgitation of bile constituents, and causes or contributes to cholestasis (Boyer, 1983; Sellinger and Boyer, 1990; Anderson, 1993). 27.3.1.1 Extrahepatic Obstructive Cholestasis 27.3.1.1.1 Structural alterations Obstruction of the common bile duct is the most obvious form of cholestasis. Pathological conditions leading mechanically to cessation of bile flow include gallstones (cholelithiasis) as well as cancer of the bile duct or pancreas head. Bile duct ligation (BDL) in the rat, an animal model for extrahepatic obstructive cholestasis, induces drastic changes in the TJs (Metz et al., 1977; De Vos and Desmet, 1978) resulting in disorganization of the network (Easter et al., 1983) with reduction of the number of strands (Rahner et al., 1996) and increased frequency of blind-ending abluminal strands (Metz et al., 1977; De Vos and Desmet, 1978) (Figure 27.3B). Similar alterations were reported for human liver cells from patients with extrahepatic cholestasis (Robenek et al., 1980b). Hepatocytes handicapped in their secretory function seem to adapt by an attempt to form new canaliculi: canalicular length is

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FIGURE 27.3 Freeze-fracture replicas of hepatocellular TJs. The canalicular membrane characterized by broken microvilli is on top of the micrographs. (A) Controls; (B) EE; (C) BDL. The regular organization of junctional strands is disturbed by cholestasis. Bar = 0.2 µm. (From Rahner, C. et al., Gastroenterology, 110, 1564, 1996. With permission.)

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almost twice the normal value after 48 h of BDL as demonstrated by stereological techniques (Rahner et al., 1996) and the number of blind-ending canaliculi that branch in right angles from the main canalicular axis and are lined by TJs displaying an extreme parallel organization of strands is increased (De Vos and Desmet, 1978). In freeze-fracture replicas, strand elements or particles are predominantly associated with the P face in aldehyde-fixed and with the E face in high-pressure-frozen cells, respectively (Staehelin, 1974; Van Deurs and Luft, 1979) (Figure 27.4). Recently, in replicas of transfected mouse fibroblasts the “tight” claudin-1 particles were shown to associate with the P face and the “leaky” claudin-2 with the E face (Furuse et al., 1998; Tsukita and Furuse, 2000). In contrast to other systems (see Chapter 2), the percentage of strand elements that is found on the opposite fracture face is not altered in aldehyde-fixed (Easter et al., 1983) or in high-pressure-frozen (Rahner et al., 1996) hepatocytes (Figure 27.4C). This makes unlikely a BDLinduced alteration of the molecular strand composition, which has been suggested to alter TJ permeability in other epithelia (Anderson and Van Itallie, 1995; Tsukita and Furuse, 2000). In normal liver, the cytoplasmic plaque protein ZO-1 that presumably is involved in signal transduction and organization of the actin cytoskeleton (see Chapter 12) outlines the canaliculi along their margins if visualized by immunocytochemistry (Anderson et al., 1989). The continuous lines become discontinuous after 48 h of BDL (Anderson et al., 1989), and immunoreactivity accumulates in the pericanalicular region, an alteration associated with increased protein and mRNA levels after 1 to 2 weeks (Fallon et al., 1993). In contrast, protein levels of the transmembrane strand component occludin decrease by 50% within 2 days and return to control values by 9 days (Fallon et al., 1995). After 2 days of BDL, the distribution of occludin parallels that of ZO-1 when both antigens become discontinuous but still colocalize. Prolonged cholestasis, however, results in dissociation with occludin remaining at the canaliculus and ZO-1 showing up in pericanalicular accumulations (Fallon et al., 1995). Examination of the cytoplasmic TJ proteins 7H6 and ZO-1, which in normal liver were in perfect register, showed that colocalization in hepatocytes but not bile duct cells became similarly discontinuous after BDL for 2 days (Kawaguchi et al., 1999). In contrast to ZO-1, protein expression levels of 7H6 decreased. In addition, in the periportal zone dissociation of signals due to partial relocation of 7H6 in discrete pericanalicular spots and downregulation of this protein were noted. This lobular gradient with more-pronounced 7H6 defects in the periportal zone is explained with the higher sensitivity of this protein to the effect of toxic bile salts to which periportal hepatocytes are more exposed. BDL-induced dissociation of integral and peripheral proteins indicates that functional integrity of the TJ depends on proper interaction between junctional components. 27.3.1.1.2 Functional alterations That impairment of structural integrity correlates with increased permeability was demonstrated by a BDL-induced 20-fold increase of HRP-positive TJs (Metz et al., 1977; Rahner et al., 1996) (Figure 27.5). This is in line with estimates of specific junctional resistance based on structural criteria (Rahner et al., 1996), which indicate a decrease by more than 80% from 48 to 8 kΩ·cm. The morphological data are paralleled by the secretion of HRP during the early peak, which was measured

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583

FIGURE 27.4 High-pressure-frozen replicas of native (chemically unfixed) TJs. The canalicular membrane is on the bottom of the micrographs. (A) Controls; (B) EE; (C) BDL. Membrane-associated particles are equivalent to strands in aldehyde-fixed specimens but are associated with the extracellular fracture face. Cholestasis does not affect the arrangement of particles within rows (= strands). Bar = 0.2 µm. (From Rahner, C. et al., Gastroenterology, 110, 1564, 1996. With permission.)

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FIGURE 27.5 TJ permeability for HRP. HRP injected into the femoral vein was used as tracer for paracellular permeability. TJs (arrows) in (A) control liver are predominantly negative, whereas cholestasis (B = BDL) greatly increases the proportion of HRP-positive junctions. Bar = 0.5 µm. (From Rahner, C. et al., Gastroenterology, 110, 1564, 1996. With permission.)

immediately after release of ligation and displayed a dramatic 6.5-fold increase (Stieger et al., 1994; Rahner et al., 1996) (Figure 27.6B). Size selectivity of TJs is also altered by BDL as shown by dextrans of various molecular weights. While dextrans up to 70 kDa are detectable in the early peak of control animals, BDL results in biliary secretion of molecules greater than 250 kDa (Rahner et al., 1996). However, the absence of the 318-kDa protein pIgA in bile after BDL (Stieger et al., 1994) indicates that the barrier, although severely impaired, is not broken down altogether but keeps functioning as a gate to some degree. Slightly defective size discrimination of TJs had already been observed in short-term biliary obstruction up to 20 min, a model that, in addition, allowed the demonstration of impaired charge selectivity by an increased output in bile of inert negatively charged solutes as compared with size-matched uncharged molecules (Cotting et al., 1989).

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FIGURE 27.6 Secretion of HRP into bile in rats with cholestasis induced by (A) EE and (B) BDL. HRP (2 mg/100 g body weight) was injected intravenously and bile was sampled at the indicated intervals. As compared with controls () cholestasis (●) increases HRP secretion during the early peak, an effect that is more pronounced after BDL than after EE treatment. (From Rahner, C. et al., Gastroenterology, 110, 1564, 1996. With permission.)

27.3.1.1.3 Effects on cell polarity BDL impairs not only the gate but also the fence function of TJs, as shown by impairment of the highly asymmetric expression of numerous domain-specific plasma membrane proteins (Durand-Schneider et al., 1987; Fricker et al., 1989; Landmann et al., 1990; Oguey et al., 1992; Stieger et al., 1994). Many but not all domain-specific antigens were shown to redistribute from the canalicular into the basolateral domain, or vice versa, most likely by lateral diffusion through damaged TJs (Stieger et al., 1994) (Figure 27.7C). Among the specific transport systems expressed in the basolateral and apical membrane domain, respectively, none was found to be translocated by BDL (Gartung et al., 1996; Landmann et al., 1998; Trauner et al., 1998). The reasons for this differential behavior are unclear but presumably include specific interactions with cytoskeletal components, which have been demonstrated paradigmatically for Na+,K+-ATPase (see Chapter 8). 27.3.1.1.4 Reversibility BDL-induced structural defects are completely reversible within 96 h after recanalization (Metz and Bressler, 1979). However, restoration of functional integrity is detected much earlier, as demonstrated by the strongly decreased early peak if ligation is released 1 h prior to injection of HRP (Stieger et al., 1994). This time interval, however, was not sufficient to reverse defective hepatocellular surface polarity appreciably (Stieger et al., 1994). 27.3.1.1.5 Conclusion BDL-induced structural, functional, and biochemical alterations are severe and impair the barrier drastically but are reversible. Although BDL reflects a relevant human pathology, it is a very complex experimental model. It is not clear at present whether injury results from increased hydrostatic pressure in a closed canalicular

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FIGURE 27.7 Effects of cholestasis on the surface polarity of hepatocytes as demonstrated by indirect immunofluorescence of aminopeptidase N, a membrane protein with canalicular domain specificity. In control liver (A) the antigen exhibits an exclusively canalicular expression, a distribution that is not altered after EE treatment (B), while after BDL (C) antigen is relocated into the basolateral domain. Arrowheads point to canaliculi. (A) shows a portal field containing a portal vein (PV) and an intrahepatic bile duct (BD). (Modified from Stieger, B. et al., Hepatology, 20, 201, 1994. With permission.)

space (Toyota et al., 1984); excessive concentration of toxic bile components including the toxic bile salt lithocholate; intrahepatic inflammation, which is a collateral consequence of BDL; or other metabolic changes. In addition, it is not known whether these parameters exert their action on the TJ directly — such as hydrostatic pressure that may force the TJ to open — or indirectly with junctional impairment as a secondary epiphenomenon. 27.3.1.2 Ethinylestradiol 27.3.1.2.1 Structural alterations Administration of ethinylestradiol (EE) is an animal model mimicking the effects of estradiol or its metabolites in cholestasis of pregnancy or that occurring occasionally after the use of oral contraceptives. This treatment induces similar (De Vos and Desmet, 1981; Robenek et al., 1982) but less profound (Rahner et al., 1996) alterations than BDL (Figures 27.3B and 27.4B). Junctional length per cubic centimeter of liver is not changed but, due to hepatocellular hypertrophy and proliferation (Hornstein et al., 1992), increases if related to the whole organ. EE does not alter the subcellular distribution of ZO-1 (Anderson et al., 1989) but induces redistribution of 7H6 in a manner similar to BDL with partial relocation to distinct spots in the pericanalicular cytoplasm (Kawaguchi et al., 1999). The 7H6 pattern differs from BDL by the absence of a lobular gradient. Protein levels of ZO-1 decrease by 10% and increase fourfold for 7H6 (Kawaguchi et al., 1999).

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27.3.1.2.2 Functional alterations A positive correlation between impaired structural integrity and increased permeability is also shown in this model by increased biliary clearance of inert solutes (Forker, 1969; Elias et al., 1983; Jaeschke et al., 1987), which is in line with a fourfold increased number of HRP-positive junctions (Rahner et al., 1996). These data are paralleled by output of HRP during the early peak, which displayed a twofold increase (Kan et al., 1989; Rahner et al., 1996) (see Figure 27.6A). Estimates of specific junctional resistance indicate a value of 22 kΩ·cm, which is half the resistance of control junctions (Rahner et al., 1996). Size selectivity of TJs is altered minimally by EE from 70 kDa in controls up to 80 kDa (Rahner et al., 1996). 27.3.1.2.3 Effects on cell polarity EE administration does not affect the domain-specific expression of plasma membrane proteins (Landmann, 1995) (see Figure 27.7B), which in BDL relocate to the opposite domain. This indicates that the TJ fence function is not affected to a significant extent. 27.3.1.2.4 Conclusion The mechanisms mediating TJ defects remain to be elucidated. They are, however, less effective than those acting in BDL since structural, functional, and biochemical alterations are less pronounced and lead to a “mild” form of cholestasis (Reichen and Simon, 1994). 27.3.1.3 Bile Acid-Induced Cholestasis: Taurolithocholate Because of their amphipathic nature, bile salts may become toxic to liver cells with increasing intracellular concentrations by inflicting, for example, damage to mitochondria (Krähenbühl et al., 1994). In rat liver, the action of lithocholic acid is strongly cholestatic. Besides its effect on the fluidity of the canalicular membrane, lithocholic acid leads to an increase of tight junctional permeability, as shown by an increase of the inulin bile to plasma ratio and by penetration of lanthanum through the TJs from the blood compartment (Vu et al., 1992). In the perfused liver, taurolithocholate leads to an increased paracellular permeability for sucrose but not for polyethyleneglycol 900 (Ballatori and Truong, 1990). This indicates that the functional integrity of TJs is only minimally affected. It is conceivable that the mechanisms resulting in these defects involve impaired mitochondrial function with concomitant ATP depletion, which may lead to a situation similar to ischemia in kidney (see Chapter 25). 27.3.1.4 Drug- and Toxin-Induced Cholestasis Drugs and xenobiotics can lead to cholestasis in susceptible patients (Zimmermann and Lewis, 1987; Feuer and Di Fonzo, 1992). This form of cholestasis may be caused by direct inhibition of the canalicular bile salt export pump (Stieger et al., 2000), which in turn results in intracellular accumulation of bile salts. 27.3.1.4.1 Cyclosporine A Cyclosporine A is a competitive inhibitor of the canalicular bile salt export pump (Stieger et al., 2000). In addition, in the hepatocyte couplet system, cyclosporin A

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disturbs the functional integrity of TJs (Roman et al., 1996). Corroborating these findings, it was demonstrated that in the isolated perfused rat liver, cyclosporin A leads to a significantly increased early peak of HRP in bile indicating a loss in TJ gate function (Lora et al., 1997a). In addition, the couplet system showed an altered distribution of pericanalicular F-actin that was prevented by coincubation with S-adenosyl-methionine (Roman et al., 1996). Taken together, the data indicate that impairment of TJs may be induced by intracellular accumulation of toxic bile salts. 27.3.1.4.2 ANIT α-Naphthylisothiocyanate (ANIT) is converted by cytochrome P-450 into a strongly cholestatic compound (Schaffner et al., 1973). Probing of paracellular permeability with sucrose in the perfused liver demonstrated a significant ANIT-induced increase (Krell et al., 1982) associated with disorganized TJ structure (Krell et al., 1987). ANIT, with time, progressively increases the permeability of the junctional barrier for markers of increasing molecular weight (Kan and Coleman, 1986). In the early peak, increased inulin appearance was detectable after 6 h, whereas changes in HRP and ovalbumin output became effective after 12 h, and pig γ-globulin after 14 h of treatment. Induction of P-450 with phenobarbitone pretreatment increased the effect of ANIT. 27.3.1.4.3 Carmustin The anticancer drug carmustine shows a delayed hepatotoxicity with functional and histopathological alterations similar to ANIT. Hence, treatment of rats with carmustin also leads to an increase in paracellular permeability for HRP, indicative of damaged TJ function (Krell et al., 1991). 27.3.1.4.4 BSP Cholephilic dyes such as bromosulfophthalein (BSP) are widely used substances for the study of hepatocellular transport of organic anions and of bile formation. If, however, BSP is perfused at high concentration through the liver, it results in an impairment of tight junctional resistance, as shown by an increased paracellular permeability for sucrose and HRP (Roma et al., 1995). This increased sucrose permeability has also been observed for other cholephilic dyes, such as rose bengal, bromcresol green, or phenol red (Roma et al., 1996), and a strong positive correlation was found between increase of biliary permeability and hydrophobicity of the dye (Roma et al., 1995; 1996). 27.3.1.4.5 Phalloidin The fungal toxin phalloidin inhibits depolymerization of actin filaments, leads to a marked increase in pericanalicular microfilaments (Gabbiani et al., 1975), and decreases bile flow (Dubin et al., 1978; Elias et al., 1980). These changes are associated with disorganization and decreased numbers of TJ strands, with increased junctional permeability for electron-dense markers, and with increased bile-to-plasma ratios of inulin and sucrose (Elias et al., 1980). In addition, cytochalasin B, a toxin depolymerizing actin filaments, also impairs bile flow and causes dilation of canaliculi and loss of microvilli (Phillips and Satir, 1988). These observations demonstrate the dependence of TJ structural integrity and barrier function on the actin cytoskeleton (see Chapter 12). Massive pericanalicular actin accumulation similar

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to that found after experimental phalloidin injury is characteristic to North American Indian childhood cirrhosis (Weber et al., 1981), a rapidly progressing form of inherited neonatal cholestasis, which may suggest impairment of the TJs. 27.3.1.4.6 Colchicine Colchicine disrupts the microtubular system of the cytoskeleton. Colchicine treatment leads to an increase of the early peak of HRP appearance in bile, indicative of damaged TJs (Kacich et al., 1983; Lowe et al., 1985).

27.3.2 OXIDATIVE STRESS Oxidative stress induced by administration of tertiary-butylhydroxide, menadione, or hydrogen peroxide in the perfused rat liver leads to reduction in glutathion content of hepatocytes. This is paralleled by an increase of the bile-to-perfusate ratio of sucrose, but to a decrease of polyethyleneglycol 900 (Ballatori and Truong, 1989). Menadione was also shown to increase paracellular permeability for HRP, which could be partially prevented by coadministration of a free-radical scavenger (Kan and Coleman, 1990). In addition, menadione also leads to an increase in intracellular Ca2+ levels, which in turn may influence junctional integrity (Kan and Coleman, 1990). By using the isolated couplet system, the effect of tBOOH on TJ integrity was related to the generation of free radicals, especially of peroxyl radicals (AhmedChoudhoury et al., 1998). Reoxygenation following ischemia is known to inflict cellular damage. Accordingly, a reoxygenation model in the perfused liver system showed a strongly increased early HRP peak in bile (Konno et al., 1992).

27.3.3 INFLAMMATORY DISEASES Colitis as induced by intracolonic instillation of trinitrobenzene sulfonic acid in rats leads to an increase of paracellular HRP secretion into bile and a dramatic increase of hepatocellular permeability for lanthanum (Lora et al., 1997). Immunofluorescence performed in these livers showed no dramatic alterations for cingulin and ZO-1. A later study, however, demonstrated a discontinuous staining for ZO-1 and dissociation of ZO-1 from 7H6 signals (Kawaguchi et al., 2000). TJs form a barrier not only for cholephilic substances between portal blood and bile, but also for toxins entering from bile into blood. Prolonged bile duct obstruction may lead to cholangitis, which eventually can lead even to septicemia. Along these lines it was demonstrated that in BDL rats bacteria may translocate from bile into blood, a process that may be facilitated by malfunctioning TJs (Karsten et al., 1998).

27.3.4 REGENERATION

AND

CELL PROLIFERATION

Two thirds hepatectomy, the classical model for liver regeneration, results in a progressive loss of TJ strands between 20 and 40 h after resection and in subsequent reassembly (Yee and Revel, 1978). Structural and functional characterization of this model at 24 h (Poucell et al., 1992) demonstrated an impaired junctional organization and a decrease in the number of parallel strands. These defects were associated with a markedly increased HRP early peak that was still charge selective (Poucell et al.,

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1992) and reversed to normal within 6 days (Hoshino et al., 1995). The data suggest that junctional elements are loosened to allow for architectural remodeling that is necessitated by cell proliferation involved in organ regeneration. Thus, it is conceivable that cell proliferation depends on downregulated TJ function, a view that is supported by the known involvement of TJ-associated proteins such as ZOs in cellular signaling pathways (Kim, 1995; Tsukita et al., 1999). Cancer is associated with rapidly dividing cells, restructuring, and altered intercellular contacts. Rat models of liver carcinoma, consequently, showed disturbed TJ morphology (Robenek et al., 1980a) associated with decreased expression of 7H6 (Zhong et al., 1994).

27.4 CONCLUDING REMARKS This chapter concentrated on animal models that simulate human diseases. Although this experimental approach is very useful for the detailed characterization of pathological conditions, its limitations are obvious. Doses used to induce a pathological effect are often higher than those given under therapeutic conditions. In addition, the precise site or sites where the different experimental procedures take effect are rarely known to sufficient detail. Therefore, it is unclear in most cases whether observed alterations of TJs are a primary pathogenetic mechanism or, rather, a secondary epiphenomenon. Indeed, impairment of TJ structural and functional integrity may represent the final and common manifestation of multiple factors exerting their effects at widely different levels. Nevertheless, animal and cell culture models will continue to play a role in the search for pathogenetic mechanisms that will be conducted with increasing emphasis on the newly identified molecular constituents. This direction of research had already started 10 years ago when a seminal study demonstrated the altered distribution pattern of ZO-1 in BDL-induced cholestasis (Anderson et al., 1989). Subsequent reports showed altered expression levels and relocation of TJ components (Fallon et al., 1995; Kawaguchi et al., 1999; 2000) and demonstrated that dissociation of cytoplasmic or of transmembrane from plaque proteins is associated with severe but not with moderate disturbance of TJs. To understand alterations of TJs in liver disease it will be necessary to implement insight obtained from other cell culture systems in the liver. These studies could address the following topics. First, cytoskeletal elements and especially actin filaments that form the thick band of subjunctional microfilaments associated with the contractile pericanalicular ring are critical for the maintenance of TJ integrity (see Chapter 12). Thus, any interference with the structural and dynamic roles of the cytoskeleton, but also with proteins associated with it, are likely to result in severe junctional impairment. A recent example of this approach is the demonstration of marked changes in perijunctional actin filament bundles associated with a strongly increased TJ permeability by activation of small G proteins from the rho family (Nusrat et al., 1995; Jou et al., 1998). Second, at the level of the TJ plaque, specific proteins establish the link between integral membrane proteins and the cytoskeleton. The MAGUK proteins ZO-1 and ZO-2, which are likely to have both structural and signaling roles, bind actin filaments as well as the cytoplasmic tails of occludin (Fanning et al., 1998) and possibly

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claudins. In addition, a variety of cytoplasmic molecules (Mitic and Anderson, 1998), including proteins involved in signaling, associate directly or indirectly with ZOs. These proteins are key candidates for the control of TJ strand assembly, maintenance, and regulation. Third, and finally, at the level of the integral membrane proteins the availability of multiple claudins with distinct tissue distribution patterns (Morita et al., 1999) allows for an epithelium-specific expression of different isoforms. The demonstration that single-transfected claudin-2 has the competence to form strands with claudin3 expressed in a neighboring cell but not with claudin-1 (Furuse et al., 1999) led to the hypothesis that heterotypic claudin-1/2 pairs might form pores within a strand (Tsukita and Furuse, 2000). It is obvious that the combination and proportion of claudin types expressed in an epithelium exert great influence on the tightness of the barrier. The observation that claudins 3 and 4 are selectively removed from junctional strands after binding Clostridium perfringens enterotoxin (see Chapter 24), a process associated with a “cholestasis-like” freeze-fracture appearance (Sonada et al., 1999), indicates that TJ barrier function may be altered or disturbed very specifically at the claudin level. Therefore, the idea that physiological as well as pathological adaptation or change of TJ permeability depends (also) on removal and insertion of transmembrane proteins with different barrier properties is open for exploration.

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Furuse, M., Sasaki, H., Fujimoto, K., and Tsukita, S. 1998. A single gene product, claudin-1 or -2, reconstitutes tight junction strands and recruits occludin in fibroblasts. J. Cell Biol., 143: 391–401. Furuse, M., Sasaki, H., and Tsukita, S. 1999. Manner of interaction of heterogenous claudin species within and between tight junction strands. J. Cell Biol., 147: 891–903. Gabbiani, G., Montesano, R., Tuchweber, B., Salas, M., and Orci, L. 1975. Phalloidin-induced hyperplasia of actin filaments in rat hepatocytes. Lab. Invest., 33: 562–569. Gartung, C., Ananthanarayanan, M., Rahman, M. A., Schuele, S., Nundy, S., Soroka, C. J., Stolz, A., Suchy, F. J., and Boyer, J. L. 1996. Down-regulation of expression and function of the rat liver Na+/bile acid cotransporter in extrahepatic cholestasis. Gastroenterology, 110: 199–209. Gautam, A., Ng, O. C., and Boyer, J. L. 1987. Isolated rat hepatocyte couplets in short-term culture: structural characteristics and plasma membrane reorganization. Hepatology, 7: 216–224. Gautam, A., Ng, O. C., Strazzabosco, M., and Boyer, J. L. 1989. Quantitative assessment of canalicular bile formation in isolated hepatocyte couplets using microscopic optical planimetry. J. Clin. Invest., 83: 565–573. Graf, J., Gautam, A., and Boyer, J. L. 1984. Isolated rat hepatocyte couplets: a primary secretory unit for electrophysiologic studies of bile secretory function. Proc. Natl. Acad. Sci. U.S.A., 81: 6516–6520. Graf, J., Henderson, M., Krumpholz, B., and Boyer, J. L. 1987. Cell membrane and transepithelial voltages and resistances in isolated rat hepatocyte couplets. J. Membr. Biol., 95: 241–254. Hardison, W. G. M., Lowe, P. J., and Shanahan, M. 1989. Effect of molecular charge on paraand transcellular access of horseradish peroxidase into rat bile. Hepatology, 9: 866–871. Hornstein, B., Stammler, L., Bianchi, L., and Landmann, L. 1992. Ethinylestradiol increases volume and decreases sinusoidal membrane surface in rat liver: a stereological analysis. Hepatology, 16: 217–223. Hoshino, M., Hirano, A., Hayakawa, T., Kamiya, Y., Ohiwa, T., Tanaka, A., Kumai, T., Katagiri, K., Miyaji, M., and Takeuchi, T. 1995. Comparative studies on bile flow and biliary lipid excretion after bile-acid loading in normal and partially hepatectomized rats. Biochem. J., 305: 367–371. Jaeschke, H., Trummer, E., and Krell, H. 1987. Increase in biliary permeability subsequent to intrahepatic cholestasis by estradiol valerate in rats. Gastroenterology, 93: 533–538. Javitt, N. B. and Arias, I. M. 1967. Intrahepatic cholestasis: a functional approach to pathogenesis. Gastroeneterology 53: 171–175. Jou, T. S., Schneeberger, E. E., and Nelson, W. J. 1998. Structural and functional regulation of tight junctions by RhoA and Rac1 small GTPases. J. Cell Biol., 141: 101–115. Kacich, R. L., Renston, R. H., and Jones, A. L. 1983. Effects of cytochalasin D and colchicine on the uptake, translocation, and biliary secretion of horseradish peroxidase and (14C)sodium taurocholate in the rat. Gastroenterology, 85: 385–394. Kan, K. S. and Coleman, R. 1986. 1-Naphthylisothiocyanat-induced permeability of hepatic tight junctions to proteins. Biochem. J., 238: 325–328. Kan, K. S. and Coleman, R. 1990. Menadione increases hepatic tight-junctional permeability. Its effect can be decreased by butylated hydroxytoluene and verapamil. Biochem. J., 270: 241–243. Kan, K. S., Monte, M. J., Parslow, R. A., and Coleman, R. 1989. Oestradiol 17β-glucuronide increases tight-junctional permeability in rat liver. Biochem. J., 261: 297–300.

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Karsten, T. M., van Gulik, T. M., Spanjaard, L., Bosma, A., van der Bergh Weerman, M. A., Dingemans, K. P., Dankert, J., and Gouma, D. J. 1998. Bacterial translocation from the biliary tract to blood and lymph in rats with obstructive jaundice. J. Surg. Res., 74: 125–130. Kawaguchi, T., Sakisaka, S., Sata, M., Mori, M., and Tanikawa, K. 1999. Different lobular distributions of altered hepatocyte tight junctions in rat models of intrahepatic and extrahepatic cholestasis. Hepatology, 29: 205–216. Kawaguchi, T., Sakisaka, S., Mitsuyama, K., Harada, M., Koga, H., Taniguchi, E., Sasatomi, K., Kimura, R., Ueno, T., Sawada, N., Mori, M., and Sata, M. 2000. Cholestais with altered structure and function of hepatocyte tight junction and decreased expression of canalicular multispecific organic anion transporter in a rat model of colitis. Hepatology, 31: 1285–1295. Kim, S. K. 1995. Tight junctions, membrane associated guanylate kinases and cell signaling. Curr. Opin. Cell Biol., 7: 641–649. Konno, H., Lowe, P. J., Hardison, W. G., Miyai, K., Nakamura, S., and Baba, S. 1992. Breakdown of hepatic tight junctions during reoxygenation injury. Transplantation, 53: 1211–1214. Krähenbühl, S., Talos, C., Fischer, S., and Reichen, J. 1994. Toxicity of bile acids on the electron transport chain of isolated rat liver mitochondria. Hepatology, 19: 471–479. Krell H., Höke, H., and Pfaff, E. 1982. Development of intrahepatic cholestasis by α-naphthylisothiocyanate in rats. Gastroenterology, 82: 507–514. Krell, H., Metz, J., Jaeschke, H., Höke, H., and Pfaff, E. 1987. Drug-induced intrahepatic cholestasis: characterization of different pathomechanisms. Arch. Toxicol., 60: 124–130. Krell, H., Fromm, H., and Larson, R. E. 1991. Increased paracellular permeability in intrahepatic cholestasis induced by carmustine (BCNU) in rats. Gastroenterology, 101: 180–188. Kullak-Ublick, G. A., Stieger, B., Hagenbuch, B., and Meier, P. J. 2000. Hepatic transport of bile salts. Semin. Liver Dis., 20: 273–292. Landmann, L. 1995. Cholestasis-induced alterations of the trans- and paracellular pathways in rat hepatocytes. Histochem. Cell Biol., 103: 3–9. Landmann, L., Meier, P. J., and Bianchi, L. 1990. Bile duct ligation-induced redistribution of canalicular antigen in rat hepatocyte plasma membranes demonstrated by immunogold quantification. Histochemistry, 94: 373–379. Landmann, L., Angermüller, S., Rahner, C., and Stieger, B. 1998. Expression, distribution and activity of Na+,K+-ATPase in normal and cholestatic rat liver. J. Histochem. Cytochem., 46: 405–410. Lora, L., Mazzon, E., Billington, D., Milanesi, C., Naccarato, R., and Martines, D. 1997a. Effects of cyclosporin A on paracellular and transcellular transport of horseradish peroxidase in perfused rat livers. Digest. Dis. Sci., 42: 514–521. Lora, L., Mazzon, E., Martines, D., Fries, W., Muraca, M., Martin, A., Dodorico, A., Naccarato, R., and Citi, S. 1997b. Hepatocyte tight-junctional permeability is increased in rat experimental colitis. Gastroenterology, 113: 1347–1354. Lowe, P. J., Kan, K. S., Barnwell, S. G., Sharma, R. K., and Coleman, R. 1985. Transcytosis and paracellular movements of horseradish peroxidase across liver parenchymal tissue from blood to bile. Effects of alpha-naphthylisothiocyanate and colchicine. Biochem. J., 229: 529–537. Lowe, P. J., Miyai, K., Steinbach, J. H., and Hardison, W. G. M. 1988. Hormonal regulation of hepatocyte tight junctional permeability. Am. J. Physiol., 255: G454–G461. Madara, J. L. and Dharmsathaphorn, K. 1985. Occluding junction structure–function relationships in a cultured epithelial monolayer. J. Cell Biol., 101: 2124–2133.

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Marcial, M. A., Carlson, L., and Madara, J. L. 1984. Partitioning of paracellular conductance along the ileal crypt-villous axis: a hypothesis based on structural analysis with detailed consideration of tight junction structure–function relationship. J. Membr. Biol., 80: 59–70. Metz, J. and Bressler, D. 1979. Reformation of gap and tight junctions in regenerating liver after cholestasis. Cell Tissue Res., 199: 257–270. Metz, J., Aoki, A., Merlo, M., and Forssmann, W. G. 1977. Morphological alterations and functional changes of interhepatocellular junctions induced by bile duct ligation. Cell Tissue Res., 182: 299–310. Mitic, L. L. and Anderson, J. M. 1998. Molecular architecture of tight junctions. Annu. Rev. Physiol., 60: 121–142. Morita, K., Furuse, M., Fujimoto, K., and Tsukita, S. 1999. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc. Natl. Acad. Sci. U.S.A., 96: 511–516. Nathanson, M. H. and Boyer, J. L. 1991. Mechanism and regulation of bile secretion. Hepatology, 14: 551–565. Nathanson, M. H. and Burgstahler, A. D. 1992. Subcellular distribution of cytosolic Ca2+ in isolated rat hepatocyte couplets — evaluation using confocal microscopy. Cell Calcium, 13: 89–98. Nusrat, A., Giry, M., Turner, J. R., Colgan, S. P., Parkos, C. A., Carnes, D., Lemichez, E., Boquet, P., and Madara, J. L. 1995. Rho protein regulates tight junctions and perijunctional actin organization in polarized epithelia. Proc. Natl. Acad. Sci. U.S.A., 92: 10629–10633. Oguey, D., Marti, U., and Reichen, J. 1992. Epidermal growth factor receptor in chronic bile duct obstructed rats — implications for maintenance of hepatocellular mass. Eur. J. Cell Biol., 59: 187–195. Phillips, M. J. and Satir, P. 1988. The cytoskeleton of the hepatocyte: Organization, relationships, and pathology, in The Liver: Biology and Pathobiology, 2nd ed., Arias, I. M., Jakoby, W. B., Popper, H., Schachter, D., and Shafritz, D. A., Eds., Raven Press, New York. 11–27. Popper, H. and Szanto, P. B. 1956. Intrahepatic cholestasis (cholangiolitis). Gastroenterology, 31: 683–700. Poucell, S., Hardison, W. G. M., and Miyai, K. 1992. Regenerative stimulus increases hepatocyte tight junctional permeability. Hepatology, 16: 1061–1068. Powell, D. W. 1981. Barrier function of epithelia. Am. J. Physiol., 241: G275–G288. Rahner, C., Stieger, B., and Landmann, L. 1996. Structure–function correlation of tight junctional impairment after intrahepatic and extrahepatic cholestasis in rat liver. Gastroenterology, 110: 1564–1578. Reichen, J. and Simon, F. R. 1994. Cholestasis, in The Liver. Biology and Pathobiology, 3rd ed., Arias, I. M., Boyer, J. L., Fausto, N., Jakoby, W. B., Schachter, D., Shafritz, D. A., Eds., Raven Press, New York, 1291–1326. Robenek, H., Döldissen, M., and Themann, H. 1980a. Morphological changes of tight junctions in the rat liver after chronic administration of N-nitrosomorpholine (NNM) as revealed by freeze-fracturing. J. Ultrastruct. Res., 70: 82–91. Robenek, H., Herwig, J., and Themann, H. 1980b. The morphologic characteristics of intercellular junctions between normal human liver cells and cells from patients with extrahepatic cholestasis. Am. J. Pathol., 100: 93–114. Robenek, H., Rassat, J., Grosser, V., and Themann, H. 1982. Ultrastructural study of cholestasis induced by long term treatment with estradiol valerate. I. Tight junctional analysis and tracer experiments. Virchows Arch. B, 40: 201–215.

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Roma, M. G., Marinelli, R. A., Crocenzi, F. A., and Garay, E. A. R. 1995. Effect of cholephilic dyes on hepatic tight junctional permeability in the rat. Biochem. Pharmacol., 50: 1079–1086. Roma, M. G., Crocenzi, F. A., and Rodriguez Garay, E. A. 1996. Does paracellular permeability play a role in cholephilic dye-induced cholestasis? Toxicol. Lett., 84: 13–22. Roma, M. G., Orsler, D. J., and Coleman, R. 1997. Canalicular retention as an in vitro assay of tight junctional permeability in isolated hepatocyte couplets: effects of protein kinase modulation and cholestatic agents. Fundam. Appl. Toxicol., 37: 71–81. Roman, I. D., Johnson, G. D., and Coleman, R. 1996. S-Adenosyl-l-methionine prevents disruption of canalicular function and pericanalicular cytoskeleton integrity caused by cyclosporin A in isolated rat hepatocyte couplets. Hepatology, 24: 134–140. Schaffner, F., Scharnbeck, H., Hutterer, F., Denk, H., Greim, H. A., and Popper, H. 1973. Mechanism of cholestasis: VII. α-Naphthyl isothiocynate-induced cholestasis. Lab. Invest., 28: 321–331. Sellinger, M. and Boyer, J. L. 1990. Physiology of bile secretion and cholestasis. Prog. Liver Dis., 9: 237–259. Sonada, N., Furuse, M., Sasaki, H., Yonemura, S., Katahira, J., Horiguhi, Y., and Tsukita, S. 1999. Clostridium perfringens enterotoxin fragment removes specific claudins from tight junction strands. Evidence for direct involvement of claudins in tight junction barrier. J. Cell Biol., 147: 195–204. Staehelin, L. A. 1974. Structure and function of intercellular junctions. Int. Rev. Cytol., 39: 191–284. Stammler, L., Reichen, J., Oehler, R., Bianchi, L., and Landmann, L. 1990. Decreased hepatocellular volume and intact morphology of tight junctions in calcium deprivation-induced cholestasis: stereological and multiple indicator dilution analysis. J. Hepatol., 10: 318–326. Stieger, B., Meier, P., and Landmann, L. 1994. Effect of obstructive cholestasis on membrane traffic and domain-specific expression of plasma membrane proteins in rat liver parenchymal cells. Hepatology, 20: 201–212. Stieger, B., Fattinger, K., Madon, J., Kullak-Ublick, G. A., and Meier, P. J. 2000. Drug- and estrogen-induced cholestasis through inhibition of the hepatocellular bile salt export pump (Bsep) of rat liver. Gastroenterology, 118: 422–430. Toyota, N., Miyai, K., and Hardison, G. M. 1984. Effect of biliary pressure vs. high bile acid flux on the permeability of hepatocellular tight junction. Lab. Invest., 50: 536–542. Trauner, M., Meier, P. J., and Boyer, J. L. 1998. Mechanisms of disease: molecular pathogenesis of cholestasis. N. Engl. J. Med., 339: 1217–1227. Tsukita, S. and Furuse, M. 2000. Pores in the wall: claudins constitute tight junction strands containing aqueous pores. J. Cell Biol., 149: 13–16. Tsukita, S., Furuse, M., and Itoh, M. 1999. Structural and signaling molecules come together at tight junctions. Curr. Opin. Cell Biol., 11: 628–633. VanDeurs, B. and Luft, J. H. 1979. Effects of glutaraldehyde fixation on the structure of tight junctions. A quantitative freeze-fracture analysis. J. Ultrastruct. Res., 68: 160–172. Vu, D. D., Tuchweber, B., Raymond, P., and Yousef, I. M. 1992. Tight junction permeability and liver plasma membrane fluidity in lithocholate-induced cholestasis. Exp. Mol. Pathol., 57: 47–61. Weber, A. M., Tuchweber, B., Yousef, I., Brochu, P., Turgeon, C., Gabbiani, G., Morin, C. L., and Roy, C. C. 1981. Severe familial cholestasis in North American Indian children — a microfilament dysfunction. Gastroenterology, 81: 653–662. Weibel, E. R. 1979. Stereological methods. Vol I. Practical Methods for Biological Morphometry. Academic Press, London, reprinted 1987.

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The Tight Junctions in the Testis, Epididymis, and Vas Deferens R.-Marc Pelletier

CONTENTS 28.1 Introduction .................................................................................................600 28.2 The Tight Junctions in the Testis ...............................................................600 28.2.1 Topographical Distribution of Intercellular Junctions in the Seminiferous Epithelium ...............................................................600 28.2.2 Physiological Consequences of the Establishment of Occluding Junctions in the Seminiferous Epithelium ....................................605 28.2.2.1 First Consequence .........................................................605 28.2.2.2 Second Consequence.....................................................606 28.2.2.3 Third Consequence........................................................607 28.2.2.4 Fourth Consequence ......................................................608 28.2.3 Synchronization of the Sertoli Cell Junctional Barrier Function is Kept in Time with the Development of the Germ Cells ..........609 28.3 Factors Susceptible to Influence the Permeability Status of the Sertoli Cell Occluding Zonules ..............................................................................610 28.3.1 The Number of Junctional Fibrils or Their Pattern within the Zonules...........................................................................................610 28.3.2 The Differential Distribution of the Junctional Particles within the Plane of the Cellular Membrane .............................................613 28.4 Proteinic Constituents of the Sertoli Cell Occluding Junctions ................614 28.4.1 The Integral Membrane Proteins...................................................614 28.4.2 Peripheral Occluding-Junction-Associated Proteins and the Subsurface or Cortical Actin in the Sertoli Cells .........................615 28.5 Mechanisms for the Assembly and Disassembly of Sertoli Cell Occluding Zonules ......................................................................................616 28.5.1 The “Zipper” Theory .....................................................................616 28.5.2 The Theory of an Intermediate Cellular Compartment within the Seminiferous Epithelium..............................................617 28.5.3 The Theory of Repetitive Removals of Membrane Segments from the Sertoli Cell Junctional Complex ...................................617

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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28.6 The Occluding Zonules Do Not Serve as an Immunological Barrier to Anti-Spleen Antibodies...........................................................................618 28.7 The Occluding Junctions in the Epididymis and Vas Deferens ................620 Acknowledgment ...................................................................................................622 References..............................................................................................................622

28.1 INTRODUCTION The testis is composed of two distinct compartments, the seminiferous tubules and the interstitial tissue. The tubules are primarily involved in producing spermatozoa. The male gametes result from many cellular divisions including a special type of cellular division, meiosis, which allows exchanges of paternal and maternal genetic information by reducing the number of chromosomes by half from diploid to haploid. The seminiferous tubules provide a “special milieu” for the successful completion of meiosis by sequestering the dividing testicular germ cells from the blood with a blood–tissue barrier. In addition to this blood–testis barrier, which separates selected generations of germ cells dividing within the testis from the rest of the body, a blood–epididymis barrier isolates the spermatozoa from the blood while they mature in the epididymis and a blood–vas-deferens barrier secludes the gametes from the blood during their transit through the vas deferens to the exterior of the body. This chapter focuses on the junctional elements that constitute these blood–tissue barriers in the male reproductive system, namely, the TJs.

28.2 THE TIGHT JUNCTIONS IN THE TESTIS 28.2.1 TOPOGRAPHICAL DISTRIBUTION OF INTERCELLULAR JUNCTIONS IN THE SEMINIFEROUS EPITHELIUM Diagrams a and b relate the various junctional membrane specializations to their actual topography along the lateral and apical membranes of a Sertoli cell. Diagram a depicts a schematic view of the histological organization of the seminiferous epithelium as it appears in electron microscopy of thin sections with the emphasis on the intercellular junctions that are established between processes of adjacent Sertoli cells and between a Sertoli cell and the many germ cells. Diagram b shows the intramembranous specializations corresponding to the intercellular junctions depicted in diagram a as they are viewed within the interior of the plasma membrane in electron microscopy of a freeze-fracture replica as it would appear if the replicated plasma membrane of one Sertoli cell were visible from the base to the apex of the cell. The seminiferous epithelium lining the tubules is inhabited by two distinct cell types, the germ cells and the somatic cells called Sertoli cells that support and nourish the germ cells. The Sertoli cells are long, cuboidal cells that typically extend from the base of the epithelium to the lumen of the tubule while sending from the cell body thin cytoplasmic processes that surround each immediately adjacent germ cell (see Diagram a). Thus, the base of any given Sertoli cell contacts the limiting membrane of the tubule through hemidesmosomes (Russell, 1977a) while the

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remaining of the body of the cell with the extending cytoplasmic processes contact adjacent Sertoli cells and germ cells through Sertoli cell–Sertoli cell and Sertoli cell–germ cell junctions (Diagrams a and b) (see reviews by Byers et al., 1993; Pelletier, 1998; Pelletier and Byers, 1992). The contact of the base of the Sertoli cells with the basement membrane matrix of the tubule in which the laminins constitute the major noncollagenous components is apparently essential to the preservation of the integrity of the seminiferous epithelium. Passive immunization with antilaminin immunoglobulin G has been reported to induce arrest of spermatogenesis and to cause changes in Sertoli cell TJs or occluding junctions in vivo (Lustig et al., 2000). In addition, it is important to emphasize that the Sertoli cell plasma membrane segments facing the germ cell are included in the continuum of the Sertoli cell junctional complex that extends over the lateral and apical membranes of the Sertoli cell (Diagrams a and b). The distinct features of the cellular junctions appearing over the membranes extending from the base to the apex of the Sertoli cell represent different moments in the assembly/disassembly of the various junctional elements forming the complex. In the adult mammalian testis, the body or “trunk” of adjacent Sertoli cells is involved in establishing a distinct type of intercellular contacts including adhering junctions (Kaya and Harrison, 1976; Russell, 1977a), gap junctions (McGinley et al., 1977; Szöllösi and Marcaillou, 1980), and TJs or occluding junctions (McGinley et al., 1979), which together constitute the Sertoli cell junctional complex. Each type of intercellular contact within the complex is illustrated in Diagrams a and b and Figures 28.1a to h, 28.2a,b, 28.3a,b. By sealing the intercellular space between Sertoli cells, occluding junctions help to build a gradient from underlying interstitial fluids to the lumenal fluids (Setchell et al., 1969). The comprehensive ultrastructural description by Dym and Fawcett (1970) showed that the complex contains areas of Sertoli cell membrane approximation of up to 10 nm called “close” junctions (Figure 28.1h), gap junctions characterized by a narrowing of the intercellular space to 2 to 4 nm (Figures 28.1c, e, f, and g), and occluding junctions, which constitute DIAGRAMS A AND B (see pages 601 and 602) Diagrams illustrating the distribution of the different types of cell junctions as they appear in electron microscopy of thin section (a) and of freeze-fracture replicas (b). (a) Each generation of germ cells (which have been shaded to facilitate their identification) — except the elongated spermatids — is lodged in a cleft that is closed at both ends by cell junctions. The junctions present along the lateral and apicolateral membranes of the Sertoli cell (which have been hatched to facilitate their identification) may involve either adjacent Sertoli cell cytoplasmic processes or a Sertoli cell and the many germ cells. JV = junction vesicles or annular profiles of junctional membranes contain ancient intercellular junctions; AV = autophagic vacuoles contain junction vesicles. (b) In freeze-fracture replicas, fusions of the outer membrane leaflet of one Sertoli cell with the outer membrane leaflet of another at the site of an occluding junction are visualized as ridges on the P-fracture face (represented in this diagram by solid lines) and corresponding grooves on the E face (not represented). The solid lines are horizontal and continuous in the occluding zonule near the basal third of the Sertoli cell; they are short and discontinuous in the focal occluding junctions. The gap junctions (represented by dotted lines or as aggregates of dots) are intercalated among the ridges. Within the continuous zonule, gap junctions are few but their number and size increase apically. (Modified from Pelletier and Byers, 1992.)

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FIGURE 28.1 (a) Thin section of junctions forming below a migrating spermatocyte. Adjacent membranes are approximated to more than 20 nm apart over most of their surface. A fuzzy coating is apparent on the membrane surface. In regions where membranes are approximated to 20 nm, the fuzzy coating is condensed into a denser line that occupies the intercellular cleft and is typical of the adhering junction (A). Sporadic, small agglomerates of 7-nm filaments are identified (f). Tissues were taken from an adult mink testis harvested during the active spermatogenic phase. Original magnification ×53,900. (b) On the right side of the micrograph, a well-developed adhering junction is identified (A). To the left of this adhering junction, facing plasma membranes are wavy and positioned more than 20 nm apart. A fuzzy material is present in the intercellular cleft; 7-nm subsurface filaments (f) are identified along the adhering junction and where new junctions are forming. Tissue taken from an adult mink testis harvested during the active spermatogenic phase. The Sertoli cell membrane segments illustrated are situated below a migrating spermatocyte. Original magnification ×73,000. (c) Thin section of junctions forming between Sertoli cells in a 10-day-old guinea pig. A gap (G) and an occluding (O) junction are identified; 7-nm subsurface filaments accompany the junctions. A fuzzy material is present in the intercellular cleft of the newly developed adhering junction (A) and also in the cleft situated on either side of the occluding junction. Original magnification ×78,730. (d) Junctions forming between rat Sertoli cells in the two-chamber assembly by peritubular cells and extracellular matrix share features that are similar to those just illustrated in vivo. The facing membranes are wavy and positioned more than 20 nm apart, and a fuzzy material is apparent in the intercellular cleft. The 7-nm subsurface filaments (f) along membrane segments where approximation of membranes and junction formation take place seem more abundant in vitro than in vivo. Original magnification ×53,900. (e) Gap and/or occluding junctions (closed arrows) are interspersed between sacculations (open arrows). Subsurface 7-nm filaments (f) typically accompany the Sertoli cell junctions. rER = cisternae of endoplasmic reticulum. Tissue taken from a mink testis harvested

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the anatomical basis of the blood–testis barrier in adult rat. Adhering junctions (i.e., adhering fascia) characterized by a widening of the intercellular space to ~20 nm (Figures 28.1a, b, d, and f) have also been reported in the complex (Pelletier and Friend, 1983; Pelletier, 1988; 1990). These Sertoli cell occluding junctions have been reported in most species including humans, monkey, dog, mink, rodent, birds, fish, amphibians, reptiles, invertebrates, insects, and nematodes (reviewed in Abraham, 1991; Pelletier and Byers, 1992). The freeze-fracture technique contributes yet another parameter to the study of cell junctions by allowing direct visualization of the interior of the cell membrane at the contact site. Thus, each occluding junction that is visible in thin section as an apparent fusion of the outer membrane leaflet of one cell with the outer membrane leaflet of another (Diagram a and Figures 28.1c, e, and f) corresponds in freezefracture replicas to a fibril made up of intramembranous particles either sitting within a shallow groove on the E fracture face (Figures 28.2a, 28.3a, 28.4, and 28.5) or aligned atop a ridge on the P fracture face (Figures 28.6 and 28.7).

28.2.2 PHYSIOLOGICAL CONSEQUENCES OF THE ESTABLISHMENT OF OCCLUDING JUNCTIONS IN THE SEMINIFEROUS EPITHELIUM 28.2.2.1 First Consequence: The Junctions Seal the Paracellular Route between All Sertoli Cells and This Forces Access to Selected Germ Cells to Take Place through the Sertoli Cells Four physiological consequences follow the establishment of occluding junctions in the seminiferous epithelium. The occluding junctions seal the paracellular route between cells and thus help create a selective barrier (Figures 28.2b and 28.3b) (see reviews by Anderson and Van Itallie, 1995; Gumbiner, 1993; Jou et al., 1998; Mitic and Anderson, 1998; Schneeberger and Lynch, 1992). When occluding zonules are established between all Sertoli cells lining the seminiferous epithelium, they collectively form a blood–tissue barrier called the blood–testis barrier. The nutriments and FIGURE 28.1 (continued) during the active spermatogenic phase. Original magnification ×72,900. (f) Adhering junctions (A) are easily recognizable because of the presence in these cellular contacts of an amorphous material condensed into a denser line bisecting the intercellular cleft. The adhering junctions are frequently interspersed between gap (G) and occluding junctions. F = 7-nm filaments; rER = cisternae of rough endoplasmic reticulum. Testicular tissue taken from an adult mink during the active spermatogenic phase. Original magnification ×45,800. (g) Gap junctions (Gap) are extensive in Sertoli cell membrane segments situated lumenal to the continuous zonule. Tissue taken from a mink testis during the active spermatogenic phase. f = 7-nm subsurface filaments; rER = cisterna of rough endoplasmic reticulum. Original magnification ×72,900. (h) A so-called close or narrow junction characterized by an approximation of adjacent cultured Sertoli cells in the two-chamber assembly by peritubular cells and extracellular matrix to about 9 to 10 nm. f = subsurface 7-nm filaments; rER = cisternae of rough endoplasmic reticulum. Original magnification ×53,900. (Parts 28.a, b, d, e, f, g, and h from Byers, S. W. and Pelletier, R.-M., in Tight Junctions, Cereijido, M., Ed., CRC Press, Boca Raton, FL, 1992. With permission.)

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FIGURE 28.2 (a) Each occluding junction visible in Figure 28.1b corresponds in this freezefracture replica to a fibril of intramembranous particles located in a groove in the E-fracture face. In the occluding zonule visible in Figure 28.1a, many narrowly spaced fibrils form a zone that encircle the Sertoli cell body. Original magnification ×38,000. (b) Only the fibrils that show no discontinuities constitute an occluding zonule and a blood–testis barrier to the entry of vascularly infused HRP. Note that the tubulobulbar complex (tb) and macular occluding junctions within the complex are permeable to the permeability tracer. Harvested from an adult guinea pig. Original magnification ×28,100. (From Pelletier, R.-M., in Male Reproduction: A Multidisciplinary Overview, Martínez-Garcia, F. and Regadera, J., Eds., Churchill Communications Europe España, Spain, 1998. With permission.)

other factors originating from the interstitial space of the testis are constrained to reach germ cells situated beyond the barrier through Sertoli cells that act as a “filter” or a “modulator.” Thus, in the adult testis, the Sertoli cell is the direct regulator of spermatogenesis because the barrier renders germ cells dependent upon Sertoli cells for their very survival; there simply is no other access to the older germ cells but through Sertoli cells. This is the fundamental functional consequence of the blood–testis barrier. 28.2.2.2 Second Consequence: Because of the Particular Topography of the Occluding Zonules Joining the Sertoli Cells, the Occluding Zonules Divide the Seminiferous Epithelium into Two Contiguous Cellular Compartments and Thus Create a Special Milieu In the adult testis, Sertoli cells bear apparent similarity in design to most epithelial cells and the occluding zonules between them divide the seminiferous epithelium into not three (Russell, 1977b; 1978), but only two (Dym and Fawcett, 1970; Pelletier and Friend, 1983; Cavicchia and Sacerdote, 1988) distinct cellular compartments: lumenal above the zonule and basal below (Pelletier, 1990). During spermatogenesis, the Sertoli cell apical cytoplasmic process expands lumenally far beyond the occluding zonule to accommodate the continual proliferation of germ cells; therefore, the blood–testis barrier is seemingly situated in the basal third of the epithelium, although the zonule is the landmark between apical and lateral Sertoli cell membranes (Pelletier, 1990). Since in the testis the occluding zonules join cytoplasmic processes of Sertoli cells that are positioned over the spermatogonia, the leptotene

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FIGURE 28.3 (a) This freeze-fracture replica shows an occluding macule. It is characterized by the fact that the junctional fibrils exist in short segments that constitute a punctual feature on the fracture face rather than a zone that surrounds the whole cellular body. This particular zonule is situated over the occluding zonule (arrow) and shows obvious signs of disassembly, such as the absence of junction particles in the grooves situated in the E-fracture face. Harvested from an adult mink during the active spermatogenic phase. Original magnification ×38,000. (b) A gap junction permeated by vascularly infused HRP is identifiable. The micrograph shows a segment of a Sertoli cell–Sertoli cell junction made up of several occluding macules, which constitutes only a punctual seal to the entry of permeability tracers and which can therefore easily be bypassed. Harvested from an adult mink during testicular regression. Original magnification ×38,000. (From Pelletier, R.-M., in Male Reproduction: A Multidisciplinary Overview, Martínez-Garcia, F. and Regadera, J., Eds., Churchill Communications Europe España, Spain, 1998. With permission.)

and the early zygotene spermatocytes, the barrier divides the seminiferous epithelium into a basal cellular compartment adjacent to the basal lamina and in contact with bloodborne substances and a lumenal cellular compartment sequestered from the blood. Meiosis is initiated in the basal compartment, but its completion takes place in the lumenal compartment away from the blood and interstitial fluids. 28.2.2.3 Third Consequence: The Junction Contributes to the Establishment and Maintenance of the Sertoli Cell Polarity Occluding junctions have traditionally served as the landmark that separates the apical above from the lateral plasma membrane below in most epithelia. By limiting access of constituents of the basolateral membrane to the apical membrane, the occluding junctions contribute to the establishment and maintenance of the cell polarity (Rodriguez-Boulan and Nelson, 1989; Cereijido et al., 1998; Yap et al., 1998). Observations made in the testes of seasonal breeders that show that the competence of the Sertoli cell occluding zonules to act as a blood barrier is coincident with the development of a lumen in the seminiferous tubule rather than with the appearance of a particular class of germ cell (Pelletier, 1986; 1990) are consistent

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with this view. In the three first physiological consequences of the establishment of occluding junctions in the seminiferous epithelium, the junctions are considered a physical barrier either to the passage of extracellular fluids or to the movement of constituents within the plane of the membrane. There is no doubt that the chemical nature of the junctional membranes will influence the movement of its constituents and the behavior of the junctions. Many authors have questioned whether junctional particles are composed of proteins or lipids (Staehelin, 1973; van Deurs and Koehler, 1979; Kachar and Reese, 1982; Pinto da Silva and Kachar, 1982). The recent identification of occludin (Furuse et al., 1993; Fujimoto, 1995; Ando-Akatsuka et al., 1996) and claudin (Furuse et al., 1998; Morita et al., 1999) (see Chapter 10 by Mitic and Van Itallie on occludin and claudin) among the constituents of the fibrils suggests an important proteinic contribution to the occluding junction without excluding a possible involvement of lipids. Studies using a two-step enzymatic method involving cholesterol esterase and cholesterol oxidase have localized cholesterol to the Sertoli cell junctional membranes (Pelletier and Vitale, 1994). The result is in agreement with the widely accepted view that plasma membranes are relatively rich in sterols (reviewed in Reinhart, 1990), but it contradicts previous reports based solely on freeze-fracture filipin cytochemistry that had suggested that junctional membranes were poor in cholesterol (Elias et al., 1979). 28.2.2.4 Fourth Consequence: Occluding Junctions Constitute Either Focal Sites of Intercellular Attachment Called Occluding Macules or a Paracellular Diffusion Barrier Called the Occluding Zonule The occluding junctions constitute sites of attachment between adjacent cells. There are no morphological features identifiable in electron microscopy of thin sections allowing one to decide when the junctions create a paracellular seal and when they constitute a site of attachment. Only the freeze-fracture replication of cellular membranes allows assessment of whether the fibrils that constitute the junction are continuous, and form an uninterrupted belt called a zonule around the cellular body (Diagram b, Figure 28.2a) or whether they are discontinuous and form interrupted insular fibrils covering small spots or macules on the plasma membrane (Diagram b, Figure 28.3a). The occluding zonule and the occluding macule are impermeable to vascularly infused permeability tracers because they both seal all or part, respectively, of the intercellular space between the plasma membrane of adjacent cells (Diagram b, Figures 28.2b and 28.3b). However, the physiological difference between the two junctional elements resides in that the occluding zonule separates apical from basolateral fluid compartments and thus forms a blood–tissue barrier, whereas the occluding macule constitutes only a punctual or focal seal that can readily be bypassed by interstitial fluids or tracers (Figures 28.2b and 28.3b). This focal contact serves more as a local of intercellular attachment than as a true paracellular diffusion barrier. For that reason, it should be emphasized that the terms “tight or occluding junction” and “occluding zonule” are not synonyms. Nowhere else is this reality better illustrated than in the testis where assembly of TJs and assembly of occluding zonules are not necessarily synchronous events. In this organ,

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focal occluding junctions (Figures 28.1c and 28.3a, c) develop between Sertoli cells during fetal life (Nagano and Suzuki, 1978; Hatier and Grignon, 1980). Around puberty, the discontinuous junctional fibrils of the focal occluding junctions become continuous and organize themselves into an impermeable occluding zonule (Figures 28.1e, f, and 28.2a, b) that encircles each supporting Sertoli cell and contributes to the creation of a selective blood–testis barrier within the seminiferous epithelium (Dym and Fawcett, 1970; Vitale et al., 1973; Pelletier and Friend, 1983). The junctional fibrils that form the occluding zonules are continuous but those of the focal occluding junctions situated above and below the zonules are discontinuous and represent moments in the assembly/disassembly of the zonule (Diagrams a and b) (Pelletier and Friend, 1983; Pelletier, 1988; 1990). This is in sharp contrast to the occluding junctions of the epididymis and deferent duct which are assembled into zonules in utero and are already impermeable by birth long before the Sertoli cell occluding zonules (Pelletier, 1994; 1995a).

28.2.3 SYNCHRONIZATION OF THE SERTOLI CELL JUNCTIONAL BARRIER FUNCTION IS KEPT IN TIME WITH THE DEVELOPMENT OF THE GERM CELLS In germ cells, mitosis involves a normal karyokinesis followed by an incomplete cytokinesis, thus resulting in the creation of up to hundreds of cells connected by intercellular bridges (Fawcett et al., 1959). Consequently, each group or syncytium of germ cells extends to contact many Sertoli cells. The synchronization of physiological activities in selected Sertoli cells to accommodate the passage or translocation of cohorts of germ cells that are not necessarily all at the same stage of development along the seminiferous tubules, and this at a very precise moment of their cellular development into the lumenal compartment of the epithelium, remains a challenging question. In mammals, the distribution of Cx43 in the tubules was reported to change in accordance with the germ cell differentiation and with the modulation of the Sertoli cell occluding zonules in a stage-dependent manner (Pelletier, 1995b). A direct association of Cx43 with the occluding junction protein ZO-1 has been reported in cardiac myocytes (Giepmans and Moolenaar, 1998; Toyofuku et al., 1998). In Leydig cells or the endothelial cells of the vessels in the interstitial compartment of the testis (Pérez-Armendariz et al., 1994; 1995; Varanda and de Carvalho, 1994) and in most epithelia of the body, Cx43 positive gap junctions are not intercalated among occluding junctional fibrils as they typically are in the Sertoli cells (Figure 28.4). An important proportion of gap junctions intercalated among Sertoli cell junctional fibrils are connexin 43 (Cx43) positive (Risley et al., 1992; Tan et al., 1996). The occluding junction excludes whereas gap junctions have a “socializing” action, two seemingly opposed functions that when mixed could result in the action of a junctional complex being synchronized with the physiological activities of the cells they join. Electrophysiological measurements and dye injection experiments have established the competence of gap junctions in cell-to-cell communications in testicular tissues (Eusebi et al., 1983; Kawa, 1987; Risley et al., 1992; Enders, 1993). It is generally agreed that the germ cells in the basal compartment must translocate beyond the junctional barrier into the lumenal compartment without

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FIGURE 28.4 Freeze-fracture replica showing a discontinuous zonule between adjacent Sertoli cells. This type of zonule is characterized by the presence of interrupted furrows (arrowheads) localized atop ridges on the P-fracture face and by the presence of gap junctions (arrows) that are typically intercalated among the furrows. Original magnification ×78,300.

causing significant leakage of bloodborne substances into the tubule. Because this process takes place at a precise moment of the germ cell development during meiosis, there must exist a mechanism to ensure that the synchronization of the Sertoli cell junctional barrier function is kept in time with the development of the germ cells. The gap junctions intercalated among junctional fibrils could help synchronize assembly/disassembly of occluding zonules established between Sertoli cells in tandem with the development of the germ cells along the seminiferous tubules. In this perspective, perhaps a cyclical phosphorylation/dephosphorylation of the Cx43 positive gap junctions could influence Sertoli cell–Sertoli cell communication and play a critical role in the synchronization of their junctional barrier function (Pelletier et al., 2001). Intercalated gap junctions are more extensive and numerous over the lumenal than over the lateral Sertoli cell membranes (Pelletier and Friend, 1983) and show different states of phosphorylation that could reflect different functional states (Pelletier et al., 2001).

28.3 FACTORS SUSCEPTIBLE TO INFLUENCE THE PERMEABILITY STATUS OF THE SERTOLI CELL OCCLUDING ZONULES 28.3.1 THE NUMBER OF JUNCTIONAL FIBRILS WITHIN THE ZONULES

OR

THEIR PATTERN

The number of fibrils forming occluding zonules can reach 100 (Figures 28.2a, 28.4, and 28.6a) and because the number of fibrils has been related to the impermeability of a junction (Pricam et al., 1974; Claude, 1978; Easter et al., 1983; Marcial et al.,

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FIGURE 28.5 Freeze-fracture replica of an occluding zonule adjoining adjacent Sertoli cells and localized close to the tubular lumen. In this micrograph, the apex of the Sertoli cells is located toward the top and the base is located toward the bottom. An apical network composed of only a few junctional strands of particles (arrow) that are associated preferentially with the E-fracture face is the only membrane specialization remaining at the end of the inactive spermatogenic phase in the adult mallard duck. Original magnification ×40,000. (From Pelletier, 1990.)

1984; Madara and Dharmsathaphorn, 1985), the Sertoli cell occluding zonule was once considered “one of the tightest junctions of the body” (Gilula et al., 1976). However, actual measurements of the degree of transepithelial resistance (TER) in the Sertoli cell occluding zonule has been reported to be only in 60 Ω·cm2, which is a value typical of leaky epithelia such as the vascular endothelium (Onoda et al., 1990). The filipin tracer technique, which exposes blockage or passage of a permeability probe in freeze-fractured membranes, helped conclude that neither the number nor the distinctive patterns of the fibrils but rather the continuity of the fibrils in the zonule dictates the permeability characteristics to the intercellular contact (Pelletier, 1990; Byers and Pelletier, 1992). In seasonal breeding birds, the number of fibrils per occluding zonule is reduced to about five during the annual testicular regression (Figures 28.5 and 28.6b); yet these zonules remain as impermeable to junctional permeability tracers as do the zonules that contain 100 fibrils during the active spermatogenic phase (Pelletier, 1990). The continual upward displacement of large numbers of migrating germ cells against the resisting impermeable occluding zonules

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FIGURE 28.6 (a) The junctional particles are no longer associated preferentially with the grooves of the E face in this freeze-fracture replica taken from an adult mink during the seasonal testicular regression. The arrow and the arrowhead, respectively, point to spherical and fibrillar particles associated with the P face. The pattern of the strands is also quite different from that illustrated during the breeding season (compare with Figure 28.2a). Original magnification ×53,900. (b) Freeze-fracture replica of a zonule adjoining two Sertoli cells in an adult duck during the seasonal testicular regression. The lumen is at the top of the micrograph. The occluding zonule is composed of only five to seven meandering rows of junctional particles that are associated preferentially with the P face (ridge). Its location and anatomical feature are reminiscent of the ones described (Staehelin, 1973; 1974) in other epithelia. Original magnification ×53,900. (From Byers, S. W. and Pelletier, R.-M., in Tight Junctions, Cereijido, M., Ed., CRC Press, Boca Raton, FL, 1992. With permission.)

joining the processes of the Sertoli cells surely creates a formidable stress on the zonules forcing them in loops and constraining the junctional membranes into gradually smaller surface areas. This stress, it is believed, could cause the junctions to proliferate and to produce redundant discontinuous fibrils. Stress has been reported to induce proliferation of junctional fibrils in a number of other systems (Huttner et al., 1982; Pitelka and Taggard, 1983; Lupu and Simionescu, 1985; Kachar and Pinto da Silva, 1991). The pattern of the fibrils within the Sertoli cell focal occluding junctions is mainly interwoven or anastomosed (Figure 28.3b), whereas in zonules the pattern is predominantly parallel (Figures 28.2a, 28.4, and 28.6a), a feature shared by oligodendrocytes in the central nervous system, the stria vascularis in the organ of Corti, the choroid plexus, and the collecting tubes of the kidney of certain species (Schnapp and Mugnaini, 1978). Gow et al. (1999) reported an interesting correlation

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between the expression of the transmembrane oligodendrocyte-specific protein claudin-11/(OSP) in the five locations and the presence of occluding junctions containing parallel arrays of junctional fibrils. Exactly what the role of claudin-11 is in conferring a typical parallel organization to the fibrils is still under investigation. In invertebrates, septate junctions are characteristically made up of parallel arrays of junctional fibrils (see Chapter 3 on invertebrates by Lane). It would be interesting to know whether some membrane particles in the junctional fibrils in birds contain claudin-11.

28.3.2 THE DIFFERENTIAL DISTRIBUTION PARTICLES WITHIN THE PLANE OF

OF THE JUNCTIONAL THE

CELLULAR MEMBRANE

Sertoli cell occluding junctions are made up of intramembranous particles that, during normal spermatogenesis, are typically associated preferentially with the E fracture face (Figures 28.2a and 28.4) (Gilula et al., 1976; Nagano and Suzuki, 1976; Pelletier and Friend, 1983) rather than with the P face as in the epididymis (Figure 28.7), vas deferens, and most other epithelia in the body. In addition, Sertoli cells do not undergo a change in the differential distribution of their junctional particles within the plane of the cell membrane following exposure to aldehyde fixatives (Nagano, 1980;

FIGURE 28.7 Freeze-fracture replica of an occluding zonule that surrounds the apex of two adjacent epididymal cells in the head of the epididymis of an adult mink obtained during the active spermatogenic phase. The strands of junctional particles (arrowhead) are typically associated preferentially with the ridges on the P-fracture face rather than with the grooves situated on the E face. Original magnification ×78,000.

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Nagano et al., 1982). These two characteristic features are also shared by occluding junctions in endothelial cells (Simionescu et al., 1975). In such seasonal breeders as the mink (Figure 28.6a) and mallard duck (Figure 28.6b), the differential distribution of particulate elements within the plane of the membrane changes during the annual reproductive cycle of the two species, and this results in junctional particles being associated preferentially with the P-fracture face (Figures 28.6a, b) during the seasonal inactive spermatogenic phase. However, the same change in the distribution of the intramembranous constituents in the two species is accompanied by junctional permeability changes in the mink (Pelletier, 1986; 1988) but not in the mallard duck (Pelletier, 1990). Species differences could account for different junctional behavior, but they would be expected to be governed by similar general principles. If the permeability characteristic of TJs would be dictated by a particular differential distribution of the junctional particles, both mink and duck occluding junctions would be expected to show similar junctional permeability changes unless the particulate junctional elements endowed with distinct physiological significance that move from one fracture face were not of the same chemical or physical nature in the two species.

28.4 PROTEINIC CONSTITUENTS OF THE SERTOLI CELL OCCLUDING JUNCTIONS 28.4.1 THE INTEGRAL MEMBRANE PROTEINS To date, integral membrane proteins of two distinct gene families have been localized in freeze-fracture replicated membranes to the particulate elements that compose the fibrils in the occluding junctions: occludin (Furuse et al., 1993; Fujimoto, 1995; Ando-Akatsuka et al., 1996; Saitou et al., 1997) and claudins 3, 4, 8, and 11 or OSP (Morita et al., 1999a, b). Given the apparent complexity of cell junctions and the diversity of their constituents, it is likely that junctional permeability changes would be dictated by a family of transmembrane proteins rather than by a single protein. Yet, the name given to the integral membrane proteins of these two gene families alludes to the notion that the proteins would seal or close the paracellular route between cells; hence the proposed terms occludin (to occlude or to close the intercellular cleft) and claudin, which instead should have been named “clauderin” from Latin claudere, which means to close. Claudine is generally used as a given name for a woman. Published reports suggest that integral proteins of each identified family constitute a small proportion of the entire population of intramembranous particles in the junctional fibrils (Furuse et al., 1993; Fujimoto, 1995; Ando-Akatsuka et al., 1996; Saitou et al., 1997; Morita et al., 1999a, b). Occludin (Byers et al., 1991) and claudin-11 (Moroi et al., 1998; Morita et al., 1999a) have both been localized to TJs in Sertoli cells. Occludin has been localized in the mouse testis and epididymis during embryonic and postnatal development by immunofluorescence confocal microscopy (Cyr et al., 1999). Contrary to the report of the absence of occludin in guinea pig Sertoli cells (Moroi et al., 1998), the integral protein has been detected in Sertoli cells of seven different species including guinea pig, but a positive correlation between the expression of occludin and the permeability status of the Sertoli

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cell occluding zonules could not be established (unpublished observation), thus casting doubt on the notion that occludin seals the paracellular route. Inactivation of both alleles of the occludin gene in mouse embryonic stem cells did not prevent the establishment of a network of normal junctional fibrils in occludin-deficient epithelial cells, suggesting the existence of yet-unidentified junction-specific proteins (Saitou et al., 1998). However, inhibition of claudin-11/OSP expression through homologous recombination in embryonic stem cells was reported to result in the absence of particulate elements in only the parallel fibrils of the Sertoli cell occluding junctions, a feature that was accompanied by sterility in male but not in female OSPnull mice (Gow et al., 1999). Although the consequences on the permeability of the Sertoli cell occluding zonules have not yet been reported in this experimental model, careful examination of the micrographs published in this report reveals that the grooves persisted in the zonules of the knockout male mice, suggesting that grooves and ridges may be regulated by families of junctional proteins other than occludin or claudin.

28.4.2 PERIPHERAL OCCLUDING-JUNCTION-ASSOCIATED PROTEINS AND THE SUBSURFACE OR CORTICAL ACTIN IN THE SERTOLI CELLS The Sertoli cell focal occluding junctions and occluding zonules units are typically made up of three constitutional elements: (1) the fusion sites joining adjacent plasma membranes, (2) the cortical actin next to the subsurface of the cells involved in the intercellular contact, and (3) the accompanying cisternae of rough endoplasmic reticulum. In the adult testis, the spatial organization or the state (filamentous vs. nonfilamentous) of the actin along the subsurface of Sertoli cells is typical. The distribution of the nonfilamentous monomeric (G-)actin follows the subsurface from the apex to the base of the Sertoli cells; however, the filamentous (F-)actin forms hexagonally packed bundles of filaments parallel to each other and to the cell surface next to the occluding junctions situated near the base and the apex of the seminiferous epithelium, but not next to those in the middle of the epithelium (Dym and Fawcett, 1970; Suárez-Quian and Dym, 1984; Vogl and Soucy, 1985; Pelletier et al., 1997). A correlation was reported between the presence of F-actin in the two locations of the epithelium and the expression of the α+ isoform of the TJ-associated protein ZO-1 (Pelletier et al., 1997), a member of the membrane-associated guanylase kinase homologue (MAGUK) family of proteins (Mitic and Anderson, 1998; Stevenson and Keon, 1998; Fanning et al., 1999). It has been shown with the immunogold technique at the electron microscope that ZO-1 localizes to Sertoli cell occluding junctions during the postnatal development and in the adult mouse testis (Byers et al., 1991). The published observation that ZO-1α– is expressed in structurally dynamic junctions and ZO-1α+ is expressed in less dynamic junctions has been suggested to reflect a differential linkage of the peripheral protein to the cytoskeleton (Balda and Anderson, 1993). The finding that ZO-1α– is predominant in adult testis where cyclic Sertoli cell occluding zonule assembly/disassembly continually takes place to accommodate the incessant translocation of germ cells into the lumenal compartment, whereas ZO-1α+ predominates in the neonatal testis, which contains

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chiefly focal occluding junctions that are not yet assembled into zonules, gives further support to this view (Pelletier et al., 1997). In addition, in the adult testis, the Sertoli cell occluding zonules are accompanied by ZO-1α+ and by F-actin near the base of the seminiferous epithelium. The focal occluding junctions are accompanied by ZO-1α– and by G-actin in the middle third of the epithelium (Pelletier et al., 1997). A differential linkage of the peripheral protein ZO-1 to the cell cytoskeleton might be dictated by different states (F/G) of the subsurface or cortical actin in the Sertoli cell, which in turn could be accompanied by distinct local junctional permeability status. Interestingly, Sertoli cell focal occluding junctions accompanied chiefly by ZO-1α– and G-actin were shown to be the only ones to resist cytochalasin D treatment (Russell et al., 1988; Weber et al., 1988), perhaps because they are more resilient than the steadier occluding zonule. The state of the junction-associated actin will likely influence the steadiness or resilience of the intercellular contact. The changes in the differential distribution of the junctional particles in the plane of Sertoli cell junctional membranes and in the permeability of the junctions that were reported to take place during the annual seasonal reproductive cycle of the mink and mallard duck were accompanied with changes in junction-associated filaments, particularly actin, suggesting a functional link between elements of the cytoskeleton and Sertoli cell junction function (Pelletier, 1986; 1988; 1990). Overexpression of the N-terminal domain of ZO-1 was reported to inhibit localization of Cx43 at cell–cell interfaces and to result in loss of electrical coupling between transfected cell pairs (Toyofuku et al., 1998). A number of peripheral proteins have been shown to be anchored to the membrane by cytoskeletal constituents such as α-spectrin (Bennett and Gilligan, 1993), ankyrin (Bennett, 1992), and 95/SAP90 (Gomperts, 1996). The actin-anchoring protein α-spectrin or fodrin (Bennett, 1990) was shown to be associated with ZO-1 (Itoh et al., 1997; Fanning et al., 1998).

28.5 MECHANISMS FOR THE ASSEMBLY AND DISASSEMBLY OF SERTOLI CELL OCCLUDING ZONULES 28.5.1 THE “ZIPPER” THEORY Although the modality of the process is still under investigation, it is widely agreed that the germ cells located in the basal compartment must migrate into the lumenal compartment to complete their meiosis and undergo cellular differentiation. The cyclic translocation of germ cells into the lumenal compartment is a well-orchestrated process that not only coincides with a precise moment during the development of the germ cells — the early zygotene spermatocyte stage in most species (Dym and Cavicchia, 1977; 1978; Pelletier, 1986; 1988) — but also takes place without significant leakage in the blood–testis barrier. In an attempt to explain this process, a few theories have been advanced. The “zipper” theory had proposed that occluding zonules between the processes of Sertoli cells that are positioned over the spermatogonia, leptotene, and early zygotene spermatocytes would “break down” while new occluding zonules would “re-form” between the Sertoli cell processes that

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extend under the migrating germ cells (Fawcett, 1975; Dym and Cavicchia, 1977; Russell and Peterson, 1985). Since no study has ever reported leakage of vascularly infused electron-opaque junction permeability tracers in the lumen even momentarily in normal animals, it is difficult to envision how Sertoli cell zonules would open without breakage of the seal between adjacent Sertoli cell plasma membranes to allow the passage of cohorts of germ cells through the intercellular space.

28.5.2 THE THEORY OF THE PRESENCE OF AN INTERMEDIATE CELLULAR COMPARTMENT WITHIN THE SEMINIFEROUS EPITHELIUM Another theory had proposed the existence of an “intermediate cellular compartment” that would be inhabited by migrating spermatocytes in transit between the basal and the lumenal compartments (Russell, 1977b). There are two obstacles to this theory. The evidence for such a compartment had been based on the inability of hypertonic solutions to induce shrinkage artifacts on adhering junctions, not on the competence of the Sertoli cell zonules to block entry into the epithelium of vascularly infused junction permeability tracers. In addition, freeze-fracture tracer (Pelletier and Friend, 1983) and electron-opaque studies (Cavicchia and Sacerdote, 1988) have consistently documented the existence of only one occluding zonule per Sertoli cell and thus provided no support to the existence of an intermediate compartment.

28.5.3 THE THEORY OF REPETITIVE REMOVAL OF MEMBRANE SEGMENTS FROM THE SERTOLI CELL JUNCTIONAL COMPLEX A third theory is proposed here. The theory proposes that the continuous upward displacement of germ cells constrains the Sertoli cell junctional membranes to proliferate into a series of loops that gradually lose their continuity with the membrane surface and create junctional vesicles that will be internalized (Diagram a). The process is accompanied by the slow translocation of spermatocytes edging into the intercellular cleft between Sertoli cells. Despite the progressive proliferation of the junctions, the occluding zonules remain situated in the basal third of the epithelium. The paracellular barrier does not leak because the seal of the intercellular space created by the occluding zonule between Sertoli cells does not open and close before and following the migration of spermatocytes as they have been proposed to do in the “zipper” theory. Instead, the stress imposed by the migration of the spermatocytes on the junctional fibrils within the occluding zonule causes them to proliferate into focal occluding junctions that fold into loops of short junctional membrane segments, which progressively lose their continuity with the Sertoli cell membrane surface and are continually removed and internalized as junctional vesicles to accommodate the transit of spermatocytes through the zonules from one cellular compartment of the epithelium to the next. Autophagic vacuoles containing the short junctional membrane segments have been reported to contain occluding, gap, and adhering junctions (Pelletier and Byers, 1992; Pelletier, 1998) and junctional proteins (Pelletier et al., 2000). The insidious progression of migrating spermatocytes within the Sertoli cell intercellular clefts occurs at a precise stage of the germ cell development, usually the zygotene stage, which lasts about 48 h in rodents

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and may vary depending on the animal species. The slowness of the germ cells edging into the intercellular cleft of Sertoli cells ensures the maintenance of a relatively close fit between adjoining plasma membranes and thus prevents excessive leakage of most bloodborne substances. Electron microscopy studies reveal that each germ cell (including the round spermatids but excluding the elongated ones) is lodged within a Sertoli cell intercellular cleft that is closed at both ends by tight, gap and adhering junctions (Diagram a) (Pelletier and Friend, 1983; Pelletier, 1986; 1988). Each generation of germ cells is secluded from the younger generation below by progressively fewer macular occluding junctions but by more gap and intermediate type of junctions. The association of the annular junctional profiles with autophagic vacuoles or with lysosomes is suggestive of two possibly distinct methods of junction disposal. Internalized junctions could either be permanently eliminated following association with autophagic vacuoles or, once processed by lysosomes, they could provide junctional elements that would be recycled for use in the assembly of new junctions. The process of removal of junctional membrane segments is repeated until all generations of germ cells (with the exception of the elongated spermatids) become lodged within a Sertoli cell intercellular cleft that is closed at both ends by diminutive occluding macules that have retained little of their initial occluding zonule barrier function. In other systems, acid phosphatase has been localized in annular profiles of internalized junctional vesicles (Larsen and Tung, 1978; Murray et al., 1981). Moreover, the internalization of surface cellular junctions and their fusion with lysosomes has been traced with the filipin-labeling technique (Risinger and Larsen, 1983).

28.6 THE OCCLUDING ZONULES DO NOT SERVE AS AN IMMUNOLOGICAL BARRIER TO ANTI-SPERM ANTIBODIES The germ cells of the testis contain autoantigens expressed at puberty. Since immunotolerance also develops around puberty, the autoantigens are tolerated in the adult provided they remain in situ. If they are introduced at another location in the body, they generate an autoimmune reaction (Mahi-Brown et al., 1988). In the normal adult mink, the blood–testis barrier was reported to be transiently leaky during the annual and seasonal reproductive cycle of the animal and to expose momentarily germ cells that contained intracellular and membrane surface autoantigens to antigen-presenting cells or lymphocytes from the blood (Pelletier, 1986; 1988). Yet, this momentary seasonal increase in the permeability of the barrier was not accompanied with an infiltration of monocytes, macrophages, or lymphocytes in the seminiferous epithelium of most mink (Pelletier, 1986; 1988). However, the transient permeability increase had been reported to coincide with an increase of 85% antisperm antibodies in the sera of both normal mink and mink that had developed a spontaneous autoimmune orchitis (Tung et al., 1984). These results have been challenged by recent measurements of antisperm antibody serum levels performed monthly during a period of 1 year by immunofluorescence microscopy and ELISA. The measurements revealed that a change in the competence of the blood–testis barrier was accompanied

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with a seven- to tenfold increase in antisperm antibody serum levels in the orchitic mink but not in the normal mink (Yoon et al., 2000). In addition, the measurements revealed that high serum levels of antisperm antibodies coincided with the periods of spermatogenic activity but not with the periods of transient permeability in the Sertoli cell zonules in the normal mink (Yoon et al., 2000). This observation suggests that the Sertoli cell occluding zonules forming the blood–testis barrier constitute an incomplete barrier to the autoantigens from the rest of the body. Additional support to the notion that the segregation of testicular antigens by the barrier is not complete is provided by the report that germ cells in the basal compartment of the seminiferous epithelium and the lamina propria of the tubules bind antibodies from anti-testis in vivo after passive transfer of the sera by intraperitoneal injections in the rat (Saari et al., 1996). Together, these results emphasize the fact that, contrary to what is generally assumed, the body is normally exposed to relatively high levels of antisperm antibodies during active spermatogenic activity without triggering an autoimmune response, because the Sertoli cell occluding zonules do not constitute an immunological barrier. The fact that most mink do not develop a spontaneous autoimmune orchitis in the face of relatively high levels of antisperm antibodies indicates that the development and maintenance of the immunotolerance in the testis requires the presence of immunosuppressive factors that prohibit a lymphocytic infiltration during periods of transient permeability increase in the blood–testis barrier and prevent the establishment of anti-germ cell autoimmune diseases. Recently identified autoantigens expressed after the manifestation of immunotolerance to self-antigens have been localized in the basal compartment situated below the blood–testis barrier and they have been shown to be accessible to circulating autoantibodies (Yule et al., 1987; 1988; Mahi-Brown et al., 1988). Clones of T and B lymphocytes that react with these antigens and that are potentially involved in the autoimmune response have been detected in normal animals (Mahi-Brown et al., 1988). The T lymphocytes have been documented to express high levels of CD95 (Fas or Apol) antigen receptors (Brunner et al., 1995; Ju et al., 1995). Activation of Fas antigen receptor by the Fas ligand (Suda et al., 1993) or by an antiFas antibody (Itoh et al., 1991) has been shown to induce apoptosis in lymphocytes and thymocytes (Ogasawara et al., 1993). The ability to express functional Fas is lacking in lpr mice, which cannot eliminate activated lymphocytes and eventually develop autoimmune diseases (Luciani and Golstein, 1994). Rodent testis is a major site of expression of the Fas ligand (Suda et al., 1993) and Fas ligand mRNA is constitutively expressed by Sertoli cells (Bellgrau et al., 1995). It was hypothesized that Sertoli cell expression of Fas ligand could not only account for the immuneprivileged sites it contributes when the cells are transplanted into the rodent kidney (Bellgrau et al., 1995), but also that it could be one of the factors that prevents an immune response by limiting access of antigen-presenting cells and lymphocytes to the intratesticular antigens when the blood–testis barrier becomes momentarily incompetent. The spontaneous development of autoimmune orchitis in only some of the mink may reflect the inherited incapacity for Sertoli cells and other testicular cells to produce immunosuppressors responsible for the regulation of the immune response rather than a defect in only the Sertoli cell zonule integrity and function.

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28.7 THE OCCLUDING JUNCTIONS IN THE EPIDIDYMIS AND VAS DEFERENS The first demonstration of the existence of a blood–tissue barrier in the testis by intravenous injections of dyes dates back to the turn of the century (Ribbert, 1904; Bouffard, 1906). By comparison, intercellular junctions in the epididymis and deferent duct have received much less attention. In electron microscopy of thin sections, the topography and morphological features of occluding junctions in the epididymis and deferent duct are similar to the ones in most epithelia in the body including the stomach and intestine. Junctional fibrils have been identified in freeze-fracture electron microscopy at the contact sites joining the apex of neighboring epididymal cells in adult guinea pig (Friend and Gilula, 1972; Greenberg and Forssmann, 1983). Contrary to the Sertoli cells, the occluding junction particles in epididymal cells are typically associated preferentially with the P-fracture face as is the case in most epithelia of the body (Figure 28.7). Fibrils surrounding the juxtalumenal cells of the epididymis and forming continuous occluding zonules that constitute the anatomical basis for a blood–epididymis barrier have been reported to be already present in the 12-day mouse embryo (Nagano and Suzuki, 1978). Occludin has been localized by immunofluorescence confocal microscopy in embryonic (days 13.5) mouse epididymis (Cyr et al., 1999). Because the number of fibrils was reported to decrease from the epididymis to the vas deferens in adult rat, it had been hypothesized that this feature could correlate with a decrease in the junctional permeability along the testicular excurrent ducts (Suzuki and Nagano, 1978a). The answer to this question varies significantly according to which junction permeability tracer is used to test the hypothesis (Figures 28.8a, b, and c). For example, tracer studies performed with lanthanum nitrate used as the junctional permeability probe concluded that the gradual development of a blood–epididymis barrier coincided with the establishment of a blood–tissue barrier in the testis around puberty (Hoffer and Hinton, 1984; Agarwal and Hoffer, 1989) and that permeability characteristics are segment specific in adult rat epididymis (Suzuki and Nagano, 1978b; Levy and Robaire, 1999; Levy et al., 1999). Conversely, tracer studies with vascular infusion of horseradish peroxidase (HRP) used as the permeability probe contradict these findings and clearly demonstrate that the blood–epididymis barrier is competent long before the blood–testis barrier and that there exists no correlation between the number or the patterns of fibrils at the site of epididymal cell occluding junctions viewed in replicated membranes and the permeability characteristics of these junctions (Pelletier, 1994; 1995a). Figures 28.8a and b show that, when lanthanum nitrate is added to aldehydes used to preserve-fix tissues and to test junction permeability, deposits of lanthanum are inconsistently associated with cell membranes and intracellular vesicles even in the presence of impermeable junctions. Because the lanthanum nitrate technique requires that the probe be added to all solutions up to dehydration to prevent the compound from being washed out from the tissues tested when exposed to solutions of pH lower than 7.8 (Revel and Karnovsky, 1967), many reports have cautioned about inconsistent staining of the extracellular space in the presence of impermeable junctions (Wolff and Schreiner, 1968; Mahrle, 1977; Pelletier, 1994). As shown in Figure 28.8b, the lanthanum nitrate tracer technique can

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FIGURE 28.8 (a) Thin section of the head of an epididymis harvested from an adult mink during the seasonal annual testicular regression. The entry of lanthanum is stopped (arrow) by an occluding zonule situated near the lumen (L) of the excurrent duct. This particular micrograph shows no lanthanum deposit associated with the cell membrane of microvilli (mv) outlining the lumen. Original magnification ×50,000. (b) Head of the epididymis harvested from an adult mink during the seasonal annual testicular regression. The arrows indicate the blocking of lanthanum by occluding zonules situated near the apex of the epididymal cells. However, deposits of lanthanum are apparent on the cell plasma membranes, on the microvilli (mv), and on the cell membranes of intracellular vesicles (v). Original magnification ×50,000. (c) Note that when vascularly infused HRP is used as a permeability tracer in place of the lanthanum, the blocking of the junctional probe through the extracellular space is clear-cut and there are no deposits on the microvilli (mv). Original magnification ×50,000.

yield false-positive results. Because the lanthanum forms a colloidal hydroxide at alkaline pH, the compound could serve as a useful marker of the extracellular space. Conversely, its presence on cell membranes may also reflect the mordanting effect on the delineation of cell membranes due to special affinities of lanthanum for particular membrane constituents. However, when HRP is vascularly infused to test the junction permeability of a tissue that is later fixed with aldehydes to follow the path of the probe, the passage or blockage of the tracer is clear-cut, and no tracer is found in the extracellular space when junctions are impermeable to the permeability probe (Figure 28.8c). Electron-opaque tracers (Pelletier, 1994) and freeze-fractured membrane replication (Pelletier, 1995a) studies showed that a lumen which had developed in utero

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in the epididymis and vas deferens was present at birth long before the formation of a lumen in the seminiferous tubules of the testis around puberty. In addition, these studies concluded that the assembly of occluding junctions into occluding zonules coincidentally with the development of a lumen in the epididymis and vas deferens resulted in the establishment of a paracellular barrier that was impermeable in the testicular excurrent ducts before than in the testis (Figure 28.8c). Furthermore, these studies showed that not only were the blood barriers in the excurrent ducts and in the testis established at different periods of development but, in the adult, the two barriers acted asynchronously. For example, in mink, there are two time intervals when the blood–testis barrier is permeable: (1) during the neonatal period and (2) during the annual seasonal testicular regression in the adult. Yet, during these two periods, although the blood–testis barrier is permeable to extracellular tracers, the blood barrier in the excurrent ducts remains impermeable (Pelletier, 1994; 1995a). In guinea pig, a continual breeder, experimental cryptorchidism in the adult results in severe damage to the testis and azoospermia; yet, a lumen persists in the excurrent duct and the blood–tissue barrier remains competent (Pelletier, 1993). These findings show that in the epididymis, just as in the testis, the presence of a lumen in the tubules dictates the impermeability of the occluding zonules, not the presence or absence of germ cells.

ACKNOWLEDGMENT The author thanks Dr. M. L. Vitale for the critical reading of the manuscript. Work performed in the author’s laboratory was supported by grants from MRC and NSERC of Canada.

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Pinto da Silva, P. and Kachar, B. 1982. On tight junction structure. Cell, 28: 441–450. Pitelka, R. D. and Taggard, B. N. 1983. Mechanical tension induces lateral movement of intramembrane components of the tight junction: study on mouse mammary cells in culture. J. Cell Biol., 96: 613–624. Pricam, C., Humbert, F., Perrelet, A., and Orci, L. 1974. A freeze-etch study of the tight junctions of the rat kidney tubules. Lab. Invest., 30: 286–291. Reinhart, M. P. 1990. Intracellular sterol trafficking. Experiencia, 46: 599–609. Revel, J.-P. and Karnovsky, M. J. 1967. Hexagonal array of subunits in intercellular junctions of the mouse heart and liver. J. Cell Biol., 33: C7–C12. Ribbert, H. 1904. Die Abscheidung intravenös injuzierten gelösten Karmens in den Geweben. Z. Allg. Physiol., 4: 201–214. Risinger, M. A. and Larsen, W. J. 1983. Interaction of filipin with junctional membranes at different stages of the junction’s life history. Tissue Cell, 15: 1–15. Risley, M. S., Tan, I. P., Roy, C., and Sáez, J. C. 1992. Cell- and stage-dependent distribution of connexin 43 gap junctions testes. J. Cell Sci., 103, 81–96. Rodriguez-Boulan, E. and Nelson, W. J. 1989. Morphogenesis of the polarized epithelial cell phenotype. Science, 245: 718–725. Russell, L. D. 1977a. Desmosome-like junctions between Sertoli and germ cells in the rat testis. Am. J. Anat., 148, 301–312. Russell, L. D. 1977b. Movement of spermatocytes from the basal to the adlumenal compartment of the rat testis. Am. J. Anat., 148: 301–312. Russell, L. D. 1978. The blood–testis barrier and its formation relative to spermatocyte maturation in the adult rat: a lanthanum tracer study. Anat. Rec., 190: 99–112. Russell, L. D. and Peterson, R. N. 1985. Sertoli cell junctions: morphological and functional correlates. Int. J. Cytol., 94: 177–211. Russell, L. D., Goh, J. C., Rashed, R. M. A., and Vogl, A. W. 1988. The consequences of actin disruption at Sertoli ectoplasmic specialization sites facing spermatids after in vivo exposure of rat testis to cytochalasin D. Biol. Reprod., 39: 105–118. Saari, T., Jahnukainen, K., and Pöllänen, P. 1996. Autoantigenicity of the basal compartment of seminiferous tubules in the rat. J. Reprod. Immunol., 31: 65–79. Saitou, M., Ando-Akatsuka, Y., Itoh, M., Furuse, M., Inazawa, J., Fujimoto, K., and Tsukita, S. 1997. Mammalian occludin in epithelial cells: its expression and subcellular distribution. Eur. J. Cell Biol., 63: 222–232. Saitou, M., Fujimoto, T., Doi, Y., Itoh, M., Fujimoto, K., Furuse, M., Takano, H., Noda, T., and Tsukita, S. 1998. Occludin-deficient embryonic stem cells can differentiate into polarized epithelial cells bearing tight junctions. J. Cell Biol., 141: 397–408. Schnapp, B. and Mugnaini, E. 1978. Membrane architecture of myelinated fibers as seen by freeze fracture, in Physiology and Pathobiology of Axons, Waxman, S. G., Ed., Raven Press, New York, 83–123. Schneeberger, E. E. and Lynch, R. D. 1992. Structure, function, and regulation of cellular tight junctions. Am. J. Physiol., 262: L647–L661. Setchell, B. P., Voglmayr, J. K., and Waites, G. M. H. 1969. A blood–testis barrier restricting passage from the blood into rete testis fluid but not into lymph. J. Physiol., 200: 73–85. Simionescu, M., Simionescu, N., and Palade, G. E. 1975. Segmental differentiation of cell junctions in the vascular endothelium. The microvasculature. J. Cell Biol., 67: 863–885. Staehelin, L. A. 1973. Further observations on the fine structure of freeze cleaved tight junctions. J. Cell Sci., 13: 763–786. Staehelin, L. A. 1974. Structure and function of intercellular junctions. Int. Rev. Cytol., 39: 191–283.

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Stevenson, B. R. and Keon, B. H. 1998. The tight junction: morphology to molecules. Annu. Rev. Cell Biol., 14: 89–109. Suárez-Quian, C. A. and Dym, M. 1984. Further observations on the microfilaments bundles of Sertoli cell junctional complexes. Ann. N.Y. Acad. Sci., 438: 476–480. Suda, T., Takahashi, T., Golstein, P., and Nagata, S. 1993. Molecular cloning and expression of the Fas ligand, a novel member of the tumor necrosis factor family. Cell, 75: 1169–1178. Suzuki, F. and Nagano, T. 1978a. Regional differentiation of cell junctions in the excurrent duct epithelium of the rat testis as revealed by freeze-fracture. Anat. Rec., 191: 503–519. Suzuki, F. and Nagano, T. 1978b. Development of tight junctions in the caput epididymal epithelium of the mouse. Dev. Biol., 63: 321–334. Szöllösi, A. and Marcaillou, C. 1980. Gap junctions between germ and somatic cells in the testis of the moth Anagasta kuehniella, Insecta: Lepidoptera. Cell Tissue Res., 213: 137–148. Tan, I. P., Roy, C., Sáez, J. C., Sáez, C. G., Paul, D. L., and Risley, M. S. 1996. Regulated assembly of connexin 33 and connexin 43 into rat Sertoli cell gap junctions. Biol. Reprod., 54, 1300–1310. Toyofuku, T., Yabuki, M., Otsu, K., Hori, M., and Tada, M. 1998. Direct association of the gap junction protein connexin-43 with ZO-1 in cardiac myocytes. J. Biol. Chem., 273: 12725–12731. Tung, K. S. K., Ellis, L. C., Childs, G. V., and Duffau, M. 1984. The dark mink: a model of male infertility. Endocrinology, 114: 922–929. van Deurs, B. and Koehler, J. K. 1979. Tight junctions in the choroid plexus epithelium. A freeze-fracture study including complementary replicas. J. Cell Biol., 80: 662–673. Varanda, W. A. and de Carvalho, A. C. 1994. Intercellular communication between mouse Leydig cells. Am. J. Physiol., 267: C563–C569. Vitale, R., Fawcett, D. W., and Dym, M. 1973. The normal development of the blood–testis barrier and the effect of clomiphene and estrogen treatment. Anat. Rec., 176: 331–344. Vogl, A. W. and Soucy, L. J. 1985. Arrangement and possible function of actin filament bundles in ectoplasmic specializations of ground squirrel Sertoli cells. J. Cell Biol., 100: 814–825. Weber, J. E., Turner, T. T., Tung, K. S. K., and Russell, L. D. 1988. Effects of cytochalasin D on the integrity of the Sertoli cell (blood–testis) barrier. Am. J. Anat., 182: 130–147. Wolff, K. and Schreiner, E. 1968. An electron microscopic study on the extraneous coat of keratinocytes and the intercellular space of the epidermis. J. Invest. Dermatol., 51: 418–430. Yap, A. S., Mullin, J. M., and Stevenson, B. R. 1998. Molecular analysis of tight junctions physiology: insight and paradoxes. J. Membr. Biol., 163: 159–167. Yoon, S. R., Vitale, M. L., and Pelletier, R.-M. 2001. Modulation of the fas system during the annual reproductive cycle in normal mink and during spontaneous autoimmune orchitis (submitted). Yule, T. D., Shepherd, K., Russell, L. D., and Tung, K. S. K. 1987. Murine testis autoantigens are not immunologically sequestered. Ann. N.Y. Acad. Sci., 513: 523–525. Yule, T. D., Montoya, G. D., Russell, L. D., Williams, T. M., and Tung, K. S. K. 1988. Autoantigenic germ cells exist outside the blood–testis barrier. J. Immunol., 141: 1161–1167.

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Relationship between Tight Junctions and Leukocyte Transmigration Alan R. Burns, David C. Walker, and C. Wayne Smith

CONTENTS 29.1 Introduction ................................................................................................629 29.2 The Multistep-Adhesion Cascade ..............................................................630 29.3 Interendothelial Clefts................................................................................631 29.4 Neutrophil Migration through Interendothelial Clefts ..............................633 29.5 Neutrophil Transmigration at Endothelial Tricellular Corners .................635 29.6 Neutrophil Adhesion to Endothelial Borders ............................................639 29.7 Transcytotic Neutrophil Migration ............................................................641 29.8 Neutrophil Transepithelial Migration and Tricellular Corners..................642 29.9 Mechanisms for Opening/Remodeling Tight Junctions ............................644 29.10 Concluding Remarks..................................................................................646 Acknowledgments..................................................................................................647 References..............................................................................................................647

29.1 INTRODUCTION An early step in the nonadaptive immune response is the accumulation of leukocytes (primarily neutrophils) at sites of inflammation. Leukocytes leave the free-flowing bloodstream by adhering to and migrating across the inflamed endothelium. In some cases (e.g., mucosal inflammation), leukocytes that enter the interstitium move toward and migrate across an inflamed epithelium. Since the majority of published studies on leukocyte transmigration focus on neutrophil emigration, this chapter will concentrate on the neutrophil. Transendothelial migration will be emphasized because of recent and important advances in the understanding of the molecular mechanisms regulating neutrophil extravasation and the role tight junctions (TJs) play in the migratory process.

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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FIGURE 29.1 Multistep-adhesion cascade and preferred pathway for neutrophil migration across an inflamed endothelium. Rolling neutrophils are captured on endothelial borders (1) after which they become activated and arrest within 5 µm of a tricellular corner (2), the site where the margins of three endothelial cells converge. Neutrophils migrate preferentially at tricellular corners by passing through TJ discontinuities (3).

29.2 THE MULTISTEP-ADHESION CASCADE In preparation for emigration, free-flowing leukocytes leave the bloodstream by adhering to the inflamed endothelium (Figure 29.1). Four classes of adhesion molecules apparently control leukocyte adhesion. They are selectins, carbohydrate ligands for the selectins, integrins, and members of the immunoglobulin superfamily. The structure and functions of these molecules have been extensively reviewed (Smith, 1993; Springer, 1994), and it is now clear that the processes of adhesion most often occur as a cascade of events driven by shear forces and signaling pathways. The three members of the selectin family constitutively express a lectin domain that interacts with carbohydrate ligands (glycoproteins or glycolipids) (Bevilacqua et al., 1991). The principal mechanism of control of this adhesive pathway resides in increases or decreases in the expression levels of the selectins or their ligands on cell surfaces. L-selectin (CD62L) is constitutively expressed on leukocytes and serves to tether leukocytes to activated endothelial cells. E-selectin (CD62E) and P-selectin (CD62P) are expressed on the surface of endothelial cells following distinct stimulation. E-selectin is newly synthesized following stimulation with cytokines such as interleukin-1β (IL-1β) or tumor necrosis factor-α (TNFα), and P-selectin is constitutively packaged in Weibel–Palade bodies and rapidly mobilized to the endothelial surface following stimulation with factors such as histamine, thrombin, and LTC4. The principal function of selectins is the primary capture of flowing leukocytes, as evidenced by the rolling of leukocytes along the endothelial surface (Jones et al., 1993). At physiological shear rates, selectins do not stop rolling leukocytes, but slow them sufficiently that two other classes of adhesion molecules can come into play. Two groups of integrins (β1 and β2) have been shown to function as secondary adhesive mechanisms interacting with members of the Ig superfamily expressed on endothelial cells following stimulation with cytokines (e.g., IL-1β). Interactions between these integrins and their ligands fail to lead to adhesion unless the leukocytes

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have been stimulated. In contrast to the selectins, control of these adhesive pathways resides not only in surface expression levels, but in avidity changes in the integrins following leukocyte activation. Even when stimulation results in high avidity of the integrins, adhesion occurs very inefficiently at physiological shear rates. Stationary adhesion under flow requires at least two adhesive steps, rolling adhesion to slow the velocity of interactions and integrin-dependent adhesion to arrest rolling cells. Stimulation of both leukocytes and endothelial cells is required for this sequence of adhesion to occur, increasing surface levels of some molecules and changing the avidity of others. IL-1β and TNFα stimulate a host of changes at the endothelium including expression of E-selectin, intercellular adhesion molecule-1 (ICAM-1, CD54), the principal ligand for the β2-integrins, LFA-1 (CD11a/CD18) and Mac-1 (CD11b/CD18), and vascular cell adhesion molecule-1 (VCAM-1, CD106, a principal ligand for the β1-integrin, VLA4 (CD49d/CD29)). Although ICAM-2 (CD102) expression is constitutive and not upregulated by cytokine treatment, it is a ligand for LFA-1. Stimulated endothelial cells produce neutrophil-activating factors (e.g., IL-8 or platelet activating factor, PAF), which stimulate avidity changes in the β1and β2-integrins and cytoskeletal activity. This is not only necessary for stationary adhesion of leukocytes to endothelial cells under conditions of flow, but for the subsequent migration of leukocytes into the extravascular space (Springer, 1994).

29.3 INTERENDOTHELIAL CLEFTS It is generally accepted that neutrophils migrate across the endothelium by penetrating the junctions that lie within the interendothelial clefts. This paradigm is based largely on the early (1960s) electron microscopic observations of Marchesi and Florey, who reported that intercellular junctions are the main sites through which neutrophils (and other leukocytes) pass from blood to tissue (Marchesi and Florey, 1960; Marchesi, 1961). Molecular evidence supporting the importance of interendothelial clefts as primary routes for leukocyte migration stems from findings that platelet–endothelial cell adhesion molecule-1 (PECAM-1) and junctional adhesion molecule (JAM) are localized to interendothelial clefts (Muller et al., 1989; Ayalon et al., 1994; Martin-Padura et al., 1998) and blocking antibodies against PECAM-1 and JAM reportedly inhibit leukocyte transendothelial migration (Muller et al., 1993; Muller, 1995; Martin-Padura et al., 1998; Del Maschio et al., 1999). Three distinct junctional complexes are found within interendothelial clefts and they are gap junctions, TJs, and adherens junctions (Dejana et al., 1995; Dejana and Del Maschio, 1995). Gap junctions are unlikely to pose a barrier to neutrophil transmigration since they occur as discrete, well-spaced plaques rather than beltlike zones within the cleft. Conversely, the beltlike design of TJs (zonula occludens) and adherens junctions (zonula adherens) has been taken as evidence that these junctions are barriers to neutrophil emigration. Even though the TJ is the most apical element of the cleft and the first structural barrier that the migrating neutrophil encounters, most studies have focused on the role of adherens junctions in neutrophil transendothelial migration (Dejana et al., 1995; Dejana and Del Maschio, 1995; Del Maschio et al., 1996; Allport et al., 1997; Bianchi et al., 1997; Gotsch et al., 1997; Moll et al., 1998).

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A factor contributing to why endothelial TJs are seldom considered to play a role in neutrophil emigration stems from the fact that neutrophil emigration occurs largely in postcapillary and collecting venules, and these vessels are considered to have poorly developed and discontinuous TJs (Bowman et al., 1992; Schnittler, 1998). But this should not be taken to imply that venular TJs are freely permeable to migrating leukocytes. On the contrary, although ultrastructural studies suggest that ~30% of venular endothelial TJs are open, the size of each pore or gap is only 30 to 60 Å (Simionescu et al., 1978a, b) and this is ~200 to 300 times smaller than the 1 to 2 µm migration pore typically observed for transmigrating neutrophils (Marchesi and Florey, 1960; Feng et al., 1998; Burns et al., 2000). Moreover, it must be kept in mind that neutrophil emigration occurs in other vessel types as well. In lung, alveolar capillaries are a principal site for neutrophil emigration and these vessels have well-developed and continuous TJs (Walker et al., 1988; Schneeberger and Lynch, 1992). At the molecular level endothelial TJs are comprised of at least two different transmembrane proteins, claudin (Furuse et al., 1998a) and occludin (Furuse et al., 1993). Claudins belong to a multigene family composed of at least 18 members (claudin-1 through claudin-18). In fibroblasts that normally lack TJs, transfection with claudin-1 or claudin-2 results in the formation of elaborate, netlike TJ strand complexes. Fibroblasts transfected with occludin alone express poorly organized TJs comprised of short strands. Claudin-1 and claudin-2 appear to be largely responsible for TJ formation, and occludin functions as an accessory molecule (McCarthy et al., 1996; Furuse et al., 1998a, b). Recent evidence suggests that claudin-5 is restricted to endothelial cells and incorporation of claudin-5 cDNA into fibroblasts results in “endothelial-like” TJs, as determined by freeze-fracture electron microscopy (Morita et al., 1999). Viewed by thin-section transmission electron microscopy, TJs appear as points of membrane fusion that presumably occur as a result of claudin–claudin and occludin–occludin interactions on adjacent cells (Furuse et al., 1993; Anderson and Van Itallie, 1995; Tsukita et al., 1996; 1999). The barrier properties (e.g., permeability to macromolecules and cells) of TJs appear to be regulated through claudin and occludin since both are linked to the cytoskeleton via a number of cytoplasmic proteins (ZO-1, ZO-2, and ZO-3) (Furuse et al., 1994; Yamamoto et al., 1997; Fanning et al., 1998; Haskins et al., 1998; Itoh et al., 1999; Tsukita et al., 1999; Wittchen et al., 1999). Experimental evidence for the importance of the C-terminal cytoplasmic domain of occludin playing a role in regulating paracellular flux is suggested by studies performed on an epithelial cell line (MDCK strain 2). Balda and colleagues (1996) showed that paracellular flux increased severalfold when MDCK cells were transfected with COOH-terminally truncated chicken occludin. Moreover, expression of COOH-terminally truncated occludin disturbed the intramembrane fence that restricts diffusion of lipids between apical and basolateral cell surface domains. Interestingly, the ultrastructural morphology of the TJs, as revealed by thin-section and freeze-fracture electron microscopy, was not altered. This latter observation supports the idea that TJs are composed of more than one structural membrane component (e.g., claudin). Adherens junctions are situated beneath the TJs and are the result of adhesive interactions between transmembrane proteins known as cadherins, which promote

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homophilic, calcium-dependent cell–cell interaction. Cadherins redistribute to the intercellular clefts when endothelial cells come in contact with each other. Although a number of cadherin subtypes (N-cadherin, E-cadherin, P-cadherin, and VE-cadherin) have been found in endothelial cells, VE-cadherin appears to play the most important role in adherens junction formation. It is found exclusively in endothelial cells and its cytoplasmic domain is linked to the cytoskeleton via a group of cytoplasmic proteins known as catenins (α-, β, and γ-catenin or plakoglobin). Adherens junction formation appears to be required for and precedes the formation of endothelial TJs (Dejana et al., 1995; Dejana and Del Maschio, 1995). PECAM-1 is a transmembrane protein and a member of the immunoglobulin supergene family. Its extracellular region is organized into six globular domains, and it engages in homophilic interactions as well as heterophilic interactions with glycosaminoglycans (GAGs) (Dejana et al., 1995). Since PECAM-1 is also found on the leukocyte cell surface, it has been suggested that homophilic interactions between leukocyte PECAM-1 and endothelial PECAM-1 may occur during transendothelial migration. This “zipper” hypothesis proposes that endothelial permeability is preserved during leukocyte migration because as the leading edge of the neutrophil (pseudopod) unzips the interendothelial junctions, the junctional contacts are replaced by endothelial PECAM-1 interactions with neutrophil PECAM-1 (Stein et al., 1997). However, studies in rats using a blocking anti-PECAM-1 antibody (Wakelin et al., 1996) or in mutant mice with a targeted deletion in PECAM-1 (Duncan et al., 1999) suggest that when PECAM-1 is blocked or absent, neutrophil transendothelial migration in response to an inflammatory stimulus is normal. Conversely, subsequent migration through the endothelial basal lamina is delayed. Although these data are consistent with a role for PECAM-1 in leukocyte extravasation, they favor a role for PECAM-1 in leukocyte migration through subendothelial extracellular matrices rather than a role in migration through interendothelial clefts. JAM, like PECAM-1, is a transmembrane protein and a member of the Ig superfamily. Its expression is confined to epithelial and endothelial cells and, unlike PECAM-1, which is primarily localized to the basolateral aspect of interendothelial clefts (Ayalon et al., 1994), immunogold electron microscopy suggests JAM has a more apical position, codistributing with TJs (Martin-Padura et al., 1998). An antiJAM antibody, BV11, inhibits monocyte emigration in a mouse model of skin inflammation (Martin-Padura et al., 1998) and monocyte and neutrophil emigration in a mouse model of experimental meningitis (Del Maschio et al., 1999). BV11 also inhibits human monocyte migration across mouse endothelial cells in vitro (MartinPadura et al., 1998). Whether JAM plays a role in human neutrophil transendothelial migration across human endothelial cells is unknown.

29.4 NEUTROPHIL MIGRATION THROUGH INTERENDOTHELIAL CLEFTS TJs have been described as forming circumferential beltlike regions of intimate contact between adjacent endothelial cells (Staehelin, 1974; Anderson and Van Itallie, 1995). Despite this potentially formidable barrier, the time required for a

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firmly adherent neutrophil to migrate across an activated endothelium is typically less than 2 min (Burns et al., 1997; Stein et al., 1997). Since neutrophils can transmigrate quickly, it has been suggested, but never proven, that endothelial TJs must open and close quickly during the migration process (Cramer, 1992). Alternatively, the idea that neutrophil migration between endothelial cells might involve proteolytic degradation of junctional proteins has been a topic of considerable interest and debate. Differences in experimental design likely account for the lack of agreement on whether neutrophil adhesion and transmigration alters the endothelial barrier function. For example, the ratio of neutrophils to endothelial cells appears to be critical as does the duration of the transmigration assay. Huang and colleagues (1988; 1993) found that at low ratios of neutrophil to endothelial cells (<5:1), albumin flux (permeability) and transendothelial electrical resistance (TER) were unaffected by neutrophil transmigration. While higher (25 to 50:1) ratios were associated with decreased TER, these ratios were deemed excessive and nonphysiological (Huang et al., 1988; 1993). Using a 10:1 ratio of neutrophils to endothelial cells, Del Maschio and colleagues (1996) reported a fourfold increase in endothelial permeability measured by horseradish peroxidase (HRP) penetration at a single time point, 60 min after the addition of neutrophils. It is not clear that this increased permeability is associated with the migration process since neutrophil migration after contact with an activated endothelial monolayer is typically complete by 10 min (Huang et al., 1988; Burns et al., 1997; Stein et al., 1997). Using real-time measurements and a ratio of neutrophils to endothelial cells of 2:1, the authors are unable to demonstrate a drop in TER during a 10-min period when neutrophils were adhering to and migrating across IL-1-treated human umbilical vein endothelial cell (HUVEC) monolayers (Burns et al., 2000). It is worth noting that since TER and paracellular permeability can vary independently (Balda et al., 1996), deciding whether TJs are degraded or breached during neutrophil transmigration requires the use of additional techniques (see below). Although several published reports show that endothelial adherens junctions undergo proteolytic degradation as a result of CD18-dependent neutrophil adhesion (Del Maschio et al., 1996; Allport et al., 1997), a subsequent study clearly demonstrates that this is not the case. Moll and colleagues (1998) found that the apparent degradation of the adherens junction protein β-catenin was artifactual and due to nonphysiological release of neutrophil proteases during sample preparation for Western blots and immunofluorescence microscopy. The authors reached a similar conclusion for TJ proteins occludin, ZO-1, and ZO-2. Although these proteins appeared to be degraded during neutrophil transendothelial migration, careful preparation of the endothelial samples for immunofluorescence microscopy and Western blot analysis showed that this degradation was artifactual (Burns et al., 2000). Moreover, using thin-section and freeze-fracture electron microscopy, TJs appear intact during and immediately following neutrophil transendothelial migration. In summary, the evidence is clear that following firm adhesion of the neutrophil to an activated endothelium, transmigration can occur rapidly (within 10 min) and need not involve widespread degradation of junctional complexes (zonula occludens and zonula adherens) or loss of endothelial barrier properties. It is still possible that neutrophil-mediated degradation of intercellular junctions is focal and occurs only

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at the site of penetration. However, in situations where the barrier properties of the monolayer are not compromised (Huang et al., 1988; 1993; Burns et al., 2000), it is difficult to understand how a focally degraded (i.e., damaged) junction reseals once the neutrophil completes its journey across the endothelium.

29.5 NEUTROPHIL TRANSMIGRATION AT ENDOTHELIAL TRICELLULAR CORNERS Another explanation for why barrier properties of the endothelium are preserved during neutrophil migration may be that the junctions remain intact. Although TJs are generally regarded as forming continuous belts around epithelial and endothelial cells (Staehelin, 1974; Anderson and Van Itallie, 1995), electron microscopic freeze-fracture images of pulmonary capillaries (Walker et al., 1994) and tracheal epithelial cells (Walker et al., 1985) show that TJs are discontinuous at tricellular corners where the borders of three cells converge (Figure 29.2). Similar discontinuities have been noted for TJs and adherens junctions in rabbit corneal endothelial cells (Barry et al., 1995). Walker and colleagues (1994) hypothesized that tricellular corners were “potential sites for the transient opening and closing of the paracellular pathway … [and] possible avenues through which white blood cells migrate during inflammatory reactions.” Using cytokine-activated HUVEC monolayers, a commonly used in vitro model of leukocyte trafficking in vivo, the authors wished to determine whether tricellular corners are preferred avenues for transmigration. It was found, in concert with previous investigator, that cultured HUVEC monolayers rarely form TJs in vitro. In vivo, endothelial TJs are located in the most apical aspect of the intercellular cleft. Hence, a model of leukocyte transmigration should include TJs that are similar to those found in vivo. It was found that exposing HUVEC to culture medium conditioned by human astrocytes increases the TER and maintains the venouslike organization of the TJ complex (Simionescu et al., 1976; Burns et al., 1997; 2000). Importantly, it was observed that the presence or absence of TJs did not affect the rate or extent of neutrophil migration under static conditions (i.e., in the absence of hydrodynamic shear). After silver staining the endothelial borders to reveal the transmigration sites (Figure 29.3), it was found that >75% of the migrating neutrophils preferred corners, even in the absence of TJs. Collectively, these observations led to the hypothesis that neutrophils migrate around endothelial TJs by crossing at tricellular corners rather than by passing through TJs that lie between two endothelial cells. Tricellular corners are also preferred sites for neutrophil firm adhesion and transmigration under physiological flow conditions. This was determined by examination of neutrophil migration across IL-1β-activated HUVEC monolayers placed in a parallel-plate flow chamber. At a wall shear stress of 2 dyn/cm2, 70% of the neutrophils migrated through tricellular corners (Gopalan et al., 2000). Moreover, immunostaining for TJ proteins (ZO-1 and ZO-2) was not altered by neutrophil adhesion and migration (Burns et al., 2000). The distance a neutrophil moved from the time it arrested to the time that it migrated was only 5.5 ± 0.70 µm (under static conditions, the distance was similar, 6.8 ± 0.9 µm) (Gopalan et al., 2000). This distance is less than one cell diameter and shows that neutrophils arrest very close to tricellular corners (Figure 29.4).

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FIGURE 29.2 TJ discontinuities at endothelial tricellular corners. Based on freeze-fracture electron microscopic analysis of pulmonary capillaries, TJs (arrowheads) are found only between pairs of adjacent endothelial cells (ECs). In (A), the flap cell (EC2) is lifted to show that TJs approach each other but are discontinuous at the tricellular corner (vertical arrow). The TJs on EC2 are mirrored on EC1 and EC3; hence, these TJs are also discontinuous (horizontal arrow). TJ discontinuities effectively define a pore (27 nm wide), the entrance of which is indicated by the vertical and horizontal arrows. In (B), a hypothetical situation is diagrammed in which the TJs slide in the plane of the membrane creating a 1 to 2 µm pore, which is wide enough to accommodate a transmigrating neutrophil.

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Distance(µm)until t ra n s m i g ra t i o n

FIGURE 29.3 Human neutrophils migrate preferentially at tricellular corners on HUVEC monolayers treated with IL-1 (10 U/ml, 4 h). In (A), after silver staining and viewing on a light microscope, endothelial borders are resolved (tricellular corners are indicated with arrows). One neutrophil is sitting on the body of an endothelial cell (1), another on top of a tricellular corner (2), and a third is transmigrating at a tricellular corner (3); a portion of the neutrophil is beneath the monolayer (arrowhead). In (B), immunostaining for ZO-1 (a TJ protein) reveals discontinuous staining at tricellular corners (arrows).

10 5

10 20 30 40 Distance rolled (µm)

50

FIGURE 29.4 Rolling neutrophils arrest on stimulated HUVEC monolayers very close (5.5 ± 0.7 µm) to the site of transmigration. HUVEC monolayers were treated with IL-1 (10 U/ml, 4 h) and mounted in a parallel-plate flow chamber. Isolated neutrophils were perfused over the monolayer at 2.0 dyn/cm2 for 11 min and videotaped using phase-contrast optics. Data points are individual neutrophils.

By using immunofluorescence microscopy, gaps were noticed in the border staining patterns for TJ proteins (occludin, ZO-1, and ZO-2) at tricellular corners (see Figure 29.3). These gaps range in size from 0.25 to 2 µm. Assuming an upper value of 2 µm for each corner and five corners/endothelial cell, tricellular corners comprise <10% of the available border area. Since neutrophil transmigration on IL-1-treated HUVEC monolayers occurs exclusively at cell borders, probability predicts that <10% of the neutrophils should migrate at tricellular corners if the

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TABLE 29.1 Comparison between Species — human (HUVEC), dog (DEC), and mouse (MEC) — for Sites of Neutrophil Adhesion and Migration on Stimulated Endothelial Monolayers Adhesion Species Human Dog Mouse

Borders 83.3 ± 8.9a 86.7 ± 3.5a 74.5 ± 5.1a,b

Cell Body 16.7 ± 8.9 13.3 ± 3.5 25.5 ± 5.1c

Migration Tricellular 77.3 ± 4.9d,e 64.0 ± 4.8d,e,f 46.6 ± 6.1e,f,g

Bicellular 22.7 ± 4.9e 26.2 ± 3.4e 43.4 ± 4.3e,h

Transcellular 0 9.3 ± 3.1i 10.0 ± 2.9i

Note: Values are mean ± SD percent of neutrophils based on four to ten separate experiments. Prior to adding neutrophils, HUVEC were stimulated with IL-1 (10 U/ml, 4 h); DEC and MEC were stimulated with LPS (30 ng/ml, 4 h). Neutrophil interactions with endothelial cells occurred in the absence of shear forces (i.e., static conditions) for 360 s (human and dog) or 700 s (mouse) prior to fixation, silver staining, and quantitation as previously described (Burns et al., 1997). a

P < 0.05 vs. same species cell body. P < 0.05 vs. DEC or HUVEC borders. c P < 0.05 vs. DEC or HUVEC. d P < 0.05 vs. same species bicellular. e P < 0.05 vs. same species transcellular. f P < 0.05 vs. HUVEC tricellular. g P < 0.05 vs. DEC tricellular. h P < 0.05 vs. DEC or HUVEC bicellular. i P < 0.05 vs. HUVEC transcellular. b

process is random (Burns et al., 1997). This value is significantly less than the observed values under static and flow conditions (77 and 70%, respectively), suggesting that neutrophil migration is not a random process. Moreover, a comparison across species (human, dog, and mouse) reveals that the preference for tricellular corners is widespread (Table 29.1). Although the percentage of neutrophils migrating at corners is significantly different between species (human > dog > mouse), in each case the number migrating at corners exceeds that predicted by random chance (i.e., ~10%). Finally, in a rabbit model of streptococcal pneumonia, there is evidence that neutrophil emigration from pulmonary capillaries occurs preferentially at tricellular corners. Using scanning electron microscopy to quantify neutrophil emigration, migration sites in the rabbit pulmonary microvasculature were examined after intrabronchial instillation of Streptococcus pneumoniae. Of 28 neutrophils observed in the process of transmigration, 14 (50%) were crossing at tricellular corners (Figure 29.5), 7 (25%) were crossing at bicellular borders, and 7 (25%) were crossing transcytotically. In summary, the in vitro and in vivo data suggest that regardless of species, 50 to 75% of neutrophil transendothelial migration occurs preferentially at tricellular corners where TJs are inherently discontinuous.

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FIGURE 29.5 Scanning electron micrographs showing neutrophil transmigration at endothelial tricellular corners. In (A), a human neutrophil migrates across an IL-1-treated (10 U/ml, 4 h) HUVEC monolayer. In (B), a rabbit neutrophil migrates across the pulmonary microvascular endothelium after intrabronchial instillation of S. pneumoniae (108 organisms, 2 h). Insets (A′ and B′) show endothelial borders (black lines) belonging to the three cells (1, 2, and 3), which define the tricellular corner in A and B, respectively.

29.6 NEUTROPHIL ADHESION TO ENDOTHELIAL BORDERS As mentioned above, tricellular corners are specialized regions of the endothelial border. That rolling neutrophils arrest within one cell diameter of a tricellular corner suggests they are guided to the corner by endothelial determinants. The cell border is a natural topographical feature of the endothelial cell, which is easily appreciated using scanning electron microscopy (see Figure 29.5). Since all cell borders begin and end at a tricellular corner, the possibility is raised that endothelial borders serve to guide neutrophils to tricellular corners. During an acute inflammatory response, endothelial P-selectin (CD62P) can mediate the initial capture of neutrophils from free-flowing blood (Smith, 1993; Nolte et al., 1994; Ley et al., 1995). P-selectin is

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FIGURE 29.6 Histamine induces neutrophil adhesion to endothelial cell borders under hydrodynamic flow conditions (2 dyn/cm2). HUVEC monolayers were mounted in a parallel plate flow chamber and perfused with isolated neutrophils in PBS-containing histamine (10–4 M). At 1 min after perfusion was started, neutrophils began rolling on the endothelium; by 4 min, large numbers of neutrophils were rolling. Fixation and silver staining at 4 min revealed that ~80% of the attached neutrophils were on endothelial cell borders.

stored in secretory granules (Weibel–Palade bodies) and is rapidly expressed on the endothelial surface after stimulation with histamine or thrombin (Bonfanti et al., 1989; Hattori et al., 1989a, b; McEver et al., 1989; Weibel and Palade, 1964). The authors wished to determine if P-selectin-dependent neutrophil capture (adhesion) occurs at endothelial cell borders. Under static or hydrodynamic flow (2 dyn/cm2) conditions, histamine (10–4 M) or thrombin (0.2 U/ml) treatment induced preferential (~80%) neutrophil adhesion to the cell borders of endothelial monolayers (Figure 29.6). Blocking-antibody studies established that neutrophil adhesion was completely P-selectin dependent. P-selectin surface expression increased significantly after histamine treatment and P-selectin immunostaining was concentrated along endothelial borders. Moreover, following histamine infusion in the guinea pig, scanning electron microscopy revealed adherent leukocytes localized to endothelial borders. The authors conclude that preferential P-selectin expression along endothelial borders may be an important mechanism for targeting neutrophil adherence to endothelial borders (Burns et al., 1999). That endothelial borders are important sites for leukocyte adhesion is supported by in vitro observations in human, dog, and mouse models of neutrophil migration across endothelial monolayers exposed to

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cytokine (IL-1, 10 U/ml for 4 h) or lipopolysaccharide (LPS, 30 ng/ml for 4 h). Table 29.1 shows that under static conditions, 75 to 85% of the adherent neutrophils localize to cell borders. Finally, recent independent in vivo observations by He and colleagues (2000), using a silver stain technique, show that adherent leukocytes preferentially (92%) adhere to endothelial borders in frog mesenteric microvessels. A related study found that in addition to storing P-selectin, Weibel–Palade bodies are also capable of storing IL-8 (Wolff et al., 1998). By using immunogold-labeled ultrathin cryosections, it was shown that IL-8 localizes intracellularly in the Golgi apparatus in cytokine-activated HUVEC monolayers. Prolonged stimulation of HUVEC with inflammatory mediators resulted in the accumulation of IL-8 in Weibel–Palade bodies, where it colocalized with von Willebrand factor. IL-8 was retained in these storage organelles for several days after the removal of the stimulus and could be released by endothelial cell secretagogues such as phorbol myristate acetate, calcium ionophore A23187, and histamine. The findings suggest that storage of IL-8 in Weibel–Palade bodies may serve as the endothelial cell “memory” of a preceding inflammatory insult, which then enables the cells to secrete IL-8 immediately without de novo protein synthesis. A logical extension of this finding is that under conditions where Weibel–Palade bodies contain IL-8, fusion with the cell surface may result in preferential P-selectin and IL-8 expression along endothelial borders (see previous paragraph). Whether coexpression of P-selectin and IL-8 occurs on borders is unknown. If it does occur, it is predicted that rolling neutrophils will be captured on the border by P-selectin and activation through IL-8 will induce arrest and firm adhesion. Collectively, these studies suggest that understanding the molecular basis for leukocyte capture on endothelial borders may be critical to understanding leukocyte trafficking into tissues.

29.7 TRANSCYTOTIC NEUTROPHIL MIGRATION Neutrophils need not take a paracellular route across the endothelium. Lymphocytes routinely migrate from blood to tissue and tissue to blood using a transcytotic pathway where the lymphocyte directly perforates the endothelial cell body (cytoplasm) (Toro and Olah, 1967; Cho and De, 1979; 1981; 1986; Azzali and Arcari, 2000). More than 40 years ago, Marchesi and Florey (1960) reported that in addition to observing neutrophil migration through interendothelial clefts, neutrophil pseudopods were sometimes observed penetrating the endothelial cytoplasm at sites immediately adjacent to the clefts (Marchesi, 1961; Marchesi and Florey, 1960). Since this did not appear to be the primary route by which neutrophils migrated across the endothelium, the biologic significance of this observation was unclear. Interestingly, recent in vivo studies appear to support a more important role for transcytotic migration. Subcutaneous injection of formyl peptide elicits rapid neutrophil emigration, which occurs almost exclusively through transcytotic pores in the guinea pig (Feng et al., 1998) and mouse (Hoshi and Ushiki, 1999). Using serial section reconstruction and transmission electron microscopy, Feng and colleagues (1998) determined that in response to formyl peptide, neutrophil migration in vivo

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(guinea pig skin) occurred through transcytotic pores (~1 µm in diameter) and the pore margin was located no closer than 2 µm from an endothelial border. This contrasts sharply to neutrophil migration across cytokine-activated HUVEC monolayers where the migration pore at a tricellular corner directly contacts three endothelial borders (see Figures 29.3 and 29.5). Since transmigration was observed within 15 min of formyl peptide injection, changes in adhesion molecule expression like ICAM-1 and E-selectin and surface presentation of chemotactic factors (e.g., IL-8) are unlikely as they require de novo mRNA and protein synthesis. Extensive changes in surface topography and density of adhesion molecules and surface presentation of chemotactic factors may be necessary for directing neutrophils to endothelial borders. The regulatory mechanism behind transcytotic migration is unknown. However, neutrophil-derived vascular endothelial growth factor (VEGF) may play a critical role in determining whether a neutrophil emigrates through interendothelial clefts or transcytotic pores. While VEGF functions as an endothelial growth and survival factor, it is also known to increase endothelial permeability. Greater than 70% of human neutrophil VEGF is localized to specific granules and, following stimulation, VEGF secretion is detected within 15 min (formyl peptide stimulation) or 30 min (TNFα stimulation) (Gaudry et al., 1997). Topical administration or subcutaneous injection of VEGF in animals induced endothelial gaps (transcytotic pores) and increased permeability within 1 to 10 min (Roberts and Palade, 1995; Feng et al., 1997) and red blood cells and platelets were observed in some pores (Feng et al., 1997). Hence, the possibility exists that in the absence of cytokine-induced endothelial surface remodeling (e.g., upregulation of ICAM-1, VCAM-1, E-selectin, IL-8, and PAF), which may be critical for targeting neutrophils to corners and interendothelial clefts, formyl peptide stimulates neutrophils to secrete VEGF creating transcytotic pores for emigration.

29.8 NEUTROPHIL TRANSEPITHELIAL MIGRATION AND TRICELLULAR CORNERS In some inflammatory lesions (e.g., mucosal inflammation), neutrophils that arrive in the interstitium move toward the basal aspect of an inflamed epithelilum and cross the epithelium by migrating along a paracellular pathway. Interepithelial junctions appear to play a critical role in neutrophil emigration, and a transcytotic migration pathway has not been observed (reviewed by Huber et al., 1998). Epithelial cell lines (MDCK and T84) are used extensively as in vitro models of neutrophil transepithelial migration. Cell monolayers are grown on transwell filters, and neutrophil transmigration is induced by placing neutrophils above the monolayer and a chemoattractant below (e.g., formyl peptide). When the monolayer is grown on the conventional side (top) of the filter, neutrophil migration proceeds in a mucosal-to-serosal direction. Neutrophils will migrate in the more physiological direction (serosal to mucosal) if the epithelial cells are grown on the opposite side (bottom) of the filter to produce an inverted monolayer (Nash et al., 1987; Parkos et al., 1991; Colgan et al., 1993). Using this model, Huber and colleagues (2000) identified a role for occludin in

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modulating neutrophil transepithelial migration. They showed that in response to formyl peptide, neutrophil migration in the mucosal-to-serosal direction increases twofold when MDCK cells express mutant occludin where the N terminus is tagged with hemagglutin (Huber et al., 2000). This is the first demonstration of a functional role for the N-terminal cytoplasmic domain of occludin. Whether occludin modulates neutrophil transepithelial migration in the serosal-to-mucosal direction remains to be determined. It has also been reported that neutrophil migration across the epithelium in a mucosal-to-serosal direction is not random and neutrophils tend to accumulate in clusters beneath the monolayer at so-called invasion sites (Nash et al., 1987; Parkos et al., 1991; Cramer, 1992). Nash and colleagues (1987) noted (data not shown) that TJ morphology as determined by freeze-fracture electron microscopy “is indistinguishable between control and experimental monolayers.” They suggested that “junctional abnormalities, if present, are highly focal.” Nonrandom invasion sites and the preservation of TJ morphology raise the possibility that neutrophils migrate preferentially at epithelial tricellular corners where TJs are discontinuous (Walker et al., 1994). To the authors’ knowledge, despite numerous publications and reviews on neutrophil migration across intestinal epithelial monolayers, the issue of tricellular corner migration has never been considered and warrants further investigation. In lung, there is suggestive evidence that epithelial tricellular corners are preferred sites for neutrophil emigration into the alveolar space. Using lavage to induce an inflammatory reaction in dog lung, Damiano et al. (1980) observed by scanning and transmission electron microscopy that neutrophils within the alveolar wall interstitium migrate preferentially across the alveolar epithelium at sites where the borders of squamous Type I pneumocytes meet cuboidal Type II pneumocytes. Similarly, in a rabbit model of streptococcal pneumonia, Walker observed neutrophils migrating between Type I and II pneumocytes (Walker et al., 1995; Behzad et al., 1996). These emigration sites are special for two reasons. First, neutrophils appear to be guided to these emigration sites by interstitial fibroblasts that form cellular bridges between the capillary endothelium and Type II pneumocytes. Neutrophils adhere to the fibroblasts and crawl to holes in the basal lamina through which fibroblasts contact Type II pneumocytes. By displacing fibroblast contacts with Type II pneumocytes, neutrophils migrate into the basolateral space of Type II pneumocytes from which they emerge onto the alveolar surface adjacent to the Type II pneumocytes. Second, the emigration sites appear to localize to epithelial tricellular corners where the borders of two Type I pneumocytes meet the border of a Type II pneumocyte (Figure 29.7). This is all the more remarkable when one considers that Type II pneumocytes comprise only 5% of the alveolar epithelial surface; the remaining 95% of the surface is occupied by Type I pneumocytes. Clearly, these data suggest that fibroblasts play a critical role in guiding neutrophils to preferred epithelial migration sites. Whether fibroblasts play a similar role in other tissues is unknown. However, it raises the issue that interstitial cells may regulate and direct leukocyte transepithelial migration. Hence, investigators using in vitro models to study leukocyte transepithelial migration may need to consider placing an interstitial environment beneath the epithelial monolayer to mimic the in vivo condition.

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FIGURE 29.7 Scanning electron micrograph showing neutrophil transepithelial migration in rabbit lung during streptococcal pneumonia. Streptococcus pneumoniae (108 organisms) was instilled into the bronchus of a rabbit. After 4 h, the lungs were removed and fixed. A neutrophil is emerging into the alveolus at a tricellular corner (arrow) where the borders between two Type I pneumocytes (1 and 2) and a single Type II pneumocyte (3) converge.

29.9 MECHANISMS FOR OPENING/REMODELING TIGHT JUNCTIONS In vitro, neutrophils can migrate quickly across endothelial cells (seconds to minutes) without affecting the barrier function of the monolayer (Huang et al., 1988; 1993; Burns et al., 2000). Neutrophil migration does not involve widespread proteolytic degradation of TJs (Burns et al., 2000) or adherens junctions (Moll et al., 1998). Although tricellular corners are preferred migration sites on activated endothelial monolayers (see Table 29.1), freeze-fracture observations on pulmonary capillaries show that the size of the pore created by the TJ discontinuity is as small as 27 nm (Walker et al., 1994). Transmission electron microscopic observations suggest that neutrophils migrate through pores that are 1 to 2 µm wide (Marchesi and Florey, 1960; Burns et al., 2000). From in vitro studies, it is also clear that a significant number of neutrophils migrate between pairs of adjacent endothelial cells (i.e., bicellularly; see Table 29.1). TJ fibrils between these cells are discontinuous, but the gaps between these fibrils are still too small (100 to 200 nm) to allow neutrophils to pass (Figure 29.8). Hence, it is hypothesized that lateral displacement of the TJ complex is necessary to create a pore wide enough (1 to 2 µm) to accommodate a transmigrating neutrophil. Displacement of the junctional complex may require the neutrophil to signal the endothelium. Activated neutrophils are known to induce a coordinate increase in endothelial cytosolic free calcium, phosphorylation of serine 19 and threonine 18 of endothelial myosin regulatory light chains, and isometric tension generation by endothelial monolayers (Hixenbaugh et al., 1997). A role for endothelial myosin

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FIGURE 29.8 Freeze-fracture replicas of HUVEC monolayers before and during neutrophil transendothelial migration. In (A), venous endothelial TJs (arrows) are clearly observed in control monolayers (arrows). In (B), the endothelium was treated with IL-1 (10 U/ml, 4 h) and fixed 5 min after the addition of isolated neutrophils in PBS. The tail-end (arrow) of a transmigrating neutrophil (N) protrudes into the lumen above the endothelial monolayer (E). At the site of penetration, the fracture plane passes through a portion of the endothelial membrane that lines the migration pore (*). An enlarged view of this membrane (inset) shows that TJs are absent. The arrow in the inset denotes the lip of the migration pore.

light-chain kinase (MLCK) is suggested by the observation that an MLCK inhibitor diminished neutrophil transmigration and inhibited endothelial F-actin formation, myosin filament formation, and MLC phosphorylation (Saito et al., 1998). Consistent with these observations is the finding that disruption of endothelial microfilaments by cytochalasin B or latrunculin A markedly inhibits monocyte transmigration (Kielbassa et al., 1998). Collectively, these data suggest that leukocyte transmigration is intimately tied to the dynamic regulation of the endothelial cytoskeleton. Since TJ transmembrane proteins occludin and claudin are anchored to the cytoskeleton through accessory cytoplasmic linker proteins (Furuse et al., 1994; Yamamoto et al., 1997; Fanning et al., 1998; Itoh et al., 1999; Wittchen et al., 1999), rearrangements in the TJ complex (e.g., displacement or loosening) are likely mediated by the endothelial cytoskeleton.

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FIGURE 29.9 Freeze-fracture replicas of guinea pig tracheal epithelium before and after mechanical stretching. Isolated tracheas were cut along the ventral midline, pinned flat on dental wax in a relaxed (A) or stretched (longitudinally and circumferentially as permitted by the underlying connective tissue; B) position, and fixed in buffered glutaraldehyde. Quantitative analysis of the freeze-fracture images showed that stretching the trachea decreased the height of the junctional complex from 0.326 ± 0.0061 µm to 0.242 ± 0.0073 µm (P = 0.01) and produced small breaks (arrows) in the TJ fibrils. Symbols: lumen (L); microvillus (M); lateral membrane (LM).

TJs may also be able to slide within the membrane. When an epithelium is mechanically stretched, freeze-fracture electron microscopy shows that the height of the TJ complex shortens by as much as 25% and small gaps appear in the fibrils (Figure 29.9). During an acute inflammatory response to cigarette smoke exposure, plasma exudate can expand epithelial tricellular corners, seemingly driving the TJs apart (Figure 29.10). Collectively, these observations suggest that active (cytoskeletal-mediated) and passive responses by the endothelium could allow the TJs to move within the membrane. The authors suggest that if this were to occur at the preferred migration site (i.e., the tricellular corner), the tripartite junctional complex would move apart as the neutrophil entered the intercellular cleft and move together as the pore closed behind the transmigrated neutrophil (see Figure 29.2). This model predicts that TJs will be absent from the endothelial membrane lining the migration pore, and this is precisely what is found by freeze-fracture electron microscopy (see Figure 29.8).

29.10 CONCLUDING REMARKS Neutrophils migrate across endothelial and epithelial cell layers during inflammatory reactions. Intercellular TJs are potential barriers through which transmigrating neutrophils must pass. Although the TJ complex is beltlike in design, it has inherent discontinuities at tricellular corners where the borders of three endothelial (or epithelial) cells converge. In certain inflammatory settings, neutrophils migrate preferentially at

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FIGURE 29.10 Tracheal epithelial tricellular corners expand and fill with plasma after cigarette smoke exposure. After a 20-min exposure to the smoke of five cigarettes the animal was sacrificed and the trachea removed and processed for thin-section transmission electron microscopy. This en face section of edematous tracheal epithelium shows a tricellular corner where the intercellular space is dilated and filled with plasma. Although TJs are still present between pairs of adjacent endothelial cells (arrows), the discontinuity between TJs at tricellular corners is enlarged (*).

tricellular corners. On activated endothelial cells, neutrophil transmigration can occur rapidly (seconds to minutes) without evidence of TJ proteolysis. Neutrophil migration at tricellular corners through TJ discontinuities provides a mechanism to explain how endothelial cells preserve their barrier function during an acute inflammatory response.

ACKNOWLEDGMENTS This work was supported by grants from the American Lung Association (ALARG068N; A.R.B.), National Institutes of Health (AI-46773; A.R.B.), and (HL-42550, ES-06091, AI-19031; C.W.S.), the Methodist Hospital Foundation (A.R.B.), and a Chao Fellowship (A.R.B.).

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Hattori, R., K. Hamilton, R. McEver, and P. Sims. 1989b. Complement proteins C5b-9 induce secretion of high molecular weight multimers of endothelial von Willebrand factor and translocation of granule membrane protein GMP-140 to the cell surface. J. Biol. Chem., 264:9053–9060. He, P., J. Wang, and M. Zeng. 2000. Leukocyte adhesion and microvessel permeability. Am. J. Physiol., 278:H1686–1694. Hixenbaugh, E., Z. Goeckeler, N. Papaiya, R. Wysolmerski, S. Silverstein, and A. Huang. 1997. Stimulated neutrophils induce myosin light chain phosphorylation and isometric tension in endothelial cells. Am. J. Physiol., 273:H981–988. Hoshi, O., and T. Ushiki. 1999. Scanning electron microscopic studies on the route of neutrophil extravasation in the mouse after exposure to the chemotactic peptide N-formyl-methionyl-leucyl-phenylalanine (fMLP). Arch. Histol. Cytol., 62:253–260. Huang, A. J., M. B. Furie, S. C. Nicholson, J. Fischbarg, L. S. Liebovitch, and S. C. Silverstein. 1988. Effects of human neutrophil chemotaxis across human endothelial cell monolayers on the permeability of these monolayers to ions and macromolecules. J. Cell Physiol., 135:355–366. Huang, A. J., J. E. Manning, T. M. Bandak, M. C. Ratau, K. R. Hanser, and S. C. Silverstein. 1993. Endothelial cell cytosolic free calcium regulates neutrophil migration across monolayers of endothelial cells. J. Cell Biol., 120:1371–1380. Huber, D., M. S. Balda, and K. Matter. 1998. Transepithelial migration of neutrophils. Invasion Metastasis, 18:70–80. Huber, D., M. S. Balda, and K. Matter. 2000. Occludin modulates transepithelial migration of neutrophils. J. Biol. Chem., 275:5773–5778. Itoh, M., M. Furuse, K. Morita, K. Kubota, M. Saitou, and S. Tsukita. 1999. Direct binding of three tight junction-associated MAGUKs, ZO-1, ZO-2, and ZO-3, with the COOH termini of claudins. J. Cell Biol., 147:1351–1363. Jones, D., O. Abbassi, L. McIntire, L. McEver, and C. Smith. 1993. P-selectin mediates neutrophil rolling on histamine-stimulated endothelial cells. Biophys J., 65:1560–1569. Kielbassa, K., C. Schmitz, and V. Gerke. 1998. Disruption of endothelial microfilaments selectively reduces the transendothelial migration of monocytes. Exp. Cell Res., 243:129–141. Ley, K., D. Bullard, M. Arbones, R. Bosse, D. Vestweber, T. Tedder, and A. Beaudet. 1995. Sequential contribution of L- and P-selectin to leukocyte rolling in vivo. J. Exp. Med., 181:669–675. Marchesi, V. T. 1961. The site of leucocyte emigration during inflammation. Q. J. Exp. Physiol., 46:115–133. Marchesi, V. T., and H. W. Florey. 1960. Electron micrographic observations on the emigration of leucocytes. Q. J. Exp. Physiol., 45:343–348. Martin-Padura, I., S. Lostaglio, M. Schneemann, L. Williams, M. Romano, P. Fruscella, C. Panzeri, A. Stoppacciaro, L. Ruco, A. Villa, D. Simmons, and E. Dejana. 1998. Junctional adhesion molecule, a novel member of the immunoglobulin superfamily that distributes at intercellular junctions and modulates monocyte transmigration. J. Cell Biol., 142:117–127. McCarthy, K., I. Skare, M. Stankewich, M. Furuse, S. Tsukita, R. Rogers, R. Lynch, and E. Schneeberger. 1996. Occludin is a functional component of the tight junction. J. Cell Sci., 109:2287–2298. McEver, R., J. Beckstead, K. Moore, L. Marshall-Carlson, and D. Bainton. 1989. GMP-140, a platelet alpha-granule membrane protein, is also synthesized by vascular endothelial cells and is localized in Weibel–Palade bodies. J. Clin. Invest., 84:92–99. Moll, T., E. Dejana, and D. Vestweber. 1998. In vitro degradation of endothelial catenins by a neutrophil protease. J. Cell Biol., 140:403–407.

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Morita, K., H. Sasaki, M. Furuse, and S. Tsukita. 1999. Endothelial claudin: claudin5/TMVCF constitutes tight junction strands in endothelial cells. J. Cell Biol., 147:185–194. Muller, W. A. 1995. The role of PECAM-1 (CD31) in leukocyte emigration: studies in vitro and in vivo. J. Leukocyte Biol., 57:523–528. Muller, W. A., C. M. Ratti, S. L. McDonnell, and Z. A. Cohn. 1989. A human endothelial cell-restricted, externally disposed plasmalemmal protein enriched in intercellular junctions. J. Exp. Med., 170:399–414. Muller, W. A., S. A. Weigl, X. Deng, and D. M. Phillips. 1993. PECAM-1 is required for transendothelial migration of leukocytes. J. Exp. Med., 178:449–460. Nash, S., J. Stafford, and J. L. Madara. 1987. Effects of polymorphonuclear leukocyte transmigration on the barrier function of cultured intestinal epithelial monolayers. J. Clin. Invest., 80:1104–1113. Nolte, D., P. Schmid, U. Jager, A. Botzlar, F. Roesken, R. Hecht, E. Uhl, K. Messmer, and D. Vestweber. 1994. Leukocyte rolling in venules of striated muscle and skin is mediated by P-selectin, not by L-selectin. Am. J. Physiol., 267:H1637–1642. Parkos, C. A., C. Delp, M. A. Arnaout, and J. L. Madara. 1991. Neutrophil migration across a cultured intestinal epithelium. Dependence on a CD11b/CD18-mediated event and enhanced efficiency in physiological direction. J. Clin. Invest., 88:1605–1612. Roberts, W. G., and G. E. Palade. 1995. Increased microvascular permeability and endothelial fenestration induced by vascular endothelial growth factor. J. Cell Sci., 108:2369–2379. Saito, H., Y. Minamiya, M. Kitamura, S. Saito, K. Enomoto, K. Terada, and J. Ogawa. 1998. Endothelial myosin light chain kinase regulates neutrophil migration across human umbilical vein endothelial cell monolayer. J. Immunol., 161:1533–1540. Schneeberger, E. E., and R. D. Lynch. 1992. Tight junctions in the lung, in Tight Junctions, Cereijido, M., Ed., CRC Press, Boca Raton, FL, 337–351. Schnittler, H. J. 1998. Structural and functional aspects of intercellular junctions in vascular endothelium. Basic Res. Cardiol., 93:30–39. Simionescu, M., N. Simionescu, and G. E. Palade. 1976. Segmental differentiations of cell junctions in the vascular endothelium. Arteries and veins. J. Cell Biol., 68:705–723. Simionescu, N., M. Simionescu, and G. E. Palade. 1978a. Open junctions in the endothelium of the postcapillary venules of the diaphragm. J. Cell Biol., 79:27–44. Simionescu, N., M. Simionescu, and G. E. Palade. 1978b. Structural basis of permeability in sequential segments of the microvasculature of the diaphragm. I. Bipolar microvascular fields. Microvasc. Res., 15:1–16. Smith, C. 1993. Endothelial adhesion molecules and their role in inflammation. Can. J. Physiol. Pharmacol., 71:76–87. Springer, T. A. 1994. Traffic signals for lymphocyte recirculation and leukocyte emigration: the multistep paradigm. Cell, 76:301–314. Staehelin, L. 1974. Structure and function of intercellular junctions. Int. Rev. Cytol., 39:191–283. Stein, B., Y. Khew-Goodall, J. Gamble, and M. A. Vadas. 1997. Transmigration of leukocytes, in Endothelium in Clinical Practice: Source and Target of Novel Therapies, Rubanyi, G. M., and V. J., Dzau, Eds., Marcel Dekker, New York, 149–202. Toro, I., and I. Olah. 1967. Penetration of thymocytes into the blood circulation. J. Ultrastruct. Res., 17:439–451. Tsukita, S., M. Furuse, and M. Itoh. 1996. Molecular dissection of tight junctions. Cell Struct. Funct., 21:381–385. Tsukita, S., M. Furuse, and M. Itoh. 1999. Structural and signalling molecules come together at tight junctions. Curr. Opin. Cell Biol., 11:628–633.

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Wakelin, M. W., M. J. Sanz, A. Dewar, S. M. Albelda, S. W. Larkin, N. Boughton-Smith, T. J. Williams, and S. Nourshargh. 1996. An anti-platelet-endothelial cell adhesion molecule-1 antibody inhibits leukocyte extravasation from mesenteric microvessels in vivo by blocking the passage through the basement membrane. J. Exp. Med., 184:229–239. Walker, D. C., A. MacKenzie, W. C. Hulbert, and J. C. Hogg. 1985. A re-assessment of the tricellular region of epithelial cell tight junctions in trachea of guinea pig. Acta Anat., 122:35–38. Walker, D., A. MacKenzie, B. Wiggs, J. Montaner, and J. Hogg. 1988. Assessment of tight junctions between pulmonary epithelial and endothelial cells. J Appl Physiol. 64:2348–2356. Walker, D. C., A. MacKenzie, and S. Hosford. 1994. The structure of the tricellular region of endothelial tight junctions of pulmonary capillaries analyzed by freeze-fracture. Microvasc. Res., 48:259–281. Walker, D. C., A. R. Behzad, and F. Chu. 1995. Neutrophil migration through preexisting holes in the basal laminae of alveolar capillaries and epithelium during streptococcal pneumonia. Microvasc. Res., 50:397–416. Weibel, E., and G. Palade. 1964. New cytoplasmic components in arterial endothelia. J. Cell Biol., 23:101–112. Wittchen, E. S., J. Haskins, and B. R. Stevenson. 1999. Protein interactions at the tight junction. Actin has multiple binding partners, and ZO-1 forms independent complexes with ZO-2 and ZO-3. J. Biol. Chem., 274:35179–35185. Wolff, B., A. R. Burns, J. Middleton, and A. Rot. 1998. Endothelial cell “memory” of inflammatory stimulation: human venular endothelial cells store interleukin 8 in Weibel–Palade bodies. J. Exp. Med., 188:1757–1762. Yamamoto, T., N. Harada, K. Kano, S. Taya, E. Canaani, Y. Matsuura, A. Mizoguchi, C. Ide, and K. Kaibuchi. 1997. The Ras target AF-6 interacts with ZO-1 and serves as a peripheral component of tight junctions in epithelial cells. J. Cell Biol., 139:785–795.

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Ocular Tight Junctions in Health, Disease, and Glaucoma Johnnie L. Underwood and Collin G. Murphy

CONTENTS 30.1 Introduction .................................................................................................654 30.2 Tight Junctions in Cornea...........................................................................655 30.3 Blood–Ocular Barriers ................................................................................657 30.3.1 The Blood–Aqueous Barrier .........................................................659 30.3.1.1 Tight Junctions in the Ciliary Body..............................659 30.3.1.2 Tight Junctions in Ciliary Epithelial Blood Vessels .....662 30.3.1.3 Tight Junctions in the Pigmented Epithelium of the Iris............................................................................662 30.3.1.4 Tight Junctions in the Iris Stroma ................................663 30.3.2 Blood–Retinal Barrier....................................................................664 30.3.2.1 Tight Junctions in the Pigmented Epithelium ..............664 30.3.2.2 Tight Junctions in Nonfenestrated Capillaries..............665 30.3.2.3 Tight Junctions in Choroidal Vasculature Endothelium...................................................................666 30.4 The Role of Tight Junctions in the Development of Glaucoma................666 30.4.1 The Aqueous Outflow System.......................................................666 30.4.2 Potential Barriers to the Flow of Aqueous Humor out of the Eye ...........................................................................................668 30.4.3 Tight Junctions in Trabecular Meshwork and Schlemm’s Canal Endothelial Cells .................................................................669 30.4.4 Hormonal Regulation of Transendothelial Fluid Flow .................669 30.4.5 Tight Junction Structure in Schlemm’s Canal Endothelial Cells ...............................................................................................671 30.4.6 The Rate of Transendothelial Fluid Flow Correlates with Changes in Tight Junctions ...........................................................672 30.4.7 ZO-1 Is Essential for Maintenance of Tight Junction Integrity and Regulation of Transendothelial Fluid Flow ...........................672 30.4.8 Patterns of Gene Expression Associated with Changes in Transendothelial Fluid Flow..........................................................673 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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30.5 Conclusion ..................................................................................................676 Acknowledgment ...................................................................................................677 References..............................................................................................................677

30.1 INTRODUCTION The eye is an extraordinary organ containing a variety of tissues, each of which contributes to the production of clear vision. Within these tissues, tight junctions (TJs) function in diverse ways to furnish barriers essential for the maintenance of normal ocular function. The roles of TJs in the tissues that maintain vision are reviewed in this chapter. The tissues within the eye are illustrated in a schematic diagram in Figure 30.1. The ciliary body, retina, and iris exhibit barrier properties that prevent blood from crossing freely into the ocular chambers. Thus, they represent part of a blood–ocular barrier that allows only clear fluid to perfuse the anterior and posterior chambers of the eye. The aqueous outflow system, which consists of the trabecular meshwork and Schlemm’s canal, is the means by which the aqueous humor produced by the ciliary body exits the eye (indicated by arrows in Figure 30.1). TJs are essential for normal corneal transparency, for aqueous humor production by the ciliary body, and for normal function of the choroid, retina, and aqueous outflow system. Malfunction of TJ barriers in most ocular tissues is associated with opacity of fluids and influx of fluids into the extracellular matrix that lead to disruption of

FIGURE 30.1 Major tissues in the eye: a schematic cross-section. TJs are important for normal function of the cornea, the ciliary body and iris, aqueous humor outflow system, retina, and choroid. The direction of aqueous humor flow is indicated by arrows. Aqueous humor is produced by the ciliary body, flows from the posterior chamber through the pupil, fills the anterior chamber, and maintains the intraocular pressure. It exits the eye through the aqueous outflow system, which includes the trabecular meshwork and Schlemm’s canal, and returns to the venous system.

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normal function. However, in open-angle glaucoma, increased resistance of the TJ barriers in the endothelial cells that line the aqueous humor outflow pathway reduces the egress of aqueous humor out of the eye and contributes to higher intraocular pressure and consequent nerve damage (Underwood et al., 1999). In the following sections, functions of TJs in specific ocular tissues are discussed, with special reference to the pathogenesis of glaucoma.

30.2 TIGHT JUNCTIONS IN CORNEA A clear cornea is essential for normal vision. Both the epithelial and endothelial cell layers contain TJ barriers that prevent free entry of large molecules into the transparent stroma (Figure 30.2). The corneal epithelium covers the anterior surface of the globe with five to seven pseudo-stratified layers (Figure 30.2). The outermost or superficial layer of the epithelium is the first line of defense against dehydration, abrasion, and infection. These cells are held firmly together by a continuous zonula occludens as demonstrated both by freeze-fracture (McLaughlin et al., 1985) and by thin-section electron microscopy studies (Hogan et al., 1971; Wang et al., 1993; Gipson and Sugrue, 1994). ZO-1 is found in an annular pattern that is associated with the TJs surrounding each of the superficial cells (Sugrue and Zieske, 1997). This epithelial layer therefore has a sealed paracellular route that provides resistance

FIGURE 30.2 TJs in the cornea. Superficial cells of the corneal epithelium are surrounded by a continuous or annular zonula occludens. Corneal endothelial cells contain focal TJs, and face the anterior chamber. Both types of TJs are indicated by the black ellipses. The relative thickness of the stroma is greater than shown, as indicated by the double-headed broken arrow. Both Bowman’s layer and Descemet’s membrane are basement membranes.

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to fluid flow and prevents entry of pathogens from outside the eye. Moreover, it confers a high resistance to the passage of ions, since its transepithelial electrical resistance (TER), measured at more than 1000 Ω·cm2 (Klyce and Wong, 1977), is close to that of typical barrier epithelia. The corneal epithelium as a whole is a plastic tissue with a rapid turnover and is replaced in its entirety approximately every week (Hanna and O’Brien, 1960; Hanna et al., 1961). The barrier in the corneal epithelium is dependent on the continuous replacement of the superficial cells that are subject to wear and tear. As superficial cells become worn or damaged, cells from the basal layer divide and move toward the surface, where new TJ connections are formed. The basal cell layer next to Bowman’s membrane does not have TJs and does not have the ZO-1 protein (Wang et al., 1993).There are also no TJs in the wing cells, but they do have ZO-1 distributed in a punctate pattern that is associated with adherens junctions that provide cell–cell and cell–matrix adhesion (Sugrue and Zieske, 1997). Experimental removal of the surface cells in vivo causes a time-dependent increase in ZO-1 expression in the lower cell layers and movement of those cells into the outer layer to reestablish the barrier function (Wang et al., 1993). The cells of the corneal endothelium, like those of the corneal epithelium, have TJs (see Figure 30.2) and adherens junctions, but also have gap junctions. The endothelium separates the aqueous humor of the anterior chamber from the corneal stroma. It allows nutrients such as glucose to enter the cornea (Thoft et al., 1971), but prevents entry of large proteins and cells that cause opacity. These two opposing functions are supported by a “pump–leak” mechanism described by Maurice nearly 30 years ago (Maurice, 1972; 1984; Waring et al., 1982; Fischbarg and Lim, 1984). According to this theory, fluid is “pumped” out of the cornea by the endothelial cells that each have approximately 3 million Na+,K+-ATPase pump sites (Geroski and Edelhauser, 1984). The “leak” could be associated with the existence of an incomplete, although often extensive and anastomosing, TJ belt such as that present in cultured bovine corneal endothelial cells as shown by freeze-fracture in a study done by Noske and Hirsch (Figure 30.3) (Hirsch et al., 1977; McLaughlin et al., 1985; Tuberville et al., 1989; Stiemke et al., 1991; Noske et al., 1994). The incomplete barrier results in a relatively low TER (transepithelial resistance) (i.e., ~20 to 73 Ω·cm2) (Lim and Fischbarg, 1981; Hodson and Wigham, 1983; Geroski and Hadley, 1992; Noske et al., 1993; Le Varlet et al., 1995), allowing tracers such as lanthanum, ruthenium red, and horseradish peroxidase (HRP) to penetrate the paracellular route with ease (Kaye et al., 1973; Leuenberger, 1973; Kruetziger, 1976; Hirsch et al., 1976; Montcourrier and Hirsch, 1985). Alternatively, the unusual distribution of TJs in the endothelial cell monolayer may provide another basis for the “pump–leak” mechanism. Corneal endothelial cells have been shown to form gaps as wide as 1.0 µm at the intersections where three cells meet (Barry et al., 1995; Petroll et al., 1999). These gaps are termed Y-junctional regions or tricellular corners and represent sites of particularly large interruptions in barrier function and therefore may be locations of enhanced permeability or the location of the “leak.” In contrast, the remaining borders, with larger concentrations of focal TJs, may provide more effective “pump” sites. The focal nature of this belt was illustrated in Figure 30.4 by ZO-1 labeling of human corneal

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FIGURE 30.3 Focal TJs. Freeze-fracture images of bovine corneal endothelial cells. Apicobasally oriented TJ strands on a fracture-P face (P) with few anastomoses and many free ends. Arrows indicate regions of the plasma membrane not barred with TJ elements. Bar = 0.5 µm. (Fromn Noske, W. and Hirsch, M., Cell Tissue Res., 245, 405, 1986. With permission.)

endothelium (Petroll, 1996). The figure shows that ZO-1 and the Y-junctional space are present on the apicolateral edge of the cells. The absence of occludin in these cells (Petroll et al., 1996; Hirase et al., 1997; Saitou et al., 1998) combined with the existence of leaky Y-junctional regions represents a major difference from classical epithelial barrier systems. Thus, this incomplete barrier and the relative “leakiness” of the corneal endothelium allows entry of nutrients in the form of small molecules into the avascular corneal tissues. It is easy to understand how disruption of TJs in this delicately balanced pump–leak system could cause accumulation of fluid and proteins that result in opacity of the stroma and impaired vision (Waring et al., 1982).

30.3 BLOOD–OCULAR BARRIERS The eye may be regarded as an extension of the brain, and like the brain, it is separated from free access to the blood by a “blood–ocular barrier” (Rubin and Staddon, 1999). Many of the same issues that arise with regard to the blood–brain barrier are applicable to its ocular counterpart. As in the brain, these barriers prevent exposure to most immune cells and circulating antibodies; consequently, many parts of the eye are immunologically privileged sites. The two major components of the blood–ocular barrier are the blood-aqueous barrier and the blood–retinal barrier. These barriers help to maintain the clear media that fills the globe and provides nutrients to the ocular tissues.

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FIGURE 30.4 Tricellular corners in human corneal endothelium are shown by ZO-1 staining in situ. Double labeling of β-catenin (a, c, e, g) and ZO-1 (b, d, f, h). Images were taken from a z-series obtained by imaging in 0.5-µm steps from the basal to apical aspect of the endothelial cells. Cross-sectional x–z projections (bottom panel) were reconstructed from the areas indicated by the horizontal lines in g and h. β-catenin formed a pericellular ring apically (e, g) and exhibited more convoluted staining basolaterally (a, c). ZO-1 was organized into a single apical pericellular ring (f, h), and only nonspecific background staining was detected basally (b). ZO-1 was nearly completely membrane associated, whereas β-catenin appeared to extend slightly from the apical membrane into the cytoplasm. Interestingly, breaks in the staining of ZO-1 were noted, with the largest gaps at the Y junction of adjacent cells (f, arrows). β-catenin also demonstrated gaps, albeit smaller ones, at some Y junctions (e, arrows). Differences in the three-dimensional organization of β-catenin and ZO-1 are more clearly demonstrated in the x–z projections (bottom panel). Cytoplasmic staining between the bright apical cell junctions in the x–z projection of ZO-1 is due to nonspecific background staining (scale bar = 15 µm). (From Petroll, W. et al., Curr. Eye Res., 18, 10, 1999. With permission.)

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FIGURE 30.5 TJs in the ciliary body. The folded region of the ciliary body is called the pars plicata and has two layers of cells joined at their apices. The pigmented epithelium lacks TJs, as do adjacent blood vessels in the stroma. Aqueous humor is produced by the nonpigmented epithelium and secreted into the posterior chamber (arrows). The nonpigmented cells have continuous apical TJs (black ellipses).

30.3.1 THE BLOOD–AQUEOUS BARRIER The blood–aqueous barrier is a complex tissue that includes the ciliary body and the neighboring iris, specifically, the nonpigmented ciliary epithelium, the ciliary endothelial blood vessels, the iris blood vessels, and the iris pigmented epithelium. This barrier prevents blood elements from entering the aqueous humor indiscriminately and at the same time allows production of aqueous humor, a cell-free fluid produced by ciliary body cells, that has a much lower protein content (<0.2%) than does plasma (~7%) (Bito, 1977; Cole, 1984). 30.3.1.1 Tight Junctions in the Ciliary Body The ciliary body lies within the vitreous space of the posterior chamber and extends around the circumference of the eye, posterior to the outer edges of the iris (see Figure 30.1). Aqueous humor is produced by the layer of nonpigmented epithelium covering the folded portion, or pars plicata, of the ciliary body, where it is secreted into the posterior chamber (see Figures 30.1 and 30.5). The annular arrangement of TJs in the nonpigmented ciliary epithelium, shown by freeze fracture (Figure 30.6), constitutes the main part of the blood–aqueous barrier and ensures that most macromolecules from the blood do not enter the aqueous humor by passive flow (Ohkuma and Nishiura, 1974; Raviola, 1974; Reale and Spitznas, 1975; Hirsch et al., 1980; 1984; Noske and Hirsch, 1986). However, the complexity of TJs in the nonpigmented ciliary epithelium is highly variable and often represented by only a small number of strands (Raviola and Raviiola, 1978; Hirsch et al., 1984; Noske and Hirsch, 1986;

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FIGURE 30.6 Ciliary epithelial TJs. Freeze-fracture images of rabbit ciliary epithelial TJs. TJs of the ciliary epithelium in a control eye. The TJ is revealed as a network of anastomosing ridges on the P face (PF) of the plasma membrane and as furrows on the E face (EF). The ridges show some discontinuities (black arrows) where in the furrows some particles or short bars may be found (white arrows). The number of parallel strands varies and the density of intramembranous particles is lower within the junctional network (asterisk) than in the corresponding Mace of the surrounding plasma membrane. (Inset) No reaction product of HRP is seen beyond the level of the TJ (arrow). GJ = gap junction. Original magnification ×75,000. (From Noske, W. and Hirsch, M., Cell Tissue Res., 245, 405, 1986. With permission.)

Noske et al., 1986). Its function has been compared with that of leaky epithelia such as rabbit gallbladder (Claude and Goodenough, 1973; Raviola and Raviiola, 1978), with a high hydraulic conductivity (Pederson and Green, 1973) and a low transport potential (Berggren, 1960; Miller and Constant, 1960). The complexity of the TJ arrays in the nonpigmented epithelium differs depending on location, with the more complex networklike arrays in the ciliary crests, and less complex ones in the “valleys” between the crests and in the pars plana (flat part of the ciliary body) (Pederson and Green, 1973). The barrier in the nonpigmented ciliary epithelium was demonstrated clearly in studies by Raviola in 1974. He showed that HRP injected into the bloodstream could easily pass through the fenestrated capillaries of the ciliary body into the ciliary pigmented epithelium and the intercellular space between nonpigmented ciliary epithelial cells, but could not pass farther than the apical ends of the nonpigmented ciliary cells because it was stopped by continuous TJs (Figures 30.5 and 30.7) (Raviola, 1974; 1977; Raviola and Raviola, 1978; Freddo and Raviola, 1982). The ciliary TJ barrier also prevents reentry of aqueous humor back into the secretory cells, ensuring unilateral accumulation of aqueous outside the basal surface of the epithelium. This accumulation or net production of aqueous humor creates the intraocular pressure required for normal ocular development and function, including the folding of the secretory epithelium itself into the pars plicata and the establishment and maintenance of normal corneal curvature (Coulombre, 1965; Bard and Ross, 1982a, b; Beebe, 1986). Intraocular pressure, maintained at a normal level, keeps the cornea from flattening against the iris (which would impede aqueous outflow) and in general provides for “mechanical stability of the ocular components”

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FIGURE 30.7 The perfusion of HRP is blocked by the TJs of the ciliary nonpigmented epithelial cells. Ciliary epithelium 10 min after injection of HRP. The tracer has penetrated the intercellular clefts between pigmented cells and those intervening between pigmented and nonpigmented cells (arrows). No reaction product is seen in the intercellular clefts between nonpigmented cells or in the posterior chamber. Two melanosomes are contained in the nonpigmented cells (asterisk). Original magnification ×9,900. (From Raviola, G., Invest. Ophthalmol., 13, 828, 1974. With permission.)

(Raviola, 1982). Many glaucoma medications, including carbonic anhydrase inhibitors, β-adrenergic antagonists, and β-adrenergic agonists, are thought to reduce intraocular pressure by decreasing the amount of aqueous humor secreted by the

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nonpigmented ciliary epithelium (Woodlief, 1980; Araie and Takase, 1981; Gharagozloo et al., 1988). Given the ultrastructure and known barrier functions of the nonpigmented epithelium, it is not surprising that ZO-1 and occludin colocalize at the TJs of the nonpigmented ciliary epithelial cells (Tserentsoodol et al., 1998). Another characteristic of cells of blood–tissue barriers, present in nonpigmented epithelium, is the glucose transporter GLUT-1, which provides an alternative mechanism for glucose entrance into these tissues (Harik et al., 1990; Takata et al., 1990; 1991; 1997). Investigators have recently found that GLUT-1 accumulates in both pigmented and nonpigmented ciliary epithelial cells, enabling the selective transfer of glucose, first from the blood to the pigmented cells and then from the nonpigmented cells into the aqueous humor across the blood–aqueous barrier (Takata et al., 1990; 1997; Tserentsoodol et al., 1998). 30.3.1.2 Tight Junctions in Ciliary Epithelial Blood Vessels Blood vessels in the nonpigmented ciliary epithelium have different barrier properties depending on their locations. Vessels situated at the bases of the ciliary processes and near the ciliary muscle have a high resistance and form part of the blood–aqueous barrier (Raviola, 1977). They contain occludin, ZO-1, and GLUT-1 (Tserentsoodol et al., 1998). Vessels located in the core stroma of the ciliary processes have little resistance and do not contribute to the barrier (Tserentsoodol et al., 1998). They are fenestrated, have noncontinuous TJs, allow penetration of tracers (Raviola, 1977), and contain ZO-1 (Tserentsoodol et al., 1998). Occludin and GLUT-1 are not present in core stromal vessels. Similarly, where the apical margins of the nonpigmented ciliary epithelial cells meet the apical margins of the pigmented ciliary epithelial cells (see Figure 30.5), only ZO-1 is found, but not occludin or GLUT-1, and it is speculated that ZO-1 is involved in holding these two cell layers together in their unusual, apex-to-apex, configuration (Tserentsoodol et al., 1998). Freeze-fracture studies of this apex-to-apex site show discontinuous TJ strands and the absence of true junctional complexes. Tracers pass easily into the intercellular space between these two cell layers (see Figure 30.7) (Raviola, 1977; Freddo, 1987). Gap and adherens junctions appear to be the main membrane specializations at the interface of these epithelia (Raviola, 1977; Raviola and Raviiola, 1978). 30.3.1.3 Tight Junctions in the Pigmented Epithelium of the Iris The iris pigmented epithelium (Figure 30.8) is a continuation of the same cell layer as the nonpigmented ciliary epithelium and constitutes another part of the blood–aqueous barrier. As in its ciliary body counterpart, the posterior pigmented epithelium is also joined apex-to-apex with the anterior myoepithelium of the iris, itself a continuation of the pigmented epithelium of the ciliary body. A comprehensive study by Freddo (1987) described the TJ barrier in the pigmented epithelium of the iris of the rhesus monkey, using tracers, freeze-fracture, and thin-section electron microscopy. Tracer coming from the fenestrated vessels of the ciliary body stroma can pass between individual anterior myoepithelial cells to reach the pigmented epithelium, but cannot flow through the paracellular space between pigmented

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FIGURE 30.8 TJs in the iris. The iris myoepitheliurn lacks TJs and is joined apex-to-apex with the iris pigmented epithelium, which has continuous apical TJs (black ellipses). Blood vessels in the iris stroma also have TJs.

epithelial cells due to the presence of apicolateral TJs. These junctions are a series of branching single or double strands of particles constituting a continuous junctional complex. The complexity of the strands is variable and similar to that reported for the ciliary nonpigmented epithelium, with the majority of TJs containing two to four strands (Raviola, 1977). One possible exception to the similarity to ciliary nonpigmented epithelium should be noted: the iris posterior pigmented epithelium, at least in the mouse eye, has ZO-1 and GLUT-1, but not occludin (Tserentsoodol et al., 1998). Thus, the association of occludin, ZO-1, and GLUT-1 in barrier epithelia may be variable and associated with different functions. 30.3.1.4 Tight Junctions in the Iris Stroma Another section of the blood–aqueous barrier is located in TJs of the vascular endothelial cells of the iris stromal vessels (see Figure 30.8) (Hogan et al., 1971). These junctions have a uniform morphology with two to eight strands, are impermeable to HRP (Raviola, 1974; Freddo and Raviola, 1982; Freddo, 1984), and are positive for ZO-1, occludin, and GLUT-1 (Tserentsoodol et al., 1998), at least in the mouse eye. Freeze-fracture images of animal tissues with experimentally induced inflammation, have shown that tracers can penetrate through the nonpigmented epithelium into the posterior chamber. These studies showed that the TJ strands of the nonpigmented epithelium are still present, albeit fragmented, irregular, and reduced in number (Yata, 1977; Noske et al., 1986; Freddo, 1987). These studies introduced the concept that susceptibility to breakdown of the blood–aqueous barrier may depend on the specific location within the ciliary nonpigmented epithelium and is perhaps related to the variability in TJ complexity mentioned above. In conjunction with similar studies of the perturbation of the blood–aqueous barrier, these results suggest that significant leakage of proteins and other macromolecules into the aqueous humor can occur without a complete disruption of the TJs.

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FIGURE 30.9 TJs in the retina and choroid. The pigmented epithelium of the retina has continuous apical TJs (black ellipses). Although larger vessels in the choroid have TJs, the choriocapillaries have large fenestrations.

30.3.2 BLOOD–RETINAL BARRIER 30.3.2.1 Tight Junctions in the Pigmented Epithelium The retina lines the posterior chamber (see Figure 30.1) and contains the blood–retinal barrier that includes the retinal pigmented epithelium and the nonfenestrated capillaries of the inner retina. The blood–retinal barrier prevents the free movement of water-soluble molecules into the retinal tissues. Consequently, retinal nutrition is dependent on specific transport systems including those that move sugars, neutral and basic amino acids, and monocarboxylic acids (Törnquist et al., 1990). The outermost (i.e., toward the outside of the eye) layer of the retina, known as the retinal pigmented epithelium (Figure 30.9), has a system of annular TJs that presents an impenetrable barrier to large molecules (Hogan et al., 1971; Raviola, 1977; Henkind et al., 1979; Cunha-Vaz, 1980). Like the nonpigmented ciliary epithelium of the blood–aqueous barrier, these cells contain occludin, the GLUT-1 transporter, and ZO-1 in an annular pattern (Takata et al., 1990; 1991; 1997; Konari et al., 1995; Tserentsoodol et al., 1998). Several in vitro models of the blood–retinal barrier have been used to examine the role of TJs in controlling ion flux and permeability. For example, chick embryo RPE cells grown in culture provide a means of examining the relationship between permeability and TJ structure. Occludin and ZO-1, induced by DMSO, form beltlike arrays around the cells which coincides with increased TER and reduced permeability (Konari et al., 1995). Monolayers of cultured bovine retinal endothelial cells also exhibit physiological similarities to the in vivo cells. They have low permeability to inulin, express significant levels of γ-glutamyl transpeptidase and alkaline phosphatase (Gillies et al., 1995), and stain positively for ZO-1 and occludin in a punctate pattern restricted to sites of endothelial cell–cell contacts (Figure 30.10) (Russ et al., 1998). The tight junctions and adherins junctional patterns exhibited by these cells were characterized in a study done by Russ et al. in which human retinal capillary

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FIGURE 30.10 Monolayers of human retinal capillary endothelial cells immunostained for occluden (A), and ZO-1 (B). Arrows indicate no staining in “y”-shaped areas of cell–cell contacts. Scale bar = 20 µm. (From Russ, P. et al., Invest. Ophthalmol. Vis. Sci., 39, 2479, 1998. With permission.)

endothelial cells were immunostained with ZO-1 and occludin (Russ, 1998). The resistance or TER in these cultures is responsive to a variety of stimuli including astrocyte-conditioned medium, which elevates the levels of ZO-1 to 160% of controls and increases the TER from 160 to >200 Ω·cm2. These models may also be useful to investigate the pathogenesis of diseases in which elevated permeability of the retinal microvasculature produces alterations in the function of the retina. 30.3.2.2 Tight Junctions in Nonfenestrated Capillaries Nonfenestrated capillaries in the inner retina are another important component of the blood–retinal barrier. Tracer studies have shown that these vessels have TJs (Shakib and Cunha-Vaz, 1966; Raviola, 1977), ZO-1, occludin, and GLUT-1 (Takata et al., 1991; 1997; Tserentsoodol et al., 1998) and are impermeable to proteins (Shakib and Cunha-Vaz, 1966; Bill et al., 1980; Pino and Thouron, 1983). However, the blood–retinal barrier in rabbits can be breached by intravitreal injections of adenosine agonists or PGE1. These agents cause retinal vessels to open at the TJs, allowing tracer molecules to pass through (Vinores et al., 1992). The finding that PGE1 increases permeability of these barriers suggests that prostaglandins could be mediators of the edema that occurs in diseases like diabetes, in which fluid accumulates in the interstitial spaces, causing the retina to become detached (Campochiaro and Ha, 1989; Kulkarni, 1990). The vascular endothelial cell barrier in the retina plays a significant role in preventing disease. The blood–retinal barrier is violated in the blood vessels of patients with diabetic retinopathy and macular degeneration (Kleine et al., 1995; Gardner et al., 1996). Loss of the barrier causes retinal detachment and fluid accumulation in the subretinal space (Bird, 1989; Törnquist et al., 1990). Moreover, recent in vitro studies suggest that vasoactive agents such as histamine increase paracellular permeability in blood vessels by reducing ZO-1 expression (Gardner, 1995; Gardner et al., 1996), a finding that may be pertinent to conditions such as diabetes, which have prolonged histamine production.

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30.3.2.3 Tight Junctions in Choroidal Vasculature Endothelium Most retinal nutrition is derived from the extensive system of capillaries in the supporting choroidal tissues and those that penetrate the retina itself. As previously mentioned, nutrients pass easily through the fenestrated endothelium of the choroidal capillaries (see Figure 30.9) and into the retinal pigmented epithelial cells where the free flow of nutrients is blocked by the TJs in the apicolateral borders (Raviola, 1977). The choriocapillaries do not have occluding junctions and are negative for occludin and GLUT-1, but positive for ZO-1 (Tserentsoodol et al., 1998). However, choroidal arteriole endothelial cells do have occluding junctions (Raviola, 1977). Tracer studies indicate that permeability of the choroidal vessels supplying the outer retina is highly variable (Peyman et al., 1971). A more recent investigation showed that the complexity of the junctional strands in choroidal vessel endothelial cells in primate eyes is related to the size of the vessel (Nagy and Ogden, 1990). This study showed that the most elaborate strands, made up of single and double rows of particles in complex networks, occurred in the large arteries and that the smaller arterioles had only single rows of particles, some discontinuities and few anastomoses. These observations support the idea that in the choroid, the larger vessels provide a more efficient barrier.

30.4 THE ROLE OF TIGHT JUNCTIONS IN THE DEVELOPMENT OF GLAUCOMA Resistance of the TJ barriers in the endothelial cells that line the aqueous humor may play an important role in regulation of aqueous humor outflow (Underwood et al., 1999). The same drugs that affect resistance and intraocular pressure in humans also increase resistance and TJs in trabecular meshwork and Schlemm’s canal endothelial cells grown in culture (Underwood et al., 1999). Identifying and studying the mechanism by which resistance is induced in the aqueous outflow pathway of the eye has been difficult because there is no really good animal model for glaucoma. In addition, the tiny pieces of tissue available from trabeculectomy specimens or autopsy eyes may be affected by prior drug therapies. Therefore, it has been difficult to determine the gene expression patterns that may lead to glaucoma and, therefore, difficult to identify those genes that may play a role in modulating fluid flow out of the anterior chamber of the eye.

30.4.1 THE AQUEOUS OUTFLOW SYSTEM The aqueous outflow system consists of the channels and tissues through which aqueous humor flows as it exits the eye through the trabecular meshwork and Schlemm’s canal and finally into the venous system (see Figures 30.1 and 30.11). Aqueous humor produced by the ciliary body (aqueous inflow) moves over the lens, below the iris, through the pupillary opening, fills the anterior chamber, and bathes the corneal endothelium (see Figure 30.1, arrows). The rate of aqueous production is ~3 µl/min (Ericson, 1958; Green and Pederson, 1973; Lee et al., 1984; Maus et al.,

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FIGURE 30.11 The aqueous outflow system. Aqueous humor flows through the tortuous, cell-lined channels of the trabecular meshwork, to the blind endings that abut the JXT. For the purpose of illustrating the flow of aqueous, the trabecular channels are somewhat straightened out in this schematic diagram and show the trabecular beams, which are made up of several different types of dense collagens. The aqueous humor passes across the trabecular cells that line these “cul-de-sacs” and the JXT, a region of loosely connected extracellular matrix (crosshatched marks). Aqueous humor then passes through the endothelial cells that line Schlemm’s canal. TJs (black ellipses) have been identified between these cells in situ. However, it is unknown whether the pathway involves only the paracellular route or includes flow through the pores in the giant vacuoles (GV). The pathway through the giant vacuole is indicated by a question mark. Note that flow across the trabecular cells is from the apical to the basal side and flow across the Schlemm’s canal endothelial cells is from the basal to the apical side.

1994). If aqueous drains from the anterior chamber at less than 3 µl/min, the pressure within the chamber (intraocular pressure, or IOP) becomes elevated, which can damage the optic nerve and cause glaucoma.

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30.4.2 POTENTIAL BARRIERS TO THE FLOW OF AQUEOUS HUMOR OUT OF THE EYE A reduction in aqueous outflow is associated with many forms of glaucoma. In angle-closure glaucoma, reduced flow is due to interference from the iris which prevents aqueous from reaching the trabecular meshwork. However, in open-angle glaucoma, there is no obvious barrier and the location of the defect is the source of great controversy (Stamper et al., 1999). Three major tissue sites could present a barrier to aqueous outflow in open-angle glaucomas (see Figure 30.11). The first site is the trabecular meshwork, which consists of a complex series of aqueous channels formed within sheets of connective tissue with densely packed collagen fibrils. These channels are completely lined with a monolayer of endothelial cells known as trabecular meshwork cells. Aqueous flows through the trabecular meshwork channels until it reaches a series of blind endings or “cul de sacs,” also covered by trabecular meshwork cells (Alvarado and Murphy, 1992). It has been suggested that the aqueous must then pass through this cellular barrier, presumably by penetrating the paracellular space (Alvarado and Murphy, 1992). Although the cellular lining of the cul-de-sacs has some discontinuities (estimated to comprise 6% of the cul-de-sac surface area) (Alvarado and Murphy, 1992), it is likely that these trabecular meshwork cells provide a significant barrier to fluid flow. The second barrier to aqueous outflow is a region of loosely woven connective tissue with a random disbursement of cells known as the juxtacanalicular tissue (JXT) (see Figure 30.11). The collagen fibrils in the JXT differ from those present in the trabecular meshwork in that they appear to be random and loosely placed whereas the collagen fibrils that make up the trabecular meshwork channels are tightly packed and dense. The difference in the density of the collagen matrix in the two areas supports the theory that fluid in the trabecular meshwork is limited to the channels and that the reduction in density in the JXT facilitates movement of the aqueous through this region with little resistance. It has been suggested that changes in the content or density of the collagen matrix of the JXT can block flow in open-angle glaucoma (Rohen, 1993; Lutjen-Drecoll et al., 1998; Lutjen-Drecoll, 1999). However, Murphy and others have presented evidence that the juxtacanalicular tissue alone provides insufficient resistance to account for the obstruction to outflow in open-angle glaucoma (Alvarado et al., 1986; Murphy et al., 1992). This issue is still being actively debated (Johnson et al., 1990; Allingham et al., 1992; Stamper et al., 1999). The final barrier in the aqueous outflow pathway is the layer of Schlemm’s canal endothelial cells (see Figure 30.11). The cells that line Schlemm’s canal form a continuous sheet, like that present in nonfenestrated circulatory vessels. These cells are very unusual, forming large outpouchings called “giant vacuoles” with openings or pores through which at least some of the aqueous is thought to exit during normal outflow (Tripathi and Boulpaep, 1989). Similar structures have been found at the site of drainage of cerebrospinal fluid in the arachnoid villus tissue of the brain (Tripathi and Boulpaep, 1989). It is not clear whether the giant vacuoles comprise transcellular pathways (formed by one cell) or paracellular openings (formed by space between adjacent cells) (Epstein and Rohen, 1991) (see question marks in Figure 30.11) Several investigators have estimated from crude measurements in the

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living monkey eye, that only 6% of the normal resistance is provided by the Schlemm’s canal endothelium (Bill, 1975; Törnquist et al., 1990). This result is not surprising, if large volumes of aqueous exit through pores in the giant vacuoles (Allingham et al., 1992; Murphy et al., 1992). Epstein and Rohen (1991) injected cationized ferritin into the anterior chamber of monkey eyes and found that it passed through the trabecular meshwork, through the juxtacanalicular tissue, and accumulated at the Schlemm’s canal endothelium. Although those authors concluded that it is not possible to determine which is the most resistant tissue in the outflow pathway because the experiment is not quantitative, it is interesting to note that the bulk of cationized ferritin binding was at the Schlemm’s canal endothelial cells and not at the trabecular meshwork cells or the juxtacanalicular matrix. These experiments support the possibility that an increase in resistance at the Schlemm’s canal endothelial cell layer could cause a significant reduction in transendothelial fluid flow and result in increased IOP and glaucoma.

30.4.3 TIGHT JUNCTIONS IN TRABECULAR MESHWORK AND SCHLEMM’S CANAL ENDOTHELIAL CELLS Thin-section electron microscopy of the tissues of the aqueous outflow pathway has shown that both trabecular meshwork and Schlemm’s canal endothelial cells have focal TJs (Hogan et al., 1971). Freeze-fracture studies of the human eye support those observations and show that both cell types have some TJ strands. Those in Schlemm’s canal endothelial cells are described as consisting of “continuous tight junctions composed of discontinuous strands that are rarely branched or anastomosed” (Figure 30.12a) (Bhatt et al., 1995) and the strands in trabecular meshwork cells are short or radiating (Bhatt et al., 1995) (Figure 30.12b). Clearly, neither cell type has TJs with the complexity found in most of the blood–ocular barrier tissues.

30.4.4 HORMONAL REGULATION OF TRANSENDOTHELIAL FLUID FLOW Transendothelial fluid flow across primary cultures of trabecular meshwork and Schlemm’s canal endothelial cells grown on permeable filter supports provide a model that simulates the aqueous outflow pathway (Polansky et al., 1984; Perkins et al., 1988; Alvarado et al., 1998; Underwood et al., 1999). Endothelial cells in this model have been shown to alter resistance in response to treatment with drugs and hormones in a manner that is similar to that observed in vivo. For example, β-adrenergic glaucoma medications, which increase aqueous outflow, also increase flow across trabecular meshwork and Schlemm’s canal endothelial cells (Alvarado et al., 1998). In addition, glucocorticoids, which induce steroid glaucoma in susceptible individuals by increasing resistance in the aqueous outflow pathway, increase resistance in trabecular meshwork and Schlemm’s canal endothelial cell barriers in vitro (Underwood et al., 1999). The resistance in steroid glaucoma, as in other types of open-angle glaucoma, is also in the region of the outflow pathway that includes the trabecular meshwork, JXT, and Schlemm’s canal (Epstein, 1986). By using this in vitro model that simulates the aqueous outflow pathway, it has been possible to begin to investigate

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FIGURE 30.12 (A) TJ strands in Schlemm’s canal endothelial cells. Freeze-fracture images of human Schlemm’s canal endothelial cells showing the variable nature of TJ strands. When more than one junctional stand was present along the length of the cell, continuous “mazelike” passages through the junctional array (previously termed slit-pores) were commonly observed (dashed line). Original magnification ×40,838. (Bhatt, 1995). (B) Tight TJ strands in trabecular meshwork cells. Freeze-fracture images of human trabecular meshwork cells showing discontinuous tight junctional strands in Figure 30.2. Junctions of trabecular endothelial cells. The organization pattern of these junctions varied extensively. One variation included isolated undulating strands. The junction again remained complementary from P face (PF) to E face (EF) (arrows). Original magnification ×47,520. (From Bhatt, K. et al., Invest. Ophthalmol. Vis. Sci., 36, 1379, 1995. With permission.)

biochemical and molecular alterations that lead to increased resistance and pathogenesis of steroid glaucoma (Underwood et al., 1999). The TER of cells in this in vitro model is low compared with epithelial cells and even to other types of endothelial cells (Renkin, 1992). Trabecular meshwork cells perfused from the apex to the base of the cell barrier, the normal physiological direction of aqueous flow at the cul-de-sacs (Figure 30.11), have been shown to have hydraulic conductivity values between 2.5 and 4.0 µl/min/mmHg/cm2 (Alvarado et al., 1997). In a recent experiment by the authors, trabecular meshwork cells with hydraulic conductivity values of 3.0 ± 0.5 had TER values of 30.0 ± 2.7 Ω·cm2. Confluent monolayers of Schlemm’s canal endothelial cells also have high transendothelial fluid flow rates but are more resistant than trabecular meshwork cells. The hydraulic conductivity of confluent Schlemm’s canal endothelial cells perfused from the apex to the base has been reported to be 1 to 2 µl/min/mmHg/cm2 (Alvarado et al., 1997). Schlemm’s canal endothelial cells with hydraulic conductivity

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levels of 1.0 µl/min/mmHg/cm2 were determined to have a TER of 47.0 ± 4.5 Ω·cm2. The direction of perfusion in these cells could be highly relevant, since aqueous humor normally flows from the apex to the base in the trabecular meshwork cells, but from the base to the apex as aqueous moves across the Schlemm’s canal endothelial cells (see Figure 30.11). The resistance afforded by this cell layer is very different in the two directions of flow; i.e., cells perfused from the base to the apex have hydraulic conductivity values that are ten times greater than cells perfused from the opposite direction. This differential resistance property of Schlemm’s canal endothelial cells may allow this endothelium to function as a one-way valve for the exit of aqueous humor toward the region of lower pressure in the aqueous veins. Thus, once the fluid passes through the Schlemm’s canal endothelial cells, reflux is unlikely.

30.4.5 TIGHT JUNCTION STRUCTURE ENDOTHELIAL CELLS

IN

SCHLEMM’S CANAL

TJs appear to play an integral role in the maintenance of these trabecular meshwork and Schlemm’s canal endothelial cell barriers. They are present primarily on the fingerlike structures that extend over adjoining cells (Figure 30.13) and form incomplete bands that allow fluid to permeate across a leaky cellular barrier (see Figures 30.12a, b) (Bhatt et al., 1995). ZO-1 is present in both cultured Schlemm’s canal endothelial and trabecular meshwork cells but the expression of ZO-1 is different in the two cell types (Underwood, 1999). Schlemm’s canal endothelial cells express both isoforms of ZO-1, whereas trabecular meshwork cells express only the smaller spliced form (Underwood et al., 1999). The presence of other TJ proteins such as occludin and cingulin has not been demonstrated in these cells, suggesting that the composition of TJ complexes may differ from that in other cell types by more than just the ZO-1 isoforms. Such differences in composition may account for the relative leakiness of these two cell types.

FIGURE 30.13 TJs between two Schlemm’s canal endothelial cells in culture. TJs were routinely located on the overlapping fingerlike extensions of the apical region of the cells. (From Underwood, J. et al., Am. J. Physiol., 277, C330, 1999. With permission.)

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30.4.6 THE RATE OF TRANSENDOTHELIAL FLUID FLOW CORRELATES WITH CHANGES IN TIGHT JUNCTIONS The maintenance of the Schlemm’s canal endothelial and trabecular meshwork cellular barriers is under hormonal regulation (Alvarado et al., 1997; Underwood et al., 1999). Dexamethasone increases resistance in monolayers of Schlemm’s canal endothelial and trabecular meshwork cells by approximately two- to threefold (Underwood et al., 1999). This increased resistance is accompanied by a concomitant increase in the frequency of TJs in both cell types. Approximately 50% of the untreated cells had TJs similar to that shown in Figure 30.13 and treatment with dexamethasone increased this frequency to 80%. In addition, the aggregate length of the occluded portions of the paracellular spaces of cells treated with dexamethasone more than doubled in the Schlemm’s canal endothelial cells and increased by 50% in trabecular meshwork cells (Underwood et al., 1999). ZO-1 was localized to the cell periphery of these cells in punctate formations similar to those observed in astrocytes (Howarth et al., 1992; Underwood et al., 1999). These correlations support the involvement of TJs in the hormonal regulation of transendothelial fluid movement across these cells.

30.4.7 ZO-1 IS ESSENTIAL FOR MAINTENANCE OF TIGHT JUNCTION INTEGRITY AND REGULATION OF TRANSENDOTHELIAL FLUID FLOW To test the hypothesis that TJs and TJ proteins are an important part of the hormonal regulation of fluid movement across Schlemm’s canal endothelial and trabecular meshwork cellular barriers, ZO-1 expression was reduced by treatment with antisense oligonucleotides. The reduction in ZO-1 expression was associated with a decrease in resistance and increased hydraulic conductivity (Figure 30.14). In these cells antisense treatment was begun prior to confluence and prior to the expression of ZO-1 (which normally begins as cells touch and become confluent) (Anderson and Van Itallie, 1995). The increase in hydraulic conductivity above untreated controls and controls treated with the complementary sense sequence showed that development of a normally resistant cellular barrier is dependent on ZO-1 expression. Reducing ZO-1 expression in these Schlemm’s canal endothelial cells did not completely inhibit the normal dexamethasone-induced reduction in hydraulic conductivity, but did prevent the cells from acquiring the same reduced level of flow that the dexamethasone-treated cells attained. The location of the TJs appears to be primarily in the “overlapping finger” regions of these cells (see Figure 30.13). These TJs may modulate or block flow through the paracellular spaces, but their function may be more complex. The rate of aqueous humor flow out of the eye is approximately 3 µl/min, as mentioned above. This relatively large volume of fluid movement may be associated with the formation of interendothelial cell gaps (Baluk et al., 1997). Examination of scanning electron micrographs of dexamethasone-treated cells suggests the possibility that shape changes could cause decreased flow. Figure 30.15 compares untreated control cells, dexamethasone-treated cells, and cells in which ZO-1 expression was blocked with antisense oligonucleotides. A few interendothelial cell gaps are present in untreated control Schlemm’s canal endothelial cells and treatment with dexamethasone causes

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ul/min/mmHg/cm

Hydraulic Conductivity

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DEX

CON+S

DEX+S

CON+A

DEX+A

FIGURE 30.14 Blocking ZO-1 expression increased transendothelial fluid flow (measured as hydraulic conductivity) in Schlemm’s canal endothelial cells. Cells treated with 10–7 M dexamethasone were concomitantly treated with antisense oligonucleotides that block ZO-1 expression for 4 days following seeding on permeable filter supports. (CON = control, DEX = dexamethasone, S = sense oligonucleotides, A = antisense oligonucleotides complementary to ZO-1. Mean and standard deviation, n = 8.) Hydraulic conductivity was measured by gravimetric analysis of media that flowed across the cell monolayers during a 10-min interval at 1 mmHg.

most of these to disappear. However, cells in which ZO-1 was blocked had a significant increase in the number and size of the gaps (four- to fivefold; p < 0.001) (Underwood et al., 1999). These results suggest that disruption of TJs caused a change in cell shape that increased interendothelial cell gaps and flow. Similar shape changes that were associated with increased transendothelial fluid flow have been observed in outflow pathway cells treated with isoproterenol and β-adrenergic agents (Alvarado et al., 1997). This dexamethasone-induced reduction in gap formation may represent an alternative mechanism for regulation of transendothelial fluid flow in Schlemm’s canal endothelial and trabecular meshwork cells.

30.4.8 PATTERNS OF GENE EXPRESSION ASSOCIATED IN TRANSENDOTHELIAL FLUID FLOW

WITH

CHANGES

TJs appear to be an essential part of the way that the body regulates transendothelial fluid flow. These results which correlate flow with TJs suggest a mechanism by which resistance can be increased in the pathogenesis of steroid-induced glaucoma and possibly other types of glaucoma as well. However, the mechanism by which regulation occurs is complex and undoubtedly includes a myriad of other molecules that interact with TJs such as cytoskeletal molecules, adhesion molecules, cytokines, and enzymes that regulate phosphorylation. To identify dexamethasone-induced changes in the pattern of expression of some of these molecules simultaneously, a microarray assay was done on trabecular meshwork cells treated with and without dexamethasone, using an 864-cDNA gene chip (Dr. Ronald Jensen, University of California–San Francisco Cancer Center). Table 30.1 shows how a variety of factors were changed in trabecular meshwork cells treated with 10–7 M dexamethasone.

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FIGURE 30.15 Shape changes in trabecular meshwork cells treated with dexamethasone or antisense oligonucleotides that block ZO-1 expression correspond to changes in transendothelial fluid flow. Confluent monolayers of trabecular meshwork cells were untreated (CON), treated with 5 × 10–7 M dexamethasone (DEX) for 5 days, or treated with 25 µM antisense oligonucleotides, which have been shown to block ZO-1 expression. Examination by scanning electron microscopy showed that control cells had a few small intercellular spaces around the edges of the cells, whereas cells treated with DEX had no obvious intercellular spaces and cells with reduced levels of ZO-1 had many large intercellular spaces. Original Magnification ×9,900.

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TABLE 30.1 Expression Microarray Analysis of Confluent Trabecular Meshwork Cells Treated with 10–7 M Dexamethasone for 7 Days Ratio Dex-Treated Cells/Controls Mean 4.55 3.38 5.0 2.3 1.0 0.5 0.7 0.4 3.85 4.06 0.2 2.99 6.21 9.69 0.60 0.09 2.18 4.51 2.7 0.3 3.79 5.07 1.3 0.35 1.76 5.19 5.27 4.8 4.4 3.2 4.3 7.6 0.43

SD 0.06 2.13 0.3 0.3 0.2 0.3 0.1 0.2 1.97 0.38 0.01 0.37 1.45 4.77 0.14 0.10 0.27 1.07 0.2 0.03 0.57 0.45 0.1 0.1 0.27 2.56 0.45 0.7 0.2 0.1 0.2 2.3 0.25

Gene Fibronectin Vimentin catenin (cadherin-associated protein), alpha 1 (102 kDa) cadherin 2, N-cadherin (neuronal) actin integrin, alpha 3 (antigen CD49C, alpha 3 subunit of VLA-3 receptor) integrin, beta 5 integrin, alpha V (vitronectin receptor, alpha polypeptide, antigen CD51) Collagen, type IV, alpha 4 Collagen, type IX, alpha 3 matrix Gla protein Collagen, type V Topoisomerase 11 Cartilage linking protein 1 Interleukin 3 receptor alpha Interleukin receptor alpha chain stromelysin nitric oxide synthase 3 (endothelial cells) protein kinase C, zeta Protein tyrosine phosphatase Fibroblast growth factor receptor 2 Fibroblast growth factor acidic Fibroblast growth factor 2 (basic) Vascular endothelial Cell Growth Factor Transforming growth factor, beta receptor II (70–80 kDa) Transforming growth factor beta 2 Epidermal growth factor receptor Tumor necrosis factor type 1 receptor associated protein Insulin-like growth factor 2 (somatomedin A) Insulin-like growth factor 2 receptor Corticosteroid binding globulin Corticotropin releasing factor receptor 1 precursor Splicing factor SRp40-1

Note: The mean values are for replicates of three and represent the fold-induction above untreated controls. DEX = dexamethasone. Microarray chips were prepared in the laboratory of Dr. Ronald Jensen, University of California–San Francisco Cancer Center, San Francisco, CA. Hybridization and analysis was done in the laboratory of Dr. Johnnie Underwood, University of California–San Francisco and Department of Surgery, VAMC–San Francisco. (RNA was extracted from cells treated for 7 days with 10–7 M dexamethasone, labeled with either Cy3. RNA from untreated cells was labeled with Cy5-dUTP. The combined labeled RNA was hybridized to the microarray slide in a solution of 25% dimethylformamide and 2X SSC for 12 h at 42°C, washed 4 min in 2X SSC and 0.1% SDS, 4 min in 1X SSC, 4 min in 0.2X SSC, and 10 sec in 0.05X SSC. The slide was scanned using an Axon laser scanner.)

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Dexamethasone induced increases in mRNA of collagen, extracellular matrix molecules, and several cytoskeletal elements. It is interesting to note that some extracellular matrix molecules were increased while others were inhibited. In addition, stromelysin was slightly increased. The significance of these few changes individually may be difficult to interpret but the simultaneous examination of a thousand or many thousands of genes from cells treated with dexamethasone may suggest a pattern of expression that can help predict what factors are involved with the development of resistance. These findings support the work of Clark et al. (1994; 1995) who have shown that changes in cross-linked actin networks are the result of changes in molecules associated with the cytoskeleton and suggested that these shape changes participate in the increased resistance associated with glucocorticoid treatment (Dickerson, 1998). Changes in the expression of type IX collagen and matrix Gla protein are interesting because for many years some investigators have found by electron microscopy that there are changes in the matrix of the juxtacanalicular tissue of patients with glaucoma (Hernandez et al., 1987; Rohen et al., 1993; Rohen and Lutjen-Drecoll, 1996; Johnson et al., 1997; Lutjen-Drecoll et al., 1998; LutjenDrecoll, 1999). The changes in the integrins and cell adhesion molecules could also be involved in the pathogenesis of glaucoma, since reduced cell–cell and cell–matrix adhesions could be related to the trabecular cell loss that has been observed in patients with glaucoma (Alvarado et al., 1981; Epstein and Rohen, 1991; Snyder et al., 1993; Zhou et al., 1998). Regulation of the enzymes associated with phosphorylation of TJ-associated molecules adds another layer of complexity to the problem of understanding the mechanisms that control fluid flow across these endothelial cell barriers. The observation that acidic FGF is induced while basic FGF is not illustrates the complex nature of the cytokine regulation. Regulation of splicing of ZO-1 has also been proposed as a possible regulatory step in control of transendothelial fluid flow and one splicing factor SRp40-1, was significantly decreased by treatment with dexamethasone. It is well known that a variety of cytokines and growth factors change in response to glucocorticoid treatment (Kleinert et al., 1996) and that many of them appear to have a direct effect on transendothelial fluid flow (Woo et al., 1996). The relationship of these factors to regulation of transendothelial fluid flow or regulation of TJs is still being worked out. However, it is interesting to note that VEGF expression, which was inhibited by dexamethasone in these trabecular meshwork cells, is associated with a decrease in occludin and an increase in permeability in retinal endothelial cells (Antonetti et al., 1998). Increased expression of VEGF is also associated with increased permeability in microvascular endothelial cells (Hippenstiel et al., 1998). Thus, the reduced permeability in these dexamethasone-treated trabecular meshwork cells may be mediated by a decrease in VEGF expression.

30.5 CONCLUSION TJs are essential for the maintenance of normal ocular function. TJs in the corneal epithelium protect the eye, and focal TJs in the corneal endothelium play an important role in the pump–leak mechanism that stabilizes the water content of the cornea. The

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ciliary nonpigmented epithelium depends on TJs for the production and one-way flow of aqueous humor. TJs in other components of the blood–aqueous barrier also contribute to the transparency of the aqueous humor. TJs in the blood–retinal barrier maintain the clarity of the transparent retinal tissues. This variety of ocular functions is achieved by the wide range of resistances generated by TJs in specific tissues. TJ regulation of fluid flow across endothelial cells in the aqueous outflow pathway is important because even though their resistance is very low compared with other endothelial and epithelial cell barriers, slight increases in resistance can lead to disease. Steroid glaucoma and other open-angle glaucomas are diseases that cause blindness in millions of people worldwide (Stamper et al., 1999). The pathogenesis of these diseases has remained a mystery partly because of the absence of a good model and partly because of the difficulty, until recently, of obtaining and growing cells from the aqueous outflow pathway. The development of an in vitro model that simulates the outflow pathway, in combination with the new microarray technology, makes it possible to correlate the physiological effects of drugs, hormones, and cytokines on the expression of hundreds or thousands of genes simultaneously. Using this approach it should be possible to identify other factors associated with increased resistance in these endothelial cells and apply that knowledge to a better understanding of the biochemistry that leads to the development of glaucoma.

ACKNOWLEDGMENT The authors thank Dianne Fristrom for her generous help with the drawings.

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31

Implications of Transport via the Paracellular Pathway on Drug Development Philip L. Smith and Chao-Pin Lee

CONTENTS 31.1 Introduction .................................................................................................685 31.2 Case Study: Development of GPIIb/IIIa Receptor Antagonist..................687 31.2.1 Approaches to Enhance Paracellular Permeability .......................688 31.2.2 Dissolution Rate Is Another Important Factor..............................689 31.2.3 Alternative Approaches for Enhancing Absorption ......................689 31.3 Conclusions .................................................................................................691 References..............................................................................................................692

31.1 INTRODUCTION Oral drug delivery remains the preferred route of delivery of therapeutic agents predominantly because of the ease of administration and, hence, patient compliance (Lee et al., 1997). Development of oral dosage forms is complicated by physiological characteristics of the gastrointestinal tract and physical considerations for both the drug and the intestinal route (e.g., regional differences in pH, epithelial and luminal enzymes, transit time within the different segments of the intestine, dissolution rate of the drug, stability of the drug within the environment of the intestine, etc.) (Lee et al., 1997; Smith et al., 1998). In developing an oral dosage formulation, the drug delivery scientist must consider the variety of routes by which a compound can be transported by the intestine and the particular characteristics of the drug, including its physicochemical as well as its pharmacokinetic and pharmacodynamic properties (Smith et al., 1992; Lee et al., 1997). As can be seen from Figure 31.1, transport of drugs across the intestinal epithelium can occur by a variety of mechanisms that exist either within the membranes of the epithelial cells (A, C, D, E, F) or by passive processes between the epithelia cells via the tight junctions (TJs) (B).

0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

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A

B

C

D

E

F

FIGURE 31.1 Routes for the transport of drugs across cellular barriers. (A) passive transcellular; (B) passive paracellular; (C and F) carrier-mediated uptake; (D and E) carrier-mediated efflux.

For hydrophilic compounds, transport across the lipid bilayer of the epithelial cell membrane may be the rate-limiting step in absorption (Thwaites et al., 1993; 1995a). There are a number of hydrophilic compounds that comprise the exceptions to this general statement, including nutrients such as amino acids (Nakanishi et al., 1994; Thwaites et al., 1995b), sugars (Tsuji and Tamai, 1996), and dipeptides (Smith et al., 1996), as well as a number of therapeutic agents including β-lactam antibiotics (Hidalgo et al., 1993; Gochoco et al., 1994), angiotensin-converting enzyme inhibitors (Yuasa et al., 1994), and the antiviral prodrug valine-acyclovir (Balimane et al., 1998; de Vrueh et al., 1998; Han et al., 1998). It has been demonstrated that these therapeutic agents traverse the lipid bilayer of the intestinal epithelial cells via a specific transporter, the peptide transporter, located within the apical cell membrane (Smith et al., 1996). Once taken up across the apical cell membrane, these compounds exit across the basolateral membrane on specific membrane carriers (Matsumoto et al., 1994; Tsang et al., 1994; Tamura et al., 1996). However, there are relatively few hydrophilic compounds that are transported via the transcellular route. For those hydrophilic compounds that cannot cross the epithelial barrier via the transcellular route, the paracellular or junctional pathway (B) is the only alternate pathway that is available to provide sufficient drug to the systemic circulation. For hydrophobic drugs, absorption can occur via a passive transcellular pathway (A). This transport pathway can be very efficient, although recently it has been shown that transporters exist within epithelial and endothelial cells that can transport drugs taken into the cell across the cell membrane back into the external medium (D, E), i.e., “recycle” these compounds back into the bathing solution from which they originated (Schinkel et al., 1996; Sparreboom et al., 1997; van Asperen et al., 1997). Although there is a significant amount of work being directed at these transporters, this chapter focuses on transport via the paracellular pathway. The following sections present an example of the development of a hydrophilic compound whose transport

Implications of Transport via the Paracellular Pathway on Drug Development 687

appears to be confined to the paracellular pathway and the implications for transport via the junctional pathway on the drug development process. This will be followed by a discussion of approaches that have been developed to address issues encountered with drug development including approaches to enhance dissolution, alter intestinal transit, and modulate the junctional pathway to provide a greater opportunity for delivery of hydrophilic compounds including peptides and proteins (Smith et al., 1992).

31.2 CASE STUDY: DEVELOPMENT OF GPIIb/IIIa RECEPTOR ANTAGONIST It has been shown that platelet aggregation and thrombus formation can lead to cardiovascular diseases (Verstraete and Zoldhelyi, 1995). Activation of the platelet and the platelet fibrinogen (GPIIb/IIIa) receptor, with subsequent binding with fibrinogen initiates platelet aggregation (Coller, 1992). Since the GPIIb/IIIa receptor plays a key role in platelet aggregation, blocking the binding of the activated GPIIb/IIIa receptor would be expected to reduce the risk of cardiovascular diseases. Researchers at SmithKline Beecham (SB) have discovered that SK&F 106760, a cyclic peptide, can bind to the GPIIb/IIIa receptor and display antiaggregatory activity in vitro (Nichols et al., 1994a). When tested in vivo, SK&F 106760 completely inhibits platelet-dependent coronary artery thrombosis in the dog following intravenous infusion (Nichols et al., 1994b). However, the bioavailability of SK&F 106760 following oral administration in dogs is low (less than 4%) (Samanen et al., 1996). Therefore, studies were designed to determine the factor(s) responsible for the low bioavailability seen following oral administration of SK&F 106760 (Samanen et al., 1997). These studies were conducted in an attempt to identify approaches to develop formulations to enhance oral absorption and increase systemic delivery of SK&F 106760. In vitro techniques for determining intestinal epithelial permeability of drugs have been established in a number of laboratories (Hidalgo, 1996; Smith, 1996). These in vitro techniques provide several advantages over in vivo studies (e.g., less compound required, ability to determine mechanisms, and higher throughput). However, it should be pointed out that there are important differences between the in vitro models and the in vivo situation. These include the lack of innervation and hence enteric and autonomic nervous system regulation, systemic blood flow, motility, and related factors (Smith, 1996). The authors have established several in vitro systems to study drug transport across the intestinal epithelial barrier. One system is derived from the early work of Ussing and co-workers who developed these techniques for the measurement of ion transport across frog skin (Ussing and Zerahn, 1951; Smith, 1996). With this system, unidirectional transport of drug molecules across segments of intestine in vitro using stripped rabbit intestinal mucosa was measured. Integrity of the intestinal tissues was assessed by concurrent determination of transport of mannitol, a passively transported marker (Dawson, 1977; Marks et al., 1991). The hexapeptide SK&F 110679, a growth hormone-releasing peptide, provides an example of the utility of this in vitro technique for studying absorption (Bowers et al., 1981). SK&F 110679 absorption was determined following intravenous or

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oral administration in rats and was found to be less than 1% (Walker et al., 1990). The mucosal (m)-to-serosal (s) and s-to-m fluxes for SK&F 110679 across rabbit jejunum and ileum are low (less than the corresponding mannitol fluxes) and similar in magnitude (Smith et al., 1994). These results suggest that the absorption of SK&F 110679 occurs by passive diffusion and that the low oral bioavailability for this drug is due to its low intestinal permeability via the paracellular pathway. When evaluated in rabbit ileal tissues, the permeabilities for SK&F 106760 were lower than mannitol, similar to the situation with SK&F 110679 (Samanen et al., 1996). Additionally, the m-to-s and s-to-m fluxes of SK&F 106760 across rabbit ileum were not pH dependent and were identical (Samanen et al., 1996). These results suggest that SK&F 106760 was not transported across rabbit ileum by a peptide transporter (Hidalgo et al., 1993; Gochoco et al., 1994; Smith et al., 1996). These results also indicate that transport of SK&F 106760 across the intestinal mucosa occurs by a passive process that is confined predominantly to the paracellular pathway. Support for this conclusion is provided by the results of an in vivo segmental absorption study in dogs (Nichols et al., 1994b). The bioavailabilities of SK&F 106760 in dogs following intraduodenal and intrajejunal administration were 3 and 6%, respectively. The limited bioavailabilities do not appear to be related to solubility since SK&F 106760 has high aqueous solubility at pH 6 to 8. The intracolonic bioavailability was below the detection limit. Since there is little hydrolysis of peptides in the colon, these results suggest the absorption of SK&F 106760 in the colon is limited and is lower than the absorption in either duodenum or jejunum. Additional support for the proposal that SK&F 106760 is transported across the intestine via the paracellular pathway is provided by the finding that the oral absorption of SK&F 106760 can be enhanced by medium-chain glycerides. Medium-chain glycerides have been demonstrated to increase intestinal permeability for a number of paracellularly transported molecules (Constantinides et al., 1994; Yeh et al., 1994).

31.2.1 APPROACHES

TO

ENHANCE PARACELLULAR PERMEABILITY

The paracellular route for transport accounts for less than 1% of the total intestinal surface area and also restricts passage of compounds via this route based on size, e.g., lisinopril and thyrotropin-releasing hormone (Chadwick et al., 1977; Thwaites et al., 1993; 1995a). Thus, compounds absorbed via this pathway may have low oral bioavailabilities because of the limited surface area for absorption as well as because of the restrictive nature of the paracellular pathway. These observations have prompted the search for safe and effective approaches to enhance the intestinal epithelial permeability of drugs. A variety of approaches have been evaluated including the use of acylcartine, medium-chain glycerides, chitosans, and other membranedisrupting agents (Swenson and Curatolo, 1992; LeCluyse et al., 1993; Artursson et al., 1994; Constantinides et al., 1994; Lindmark et al., 1995; Schipper et al., 1997; 1999; LeCluyse and Sutton, 1997). For many of these approaches, issues related to safety (e.g., intestinal epithelial damage and the potential for simultaneous absorption of toxic agents) and efficacy (e.g., formulation of the enhancer and drug in such a manner that they remain colocalized during transit through the gastrointestinal

Implications of Transport via the Paracellular Pathway on Drug Development 689

tract) have precluded their development into a commercial dosage form. Schipper and co-workers (1999) have recently described the effects of a variety of chitosans with differing molecular weights and degrees of acetylation on the in vitro permeability of atenolol in confluent monolayers of Caco-2 cells and on the absorption of atenolol from perfused rat ileum in vivo. Although chitosan and its derivatives enhanced the permeability of atenolol in Caco-2 cell monolayers, the chitosans did not produce sufficient absorption in vivo to warrant further development. Further studies with the perfused rat model demonstrated that the failure of the chitosans to enhance atenolol absorption was related to mucus secretion by the intestine. Hence, the failure of chitosans to enhance permeability was attributed to the inability of chitosan to bind to the cell membrane in the presence of mucus (Schipper et al., 1999). Thus, although the in vitro results are encouraging, there are a significant number of obstacles to overcome for these approaches to achieve commercial success. Alternative approaches to enhance intestinal permeability have focused on modulation of epithelial barrier function through regulation of existing biochemical pathways, e.g., protein kinase C (PKC)-related polymerization of actin filaments (Fasano et al., 1995; 1997; Fasano and Uzzau, 1997). This topic is more fully addressed in Chapter 32 by Fasano. Thus, there are currently no approaches to enhance epithelial permeability of the intestine on the market.

31.2.2 DISSOLUTION RATE IS ANOTHER IMPORTANT FACTOR In addition to transport across the intestinal mucosa, optimal drug absorption is dependent on dissolution of the compound within the absorption window of the intestine. Therefore, it is important to determine the pH-dependent solubility of the drug of interest. One common approach employed to enhance the dissolution of poorly soluble drugs is to reduce the particle size of the drug substance, thereby increasing the total surface area (Shaw and Carless, 1974; Ridolfo et al., 1979; Shastri et al., 1980). For example, a study was performed to evaluate the effect of particle size reduction on the absorption of digoxin. Reduction of digoxin particle size increased the surface area and presumably the rate of dissolution. The mean particle size of digoxin was reduced from 22 to 12 and 3.7 µm. Reduced particle size preparations of digoxin were dosed to humans. As predicted, results from this study demonstrated more rapid and complete absorption of digoxin with the smaller particle size formulations (Shaw and Carless, 1974). Based on these finding with digoxin, reduction of the particle size of poorly water-soluble drugs may provide improved absorption, especially at higher doses.

31.2.3 ALTERNATIVE APPROACHES

FOR

ENHANCING ABSORPTION

In addition to formulation approaches to enhance the oral bioavailability of the peptide GPIIb/IIIa antagonist, chemists at SB have also developed nonpeptide analogues of SK&F 106760 with the hope that these nonpeptide structures would serve as templates in the design of GPIIb/IIIa antagonists with higher oral bioavailability (Ku et al., 1994). Using the X-ray crystal structure of an analogue of SK&F 106760

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FIGURE 31.2 Nonpeptide analogues of SK&F 106760 employed to evaluate structure–transport relationships for GPIIb/IIIa antagonists.

together with the solution nuclear magnetic resonance (NMR) data for SK&F 106760, medicinal chemists at SB developed SB 207448 (Figure 31.2), a potent nonpeptide GPIIb/IIIa antagonist. SB 207448 contains the benzodiazepine template, an amidino group to mimic the Arg side chain, and a carboxylic acid to mimic the Asp side chain. The in vitro antiaggregatory activity of SB 207448 is comparable with that of SK&F 106760 (Bondinell et al., 1994). However, when SB 207448 was dosed intraduodenally in dogs, no detectable inhibition of ex vivo platelet aggregation was observed, suggesting poor bioavailability presumably due to poor absorption (Samanen et al., 1996). Results from permeability studies using rabbit ileum also suggested that SB 207448 has very low intestinal permeability. These findings led to the conclusion that the oral bioavailability of SB 207448 is limited by its poor intestinal permeability. In vitro permeability screening of benzodiazepine-containing GPIIb/IIIa receptor antagonists was initiated as a strategy for identifying molecules with potential to have good in vivo absorption. A structure–transport relationship was developed based on systematically changing the ileal permeability of the molecule by changing the structure (Lee et al., 1995). In vitro permeability of benzodiazepine-containing GPIIb/IIIa antagonists, such as SB 209751, SB 208433, and SB 209499, across rabbit intestinal mucosa was studied to determine the significance of the charge on the molecule to intestinal permeability (Figure 31.3). The results suggest that using an ethyl ester to eliminate the negative charge on the carboxylic acid at pH 7.4 can enhance the permeability of these benzodiazepine-containing GPIIb/IIIa antagonists by twofold. However, when the amidino group at the amino terminal is removed, permeability increased more than 400-fold. This result indicates that the intestinal permeability of benzodiazepine-containing GPIIb/IIIa antagonists can be improved by modifying the functional groups on these molecules. This may result from a change in the transport route from paracelluar to transcellular as a result of the changes in the molecules, since they have now become more lipophilic. Additional evidence that the oral absorption of nonpeptide GPIIB/IIIa antagonists can be improved by modifying their structures comes from the results of an in vivo study of SB 208651, an N-methyl analogue of SB 207448 (see Figure 31.2) (Samanen et al., 1996). In contrast to the results with SB 207448, intraduodenal

Implications of Transport via the Paracellular Pathway on Drug Development 691

FIGURE 31.3 Structures of GPIIb/IIIa antagonists used to evaluate routes and mechanisms of intestinal absorption.

administration of SB 208651 in dogs yielded approximately 10% bioavailability as determined by ex vivo pharmacological activity. The results from in vitro permeability studies for SB 208651 support the conclusion that the intestinal permeability of SB 208651 is higher than that of SB 207448, which leads to a higher oral absorption and bioavailability.

31.3 CONCLUSIONS The drug development process is resource intensive (Smith, 1997). Low oral bioavailability can increase the complexity of a development program as a result of concerns regarding inter- and intrasubject variability and safety. With low oral bioavailability, the possibility of delivering an inappropriate dose, especially with drugs having a narrow therapeutic margin, is a major concern. Hence, significantly more effort in terms of defining the absorption profile, food effects, drug interactions, and potential for disease to alter absorption can require significantly longer clinical trials and development time. In selecting drug candidates for further development, it is helpful to know the mechanisms of absorption since, as shown in the case of SK&F 106760, transport via the paracellular pathway may limit bioavailability. Thus, the formulation scientist must apply a variety of approaches to understand the mechanisms involved in absorption of a drug candidate to be able to provide a dosage form that will provide optimal dissolution and absorption. In the future, an understanding of the molecular structure of the TJs will help in developing strategies to enhance paracellular drug delivery.

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32

Pathological and Therapeutic Implications of Macromolecule Passage through the Tight Junction Alessio Fasano

CONTENTS 32.1 Introduction .................................................................................................698 32.2 Pathological Conditions Associated with TJ Dysfunction.........................698 32.2.1 TJ Dysfunction in the Blood–Brain Barrier .................................699 32.2.2 TJ Dysfunction in the Intestinal Epithelium.................................701 32.2.2.1 Ankylosing Spondylitis .................................................702 32.2.2.2 Insulin-Dependent Diabetes Mellitus............................703 32.2.2.3 IgA Nephropathy...........................................................703 32.2.2.4 Multiple Sclerosis..........................................................704 32.2.2.5 Celiac Disease ...............................................................704 32.2.2.6 Inflammatory Bowel Diseases.......................................704 32.2.2.7 Conclusion .....................................................................705 32.3 Therapeutic Use of TJ Modulation ............................................................705 32.3.1 Blood–Brain Barrier Delivery .......................................................705 32.3.2 Transmucosal Delivery ..................................................................706 32.3.3 Transdermal Delivery ....................................................................707 32.3.4 Oral Delivery .................................................................................707 32.4 The Zonulin System....................................................................................708 32.4.1 Physiology of the Zonulin System................................................709 32.4.2 Pathology of the Zonulin System..................................................711 32.4.3 Therapeutical Use of the Zonulin System ....................................714 32.5 Conclusions .................................................................................................717 References..............................................................................................................717

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32.1 INTRODUCTION The paracellular route is the dominant pathway for passive solute flow across both epithelial and endothelial barriers, and its permeability depends on the regulation of intercellular tight junctions (TJs), also known as the zonula occludens (ZO) (for more details on the structure and functions of TJs, see chapters 2, 4, and 5). As a barrier between apical and basolateral compartments, TJs selectively control the passive diffusion of ions and small, water-soluble solutes through the paracellular pathway, thereby counterregulating any gradients generated by transcellular pathways (Diamond, 1977). Variations in transepithelial conductance can usually be attributed to changes in the permeability of the paracellular pathway, since the resistance of the cells’ plasma membrane is relatively high (Madara, 1989). The TJ represents the major barrier within this paracellular pathway, and the electrical resistance of epithelial and, to a lesser extent, endothelial tissues seems to depend on the number of transmembrane protein strands and their complexity within the TJ, as observed by freeze-fracture electron microscopy. A century ago, TJs were conceptualized as a secreted extracellular cement forming an absolute and unregulated barrier within the paracellular space (Cereijido, 1992). Biological studies of the past several decades (amply reviewed in this book) have shown that TJs are dynamic structures subjected to structural changes that dictate their functional status under a variety of developmental (Revel and Brown, 1976; Magnuson et al., 1978; Schneeberger et al., 1978), physiological (Gilula et al., 1976; Sardet et al., 1979; Mazariegos et al., 1984; Madara and Pappenheimer, 1987), and pathological circumstances (Milks et al., 1986; Nash et al., 1988; Shasby et al., 1988). To meet the many diverse physiological challenges to which the epithelial and endothelial barriers are subjected, TJs must be capable of rapid and coordinated responses. This requires the presence of a complex regulatory system that orchestrates the state of assembly of the TJ multiprotein network. While knowledge on TJ ultrastructure (see Chapter 2 and Chapters 10 through 12) and intracellular signaling events (Chapters 15 and 17) has significantly progressed during the past decade, relatively little is known about their pathophysiological regulation secondary to extracellular stimuli. Therefore, the intimate pathogenic mechanisms of diseases in which TJs are affected and the utility of TJ modulation for drug delivery both have remained unexplored owing to limited understanding of the extracellular signaling involved in TJ regulation (see also Chapter 16). This chapter reviews the current information on the pathological consequences of an aberrant TJ permeability to macromolecules and the use of these intercellular structures for the delivery of poorly bioavailable drugs, such as proteins and peptides. Further, the discovery of a novel extracellular signaling of TJ regulation, its involvement in disease pathogenesis, and its use for drug and antigen delivery are discussed.

32.2 PATHOLOGICAL CONDITIONS ASSOCIATED WITH TJ DYSFUNCTION TJ dysfunction, primary or secondary to degenerative or inflammatory processes, has been described in several pathological conditions. This chapter specifically

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FIGURE 32.1 BBB model. The BBB is composed of four main structures (from outside to inside): endothelial cell, pericyte, astrocyte, and neuron. Note how the endothelial cells with their TJs represent the only barrier limiting macromolecule flux from the bloodstream to the brain, and vice versa. (From Prokai, L., Prog. Drug Res., 51, 95, 1998. With permission.)

focuses on some of the diseases of either the brain or the intestine in which the TJ defect is described as the primary pathogenetic factor leading to structural and functional changes that characterize these clinical conditions.

32.2.1 TJ DYSFUNCTION

IN THE

BLOOD–BRAIN BARRIER

The blood–brain barrier (BBB) is composed of capillary endothelial cells, microvascular pericytes, astrocyte foot processes, and perivascular microglial cells (Figure 32.1). The intercellular TJ of endothelial cells represents the only anatomical barrier that interfaces between blood and the brain and, therefore, is the limiting structure to the passage of possible harmful substances (including polar and lipidinsoluble moieties such as peptides) from the bloodstream to the central nervous system (CNS). A detailed review of the physiological regulation of the BBB TJ is outlined in Chapter 19. Several diseases are characterized by BBB dysfunction (Partridge, 1998); however, multiple sclerosis (MS) is the most extensively studied pathological condition associated with BBB impairment and, therefore, is the focus of this section. MS is a chronic, inflammatory demyelinizing disorder of the CNS and represents the most common cause of acquired neurological dysfunction in young adults. Although the etiology remains unknown, there is strong circumstantial evidence to support the conclusion that it is mediated by an autoimmune attack directed against CNS myelin, with subsequent formation of scar tissue (plaques) typical of the disease. Focal breakdown of the BBB is a well-established feature of MS and represents the earliest sign of lesion formation (Figure 32.2). It has been shown that trypan blue, a marker that is typically excluded by the BBB, escapes from cerebral

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FIGURE 32.2 Electron micrograph of an inflamed venule from a patient with MS. The curved arrow points to an interendothelial cell TJ that has been split apart. Original magnification ×13,000, inset ×30,000. (From Brosnan, C. F. and Cluadio, L., in Introduction to the Blood–Brain Barrier, Partridge, W. M., Ed., Cambridge University Press, London, 1998. With permission.)

vessels in the vicinity of plaques after death (Broman, 1964). More recently, gadolinium-enhanced magnetic resonance imaging (MRI) has demonstrated that alterations in BBB function precede the onset of clinical signs and demyelinization (Kermode et al., 1990; Davie et al., 1994). Under normal circumstances, the BBB restricts the access of components of the immune system to the CNS compartment. During immune-mediated and inflammatory events, however, the competency of TJ BBB is lost, leukocyte, cytokines, and serum proteins (i.e., complement) invade the CNS (Compston, 1990; Selmaj et al., 1991), and vasogenic edema develops (see Figure 32.2). In these inflamed areas, the TJ has been found permeable to tracers that can be seen to cross throughout the junctional cleft. Thus, it seems that factors released by inflammatory cells can damage the endothelial TJs making them more permeable to vascular solutes and cells that then can reach the CNS parenchyma (see Figure 32.2). In cell culture, this phenomenon has been observed in endothelial cells exposed to tumor necrosis factor-alpha (TNFα) and interferon-gamma (IFNγ) (Brosnan, 1998). In conclusion, the correlation between the clinical presentation of MS and assessment of BBB function suggests that clinical relapse and progressive disease are associated with breakdown of the BBB. These findings may be of pathogenetic importance with respect to the access of myelinotoxic factors to the CNS and may provide the rationale to target endothelial TJ competency for the treatment of this devastating disease.

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32.2.2 TJ DYSFUNCTION

IN THE INTESTINAL

701

EPITHELIUM

The intestinal epithelium is the largest mucosal surface that provides an interface between the external environment and the mammalian host. Under physiological circumstances, the intestine represents the primary site for active transport of fluid and electrolytes from the gut lumen through the transcellular pathway. The paracellular pathway, however, serves as the predominant route for passive transepithelial solute flow. Healthy, mature gut mucosa with its intact TJs serves as the main barrier to the passage of macromolecules. Further, the intestinal barrier functions as the major organ of defense against foreign antigens, toxins, and macromolecules entering the host via the oral/enteric route. During such healthy state, quantitatively small but immunologically significant fractions of antigens cross the defense barrier. These antigens are absorbed across the mucosa along two functional pathways. The vast majority of absorbed proteins (as much as 90%) crosses the intestinal barrier through the transcellular pathway (Figure 32.3), followed by lysosomal degradation that

FIGURE 32.3 Schematic representation of the three transepithelial cellular pathways. (a) Transcellular active transport; (b) transcellular passive transport; (c) paracellular transport. The carrier-mediated, transcellular active transport is mainly limited to small molecules, such as sugars and amino acids, with only a minimal amount of intact proteins crossing the apical membrane by pinocytosis. However, particles absorbed through epithelial or endothelial cells are subject to degradation by lysosomes and, therefore, are less efficiently absorbed. The other two pathways are theoretically available for oral delivery of drugs and vaccines because they do not require the presence of specific carriers for the transepithelial transport of molecules. The paracellular pathway may be used for delivery of drugs and peptides by modulating the permeability of TJs. (From Fasano, A., Trends Biotech., 16, 152, 1998. With permission.)

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converts proteins into smaller, nonimmunogenic peptides. The remaining portion of peptides is transported as intact proteins, resulting in antigen-specific immune responses. This latter phenomenon utilizes the paracellular pathway (see Figure 32.3) that involves a subtle but sophisticated regulation of intercellular TJs that leads to antigen tolerance. When the integrity of the TJ system is compromised, such as during prematurity or exposure to radiation, chemotherapy, and/or toxins (see Chapters 23 and 24), an immune response to environmental antigens (including autoimmune diseases and food allergies) may develop. The specific cells that are important for this immune response lie in close proximity to the luminal antigens and account for as much as 80% of all immunoglobulin-producing cells in the body (Brandtzaeg et al., 1989). Another important factor for intestinal immunological responsiveness is the major histocompatibility complex (MHC). Human leukocyte antigen (HLA) class I and class II genes are located in the MHC on chromosome 6. These genes code for glycoproteins, which bind peptides, and this HLA–peptide complex is recognized by certain T-cell receptors in the intestinal mucosa (Bjorkman et al., 1987; Cuvelier et al., 1987). Susceptibility to at least 50 diseases has been associated with specific HLA class I or class II alleles. This chapter specifically focuses on those clinical conditions in which the involvement of intercellular TJs has been clearly demonstrated. A common denominator of these diseases is the presence of several preexisting conditions leading to an autoimmune process. The first is a genetic susceptibility for the host immune system to recognize, and potentially misinterpret, an environmental antigen presented within the gastrointestinal (GI) tract. Second, the host must be exposed to the antigen. Finally, the antigen must be presented to the GI mucosal immune system following its paracellular passage (normally prevented by the TJ competency) from the intestinal lumen to the gut submucosa (Bjarnason et al., 1986; Wendling et al., 1992) In all cases, increased permeability appears to precede disease and causes an abnormality in antigen delivery that triggers the multiorgan process leading to the autoimmune response. 32.2.2.1 Ankylosing Spondylitis Ankylosing spondylitis (AS) is a common and highly familial rheumatic disorder that typically affects young and middle-aged adults and is characterized by stiffness and pain in the back. The link between increased intestinal permeability and AS has been clearly established (Smith et al., 1985; Casaks et al., 1990; Mielants et al., 1991; Wendling et al., 1992) Using different markers of TJ permeability, two independent studies (Martinez-Gonzales et al., 1994; Vaile et al., 1999) found an increased intestinal permeability in both patients with AS and their relatives. These changes precede the clinical manifestations of the disease (Martinez-Gonzalez et al., 1994; Vaile et al., 1999), suggesting a pathogenic role of TJ dysfunction in AS. Since the vast majority of patients with AS are treated with nonsteroidal anti-inflammatory drugs (NSAIDs), it is conceivable to hypothesize that the TJ incompetency is secondary to the enteropathic effects of the drugs. However, the observations that asymptomatic, nontreated relatives show the same intestinal barrier defect and, more importantly, that the increased permeability affects the small intestine but not the

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gastric mucosa (where NSAIDs exert a significant damaging effect) (Vaile et al., 1999), both suggest that preexisting, genetically determined small bowel TJ permeability abnormalities with subsequent altered antigen exposure, are important factors in the development of AS. 32.2.2.2 Insulin-Dependent Diabetes Mellitus Insulin-dependent diabetes mellitus (IDDM; type I) is an autoimmune condition, sometimes associated with diseases that are characterized by marked immunological features, such as celiac disease (CD) and thyroiditis (Collin et al., 1989; Maki et al., 1995). GI symptoms in diabetes mellitus have been generally ascribed to altered intestinal motility (Feldman and Schiller, 1983) secondary to autonomic neuropathy (Ellenberg, 1963). However, other studies suggest that an increased permeability of intestinal TJs is responsible for both the onset of the disease and the GI symptoms that these patients often experience (Mooradian et al., 1986; Cooper et al., 1987; Carratù et al., 1999) This hypothesis is supported by a recent study performed on a spontaneously diabetic animal model (Meddings et al., 1999). The authors of this study showed an increased permeability of the small intestine (but not of the colon) of BB/Wor diabetic-prone rats that precedes by at least a month the onset of diabetes. Further, histological evidence of pancreatic islet destruction was absent at the time of increased permeability but clearly present at a later time (Meddings et al., 1999). Therefore, those authors presented evidence that increased permeability occurred before either histological or overt manifestation of diabetes in this animal model. Finally, the use of hydrolyzed casein diet significantly reduced the incidence of diabetes without altering the small intestinal permeability, suggesting a diet-independent, genetically determined early event responsible for the opening of intercellular TJs. 32.2.2.3 IgA Nephropathy Immunoglobulin A nephropathy (IgA NP) is the most common form of primary glomerulophritis whose pathogenesis seems related to an immune complex deposit of IgA granules in the glomeruli (Freehally, 1988; Emancipator and Lamb, 1989; Williams, 1993; Emancipator, 1994). Among other mechanisms, the increased absorption of food antigens has been described as a possible etiological cause of the disease (Nagy et al., 1988; Coppo et al., 1990). In both children (Davin et al., 1988) and adults (Rostoker et al., 1993) with IgA NP, increased intestinal permeability, which may cause chronic antigen entry with constant antibody production, has been demonstrated. Kovacs et al. (1996) have demonstrated that patients with IgA NP with increased intestinal TJ permeability showed a significant decrease in creatinine clearance and an increase in clinical manifestations of the disease (i.e., proteinuria and hematuria) as compared with those with a normal permeability, suggesting a pathogenic role of intestinal TJ permeation in IgA NP. It is known from animal experiments that intestinal hyperpermeability, which may be caused by the damaging effect of uremic toxins on the intestinal mucosa, is common in uremia

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(McNair and Olsen, 1974; Sterner et al., 1980). However, human studies showed that a significantly increased intestinal TJ permeability also occurred in nonazotemic patients (Kovacs et al., 1996). This observation confirmed that the increased intestinal TJ permeability is a primary defect and, therefore, may play a pathogenic role in IgA NP independently of the effect of uremia. Further, the deterioration in kidney function in patients with IgA NP with increased intestinal permeability seemed to be faster over a 4-year period compared with patients with IgA NP with normal permeability, suggesting that alteration of intestinal TJ competency may be considered a bad prognostic sign (Kovacs et al., 1996). 32.2.2.4 Multiple Sclerosis In addition to increase in BBB permeability (see above), patients with MS may also experience an increased permeability of intestinal TJs. Yacyshyn and co-workers (1996) have demonstrated that 25% of patients with MS studied had an increased intestinal permeability. Interestingly, 40% of these patients were coincidentally affected by inflammatory bowel diseases (Yacyshyn et al., 1996), clinical conditions in which TJ competency is decreased (see Chapter 26). The fact that patients with MS (Yacyshyn et al., 1996), Crohn’s disease (Yacyshyn and Pilarski, 1993), and AS (Vaile et al., 1999), all present an increased number of peripheral B cells exhibiting CD45RO, a marker of antigen exposure, further supports the concept of preexisting, genetically determined small intestinal permeability abnormalities with subsequent altered antigen exposure as a pathogenic factor common to these diseases. 32.2.2.5 Celiac Disease Celiac disease (CD) is an autoimmune enteropathy triggered by the ingestion of gluten in susceptible individuals (Not et al., 1998). The disease is associated with HLA alleles DQA1*0501/DQB1*0201, and in the continued presence of gluten the disease is self-perpetuating (Sollid and Thorsby, 1993). Although initially thought to be relatively uncommon, it has now been found in 0.4% of the general population (Catassi et al., 1995; Not et al., 1998), often presenting with vague, non-GI symptoms (Hin et al., 1999). During the active phase, a characteristic IgA autoantibody recognizing the endomysium is produced. Early in the disease, TJs are opened (Madara and Trier, 1980; Schulzke et al., 1998) through a recently described mechanism (Fasano et al., 2000) and severe intestinal damage ensues (Schulzke et al., 1998). More-detailed information about this clinical condition is discussed in Chapter 26. 32.2.2.6 Inflammatory Bowel Diseases Crohn’s disease and ulcerative colitis (UC) are inflammatory diseases involving the GI tract in which abnormal paracellular permeability defects precede the development of both syndromes and, therefore, appear to play an important role in disease pathogenesis (Hollander et al., 1986; Yacyshyn and Meddings, 1995; Schmitz et al., 1999). These clinical conditions will not be discussed further, since they have been amply covered in another part of this book (see Chapter 26).

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32.2.2.7 Conclusion In conclusion, there now appears to be a spectrum of diseases that are associated with intestinal presentation of environmental antigens in the context of abnormal GI TJ permeability. The type of disease developed by the host is probably dictated by the genetic background of the host and/or the antigen(s) presented to the mucosal immune system. The abnormal TJ permeability appears to be a common denominator in these diseases and may be responsible for the chronic, pathological passage of luminal antigens to the mucosal immune system, a process possibly involved in the autoimmune etiology characteristic of these syndromes.

32.3 THERAPEUTIC USE OF TJ MODULATION Although the chronic, uncontrolled passage of antigens through biological barriers may be cause of disease, the controlled, reversible passage of macromolecules through intercellular TJs may represent an extremely powerful alternative strategy for the delivery of macromolecules normally not absorbed through these barriers. Theoretically, three transepithelial/endothelial pathways are available for the passage of molecules from one body compartment to another (see Figure 32.3): (1) transcellular (i.e., through the cell) carrier-mediated active or facilitated transport; (2) transcellular passive transport; and (3) paracellular (i.e., between adjacent cells) transport. With the exception of those molecules that are transported by active or facilitated transcellular mechanisms, the absorption of large hydrophilic macromolecules is mainly limited to the paracellular pathway (Lee et al., 1991). Under normal conditions, however, this pathway is restricted to molecules with molecular radii <11Å and, therefore, is not accessible to large compounds. The utility of TJ modulation for drug delivery has remained long unexplored owing to limited understanding of TJ physiology and the lack of substances capable of increasing TJ permeability, without irreversibly compromising either epithelial or endothelial integrity and function (Lee et al., 1991; Citi, 1992; Hochman and Artursson, 1994). This chapter reviews drug delivery systems that utilize the modulation of intercellular TJs as the primary strategy to increase drug bioavailability.

32.3.1 BLOOD–BRAIN BARRIER DELIVERY The microvasculature of the CNS behaves as a continuous lipid bilayer, with intercellular TJs preventing the passage of polar, lipid-insoluble substances such as peptides. Therefore, the competency of the TJs of the BBB represents the major obstacle to the delivery to the CNS of peptide-based drugs that are potentially useful for the treatment of diseases affecting the brain and the spinal cord. Several approaches have been taken to overcome this obstacle, including invasive (Prokai, 1998) and chemical (Begley, 1996) procedures. However, these strategies have been hampered by substantial limitations, including increased risk of infections, discomfort for the patients, and chemical stability of biologically manipulated compounds (Prokai, 1998). Therefore, the alteration of the BBB seems to represent a more acceptable choice. However, lack of information on how this barrier is physiologically regulated leaves the reversible

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disruption of the BBB as the only available strategy. Intracarotid infusion of hypertonic solutions of mannitol, arabinose, lactamide, saline, urea, glycerol, and radiographic contrast agents shrinks the brain capillary endothelial cells, resulting in transient opening of TJs, so facilitating the transport of molecules that otherwise cannot cross the BBB (Prokai, 1998). Vascular permeability to macromolecules transiently increases following infusion of hypertonic mannitol, after which it returns to preinfusion levels within 2 h (Prokai, 1998). Thus far, osmotic modification of the BBB has been studied most commonly as a method for improving brain tumor therapy (Gumerlock, 1992), whereas brain delivery of peptides by this method has not been specifically addressed. However, the considerable toxic effect of this procedure should be taken into account, since this system can lead to inflammation, encephalitis, and to the incidence of seizure in as high as 20% of the procedures (Prokai, 1998).

32.3.2 TRANSMUCOSAL DELIVERY The body contains a variety of mucosal surfaces that have been targeted as potential sites for the noninvasive delivery of macromolecular drugs (Sayani and Chien, 1996), including pulmonary, nasal, buccal, rectal, and vaginal tissues. The transmucosal route has the advantage of bypassing first-pass metabolism; however, it presents several limitations, including the enzymatic barrier (most mucosal surfaces secrete enzymes that would degrade proteins and peptides) and the presence of intercellular TJs, which prevents the passage of large hydrophilic molecules. Thus, enzyme inhibitors and permeation enhancers are usually combined with therapeutic drugs to promote absorption. The pulmonary route of drug delivery is well established in the treatment of lung diseases such as asthma. In recent years, this technology has progressed so that inhalation can be potentially used to deliver protein drugs to the systemic circulation (Yu and Chien, 1997). However, despite the advantage of a significant surface area, this approach suffers the limitation of nonreproducible placement of the drug at the site of absorption in the alveoli. Further, the bioavailability in humans of drugs coadministered with enzyme inhibitors and penetration enhancers affecting the phospholipid bilayer of alveolar cells seems to be relatively low (Adjei and Garren, 1992). Therefore, the need for more efficient lung delivery systems has refocused the attention to the modulation of intercellular TJs as a potential alternative; however, data on this approach are currently scanty (Leone-Bay et al., 2000). The nasal route is probably the most popular alternative to the oral route, because, as for the oral delivery, self-administration is simple and convenient. Also, the nasal mucosa is highly vascularized and provides a fertile surface for drug absorption. As for other mucosal surfaces, the most important limitation to drug delivery via the nasal route is the enzyme degradation and the ability of the intercellular TJs of the nasal mucosa to prevent the transport of any molecule larger than 1 kDa (LeoneBay et al., 2000). Nasal absorption of leuprolide has been demonstrated in healthy humans using cyclodextrans and ethylenediaminetetracetic acid (EDTA), a calcium chelator that destroys TJs (Adjei and Garren, 1992) as delivery agents.

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32.3.3 TRANSDERMAL DELIVERY The skin provides an attractive and readily accessible site for drug delivery, particularly because, like the transmucosal route, it avoids first-pass metabolism. The major limitation of this route is the low penetration rate of substances through the skin and the local irritation caused by the currently used drug delivery systems. Although the use of liposomes and niosomes as delivery agents have been extensively studied (Artmann et al., 1990; du Pleiss et al., 1992), limited data are currently available on the use of TJ modulation for skin delivery (Leone-Bay et al., 2000).

32.3.4 ORAL DELIVERY The oral route represents one of the most attractive and acceptable routes for the administration of therapeutic compounds. The oral delivery of drugs and proteins would avoid the pain and discomfort associated with injections and would also eliminate the possibility of infections caused by the possible reuse of needles. Moreover, oral formulations are less expensive to produce, because they do not need to be manufactured under sterile conditions. Over the past few years, there has been an explosion in research aimed at creating new oral drug-delivery systems. This research has been fueled by unprecedent challenges, such as the need to deliver new, more complex drugs (e.g., proteins, hormones, etc.) that are becoming available through recombinant DNA technology. Thus, considerable attention has been directed at finding ways to increase the intestinal permeability of these compounds (Lee et al., 1991; Fasano, 1998a, b). However, the intestinal absorption of numerous compounds routinely used for the treatment of common diseases is profoundly limited by their physicochemical characteristics. These limitations are the consequence of the function of the GI tract that, by design, is to degrade macromolecules such as proteins to their basic elements (i.e., amino acids) in order to be absorbed. Acid-induced hydrolysis in the stomach, enzymatic activity throughout the GI tract, intestinal peristalsis, and bacterial fermentation in the large intestine, all contribute to the degradation of proteins and peptides. Taken together, these physiological functions and characteristics of the GI tract result in the very low bioavailability of proteic drugs. Therefore, it is not surprising that the oral delivery of macromolecules was thought to be impossible or, at least, extremely difficult. Attempts to increase paracellular transport by loosening intestinal TJs have been hampered by unacceptable side effects induced by the potential absorption-enhancing agents so far tested (Lee et al., 1991; Citi, 1992; Hochman and Artursson, 1994). For the most part, these agents fall into two classes — calcium chelators and surfactants (Hochman and Artursson, 1994). Both have properties that limit their general utility for promoting the absorption of various molecules. In the case of calcium chelators, Ca2+ depletion induces global changes in the cells, including disruption of actin filaments, disruption of adherent junctions, and diminished cell adhesion (Citi, 1992). In the case of surfactants, the potential lytic nature of these agents may cause exfoliation of the intestinal epithelium, irreversibly compromising its barrier functions (Hochman and Artursson, 1994). Chitosan, a mucoadhesive

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polymer, has been recently described as a potentially valid alternative (Dodane et al., 1999; Thanou et al., 2000). Using Caco-2 monolayers as an in vitro model of the intestinal barrier, Dodane et al. (1999) have shown that chitosan caused a reversible, time and dose-dependent decrease in transepithelial electrical resistance. The effect of chitosan on TJs was confirmed by an increased permeability to mannitol and the displacement from the junctional complex of the tight junctional proteins ZO-1 and occludin (Dodane et al., 1999). However, chitosan treatment appeared to perturb the cell plasma membrane as demonstrated by the increased release of lactate dehydrogenase (Dodane et al., 1999). Further, at physiological pH, chitosan is insoluble and, therefore, ineffective (Thanou et al., 2000). This limitation can be overcome by using N-trimethyl chitosan chloride (TMC), a permanently quaternized derivative with improved aqueous solubility compared with native chitosan (Thanou et al., 2000). However, high charge density was necessary for TMC to improve the TJ permeability substantially in CaCo2 monolayers (Thanou et al., 2000).

32.4 THE ZONULIN SYSTEM In recent years much has been discovered about the structure, function, and regulation of TJs. However, the precise mechanism(s) through which they operate are still incompletely understood. Several microorganisms have been shown to exert a cytophatic, pathological effect on epithelial cells that involves the cytoskeletal structure and TJ function in an irreversible manner. These bacteria alter the intestinal permeability either directly (i.e., EPEC) or through the elaboration of toxins (i.e., Clostridium difficile, Bacteroides fragilis; for a more complete review see Chapter 23). A more physiological mechanism of regulation of TJ permeability has been proposed for the zonula occludens toxin (Zot) elaborated by Vibrio cholerae (Fasano et al., 1991; Baudry et al., 1991). The author’s group has recently reported that Zot possesses multiple domains that allow a dual function of the protein as a morphogenetic phage peptide for the V. cholerae phage CTXφ and as an enterotoxin that modulates intestinal TJs (Uzzau et al., 1999). The discovery of Zot has shed some light on the intricate mechanisms involved in the modulation of the intestinal paracellular pathway. Zot action is mediated by a cascade of intracellular events that lead to a protein kinase C (PKC)-dependent polymerization of actin microfilaments strategically localized to regulate the paracellular pathway (Figure 32.4) (Fasano et al., 1995). These changes are a prerequisite to opening of TJs and are evident at a toxin concentration as low as 1.1 × 10–13 M (Fasano et al., 1995). The toxin exerts its effect by interacting with the surface of enteric cells. By using immunofluorescence binding studies, the author’s group has shown that Zot binding varies within the intestine, being detectable in the jejunum and distal ileum, but not in the colon, and decreasing along the villous–crypt axis (Fasano, 1997). This binding distribution coincides with the regional effect of Zot on intestinal permeability (Fasano et al., 1997) and with the preferential F-actin redistribution induced by Zot in the mature cells of the villi (Fasano et al., 1995), suggesting that the regional effect of Zot is associated with its binding of a surface receptor or receptors whose distribution varies within the intestine and along the villous–crypt axis. These data also suggest that the expression of this receptor(s) is upregulated during enterocyte differentiation.

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This hypothesis is supported by the observation that human intestinal epithelial CaCo2 cells (which resemble the mature absorptive enteric cell of the villi), but not cryptlike T84 cells, express this receptor(s) on their surface (Uzzau et al., 2001). The paucity of Zot binding in the crypt area may also reflect the fact that this region is already leaky as compared with the more mature epithelium of the tip of the villi (Marcial et al., 1984) and thus might not need to express a significant amount of a putative receptor(s) involved in TJ regulation. Taken together, these data showed that Zot regulates TJs in a rapid, reversible, and reproducible fashion, and probably activates intracellular signals that are operative during the physiological modulation of the paracellular pathway (see Figure 32.4). Based on this observation, it is postulated that Zot may mimic the effect of a functionally and immunologically related endogenous modulator of epithelial TJs. The combination of affinity-purified anti-Zot antibodies and the Ussing chamber assay allowed identification of an intestinal Zot analogue that Wang et al. named zonulin (Wang et al., 2000). When zonulin was studied in a nonhuman primate model, it reversibly opened intestinal TJs. Since V. cholerae infections are strictly confined to the GI tract, an exclusively intraintestinal role for zonulin was anticipated. Surprisingly, this protein(s) was detected in a wide range of extraintestinal tissues and has now been purified from human intestine, heart, and brain (Wang et al., 2000). This author’s group has provided evidence that the zonulins comprise a family of tissue-specific regulators of TJs (Fasano, 2000). Each family member has a molecular weight of 47 kDa, a distinct N-terminal receptor-binding motif that confers tissue specificity, and a C-terminal domain probably involved in the rearrangement of cytoskeletal elements functionally connected to intercellular TJs. Amino acid substitution within the N-terminal binding motif identified three amino acid residues that dictate tissue specificity (Di Pierro, 2000). On the basis of this widespread tissue distribution, it is unclear whether or not the zonulin system exerts a systemic and/or a local permeabilizing control on endothelial and epithelial cells. However, the observations that human tissues other than the intestine express zonulin receptors (Lu et al., 2000), that zonulin receptors purified from different human organs have distinct N termini, (Lu et al., 2000), and that the zonulin family members exert a tissue-specific effect on TJ permeability (Fasano, 2000), all support paracrine/autocrine regulation. The zonulin receptor family might provide regional tissue responsiveness to a given stimulus on the basis of local requirements. This restriction of TJ disassembly to a specific organ system could prevent deleterious consequences in other anatomical sites (e.g., intestine vs. BBB).

32.4.1 PHYSIOLOGY

OF THE

ZONULIN SYSTEM

The physiological role of the zonulin system remains to be established; however, it is likely that this system is involved in several functions, including TJ regulation responsible for the movement of fluid, macromolecules, and leukocytes between body compartments. Another possible physiological role of the zonulins is, at least in the intestine, the protection against microorganism colonization (El Asmar et al., 2000). In the absence of enteric infections, the mammalian small intestine is virtually sterile. The colonization of the proximal gut by enteric microorganisms (even without

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FIGURE 32.4

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apparent mucosal damage or elaboration of specific toxins) may lead to a leaky intestine (Isolauri et al., 1986); however, the mechanism(s) by which this disturbed physiological regulation of the intestinal TJ permeability secondary to proximal bacterial contamination occurs remains unclear. The author’s group has recently provided evidence that both normal enteric bacterial flora isolates (well characterized for not harboring any known pathogenic traits) and pathogenic bacteria induce alteration of TJ competency, as suggested by changes in epithelial resistance and increased passage of inulin (El Asmar et al., 2000). These changes were mirrored by the concomitant expression of zonulin in organ culture systems and occurred even when bacteria were killed by gentamicin treatment (El Asmar et al., 2000). These results suggest that the presence of enteric microorganisms in the small intestine (but not in the colon, where the zonulin system is not operative; Fasano, 1991; 1995, 1997) induces a host-dependent mucosal response that leads to the luminal secretion of zonulin. The role of zonulin in the bacteria-induced impairment of the intestinal barrier function is sustained by the observation that zonulin is detected in organ culture supernatants only when exposed to bacteria and by the blocking effect of zonulin inhibitors on these intestinal barrier changes (El Asmar et al., 2000). The fact that the interaction of bacteria with the intestinal mucosa induces zonulin release, irrespective of their pathogenic traits or viability, can be interpreted as a bacteria-independent mechanism of defense of the host that reacts to the abnormal presence of microorganisms on the surface of the small intestine. Following the zonulin-induced opening of TJs, water is secreted into the intestinal lumen following hydrostatic pressure gradients (Fasano et al., 1997) and bacteria are “flushed out” from the small intestine.

32.4.2 PATHOLOGY

OF THE

ZONULIN SYSTEM

Given the complexity of both cell-signaling events and intracellular structures involved in the zonulin system, it is not surprising that this pathway may be affected when the physiological state of epithelial and/or endothelial cells is dramatically changed, as it is in many of the autoimmune diseases in which TJ dysfunction appears to be the primary defect (see above). To explore this possibility, the author’s group focused its studies on CD and IDDM, two autoimmune conditions in which the finely tuned regulation of intestinal TJ permeability is lost (see above). FIGURE 32.4 Proposed zonulin/Zot intracellular signaling leading to the opening of intestinal TJs. Zonulin and Zot interact with the same specific surface receptor (1) whose distribution within the intestine varies. The proteins are then internalized and activate phospholipase C (2) that hydrolyzes phosphatidyl inositol (3) to release inositol 1,4,5-tris-phosphate (PPI-3) and diacylglycerol (DAG) (4). PKCα is then activated (5), either directly (via DAG) (4) or through the release of intracellular Ca2+ (via PPI 3) (4a). PKCα catalyzes the phosphorylation of target protein(s), with subsequent polymerization of soluble G-actin in F-actin (7). This polymerization causes the rearrangement of the filaments of actin and the subsequent displacement of proteins (including ZO-1) from the junctional complex (8). As a result, intestinal TJs become looser. (From Fasano, A., Trends Biotech., 16, 152, 1998. With permission, modified.)

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FIGURE 32.5 In situ zonulin expression in intestinal biopsies from patients with CD and controls. Zonulin expression in intestinal tissues was evaluated by immunofluorescence microscopy of tissue sections incubated with affinity-purified anti-Zot antibodies followed by FITC-conjugated goat antirabbit antibody. Zonulin staining was increased in tissues from patients with acute CD (right panel) compared with the controls (left panel). This increased zonulin expression consistently appeared in a diffuse reticular pattern (see arrows) that was most evident toward the tip of the villi (V), progressively decreasing along the villous–crypt axis, and disappearing in the crypt region (C). The preferential zonulin overexpression in the villi coincides with the distribution of the zonulin/Zot receptor within the villous–crypt axis (Fasano, 1995; 1997). Original magnification ×20.

To determine whether zonulin is perturbed during the acute phase of CD, intestinal tissues from patients with active CD and non-CD controls were probed for zonulin expression (Fasano et al., 2000). Immunofluorescence analysis of CD tissues revealed increased zonulin expression within the intestinal submucosa with a characteristic reticular pattern that was consistently absent in control tissues (Figure 32.5). Quantitative immunoblotting of intestinal tissue lysates from patients with active CD confirmed increased zonulin protein compared with control tissues (Fasano et al., 2000). Zonulin upregulation was further substantiated by testing the serum zonulin level in a large number of patients with CD (T. Not and A. Fasano, unpublished). The increased expression of zonulin in the face of TJ disassembly might permit zonulin presentation to the submucosal gut immune system. Accordingly, an ELISA was used to detect antizonulin antibodies in the sera of patients with CD and controls. Antizonulin IgG was not increased in patients with CD compared with controls. In contrast, antizonulin IgA was elevated in the sera of 25 of 117 (21.4%) patients with CD during the acute phase of the disease but in none of the 30 patients in remission (Fasano et al., 2000). Only 9 of 163 (5.5%) healthy subjects had a minimally but significantly elevated antizonulin IgA titer (Fasano et al., 2000). The incidence of antizonulin antibodies during the acute phase of CD is consistent with the incidence of other autoantibodies described in CD (Ventura et al., 1999). There was a significant correlation between serum levels of zonulin and antizonulin IgA antibodies (Figure 32.6). In seven patients with CD followed longitudinally, the elevated antizonulin IgA returned to normal after 3 to

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FIGURE 32.6 Correlation between serum zonulin levels and circulating antizonulin antibodies. A clear correlation between zonulin serum levels and antizonulin IgA antibody titers was found in sera of patients with CD during the acute phase of the disease.

6 months asymptomatic remission on a gluten-free diet (Fasano et al., 2000). The TJ derangements occurring in CD are more pronounced in villous enterocytes (Schulzke et al., 1998) and, therefore, coincide with the zonulin receptor distribution along the GI tract (Fasano et al., 1997). These findings, together with the observation that zonulin is overexpressed during the acute phase of CD, suggest that this protein contributes to CD pathogenesis by increasing TJ permeability typical of the early stages of this clinical condition. It has been recently reported that untreated CD predisposes to autoimmune disorders such as IDDM, Hashimoto’s thyroiditis, autoimmune hepatitis, and connective tissue diseases (Ventura et al., 1999). One could hypothesize that zonulin opens small intestinal TJs during the early stage of CD, and permits entry of putative allergens into the intestinal submucosa where an autoimmune response is elicited. Alterations in intestinal TJ permeability have also been shown to be one of the preceding pathophysiological changes associated with the onset of IDDM (Mooradian et al., 1986; Cooper et al., 1987; Carratù et al., 1999; Meddings et al., 1999). To establish whether zonulin may be responsible, at least in part, for these early changes, an established rat model of IDDM was used (Fasano et al., 1997; Meddings et al., 1999). Two genetic breeds, i.e., BB/Wor diabetic-prone and diabetic-resistant rats, were studied to determine whether they exhibited significant changes in intraluminal secretion of zonulin and intestinal permeability. No difference in intraluminal zonulin concentration was observed in these two groups of animals before age 40 days (Watts et al., 2000b). Thereafter, a fourfold increase in intraluminal zonulin was observed in diabetic-prone rats, whereas no significant increment was detected in diabetic-resistant animals (Watts et al., 2000b). This increase in intraluminal zonulin was found: 1. To be age-related; 2. To be detectable only in the small intestine (i.e., jejunum and ileum), but not in the colon;

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3. 4. 5. 6.

To correlate with an increase in intestinal permeability of the intestine; To precede the onset of diabetes by at least 3 to 4 weeks; To remain high in these diabetic-prone rats; and To correlate with the progression toward full-blown diabetes (Watts et al., 2000b).

Of note, those diabetic-prone rats that did not have an increase in intraluminal zonulin did not progress to diabetes (Watts et al., 2000b). Thus, these results suggest that zonulin can be responsible for the early permeability changes (and, therefore, for the pathogenesis of IDDM) already described in this animal model (Meddings et al., 1999) and confirmed by these studies. This hypothesis is further supported by the observation from both Watts et al. (2000b) and other investigators (Meddings et al., 1999) that the intestinal permeability changes in these diabetic rats are confined to the small intestine, paralleling the regional distribution of the zonulin intestinal receptor (Fasano et al., 1997) and, therefore, of the site in which the zonulin system is operative.

32.4.3 THERAPEUTICAL USE

OF THE

ZONULIN SYSTEM

Considering the limitations of the TJ modulators currently tested for drug delivery (see above), it was reasonable to explore whether findings from basic research on the zonulin system could be applied to developing new approaches to the enhancement of drug absorption via the regulation of intercellular TJs. So far, the main source of zonulin remains its biochemical purification from human cadavers. Therefore, Zot was used as a valid alternative to utilize the zonulin system for drug delivery, since both proteins share the same binding motif to engage to the zonulin receptor (Di Pierro et al., 2001) and, therefore, activate the same intracellular signaling leading to the modulation of intercellular TJs (Wang et al., 2000). Since Zot represents the prokaryotic analogue of zonulin, it displays multiple characteristics that might make it a promising tool to enhance drug and peptide transport through epithelial mucosa. First, Zot is not cytotoxic and does not affect the viability of the intestinal epithelium ex vivo (Fasano et al., 1991; 1997); second, it does not completely abolish the intestinal transepithelial resistance (Fasano et al., 1991; 1995; 1997); third, it interacts with a specific intestinal receptor whose regional distribution within the intestine varies; fourth, Zot is not effective in the large intestine, where the presence of the colonic microflora can be potentially harmful if the mucosal barrier is compromised (Fasano and Uzzau, 1997; Fasano et al., 1997); fifth, it does not induce acute or chronic systemic side effects when orally administered (Fasano et al., 1997); and, finally, it induces a reversible increase in tissue permeability (Fasano et al., 1991; 1995; 1997). To establish the efficacy of Zot as an intestinal-absorption enhancer, insulin and IgG were selected as drugs to be orally delivered. This choice was based on the relative size, structure, biological activities, and therapeutic relevance of these proteins. In vitro experiments in the rabbit ileum demonstrated that Zot (1.1 × 10–10 M) reversibly increases the intestinal absorption of both insulin (by 72%) and IgG (by 52%) in a time-dependent manner (Fasano and Uzzau, 1997). The permeabilizing

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effect peaked at 80 min and was completely reversible within 20 min after the withdrawal of the molecule. Similar results were obtained when zonulin was used as an insulin delivery enhancer (A. Fasano, unpublished). When tested in the intact host by using the rabbit in vivo perfusion assay, Zot increased the passage of insulin across both the jejunum and distal ileum tenfold, whereas no substantial changes were observed in the colon (Fasano and Uzzau, 1997). Similar results were obtained with IgG, for which Zot induced twofold and sixfold increases in IgG absorption in the jejunum and ileum, respectively. Again, no increases in absorption were detected in the colon (Fasano and Uzzau, 1997). To evaluate the bioactivity of insulin after enteric coadministration with Zot, the hormone was orally administered to acute type-1-diabetic male rats with or without Zot, and the blood glucose levels of the rats were serially measured. After oral administration of insulin alone, given at doses between 5 and 30 IU, blood-glucose levels of treated animals were not appreciably lowered (Fasano and Uzzau, 1997). By contrast, when insulin (at doses as low as 10 IU) was orally coadministered with Zot, a significant reduction in blood-glucose concentration was observed (Figure 32.7). This decrement was comparable with that seen with a conventional

Blood glucose decrement (mg/dl)

75 25 -25 -75 -125 -175 -225 0

60

120

180

240

300

360

Time (min) FIGURE 32.7 In vivo effect of Zot on insulin oral delivery in an animal model of diabetes. Effect of oral insulin (10 IU), alone () or in the presence of Zot (5 µg) (▫) on serum glucose in diabetic rats. The coadministration of Zot induced a reduction in blood-glucose concentration comparable with that seen with a conventional dose of insulin subcutaneously administered (●), and the blood-glucose levels returned normal by 6 h after administration. Bloodglucose levels of untreated animals () and animals treated with oral Zot alone (∆) are shown for comparison. These results would anticipate a relative oral insulin bioavailability of 20% when coadministered with Zot. (From Fasano, A. and Uzzau, S. J. Clin. Invest., 99, 1158, 1997. With permission.)

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dose of insulin subcutaneously administered (range 1.2 to 2.4 IU); the blood-glucose level returned to normal within 6 h after administration (Figure 32.7). None of the animals treated with insulin + Zot experienced diarrhea, fever, or other systemic symptoms, and no structural changes could be demonstrated in the small intestine on histological examination (Fasano and Uzzau, 1997). These results demonstrate that coadministration of Zot with biologically active ingredients enhances intestinal absorption of the active molecule, and that this enhancement is effective for both relatively small (5.7 kDa: insulin) and large (140 to 160 kDa: IgG) molecules (see Figure 32.4). Furthermore, experiments in diabetic rats demonstrate that orally-delivered insulin retains its biological activity without provoking severe hypoglycemia within the range of the insulin administered to the animals, that is, up to 15 times more than the effective parenteral insulin dose. These findings might have important practical implications, since the insulin therapeutic index (i.e., the ratio between the median toxic dose and the median therapeutic dose) of insulin is relatively low. To establish whether the zonulin system is operative also in primates, both in vitro and in vivo experiments were conducted. Initially, different intestinal segments of healthy rhesus monkeys (Macaca mulatta), including jejunum, ileum, and colon, were mounted in Ussing chambers and exposed to purified Zot (10–9 M) added to the mucosal side of the tissue. Zot induced a significant reduction of Rt in the jejunum and distal ileum, while the colon remained unaffected (Wang et al., 2000). This effect occurred within 60 min of the addition of Zot, reached its plateau after 2 h, and was readily reversible once Zot was withdrawn from the reservoir of the chamber. Based on these results, the author’s group explored the passage of peptide molecules through the monkey small intestines in vitro mounted in Ussing chambers. Both 1deamino-8-D-arginine vasopressin (desmopressin) (Lundin and Fasano, personal communication) and insulin (Watts et al., 2000a) were used as markers to investigate further Zot as a regulator of intestinal permeability in primates. The coadministration of Zot significantly increased the apparent permeation coefficients of desmopressin in the ileum and, to a lesser degree, in the jejunum. Similarly, Zot increased monkey intestinal absorption of insulin compared with the untreated control in a timedependent manner both in the jejunum and in the ileum (Watts et al., 2000a). Zot was also tested in an in vivo primate model of diabetes mellitus. Insulin was intragastrically administered to diabetic monkeys either alone or in combination with increasing amounts of Zot. Oral coadministration of Zot and insulin decreased the blood-glucose levels in a dose-dependent manner. Measurements of blood-insulin levels revealed that insulin bioavailability increased from 5.4% in controls to 10.7 and 18% when Zot 2 and 4 µg/k were coadministered, respectively (Watts et al., 2000a). Combined together, these results demonstrate that the permeabilizing effect of the zonulin system also occurs in primates and that the kinetics and the regional effects of this system are similar to those observed in the rabbit model. The use of the zonulin system for macromolecules delivery was also explored in districts other than the intestine (Di Tommaso et al., 1999). Intranasal immunization of mice with either ovalbumin (Di Tommaso et al., 1999) or tetanus toxoid (TT) (M. T. De Magistris and A. Fasano, unpublished) induced a mucosal-specific

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immune response that strongly increased when these molecules were coadministered with Zot. When mice were challenged with tetanus toxin, 100% immunoprotection was observed in animals previously immunized with Zot + TT, whereas 0% survival was recorded among mice that received only TT (M.T. De Magistris and A. Fasano, unpublished).

32.5 CONCLUSIONS The paracellular pathway was once considered to be exclusively the route for passive, unregulated passage of water, electrolytes, and small molecules. Its contribution to the general economy of transepithelial transports was, therefore, judged to be simply secondary to the active, transcellular transport processes. It is now becoming apparent that the elements that govern this pathway; i.e., the TJs, are extremely dynamic structures involved in developmental, physiological, and pathological circumstances. An increased number of autoimmune diseases are now described whose pathogenesis is associated with a primary dysfunction of intercellular TJs. These same structures, however, are used to develop innovative strategies for the delivery of macromolecules normally not absorbed through biological barriers. The discovery of the zonulin system has shed some light on the intricate pathophysiological regulation of intercellular TJs that, however, remains far from being completely addressed. It is conceivable that the zonulins participate in the physiological regulation of intercellular TJs not only in the small intestine, but also throughout a wide range of extraintestinal epithelia (e.g., the tracheobronchial tree and the renal tubule) as well as the ubiquitous vascular endothelium, including the BBB. Dysregulation of this conceptual zonulin model may contribute to disease states that involve disordered intercellular communication, including developmental and intestinal disorders leading to autoimmune disease (i.e., CD and IDDM), tissue inflammation, malignant transformation, and metastasis. This same system can offer the opportunity of targeted, tissue-specific delivery of macromolecules and drugs currently engineered by recombinant DNA techniques or that will become available through the Human Genome Project.

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Nash, S., Stafford, J., and Madara, J. L. 1988. The selective and superoxide-independent disruption of intestinal epithelial tight junctions during leukocyte transmigration, Lab. Invest., 59, 531. Not, T., Horvath, K., Hill, I. D., Partanen, J., Hammed, A., Magazzu, G., and Fasano, A. 1998. Celiac disease risk in the USA: high prevalence of antiendomysium antibodies in healthy blood donors, Scand. J. Gastroenterol., 33, 494. Partridge, W. M. 1998. Introduction to the Blood–Brain Barrier, Cambridge University Press, Cambridge, UK. Prokai, L. 1998. Peptide drug delivery into the central nervous system, Prog. Drug Res., 51, 95. Revel, J. P. and Brown, S. S. 1976. Cell junctions in development with particular reference to the neural tube, Cold Spring Harbor Symp. Quant. Biol., 40, 443. Rostoker, G., Wirquin, V., Terzidis, H., Petit-Phar, M., Chaumette, M. T., Delchier, J. C., Belghitis, D., Lang, P., Dubert, J. M., Meignan, M., Lagrue, G., and Weil, B. 1993. Mucosal immunity in primary glomerulonephritis, III. Study of intestinal permeability, Nephron, 63, 286. Sardet, C., Pisam, M., and Maetz, J. 1979. The surface epithelium of teleostean fish gills: cellular and junctional adaptations of the chloride cell in relation to salt adaptation, J. Cell Biol., 80, 96. Sayani, A. P. and Chien, Y. W. 1996. Systemic delivery of peptides and proteins across absorptive mucosae, Crit. Rev. Ther. Drug Carrier Syst., 13, 85. Schmitz, H., Barmeyer, C., Fromm, M., Runkel, N., Foss, H. D., Bentzel, C. J., Riecken, E. O., and Schulzke, J. D. 1999. Altered tight junction structure contributes to the impaired epithelial barrier function in ulcerative colitis, Gastroenterology, 116, 301. Schneeberger, E. E., Walters, D. V., and Olver, R. E. 1978. Development of intercellular junctions in the pulmonary epithelium of the foetal lamb, J. Cell Sci., 32, 307. Schulzke, J. D., Bentzel, C. J., Schulzke, I., Riecken, E. O., and Fromm, M. 1998. Epithelial tight junction structure in the jejunum of children with acute and treated celiac sprue, Pediatr. Res., 43, 435. Selmaj, K., Raine, C. S., Cannella, B., and Brosnan, C. F. 1991. Identification of lymphotoxin and tumor necrosis factor in multiple sclerosis lesions, J. Clin. Invest., 87, 949. Shasby, D. M., Winter, M., and Shasby, S. S. 1988. Oxidants and conductance of cultured epithelial cell monolayers: inositol phospholipid hydrolysis, Am. J. Physiol., 255, 24, C781. Smith, M. D., Gibson, R. A., and Brooks, P. M. 1985. Abnormal bowel permeability in ankylosing spondylitis and rheumatoid arthritis, J. Rheumatol., 12, 299. Sollid, L. and Thorsby, E. 1993. HLA susceptibility genes in celiac disease: genetic mapping and role in pathogenesis, Gastroenterology, 105, 910. Sterner, G., Asp, N., Dahlquist, A., Denneberg, T., and Lindberg, T. 1980. Small intestinal dipeptidases and disaccharidases in experimental uremia in rats, Nephron, 26, 149. Thanou, M. M., Kotzé, A. F., Scharringhausen, T., Luessen, H. L., de Boer, A. G., Verhoef, J. C., and Junginger, H. E. 2000. Effect of degree of quaternization of N-trimethyl chitosan chloride for enhanced transport of hydrophilic compounds across intestinal Caco-2 cell monolayers, J. Controlled Release, 64, 15. Uzzau, S., Cappuccinelli, P., and Fasano, A. 1999. Expression of Vibrio cholerae zonula occludens toxin and analysis of its subcellular localization, Microb. Pathog., 27, 377. Uzzau, S., Lu, R., Wang, W., Fiore, C., and Fasano, A. 2001. Purification and preliminary characterization of the zonula occludens toxin receptor from human (CaCo2) and murine (IEC6) intestinal cell lines, FEMS. Vaile, J. H., Meddings, J. B., Yacyshyn, B. R., Russell, A. S., and Maksymowych, W. P. 1999. Bowel permeability and CD45RO expression on circulating CD20+ B cells in patients with ankylosing spondylitis and their relatives, J. Rheumatol., 26, 128.

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Ventura, A., Magazzu, G., and Greco, L. 1999. Duration of exposure to gluten and risk for autoimmune disorders in celiac patients, Gastroenterology, 117, 303. Wang, W., Uzzau, S., Goldblum, S. E., and Fasano, A. 2000. Human zonulin, a potential modulator of intestinal tight junctions, J. Cell Sci., 24, 4435. Watts, T. L., Alexander, T., Hansen, B., and Fasano, A. 2000a. Utilizing the paracellular pathway: a novel approach for the delivery of oral insulin in diabetic rhesus macaques, J. Invest. Med., 48, 185. Watts, T., Not, T., Berti, I., El Asmar, R., Di Pierro, M., Margaretten, K., and Fasano, A. 2000b. Alterations in zonulin secretion parallels changes in intestinal permeability and precede the onset of diabetes in BB/Wor diabetic rats, J. Invest. Med., 48, 184. Wendling, D., Bidet, A., and Guidet, M. 1992. Evaluation de la perméabilité intestinale au cours de la spondylarthrite ankylosante par le test au 51Cr-EDTA, Rev. Esp. Reumatol., 19, 253. Williams, D. G. 1993. Pathogenesis of idiopathic IgA nephropathy, Pediatr. Nephrol., 7, 303. Yacyshyn, B. R. and Meddings, J. B. 1995. CD45RO expression on circulating CD19+ B cells in Crohn’s disease correlates with intestinal permeability, Gastroenterology, 108, 132. Yacyshyn, B. R. and Pilarski, L. M. 1993. Expression of CD45RO on circulating CD19+ B-cells in Crohn’s disease, Gut, 34, 1698. Yacyshyn, B., Meddings, J., Sadowski, D., and Bowen-Yacyshyn, M. B. 1996. Multiple sclerosis patients have peripheral blood CD45RO+ B cells and increased intestinal permeability, Dig. Dis. Sci., 41, 2493. Yu, J. and Chien, Y. W. 1997. Pulmonary drug delivery: physiologic and mechanistic aspects, Crit. Rev. Ther. Drug Carrier Syst., 14, 395.

33

Functions of OSP/Claudin11-Containing Parallel Tight Junctions: Implications from the Knockout Mouse Cherie M. Southwood and Alexander Gow

CONTENTS 33.1 33.2 33.3 33.4 33.5

Introduction .................................................................................................723 Transmembrane Tight Junction Proteins ....................................................724 The Claudin Family of Integral Membrane Proteins .................................725 Claudin Family Members Regulate Paracellular Diffusion .......................728 Claudin 11...................................................................................................728 33.5.1 Tight Junctions of the Stria Vascularis..........................................730 33.5.2 Tight Junctions of Sertoli Cells.....................................................731 33.5.3 Tight Junctions of Central Nervous System Myelin ....................734 33.6 Is The Function of Claudin-11 Conserved in Different Tissues?..............737 Acknowledgments..................................................................................................740 References..............................................................................................................740

33.1 INTRODUCTION The characterization of tight junctions (TJs) has primarily been the province of morphologists and physiologists since their initial recognition as distinct intercellular bridges between epithelial cells (Cereijido, 1991). Scientists in these fields have described the localization, physical appearance, and morphological and functional diversity of TJs in exquisite detail and have determined how these structures interact with and complement other intercellular junctions in vivo and in cultured epithelial cell monolayers. Furthermore, these studies have yielded a comprehensive overview of the armamentarium of extracellular barriers available to multicellular organisms for excluding unwanted solutes from the environment while maintaining open endocytic and exocytic pathways for the organism to import nutrients and export metabolic waste. Over the last 10 years, biochemical and molecular techniques have been increasingly applied to the analysis of TJ structure/composition and these studies have led 0-8493-2383-5/01/$0.00+$1.50 © 2001 by CRC Press LLC

723

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to a quantum leap in the understanding of how multicellular organisms effectively isolate their internal environment from the external milieu. First and foremost, debate over the identity of the mediators of TJs — be they principally proteinaceous or predominantly nonbilayer-phase lipids — has been settled in favor of proteins. Second, the purification of many of these proteins and subsequent cloning and sequencing of the encoding genes has provided a clear view of the molecular basis of TJ structure and function. Conceptually, TJ structural proteins can be divided into three categories: membrane-associated cytoplasmic proteins, such as ZO-1, ZO-2, and ZO-3; transmembrane proteins, including occludin and the claudin family; and membrane-associated extracellular proteins, possibly including extracellular proteases and protease inhibitors. Currently, there is little evidence in support of the association of such extracellular proteins to TJs. Early biochemical fractionation schemes were very successful for the purification of over a dozen cytoplasmic protein components, and the subsequent application of molecular techniques enabled rapid identification of the genes encoding these proteins. However, the functional characterization of several proteins has proved relatively difficult, in large part because of the lack of suitable tools with which to dissect function (see Chapters 11 and 12). On the other hand, the identification and purification of transmembrane components of TJs has been an arduous task and until recently has been somewhat disappointing (Tsukita and Furuse, 1999). Nonetheless the application of molecular techniques, particularly gene ablation by homologous recombination in embryonic stem cells and the study of naturally occurring mutations, has clearly demonstrated the relative importance of these proteins in mediating the physical TJ barrier in the paracellular space (Saitou et al., 1998; Gow et al., 1999; Simon et al., 1999; Hirano et al., 2000).

33.2 TRANSMEMBRANE TIGHT JUNCTION PROTEINS The simple fact that TJs occlude the paracellular space has prompted expectations that the proteins that directly mediate these diffusion barriers will be present in the zonula occludens of many, if not all, cell types and will traverse the bilayer to connect with the cytoskeleton. Without doubt, the most significant contribution to the identification of such integral membrane proteins has come from Tsukita’s laboratory. Occludin was the first of these proteins to be identified (Furuse et al., 1993), and is a polytopic membrane protein predicted to have four transmembrane domains with the amino- and carboxyl-termini exposed to the cytoplasm. Although occludin is incorporated into the intramembranous TJ fibrils of most epithelial cells, recent studies in knockout mice demonstrate that this protein is unlikely to mediate the physical paracellular barrier of TJs (Saitou et al., 1998; Gow et al., 1999). Importantly, TJ assembly in cultured epithelial cell monolayers is disrupted by interfering with occludin function in various ways, which suggests that this protein plays an important role in regulating TJ formation (Balda et al., 1996; Sakakibara et al., 1997; Wong, 1997; Wong and Gumbiner, 1997; Lacaz-Vieira et al., 1999). A large number of other integral membrane proteins have recently been colocalized to TJs, including Junctional Adhesion Molecule, Vesicle-Associated Protein-33, and particularly members of the claudin family, and details about the expression of these proteins in different cell types as well as their contributions to TJ function are now emerging

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(Furuse et al., 1998a, b; Martin-Padura et al., 1998; Gow et al., 1999; Lapierre et al., 1999; Morita et al., 1999a, b; Simon et al., 1999; Liu et al., 2000; Palmeri et al., 2000).

33.3 THE CLAUDIN FAMILY OF INTEGRAL MEMBRANE PROTEINS The initial classification of two related proteins, claudins 1 and 2, as components of TJs (Furuse et al., 1998b) prompted a procession of publications detailing novel family members with broad and overlapping expression patterns (Furuse et al., 1998a; Morita et al., 1999a, b; Simon et al., 1999; Swisshelm et al., 1999). Several mammalian claudin cDNAs were known previously, but their cellular functions and subcellular localization to TJs were not initially appreciated (Briehl and Miesfeld, 1991; Bronstein et al., 1996; Katahira et al., 1997; Sirotkin et al., 1997). Currently, the family stands at 20 members in mice and humans, although there is evidence of an additional nine novel humans genes from homology searches of the Genbank high-throughput genomic sequence database. Three of these novel genes are represented as ESTs in the dbest database and reside on human chromosomes 1, 13, and 19 (Southwood and Gow, unpublished observations). In addition, several family members have been identified from the open reading frames of cDNAs expressed in other chordates including zebrafish, Xenopus, and ascidians. TJs have not been identified in Drosophila from morphological examination or from Genbank homology searches to find genes orthologous to occludin or claudins, despite the fact that these structures are found in other insect species (see Chapter 3); in Drosophila, pleated-septate junctions appear to occlude the paracellular space (Baumgartner et al., 1996). With such a large number of claudin family members described, an important task over the coming years will be to integrate the molecular data to identify subfamilies on the bases of evolutionary conservation, the morphology of the intramembranous fibrils, and the degree of leakiness of junctional complexes formed by each claudin, as well as amino acid and gene structure similarities. An analysis of gene organization effectively divides the claudin family into two groups: those genes containing introns vs. those genes comprising a single exon (Table 33.1). In genes with introns, the structures of the 5′ exons appear to be the most highly conserved; the first coding exons of CLAUDINS 1, 7, 10, 11, 15, 18, and 19 are 211 to 229 base pairs in length. These sizes correspond to between 70 and 76 codons plus the first base pair of the following codon before the splice-donor site. The nomenclature in Table 33.1 used to represent these sizes is 701/3 to 761/3 nucleotide triplets, where the fractional value denotes the nucleotide of the final triplet that precedes the exon boundary. The length of the second coding exon in each gene is also conserved, and is either 54 or 55 nucleotide triplets. Note that because of the split codon in the first coding exon of each gene, the codon preceding the splice-donor site in the second coding exon is also split after the first nucleotide. The gene structure of the intron-containing gene, CLAUDIN 16, differs from the other genes of this class in that the lengths of the first and second coding exons are unique; however, the third coding exon is 55 nucleotide triplets in length. Alternatively spliced exons have not been observed for any CLAUDIN gene, although the

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TABLE 33.1 The Gene Structures of Mouse and Human Claudin Genes No. of Nucleotide Triplets per Coding Exon Claudin No. of Gene Species Exons 1a Hu 4 2 Hu/Ms 1 3d Hu/Ms 1 4d Hu/Ms 1 5 Hu/Ms 1 Hu/Ms 1 6b 7 Hu 5 8c Hu/Ms 1 9b Hu/Ms 1 10 Hu 5 11 Hu/Ms 3 12 Hu/Ms 1 13d Ms 1 14 Hu/Ms 1 15 Hu 5 16a Hu 5 17c Hu/Ms 1 Hu 5 18e 19 Hu 4 20 Hu 1

1st 741/3 230 220 209/210 218 220 741/3 225 217 731/3 751/3 244 211 239 701/3 108 224 731/3 741/3 219

2nd 55

3rd 281/3

4th 531/3

5th

55

281/3

531/3

54 55

271/3 762/3

36

371/3

55 341/3

271/3 55

39 64

341/3 432/3

55 55

391/3 281/3

37 531/3

561/3

Human Gene Human Acc No. Chromosome AC009520 3q28-29 AL158821 Xq22.3-23 7q11.23 AC023010 7q11.23 AC000071 22q11.2 AC004643 16p13.3 AC003688 17 AP001846 18q22 AC004643 16p13.3 AL139376 13 AC008041 3q26.2-26.3 AC006153 7q2 7q11.23 AC000005 21 AC006329 7 AC009520 3q28-29 AP001846 18q22 AC016252 3 AL136383 1 AL139101 6

Hu, human; Ms, mouse. a,b,c Several human genes are located in close proximity to each other. d Five mouse claudin genes, including claudins 3, 4, 13 and two novel claudins, are in close proximity to each other and map to a single contig from mouse chromosome 5 (Genebank Acc# AC079938). Currently, it is not known if the genomic organization is similar at human chromosome 7q11.23, where CLAUDIN 3 and CLAUDIN 4 lie in close proximity; however, mouse chromosome 5 and human chromosome 7q11 are largely syntenic. e An open reading frame containing 731/3 codons and a canonical splice donor site lies upstream of the CLAUDIN 18 mRNA published in Genbank, suggesting that this gene has two promoters. However, ESTs containing the upstream exon have not been found.

presence of an exon (complete with splice-donor site) 5′ to the canonical exon 1 of CLAUDIN 18 (Genbank Acc# AF221069) is noted. Both exons 1 contain an open reading frame of 73 1/3 codons, share 71% identity (88% similarity), and are in-frame with the open reading frame of exon 2. However, the possibility that this novel exon may reflect the serendipitous presence of a pseudogene cannot formally be excluded because its sequence is not currently represented in the Genbank database as a cDNA or an Expressed Sequence Tag. Phylogenetic analysis of the primary structures of the claudins also suggests that family members encoded by intronless genes are most closely related to each other.

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FIGURE 33.1 Phylogenetic guide tree for the claudin family. The primary structures of 20 claudins were aligned using a CLUSTAL W analysis to demonstrate the similarities between intronless genes and those with multiple introns. To generate these data, an analysis of 10,000 random bootstrap trials was performed using default values for all variables and a correction for multiple substitutions (Thompson et al., 1994; 1997). Similar guide trees were obtained whether or not alignment positions with gaps were included or excluded in the bootstrap trials. Amino acid sequences used were from humans except for the inclusion of mouse claudin-13 in lieu of the sequence for the orthologous human protein. Terminal branches of the tree (bottom) have been trimmed for ease of presentation, which eliminates the significance of branch lengths.

Figure 33.1 shows a guide tree generated from a CLUSTAL W analysis (Thompson et al., 1994) that was obtained by compiling the results of 10,000 randomly generated bootstrap trials. The 12 claudins on the left of the tree are encoded by intronless genes and appear to be most closely related to a single gene that diverged from eight genes containing multiple introns (right). Branch lengths near the root of the tree (top) reflect the relative time since the divergence of each pair of proteins; however, the terminal branch lengths of the tree (bottom) have been trimmed for convenience and their lengths do not reflect divergence time. One interpretation of this analysis is that the primordial claudin gene contained introns and, during a course of multiple gene duplication and divergence events, gave rise to an intronless gene, which itself underwent gene duplications and divergence. There are several instances of the clustering of CLAUDIN genes; CLAUDINS 1 and 16, 3 and 4, 6 and 9, 8 and 17 are located on different chromosomes in pairs, with less than 150 kb of intervening sequence. Furthermore, human CLAUDIN 11 (3q26.2-26.3) maps nearby to CLAUDINS 1 and 16 (3q28-29). In mouse, a single 78-kb contig (Genbank Acc# AC079938) contains three claudin genes — 3, 4, 13 — and two apparent pseudogenes (interrupted open reading frames). This region of mouse chromosome 5 is syntenic to human chromosome 7q11, which may have a similar genomic structure; indeed, CLAUDINS 3 and 4 map to human 7q11. Certainly, such tight spatial relationships suggest close evolutionary ties stemming from gene duplication, as does the presence or absence of introns in each gene of the clusters; however, it remains to be determined how significantly the expression patterns and functions of these genes have diverged during evolution.

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33.4 CLAUDIN FAMILY MEMBERS REGULATE PARACELLULAR DIFFUSION Rather than serving as impermeable barriers to the paracellular diffusion of large and small molecules alike, the TJs of many so-called leaky epithelia permit the transepithelial movement of water and particular small-molecular-weight molecules while impeding the diffusion of others to varying degrees. Electrophysiological data have led to the widely accepted view that paracellular diffusion is a qualitatively selective process whereby permeability is determined by the number of intramembranous fibrils in the junctional complex (Claude and Goodenough, 1973) as well as the size and charge of ions passing through relatively large pores in the junctions (reviewed in Reuss, 1991). This diffusion promiscuity contrasts to the demonstration that ionic channels generally utilize small-diameter pores as highly selective filters for the diffusion of single ion species across a bilayer. Another aspect of the new understanding about the functions and properties of TJs stems from molecular studies of patients with primary hypomagnesemia (Simon et al., 1999). As discussed in detail in Chapter 22, Mg2+ retrieval from the renal filtrate in the thick ascending loop of Henle occurs predominantly via paracellular diffusion down an electrochemical gradient. However, point mutations throughout the coding region of the CLAUDIN 16 gene render patients virtually incapable of resorbing Mg2+ in the kidney and cause severe depletion of this ion in the blood, among other abnormalities. An intriguing property of the ascending loop, which is an example of a tight epithelium, is the specificity with which the epithelial cells resorb Mg2+ but not other ions. Moreover, very little water moves across the epithelia in this region, suggesting that TJs of the ascending loop exhibit a degree of specificity for Mg2+, which may be more akin to an ion channel than has previously been appreciated (Wong and Goodenough, 1999).

33.5 CLAUDIN 11 Evidence of the evolutionary divergence of paralogous claudin genes is most apparent at the 3′ ends of the coding regions, where there is little discernible homology at the amino acid level. Nevertheless, the importance of the primary structure at the carboxyl termini of the claudins is suggested by the high degree of conservation of the amino acid sequence (>80% identity) between orthologous mouse and human proteins. Furthermore, the fact that cytoplasmic TJ proteins, such as ZO-1, ZO-2, and ZO-3, bind to the carboxyl termini of claudins (Itoh et al., 1999) suggests that this region may be important for organizing TJs with specific transmembrane components in different locations. At least for CLAUDIN 11, the gene structure between mouse and human has also remained constant during evolution (Figure 33.2); not only are the number of codons in each exon identical, but the sizes of introns 1 and 2 are unchanged between the two species at 4 and 9 kb, respectively. To investigate the function of claudin-11 in detail, the authors disrupted the gene encoding this protein in mice by homologous recombination in embryonic stem cells (Gow et al., 1999). Essential features of the targeting construct are shown in Figure 33.2,

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FIGURE 33.2 Comparison of the structures of the human and mouse CLAUDIN 11 genes. Each gene has three coding exons (black rectangles), containing the same number of codons. The sizes of the introns are also conserved. The targeting construct used to generate claudin 11-null mice is shown. This construct is centered around exon 1 and replaces the coding sequence with the bacterial β-galactosidase open reading frame and an antibiotic-resistance gene, PGKneo, as a selectable marker. The arrowheads at either end of this marker gene represent loxP sites used for its excision if necessary. “X” represents the positions of hypothetical crossover and integration sites between the targeting construct and the endogenous claudin 11 gene.

where the coding region for claudin-11 in exon 1 is replaced by the coding region for the bacterial enzyme, β-galactosidase (lacZ), and that a PGKneo gene flanked by loxP sites (arrowheads) lies immediately downstream of the lacZ open reading frame. The utility of this construct is that transcription from the claudin 11 promoter generates β-galactosidase, which is not represented in wild-type mice and can easily be detected using a simple histochemical stain in heterozygous (i.e., one copy of wild-type claudin 11 gene and one copy of the mutated gene) or homozygous mice (i.e., two copies of the mutated gene; claudin 11-null). Histochemical staining for β-galactosidase in knockout mice reveals broad expression of claudin 11 in embryos from E8.5 to birth, and is most widespread at E14.5 where staining is found in the leptomeninges surrounding the brain, cranial nerves, and spinal cord; the nasal epithelium; the hair follicles of the vibrissae; the membranous labyrinth of the vestibulocochlear apparatus; the perioptic mesenchyme; the foregut; the epaxial muscles; the mesonephric ducts; and the urogenital tract (Gow et al., 1999). Importantly, the expression of β-galactosidase in these tissues is consistent with in situ hybridization data using the claudin-11 cDNA as a probe (Bronstein et al., 2000). Assuming that claudin-11 functions only in the capacity of regulating paracellular diffusion, it is surprising that TJs are established in so many tissues during development and that the observed phenotypes in the knockouts are not particularly dramatic. Nonetheless, defects have been identified in the inner ear, testis, and central nervous system and it is speculated that claudin-11-containing TJs may perform a similar function in each of these tissues. In more general terms, similarities in the freeze-fracture morphology of TJ fibrils from these tissues — namely, linear, rarely branching arrays of intramembranous particles (Gow et al., 1999) — suggest a close relationship between ultrastructural appearance and the molecular composition of most TJs throughout the body. Thus, these features may be important indicators of comparable function in different tissues (Gow et al., 1999).

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FIGURE 33.3 Expression of claudin-11 in the inner ear. (A) The vestibular semicircular canals are interconnected with the auditory cochlear duct via the utricle and the saccule, and are filled with endolymph. An additional compartment filled with perilymph is connected to the oval and round windows at each end. Crosshatching represents the temporal bone. (B) Transverse section from A shows the three compartments of the cochlear duct. The central scala media contains the organ of Corti and is filled with perilymph secreted by the stria vascularis in the spiral ligament. Note the dark β-galactosidase staining in this section, which shows the basal cell layer of the stria. The intermediate and marginal cell layers of the stria are unstained.

33.5.1 TIGHT JUNCTIONS

OF THE

STRIA VASCULARIS

The vestibulocochlear apparatus in mammals comprises the membranous and bony labyrinths within the lateral petrous bone of the cranial vault (Figure 33.3A). The membranous labyrinth (the Scala media) encloses a compartment running the length of the inner ear from the semicircular canals to the apex of the spiral cochlear duct and is filled with an extracellular fluid, the endolymph. Different in composition from most extracellular fluids, the endolymph contains a high concentration of K+ (~150 mM in rat; Salt and Konishi, 1986) and a low concentration of the other major cation, Na+ (~1 mM in rat; Salt and Konishi, 1986). Endolymph is largely secreted into the scala media by the stria vascularis, a specialized epithelium embedded in the lateral wall (spiral ligament) of the cochlear duct, which maintains the Scala media at a positive potential of 80 to 100 mV compared with surrounding compartments (Figure 33.3B). This endocochlear potential is critical for the normal function (reviewed in Salt and Konishi, 1986) of hair cells in the organ of Corti, which constitutes the mechanosensory apparatus for signal transduction to the brain. Two compartments flank the Scala media on either side, the Scalae vestibuli and tympani, which are joined at the apex of the cochlear duct and are filled with perilymph. Perilymph is similar in composition to most other extracellular fluids, with a high

Functions of OSP/Claudin-11-Containing Parallel Tight Junctions

731

concentration of Na+ (~150 mM) and a low concentration of K+ (~4 mM), and is continuous with cerebrospinal fluid in the subarachnoid space via the cochlear aqueduct. As shown by β-galactosidase histochemistry in Figure 33.3B, claudin-11 is normally expressed in a restricted region of the spiral ligament in the adult cochlear duct, which reveals the basal cell layer of the stria vascularis. TJs of the basal cells act in concert with a second TJ network of the marginal cell layer to isolate the intermediate compartment (Luciano et al., 1995). The isolation of this cellular compartment is believed to be crucial for establishing the endocochlear potential, although the molecular details are at present unclear (Salt et al., 1987; Souter and Forge, 1998; Takeuchi and Ando, 1998). Evidence from ultrastructural and immunocytochemical analyses suggests that marginal cells in the stria utilize a Na+,K+ATPase pump to resorb K+ from the intermediate compartment for secretion into the Scala media (Xia et al., 1999). Moreover, gene targeting experiments demonstrate that the absence of K+-channel or Na+,K+,Cl–-cotransporter activities in the marginal cells largely abolishes the secretion of endolymph by the stria, which results in a collapse of the membranous labyrinth, degeneration of hair cells in the organ of Corti, vestibular abnormalities, and profound deafness (Vetter et al., 1996; Delpire et al., 1999; Dixon et al., 1999; Flagella et al., 1999). The majority of the K+ reaching into the stria vascularis originates in the perilymph, and recent studies indicate that a network of diverse cell types from the organ of Corti to the spiral ligament are coupled by connexin 26-containing gap junctions to channel K+ into the stria (reviewed in Kikuchi et al., 2000). As anticipated from the β-galactosidase staining in the cochlea, TJs of the stria basal cell layer are absent in claudin 11-null mice (Gow et al., 1999) and studies are currently under way to determine if these mutants exhibit hearing deficits. However, it is clear from the initial analyses that overt vestibular phenotypes, such as the circling behavior of Na+,K+,Cl–-cotransporter knockout and shaker-with-syndactylism mice (Madara et al., 1993; Dixon et al., 1999; Flagella et al., 1999), are not observed in the claudin 11-null mice; thus, the absence of basal cell TJs in the stria may confer a relatively mild defect. Precedence for such a notion comes from the characterization of Brn4 knockout mice, in which spiral ligament fibrocytes in the cochlear duct involved in K+ recycling from the perilymph to the stria vascularis are morphologically abnormal, resulting in profound deafness in adult animals with no discernible vestibular defects (Minowa et al., 1999).

33.5.2 TIGHT JUNCTIONS

OF

SERTOLI CELLS

In many mammalian species, somatic Sertoli cells and cells of the primordial germ cell lineage constitute the stratified epithelium, which is organized into seminiferous tubules in the testis. Sertoli cells are highly polarized, have basally located nuclei, span the thickness of the epithelium, and serve as support cells for the proliferation and differentiation of the male gametes. On the other hand, spermatogonia are comparatively small cells that form the stem cell population of the germ cell lineage and, similar to Sertoli cells, are attached to the basement membrane. The differentiation of spermatogonia is a continuous process that can be divided into two phases

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on the basis of cell cycle-related events and spatial organization. The first phase involves proliferation for self-renewal of the stem cell population and the production of spermatocyte precursor cells, which occurs close to the basement membrane. The second phase includes centripetal displacement of primary spermatocytes toward the lumen of the seminiferous tubule, meiotic cell division, and differentiation into spermatids. During this phase, the developing spermatids are heavily dependent on Sertoli cells to serve as attachment surfaces; to produce the luminal fluid of the seminiferous tubule; and to provide a sealed, regulated environment to meet the ionic and hormonal requirements of the developing germ cells. TJs between Sertoli cells form the cellular basis of the sealed adluminal compartment in the seminiferous tubule and constitute the major component of the blood–testis barrier (Dym and Fawcett, 1970; Gilula et al., 1976). These intercellular junctions are among the most extensive and impermeable that have been characterized and are described in detail in Chapter 28. Indeed, this impermeability is highlighted by differences in the ionic composition of the seminiferous fluid compared with that in the interstitial space. In particular, the adluminal fluid contains a high concentration of K+ (~50 mM; Waites and Gladwell, 1982) and a low concentration of Na+ (~110 mM; Waites and Gladwell, 1982). Thus, seminiferous fluid exhibits some similarities to endolymph in the inner ear, although the adluminal electrical potential is a modest –1 mV (Cuthbert and Wong, 1975). Early studies of the testis indicated that the differentiation of germ cells is critically dependent on Sertoli cell TJs and, although the molecular basis of this dependence has not been established, the maturation of Sertoli cell TJs is strongly correlated with the commencement of spermatogenesis in many continuous- and seasonal-breeding species (Gilula et al., 1976). In addition to generating a regulated extracellular environment in the adluminal compartment, the blood–testis barrier is widely believed to conceal developing spermatocytes from immune surveillance. Spermatogenesis in many mammals begins long after the perinatal establishment of “self-tolerance” by the immune system. Conceivably, a consequence of the expression of sperm-specific genes during sexual maturation is that these cells could be regarded as foreign, thereby leading to autoimmune attack and destruction of the germ cell lineage. However, evidence obtained over the last decade indicates that the immune-privileged status of seminiferous tubules is provided by Sertoli cells in the form of secreted immunosuppressive compounds to quell local autoimmune responses (Yule et al., 1990; Turek et al., 1996). Freeze-fracture preparations of the seminiferous tubules from claudin 11-null mice demonstrate that Sertoli cell TJs in these animals are completely absent, although the occluding junctions between endothelial cells in the testis are morphologically normal (Gow et al., 1999). A major consequence of the absence of the blood–testis barrier is azoospermia, and at the light microscope level few spermatocytes have been observed beyond the leptotene/pachytene stage (Figure 33.4) which is the point at which germ cells reach the adluminal compartment (Leblond and Clermont, 1952). Furthermore, arrested spermatogenesis leads to a retrogression of the seminiferous epithelium, and by 7 months after birth most tubules in the testis of knockout mice are markedly hypocellular. Nonetheless, Sertoli cells and spermatogonia are present and appear morphologically normal by light microscopy.

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FIGURE 33.4 Pathology in the testes of 7-month-old claudin 11-null mice. (A) Seminiferous tubules in wild-type mice exhibit a well-organized stratified epithelium and a prominent lumen containing the filamentous tails of spermatids (arrowheads). (B) Seminiferous tubules from claudin 11-null mice are reduced in size and hypocellular. The epithelia are disorganized and well-defined lumina are rarely observed. Germ cells in the early stages of spermatogenesis, up to leptotene and pachytene stages, are present in most tubules but more-differentiated cells are absent. There is a marked increase in the Leydig cell population.

Outside the seminiferous tubules, secondary pathological changes are also evident in adult mice: a dramatic increase in the Leydig cell population is apparent and the tunica albuginea is conspicuously thickened. Importantly, there is no evidence of T-cell infiltration into the testis, and autoimmune orchiditis has not been observed at any age in the knockout animals. Furthermore, antispermatocyte antibodies are not detected in serum from these animals, which argues strongly against any involvement of the immune system in the phenotype. From these data, the authors conclude that Sertoli cell TJs are critical for spermatogenesis but play little or no role in protecting spermatocytes from immune surveillance (Gow et al., 1999).

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33.5.3 TIGHT JUNCTIONS

Tight Junctions OF

CENTRAL NERVOUS SYSTEM MYELIN

The central nervous system can be conveniently divided into two regions on the basis of color: gray matter contains neuronal and glial cell bodies, whereas white matter is devoid of neuronal cell bodies and comprises mostly myelinated axons and glial cells. The oligodendrocyte is the glial cell that synthesizes the lipid-rich myelin sheath from which white matter derives its color and a single cell can ensheath between one and several dozen axons (reviewed in Morell and Quarles, 1999; Peters et al., 1991). Myelin sheaths in the central nervous system are synthesized in segments, or internodes, that are several hundred micrometers in length and are elaborated by single flattened processes that extend from oligodendrocyte cell bodies. Upon contacting an axon, each cell process expands into a large cytoplasm-filled membrane sheet as it spirally wraps around an axon ten times or more. After one or two wraps, compaction of the myelin membrane commences and the cytoplasm is largely extruded from central regions of the membrane sheet back to the cell body. When viewed in cross section under the electron microscope, the spiral nature of the wrappings can be appreciated by following the course of the dark or light repeating lines in the sheath (reviewed in Raine, 1984). Other morphological features of central nervous system myelin that are visible by electron microscopy in transverse sections of myelin sheaths include Schmidt–Lanterman incisures (Blakemore, 1969; Hildebrand, 1971), which are cytoplasm-filled channels within the compacted regions of large-diameter myelin sheaths (these structures are more common in peripheral nervous system myelin), and a series of linear structures that often span the thickness of a sheath, appropriately named the radial component. When purified biochemically, the radial component has been shown to contain several known myelin proteins and lipids, but none that is a specific marker of this structure (Kosaras and Kirschner, 1990; Karthigasan et al., 1996; Yamamoto et al., 1999). Importantly, the localization and appearance of the radial component corresponds to linear rows of intramembranous particles visualized in freeze-fracture preparations of central nervous system myelin (reviewed in Schnapp and Mugnaini, 1978). The freeze-fracture data suggest that the radial component represents TJs throughout the myelin sheath, although transmission electron microscopy reveals considerable morphological differences from TJs between epithelial cells (Kosaras and Kirschner, 1990). On the other hand, the fact that claudin-11 has been localized to the radial component and the paranodal loops by immunocytochemistry, immunoelectron microscopy, and immunofreeze fracture lends considerable additional support to the identification of these structures as occluding junctions (Gow et al., 1999; Morita et al., 1999b; Arroyo and Scherer, 2000; Bronstein et al., 2000). Myelin sheaths along nerve fibers are punctuated on either side by short segments of unmyelinated axon on the order of 1 µm in length, called the nodes of Ranvier. The specialized region of myelin juxtaposed to a node of Ranvier is known as paranodal myelin and represents the lateral edge of the myelin membrane sheet where cytoplasm remains after compaction is completed (Figure 33.5). With each spiral turn of myelin, the lateral edge adheres to the axonal membrane closer and closer toward the node of Ranvier, and in longitudinal sections the paranodal myelin

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FIGURE 33.5 The relationship between a central nervous system myelin sheath and an axon. (A) Schematic of part of the myelin sheath in the region of the paranode (right), which has been cut away to show salient features: the axon; the lateral myelin loops, which are interconnected by TJs; the radial component, which often spans the myelin from the inner to outer loops; and the intramyelinic space, which is delimited by the TJs in the sheath. In addition, a portion of the outer turn of the myelin sheath has been cut away to show the spiral course traveled by the radial component along the internode. The radial component is also observed in regions of the sheath other than between the inner and outer loops (not shown here for simplicity). (B) The view of a myelin sheath that has been unraveled to reveal its overall shape, and to indicate the locations of the cytoplasm-filled inner, outer, and lateral myelin loops. Sets of TJ strands through the myelin are represented by the black lines around the edge and through the internode. For clarity, these junctions are positioned in the central region of the cytoplasmic channels rather than in their true locations along the outside edges. Note that the TJs are continuous around the perimeter of the myelin. The radial component is shown as rows of TJs roughly parallel with the inner and outer loops, which may be continuous with the paranodal junctions (dashed lines).

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membrane appears as a series of cytoplasm-filled loops where the outer turn of the membrane defines the boundary of the node. A major function of the internodal myelin is to insulate electrically most of the axonal membrane, whereas at the nodes of Ranvier the paranodal myelin serves to cluster the voltage-gated Na+ channels, which depolarize this membrane segment in the propagation of saltatory conduction. The importance of the structural organization of paranodal myelin for nerve conduction has been known for decades, but only recently have molecular studies identified several of the adhesive molecules that stabilize the sheath in this region (reviewed in Arroyo and Scherer, 2000; Popko, 2000). Three junctional complexes have been observed by transmission and freeze-fracture electron microscopy in central nervous system myelin (reviewed in Rosenbluth, 1990; Schnapp and Mugnaini, 1978): axoglial junctions, adherens junctions, and occluding junctions. Axoglial junctions are reminiscent of the pleated septate junctions in Drosophila (Baumgartner et al., 1996; Einheber et al., 1997), and bridge the extracellular space between the myelin sheath and the axon (i.e., a heterotypic junction). Recent studies indicate that caspr/neurexin IV mediates these junctions in mammals (Baumgartner et al., 1996; Einheber et al., 1997; Bhat et al., 2000). On the other hand, the adherens and occludens junctions in myelin are rather unusual in the sense that they connect the juxtaposed surfaces of the same cell (i.e., an autotypic junction) and may have been adapted by oligodendrocytes to perform novel functions. Indeed, the extraordinary nature of the oligodendrocyte itself is emphasized because it is one of the only known cell types that does not secrete a basal lamina but nonetheless makes extensive use of epithelial junctions in myelin sheaths and on the cell body (Rash et al., 1997). The function of occluding junctions in myelin remains unclear, despite the fact that more than 20 years have passed since these structures were initially found to circumscribe the myelin sheaths of both oligodendrocytes and Schwann cells (reviewed in Schnapp and Mugnaini, 1978). TJs in peripheral nervous system myelin occupy positions analogous to those in the central nervous system (Mugnaini and Schnapp, 1974), but do not contain claudin-11 (Bronstein et al., 1996). From early studies in the central nervous system, TJs in myelin were thought to impart structural stability on the internode (Tabira et al., 1978; Nagara and Suzuki, 1982; Yamamoto et al., 1999) and to conceal myelin-specific components from immune surveillance, which might otherwise provoke an autoimmune response (Mugnaini and Schnapp, 1974). However, data from claudin 11-null mice do not support either of these hypothesized roles: phagocytosis of myelin sheaths or autoimmune-mediated demyelination are not observed in the central nervous system of knockout mice; myelin compaction, period, and thickness are normal; and myelin proteins are present in knockout brains at wild-type levels. The fact that claudin 11-null mice live a normal lifespan without evidence of the neurodegeneration observed in other myelin mutants with advancing age (Mastronardi et al., 1993; Kagawa et al., 1994; Readhead et al., 1994; Griffiths et al., 1998; Stecca et al., 2000) suggests that claudin-11 does not play a role in myelin stability — the axoglial and adherens junctions are more likely to perform this function, as has been suggested from other studies (Fannon et al., 1995; Bhat et al., 2000). Rather, myelin TJs may serve in a capacity more akin to those in epithelial

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cell layers in which they regulate the permeability of the paracellular space (Figure 33.6). In this light, occluding junctions are continuous around the perimeter of the myelin sheath and delimit a small extracellular intramyelinic compartment. This compartment is separated from the periaxonal space — which is located between the myelin sheath and the axon — and from the general (including the nodal) extracellular space by the TJs that seal the inner and outer myelin loops (Mugnaini and Schnapp, 1974). The effect of this seal is to limit the extracellular current through myelin and to assure that the myelin sheath acts as a capacitor of high resistance. The absence of this seal would result in the generation of an internodal resistive shunt current through the myelin adjacent to a depolarizing node of Ranvier. One undesirable outcome stemming from internodal shunt currents is reduced saltatory conduction velocity (Ritchie, 1984), the extent of which increases with the number of nodes of Ranvier along the axon. Accordingly, long fiber tracts are most severely affected by abnormal conduction such as those innervating caudal regions of the animal. Indeed, the neurological phenotype of claudin 11-null mice is consistent with conduction defects; these animals exhibit increased latency times of evoked potentials in the visual cortex, in response to strobe flashes, and persistent hind limb weakness. Furthermore, the transient intention tremors observed in weanling knockout mice may reflect a diminished competence of thin myelin sheaths to insulate axons during myelinogenesis (Gow et al., 1999).

33.6 IS THE FUNCTION OF CLAUDIN-11 CONSERVED IN DIFFERENT TISSUES? Although the understanding of claudin-11 function in embryos is currently limited, the investigation of adult phenotypes in claudin 11-null mice have provided considerable details about the functions of TJs in the testis and central nervous system myelin. Furthermore, doubt has been cast over widely accepted views concerning the functions of the occluding junctions in several tissues, particularly those purporting an involvement in the generation of immune-privileged compartments. With the identification of around two dozen members of the claudin family, and the likelihood that all of these proteins mediate the physical paracellular barrier of TJs in different tissues, a major challenge will be to define the properties of each family member in molecular terms. Undoubtedly, these data will clarify many mysteries underlying the diverse morphological and physiological characteristics of TJs throughout the body. Indeed, the delineation of claudin-16 as a paracellular Mg2+ filter is ample precedence (Simon et al., 1999; Hirano et al., 2000). Despite the absence of detailed functional data for claudin-11 can one, nonetheless, construct a usable hypothesis to focus future efforts? By considering several aspects of TJ function that are common to two adult tissues expressing claudin-11 — namely, the testis and the inner ear — a rudimentary hypothesis can be framed that addresses the function of the occluding junctions between Sertoli cells and basal cells of the stria vascularis. Thus, it is posited that claudin-11 parallel TJs constitute a paracellular barrier that is impermeable to K+ and functions to delimit compartments in which the concentration of this ion in the extracellular milieu vastly exceeds that in the interstitial fluids of surrounding compartments. Although these junctions

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FIGURE 33.6 Internodal resistive shunt currents generated in the absence of myelin TJs. (A) A longitudinal section through a myelinated axon to show a myelin sheath and flanking nodes of Ranvier. For clarity, the periaxonal and intramyelinic spaces are drawn disproportionately large, and only the two innermost myelin lamellae are represented. The paranodal myelin loops (hatched) in wild-type mice (top) are interconnected by TJs and adhere to the axon via axoglial junctions. Sodium ions moving through ion channels at a depolarizing node of Ranvier (left) displace K+ in the region to generate a capacitive shunt current at the adjacent node (right). The Na+ ions at the external surface of this node are displaced from the membrane and complete a circuit with the depolarizing node along the outside of the myelin sheath. However, in the absence of myelin TJs (bottom), some of the displaced K+ at the depolarizing node activate internodal K+ channels in the axonal membrane and K+ can flow into the periaxonal and intramyelinic spaces to generate a resistive shunt current, which spirals through the myelin and completes the circuit with the depolarizing node. Consequently, less current is available at the adjacent node, which increases the time taken to drive those Na+ channels to threshold and depolarizes the node, i.e., conduction velocity is slowed. (B) Cross section through the internode of a myelinated axon to show more clearly the spiral path taken by the resistive shunt current (dashed spiral arrow) through the connected periaxonal and intramyelinic spaces in claudin 11-null mice (left). Only the first turn of the multilamellar membrane is represented (thick spiral line). In wild-type mice (right), these shunt currents are not normally generated because the continuity of the TJs around the inner, outer, and paranodal myelin loops electrically seal the intramyelinic space and ensure that the myelin sheath acts as a capacitor of high resistance.

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may also be impermeable to Na+, this ion may be excluded from traversing the TJs because of unfavorable electrochemical gradients, which are analogous to those governing the exclusion of Na+ from intracellular compartments. In claudin 11-null mice, the absence of even the remnants of intramembranous fibrils in freeze-fracture replicas from Sertoli cells and stria basal cells suggests that neither occludin nor other claudin family members are significant components of the TJs between these cell types (Gow et al., 1999). One interpretation of this observation is that claudin-11 exhibits unique properties that cannot be recapitulated by other proteins. In accordance with this hypothesis, one might reasonably expect that claudin-11 junctions isolate K+-rich fluids in the lumina of epithelia in addition to seminiferous tubules and stria basal cells, and experiments are under way to survey different tissues for such evidence. In any case, with this basic hypothesis in hand, one can now expand its framework to accommodate the function of claudin-11 in central nervous system myelin sheaths. As discussed in Section 33.5.3, the principal function of the myelin sheath is to minimize shunt currents across the axonal membrane at all points outside of the nodes of Ranvier. To fulfill this role, myelin must be electrically sealed at the inner and outer edges of the spiral membrane sheet, at the lateral edges of each turn of membrane around the axon, and between the myelin sheath and the axon. Studies from several laboratories (Mugnaini and Schnapp, 1974; Einheber et al., 1997; Morita et al., 1999b; Arroyo and Scherer, 2000) show that claudin-11 and caspr/neurexin IV are appropriately targeted in myelin to establish these barriers and, thereby, serve to delimit two small compartments inside the myelin/axon unit, called the intramyelinic space and the periaxonal space (see Figure 33.6). Consistent with this hypothesis, it is envisaged that claudin-11 TJs in myelin serve to increase the electrical resistance of this membrane because of their relative impermeability to K+. In this vein, it is noted that voltage-gated K+ channels in neurons are almost entirely spatially restricted to the internodal region of the axon (reviewed in Arroyo and Scherer, 2000) and do not normally pass significant outward current because of the low capacitance of the myelin sheath (reviewed in Ritchie, 1984). However, strong K+ currents from these concealed channels are detectable if the organization of the paranode is disrupted by acute paranodal demyelination (Chiu and Ritchie, 1980) or after repetitive stimulation of the nerve (Moran and Mateu, 1983). A likely explanation for K+-channel activity under these conditions is that the axoglial and TJs at the paranodes are disrupted, which enables internodal resistive shunt currents to form within the internode as each node of Ranvier is depolarized. Therefore, the authors contend that the absence of these shunts under normal conditions is due, in no small measure, to the impermeability of the paranodal junctions to K+. Indeed, direct evidence of a requirement for K+-impermeable barriers around axons has been obtained in Drosophila (Baumgartner et al., 1996). In the absence of neurexin, Drosophila first-instar larvae fail to hatch as a result of reduced motor neuron activity and muscle contraction. However, signal transduction is restored if neurons dissected from these animals are bathed in solutions with a low concentration of K+. The fact that hemolymph is a K+-rich fluid demonstrates that a major function of the pleated septate junctions in Drosophila is to provide a barrier to the movement of K+ across the hemolymph–nerve barrier, and it is reasonable to

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assume that the function of neurexin IV has been conserved in vertebrates (Bhat et al., 2000). The fact that the periaxonal space and the intramyelinic space are separated by TJs along the inner turn of the myelin membrane strongly suggests that the occluding junctions must also be impermeable to K+ to avoid the generation of internodal shunt currents during saltatory conduction. Therefore, it is likely that oligodendrocytes utilize TJs in myelin comprising claudin-11 for two major reasons: to increase the electrical resistance of this insulating membrane, and to delimit the intramyelinic space with a diffusion barrier that is impermeable to K+. In conclusion, characterization of mice lacking the claudin 11 gene has revealed phenotypes in several tissues, which when considered together, suggests a conserved function for parallel TJs in the inner ear, testis, central nervous system myelin, and possibly in other tissues where these junctions are present. Thus, it is speculated that a major function of claudin-11 is to minimize the paracellular diffusion of K+ between compartments. The widespread expression of claudin-11 implies that high concentrations of K+ play important roles in many cell types, not only for sensorineural transmission by hair cells in the vestibulocochlear apparatus, but also in less well characterized processes such as spermatogenesis in the testis and saltatory conduction in the central nervous system. It is currently an open question whether claudin-11 plays important roles during development and, if so, what purposes confined K+-rich fluids in the embryo might serve.

ACKNOWLEDGMENTS The authors are deeply indebted to Dr. Victor Friedrich, Jr., for sharing invaluable insights into myelin ultrastructure and electrophysiology and to Dr. Robert Lazzarini for his critical review of the manuscript. This work was supported in part by grants from the National Multiple Sclerosis Society of America (RG2891A1) and the Human Frontiers Science Program (RG318/97).

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Index A A549, 359 ABC transporter, see ATP-binding cassette transporter Accesory cells, 447 N-Acetyl-β-D-glucosaminidase, 405 N-Acetylneuraminic acid, 434 Actin, 55, 404 -anchoring protein, 616 cortical, 615 cosedimentation, 275 cytoskeleton, 9, 153, 168, 169, 170, 176, 186, 503, 578 filaments, 175, 202, 590 junction-associated, 616 microfilaments, 708 permeability, subsurface or cortical, 616 polymerization, 176 remodeling, 129 Actin–myosin cytoskeleton, organization and regulation of tight junction by, 265–284 cytoskeleton and paracellular permeability, 267–271 bacterial pathogens, 269–270 cytochalasins, 267–269 pharmacological agents, 269 physiological agents, 270–271 small GTP-binding proteins, 270 functional significance of direct interactions between cytoskeleton and TJ proteins, 274–277 clues from invertebrate model systems, 277 de novo assembly of tight junctions, 275–277 regulation of paracellular permeability, 274–275 interaction of actin cytoskeleton with molecular components of tight junction, 272–274 organization of F-actin at tight junction of epithelial cells, 266–267 possible mechanisms of cytoskeletal action on tight junction organization and function, 271–272

Actomyosin contraction, 266, 270, 271 Acute myeloid leukemias, 241 Acute renal failure, 535 AD, see Alternative domain Adaptor proteins, 151 Adenylate cyclase, 9 ADH, see Antidiuretic hormone Adherens junctions (AJ), 168, 185, 233, 417, 427, 431, 736 blood–brain barrier, 420 cadherin-based, 287 neutrophil transendothelial migration, 631 protein, degradation of, 634 Adhesion(s) cell–matrix, 676 molecules, 222, 630 zipper formation, 176 β-Adrenergic glaucoma medications, 669 AF6, 21, 23 Afadin, 131, 241, 272 AF-6/afadin, 274, 277 AIDS, 565 Airway epithelia, 499 AJ, see Adherens junctions Alkaline phosphatase, 664 Alkali sulfates, TJs induced by, 471 Alternative domain (AD), 246 Amanita phalloides, 202 Amiloride, 65 Aminophospholipid translocase, 322 ANIT, see α−Naphthylisothiocyanate Ankylosing spondylitis (AS), 702 Ankyrin, 179 -binding domain, 186 -binding modules, 167 /fodrin cytoskeleton, 168 Antibiotic-associated diarrhea, 519 Antibodies, anti-sperm, 618, 733 Antidiuretic hormone (ADH), 64, 485 Antimycin A, 269 Antispermatocyte antibodies, 733 AP-1, 167 Apical crypt, 449 Apical cytoskeleton, 509 Apical domain, 350 Apical junctional complex, 266, 418, 419, 427 see also Tight junction, Adherens junctions

747

748 Apical membranes, 314 Apical sorting receptor, 151 Apical targeting, 167 Apicolateral TJs, 663 Apoptosis, 399, 401, 570 in HT29 cells, 475 of mammary epithelial cells, 406 signal-regulating kinase (ASK1), 478 Aprotinin, 354 Aquaporin, 77 Aquaporin-1 gene knockout mice, 208 Aqueous humor, 660, 666, 671 flow, 672 transparency of, 677 Aqueous outflow system, 654, 666 Arachnids, 40, 43, 54 Armadillo family, 170 Arthropods TJs, microfilaments of, 55 AS, see Ankylosing spondylitis Asbestos, 356 Ascidians, 725 ASK1, see Apoptosis signal-regulating kinase Aspirin, 355 Astrocyte conditioned medium, 423, 424, 432 Asymmetric cues, 171 ATP, 467 -binding cassette (ABC) transporter, 323 -dependent transport proteins, 576 depletion, 269, 277, 278, 545 Atypical protein kinase C, 242 Auditory cochlear duct, 730 Autoimmune diseases, 702 Axoglial junctions, 736 Axons, myelinated, 734, 738 Azoospermia, 732 Azurocidin, 359, 360

B Bacillus polymyxa, 356 Backleak, 535 Bacterial endotoxins, 406 Bacterial metalloproteases, 501, 502 Bacterial/permeability-increasing protein, 359 Bacterial toxins, 12 Bacterial translocation, 499, 501 Bacteriodes fragilis, 270, 708 Bacteroides fragilis enterotoxin (BFT), 496, 497 BAI, see Brain angiogenesis inhibitor Basal cells, function of occluding junctions between Sertoli cells and, 737 Basement membrane, 168, 350 Basolateral domain, 350 Basolateral membrane, 314, 499, 500

Tight Junctions Basolateral sorting, 167 Basolateral vesicles, docking and fusion of, 178 BBB, see Blood–brain barrier B-cadherin, 420 BDL, see Bile duct ligation Benzodiazepine, 690 BFA, see Brefeldin A BFT, see Bacteroides fragilis enterotoxin Bi-ionic potentials, 72, 82 Bile acids, 576 Bile duct ligation (BDL), 580, 582, 586 7H6 and, 582 -induced cholestasis, ZO-1 in, 590 occludin and, 582 reversibility, 585 ZO-1 and, 582 Bile-to-plasma ratios, 588–589 Bile plugs, 580 Bile salts, 355 Bile secretion, 576 Biliary clearance, 578 Blastocyst, 286, 285 cavitation, 291 formation, 291, 295 Blastula formation, 293, 295 Blind loop(s) self-filling, 562 syndrome, 562 Blood–aqueous barrier, 659, 662, 663, 677 Blood-to-bile ratio, 578 Blood–brain barrier (BBB), 48, 49, 54, 415, 416, 699 adherens junctions, 420 anatomy, 417 cell culture model, 422 culture models of endothelial, 422 delivery, 705 development, 420 molecular basis, 419 TJ dysfunction in, 699 protein composition, 419 Blood–brain barrier, unique aspects of, 415–443 development, 420–436 endothelia, 420–425 epithelia, 425–436 protein composition, 419–420 structure of different regions, 416–419 endothelia, 416–417 epithelia, 417–419 Blood–brain permeability barrier, 47 Blood–milk potential, 398, 406 Blood–ocular barrier, 654, 657 Blood–retinal barrier, 49, 657, 664, 665, 677 Blood–testis barrier, 49, 618, 732

Index Blood–tissue barrier, 416, 608 Blow fly, 54 Bovine chronic interstitial nephritis, 488 Bowman’s membrane, 656 BPI, 359 Brain angiogenesis inhibitor (BAI), 240 leptomeninges surrounding, 729 Brefeldin A (BFA), 155 Bridge function, of TJ, 199 Bromocryptine, 401 Bromodomain, 247 Bromosulfophthalein (BSP), 588 Brush-border myosin, 243 BSP, see Bromosulfophthalein

C Ca2+ activated cell contacts, 171 depletion, 466 -independent cell adhesion molecules, 173 Cable analysis, 65 Caco-2, 462, 471, 508, 521, 522, 566 cell line, 337 drug delivery model, 698 intestinal epithelial cell line, 342 Cadherin(s), 122, 429, 633 -5, 420 adhesion, 295 –catenin complex, 131 E-, see E-cadherin Caenorhabditis elegans, 241, 278, 374 Calcitonin, 485 Calcium ionophore A23187, 641 ions, 466 switch, 269 Calliphora, 54 Calmodulin (CaM), 9, 171 CaM, see Calmodulin cAMP, 424, 432 Canalicular bile salt export pump, 587 Canalicular permeability, 578 Canaliculi, 576, 577 Cancer, 590 Canoe, 241, 277 CAP-57, 359, 360 Capacitive shunt current, 738 Capillaries, 666 Capillary wall, 350 Carmustin, 588 Caspr/neurexin IV, 736 Cathepsin G, 359

749 Cationic proteins, 359, 362 α-Catenin, 129, 177, 236, 273, 404 β-Catenin, 127, 180, 181, 245, 296, 404, 420, 658 γ-Catenin, 420 Caveolin-1, 187 CD, see Celiac disease Cdc42, 129, 243 Celiac disease (CD), 406, 703, 704, 711 Celiac sprue, 559 dilution potential, 563 intestinal permeability, 560 tight junction changes, 559 ultrastructural changes, 561 Cell(s) adhesion, see Cell–cell junctions and cell adhesion, general themes in adhesion molecules, 676 barrier, vascular endothelial, 665 basal, 731 CaCo-2, 517, 566 –cell junctions, in gene expression and differentiation, 380 cycle, G1 to S-phase transition during, 384 embryonic stem, 216 endothelial choroidal arteriole, 666 permeability in microvascular, 676 Schlemm’s canal, 668, 669, 671, 673 gaps, interendothelial, 672 germ, lineage, 731 glial, 40, 47 growth, 384 HT-29/B6, 565, 568, 570 HT-29cl.19A, 566 L, 523 Leydig, 733 loss, trabecular, 676 Madin–Darby canine kidney, 200, 215, 525, 526, 538, 632 –matrix adhesions, 676 motility, 170 nonpigmented, 659 perineurial, 41 polarity, 171, 182, 509, 585 epithelial, 172 loss of, 536 proliferation, 589 retinal capillary endothelial, 665 Sertoli, 731, 732, 739 spermatocyte precursor, 732 T-, 733 T84, 566, 568 tracheal epithelial, 635 Vero, 522

750 Cell–cell junctions and cell adhesion, general themes in, 121–144 desmosomes, 131–135 desmosomal cadherins, 131–132 desmosomal cytoplasmic interactions, 132–133 intermediate filament binding proteins, 133–135 molecular structure and assembly of adherens junctions, 122–131 actin remodeling and cell compaction, 129 adhesive trans-dimerization of cadherins, 125 cadherins, 122–123 lateral cadherin cis-dimer formation in plane of membrane, 123–124 role of adherens junction in initiating cell–cell contact, 123 role of cadherin juxtamembrane domain and p120 in lateral dimerization and clustering, 125–128 role of catenins in harnessing of cytoskeleton, 128–129 plakoglobin, 133 regulations of cell adhesion, 129–131 role for junctional proteins in signal transduction and gene regulation, 135–136 Cell surface lipid polarity, tight junctions and, 305–332 generation of cell surface lipid polarity, 316–323 lipid synthesis, 316–317 lipid transport and sorting, 317–318 monomeric lipid transport and cell surface lipid polarity, 321–322 transmembrane lipid asymmetry and cell surface lipid polarity, 322–323 vesicular traffic to apical and basolateral plasma membrane domains, 318–321 lateral lipid macrodomains not stabilized by cell–cell contacts, 314–316 myelin and neuronal cells, 314–315 other polarized cells, 316 spermatozoa, 315–316 lipids heterogeneously distributed over cellular membranes, 306–308 lipid heterogeneity within single membrane, 307–308 lipids of different cellular membranes, 306–307 major lipid classes in mammalian membranes, 306

Tight Junctions surface polarity of lipids, 308–310 apical lipid composition, 308–309 epithelial surface domains, 308 protective functions of glycosphingolipids, 309–310 tight junctions, 310–314 how tight junctions affect lipid diffusion, 312–314 lateral diffusion, 310 structure, 310–312 Cellular membrane, distribution of junctional particles within plane of, 613 Cellular pathway, 64 Centipedes, 49 Central nervous system (CNS), 98, 415, 699, 729 avascular, 47 endothelia, 416 epithelia, 416 myelin, 699, 734, 740 Cephalochordates, 53 CFTR, see Cystic fibrosis transmembrane conductance regulator Charge selectivity, 578, 579, 584 Chitosan, 707, 708 Chloride, 405, 406 cell(s), 447 cultures without, 453 cultures with, 453 freshwater, 451 seawater teleost, 447, 450 secretion, current model for, 449 permeability to, 397 Cholangitis, 589 Cholelithiasis, 580 Cholera toxin (CT), 475, 501, 520 Cholestasis, 580, 581, 583, 586 bile acid–induced, 587 drug-induced, 587–589 ethinylestradiol, 586–587 extrahepatic obstruction, 580–586 inflammatory disease, 589 oxidative stress, 589 toxin-induced, 587–589 ZO-1 in BDL-induced, 590 Cholesterol, 306, 608 Choriocapillaries, 666 Choroidal arteriole endothelial cells, 666 Choroidal vasculature endothelium, 666 Choroid plexus development, culture models, 425 Chronic interstitial nephritis with fibrosis (CINF), 488 Cigarette smoke exposure, 646 Ciliary body, 654, 659 Ciliary epithelium, 360, 660, 661

Index Ciliary nonpigmented epithelium, 677 CINF, see Chronic interstitial nephritis with fibrosis Cingulin, 23, 168, 272, 277, 375 assembly, 288, 294 binding sites, 289 Cirrhosis, North American Indian childhood, 589 c-Jun, 478 Class V myosins, 241 Clathrin adaptor complex, 167 Claude Hypothesis, 32, 62, 77, 105, 221, 489 Claudin(s), 8, 75, 168, 179, 236, 523, 632, see also Occludin and claudins carboxyl termini of, 728 family, 725 freeze-fracture characteristics of, 218 gene(s), structure, 725, 726 identification of, 216 interaction of with protein, 224 intron containing, 725, 727 mediation of specific paracellular fluxes in vivo by, 483–492 multiple, 591 phylogenetic analysis of, 726 normal Mg2+ homeostasis, 484–486 transepithelial electrical resistance and, 222 paracellin-1 and paracellular channels, 489–490 positional cloning of gene causing recessive renal hypomagnesemia, 487–489 recessive renal hypomagnesemia with hypercalciuria and nephrocalcinosis, 486–487 single exon, 725, 727 structure–function properties of, 98 Claudin-1, 240, 288, 291, 407, 523, 555 Claudin-2, 524 Claudin-3, 523 Claudin-4, 407, 523 Claudin-5, 419, 421, 524 Claudin-6, 523 Claudin-7, 523 Claudin-8, 523 Claudin-10, 524 Claudin-11, 728, see OSP/claudin-11-containing parallel tight junctions, functions of claudin-11-null mice, neurological phenotype of, 737 Claudin-14, 523 Claudin-16, 8, 13, 725 bovine chronic interstitial nephritis, 488 human recessive renal hypomagnesemia, 488 positional cloning, 487 selective paracellular ion conductance, 489

751 Caudin-18, 726 CLAUDIN genes, clustering of, 727 Cl–/HCO 3– exchanger, 179 Clostridium botulinim toxin, 496 difficile, 708 toxin, 495, 564 toxin A, 269 toxin B, 269 perfringens enterotoxin (CPE), 98, 407, 518 receptors, 520, 522 -containing complexes, 521 -induced cytotoxicity, 521 Clostridium perfringens enterotoxin, interactions between tight junction proteins and, 517–532 consequences of CPE, 525–526 CPE interaction with tight junction proteins, 522–525 CPE interaction with occludin, 524–525 early studies of CPE receptors, 522–523 identification of claudins 3 and 4 as functional CPE receptors, 523–524 CPE mechanism of action, 519–522 CPE effects on gastrointestinal tract, 519–521 CPE effects on mammalian cells, 521–522 current model for CPE action, 527–529 importance of CPE-induced changes in tight junction structure for CPEassociated gastrointestinal disease, 526–527 role of CPE in human gastrointestinal disease, 518–519 Clustering, 180 CNS, see Central nervous system Coiled-coil region, 244 Colchicine, 475, 589 Collagen, 676 Colloid-osmotic equilibrium, 521 colo 316 cells, 473 Colon adenocarcinoma, 461 Comma 1D cells, 409 Compaction, 287, 288 Conductance measurements, local, 570 paracellular, 66 scanning technique, 570 transcellular, 66 Connexin, 731 Connexin-43, 236, 609 Continuous tight junctions, 669 Cornea, tight junctions in, 655

752 Corneal endothelium, 658, 676 Corneal epithelium, 656 Cortactin, 236, 273, 276 Cortical cytoskeleton, 266 CPE, see Clostridium perfringens enterotoxin Cranial nerves, 729 Crohn’s disease, 558, 704 Cross-linking, induced by glutaraldehyde fixation, 53 Crustacea, 49 Crypt–villus axis, TJ morphology, 562 CstF-64, 245 CT, see Cholera toxin C3 transferase, 270 Cul de sacs, 668 Cyclic AMP, 475 Cycloheximide, 409, 468 Cyclosporin A, 587 Cystic fibrosis transmembrane conductance regulator (CFTR), 450 Cytochalasin(s), 267 B, 588 D, 202, 268, 269 TJ formation inhibited by, 475 Cytokine(s), 357, 630 -activated HUVEC monolayers, 635 knockout models, 565 regulation, 676 Cytolytic proteins, 357 Cytoplasmic plaque proteins, tight junction, 231–264 architecture of cytoplasmic plaque of TJ, 251–252 cytoskeletal proteins, 248–249 actin, 248–249 spectrin, 249 GTP-binding proteins and protein kinases, 249–250 G proteins, 250 protein kinases, 250 Rab proteins, 249–250 non-PDZ and other TJ plaque proteins, 243–248 ASH1, 247 BG9.1 antigen, 220-kDa protein, 7H6, 19B1, Sec6/8, 248 cingulin, 243–244 protein 4.1R, 247–248 symplekin, 244–246 ZONAB, 246–247 nuclear localization of TJ plaque proteins, 252–253 phosphorylation of TJ plaque proteins, 251 role of TJ cytoplasmic plaque proteins in TJ function, 252

Tight Junctions TJ plaque proteins containing PDZ domains, 232–243 AF-6, 241–242 ASIP/PAR-3 and PAR-6, 242–243 MAG-1/BAP1, 240–241 ZO-1, 232–237 ZO-2, 237–240 ZO-3, 240 TJ plaque proteins and disease, 253–254 Cytoskeleton, 44, 232 apical actin, 498 components, 46, 202 cortical, 266 elements, 55, 676 regulation, 494 Cytotoxicity, 356, 521

D dbpA, see DNA-binding protein A Deafness, and claudin-14 mutation, 731 Defensins, 359 Demyelination, 736, 739 2-Deoxyglucose, 269 Desmocollins, 131 Desmogleins, 131 Desmoplakins, 134 Desmosomal cadherins, 131 Desmosomes, 131, 168, 287 Detergent-insoluble glycolipid-enriched membranes (DIGs), 28 Detergent-insoluble membrane rafts, 307 Detergent-resistant membrane fragments (DRMs), 28 Developmental assembly, tight junction, 285–303 regulation of tight junction assembly during early Xenopus development, 293–295 tight junction proteins as regulators of cell differentiation during development, 295–296 trophectoderm model of tight junction assembly, 286–293 cell adhesion and regulation of tight junction assembly, 292–293 molecular maturation of tight junction during trophectoderm differentiation, 287–292 Developmental studies, 46 Dexamethasone, 401, 409, 406, 672, 673 Diarrhea, 519 antibiotic-associated, 519 infectious, 564

Index leak flux, 554 sporadic, 519 Dicentrarchus labrax, 453 Diffusion, 206 barrier, 47, 310 exchange, 106 potentials, 72 DIGs, see Detergent-insoluble glycolipidenriched membranes Dileucine-dependent motifs, 167 Dilution potentials, 72, 74 Dinitrophenol (DNP), 474 Dissolution, of poorly soluble drugs, 698 dlg tumor suppressor gene product, 234 DMEM, see Dulbecco Modified Eagle’s Medium DNA-binding protein A (dbpA), 246 DNP, see Dinitrophenol Docking, 177, 178 DRMs, see Detergent-resistant membrane fragments Drosophila, 171, 725 fat facets gene, 242 first-instar larvae, 739 melanogaster, 234 ash1, 247 protein, 241 tamou gene, 236 Na+,K+-ATPase, 179 septate junctions in, 736, 739 tumor suppressor, 156, 383 Drug delivery, oral, 685, 707 pulmonary route of, 706 TJ modulation for, 705 transdermal, 707 transmucosal, 706 via nasal route, 706 Drug development, implications of transport via paracellular pathway on, 685–695 alternative approaches for enhancing absorption, 689–691 approaches to enhance paracellular permeability, 688–689 dissolution rate, 689 Drug-induced cholestasis, 587 Dulbecco Modified Eagle’s Medium (DMEM), 462, 464 Duodenum, 558

E Early peak bile secretion, 584, 585 E-cadherin, 157, 168, 355, 404, 497 adhesion, 296 /catenin

753 adhesion system, 287 protein complex, 286 -mediated cell–cell contacts, 178 newly synthesized, 181 ECM, see Extracellular matrix ECP, see Eosinophil cationic protein Edge effect, 66 EDN, see Eosinophil-derived neurotoxin EDTA, see Ethylenediaminetetracetic acid EE, see Ethinylestradiol EF, see Exoplasmic face E-fracture face, 23, 42, 421 EGTA, 405 EHEC, 507 Elastase, 355, 499 Electrical potential, measurement of, 397 Electrical resistance, 100, 220 Electrical transepithelial potential, 576 Electrogenic Cl– transport, 70 Electromotive forces (EMF), 63 Electron microscope (EM), 39 Electron microscopy, freeze-fracture, 213, 402, 555, 632 Electron-opaque tracers, 621 ELISA, 618 EM, see Electron microscope Embryogenesis, actin cytoskeleton regulation during, 277 Embryonic stem (ES) cells, 216, 728 EMF, see Electromotive forces EnaC, 353 Endocochlear potential, 730 Endocytosis, 473 Endoplasmic reticulum (ER), 146, 316 Endothelial blood-brain barrier, culture models of, 422 Endothelial cell(s) choroidal arteriole, 666 line, 423 permeability in microvascular, 676 Schlemm’s canal, 668, 669, 671, 673 Endothelial junctions, factors regulating, 424 Endothelial monolayers, isometric tension generation by, 644 Endothelial TJs, 12 Endothelial tricellular corners, 636 Endothelium, 416 corneal, 676 development, environmental interactions during, 421 junctional complexes, 417 Entamoeba histolytica, 498 Enterocyte effacement, locus of, 504 Enteropathogenic Escherichia coli (EPEC), 270, 504, 506

754 Eosinophil(s), 358 cationic protein (ECP), 358 -derived neurotoxin (EDN), 358 peroxidase (EPO), 358 proteins, 362 Epaxial muscles, 729 EPEC, see Enteropathogenic Escherichia coli Ependyma, 419 EphB3, 242 Eph-related receptor tyrosine kinases, 242 Ephrins, 243 Epidermal growth factor, 278 Epididymis, see Testis, epididymis, and vas deferens, tight junctions in Epithelial cell polarity, 172 Epithelial cells, protein targeting pathways and sorting signals in, 145–164 apical and basolateral sorting signals directing polarized protein trafficking, 147–150 apical sorting signals interacting with specialized lipid microdomains, 150–151 basolateral sorting signals interacting with specialized cytoplasmic adaptors, 151–152 control of polarity by small GTPases, 155–156 generation and maintenance of polarity in epithelial cells, 145–146 intracellular sorting and polarized delivery of proteins to cell surface, 146–147 passive mechanisms accounting for polarized distribution of membrane and cytoskeletal proteins, 157–158 role of actin cytoskeleton in Golgi exit and arrival at cell surface, 153 role of microtubule motors in exit from TGN and transport to plasma membrane, 152–153 tight junction/zonula adherens, 156–157 Epithelial–mesenchymal transitions, 122, 383 Epithelial migration sites, role of fibroblasts in guiding neutrophils to preferred, 643 Epithelial permeability, see Extracellular macromolecules, epithelial permeability modulated by Epithelial polarity, biogenesis of tight junctions and, 165–195 biogenesis of epithelial cell polarity, 171–179 assembly of cytoskeleton proteins and signaling complexes at sites of cell contacts, 175–177

Tight Junctions external signals inducing cell surface asymmetry, 171–175 structures specifying targeting and retention of membrane proteins synthesized de novo, 177–179 cytoskeleton, 170 dynamics of cell membrane polarity, 182 epithelial cell adhesion, 168–170 cell–cell contacts, 168–169 cell–cell contacts and signaling, 169–170 cell–extracellular matrix contacts, 169 polarity of adherens junction proteins, 181 polarity-maintaining mechanisms, 166–168 selective stabilization, 167–168 selective targeting, 167 polarity of tight junctions, 179–1781 relationship between tight junctions and adherens junctions, 182–188 cell junction proteins recruited to initial lateral membrane, 185 epithelial contacts and polarity maintained by selective targeting and stabilization, 187–188 establishment of structural and molecular asymmetry at cell surface, 182–183 membrane proteins stabilized by membrane skeleton, 186–187 role of Sec6/8 complex in initial stage of cell–cell contact, 183 TJ as targeting patch, 187 ubiquity of polarity, 166 Epithelial polarization, process of triggered by extracellular Ca2+, 182 Epithelial regions, development of, 418 Epithelial resistance, 555, 556 Epithelial tight junctions, 525 Epithelium as barrier, 350 of choroid plexus, 416, 419 ciliary, 360, 660, 661 corneal, 656 development, 425 intestinal, 406, 521, 701 leaky, 4, 68, 69, 102 low resistance of, 4 mammary, 408 nasal, 729 nonpigmented, 663 opercular, 449 paracellular ionic permeability ratios of, 75 pigmented, 659, 664 seminiferous, 600 tight, 68, 102

Index EPO, see Eosinophil peroxidase Equivalent-circuit analysis, 63 ER, see Endoplasmic reticulum ErbB-2, 246, 384 ES cells, see Embryonic stem cells Escherichia coli, 341 E-selectin, 630, 642 Ethanol, 355 Ethinylestradiol (EE), 586, 586 Ethylenediaminetetracetic acid (EDTA), 706 Evanescent field microscopy, 157 Evolutionary conservatism, 725 Exchange diffusion, 106 Exocyst, 156, 180 Exocytosis, 187 Exogenous molecules, 40 Exoplasmic face (EF), 459 Exotoxin A, 499 Extracellular fluids, passage of, 608 Extracellular macromolecules, epithelial permeability modulated by, 349–366 alteration by proteases, 353–355 pathophysiological role of proteases, 354–355 physiological role of proteases, 353–354 cationic proteins, 362–363 epithelium as barrier, 350–353 modulation by cellular constituents, 360–362 histones, 361–362 protamine, 360–361 modulation by leukocytes, 357–360 eosinophil proteins, 358–359 neutrophil proteins, 359–360 xenobiotics, 355–357 bacterial, 356–357 nonbacterial, 355–356 Extracellular matrix (ECM), 168, 676 Extracellular pathway, 352 Extramacrochaetae, 236 Eye, anterior chamber of, 666

F F-actin formation, 645 paracellular permeability increased by redistributing, 510 FAK, see Focal adhesion kinase Fam deubiquinating enzyme, 242 Fascia adhaerens, 3 Fascia occludens, 460, 462 Fast freezing, 44

755 Feedback inhibitor, 405 Fence function, 199, 202, 578 Fer, 170 Fibril(s) arthropod TJs associated with, 46 freeze-fracture, 219 junctional, 609, 610, 612 morphology of intramembranous, 725 subunits, asymmetric, 44 Fibroblast(s) expression, 223 L-cell, 225 role of in guiding neutrophils to preferred epithelial migration sites, 643 Filipin, 473, 611 Filopodia, 183 Flippase, 322 Fluid convection, 206 Fluid/electrolyte absorption, 520 secretion, 520 transport alterations, 520 Fluorescence emission intensity ratio, 203 Fluorescence resonance energy transfer (FRET), 202 Fluorescent tracers, 398 Focal adhesion kinase (FAK), 170 Fodrin, 616 /ankyrin membrane skeleton, 186 -based membrane skeleton, 178 Food allergies, 702 Foodborne human gastrointestinal diseases, 519 Foregut, 729 Formyl peptide, 641 Forskolin, 475 Forssman antigen, 312 Fracture face, 43, 620 labeling technique, 26 plane, 44 Fragilysin, 270 Freeze-cleaved replicas, 41 Freeze fracture, 91, 421, 427 complexity of junctional strands, 424 electron microscopy, 213, 402, 473, 555, 632 images, of mammary epithelium TJs, 403 P face, 424, 432 replicas, 459, 577 of guinea pig tracheal epithelium, 646 of hepatocellular TJs, 581 strand complexity, 432 studies, of freshwater chloride cells, 451 techniques, 23, 462

756 Freshwater chloride cells, freeze-fracture studies of, 451 environment, 446 FRET, see Fluorescence resonance energy transfer Frog skin, 1 Fundulus heteroclitus, 449, 450, 451, 452 Fusion of basolateral vesicles, 178 machinery, 156

G Gα12, 23 Gαi-2, 250 Gαo, 250 -Galactosidase, 729 Galactosylceramide (GalCer), 306 GalCer, see Galactosylceramide Gallbladder, Necturus, 360 Gap junction(s), 40, 45, 416, 419, 427, 731 Cx43 positive, 609 particles, 44 Gastrointestinal disease, 518, 519 Gate function, 199, 202, 576 GDNF, see Glial-derived neurotrophic factor GEMs, see Glycolipid-enriched membranes Gene expression and differentiation, cell–cell junctions in, 380 regulation, 135 Genistein, 565 Germ cell(s) access to through Sertoli cells, 606 development of, 609 lineage, 731 passage or translocation of cohorts of, 609 GFP, see Green fluorescent protein GFR, see Glomerular filtration rate Giant vacuoles, 668 Gitelman’s syndrome, 487 GL, see Glycerophospholipids Glaucoma medications, 661, 669 open-angle, 669 role of tight junctions in development of, 666 steroid-induced, 669, 673 Glaucoma, ocular tight junctions in health, disease, and, 653–683 blood–ocular barriers, 657–666 blood–aqueous barrier, 659–663 blood–retinal barrier, 664–666

Tight Junctions role of tight junctions in development of glaucoma, 666–676 aqueous outflow system, 666–667 hormonal regulation of transendothelial fluid flow, 669–671 patterns of gene expression associated with changes in transendothelial fluid flow, 673–676 potential barriers to flow of aqueous humor out of eye, 668–669 rate of transendothelial fluid flow correlates with changes in tight junctions, 672 tight junctions in trabecular meshwork and Schlemm’s canal endothelial cells, 669 tight junction structure in Schlemm’s canal endothelial cells, 671 ZO-1, 672–673 tight junctions in cornea, 655–657 GlcCer, see Glucosylceramide Glia, 48 Glial cell(s) layer, 40 modified, 47 Glial-derived neurotrophic factor (GDNF), 424 Glomerular feedback, 535 Glomerular filtration rate (GFR), 535 Glomeruli, 251 Glucagon, 485 Glucocorticoids, 396, 400, 408, 669 Glucosamine, 434 Glucose transporter, 662 Glucosylceramide (GlcCer), 306 GLUT-1, 662, 664 γ-Glutamyl transpeptidase, 664 Glycerophospholipids (GL), 306 Glycocalyx, 308, 350, 351 Glycolipid-enriched membranes (GEMs), 28 Glycosaminoglycan layer, 341 Glycosylphosphatidylinositol (GPI) anchors, 167 Golgi apparatus, 152 Golgi complex, 152 trans-Golgi network (TGN), 146, 167 GPIIB/IIIa, 687 GPI anchors, see Glycosylphosphatidylinositol anchors G protein(s), 478 heterotrimeric, 370 receptors, 9 Green fluorescent protein (GFP), 209 Growth hormone–releasing peptide, 687 GST fusion protein, 246

Index GTPase(s) in ischemia, 545 rho family, 155, 170, 176 small, 370, 372 GTP-binding proteins, 175, 368 Guanylate kinase (GUK), 168, 234 homologies, 419 phylogenetic comparisons, 419 GUK, see Guanylate kinase

H 7H6, 23 HC11, 409 Heavy meromyosin (HMM), 46 Helicobacter pylori, 507, 508 Hemolymph composition, regulation of, 53 –nerve barrier, 739 Henle’s loop, 360 Hensin, 173 Hepatectomy, 589 Hepatocyte growth factor/scatter factor (HGF/SF), 296 Hepatocytes, 576, 577 Heterotrimeric G proteins, 369, 370 HGF/SF, see Hepatocyte growth factor/scatter factor Histamine, 406, 640, 641 Histone H4, 363 Histones, 361 Histopathological damage, 521 HIV enteropathy, 559, 564, 565 HLA, see Human leukocyte antigen HMM, see Heavy meromyosin Homotypic interactions, 182 Hormonal regulation of fluid movement, 672 of pregnancy to lactation transition, 398 Horseradish peroxidase (HRP), 11, 420, 579, 620, 621 early peak, 588, 589 inulin, 432 mannitol, 432 permeability, 584 Horseshoe crabs, 49 Host–pathogen interactions, 493 HRP, see Horseradish peroxidase HT29, 461 /B6 cells, 565, 568, 570 cl.19A cells, 566 differentiation and apoptosis induced by microtubule inhibitors, 475

757 HuASH1, 385 Human leukocyte antigen (HLA), 702 Human umbilical vein endothelial cell (HUVEC) monolayers, 634 cytokine-activated, 635 human neutrophil migration at tricellular corners on, 637 Huntingtin, 244 HUVEC monolayers, see Human umbilical vein endothelial cell monolayers Hydraulic conductivity, 672 Hydrostatic pressure, 586 Hypermagnesemia, 486 Hyperregenerative transformation, small intestinal mucosa, 560 Hypertonic salt solutions, 462 Hypomagnesemia, 728 Hypophysectomy, 401 Hysterectomy, 401

I ICAM-1, see Intercellular adhesion molecule-1 ICM, see Inner cell mass Id-1, 409 IDDM, see Insulin-dependent diabetes mellitus IF, see Intermediate filaments IFNδ, see Interferon-δ IgA NP, see Immunoglobulin A nephropathy IHRC, see Isolated rat hepatocyte couplets IL-8, Weibel–Palade bodies capable of storing, 641 Ileum, 519 Immune-privileged components, 737 Immune surveillance, 732, 736 Immunocytochemistry, 403 Immunofluorescence microscopy, 200, 618 Immunoglobulin A nephropathy (IgA NP), 703 superfamily, 630, 633 Immunogold labeling, of Forssman glycolipid, 313 Immunolabeling, see Ultrastructure and immunolabeling of tight junction Immunological barrier, to anti-sperm antibodies, 618 Impedance analysis, 554 IMPs, see Intramembranous particles Indirect immunofluorescence microscopy, 408 Infectious diarrhea, 564 Inflammation, freeze-fracture images of animal tissues with, 663 Inflammatory bowel diseases, 704

758 Inflammatory cascade, Yersinia, 510 Inflammatory diseases, 589 Inner cell mass (ICM), 286, 287 Inner ear, 729, 737, 740 Inositolphospholipids (PI), 306 Insecta, 49 Insects, 40, 41 Insulin, 485, 714 Insulin-dependent diabetes mellitus (IDDM), 703, 711, 713 Integrin, 630, 675, 676 focal adhesion mediated by, 168 two groups of, 630 β1-Integrin, 509 Intercalated cells, of collecting tubule, 173 Intercalated disks, 122 Intercellular adhesion molecule-1 (ICAM-1), 631 Interendothelial cell gaps, 672 Interferon-γ (IFNγ), 269, 406, 566, 567, 568, 700 Intermediate cellular compartment, theory of presence of, 617 Intermediate complex, ~135 kDa, 522 Intermediate filaments (IF), 170 Interstitial space, 398 Intestinal barrier defect, 702 Intestinal epithelia, 406 Intestinal epithelium desquamation of, 521 TJ dysfunction in, 701 Intestinal inflammation, tight junctions in, 553–574 infectious intestinal diseases and epithelial barrier function, 564–565 HIV enteropathy, 564–565 infectious diarrhea, 564 lack of tight junction changes in short bowel syndrome, 563–564 mechanisms of tight junction alteration in ulcerative colitis, 557 tight junction changes in blind loop syndrome, 562–563 tight junction changes in celiac sprue, 559–562 diarrhea, 562 intestinal permeability, 560 tight junction modulators, 561 tight junction structure, 560–561 tight junction changes in Crohn’s disease, 558–559 intestinal permeability in Crohn’s disease, 558 tight junction molecules in Crohn’s disease, 559 tight junction morphology in Crohn’s disease, 558–559

Tight Junctions tight junction changes in ulcerative colitis, 554–557 epithelial permeability in ulcerative colitis, 554–555 freeze fracture electron microscopy, 555 tight junction molecules in ulcerative colitis, 555 tight junction down-regulation by proinflammatory cytokines, 565–570 interferon-γ in T84 cells, 567–568 occludin promoter activity in response to proinflammatory cytokines, 568–570 TNFα in CaCo-2, T84, and HT-29cl.19A cells, 566–567 TNFα in HT-29/B6 cells, 565–566 tight junctions and apoptosis, 570–571 Intestinal permeability, 560, 708 Intestinal villi, 253 Intestine, 12, 701 absorption of nutrients in, 333 compound transport by, 685 Intracellular calcium, 278 Intracellular signaling, in classical and new tight junction functions, 367–394 cell–cell junctions in gene expression and differentiation, 380–385 cell–cell junctions and transcription factors, 383–385 Ras-mediated transformation and junctional complex, 380–383 experimental systems, 368–369 GTP-binding protein pathways, 369–373 heterotrimeric G proteins, 369–372 small GTPases, 372–373 phosphorylation of tight junction proteins, 375–379 serine/threonine phosphorylation, 375–377 tyrosine phosphorylation, 377–379 protein kinase A pathway, 374 protein kinases C pathway, 373–374 regulation of paracellular permeability by multiple signaling pathways, 379–380 tight junction functions, 367–368 Intramembranous particles (IMPs), 40, 734 Intramembranous patterns, 41 Intraocular pressure (IOP), 660, 667 Intron-containing claudins, 725, 727 Inulin, 11, 432 [3H]inulin, 408

Index Invasin, 509 Invertebrates, tight junctions in, 39–59 assembly of arthropod tight junctions during development, 54 biochemistry of invertebrate tight junctions, 54–55 comparisons between tight junctions in invertebrates, lower chordates, and vertebrates, 51–54 distribution of tight junctions among invertebrates, 40–41 fine structure features of tight junctions, 41–47 coexistence with other junctions, 45–46 cytoskeletal associations, 46–47 in replicas after conventional fixing and cryoprotection, 41–44 in replicas after rapid freezing with no fixation, 44 in thin sections, 41 invertebrate groups that possess tight junctions, 49 junctions peculiar to invertebrates with tight junction-like characteristics, 49–51 reticular septate junctions, 51 retinular junctions, 51 smooth septate junctions, 49–51 models of invertebrate tight junctions, 48 physiological roles, 47–48 cell–cell adhesion, 48 permeability barriers, 47–48 Involution, apoptosis during, 401 Ion(s) channels, 171, 728 permeability, 75 selectivity, 72 TJ discriminating among, 7 IOP, see Intraocular pressure IQGAP, 129 Iris, 654 pigmented epithelium of, 662 stroma, 663 Ischemia actin cytoskeleton, 541 cadherin, 544 GTPase family members in, 545 loss of lipid polarity, 538 models ATP depletion with 2-deoxyglucose, 535 renal artery clamp, 535 Isoelectric point, 7 Isolated rat hepatocyte couplets (IRHC), 579

759

J Jagged, 242 JAM, see Junction adhesion molecule Jaspis johnstoni, 509 Junction adhesion molecule (JAM), 21, 168, 289, 312, 631, 633 coimmunoprecipitation, 236 -mediated homophyllic adhesion, 240 assembly, 40 -associated actin, 616 complexes, endothelia, 417 depth, 578 fibrils, 609, 610 number of or pattern within zonules, 610 proliferation of, 612 particles, distribution of within plane of cellular membrane, 613 permeability, 614, 616, 620 proteins, proteolytic degradation of, 634 Juxtacanalicular tissue (JXT), 668, 669 JXT, see Juxtacanalicular tissue

K Kallikrein, 354 Kidney, ischemia-induced tight junction dysfunction in, 533–551 kidney, 533–535 renal ischemia, 535–536 tight junction function distrupted by renal ischemia, 536–538 tight junction regulation and ischemia, 538–545 actin cytoskeleton dysfunction and tight junction regulation, 539 GTPase regulation and tight junction dysfunction, 540–541 other regulatory pathways, 541–545 protein kinases and tight junction dysfunction, 539–540 Knockout mouse, see OSP/claudin-11-containing parallel tight junctions, functions of K-RasB, 240

L α-Lactalbumin, 396 Lactation, 395, 410 initiation of, 399 transition between pregnancy and, 396

760 Lactogenesis, 396 Lactose, 405, 407 lacZ, 729 Laminins, 603 Lampreys, 454 Lanthanum nitrate, 620 Large complex, containing CPE ~155 kDa, 522 ~200 kDa, 522 Lateral intercellular spaces (LIS), 63, 203, 207 L-cells, 224, 241, 523 LDH release, 356 Leak flux diarrhea, 554 Leaky epithelia, 4, 68, 69, 102 Lebistes reticulatus, 451 LEF-1, 170 Leptotene/pachytene, 732 Leucine amino peptidase, 187 -rich-repeat protein, 187 Leukocyte(s), 357 adhesion, endothelial borders as sites for, 640 trafficking, 635 Leukocyte transmigration, relationship between tight junctions and, 629–652 interendothelial clefts, 631–633 mechanisms for opening/remodeling tight junctions, 644–646 multistep-adhesion cascade, 630–631 neutrophil adhesion to endothelial borders, 639–641 neutrophil migration through interendothelial clefts, 633–635 neutrophil transepithelial migration and tricellular corners, 642–644 neutrophil transmigration at endothelial tricellular corners, 635–638 transcytotic neutrophil migration, 641–642 Leydig cell, 733 Lipid(s) asymmetry, 322 diffusion, 312 distribution, 209 microdomain, 150, 320 polarity, 308 generating, 316 loss of, 538 maintaining, 312 polarized delivery of to cell surface, 146 probes, flip-flop of, 209 rafts, 187, 320 sorting, 317 transfer proteins, 322 transport, 317 Lipopolysaccharide (LPS), 406

Tight Junctions Liquid-junction potentials, 74 LIS, see Lateral intercellular spaces Liver disease, tight junctions in, 575–597 hepatocellular tight junctions, 578–580 morphology, 578 probing of tight junctions in liver, 578–580 role of tight junctions in bile secretion, 576–578 tight junctions in liver disease, 580–590 cholestasis, 580–589 inflammatory diseases, 589 oxidative stress, 589 regeneration and cell proliferation, 589–590 LLC-PK1, 356 Local conductance measurement, 570 Loop of Henle, 728 Lovastatin, 28 LPS, see Lipopolysaccharide L-selectin, 630 Lumen, 622 Luminal domain, 350 LY294002, 409 Lysine-rich sequence, 248

M Macaca mulatta, 716 Macromolecule passage through tight junction, pathological and therapeutic implications of, 697–722 pathological conditions associated with TJ dysfunction, 698–705 blood–brain barrier, 699–700 intestinal epithelium, 701–705 therapeutic use of TJ modulation, 705–708 blood–brain barrier delivery, 705–706 oral delivery, 707–708 transdermal delivery, 707 transmucosal delivery, 706 zonulin system, 708–717 pathology, 711–714 physiology of, 709–711 therapeutical use of, 714–717 Macromolecules, passage through tight junction, 396 Macula adhaerens, 3, 20 Madin–Darby canine kidney (MDCK) cells, 171, 200, 215, 355, 371, 407, 525, 632 model, 538 monolayers, 461 sheets, 468 Magnetic resonance imaging (MRI), 700

Index MAGUK, see Membrane-associated guanylate kinase Major basic protein (MBP), 358, 363 Major histocompatibility complex (MHC), 702 Mammary culture systems, 408 Mammary epithelium, 396, 397, 399, 402, 406, 408 Mammary gland, regulation of tight junction permeability in, 395–414 hormonal regulation of tight junction permeability in in vitro mammary systems, 408–410 glucocorticoids, 408–409 prolactin, 409–410 in vivo regulation of mammary tight junctions by milk stasis, mastitis, and oxytocin, 405–408 mastitis, 406–407 milk stasis, 405–406 oxytocin, 407–408 perspective, 410–411 role for TGF-β, 410 tight junction permeability in mammary glands of pregnant and lactating animals, 397–399 transition from pregnancy to lactation, 399–405 hormonal regulation, 399–402 mechanism of tight junction closure, 402–405 Manduca sexta, 54 Mannitol, 11, 406, 432 [14C]mannitol, 408 [3H]mannitol, 252 MAPK, see Mitogen-actived protein kinase MARCKS, see Myristoylated alanine-rich C kinase substrate Mast cell, 461 Mastitis, 399, 406 MBP, see Major basic protein MDCK cells, see Madin–Darby canine kidney cells Meiosis, initiation of in basal compartment, 607 MEK1, see Mitogen-activated protein kinase kinase Membrane(s) apical, 314 -associated guanylate kinase (MAGUK), 11, 168, 232, 295, 383, 590 basement, 168, 350 basolateral, 314, 499, 500 claudin-1, 288 components, diffusion barrier to, 310 detergent-insoluble glycolipid-enriched, 28 domain, 314

761 glycolipid-enriched, 28 interdigitations, 40 lipids of, 306 microdomains, specialized, 187 presumptive junctional, 53 Mena, 177 Mesonephric ducts, 729 Metalloproteinases, 32 Methyl-β-cyclodextrin, 28 3-O-Methylglucose, 434 MF, see Microfilaments Mg2+, 728 MHC, see Major histocompatibility complex Microarray assay, 673, 675 Microbial pathogens, affecting tight junctions, 493–515 access to basolateral ligands by pathogeninduced alterations in tight junctions, 509–510 disruption of tight junction barrier by pathogen-stimulated signaling events, 503–509 Helicobacter pylori, 507–508 pathogenic Escherichia coli, 504–507 Rotavirus, 508–509 Vibrio cholera, 503–504 disruption of tight junctions by bacterial proteases, 498–503 Porphyromonas gingivalis, 499–501 Pseudomonas aeruginosa, 498–499 Vibrio cholera, 501–503 microbial disruption of tight junctions via targeting of cytoskeleton, 495–498 Bacteroides fragilis, 496–497 Clostridium botulinim toxin, 496 Clostridium difficile toxins, 495–496 Entamoeba histolytica, 498 Microfilaments (MF), 170 Microtubules (MTs), 46, 152, 170, 475 Microvascular endothelial cells, permeability in, 676 Milk components, 395 composition, 397, 401, 407 damaged cells in, 405 ejection, 407 potassium, 407 secretion, 408, 411 stasis, 405 TJ effects on composition, 405 yield, 405, 407 Millipedes, 49 Mitochondria, 361, 447 Mitogen-actived protein kinase (MAPK), 31, 544

762 Mitogen-activated protein kinase kinase (MEK1), 31, 251 ML-7, 341 ML-9, 341 MLC, see Myosin light chain MLCK, see Myosin light chain kinase MLC20 phosphorylation, 504 Model(s) cytokine knockout, 565 drug delivery, 698 invertebrate tight junction, 48 ischemia, 535 MDCK cell, 538 offset double-fibril, 48 streptococcal pneumonia, 638 trophectoderm, 292, 293 Modified glial cells, 47 Monocyte transmigration, inhibition of, 645 Monomeric exchange, 322 Monomeric G proteins, 369 Monosaccharide/mannitol permeability, of cultured RPE, 434 Moth TJ, developmental studies in, 54 Motor proteins, 152 MRI, see Magnetic resonance imaging MRP-8, 406 MS, see Multiple sclerosis MTs, see Microtubules Müller cells, 418, 422 Multicellular complexes, 447 Multiple sclerosis (MS), 699, 704 Multistep-adhesion cascade, 630 Myelin sheath, 734, 738 tight junctions, 734 Myelinogenesis, 737 Myoepithelial cells, 407 Myosin brush-border, 243 nonmuscle, 249 Myosin light chain (MLC), 270 pathway, 370 phosphorylation, 372, 494 Myosin light chain kinase (MLCK), 203, 271, 340, 342, 504, 644–645 Myriapoda, 49 Myristoylated alanine-rich C kinase substrate (MARCKS), 503

N Na-dependent transport proteins, 576 Na+-glucose cotransporter, 334 Na+/H+ exchanger, 71, 339

Tight Junctions Na-independent transport proteins, 576 Na+,K+-ATPase, 64, 168, 171, 536 anchoring, 175 polarity, 179, 182 Na–K–2cl cotransporter, 485 Na+-nutrient cotransport, physiological regulation of tight junction permeability by, 333–347 cytoskeletal regulation of tight junction, 341–343 distal events in Na+-glucose cotransport, 339–341 dynamic regulation of tight junction permeability, 334–337 in vivo evidence for regulation of paracellular absorption by Na+-G cotransport, 336–337 physiological regulation of tight junction permeability in mammalian small intestine, 334–336 proximal signals linking Na+-glucose cotransport to myosin light chain phosphorylation, 337–339 unified model of Na+-nutrient cotransport–dependent tight junction regulation, 343 α-Naphthylisothiocyanate (ANIT), 588 Nasal epithelium, 729 N-cadherin, 420 NCAM, see Neural cell adhesion molecule Nectin, 131 Necturus gallbladder, 7, 65, 79, 206, 360 Nephrin, 108 Neural cell adhesion molecule (NCAM), 150, 157 Neurexin, 242, 739 Neurodegeneration, 736 Neuroepithelium, 182, 417, 418 Neuroglian, 186 Neutrophil(s), 359 activated, 644 adhesion, 640 emigration, formyl peptide and, 641 migration, 630 across HUVEC monolayers, 634 through interendothelial clefts, 633 mucosal-to-serosal direction of, 643 transcytotic, 641 P-selectin and, 639 rolling, 630 transendothelial migration, adherens junctions in, 631 transepithelial migration of, 509, 642 transmigration, 635, 647 NF-IL6, see Nuclear factor interleukin-6

Index NMR, see Nuclear magnetic resonance Nocodazole, 475 Nodes of Ranvier, 734 Nonmuscle myosin, 249 Nonpigmented ciliary epithelial cells, 660 Nonpigmented epithelium, 663 Nonreceptor tyrosine kinases, 9 Nonselective cation channel, 354 Nonsteroidal anti-inflammatory drugs (NSAIDs), 702, 703 Noradrenaline, 422, 432 North American Indian childhood cirrhosis, 589 Notch ligand, 242 NSAIDs, see Nonsteroidal anti-inflammatory drugs N-terminal kinase, 478 Nuclear factor interleukin-6 (NF-IL6), 568 Nuclear localization signals, 252 Nuclear magnetic resonance (NMR), 690 Nucleic acid–binding protein, 236

O OAK, see Occludin-associated kinase Occludin, 8, 75, 185, 187, 272, 273, 291, 375, 404, 424, 497, 505, 555, 614, 662, 724 -associated kinase (OAK), 377 BDL and, 582 cytoplasmic domain of, 430 decrease in, 676 degradation, 501 dephosphorylation, 504, 506 dissociation of from membrane, 509 downregulation, 295 expression of, 421 gene structure promoters, 568 identification of, 214 interaction of with protein, 224 localization of, 201 mouse embryo, 294 neutrophil transepithelial migration and, 643 overexpression of, 223 paracellular permeability increased by redistributing, 510 phosphorylation, 168, 225, 376, 378 promoter, 568 reliance of on basolateral signal, 179 in RPE, 431 structure–function properties of, 95 transepithelial electrical resistance and, 221 Xenopus embryo, 294 Occludin 1B, 215, 226 Occludin and claudins, 213–230

763 claudins, 216–220 distribution of, 218–220 freeze-fracture characteristics of, 218 identification of, 216 sequence comparisons of, 216–218 functional analysis of, 220–224 occludin and claudins as cell–cell adhesion molecules, 222–224 transepithelial electrical resistance, 220–222 interactions of occludin and claudins with other proteins, 224–225 occludin, 214–216 distribution of, 215–216 freeze-fracture characteristics of, 215 identification of, 214 phosphorylation as possible determinant of fibril organization, 225–227 sequence of, 214–215 Occluding junctions, 1, 608, 736, 737 Offset double-fibril model, 48 Okadaic acid, 409 Oligodendrocyte, 734 Oligodendrocyte-specific protein (OSP), 98 Oncorhynchus mykiss, 451, 453 Open-angle glaucoma, 669 Open meshworks, arachnid, 53 Opercular epithelia, 449 Optical methods, for tight junction study, 199–212 previous applications of light microscopy, 200–208 cytoskeleton–tight junction interactions, 202–203 immunofluorescence microscopy, 200–202 solute permeability of tight junction, 203–205 water flow across tight junction, 205–208 prospects for light microscopic methods in study of tight junctions, 208–209 lipid distribution, 209 protein–protein interactions using FRET, 208–209 tight junction dynamics studied with GFP-labeled proteins, 209 Oral dosage formulation, development of, 685 Oral drug delivery, 685 Oreochromis mossambicus, 451 Organ of Corti, hair cells in, 730 Orthologous claudin genes, 728 Osmoregulation, patterns of in seawater and fresh water, 446 Osmotic gradient, 576 Osmotic water transport, 71

764 OSP, see Oligodendrocyte-specific protein OSP/claudin-11-containing parallel tight junctions, functions of, 723–745 CLAUDIN 11, 728–737 tight junctions of central nervous system myelin, 734–737 tight junctions of Sertoli cells, 731–733 tight junctions of stria vascularis, 730–731 claudin-11 conserved in different tissues, 737–740 claudin family of integral membrane proteins, 725–727 paracellular diffusion regulated by claudin family members, 728 transmembrane tight junction proteins, 724–725 Ouabain, 204 Ovariectomy, 400, 405 Oxidative stress, 589 Oxytocin, 396, 407

P p120, 125, 127 PAF, see Platelet activating factor PAMR, see Perijunctional actomyosin ring PAR-3, 243 Paracellin-1, see Claudins, mediation of specific paracellular fluxes in vivo by Paracellular barrier, physical, 724 Paracellular channels, 489 Paracellular diffusion barrier, 608 Paracellular Mg2+ flux, see Claudins, mediation of specific paracellular fluxes in vivo by Paracellular pathway, 350, 352, 450, 686 calcium reabsorption, 485, 486 magnesium reabsorption, 485, 486 passage of tracers through, 100 role of in kidney function, see Claudins, mediation of specific paracellular fluxes in vivo by Paracellular permeability, 252, 379, 526, 580, 584, see also Macromolecule passage through tight junction, pathological and therapeutic implications of Paracellular resistance, 68 Paracellular route diffusion of solutes between cells via, 19 limited by TJ, 7 Paracellular water transport, 78

Tight Junctions Parallel-plate flow chamber, 635 Parallel tight junctions, see OSP/claudin-11containing parallel tight junctions, functions of Paralogous claudin genes, 728 Paranodal junctions, 735 Paranodal loops, 734 Parathyroid hormone (PTH), 485 Pars plana, 660 Pasteurella multocida, 372 Pathogenic viruses, 362 Pathogens, see Microbial pathogens, affecting tight junctions P-cadherin, 420 PD09805, 409 PDZ, 187 -binding motif, 224 homologies, 419 phylogenetic comparisons, 419 proteins, 167 PDZ2, ZO-1 homologies, 419 phylogenetic comparisons, 419 PDZ3, ZO-1 homologies, 419 phylogenetic comparisons, 419 PE, see Phosphatidylethanolamine PECAM-1, see Platelet–endothelial cell adhesion molecule-1 PEG, see Polyethylene glycol Pemphigus foliaceus, 131 Pemphigus vulgaris, 131 Peptide transporter, 686 Periaxonal space, 737 Pericytes, 416, 417, 422 Perijunctional actin–myosin ring, 202 Perijunctional actomyosin ring (PAMR), 266, 271, 494 contraction of, 340 linkage between TJ permeability and, 340 Perineurial cells, 41, 44, 48 Perineurial layer, TJs of developing blow fly, 54 Perineurium, 40, 41, 47 Periodontal diseases, 499 Peripheral proteins, 616 Permeability alterations, 521 barriers, 48, 49 canalicular, 578 epithelial, 401 HRP, 584 intestinal, 560, 708 ionic, 75 junctional, 614, 616

Index measurement in vivo, 397 paracellular, 252, 379, 526, 580, 584, 688 solute, 203 subsurface or cortical actin, 616 TJ, 340, 400, 407 tracer, junction, 620 water, 77, 79 Permeability to ions and water, tight junction, 61–88 electrical resistance of tight junctions, 63–72 epithelia distinguished from ratio of paracellular to transcellular conductance, 66–69 junctional electrical resistance as measure of junctional ion permeability, 63–66 junctional location of high-conductance pathway in leaky epithelia, 71–72 roles of paracellular pathway in leaky epithelia, 69–71 junctional ion selectivity, 72–74 junctional water permeability, 77–82 estimates of juntional water permeability, 79–81 junctional permeation of large hydrophilic solutes, 79 relationship between junctional ion and water permeability, 78–79 solute–solvent coupling and electrokinetic phenomena, 81–82 mechanisms of junctional ion permeation, 74–77 junctional pores, 74–75 relationship of junctional depth and junctional permeability, 77 solute charge and size, 75–77 PF, see Protoplasmic face P face, 23, 41 P-fracture face, 421 PF ridges, 44, 48 Phalloidin, 202, 588 PHD finger, 247 Phorbol myristate acetate, 641 Phosphatase inhibitors, 425, 505 Phosphatidylethanolamine (PE), 306 Phosphatidylserine (PS), 306 Phospholipase C (PLC), 9, 171, 478 Phosphorylation, 113, 425 PI, see Inositolphospholipids Pia, TER measured in, 421 Pigmented epithelium, 659, 662, 664 PI-3 kinase, 409 PKA, see Protein kinase A PKCλ, see Protein kinase C

765 PKCζ, 242 PKC, see Protein kinase C zeta Placental lactogen, 401 Plakoglobin, 133 Plakophilin, 134 Plakophilin-2, 245 Plasma membranes, 521, 608 Plasmin, 354 Platelet activating factor (PAF), 631 –endothelial cell adhesion molecule-1 (PECAM-1), 631, 633 PLC, see Phospholipase C Pleated septate junctions, 50 Podocyte, 108 Polar expression, 576 Polychaetoid, 277 Polyethylene glycol (PEG), 558 Polylysine, 361 Polymyxin B, 356, 363 Polyproline, 241 Porcine kidney epithelial cell line, 373 Porelike structure, of ~155 kDa complex, 529 Porphyromonas gingivalis, 497, 499 Potassium, 405 p55 protein, 234 Precursor particle, 46 Pregnancy, 396, 397, 398, 410 Presumptive junctional membrane, 53 Primary barrier defect, 558 Profilin, 274 Progesterone, 396, 399, 400 Proinflammatory cytokines, 557, 565 Prolactin, 396, 400, 401, 409 Prostaglandin F2α, 402 Protamine, 360, 363 Protamine sulfate, 251 Protease(s), 11, 353 inhibitors, 469, 471 pathophysiological role of, 354 Proteases, tight junctions and, 459–482 degradation of tight junctions, 473–475 effects of proteases, 461 tight junction formation and cell differentiation, 475–478 tight junctions in human colon adenocarcinoma, 461–473 calcium ions, 466–467 effects of trypsin on formation of tight junctions in HT29 cells, 462–463 effects of various proteases on tight junction formation, 468–469 effect of temperature, 464–466 energy requirement, 467

766 primary tumors and derived cell lines, 461–462 protease inhibitors, 469–472 protein synthesis, 468 separation of induction and assembly of tight junctions, 464 Protein(s), 687 4.1, 266, 273, 276 actin-anchoring, 616 adaptor, 151 adherens junction, 634 ATP-dependent, 576 bacterial/permeability-increasing, 359 basolateral, 157 Caenorhabditis elegans kinesin-like, 241 cationic, 359, 362 composition, blood–brain barrier TJs, 419 constituting targeting site, 178 cytolytic, 357 cytoskeletal, 248 Drosophila melanogaster, 241 eosinophil, 362 GST fusion, 246 GTP-binding, 175, 368 guanylate kinase, 168 heterotrimeric G, 369, 370 Id-1, 409 interaction of occludin with, 224 kinase A (PKA), 207, 371, 374, 565 kinase C (PKC), 9, 23, 171, 269, 292, 370, 373, 409, 478, 698 activation of, 508 calcium-dependent, 503 -dependent polymerization, of actin microfilaments, 708 inhibitors, 504 zeta (PKCζ), 242 leucine-rich-repeat, 187 lipid transfer, 322 MAGUK, 590 mononeric G, 369 motor, 152 Na-dependent transport, 576 Na-independent transport, 576 nucleic acid–binding, 236 oligodendrocyte-specific, 98 p55, 234 PDZ, 167 peripheral occluding-junction-associated, 615 phosphorylation of tight junction, 375 polarized delivery of to cell surface, 146 –protein interactions, using FRET, 208 proteolytic degradation of junctional, 634 Rab, 177 Rho GTP-binding, 495

Tight Junctions scaffolding, 272 Sec1/munc-18 related, 177 selective stabilization of membrane, 179 signaling, 175, 236 small G, 369 sorting, 166 synthesis, 468 Tamm Horsfall, 488 targeting pathways, see Epithelial cells, protein targeting pathways and sorting signals in transmembrane, 633, 724 YopE, 270 ZO, 224 Protoplasmic face (PF), 459 PS, see Phosphatidylserine P-selectin, 630, 639 neutrophils and, 639 storage of in secretory granules, 640 Pseudogene, 726 Pseudomonas aeruginosa, 356, 498 aeruginosa exotoxin A, 499 elastase, 355, 499 Pseudo-solvent drag, 81 PTA1, 246 PTH, see Parathyroid hormone Pulmonary capillaries, 635, 636 Pump–leak mechanism, 656, 676 Puncta distribution, 176 Punctate appositions, 41, 49 Pupal metamorphosis, 54 Puromycin, 468 Putricine, 362

R Rab 13, 23, 289 Rab 3B, 10, 23 Rab proteins, 177 Rac 1, 129 Rac GTPase, 270 Raft(s), 150 detergent-insoluble membrane, 307 lipid, 320 pathway, 321 sphingolipid–cholesterol, 498 Ras, 478 Rat Y-box binding protein-a (RYB-a), 246 R-cadherin, 420 Red cell trapping, 545 Regurgitation, of bile constituents, 576 Renal cortical collecting duct, 206 Renal ischemia, 535

Index decreased dextran clearance, 537 loss of cell polarity, 536 molecular changes, 537 TJ disruption, 536 Renal proximal tubule, 206 Resistive shunt current, 738 Reticular septate junctions, 51 Retina, 654, 664 Retinal capillary endothelial cells, 665 Retinal conditioned medium, 431, 432, 433 Retinal pigment epithelium (RPE), 146, 182, 416, 419 culture cell model, 426 culture model of chick, 431 development of, 426 junctions factors regulating, 432 ion selectivity of, 436 ratio of monosaccharide/mannitol permeability of cultured, 434 regulation of, 432 structure of, 426 ZO-1 expression, 436 Retinular junctions, 51 Rho A activity, 127 Rho family, GTPases of, 176 Rho GTPase, 270, 496 Rho GTP-binding proteins, 495 Rotavirus, 508 Rough endoplasmic reticulum, cisternae of, 615 RPE, see Retinal pigment epithelium RU486, 400 RYB-a, see Rat Y-box binding protein-a

S S1, see Subfragment-1 Salmo gairdneri, 451 Saltatory conduction, 736, 740 Scaffolding proteins, 272 Scalae vestibuli, 730 Scanning electron microscopy, 639, 643 Schlemm’s canal endothelial cells, 668 Schmidt–Lanterman incisures, 734 Scorpions, 40 Scrib, 187 Seawater -adapted teleosts, 452 environment, 446 teleost chloride cell, 447, 450 Sec6/8, 10, 23, 178, 183 Sec1/Munc-18 related proteins, 177 Secretory pathway, 319 Selective stabilization, 167, 179

767 Selective targeting, 167 Self-filling blind loops, 562 Seminiferous epithelium, 600 Seminiferous tubule, lumen in, 607 Septate junctions, 39, 40, 44, 53 arthropod, 46 Drosophila, 736, 739 pleated, 50 reticular, 51 smooth, 49 structural features of, 49 Sequences, genetic analysis looking for homologous, 55 Serine 19, phosphorylation of, 644 Serine protein kinase, 236 Serosal domain, 350 Sertoli cell(s), 739 access to older germ cells through, 606 cell junctions between membranes of, 600 as filter, 606 freeze-fracture observations of, 610 function of occluding junctions between basal cells and, 739 junctional barrier, synchronization of, 609 as modulator, 606 molecular composition of, 614–615 occluding junctions, proteinic constituents of, 614 occluding zonules, mechanism for assembly and disassembly of, 616 sealing of paracellular route between, 605 seminiferous tubules and, 600 tight junction, 731 assembly of, 616–617 development of, 605 as immunological barrier, 618 polarity, 607 Serum albumin, 398 SET, 247 SH3 homologies, 419 phylogenetic comparisons, 419 Shigella flexneri, 509 Short bowel syndrome, 563, 564 Short-circuit currents, 4, 64 Shunt pathway, 64, 68 Side-to-side interactions, 405 Sigmoid colon, 556 Signaling proteins, 175, 236 Signal transduction, 135, 409 centers, 168 events, 498 Single exon claudins, 725, 727 Size selectivity, 578, 584, 587 SL, see Sphingolipids

768 S1 labeling, 55 SM, see Sphingomyelin Small complex, ~90 kDa, 522 Small G proteins, 369 Small GTPases, 372 Smooth septate junctions, 49 SNAREs, 177, 321 Sodium, 405, 406 flux, 204 glucose transporter, 270 permeability to, 397 Solute permeability, of tight junction, 203 Solvent drag, 71, 81, 334 Sorting motifs, tyrosine-based, 181 signals, AP1, 151 Soybean trypsin inhibitor (STI), 464 Space of Disse, 576, 577 Spectrin, 179, 266, 273 α-Spectrin, 616 Spermatocyte precursor cells, 732 Spermatogenesis, 732, 740 Spermatozoa, 315 Spermidine, 362 Spermine, 362 Sphingolipid–cholesterol rafts, 498 Sphingolipids (SL), 306 Sphingomyelin (SM), 306 Spiders, 40 Spinal cord, 729 Sporadic diarrhea, 519 Src, 167, 170, 236 S-SCAM, 241 S1 subfragment, 46 Steroid-induced glaucoma, 669, 673 Sterols, 608 STI, see Soybean trypsin inhibitor Streptococcal pneumonia, 638, 643 Streptococcus pneumoniae, 638 Stress fibers, 175 Stria vascularis, 730, 737 Structure and function, relationship between tight junction, 89–119 changes in phosphorylation state of tight junction components, 113 electric circuit analysis as tool for predicting epithelial resistance, 109–113 cells differing in TER, 110 epithelia derived from different animal species, 110 establishment of epithelial monolayers, 110 mixed monolayers formed by cells derived from different animal species, 111–113

Tight Junctions natural epithelia formed by different types of cells, 109–110 experimental modification of tight junction permeability, 113 gate function of tight junctions, 100–107 classification of epithelia as tight or leaky, 102–103 electrical resistance of tight junction, 100–102 morphological aspects affecting TER, 103–104 passage of tracers through paracellular pathway, 100 relationship between specific resistance of tight junction and number of strands, 104–107 molecular fence and paracellular gate functions of tight junctions, 99–100 molecular nature of tight junction strands, 93–99 claudins, 98–99 occludin, 95–98 presence of pores or channels within tight junction strands, 107–109 nephrin, 108 paracellin, 108–109 ultrastructural features of tight junctions, 90–93 freeze fracture, 91–93 thin section, 90–91 Subepithelial resistance, 555, 556 Subfragment-1 (S1), 243, 249 [14C]-sucrose, 399, 407 Symplekin, 21, 23, 244

T TAL, see Thick ascending limb of Henle’s loop Tamm Horsfall Protein (THP), 488 Tamou, 236 Tannic acid staining, 46 Targeting patches, 177 site, proteins constituting, 178 Taurolithocholate, 587 Taxol, 475 T-cell infiltration, 733 T84 cells, 240, 359, 462, 566, 568 Teleost(s) chloride cell freeze fracture, 451 seawater, 447, 450

Index fresh water–adapted animals, 452 seawater-adapted, 452 Teleost chloride cell tight junctions, 445–458 aquatic environment, 445–446 chloride cell in fresh water–adapted teleosts, 451–452 chloride cell in nonteleost species, 455 chloride cell in seawater-adapted teleosts, 447–449 insights from isolated epithelia containing chloride cells, 452–453 dynamic nature of chloride cell–chloride cell tight junction in seawateradapted teleosts, 452 studies on isolated teleost epithelia from fresh water–adapted animals, 452–453 ion absorption mechanism in teleost gill, 450–451 model for sodium chloride secretion by seawater-adapted teleost chloride cell, 449–450 new observations from gill tissue in culture, 453–454 cultures with chloride cells, 453 cultures without chloride cells, 453–454 patterns of osmoregulation in seawater and in fresh water, 446–447 freshwater environment, 446–447 seawater environment, 446 TEM, see Transmission electron microscopy TER, see Transepithelial electrical resistance Terminal bar, 1, 3, 20, 122 Testis, epididymis, and vas deferens, tight junctions in, 599–628 factors susceptible to influencing permeability status of Sertoli cell occluding zonules, 610–614 differential distribution of junctional particles within plane of cellular membrane, 613–614 number of junctional fibrils, 610–613 immunological barrier to anti-sperm antibodies, 618–619 mechanisms for assembly and disassembly of Sertoli cell occluding zonules, 616–618 theory of presence of intermediate cellular compartment within seminiferous epithelium, 617 theory of repetitive removal of membrane segments from Sertoli cell junctional complex, 617–618 zipper theory, 616–617

769 occluding junctions in epididymis and vas deferens, 620–622 proteinic constituents of Sertoli cell occluding junctions, 614–616 integral membrane proteins, 614–615 peripheral occluding-junction-associated proteins and subsurface or cortical actin in Sertoli cells, 615–616 tight junctions in testis, 600–610 physiological consequences of establishment of occluding junctions in seminiferous epithelium, 605–609 synchronization of Sertoli cell junctional barrier function, 609–610 topographical distribution of intercellular junctions in seminiferous epithelium, 600–605 Tetanus toxoid (TT), 716 TGF-β, 410 TGN, see trans-Golgi network Therapeutic manipulation blood–brain barrier, 705 dermal barrier, 707 mucosal barrier, 706 oral delivery, 707 Thick ascending limb of Henle’s loop (TAL), 455 Thin sections, appearance of TJ in, 90 THP, see Tamm Horsfall Protein Threonine 18, phosphorylation of, 644 Thrombin, 640 Tight epithelia, 68, 102 Tight junction (TJ), 1, 3, 19, 39, 90, 166, 213, 232, 286, 310, 333, 350, 415, 446, 459, 484, 534, 629, 654, 698, 723 ability of to slide within membrane, 646 apicolateral, 663 assembly, 170, 464 mouse embryo, 288 Xenopus, 293 -associated molecules, phosphorylation of, 676 bacterial toxins, 269 barrier disruption of by pathogen-stimulated signaling events, 503 function, 168, 203, 576 regulation of, 494 bile secretion, 576–578 blind loop syndrome, 562 celiac sprue, 559 central nervous system myelin, 734 cholestasis, 580–589 ciliary body, 659

770 coexistence of with gap junction, 45 continuous, 669 cornea, 655 Crohn’s disease, 558 cytochalasins, 267 cytoskeletal-mediated disruption of by microbial pathogens, 495 degradation of, 473, 474 developmental studies in moth, 54 dysfunction, 539 in blood–brain barrier, 699 in intestinal epithelium, 701 electrical resistance of, 63, 100 endothelial-like, 632 epithelial, 525 extrahepatic obstruction, 580–586 fence function of, 168, 585 fibrils disintegration of, 525 dramatic fragmentation of, 525 formation, 180, 464, 475 fracturing characteristics of, 53 functions, see Intracellular signaling, in classical and new tight junction functions GTP-binding proteins, 369 heterotrimeric G proteins, 369 HIV enteropathy, 564 HRP-positive, 582 infectious diarrhea, 564 ischemia, effects of, 535 isoelectric point of, 7 kisses, 267 lipid polarity, maintaining, 312 liver disease, 580 loose network of particle rows in tunicate, 53 mechanisms for opening/remodeling, 644 microbial toxins disrupting, 498, 499 microfilaments of arthropod, 55 modulation, therapeutic use of, 705 molecular fence function of, 99 morphology, crypt–villus axis, 562 morphometry, ulcerative colitis, 556 noncontinuous, 662 paracellular gate function of, 99 paracellular route limited by, 7 particles, 41 permeability, 340, 400, 407, see also Mammary gland, regulation of tight junction permeability in linkage between perijunctional actomyosin ring and, 340 milk secretion and, 411 oxytocin and, 408

Tight Junctions plaque proteins, 253 pores, 434–436 protein kinase A, 374 protein kinase C, 373 proteins phosphorylation of, 375 transmembrane, 724 proteolysis, 647 regulation, 369, 373, 374, 408, 507 relationship between specific resistance of and number of strands, 104 resistance, 579, 580 ridges, 48 sealing, 410 Sertoli cell, 605, 607, 618, 731, 732 short bowel syndrome, 563 size selectivity of, 587 solute permeability of, 203 strand, 107, 403 stria vascularis, 730 structure, 493, 525 transcription factors, 383 transport of drugs by processes between epithelia cells and, 685 ulcerative colitis, 554 water flow across, 205 zonulin-induced opening of, 711 Tight junction, evolution of ideas on, 1–18 biosynthesis and assembly of tight junction, 11 definition of tight junction, 7–9 new roles in signaling, 11–12 not always so tight, 4–7 role of tight junctions in human disease, 12–13 terminal bar, 1–4 tight junctions and apical/basolateral polarity, 9–11 tight junctions in special situations, 12 Tissue culture, 408 TJ, see Tight junction TMC, see N-Trimethyl chitosan chloride TNF, see Tumor necrosis factor TNFα, see Tumor necrosis factor-alpha Toad urinary bladder epithelium, 64 Toxin-induced cholestasis, 587 Trabecular cell loss, 676 Trabecular meshwork, 666, 668, 671 Tracer molecules, 41 Tracheal epithelial cells, 635 Transbilayer distribution, 317 Transcellular pathway, 19 Transcellular resistance, 68 Transcription factors, 170, 383

Index Transendothelial fluid flow, 669, 676 Transendothelial migration, 629 Transepithelial electrical resistance (TER), 28, 90, 95, 102, 220, 268, 368, 407, 493, 502, 534, 578, 665 claudin and, 222 increase of by HUVEC exposure to culture medium, 635 measurement of, 656 morphological aspects affecting, 103 neutrophil transmigration and, 634 occludin and, 221 Transepithelial resistance, 68, 252, 408 Transmembrane proteins, see Occludin and claudins Transmembrane tight junction proteins, 724 Transmission electron microscopy (TEM), 90, 643 Transport number effect, 66 Transport vesicles, TGN-derived, 177 Tricellular corners, 656 N-Trimethyl chitosan chloride (TMC), 708 trithorax transcription factor, 247 Trophectoderm, 286 differentiation, 287, 288, 292 impermeable, 291 model, 292, 293 Trypsin, 353, 462 t-SNAREs, 156 TT, see Tetanus toxoid Tumor necrosis factor (TNF), 406 Tumor necrosis factor-alpha (TNF), 269, 565, 566, 570, 631, 642, 700 Tunicates, 40, 53 Two-membrane hypothesis, 66 Type II pneumocytes, 643 Type III secretory system, 504 Tyrosine -based sorting motifs, 181 -dependent motifs, 167 kinase(s), 249, 478 Eph-related receptor, 242 nonreceptor, 170 phosphatase, 249 phosphorylation, 168, 278

U UC, see Ulcerative colitis Ulcerative colitis (UC), 554, 704 aberrant strands in, 555 proinflammatory cytokines, 557 strand discontinuities in, 555

771 tight junction morphometry in, 556 ultrastructural changes, 555 Ultrastructure and immunolabeling of tight junction, 19–37 components of junctional complex, 20–21 morphology of tight junctions in freezefracture replicas and immunolocalization of tight junction proteins in tight junction strands, 23–26 structure–function correlations, 32 tight junctions and lipid environment, 26–30 ultrastructure of developing epithelial tight junctions, 30–31 ultrastructure of right junction and immunolocalization of tight junction proteins, 21–23 Umbilical cord–derived cell line, 423 unc-104, 241 Unstirred water layer, 350 Urinary bladder, 360 Urogenital tract, 729 Urokinase, 354 Ussing chamber, 67

V VACs, 187 VAP-33, 21, 23, 187 Vascular cell adhesion molecule-1 (VCAM-1), 631 Vascular endothelial cell barrier, 665 Vascular endothelial growth factor (VEGF), 379, 423, 642, 676 Vas deferens, see Testis, epididymis, and vas deferens, tight junctions in VASP, 177 VCAM-1, see Vascular cell adhesion molecule-1 VE-cadherin, 420, 633 Vectorial transport, 576 VEGF, see Vascular endothelial growth factor Vero cells, 522 Vesicular traffic, to plasma membrane domains, 318 Vestibular semicircular canals, 730 Vestibulocochlear apparatus, 740 Vibrio cholerae, 564, 708 hemagglutinin protease, 501 zonula occludens toxin, 270 Vibrissae, hair follicles of, 729 Vinculin, 129, 177, 245 Vitamin A, 461 v-SNAREs, 156

772

W Water flow, across tight junction, 205 permeability, 77, 79 transport osmotic, 71 paracellular, 78 Weibel–Palade bodies, 630, 640, 641 Western blotting, 408 Wingless/Wnt signaling pathway, 170 Wnt pathway, 383 Wnt signal, 123 Wortmannin, 409 WW domains, 241

X Xenobiotics, 355 Xenopus, 171, 223, 460, 725 blastula formation, 286 embryo, 293 laevis, 113

Y Yersinia enterocolitica, 510 pseudotuberculosis, 270, 509 Y junction, 656, 658 YopE protein, 270

Z ZAK, see ZO-1-associated protein kinase Zebrafish, 725 Zinc metalloprotease, 499 Zipper theory, 616 ZO, see Zonula occludens ZO-1α –, see Zonula occludens-1 ZO-1α +, 234, 615 ZO-1+, 234, 615 ZO-2, 11, 168, 180, 236, 272, 273, 375, 498, 632

Tight Junctions downregulated, 555 homologies, 419 phylogenetic comparisons, 419 steady-state level of, 430 ZO-3, 11, 168, 180, 272, 632 homologies, 419 phylogenetic comparisons, 419 ZONAB, see ZO-1 associated nucleic acidbinding protein Zonula adherens, 293 Zonula occludens (ZO), 3, 20, 698 Zonula occludens-1 (ZO-1), 11, 129, 168, 180, 185, 272, 289, 408, 421, 424, 427, 461, 497, 498, 507, 590, 632, 655, 665, 708 antibodies binding ZO-1LP and, 430 associated nucleic acid-binding protein (ZONAB), 21, 23, 236, 246, 383 -associated protein kinase (ZAK), 23, 236, 376 BDL and, 582 clustering of transmembrane junctional proteins by, 237 downregulated, 555 expression, 436, 671, 672, 674 homologies, 419 paracellular permeability increased by redistributing, 510 pattern of in BDL-induced cholestasis, 590 phosphorylation, 379 phylogenetic comparisons, 419 proline-rich region of, 236 relocation, 295 scaffolding function of, 237 staining, 404 subcellular distribution of, 586 Zonula occludens toxin (ZOT), 270, 502, 502, 503, 561, 564, 708 Zonule(s) assembly/disassembly of, 609 discontinuous, 609 pattern of junctional fibrils within, 610 Zonulin, 561, 709 Zonulin system, 708 pathology of, 711 therapeutical use of, 714 ZOT, see Zonula occludens toxin Zyxin, 177