International Review of Cytology, Vol. 264 [1st ed.] 0123742633, 9780123742636, 9780080555119

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V O LU M E

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S I X T Y

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INTERNATIONAL REVIEW OF

CYTOLOGY A Survey of Cell Biology

INTERNATIONAL REVIEW OF CYTOLOGY Series Editors

GEOFFREY H. BOURNE JAMES F. DANIELLI KWANG W. JEON MARTIN FRIEDLANDER JONATHAN JARVIK

1949–1988 1949–1984 1967– 1984–1992 1993–1995

Editorial Advisory Board

ISAIAH ARKIN EVE IDA BARAK PETER L. BEECH HOWARD A. BERN ROBERT A. BLOODGOOD DEAN BOK HIROO FUKUDA RAY H. GAVIN SIAMON GORDON MAY GRIFFITH WILLIAM R. JEFFERY KEITH LATHAM

WALLACE F. MARSHALL BRUCE D. MCKEE MICHAEL MELKONIAN KEITH E. MOSTOV ANDREAS OKSCHE THORU PEDERSON MANFRED SCHLIWA TERUO SHIMMEN ROBERT A. SMITH WILDRED D. STEIN NIKOLAI TOMILIN

V O LU M E

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S I X T Y

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INTERNATIONAL REVIEW OF

CYTOLOGY A Survey of Cell Biology EDITED BY

KWANG W. JEON Department of Biochemistry University of Tennessee Knoxville, Tennessee

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Front Cover Photograph: Courtesy of T. Nakano and K. Goto, Department of Anatomy and Cell Biology, Yamagata University School of Medicine, Japan

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This book is printed on acid-free paper. Copyright # 2007, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2007 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0074-7696/2007 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail: permissions@elsevier. com. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Support & Contact’’ then ‘‘Copyright and Permission’’ and then ‘‘Obtaining Permissions.’’ For information on all Elsevier Academic Press publications visit our Web site at www.books.elsevier.com ISBN: 978-0-12-374263-6

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CONTENTS

Contributors

1. Function and Evolution of the Vacuolar Compartment in Green Algae and Land Plants (Viridiplantae)

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Burkhard Becker 1. Introduction 2. Structure and Function of Vacuoles in Embryophytes 3. Structure and Function of Vacuoles in Green Algae 4. Evolution of Vacuolar Compartments in Plants 5. Concluding Remarks Acknowledgments References

2. Cell Biology and Pathophysiology of the Diacylglycerol Kinase Family: Morphological Aspects in Tissues and Organs

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Kaoru Goto, Yasukazu Hozumi, Tomoyuki Nakano, Sachiko S. Saino, and Hisatake Kondo 1. 2. 3. 4.

Introduction Molecular Heterogeneity Gene Expression in the Brain Morphological Analysis of the Subcellular Localization in Tissues and Organs 5. Pathophysiological Implications in Animal Studies 6. Concluding Remarks Acknowledgments References

3. Structure and Function of Desmosomes

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¨fer, Reinhard Windoffer, Sergey Troyanovsky, and Bastian Holtho Rudolf E. Leube 1. Introduction 2. Morphology 3. Molecular Architecture

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4. Biogenesis 5. Dynamics 6. Imbalance of Desmosomal Protein Synthesis in Transgenic Mice 7. Interplay Between Desmosomes and Other Cell Components 8. Desmosomes and Disease 9. Concluding Remarks Acknowledgments References

4. Subepithelial Fibroblasts in Intestinal Villi: Roles in Intercellular Communication

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Sonoko Furuya and Kishio Furuya 1. Introduction 2. Morphological Features of Subepithelial Fibroblasts 3. Receptors in Subepithelial Fibroblasts 4. Gap Junction Communication 5. Mechanosensitive Networks via ATP Receptors 6. Roles of Subepithelial Fibroblasts in the Villi 7. Concluding Remarks References

5. Syndrome of Aluminum Toxicity and Diversity of Aluminum Resistance in Higher Plants

166 168 185 191 195 202 211 211

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Jian Feng Ma 1. Introduction 2. Syndrome of Aluminum Toxicity 3. Aluminum Resistance 4. Beneficial Effect of Aluminum on Plant Growth 5. Concluding Remarks Acknowledgments References Index

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CONTRIBUTORS

Burkhard Becker Botanical Institute, University of Cologne, 50931 Ko¨ln, Germany Sonoko Furuya Section of Brain Structure, Center for Brain Experiment, National Institute for Physiological Sciences, Okazaki 444-8585, Japan Kishio Furuya Cell Mechanosensing Project, ICORP/SORST, Japan Science and Technology Agency, Nagoya 466-8550, Japan Kaoru Goto Department of Anatomy and Cell Biology, Yamagata University School of Medicine, Yamagata 990-9585, Japan ¨fer Bastian Holtho Department of Anatomy and Cell Biology, Johannes Gutenberg University, 55128 Mainz, Germany Yasukazu Hozumi Department of Anatomy and Cell Biology, Yamagata University School of Medicine, Yamagata 990-9585, Japan Hisatake Kondo Division of Histology, Department of Cell Biology, Tohoku University Graduate School of Medical Science, Sendai 980-8575, Japan Rudolf E. Leube Department of Anatomy and Cell Biology, Johannes Gutenberg University, 55128 Mainz, Germany Jian Feng Ma Research Institute for Bioresources, Okayama University , Kurashiki 710–0046, Japan Tomoyuki Nakano Department of Anatomy and Cell Biology, Yamagata University School of Medicine, Yamagata 990-9585, Japan

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Sachiko S. Saino Department of Anatomy and Cell Biology, Yamagata University School of Medicine, Yamagata 990-9585, Japan Sergey Troyanovsky Department of Internal Medicine (Dermatology), Washington University Medical School, St. Louis, Missouri 63110 Reinhard Windoffer Department of Anatomy and Cell Biology, Johannes Gutenberg University, 55128 Mainz, Germany

C H A P T E R

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Function and Evolution of the Vacuolar Compartment in Green Algae and Land Plants (Viridiplantae) Burkhard Becker

Contents 2 3 3 7 8 10 10 12 13 17 18 18

1. Introduction 2. Structure and Function of Vacuoles in Embryophytes 2.1. Types and functions of vacuoles 2.2. Structure and development of vacuoles 2.3. Protein targeting to vacuoles 3. Structure and Function of Vacuoles in Green Algae 3.1. Types and functions of vacuoles 3.2. Development of vacuoles 4. Evolution of Vacuolar Compartments in Plants 5. Concluding Remarks Acknowledgments References

Abstract Plant vacuoles perform several different functions and are essential for the plant cell. The large central vacuoles of mature plant cells provide structural support, and they serve other functions, such as protein degradation and turnover, waste disposal, storage of metabolites, and cell growth. A unique feature of the plant vacuolar system is the presence of different types of vacuoles within the same cell. The current knowledge about the vacuolar compartments in plants and green algae is summarized and a hypothesis is presented to explain the origin of multiple types of vacuoles in plants. Key Words: Plant vacuole, Green algae, Protein targeting, Turgor pressure, Autophagy. ß 2007 Elsevier Inc.

Botanical Institute, University of Cologne, 50931 Ko¨ln, Germany International Review of Cytology, Volume 264 ISSN 0074-7696, DOI: 10.1016/S0074-7696(07)64001-7

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2007 Elsevier Inc. All rights reserved.

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1. Introduction Plant vacuoles are large single-membrane–bounded compartments within the cytoplasm of a cell that function in several different ways (Marty, 1999). The vacuole is essential for the viability of the plant cell (Rojo et al., 2001). In mature plant cells, vacuoles tend to be very large (80% or more of the cell volume), occupy a central position, and are extremely important in providing structural support, as well as serving functions such as storage, waste disposal, protection, and growth (Marty, 1999). Vacuoles in animal cells, however, tend to be much smaller, and are more commonly used to store materials temporarily or to transport substances. The central vacuole in a plant cell is enclosed by a membrane termed the tonoplast, which is part of the endomembrane system of the cell (i.e., the vacuole is linked to other compartments of the endomembrane system by vesicular transport) (Surpin and Raikhel, 2004). The large central vacuole develops as the cell matures by fusion of smaller vacuoles, which are derived from the endoplasmic reticulum and/or Golgi apparatus. Because the central vacuole is highly selective in transporting materials through its membrane, the chemical composition of the vacuolar solution (termed the cell sap) differs markedly from that of the surrounding cytoplasm and varies among different cell types (Marty, 1999). For example, some vacuoles contain pigments that give certain flowers their characteristic colors. The central vacuole also contains plant wastes that taste bitter to insects and animals, while developing seed cells use the vacuole as a repository for protein storage. The central vacuole stores salts, minerals, and nutrients; helps in plant growth; and plays an important structural role for the plant. Under optimal conditions, vacuoles are filled with water to the point that they exert a significant pressure against the cell wall (turgor pressure). This turgor pressure is cell specific in regulation (Findlay, 2001) (e.g., in the guard cells of leaf stomata, changes in turgor are used to open and close the stomata). In addition, the turgor pressure helps to maintain the structural integrity of the plant, along with the support from the cell wall and enables the plant cell to grow much larger without having to synthesize new cytoplasm. In most cases, the plant cytoplasm is confined to a thin layer positioned between the plasma membrane and the tonoplast, yielding a large ratio of membrane surface to cytoplasm (Weibe, 1978). Plant vacuoles are also important for their role in molecular degradation and storage. Sometimes these functions are carried out by different vacuoles in the same cell, one serving as a compartment for breaking down materials (similar to the lysosomes found in animal cells), and another storing nutrients, waste products, or other substances (Marty, 1999). Here, I will review advances in our understanding of the plant vacuole, concentrating on work published in 2005 and 2006. I will then briefly

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discuss the structure and function of vacuoles in green algae. The last section will focus on the origin of the vacuolar system in plants and algae.

2. Structure and Function of Vacuoles in Embryophytes 2.1. Types and functions of vacuoles It is a common belief that most plant cells contain only the single large central vacuole described above. Therefore, it was a big surprise when Paris et al. (1996) showed that two different types of vacuoles were present within the same cell performing different functions (Fig. 1.1A). Since 1996 several studies have shown that plant cells contain multiple types of vacuoles with distinct functions (Di Sansebastiano et al., 2001; Epimashko et al., 2004; Jauh et al., 1998, 1999; Park et al., 2004). Up to three separate distinct types of vacuoles have been reported within a single plant cell ( Jauh et al., 1999). Generally, vacuoles are divided into two categories. Lytic vacuoles (see LVs, Fig. 1.1A) are acidic compartments and are rich in hydrolases. LVs are considered as equivalent to the animal lysosome and are recognized by the presence of g-TIP (the g-isoform of tonoplast intrinsic proteins, a member of the large glyceroaquaporin protein family present in plants). Protein storage vacuoles (see PSVs, Fig. 1.1A) are most often found in storage organs and are characterized by the presence of a-TIP (Jauh et al., 1999), but have recently also been reported in mesophyll cells (Park et al., 2004). The presence of separate vacuoles of distinct function in seed plants is in marked contrast to animal and fungal cells that contain only lysosomes or a single vacuole (see also the discussion of this question by Robinson et al., 2005). In the past 2 years, progress has been made in understanding most aspects of vacuolar function. The following is a brief summary of major recent findings regarding vacuolar function. 2.1.1. Turgor The structural importance of the plant vacuole is related to its ability to control turgor pressure. Turgor pressure dictates the rigidity of the cell and is associated with the difference between the osmotic pressure inside and outside of the cell. The role of aquaporins in water relations of the vacuole is a hot topic in plant research. Since the progress in this field has been reviewed several times in the past years, the reader is referred to the reviews by Hachez et al. (2006) and Luu and Maurel (2005) for further information. 2.1.2. Storage Plant vacuoles store a large variety of chemical compounds. The protein targeting of storage protein to specialized vacuoles (protein storage vacuoles) and the processing inside the vacuoles are discussed in Section 2.2.

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Figure 1.1 (A) The endomembrane system of a plant cell.The vesicular transport pathways to the vacuole are shown. Uncharacterized pathways expected to be present are in gray. Pathways possibly involving vacuolar-sorting receptors (VSRs) are colored according to the suggested functions (Masclaux et al., 2005) for the three phylogenetic groups of ArabidopsisVSRs: green, lytic vacuole; red, protein storage vacuole; and blue, endocytotic pathway. (B) Phylogenetic relationships of VSRs from Arabidopsis and a few other selected angiosperms.The color coding is the same as in (A). (Modified from Masclaux et al. [2005].)

In addition, the large central vacuole is known to store several other small molecules. Stored metabolites and chemicals can either serve as a cellular pool, on which the cell can rely during starvation, or as a detoxification step to prevent interference of chemicals with cellular function. It has long been known that nitrate, the principal nitrogen source of most plants, can accumulate in large quantities in certain plants (e.g., of the

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families Chenopodiaceae, Poaceae, Brassicaceae, and Asteraceae) (Martinoia et al., 1981). Most of the nitrate is stored in the vacuole. However, the mechanism of uptake into the vacuole has remained elusive for years. More recently, De Angeli et al. (2006) provided evidence that nitrate is transported into the vacuole using a proton antiport mechanism and localized a member of the voltage-gated chloride channel protein family (AtCLCa) to the tonoplast, confirming its involvement in nitrate transport as earlier suggested (Geelen et al., 2000). Several other vacuolar carrier systems have already been characterized at the molecular level, and recent work has added to this list, including Zn (Elbaz et al., 2006), Al (Kochian et al., 2005), Na (Yamaguchi et al., 2005), Mn (Pittman, 2005), Ca (Pittman et al., 2005), Fe uptake systems (Kim et al., 2006), and a novel monosaccharide transporter (Wormit et al., 2006). Detoxification of xenobiotics generally occurs in four steps: activation of a xenobiotic, formation of a GSH-S-conjugate (GSX), sequestration (in plants: into the vacuole), and degradation of GSX (Sandermann, 1994). In mammals GSX is degraded by g-glutamyltransferases (GGTs) and L-cysteinglycinyldipeptidase (DPase) reactions (Meister and Anderson, 1983). Whereas the first three steps are well characterized in plants (Foyer et al., 2001; Rea, 1999), degradation of GSX by GGT and DPase has not been demonstrated in plants so far. Nakano et al. (2006) have now demonstrated the presence of GGT and DPase activity in isolated vacuoles from radish cotyledons suggesting that the complete pathway for detoxification is conserved between animals and plants. The proteome of the tonoplast from Arabidopsis thaliana has been characterized (Carter et al., 2004; Sazuka et al., 2004; Shimaoka et al., 2004; Szponarski et al., 2004). In addition to proteins known to be present (VATPase, Hþ transporting PPase, TIPs), several proteins with unknown or unexpected functions were detected. Additional work will be required to analyze the relevance of their presence in the isolated tonoplast fractions. However, the list of tonoplast proteins is far from complete, as revealed by a new proteomic study. Endler et al. (2006) identified 40 additional proteins in the tonoplast fraction isolated from barley leaves that were not detected in the four proteomic studies of the tonoplast of A. thaliana. Among the new proteins identified by Endler et al. (2006) is the first tonoplast sucrose transporter (HvSUT2). Arabidopsis mesophyll cells possess a homologue of HvSUT2 (AtSUT4) on their tonoplast membrane. Sucrose is mainly stored in the vacuole of plant cells. In leaves, sucrose is transported to the vacuole during the light period. At night, sucrose is then transported to the phloem for transport to other tissues (Kaiser and Heber, 1984). Salt stress is a major problem in agriculture. About 20% of the world’s cultivated lands are affected by salinity (Chrispeels and Sadava, 2003). To cope with high salinity, halophytes have evolved mechanisms to protect their cells from the detrimental effects of salts on plants. Among other

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adaptations (increased cytosolic concentrations of compatible solutes), halophytes sequester sodium to vacuoles to increase vacuolar osmolarity and keep sodium away from the sites of metabolism. Sodium is transported into vacuoles using an Naþ/Hþ antiporter. The Naþ/Hþ antiport is driven by the electrochemical proton gradient generated by V-ATPase and V-PPase. From these two the latter enzyme activity has been shown to be increased during salt stress in Salicornia bigelovii (Parks et al., 2002). More recently, Guo et al. (2006) cloned the V-PPase from the halophyte Suaeda salsa. When the V-PPase from S. salsa was expressed in Arabidopsis, increased salt and drought tolerance were observed (Guo et al., 2006). 2.1.3. Autophagy and vacuole-mediated cell death Autophagy is the nonselective uptake of large portions of the cytosol and/or organelles by encapsulating cellular material with membranes, transporting the material to the vacuole, and degradation in the vacuole. Autophagy is an important mechanism for protein turnover, as well as a universal reaction to cell starvation. The process is best characterized in yeast (see Klionsky et al., 2003 for a recent list of autophagy-related genes). Autophagic transport to the vacuole occurs by two morphological distinct but mechanistically overlapping pathways: microautophagy and macroautophagy (Reggiori and Klionsky, 2002). In microautophagy the material to be degraded is directly taken up by the lysosome/vacuole. In macroautophagy the material to be degraded is first engulfed by membranes separate from lysosomes/vacuoles and then transported to the vacuole. Two different autophagic pathways have been demonstrated to occur in plant cells (Toyooka et al., 2001). Degradation of starch granules in mung bean cotyledons is clearly similar to microautophagy (Toyooka et al., 2001), whereas the uptake of cytosol and mitochondria involved a mechanistically different second pathway. The presence of a macroautophagic pathway in plants has recently been demonstrated by Toyooka et al. (2006). In addition to a role in mobilization of nutrients during germination, autophagy constitutively takes place in root tips (Inoue et al., 2006) and is also involved in other processes. Thompson et al. (2005) reported that autophagy plays an essential role in nutrient recycling in Arabidopsis. Plants lacking the autophagic pathway display early senescence and are hypersensitive to carbon and nitrogen starvation (Thompson et al., 2005). Stroma proteins of the chloroplast have been shown to be transported to the vacuole (Chiba et al., 2003); however, whether this involves autophagy or another mechanism to be discovered is an open question . Plants exhibit programmed cell death (PCD) during plant development (e.g., floral organs, Rogers, 2006) or as a response to pathogens. Although some mechanisms underlying PCD are thought to be conserved between plants and animals, a key feature of PCD in animals is the dependence on caspase protease activity. Up to now, the presence of caspase activity in plants has remained elusive. Hatsugai et al. (2004) reported that a vacuolar processing

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protease (VPE), which is structurally unrelated to caspase, has a caspase-1 activity (see Hatsugai et al., 2006 for a detailed comparison of VPE with caspases). Furthermore, this enzyme is essential for virus-induced responses that involve PCD (Hatsugai et al., 2004), indicating that PCD in plants is mediated by the vacuole. Recently it has been hypothesized that an autophagic pathway is required for executing PCD (Seay and Dinesh-Kumar, 2005).

2.2. Structure and development of vacuoles Fluorescent probes (e.g., green fluorescent protein [GFP]) have been used for more than a decade to address biological questions. However, the focus of most studies using GFP-tagged vacuolar proteins (e.g., Czempinski et al., 2002; Kataoka et al., 2004; Kutsuna and Hasezawa, 2002; Mitsuhashi et al., 2000; Reisen et al., 2003; Ueoka-Nakanishi et al., 2000; Yano et al., 2004) was to localize the protein and not to study the dynamics or development of the vacuole. Only a few studies have addressed the structure and dynamics of the vacuolar system. The structure of the vacuolar system of germinating pollen was investigated by Hicks et al. (2004) using a g-TIP-GFP construct. In germinating pollen the vacuolar system consists of elongated (tubular) vacuoles with highly mobile cytoplasmic invaginations. Hicks et al. (2004) also investigated the effect of the vacuoless1 mutation (vcl1) on the structure of the vacuole in the male gametophyte. Vcl1 was shown to be essential for biogenesis of the vacuole in the embryo. Inactivation of both copies of VCL1 caused the accumulation of small vesicles and autolysosomes in the cells and led to lethality at the torpedo stage of embryogenesis (Rojo et al., 2001). Surprisingly, vcl1 did not affect the vacuolar structure in the male gametophyte, although it affected the fertility of the male gametophyte (Hicks et al., 2004). The structure of the vacuolar system is also affected by environmental signals. Irani and Grotewold (2005) demonstrated that light affected the morphology of the vacuolar system in BMS (Black Mexican Sweet, a maize cell line) cells. In maize anthocyans are synthesized after induction by light. Using a genetically engineered cell line expressing the biosynthetic enzymes constitutively (using the CaMV 35S promoter) and leading to accumulation of anthocyans in the dark, changes in cell pigmentation in BMS cells upon transfer to light were observed (Irani and Grotewold, 2005). Subsequently, it could be demonstrated that the changes in cell pigmentation were not due to different cellular concentrations of anthocyans. Instead, several small anthocyan-accumulating vacuoles fused to a large central vacuole upon transferring BMS cells to light (Irani and Grotewold, 2005), indicating that light is regulating the structure of the vacuolar system in this cell line. The response of the vacuole to osmotic changes is well known (plasmolysis and deplasmolysis). While the tonoplast protein complexes have been well

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studied during this process, the tonoplast itself is less well described. Reisen et al. (2005) have determined the three-dimensional structure of the vacuolar system upon (acclimation to) osmotic stress. During plasmolysis the tonoplast of the central vacuole folds, but no vacuolation or vesiculation of the vacuole was observed. When the cells were allowed to acclimate to the osmotic stress, the large central vacuole was converted into a complex vacuolar network with an increased tonoplast surface area upon osmotic stress (Reisen et al., 2005). Mimicking desiccation, polyethylene glycol (PEG) treatment of cells led to spherical structures composed of tonoplast material inside the vacuoles. Again no vacuolation or vesiculation of the tonoplast was observed as reported earlier by Chang et al. (1996) and others. Actin filaments have been shown to be important for maintaining the structure of the large vacuoles in tobacco BY-2 cells (Higaki et al., 2006). Disruption of the actin cytoskeleton caused the formation of small spherical vacuoles (Higaki et al., 2006). Changes in the structure of the vacuolar system are also important for the stomata opening in plants. Gao et al. (2005) reported that in the stomata cells of Vicia faba the vacuolar system consists of several small vacuoles when the stomata are closed. During opening the small vacuoles fuse to large vacuoles. Furthermore, a mutation in the SGR3 gene, which is involved in vacuolar fusion, leads to retardation of the stomata opening, indicating that vacuolar fusion is important for the stomata opening process. As an alternative approach to studying the structure and function of the vacuole in living cells, Dubrovsky et al. (2006) suggested using Neutral Red as a probe to investigate the structure of vacuoles with laser-scanning microscopy.

2.3. Protein targeting to vacuoles Protein trafficking to vacuoles in land plants is highly complex, due to multiple types of vacuoles (lytic and storage vacuoles) occurring in plants. A comparison with the animal and yeast lysosomal/vacuolar system was recently presented by Robinson et al. (2005). Recent research has concentrated on the vacuolar-sorting receptors (VSRs). Soluble vacuolar proteins bind to the VSR in the Golgi complex and are transported to the vacuole. Upon arrival at the vacuole or a prevacuolar compartment, the complex of cargo protein and VSR dissolves and the VSR are retrieved to the Golgi complex. So far vacuolar-sorting receptors have been investigated in detail only from pumpkin (PV72), pea (BP-80), and Arabidopsis (AtELP). All are type I integral-membrane proteins with EGF-like motifs in their lumenal domain. Vacuolar sorting receptors such as BP-80 are concentrated on post-Golgi membranes (Li et al., 2002) and are constitutively retrieved from the prevacuolar compartment to the Golgi apparatus by a saturable mechanism (Da Silva et al., 2005). The targeting of BP-80 was investigated by a mutational analysis (Da Silva et al., 2006). Da Silva et al. (2006) showed

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that the several amino acid motifs in the cytoplasmic tail and the transmembrane domain are required for proper targeting of BP-80. Pea BP-80 and Arabidopis AtELP are thought to be involved in transport to the lytic vacuole. In Arabidopsis, VSRs form a small protein family with seven members. Masclaux et al. (2005) suggested that the three phylogenetic branches observed for the seven VSRs from Arabidopsis (Masclaux et al., 2005; Paris and Neuhaus, 2002; Shimada et al., 2003) reflect different functions of the VSRs; they have tentatively assigned the three branches to the lytic, storage vacuole, and a putative endocytotic pathway (Fig. 1.1). However, Park et al. (2005) have identified AtRMR1 (a unique receptor-like protein of the ReMembR-H2 [RMR] protein family) as a VSR involved in transport to the storage vacuoles. This suggests that either structurally different receptors are involved in the same transport pathway or the interpretation of VSR phylogeny by Masclaux et al. (2005) is not correct. In animals and fungi, lysosomal/vacuolar trafficking depends on the retrograde transport of the vacuolar-sorting receptor to the Golgi complex. Several proteins forming a retromer complex have been implicated in this process. Now Shimada et al. (2006) report the characterization of the first plant mutant in a retromer component. As might be expected the mutant fails to accumulate 12S globulin and 2S albumin in the storage vacuole. Instead these proteins are secreted. Apparently, the lack of a vacuolarsorting receptor in the Golgi complex, due to failure to recycle the receptor, causes the misrouting of these proteins (Shimada et al., 2006). Other components of the complex machinery delivering proteins to the vacuoles were recently characterized. Proteins containing an epsin N-terminal homology (ENTH) domain have been identified as playing a critical role in various vesicular transport steps. The ENTH domain specifically binds phosphatidylinositols (different ENTH domains have a different lipid specificity) and is thought to be responsible for targeting these proteins to specific compartments and to assist in the formation of clathrin-coated vesicles by introducing curvature to the membrane (Legendre-Guillemin et al., 2004). Now Song et al. (2006) report that Arabidopsis epsin 1 interacts with clathrin and the adaptor-1 complex and plays an important role in vacuolar trafficking of soluble proteins. Other proteins involved in vacuolar trafficking currently also being investigated include a GTPase-activating protein in rice (Heo et al., 2005) and the role of specific syntaxin isoforms in the vacuolar system (Foresti et al., 2006). Although the basic characteristics of plant vascular sorting signals (VSS) were worked out several years ago (Vitale and Raikhel, 1999), identifying N-terminal, internal, and C-terminal VSS in several proteins, research on vacuolar-sorting signals in vacuolar proteins is still going on. VSS were recently investigated in soybean 11S globulin (Maruyama et al., 2006). Similar to the situation for BP-80 (see previous), multiple sorting also exists in the 11S globulin of soybean. In contrast, targeting of proConA to the

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vacuole depended on a new nine-amino acid-containing C-terminal propeptide, when proConA was expressed in tobacco (Claude et al., 2005). In addition, a recent investigation of VSS function in proricin showed that the position of the VSS is important for correct function of the VSS to the storage vacuole ( Jolliffe et al., 2003). Most proteins are transported via the Golgi apparatus to the vacuoles; however, some storage proteins follow other routes—the direct endoplasmic reticulum (ER) to the vacuole pathway or the autophagy pathway. Oufattole et al. (2005) have now identified an amino acid sequence PIEPPPHH directing a membrane protein to the ER to the vacuole pathway. They showed that transport depends on a putative receptor AtSRC2, which binds the sequence PIEPPPHH and is required for internalization of the ER-derived transport vesicle into the vacuole (Oufattole et al., 2005). Many plant proteins undergo proteolytic processing during their transport from the Golgi to the vacuoles. Otegui et al. (2006) have now analyzed this process in detail in Arabidopsis. They show that storage proteins and processing enzymes are packaged in separate vesicles in the Golgi. Both vesicles seem to fuse into a prevacuolar compartment, where the processing of the 2S albumin starts.

3. Structure and Function of Vacuoles in Green Algae 3.1. Types and functions of vacuoles It was 16 years ago that Domozych (1991) reviewed this topic in this series. At that time he wrote: ‘‘Little is known about the ‘non-contractile’ or cytoplasmic vacuoles of green algae, especially about their origins and functions. Because of their common occurrence in most green algae, it may be assumed that they are important in turgor control or waste storage, similar to the vacuole in higher plants’’(Domozych, 1991). In agreement with this statement the transport capacities of the tonoplast membranes in green algae were found generally to be similar to land plants (Bethmann et al., 1995; Heidecker et al., 1999, 2003a,b; Mimietz et al., 2003; Raven, 1989). Progress in this area is still slow; however, in the following I want to highlight some recent progress in this research area, but it is not the aim of this review to summarize all the work published since Domozych published his review in 1991. 3.1.1. Contractile vacuole The structure of contractile vacuoles has now been described in some detail in the two chlorophyte algae Chlamydomonas (Luykx et al., 1997a,b) and Scherffelia (Becker and Hickisch 2005, Fig. 1.2). The structure of the

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Figure 1.2 Electron micrograph of a Scherffelia dubia cell.The contractile vacuole (CV) and the polyphosphate storing vacuoles (V) are indicated. C, chloroplast; F, flagellum; G, Golgi stack; N, nucleus.

contractile vacuole in both organisms is very different. In Chlamydomonas, the large round vacuole typically observed shortly before it discharges its content into the medium develops from small vacuoles by membrane fusion (Luykx et al., 1997b). In contrast, in Scherffelia, the large round vacuoles develop from a membranous reticulum (Becker and Hickisch, 2005). From this and other work (Allen and Naitoh, 2002; Patterson, 1980) it is now clear that there is considerable variation in the structure of the contractile vacuole in protists. Whether the CVs use the same or similar mechanisms for their function cannot be answered today. Further work is required on structurally different CVs to address this problem. 3.1.2. Other vacuoles Many algae contain intracellular granules, which stain with basic dyes. These granules have been referred to as volutin or metachromic granules, and it is generally agreed that the volutin granules are storage vacuoles containing polyphosphate. Recently, this has been confirmed by EDAX analysis and

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biochemical analysis of isolated granules from Chlamydomonas (Komine et al., 2000). Since these polyphosphate-containing vacuoles possess a proton pumping pyrophosphatase (Ruiz et al., 2001), V-ATPase (Ruiz et al., 2001) and acid phosphatase (Matagne et al., 1976), the polyphosphatecontaining vacuoles of Chlamydomonas represent the lytic compartment in Chlamydomonas. These vacuoles are apparently also involved in degradation of plastidic proteins (Park et al., 1999).

3.2. Development of vacuoles In many large unicellular or multicellular algae, the cells contain a large central vacuole. Whereas the transport capacities of the tonoplast membrane have often been investigated, little is known about the development and dynamics of the large vacuoles in algae. Changes in the structure of the vacuolar system during the life cycle have been investigated only in Acetabularia (Ngo et al., 2005) using Neutral Red and light microscopy. During development the large central vacuole increases in size from 10 mm to 35 mm. Local application of the dye was used to investigate the connectivity of the large central vacuole within the thallus. Interestingly, the dye moved at different rates through different regions of the central vacuole indicating that the internal structure of the various regions of the central vacuole are different. It was concluded that the central vacuole of Acetabularia is a ramified polar organelle with a gellike sap. The morphology of the central vacuole is actively remodeled during development (Ngo et al., 2005). Cell growth is among other factors controlled by the TOR kinase in eukaryotes (Inoki et al., 2005). In contrast to land plants and similar to other eukaryotes, the TOR kinase of Chlamydomonas is inhibited by rapamycin (Crespo et al., 2005). Treatment of Chlamydomonas cells with rapamycin affected the structure of the vacuolar system possibly due to enhanced autophagy (Crespo et al., 2005), suggesting that the structure of the vacuolar system is also regulated during cell growth in small unicells. Environmental parameters (e.g., carbon dioxide, heavy metals) affect the number and function of vacuoles. Sasaki et al. (1999) reported an increase in the number of vacuoles and the activity of tonoplast proteins in Chlorococcum littoreale when cells were incubated under extremely high carbon dioxide concentrations. In another study (Nishikawa et al., 2003) it was found that at high concentrations of heavy metals (especially cadmium) the number and volume of vacuoles increased in Chlamydomonas acidophilum. Nothing is known about protein trafficking to the vacuole in green algae.

Plant Vacuoles

13

4. Evolution of Vacuolar Compartments in Plants Before discussing the evolution of the vacuolar system in plants, let me briefly summarize our current knowledge on the evolution of plants (Fig. 1.3). At the present time, there is a broad consensus that primary plastids evolved only once from a cyanobacterium that was taken up by a eukaryotic flagellate. Glaucophytes (Glaucocystophyceae), red algae (Rhodoplantae), and green algae and land plants (Viridiplantae), which dominate many of today’s ecosystems, are the descendants of this singular event (Keeling, 2004). The Viridiplantae are grouped into two phyla: the Chlorophyta, which include the Chlorophyceae, the Ulvophyceae, the Trebouxiophyceae, and most prasinophytes (scaly green flagellates); and the Streptophyta, which include a small group of freshwater algae known as Charophyceae, the scaly flagellate Mesostigma, and the embryophytes (Lewis and McCourt, 2004). Most likely, the Chlorophyta and Rhodophyta evolved in a marine environment, whereas the Streptophyta originated in a freshwater or brackish habitat (Falkowski et al., 2004; Simon et al., 2006). The vacuole is part of the eukaryotic endomembrane system, and many of the molecular components required for the biogenesis and maintenance of the endomembrane system are very well conserved between different eukaryotes (Becker and Melkonian, 1996). Therefore, the last common ancestor of the Viridiplantae inherited a typical eukaryotic endomembrane system. A general evolutionary trend in plants is the formation of a large central vacuole that allows the cell volume to increase without the need to invest in cytoplasm and other organelles. However, this is not a development specific to plants, as a large central vacuole is also found in various other organisms, (e.g., fungi and heterokont algae). The formation of a large central vacuole is observed in several phylogenetic groups within the Viridiplantae (e.g., chlorophytes, ulvophytes, and charophytes), which evolved independently from a unicellular flagellate, and, therefore, the central vacuole probably evolved several times independently even within the Viridiplantae. In many plants and green algae the vacuole adapted to specialized functions (e.g., storage of special compounds such as anthocyanins in the leaf epidermis of many angiosperms). In other cases vacuoles serve the same functions in plants and green algae; however, they use different mechanisms to perform this function. One example of this phenomenon is the storage of phosphorus in the vacuole. In algae, phosphorus is generally stored as polyphosphate, whereas in angiosperms, phosphorus is stored as phytic acid in the vacuole (Mitsuhashi et al., 2005).

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Chlorophyta: Chlorophytes Ulvophytes Trebouxiophytes Prasinophytes

Streptophyta: Mesostigma Charophytes Embryophytes

Viridiplantae

Glaucocystophyceae

Last common ancestor

Rhodoplantae

Endocytosis of a cyanobacterium

Heterotrophic flagellate

Figure 1.3 Evolution of plants. Primary plastids evolved once from a cyanobacterium taken up by a heterotroph flagellate. Glaucophytes, red algae, and green algae (and embryophytes) are the descendants of this unique event. Within the green algae two major evolutionary lines are observed: the Chlorophyta, which include most green algae (e.g., Chlorella, Chlamydomonas, Ulva); and the Streptophyta, which include Mesostigma, a small group of freshwater algae known as the Charophyceae (e.g., Charales, Coleochaetales, and desmids), and the embryophytes.

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As previously indicated, the major differences between the vacuolar system of plants and the vacuoles or lysosomes of fungi and animals is the presence of different types of vacuoles within a single plant cell. How can we explain this difference? It is important to note that seed plant cells are not unique in containing separate vacuoles with distinct functions. Many freshwater cell wall-less protists contain a contractile vacuole (CV) involved in osmoregulation and an acidic (lytic?) vacuole that might also serve a storage function (e.g., polyphosphate in algae) (see Fig. 1.2). Life on earth evolved in a marine environment. Organisms from several evolutionary lines invaded the freshwater habitat and were faced with the problem of water uptake by osmosis in a freshwater environment. Faced with a hypotonic medium, CVs evolved several times, most likely independently, using the same basic mechanisms but structurally unique solutions to the problem (Allen and Naitoh, 2002; Patterson, 1980). Some osmotolerant freshwater protists lose their contractile vacuoles when transferred into a hypertonic medium (Allen and Naitoh, 2002). Thus, protists seem to be able to ‘‘switch between a CV-containing and CV-less life style’’ (Fig. 1.4). The evolution of multicellularity took place several times on earth (e.g., animals, fungi, streptophytes [charophyte algae and land plants], red algae, brown algae). In this context it is remarkable that within the above-mentioned groups only

Animals

Marine

Chlorophytes

Streptophytes

Freshwater

Figure 1.4 Evolution of the vacuole. Life started in a marine environment.When protists invaded the freshwater habitat contractile vacuoles developed. Streptophytes are the only major evolutionary line that evolved in a freshwater environment. Freshwater protists and seed plants are the only known organisms having different types of vacuoles. Green, chloroplasts; orange, acidic (lytic) vacuole (lysosome); blue, vacuole involved in osmoregulation; gray, nucleus with nucleolus. For simplicity, heterotroph freshwater protists have been omitted.

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streptophytes made the transition to multicellularity in a freshwater environment (however, note that the question of whether fungi evolved in a marine or freshwater habitat is still not settled [ James et al., 2006]). Therefore, I propose that the separate vacuoles found in seed plants may be derived from the different types of vacuoles present in the last cell wallless ancestor of the streptophyte algae (see Fig. 1.4). How can we obtain proof for such a hypothesis? Distinct types of vacuoles in seed plants can be identified by the TIP isoform present on the tonoplast membrane: a-TIP is associated with the protein storage type of vacuoles (Park et al., 2004), gTIP is associated with the lytic type of vacuoles ( Jauh et al., 1999), and dTIP is found in pigment-containing vacuoles ( Jauh et al., 1998). TIPs belong to the aquaglyceroporin protein superfamily and represent a plantspecific subfamiliy. It has been suggested that diversification of plant aquaporins into the PIP, TIP, NIP, and SIP subfamilies preceded the divergence of bryophytes and tracheophytes (Borstlap, 2002). However, preliminary evidence indicates that differentiation of TIP isoforms might have occurred later as all moss-specific TIPs form an independent lineage in phylogenetic trees and do not cluster together with seed plant TIPs (Borstlap, 2002). In addition, the genome (http://genome.jgi-psf.org/chlre2/chlre2.home. html) of the unicellular freshwater chlorophyte Chlamydomonas probably contains only one functional aquaporin, and green algal genomes and ESTs do not contain any evidence of aquaporins of the TIP subfamily (unpublished observations). Thus, TIPs might not be the right marker to address this question. Every cell maintaining two different types of vacuoles is faced with the problem of protein targeting of vacuolar proteins to their different destinations. In seed plants proteins use different targeting signals and pathways for transport to a lytic or a protein storage type of vacuole (Vitale and Galili, 2001). So far, the latter has been reported only for seed plants. Whether these pathways are present in other streptophyte groups has never been addressed to my knowledge. If the hypothesis presented above is correct, the two pathways might be conserved within streptophytes. To address this question I tried to detect homologues of VSRs (and RMRs), storage proteins, and the plant-specific thiol protease aleurain within the Viridiplantae using BLAST analysis (Table 1.1). Homologues of VSRs and an aleurain type of thiol protease were found in all green algae, whereas the RMR-type receptor appears to be restricted to embryophytes. Because significant hits outside the Viridiplantae were not observed for any of these molecular markers, the molecular markers represent true innovations of the Viridiplantae. Aleurain and its sorting receptor VSR date back before the separation of the chlorophyte and streptophyte evolutionary line, and the RMR-type receptor, which is involved in traffic to the PSV, is clearly present in liverworts and bryophytes and might indicate the presence of PSV in embryophytes. Whether green algae contain a PSV is currently an open

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Table 1.1 Evolution of the vacuolar systema

a

b c

Protein

Chlorophytes

Streptophyte algaeb

VSR RMR Aleurain Storage proteinsc

þ

þ

þ

þ

Bryophytesb

Fernsb

þ þ þ

þ þ þ þ

The results of a BLAST analysis using angiosperm storage proteins, aleurain, VSR, and RMR-type receptor proteins (NCBI and JGI database using an expect threshold 30 mm) neurons conduct tactile and deep sensations via myelinated A-fibers. On the other hand, small-diameter (12 weeks) phenotype in skin and hair: flaky skin with pustules containing inflammatory cells and thinning of hair with abnormal hair follicles – altered terminal differentiation: suprabasal K14, increased filaggrin, loricrin, involucrin – widened intercellular spaces – localized progressive ventral alopecia with considerable variability: degenerated hair follicles (utriculi) filled with sebum and keratinized dermal cysts presenting ectopic K1 and loricrin expression indicative of interfollicular transdifferentiation – delayed ventral and dorsal hair growth after plucking – thickened epidermis in regions of alopecia: acanthosis, hypergranulosis, hyperkeratosis, increased basal and suprabsal keratinocyte proliferation with suprabasal K14 expression and strong K6 expression – altered keratinocyte differentiation – increased b-catenin–dependent transcriptional activity leading to increased cyclin D1 levels – no apparent phenotype

Reference

Hardman et al., 2005

Henkler et al., 2001

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implantation in this instance. Only 3% of homozygous mutants could be detected in E2.5 embryos at the 8 to 16 cell morula stage that were obtained from a heterozygous intercross. This demonstrates convincingly that Dsc3 fulfills crucial nondesmosomal functions, at least during this early developmental phase. One possibility is that its presence regulates compaction in some unknown way. Similarly, a nondesmosomal function is also likely for Dsg2, since its deletion prevented survival of embryonal stem cells that are derived from desmosome-free inner mass cells but did not induce defects in the desmosome-positive trophectoderm layer (Eshkind et al., 2002). The provocative conclusion is that Dsg2 also fulfills, at least under certain circumstances, nondesmosomal functions that are essential for cell proliferation. The elucidation of the molecular mechanism that leads to the decrease in PP2 levels in DSG2þ/ embryonal stem cells and its consequences on cytoskeletal organization and gene expression may help to unravel this mystery. It will be of interest to find out whether similar changes occur in the absence of Dsc3. The slightly later embryonic lethality of DP-deficient mice at E6.5 appears to be caused by different defects (Gallicano et al., 1998). A trophectoderm is formed, implantation takes place, but extraembryonic tissues do not develop properly and an overall defect in proliferation occurs. Remarkably, desmosomal-like structures were still detectable in embryonal endoderm and the ectoplacental cone, albeit at reduced number and size resulting in a collapse of the keratin filament network. Formally, it has not been excluded that partial transcripts of the mutant DSP gene are still generated giving rise to amino-terminally deleted polypeptide mutants and thereby explaining the surprising residual desmosome formation. Another bottleneck was identified for DP function by tetraploid rescue of the extraembryonic defects. In this case, lethal postgastrulation defects were noted around E10 affecting heart function, neuroepithelium, skin, and capillaries (Gallicano et al., 2001). Knockdown experiments further showed that the presence of DP is important for tube formation by investigating capillary formation in vitro (Zhou et al., 2004). To examine DP function in adult skin, epidermis-specific knockout animals were prepared (Vasioukhin et al., 2001). Although the number of desmosomes was not significantly altered, they lacked keratin filaments, and mechanical stress led to intercellular blister formation. Cultivating keratinocytes from these animals further revealed defects in epidermal sheet formation demonstrating that DP is needed for functional desmosome assembly and, even more, reinforcement of stable intercellular adhesion. Inactivation of the plakoblobin-encoding JUP gene and the plakophilin 2-encoding PKP2 gene induced embryonic lethality at E10.5 primarily due to heart defects (Bierkamp et al., 1996; Grossmann et al., 2004; Isac et al., 1999; Ruiz et al., 1996). The absence of these arm-repeat polypeptides led to redistribution of DP into granular aggregates and other junctions.

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Figure 3.12 Microscopy of reduced desmosomal adhesion in DSG3 knockout mice (DSG3-KO; A, B, D) and pemphigus vulgaris patients (PV; C). Light microscopy of hematoxylin and eosin-stained sections of epidermis reveals intraepidermal blister formation (A, B). By electron microscopy half-desmosomes are readily apparent in DSG3 knockout mice (B, D) with adhering desmoglea (arrows in D) and large intercellular spaces (double arrows in C). Bars: A, 40 mm (same magnification in C); 0.5 mm in B; 50 nm in D. (The figures are taken from Figs.4E and G and 5B and C of Koch et al.,1997, by copyright permission of The Rockefeller University Press.)

In addition, skin blistering was noted in PG mutants with a reduced number of morphologically abnormal desmosomes and retracted intermediate filaments (Bierkamp et al., 1996). It appears that in the absence of PG b-catenin can take over some of its functions, since it was localized to the residual desmosomes in this situation (Bierkamp et al., 1999). On the other hand, PG cannot fully compensate for the loss of b-catenin, which induced ectoderm defects during the gastrulation stage and subsequent lethality (Haegel et al., 1995; Huelsken et al., 2000), although it is upregulated in heart upon b-catenin depletion to maintain apparently normal cardiac structure and function (Zhou et al., 2007). Similarly, different phenotypes were elicited in X. laevis. Depletion of PG resulted in a partial loss of adhesion, and a loss of embryonic shape, but did not affect dorsal signaling (Kofron et al., 1997), whereas downregulation of b-catenin inhibited dorsal mesoderm induction in early embryos (Heasman et al., 1994). In addition, overexpression of PG in X. laevis induced anterior axis duplication upon nuclear accumulation (Karnovsky and Klymkowsky, 1995). In contrast to the strong phenotypes induced by the absence of obligatory desmosomal components, gene ablation of other components has

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generally led to less severe defects. In the case of the desmosomal cadherins, pathologies were mostly noted in skin and its appendages (DSC1, DSG3) as well as in mucous epithelia (DSG3) as detailed in Table 3.2. The phenotypes correlate well with the known expression patterns (Chidgey et al., 2001; Koch et al., 1997, 1998). They also highlight the important contribution of these adhesion molecules to mechanical stability of desmosomes, since split half-desmosomes were seen at the cell surface in blistered skin (Fig. 3.12). It is of note, however, that alterations result not only from reduced cell–cell adhesion but also include changes in cell proliferation (Chidgey et al., 2001). Cooperative effects were observed between desmosomal and classical cadherins (Lenox et al., 2000). Surprisingly little phenotypic changes were noted in a DSC1 deletion mutant lacking the Dsc1a- and Dsc1b-specific regions including the PG-binding site in Dsc1a (Cheng et al., 2004). Finally, depletion of the facultative plakins resulted in variable deficiencies in desmosome-bearing tissues ranging from skin blistering and disintegration of the intercalated discs in myocardium in the case of plectin (Andra et al., 1997), a slight defect in epidermal barrier formation for envoplakin (Maatta et al., 2001), to the absence of detectable dysfunctions for periplakin (Aho et al., 2004). Similarly, no defects were noted in desmoyokin / mice (Kouno et al., 2004).

6.2. Overproduction and ectopic synthesis of wild-type and mutant desmosomal proteins Orthotopic overexpression and ectopic production of desmosomal components as well as expression of mutant desmosomal components have been achieved by injecting specific gene constructs into the male pronuclei of murine zygotes and by examining the transgenic offspring. The resulting complex phenotypes are listed in Table 3.2. Although some of the reports contradict each other in part, several important conclusions can be drawn from these experiments. Dysbalance of desmosomal protein synthesis affects tissue homeostasis by altering tissue differentiation with coincident weakening of cell–cell adhesion and by altering proliferation. While the adhesive defects were expected, although they turned out to be rather mild-natured and affected primarily hair (Allen et al., 1996; Charpentier et al., 2000; Hardman et al., 2005; Merritt et al., 2002), changes in proliferation were much more difficult to understand. Reduced proliferation was noted in situations of PG overexpression (Charpentier et al., 2000), whereas increased proliferation was observed in mice producing elevated levels of Dsg2 (Brennan et al., 2007), or mice synthesizing either amino-terminally deleted Dsg3 or full length Dsg3 and Dsc3a/b in suprabasal epidermal cell layers (Allen et al., 1996; Hardman et al., 2005; Merritt et al., 2002). The most plausible explanation is that the levels of available PG affect cell proliferation via its suppressor function on c-MYC gene expression

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(Williamson et al., 2006). Such a dysbalance may also be the underlying reason why reduced proliferation has been reported in Dsg2 and DP knockout mice (Eshkind et al., 2002; Gallicano et al., 1998) but does not readily explain the increased proliferation in Dsc1-deficient animals (Chidgey et al., 2001). The complex crosstalk among the various plaque polypeptides of different junctions and their partially opposing effects on gene transcription in different cell types, all of which relies on specific stoichiometric ratios, remain to be elucidated to provide a molecular understanding of the underlying pathophysiological mechanisms. In addition, it is becoming clear that desmosomal cadherins present isotype-specific functions. In accordance, severe though variable defects were induced by increased suprabasal production of Dsg3 and Dsc3 that are normally restricted to the more basal compartment of the epidermis (Elias et al., 2001; Hardman et al., 2005; Merritt et al., 2002), or ectopic production of Dsg2 in suprabasal keratinocytes (Brennan et al., 2007). These situations also highlight the importance of the relative quantitative levels of desmosomal cadherins for epithelial proliferation and differentiation.

7. Interplay Between Desmosomes and Other Cell Components 7.1. Crosstalk with adherens junctions Junction formation appears to be organized in a hierarchical fashion with respect to temporal and spatial coordination. Therefore, adherens junctions initiate cell–cell contacts that are later stabilized by desmosomes. During embryonic development, desmosomes are established only after adherens junctions are formed (see above). Similarly, the same order was observed in cultured MDCK cells employing the calcium switch system in combination with specific inhibitory antibodies (Gumbiner et al., 1988). Also, cadherin function has been shown to be essential for desmosome assembly in keratinocytes (Amagai et al., 1995a; Lewis et al., 1994; Wheelock and Jensen, 1992). Detailed analysis of epithelial sheet formation revealed that this process starts with zippering of actin-containing junctions at the tips of early membrane contacts and that desmosome formation originates in flanking regions where cells were in apposition (Vasioukhin et al., 2000). Important molecular regulators of desmosome formation may be PG, p0071, and even b-catenin, which are, at least under certain circumstances, components of both types of adhaerens junctions (Hatzfeld et al., 2003; Lewis et al., 1997; Palka and Green, 1997; Setzer et al., 2004). In addition, PPs may be of particular importance, because, in addition to their dual junctional localization, they affect actin organization (Chen et al., 2002; Hatzfeld et al., 2000). Furthermore, signaling mechanisms may contribute

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to the crosstalk as suggested by the observation that a-catenin-mediated junction assembly can be bypassed by PKC to establish junctional complexes in colon carcinoma cell lines (van Hengel et al., 1997). Specifically, it has been shown that PG is required in a complex with E-cadherin to initiate desmosome assembly in A-431 cells (Lewis et al., 1997). Similarly, introduction of PG into SCC9 cells lacking PG and E-cadherin led to an increase in expression and stability of N-cadherin and a decrease in the level and stability of b-catenin, which in turn induced desmosome formation (Parker et al., 1998). An interesting example of crosstalk between desmosomes and adherens junctions in the context of overall cell behavior was provided by analyses of p0071/PP4 (Setzer et al., 2004). Overexpression of this polypeptide in A-431 cells resulted in increased adherens junction assembly and reduced desmosome assembly that were accompanied by keratin filament retraction. These cells exhibited reduced migration in an in vitro wounding system without noticeable alterations in overall adhesive strength. It was suggested that PP4 regulates PG availability. Experimental evidence has also been accrued to suggest that PP2 mediates crosstalk between desmosome formation and b-catenin signaling. It was shown that PP2 binds to b-catenin in vitro and that upregulation of PP2 enhances b-catenin signaling (Chen et al., 2002). On the other hand, PPs have been identified as Dsc-binding partners (Bonne et al., 2003). Hence, the increased levels of b-catenin transcriptional activity observed in mice producing increased levels of Dsc3 under the K1 promoter suggest a link between both via a PP signaling activity (Hardman et al., 2005). Taken together, it appears that each arm-molecule contributes in a specific way to the delicate balance between adherens junctions and desmosomes: p120ctn together with p0071 increase the assembly and stability of adherens junctions, whereas p0071 negatively affects desmosome assembly and stability. On the other hand, PP1 increases desmosomal assembly and stability (Hatzfeld et al., 2000; Sobolik-Delmaire et al., 2006; Wahl, 2005). In out-of-balance situations arm-proteins may substitute for each other. Therefore, b-catenin has been localized to desmosomes of PG knockout mice (Bierkamp et al., 1999). Conversely, disruption of desmosomal functions also weakens adherens junctions. This has been noted in DP-deficient keratinocytes (Vasioukhin et al., 2001). Here, DP was shown to be essential for the maturation of adhaerens junctions by clamping down the transient zippering ‘‘courtship’’ of classical cadherins and thereby promoting the maturation of puncta adhaerentia-type junctions and affecting cortical actin remodeling. Similarly, cells producing a DP mutant lacking its rod and IF-binding domains presented reduced mechanical resilience despite the continued presence of other adherens junctions (Huen et al., 2002). Crosstalk between desmosomes and adherens junctions also includes the transmembrane complexes. In support of this, the presence of Dsgs in

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nondesmosomal junctions has been reported in embryonal stem cells (Eshkind et al., 2002). In addition, Dsgs were found in association with b-catenin in PG-deficient keratinocytes (Bierkamp et al., 1999). Desmosomal cadherins may even associate in cis with E-cadherin as suggested by in vitro observations in low calcium conditions (Troyanovsky et al., 1999). Also, altered adherens junctions were found in transgenic mice synthesizing amino-terminally deleted Dsg3 (Allen et al., 1996). Another recent report provided evidence for the presence of an N-cadherin–catenin–vimentin complex that appears to promote cell–cell adhesion in fibroblast L-cells (Kim et al., 2005).

7.2. Crosstalk with cytoskeletal filaments 7.2.1. Intermediate filaments Even though intermediate filaments and desmosomes are attached to each other, they are not essential for the morphogenesis of the other. Therefore, desmosomes are formed in the absence of an intact intermediate filament cytoskeleton (Baribault and Oshima, 1991; Magin et al., 1998) and keratin filament networks exist in the absence of desmosomes, although they are usually collapsed around the nucleus and do not withstand mechanical stress (Gallicano et al., 1998; Troyanovsky et al., 1993, 1994a; Vasioukhin et al., 2001). Interestingly, desmosome-anchored intermediate filaments are more resistant to phosphatase inhibitors and are more long lived than nonanchored filaments, suggesting that they acquired specific properties making them particularly stable and protecting them against disassembly (Strnad et al., 2001, 2002; Windoffer et al., 2004). Desmosomes are capable of anchoring different types of intermediate filaments. Keratin filaments associate in epithelial cells, desmin filaments in cardiomyocytes, and vimentin in meningeal cells (see Fig. 3.6). The different filaments bind to the carboxy-terminus of the plakins desmoplakin, plectin, and periplakin (Fontao et al., 2003; Karashima and Watt, 2002; Kazerounian et al., 2002; Nikolic et al., 1996; Stappenbeck and Green, 1992) but use different binding motifs (Stappenbeck et al., 1993) and exhibit different binding affinities (Meng et al., 1997). 7.2.2. Actin filaments and microtubules The organization of the actin filament cytoskeleton has a major impact on desmosome formation as shown by drug-induced actin filament disassembly that resulted in compromised stability, assembly, and spatial organization of desmosomal components at the plasma membrane (Pasdar and Li, 1993). Furthermore, actin filaments have been implicated in the delivery of DP-PP2–containing particles to the plasma membrane (Godsel et al., 2005). A connection between desmosome dynamics and actin filament organization may also be mediated by p0071, which is essential for Rho

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A-mediated contractile ring formation during cytokinesis (Wolf et al., 2006). A remarkable dual function was also described for periplakin, which binds to actin via its head-rod domain and to intermediate filaments by its carboxyterminus, which can be separated in vivo by specific caspase 6-mediated cleavage (Kalinin et al., 2005). On the other hand, intact microtubules appear to be dispensable for desmosome formation (Pasdar et al., 1992). A rather unexpected finding was, however, reported recently (Lechler and Fuchs, 2007). It was shown that DP is essential for the cell type–specific organization of the microtubule system in suprabasal keratinocytes. The effect was attributed to binding of DP to the microtubule-anchoring protein ninein that becomes relocalized in suprabasal cells from a centrosomal to a junctional position around which microtubules reorient. These findings extend much earlier observations in which the plus end microtubule-binding protein CLIP170 was localized to desmosomes in polarizing MDCK cells (Wacker et al., 1992).

7.3. Crosstalk with the nucleus 7.3.1. Shuttling of desmosomal proteins It has been established that the desmosomal arm-repeat–containing polypeptides also reside in the nucleus (Bonne et al., 1999; Hu et al., 2003; Klymkowsky, 1999; Schmidt et al., 1997), where they can interact with the transcriptional apparatus to affect gene expression. In this way, direct regulatory mechanisms appear to exist that couple tight cellular adhesion to proliferation and may thus represent part of the molecular machinery that is responsible for the long-known contact inhibition, a basic property of nontransformed cells. One of the most interesting questions in this context concerns the exact origin of junctional proteins in the nucleus: Are they directly delivered from the endoplasmic reticulum to the nucleus or do they first pass through cell–cell junctions? Several observations suggest that the nuclear pool of b-catenin is fueled from the cadherin–catenin complex (Gottardi and Gumbiner, 2004; Klingelhofer et al., 2003). It is of note that PG exhibits weaker binding to the transcription factors LEF1 and TCF4 than b-catenin (Maeda et al., 2004; Williams et al., 2000) and that there are significant differences in the transactivation capacity of both (Hecht et al., 1999; Simcha et al., 1998). In contrast to b-catenin, PG decreases rather than increases the affinity of TCF4 and LEF1 for DNA (Miravet et al., 2002; Zhurinsky et al., 2000). It is therefore likely that PG antagonizes, at least in part, b-catenin, as also suggested by recent observations on the suppression of c-MYC gene expression by PG, which is in contrast to its activation by b-catenin (Williamson et al., 2006). The situation becomes even more complicated when considering the binding of PP2 to b-catenin, which upregulates b-catenin/TCF signaling (Chen et al., 2002).

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The precise molecular signals and mechanisms that determine nuclear import and export are not known. In the case of PP2 it was shown that S28 phosphorylation by the serine kinase C-TAK-1 facilitates binding to 14-3-3, thereby preventing nuclear entry (Muller et al., 2003). 7.3.2. Signal transduction It should be kept in mind that the function of cell adhesion molecules extends beyond the mere mechanical coupling of cells engaging in processes of differentiation and proliferation and thereby facilitating tissue homeostasis. They are positioned at the extracellular to intercellular interface and might therefore be important mediators of signal transduction either outside-in or inside-out. Very little is known, however, about such sensing functions. Yet, the observation that anti-Dsg3–containing sera from patients with the autoimmune blistering disease pemphigus vulgaris leads to Dsg3 phosphorylation (Aoyama et al., 1999) concomitant with a transient increase in intracellular calcium and PKC activity (Seishima et al., 1995) supports the notion of information transmission from the outside to intracellular signaling. Autoantibodies directed against Dsg3 also resulted in p38 MAPK activation and HSP27 phosphorylation (Berkowitz et al., 2005). Remarkably, all these changes take place prior to antibody-induced cell separation, suggesting that the specific antibodies do not simply interfere with cell–cell adhesion by steric hindrance (see also Waschke et al., 2005) but elicit specific intracellular signaling events. Consequently, inhibition of p38 MAPK signaling prevented the antibody-dependent keratin filament retraction and actin reorganization (Berkowitz et al., 2005). An important proof of principle was made by coinjecting PV antibodies together with specific p38 MAPK inhibitors intradermally to demonstrate that skin splitting can be prevented effectively in a living animal (Berkowitz et al., 2006). Furthermore, one of the crucial upstream components in this cascade may be the plaque protein PG whose absence also prevented PV IgG-dependent keratinocyte splitting in vitro (Caldelari et al., 2001). Another recent study implicated Rho A as a downstream target of the p38 MAPK-dependent cascade (Waschke et al., 2006). It was shown that Rho A activation and p38 MAPK inactivation inhibited PV- and PF-IgG–dependent splitting as well as cytoskeletal reorganization in an ex vivo human skin model and between HaCaT keratinocytes. Much remains to be done to work out the precise sequence of events. Even more, it will be a major challenge to determine whether these events are directly or indirectly linked to the desmosomal adhesion-dependent regulation of cell survival and proliferation. In particular, the role of PG as an important suppressor of c-MYC transcription has been acknowledged recently (Williamson et al., 2006), a mechanism that would also explain why its absence has an antiapoptotic effect in keratinocytes (Dusek et al., 2006b). Similarly, the depletion of Dsg2 and/or of Dsc3

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in knockout mice could exert its antiproliferative effects by setting PG free to suppress cell proliferation (Den et al., 2006; Eshkind et al., 2002). Conversely, desmosomal dynamics are affected by signaling molecules. The best case so far is the wounding-induced destabilization of desmosomes. It has been shown that desmosomes transit from a calcium-insensitive to a calcium-sensitive state that appears to be mediated by PKC (Wallis et al., 2000). Since the reversion to calcium independence does not rely on additional desmosomal components and occurs spontaneously in confluent cultures, it has been proposed by Garrod and colleagues (2005) that it is primarily associated with conformational changes possibly mediated by altered phosphorylation and resulting in hyperadhesion whose morphological correlate is the presence of a midline. Hyperadhesion is proposed to be the primary condition in healthy epithelia in vivo. Evidence for this model was provided by examining the mechanical stability of confluent HaCaT keratinocytes with hyperadhesive desmosomes presenting increased mechanical resilience (Kimura et al., 2006).

8. Desmosomes and Disease Much has been learned about desmosomal functions in human diseases that are associated with desmosomal malfunctions and are caused by different pathogenetic mechanisms due to either genetic defects, autoantibodies, bacterial toxins, or to malignant transformation. Table 3.3 lists the affected desmosomal polypeptides together with brief summaries of major symptoms. Since it is impossible to cover the entire literature on the topic, we will restrict the discussion to a few selected aspects.

8.1. Genetic diseases In the past few years there has been an exponentially growing list of monogenic human diseases that are caused by mutations in genes coding for desmosomal proteins (Kottke et al., 2006; McGrath, 2005; McGrath and Wessagowit, 2005). These include the desmosomal cell adhesion molecules as well as desmosomal plaque components and are associated with two major disease phenotypes: those that affect primarily the epidermis and its appendages and those whose clinical symptoms become manifest as cardiac dysfunction. Although the underlying genetic defects are known, much needs to be learned about the molecular pathogenetic mechanisms. In skin, desmosomal deficiencies caused either by DSG1 or DSP mutations lead to hyperkeratosis that often occurs in the form of prominent bands on palms and soles. Hence, these diseases are referred to as striate palmoplantar keratoderma (SPPK; Alcalai et al., 2003; Armstrong et al.,

124 Table 3.3 Summary of genetic diseases that are caused by distinct desmosomal gene mutations (Holthoefer et al., 2007) Type

Mutated gene ^ Inheritance

Clinical symptoms

Striate palmoplantar keratoderma (SPPK1) Striate palmoplantar keratoderma (SPPK2)

DSG1 – dominant

- hyperkeratotic bands on palms and soles

DSP – dominant and recessive

- hyperkeratotic bands on palms and soles (may be generalized) - varying degrees of ultrastructural alterations in desmosomes and intermediate filament organization - woolly hair and hair loss - nail dystrophy - arrhythmic right ventricular cardiomyopathy - dysplasia of skin, hair, nails - skin fragility with thickening of epidermis - trauma-induced epidermal erosions and blistering - impaired wound healing due to reduced migration - ultrastructural and molecular alterations of epidermal desmosomes

Ectodermal dysplasia skin fragility syndrome

PKP1 – recessive

Balding mouse (balJ, balpas)

DSG3 – recessive

Human: localized autosomal recessive hypotrichosis (LAH)

DSG4 – recessive

Mouse/rat: lanceolate hair phenotype (lah; Iffa Credo ‘‘hairless’’ rat) Arrhythmogenic right ventricular cardiomyopathy

DSG2 – dominant PKP2 – dominant

Naxos disease

JUP (plakoglobin) – recessive

- epidermal acantholysis ! cutaneous erosions - mucous membranes ! runting at d 8–10 - patchy alopecia - hypotrichosis of scalp, trunk, and extremities sparing facial, pubic, axillary hair - brittle, dystrophic hair: abnormal hair shaft with nodules and lance heads (trichorrhexia nodosa), impaired pigmentation, severe disturbance of inner root sheath - thickened interfollicular epidermis with hyperkeratosis and acanthosis - arrhythmogenic right ventricular cardiomyopathy - atrophy of right ventricular myocytes and replacement with fatty or fibrofatty tissue - arrhythmogenic right ventricular cardiomyopathy - striate palmoplantar keratoderma - woolly hair

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1999; Basso et al., 2006; Hunt et al., 2001; Kljuic et al., 2003b; Norgett et al., 2000; Pilichou et al., 2006; Rickman et al., 1999; Whittock et al., 1999). The hyperkeratosis is paired with ultrastructural alterations in desmosomes and widening of intercellular spaces (Armstrong et al., 1999; Milingou et al., 2006; Whittock et al., 1999). The severity of disease is generally more restricted in the case of DSG1 mutations corresponding to the type 1 SPPK, occasionally presenting only focal hyperkeratoses instead of the conspicuous striations (Milingou et al., 2006). In contrast, the DSP-caused type 2 SPPK often affects the entire body surface and goes along with defects in hair, nails, and, most notably, the heart. Corresponding differences in severity were also noted by electron and immunofluorescence microscopy revealing that desmosome number, size, and morphology are severely altered and changes in keratin filament organization and composition take place (Wan et al., 2004). The phenotypic differences between the two SPPK types are most likely accounted for by the partial compensation of Dsg1 deficiencies through Dsg3, whereas no such redundancy exists for DP. Interestingly, a pedigree was recently described with a mutation in the amino-terminal domain of DP (S299R) disrupting a putative phosphorylation site that may be implicated in PG binding and was exclusively associated with cardiac symptoms in the form of arrythmogenic right ventricular cardiomyopathy (ARVC; Rampazzo et al., 2002). This finding suggests that DP functions can be separated into those that are essential in the heart and rely primarily on molecular interaction with PG and those that are needed for intermediate filament anchorage with an essential contribution to skin integrity. The latter conclusion is also supported by another DSP mutation leading to pathological manifestations in the skin in the form of lethal acantholysis ( Jonkman et al., 2005). The underlying molecular defects were compound heterozygous mutations of DSP in which both mutated alleles had an intact amino-terminus but lacked the carboxy-terminal keratin-binding sites, thereby completely abrogating intermediate filament anchorage although desmosomes were still formed. The ectodermal dysplasia skin fragility syndrome is another type of epidermal disease that is caused by mutations in a gene encoding a desmosomal component. PKP1 has been identified as the major molecular target in this recessively transmitted affliction (Hamada et al., 2002; McGrath et al., 1997; South, 2004; Sprecher et al., 2004; Whittock et al., 2000). By immunohistology and electron microscopy desmosomal size and frequency were shown to be significantly reduced in the lower suprabasal layers (McMillan et al., 2003). Although desmosome number was also reduced in the upper suprabasal layers, their size was considerably increased in comparison to control skin. In all suprabasal layers poorly developed inner and outer plaques were seen. In addition, a widening of intercellular spaces, reduced midlines, and perturbed intermediate filament organization with anchorage defects and prominent perinuclear aggregates were noted

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(Hamada et al., 2002; McGrath et al., 1997; South, 2004; Sprecher et al., 2004). The skin fragility was paired with thickening of the epidermis and, most notably, an increased sensitivity to trauma paired with compromised wound healing. It was proposed that reduced migration is responsible for the delayed wounding response (South et al., 2003). A skin phenotype with pronounced alterations in hair was described in the balding mouse strains that carry mutations in the DSG3 gene (Pulkkinen et al., 2002). They present prominent acantholysis which, in contrast to the aforementioned diseases, is not accompanied by pronounced hyperkeratosis. These blisters are therefore considered to be a direct consequence of compromised adhesion, although we are not aware of any detailed ultrastructural analyses. As expected, the alterations are not restricted to the epidermis but extend to hair and oral mucosa, reflective of the broad distribution pattern of Dsg3. The adhesion defects, which result in impaired hair anchorage, are most notable in these mice, which are referred to as balding (bal) mice (Montagutelli et al., 1997). Almost identical defects are seen in DSG3 knockout mice (Koch et al., 1998). A different phenotype was described for DSG4 mutants that are the cause of localized autosomal recessive hypotrichosis (LAH) in humans (Kljuic et al., 2003a; Moss et al., 2004; Schaffer et al., 2006; Zlotogorski et al., 2006) and are observed in the lanceolate mouse (Kljuic et al., 2003a) and rat (Bazzi et al., 2004; Jahoda et al., 2004; Meyer et al., 2004). Most prominent are the distinct hair abnormalities that result in dystrophic alopecia and are accompanied by variable degrees of follicular hyperkeratosis. The hair is short, dysmorphic, and brittle with characteristic lance heads and alterations in the hair shaft as well as in inner and outer root sheath. The defects demonstrate that the adhesion dysfunction that most likely gives rise to the typical swelling of the hair shaft and hair loss is also coupled with impaired proliferation in the hair matrix and abnormal differentiation in the precortex zone (Bazzi et al., 2004; Jahoda et al., 2004; Kljuic et al., 2003a). It was therefore proposed that Dsg4 acts as a regulator of the transition from proliferation to differentiation (Kljuic et al., 2003a). In addition, mutation of the facultative desmosomal protein corneodesmosin that is synthesized in the upper suprabasal layers of the epidermis and in the inner root sheath of the hair follicle gives rise to the autosomal dominant disorder hypotrichosis simplex, which is characterized by reduced cell–cell adhesion in the inner root sheath and an accumulation of cytoplasmic aggregates (Levy-Nissenbaum et al., 2003). Naxos disease has long been known as a syndrome in which ARVC is coupled with palmoplantar keratoderma and woolly hair (Protonotarios et al., 1986). Originally, defects in the PG-encoding JUP gene were described as the genetic cause of this recessive disease (McKoy et al., 2000; Protonotarios et al., 2001, 2002). Interestingly, reduced connexin 43 staining can be discerned early and a significant decrease of intercalated discs and ultrastructurally identifiable gap junctions becomes evident (Kaplan et al.,

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2004) providing an explanation for the observed alterations in intracardiac conduction. Recently, it was found that DSP mutations lead to very similar disease phenotypes acting both in a dominant and recessive fashion (Alcalai et al., 2003; Norgett et al., 2000; Rampazzo et al., 2002; Yang et al., 2006). Quite unexpected was the identification of multiple mutations throughout the DSG2 gene in ARVC (Awad et al., 2006; Basso et al., 2006; Pilichou et al., 2006; Syrris et al., 2006; Tsatsopoulou et al., 2006). In this instance, the dominantly inherited disorder exhibits partial penetrance and is also characterized by cardiac disease with ventricular tachyarrhythmias. Morphologically, an atrophy of ventricular cardiomyocytes and fibrofatty replacement are pathognomonic for all types of ARVC. This again shows that a desmosomal disease phenotype is not simply due to compromised adhesion but is instead the consequence of altered differentiation and cell survival. Given the similarities to the aforementioned cardiac disease phenotypes observed in DSP and JUP mutations, it may be postulated that these factors contribute to the same pathway. Furthermore, PKP2 mutations were reported with almost identical clinical symptoms (Antoniades et al., 2006; Basso et al., 2006; Gerull et al., 2004; Tsatsopoulou et al., 2006), indicating that all factors are intertwined. Therefore, a unique possibility exists to genetically sort out the hierarchy of the cascade of these various polypeptides in the pathogenesis of ARVC by examining their levels and localization in the different genetic backgrounds. Along this line, the observed downregulation of PP2 in DSG2þ/ embryonal stem cells (Eshkind et al., 2002) can be taken as an indication that Dsg2 is upstream of PP2. Another peculiarity is that DSC2 has not been implicated in ARVC so far, possibly indicating that it lacks the specific signaling function of Dsg2. The Dsg2specific RUD region (see Fig. 3.5) would be a potential module to mediate such functions. Finally, it is puzzling why the DSG2 mutations manifest primarily in the heart and not in liver, colon, or other simple epithelia whose predominant, if not exclusive, Dsg isoform is Dsg2 (see Table 3.1). Maybe the long-lived nature of cardiomyocytes allows the full development of a clinically relevant phenotype whereas the high turnover in other tissues prevents it. Detailed examination of the various Dsg2-positive epithelia may reveal more subtle alterations. Mutations in the plakin plectin do not lead to desmosomal defects, suggesting that this molecule does not fulfill essential desmosomal functions. Instead it is primarily involved in hemidesmosomal cell–extracellular matrix adhesion in the skin thus leading to junctional epidermolysis bullosa (McLean et al., 1996; Pfendner et al., 2005; Smith et al., 1996). The evidence for a direct contribution of desmosomal gene mutation to cancer is still surprisingly scarce despite the overwhelming evidence for such a role of the related b-catenin in several malignancies, most notably those that arise from the large intestine (Behrens, 2005; Giles et al., 2003). The strongest case has been presented for PG: The presence of an allelic

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variation of the JUP gene was shown to be associated with a predisposition to familial breast and ovarian cancer and loss of heterozygosity (Aberle et al., 1995). In addition, a mutation of S28, a potential phosphorylation site, was identified in a gastric cancer (Caca et al., 1999).

8.2. Autoimmune diseases The examination of several autoimmune diseases that are caused by antibodies directed against desmosomal cadherins has contributed significantly to the understanding of the importance and the mechanisms of desmosomal adhesion. Since numerous reviews have been published on the topic (Bystryn and Rudolph, 2005; Kottke et al., 2006; Payne et al., 2004), we will only summarize a few well-established aspects of these diseases. A common feature of these life-threatening diseases is the formation of blisters that are prone to superinfection (Fig. 3.13). While Dsg3 autoantibodies result in acantholysis of lower suprabasal cell layers in the epidermis and in erosions of the oral mucosa (Figs. 3.12 and 3.13), only superficial and skin-restricted blister formation is elicited by Dsg1 autoantibodies. These differences have been known for a long time and led to the distinction of two major forms of pemphigus, namely the Dsg 3-related PV and Dsg 1-related pemphigus foliaceus (PF) that occurs also endemically as the Brazilian fogo selvagem. The desmoglein compensation theory states that both Dsgs can compensate for each other to maintain adhesion. Therefore, the lesions in PV arise in the deepest epidermal layers that lack Dsg1, leading to the notorious ‘‘tombstone-like’’ appearance (Fig. 3.13D and E). Conversely, blisters occur in PF in the most superficial layers where there is no Dsg3. This argument is further supported by experiments in which DSG3 / animals were treated with anti-Dsg1 antibodies and suprabasilar blisters developed instead of subcorneal blisters (Mahoney et al., 1999). Furthermore, regional differences in lesion formation in the skin and oral mucosa are most likely due to the different distribution of Dsg1 and Dsg3 (see Table 3.1). It is not clear, however, why Dsg2 is not able to make up for the loss of Dsg3 in PV. The pathophysiology of PV and PF has been worked out in some detail. Specific autoantibodies that are directed against distinct domains of desmosomal cadherins have been isolated from affected patients where they occur bound to the surface of epidermal cells and in circulating body fluids (e.g., Allen et al., 1993; Amagai et al., 1991, 1994; Eyre and Stanley, 1987, 1988; Ishii et al., 1997; Koulu et al., 1984; Olague-Alcala et al., 1994; Stanley et al., 1986; Fig. 3.13C). Various transfer assays demonstrated that these isolated antibodies are sufficient to induce blister formation (Amagai et al., 1992; Anhalt et al., 1982; Mahoney et al., 1999; Rock et al., 1989). Most convincingly, passive transfer of affinity-purified anti-Dsg1-antibodies into neonatal mice led to subcorneal, epidermis-restricted acantholysis

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Figure 3.13 Alterations in patients suffering from pemphigus vulgaris (A, B, D, E) and pemphigus foliaceus (C). Note the blister formation in various parts of the skin and oral mucosa of the patient shown in A and B independent of mechanical stress. The affected areas have a tendency for bacterial superinfection. Patient sera contain autoantibodies that elicit a plasma membrane staining in the epidermis corresponding to desmosomes (C). In pemphigus foliaceus staining is strongest in suprabasal cell layers (C) and in pemphigus vulgaris in intermediate cell layers.The histology of affected areas reveals intraepithelial blister formation due to loss of cell^cell adhesion either in the upper cell layers in the case of pemphigus foliaceus (not shown) or in the lower cell layers along with formation of basal‘‘tombstones’’ in the case of pemphigus vulgaris (D, E). Bars:50 mm in C, E;100 mm in D. (The original images were kindly provided by Dr. C. Sunderk€ otter and were prepared by the photolaboratory of the Clinic and Polyclinic of Dermatology, University Mˇnster, thanks to Ms. J. Bˇckmann, P.Wissel, and Dr.T. Luger.)

without mucous membrane involvement (Amagai et al., 1995b; Rock et al., 1989). Conversely, antibodies against the extracellular domain of Dsg3 caused suprabasilar blisters as well as acantholysis in mucous membranes of neonatal mice (Amagai et al., 1992). Furthermore, all pathogenic antibodies

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could be depleted from either PV or PF sera by adsorption to the extracellular domains of Dsg3 and Dsg1, respectively (Amagai et al., 1994, 1995b; Memar et al., 1996). A highly sophisticated adoptive PV mouse transfer model was recently described: Splenocytes were isolated from DSG3-/mice that had been immunized with recombinant Dsg3 and were introduced into RAG2 / immunodeficient mice (Amagai et al., 2000b). As expected, the recipient mice developed typical symptoms of PV. Thus, there is little doubt that anti-Dsg antibodies induce skin blistering. Two aspects are, however, the subject of intense discussion: (1) What is the exact molecular mechanism that leads to skin blistering? (2) How do other autoantibodies that are frequently observed in PV and PF patients contribute to the disease? For a detailed discussion of these topics, see Amagai et al. (2006). Two major alternative pathomechanisms have been discussed in the past: (1) direct inhibition of adhesion by antibodies and/or (2) involvement of intracellular signaling processes. The first is supported by observations showing that the most potent pathogenic PV monoclonal antibodies interact with the functionally important N-terminal adhesive interface (Tsunoda et al., 2003). Release of desmosomal cadherins into small clusters that are readily internalized and the formation of desmosomal halves at the cells surface are considered to be direct consequences of antibody binding (Calkins et al., 2006; Sato et al., 2000). Although pathogenic autoantibodies may not be able to penetrate inside desmosomes, it can be assumed that they are able to block turnover of desmosomal components, especially taking the high turnover rates noted in cultured systems into consideration (Gloushankova et al., 2003; Windoffer et al., 2002). The second pathogenetic mechanism is primarily supported by in vitro observations. Addition of IgG from PV sera to cultured keratinocytes caused a transient increase in intracellular calcium and inositol 1,4,5-triphosphate (IP3) (Seishima et al., 1995). This response was inhibited by an inhibitor of phospholipase C (PLC), suggesting that the latter is involved in PV-IgG-induced inositol bisphosphate hydrolysis to generate IP3 (Esaki et al., 1995). PLC also produces diacylglycerol, which activates PKC, of which certain isoforms were increased and translocated to a particulate cytoskeletal fraction within 30 sec of PV-IgG supplementation (Osada et al., 1997). PV-IgG also induced phosphorylation of Dsg3 and dissociation of plakoglobin from Dsg3 in cultured keratinocytes (Aoyama et al., 1999). Interestingly, keratin retraction from cell–cell contact sites was observed that required PGmediated signaling (Caldelari et al., 2001) and involves p38 MAPK and Rho A (Berkowitz et al., 2005, 2006; Waschke et al., 2006). It should be stressed that activation of intracellular pathways precedes cell splitting and desmosomal cadherin disengagement, which is not completely abolished by antibody binding (Waschke et al., 2005). Eventually, signaling may lead to apoptosis (Arredondo et al., 2005; Wang et al., 2004). Clearly, antibody

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binding itself does not instantaneously lead to splitting, which is temporally and spatially regulated (Shimizu et al., 2004). The ongoing controversy about the pathogenesis of PF and PV is even more complicated by the fact that patients develop, in addition to anti-Dsg1 and anti-Dsg3, a large spectrum of autoantibodies against cell surface molecules such as other Dsgs, all Dscs, E-cadherin, collagen XVIII, several subunits of the nicotinic acetylcholine receptors, and annexins, to name but a few (Amagai et al., 2006). Obviously, anti-E-cadherin antibodies, for example, act synergistically enhancing the antiadhesion effect of anti-Dsg antibodies (Evangelista et al., 2006). Also, these additional autoantibodies may be pathogenic on their own as shown for antiacetylcholine receptor antibodies both in vitro and in vivo (Nguyen et al., 2000). Similarly, it was recently reported that activation of the EGF receptor and Src kinases can still be elicited in Dsg-depleted keratinocytes by PV-IgG (Chernyavsky et al., 2007). The involvement of Dscs in autoimmune diseases has been much less explored. It is likely linked to IgA-pemphigus, a vesiculopustular dermatosis in which Dsc antibodies were identified. Although initial observations suggested that different subtypes could be correlated with immunoreactivities directed either against Dsc3 or Dsc1 (Hashimoto et al., 1996), more stringent assays could confirm the presence of only Dsc1-specific autoantibodies (Hashimoto et al., 1997). Furthermore, Dsc antibodies were also observed in Hallopeau-type pemphigus vegetans (Hashimoto et al., 1994), in pemphigus herpetiformis (Kozlowska et al., 2003), and also in PF and PV (Dmochowski et al., 1993, 1995), although these findings may not be related directly to the etiology of these diseases.

8.3. Bacterial toxins The importance of Dsgs for epidermal integrity became even more apparent when the target of Staphylococcus aureus toxins was identified as Dsg1 (Amagai et al., 2000a, 2002; Hanakawa et al., 2002). S. aureus infections are among the most common bacterial infections in children. These patients usually present with bullous impetigo that may, especially in young children but also in immunocompromised adults, develop into a generalized and life-threatening form, the so-called staphylococcal scalded skin syndrome (SSSS). In this case, subcorneal epidermal blisters appear over the entire body surface. The diseasecausing exfoliative toxins A, B, and D are serine proteases, all of which with very high specificity exclusively cleave Dsg1 after glutamic acid residue 381, which is positioned between extracellular domains 3 and 4 (see Fig. 3.5; Amagai et al., 2002; Hanakawa et al., 2002). The resulting symptoms phenocopy those observed in PF, namely cell–cell separation in the cell layers beneath the stratum corneum. In this way, bacteria open up a specific niche that is used for spreading in the otherwise hardly penetrable skin.

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8.4. Cancer The inverse interrelationship between cell–cell adhesion and malignant transformation is well recognized, and reduced cell–cell adhesion is considered to be one of the hallmarks of malignant tumors favoring formation of metastasis. The relevance of this concept for desmosomal adhesion is supported by observations of desmosome-free fibroblasts producing desmosomal components (Dsc1a and b, Dsg1, and PG) that not only exhibited increased adhesion but also reduced invasion in an in vitro assay (Tselepis et al., 1998). On the other hand, components of certain adhesion structures, most notably of desmosomes, have served as reliable tumor markers in histodiagnosis, especially in the distinction of epithelial tumors and meningiomas (Akat et al., 2003; Moll et al., 1986). Downregulation of desmosomal components has been reported for a number of different tumors such as squamous cell carcinoma of the head and neck (Bosch et al., 2005), mouth (Depondt et al., 1999; Harada et al., 1992; Hiraki et al., 1996; Shinohara et al., 1998), pharynx (Depondt et al., 1999), esophagus (Natsugoe et al., 1997), and skin (Krunic et al., 1998; Tada et al., 2000), and was seen in urothelial carcinomas (Conn et al., 1990). Reduced Dsg3 was also noted in breast cancer (Klus et al., 2001; Oshiro et al., 2003). In gastric cancer reduced and abnormally distributed Dsg2 was observed (Biedermann et al., 2005; Yashiro et al., 2006). Interestingly, a significant reduction in Dsgs was noted in comparisons of low- and high-grade intraepithelial lesions of the uterine cervix (Alazawi et al., 2003; de Boer et al., 1999). An absence of downregulation, however, was observed in colorectal carcinomas (Collins et al., 1990), and even an upregulation of Dsg2 was described in squamous cell carcinomas with a positive correlation to high-risk tumors (Kurzen et al., 2003). Dsc, Dsg, and DP expression was reported to be inversely correlated with differentiation status, invasiveness, and lymph node metastasis in oral squamous cell carcinomas (Hiraki et al., 1996; Shinohara et al., 1998). More detailed analyses suggested that the reduction of desmosomal components is most prominent in the respective invasion front (i.e., in a region where cells migrate into the surrounding connective tissue) (Depondt et al., 1999; Hiraki et al., 1996; see, however, Kurzen et al., 2003) and is related to more aggressive types of tumors some of which undergo epithelial–mesenchymal transition (for desmosome dissociation during early stages of epithelial–mesenchymal transition, see Boyer et al., 1989; Savagner et al., 1997). The direct molecular linkage between cell adhesion and altered gene expression in tumors has been elucidated, at least in part, for b-catenin (Behrens, 2005; Giles et al., 2003). b-Catenin is a major structural protein of adherens junctions as part of the linker system coupling the transmembrane cadherin adhesion molecules to the actin cytoskeleton. Nonjunctional b-catenin is usually rapidly degraded by the ubiquitin–proteasome system

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involving the serine/threonine kinase GSK 3b and the axin/APC multiprotein complex. The amino-terminally serine-phosphorylated b-catenin is recognized by the ubiquitin ligase b-TrCP. Stabilization of cytoplasmic b-catenin by wnt signaling or in tumors through a number of different mechanisms results in nuclear translocation where b-catenin interacts with LEF/TCF transcription factors leading to transactivation of LEF/TCF target genes, which include the dorsalizing genes Siamois, Twin, and Xn3, the protooncogene c-MYC, cyclin D1, fibronectin, and the matrix metalloprotease matrilysin. b-Catenin mutations occur frequently and are among the most common in colorectal carcinomas. The situation is much more complicated for the related PG, which, as pointed out above, differs in several ways from b-catenin and is rarely mutated in tumors. Most observations suggest, however, that it may function as a tumor suppressor. In accordance, reduced PG synthesis has been noted in tumors and metastatic lesions of renal cells (Buchner et al., 1998), oral and pharyngeal squamous cell carcinomas (Depondt et al., 1999), esophageal carcinomas (Nakanishi et al., 1997), prostate cancer (Shiina et al., 2005), and skin carcinomas (Tada et al., 2000). Furthermore, JUP mutations have been identified in gastric cancer (Caca et al., 1999) and loss of PG has been observed during cancer progression (Aberle et al., 1995; Amitay et al., 2001). Loss of heterozygosity has been reported in some sporadic breast, ovarian, and prostate cancers (Aberle et al., 1995; Shiina et al., 2005). The tumor suppressor function is also supported by experiments in which PG was overexpressed in various cell lines leading to inhibited cell growth and reduced tumorigenicity (Simcha et al., 1996; Winn et al., 2002). Similarly, de novo expression of PG in SCC9 squamous cell carcinoma cells led not only to epidermoid differentiation with desmosome formation but also to a decreased growth rate and increased matrix adhesiveness (Parker et al., 1998). Furthermore, overproduction of PG in skin of transgenic mice resulted in suppression of epithelial proliferation and hair growth (Charpentier et al., 2000) in line with its proposed function as a key suppressor of c-MYC in skin (Williamson et al., 2006). On the other hand, PG protects keratinocytes from apoptosis possibly by an increase in the antiapoptotic molecule BCL-XL in keratinocytes (Dusek et al., 2006b). Furthermore and in contrast to the aforementioned studies, an oncogenic potential of PG was proposed by others. Thus a strong transforming capacity was described for PG in RK3E epithelial cells (Kolligs et al., 2000) and overexpression of PG in squamous cell carcinoma cells of line SCC9 resulted in uncontrolled growth and foci formation with inhibition of apoptosis and induction of BCL-2 expression (Hakimelahi et al., 2000). The discrepancies may be explained by cell type–specific differences and indirect effects of PG on b-catenin activity. Several studies investigated the distribution of PPs in various types of carcinomas (Furukawa et al., 2005; Mertens et al., 1999; Moll et al., 1997;

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Papagerakis et al., 2003; Schwarz et al., 2006). In squamous cell carcinomas an inverse relationship between the degree of malignancy and synthesis of PP1 and PP3 was observed (Schwarz et al., 2006). Interestingly, p0071 appeared to be associated with tumor growth, exhibiting an inverse relationship to tumor size (Papagerakis et al., 2003). PP2 antibody staining was reported to be generally weak or absent in these tumors by some (Mertens et al., 1999; Schwarz et al., 2006) but not by others who found a positive correlation with metastasis formation (Papagerakis et al., 2003). PP2 was, however, consistently detectable in adenocarcinomas (Mertens et al., 1999; Schwarz et al., 2006). In hepatocellular carcinoma PP2 was the only PP except for limited PP1-positive foci with nuclear reactivity (Schwarz et al., 2006). In most other adenocarcinomas, with the exception of prostate cancer, PP2 was coexpressed with PP3, whereas PP1 was usually absent (Schwarz et al., 2006). Interestingly and somewhat in contrast to other aforementioned reports, PP3 was shown to be elevated in all non–small-cell lung carcinomas including adenocarcinomas and squamous cell carcinomas, and it was proposed to be a useful prognostic marker (Furukawa et al., 2005). Whether PPs themselves affect tumor development and progression in an isotype-specific fashion as suggested by some (Furukawa et al., 2005; Schwarz et al., 2006) and how this relates to their specific subcellular localization remain to be shown.

9. Concluding Remarks The current review attempted to provide a broad overview of the molecular diversity of desmosomes and their functions as tissue stabilizers but also attempted to point out current developments that assign additional functions to desmosomes concerning tissue differentiation and proliferation. In the coming years it will be necessary to meet the challenge to work out the molecular details of the complex interactions that occur not only within desmosomes but that also take place with outside partners to identify upstream and downstream regulators whose intricate balance affects the basic cellular machinery resulting in finely tunable shifts between differentiating and proliferating activities.

ACKNOWLEDGMENTS We wish to thank Dr. Holger Jastrow (this institute) for providing Figs. 3.1B and C, Dr. Ju¨rgen Kartenbeck (German Cancer Research Center, Heidelberg, Germany) for Figs. 3.6 and 3.7D, Dr. Peter Koch (Baylor College of Medicine, Houston, TX) for contributions to Fig. 3.12, and Dr. Cord Sunderk€ otter (Clinic and Polyclinic of Dermatology, University

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Mu¨nster, Germany) for images presented in Fig. 3.13. The authors are also grateful for the expert technical help of Ursula Wilhelm. The work was supported by the Deutsche Krebshilfe and the German Research Council.

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C H A P T E R

F O U R

Subepithelial Fibroblasts in Intestinal Villi: Roles in Intercellular Communication Sonoko Furuya* and Kishio Furuya†

Contents 1. Introduction 2. Morphological Features of Subepithelial Fibroblasts 2.1. Cell cycle, origin, and proliferation 2.2. Morphology of subepithelial fibroblasts in the small intestine 2.3. Culture of subepithelial fibroblasts 3. Receptors in Subepithelial Fibroblasts 3.1. Receptors detected by immunohistochemistry and in situ hybridization 3.2. Receptors detected by Ca2þ measurements in culture 4. Gap Junction Communication 4.1. Morphology of gap junctions in situ and in culture 4.2. Dye coupling between adjacent cells in culture 4.3. Permeability changes measured by the FRAP method 5. Mechanosensitive Networks via ATP Receptors 5.1. Mechanical stimulations evoke Ca2þ responses and ATP release in cultured subepithelial fibroblasts 5.2. Changes in mechanosensitivity with cell shape 5.3. Contractility of subepithelial fibroblasts 5.4. Propagation of Ca2þ signals from subepithelial fibroblasts to neural cells 6. Roles of Subepithelial Fibroblasts in the Villi 6.1. Regulation of the barrier/sieve function 6.2. Contractile mechanical frame and motility of the villi 6.3. Mechanosensors in the villi 6.4. Other signal transduction in the villi 7. Concluding Remarks References * {

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Section of Brain Structure, Center for Brain Experiment, National Institute for Physiological Sciences, Okazaki 444-8585, Japan Cell Mechanosensing Project, ICORP/SORST, Japan Science and Technology Agency, Nagoya 466-8550, Japan

International Review of Cytology, Volume 264 ISSN 0074-7696, DOI: 10.1016/S0074-7696(07)64004-2

#

2007 Elsevier Inc. All rights reserved.

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Abstract Ingestion of food and water induces chemical and mechanical signals that trigger peristaltic reflexes in the gut. Intestinal villi are motile, equipped with chemosensors and mechanosensors, and transduce signaling to sensory neurons, but the exact mechanisms have not yet been elucidated. Subepithelial fibroblasts located under the villous epithelium form contractile cellular networks via gap junctions. The networks ensheathe lamina propria and are in close contact with epithelium, neural and capillary networks, smooth muscles, and immune cells. Unique characteristics of subepithelial fibroblasts have been revealed by primary cultures isolated from rat duodenal villi. They include rapid reversal changes in cell shape by cAMP reagents and endothelins, cell shape– dependent mechanosensitivity that induces ATP release as a paracrine mediator, contractile ability, and expression of various receptors for vasoactive and neuroactive substances. Herein, we review these characteristics that play a key role in the villi. They serve as a barrier/sieve, flexible mechanical frame, mechanosensor, and signal transduction machinery in the intestinal villi, which are regulated locally and dynamically by rapid cell shape conversion. Key Words: Subepithelial fibroblast, Mechanosensor, Intestinal villi, ATP release, P2Y, Endothelin, Gap junction. ß 2007 Elsevier Inc. Abbreviations: BMP, bone morphogenetic protein; CBX, carbenoxolone; CGRP, calcitonin gene-related peptide; COX-1, COX-2, cyclooxygenase-1, -2; CSF, colony-stimulating factor; dBcAMP, dibutyryl cyclic adenosine monophosphate; ECM, extracellular matrix; EGF, epidermal growth factor; ET, endothelin; FCS, fetal calf serum; FRAP, fluorescence recovery after photobleaching; GABA, g-aminobutyric acid; HGF, hepatocyte growth factor; IGF, insulin-like growth factor; IL-1b, interleukin-1b; INF-g, interferon-g; KGF, keratinocyte growth factor; LPS, lipopolysaccharide; NE, norepinephrin; NGF, nerve growth factor; NK, neurokinin; PDGF-A, PDGF-B, platelet-derived growth factor-A, -B; PDGFR-a, platelet-derived growth factor receptor-a; PGE2, prostaglandin E2; PMA, phorbol myristate acetate; RT-PCR, reverse transcription polymerase chain reaction; SCF, stem cell factor; SEM, scanning electron microscopy; a-SMA, a-smooth muscle actin; SMM, smooth muscle myosin; TEM, transmission electron microscopy; TGF-a, TGF-b, transforming growth factor-a, -b; TNF-a, tumor necrosis factor-a; VIP, vasoactive intestinal peptide.

1. Introduction The gastrointestinal tract is not only a digestive and immune organ but also a mechanosensory organ (Furness et al., 1999). The ingestion and digestion of food and water may give rise to chemical and mechanical signals that induce peristaltic reflexes in the gut (Buchan, 1999; Cooke et al., 2003;

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Furness, 2000; Furness et al., 1999, 2004; Grundy, 2000; Ho¨fer et al., 1999; Holzer et al., 2001; Kunze and Furness, 1999). Even in the isolated small intestine, distention caused by intraluminal hydrostatic pressure elicits peristalsis (Bu¨lbring et al., 1958; Tsuji et al., 1992). Mechanosensors that detect changes in tension from touch, stretch, pressure, and shear stress seem to exist in the mucosa, submucosa, and muscle layers including the myenteric plexus (Cooke et al., 2003; Furness et al., 2004). It is generally thought that enterochromaffin cells, epithelial cells, and neural cells are mechanosensor cells (Cooke et al., 2003). However, the mechanosensing machinery has not yet been fully elucidated. In the gastrointestinal tract, a cellular network of subepithelial fibroblasts exists under the basal lamina of the epithelium from the esophagus to the anus. They show fibroblast-like features communicated via gap junctions, rich in actin and myosin, and are enveloping lamina propria (Desaki et al., 1984; Gu¨ldner et al., 1972; Joyce et al., 1987; Komuro, 1990; Komuro and Hashimoto, 1990; Marsh and Trier, 1974a; Pitha, 1968). Gu¨ldner et al. (1972) called them myofibroblasts because of the similarity of their filaments to those of smooth muscle. In the core region of the villus, fibroblast-like cells extend several slender processes to smooth muscles and capillaries (lamina propria fibroblasts) ( Joyce et al., 1987). Subepithelial fibroblasts and lamina propria fibroblasts communicate to each other and form three-dimensional cellular networks in the villi, which are in close contact with nerve varicosities, capillaries, smooth muscles, and immune cells in the lamina propria (Desaki et al., 1984; Gu¨ldner et al., 1972; Komuro, 1990; Komuro and Hashimoto, 1990; Toyoda et al., 1997). Based on their anatomical location, the cellular network of subepithelial fibroblasts is supposed to not only act as a skeleton for villi and influence the absorption efficiency and transportation of nutrients by possible contraction (Desaki et al., 1984; Gu¨ldner et al., 1972; Komuro, 1990) but also to play a pivotal role by mediating signal transduction in the villi. Subepithelial fibroblasts secrete components of basal lamina and function as mechanical barriers in cooperation with the epithelium. They also play major roles in the regulation of proliferation, migration, differentiation and transepithelial resistance of epithelial cells, inflammatory responses, and repair responses to injury by secretion of proteases, growth factors, and cytokines such as TGF-b, TGF-a, KGF, HGF, epimorphin, EGF, IGF, IL-1b, TNF-a, and PGE2 (Powell et al., 1999a,b, 2005). More recently, the molecular basis of these functions in morphogenesis, in immune responses, and in many disease states has been elucidated. We were interested in subepithelial fibroblasts of intestinal villi after first finding that subepithelial fibroblasts possess abundant ET receptors using electron microscopic autoradiography (Furuya et al., 1990, 1991). To reveal the physiological roles of subepithelial fibroblasts in the villi, we established

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a primary culture system of subepithelial fibroblasts isolated from rat duodenal villi (Furuya and Furuya, 1993). Cultured subepithelial fibroblasts keep cellular networks and show unique characteristics such as rapid changes in cell shape from flat to stellate and from stellate to flat depending on intracellular cAMP levels, the expression of various kinds of receptors for vasoactive and neuroactive substances including ETs and ATP, cell shape– independent gap junction permeability, contractile ability, and highly mechanosensitive networks that release ATP (Furuya and Furuya, 1993; K. Furuya et al., 1994, 2005, 2006; S. Furuya et al., 2006). These findings implied hidden but important roles of subepithelial fibroblasts in intestinal villi. In particular, we presumed that the cell shape changes are essential in every function of subepithelial fibroblasts. The structure and the function of the gut differ along the villus–crypt and the duodenum–colon axes. In the small intestine, the cell shapes of subepithelial fibroblasts are quite different in three regions: the crypt, the lower area of the villus, and the upper area of the villus (Desaki and Shimizu, 2000). In the lower area of the villus, cells are flat with broad cell processes, but in the upper area, cells are stellate with several thin processes, suggesting different functions along the villus–crypt axis. The intestinal villi are not simple amplifiers of the mucosal surface to absorb nutrients, but rather are functional surfaces to achieve both a smooth transfer and a long-term folding of the contents. It has long been known that intestinal villi are moving spontaneously (Hambleton, 1914; Lee, 1971; Nanba et al., 1970; Womack et al., 1987, 1989). In the dog duodenum, intestinal villi are repeating the rapid contraction (shortened from one-half to one-fourth of its full length) and the slower extension (Nanba et al., 1970). It is partly under neural control; however, it is still not known how these flexible and graceful properties of villi are achieved. In this chapter, we review the morphological features and the unique characteristics of subepithelial fibroblasts in intestinal villi and discuss the roles of subepithelial fibroblast networks as a barrier/sieve, flexible mechanical frame, mechanosensor, and signal transduction machinery in the intestinal villi, which are likely regulated locally and dynamically in the villi by rapid cell shape changes and cell shape–dependent mechanosensitivities.

2. Morphological Features of Subepithelial Fibroblasts The intestinal villi vary in shape depending on the regions and ages of animals. Human intestinal villi are 0.5 to 1.5 mm in height, and their appearance varies from a broad leaf like shape in duodenal villi, a tall and thin

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shape in jejunal villi, and a short and broad shape in ileal villi (Rubin, 2003). Rat intestinal villi are tongue shaped, less than 0.5 mm in height and number about 40/mm2 in rat jejunum (Komuro and Hashimoto, 1990). Under the epithelium, subepithelial fibroblasts sheathe villous projections and glandular invaginations.

2.1. Cell cycle, origin, and proliferation In the intestinal villi, epithelial cells and underlying fibroblasts proliferate in the crypts. Postmitotic fibroblasts migrate to the upper area of the villi in parallel and in approximate synchrony with epithelial cells (Marsh and Trier, 1974b; Parker et al., 1974). The epithelial cell cycle from the site of the final division in the crypt to the point of the exfoliation from the villus tip generally takes 2 to 7 days (Brittan and Wright, 2002; Wright and Alison, 1984). In the rabbit jejunum, subepithelial fibroblasts stay in the upper part of the villi longer than epithelial cells, and apoptotic cells are phagocytosed by macrophages in the lamina propria of the intestinal villi (Parker et al., 1974). Subepithelial fibroblasts and epithelial cells exert reciprocal inductive interactions in the formation of the villus–crypt unit and in cell proliferation, migration, and differentiation. Signals of epithelial–mesenchymal crosstalk involved Wnt, Notch, BMP, and PDGF signaling pathways (Brittan and Wright, 2002, 2004; Clatworthy and Subramanian, 2001; Crosnier et al., 2006; Leedham et al., 2005; Moore and Lemischka, 2006). Moreover, growth factors secreted from intestinal myofibroblasts such as EGF, IGF, TGF-b, HGF, and epimorphin are involved in the regulation of epithelial cell proliferation and differentiation (Potten et al., 1995). As for the origin of stem cells, bone marrow stem cells have recently been reported to transdifferentiate into subepithelial fibroblasts in the small intestinal villi and colon of mice after irradiation and bone marrow transplantation (Anjos-Afonso et al., 2004; Brittan and Wright, 2004; Brittan et al., 2002; Leedham et al., 2005). The proliferation of subepithelial fibroblasts is essentially regulated by platelet-derived growth factor (PDGF) (Betsholtz et al., 2001; Brittan and Wright, 2002). PDGF-A expressed in the intestinal epithelium acts by paracrine signaling through its mesenchymal receptor, PDGFR-a. Results from PDGF-A or PDGFR-a knockout mice indicate that PDGFA is essential for the normal proliferation and differentiation of PDGFR-aexpressing mesenchymal cells (Karlsson et al., 2000). In cultured colonic myofibroblasts, PDGF-AA, PDGF-BB, EGF, and FGF ( Jobson et al., 1998), but not endothelin (Kernochan et al., 2002) stimulated [3H]thymidine incorporation and cellular proliferation.

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2.2. Morphology of subepithelial fibroblasts in the small intestine 2.2.1. Scanning electron microscopic observations In specimens macerated with osmic acid (Komuro, 1985), HCl (Desaki and Shimizu, 2000; Desaki et al., 1984; Takahashi-Iwanaga and Fujita, 1985), NaOH (Toyoda et al., 1997), or enzymatic digestion (Komuro and Hashimoto, 1990), the epithelium, basal lamina, and dense reticular collagen fibrils underlying the basal lamina are successfully removed. Basal lamina contains numerous small pores in the upper two-thirds of the villi except at the very top part, and these pores are called fenestrations (Komuro, 1985). About 500 fenestrations are present on each side of a tongue-shaped villus; they are 0.5 to 5 mm in diameter, with the average being 3 mm (Komuro, 1985). These fenestrations are fewer and smaller at the base of the villi, and absent in the crypts (Komuro, 1985; Komuro and Hashimoto, 1990). Under the basal lamina, collagen fibrils and subepithelial fibroblasts form a reticular sheet that contains numerous pores ranging from 3 to 7 mm in diameter (called foramina), and approximately 400 to 800 foramina are found on each side of a villus (Toyoda et al., 1997). After removal of collagen fibrils by HCl hydrolysis, extensive cellular networks of fibroblast-like cells appear that are composed of flattened and multipolar cells with branched processes over the microvascular tree (Desaki and Shimizu, 2000; Desaki et al., 1984). The features of the cellular network are different in three regions: the upper two-thirds of the villus, the lower one-third of the villus, and the intestinal gland (crypts of Lieberku¨hn), as shown in Fig. 4.1 (Desaki and Shimizu, 2000; Komuro and Hashimoto, 1990). Subepithelial fibroblasts in the upper two-thirds of the villi appear to be stellate with several slender processes. Subepithelial fibroblasts change shape and become more stellate with a gradient to the top of the villus. These numerous slender processes contact with each other, and form a cellular sieve consisting of a large number of circles with various diameters from 0.3 to 5 mm, with the average being 3 mm (Desaki and Shimizu, 2000; Komuro and Hashimoto, 1990; Toyoda et al., 1997). These pores (fenestrations of the basal lamina, foramina of the reticular sheet, and cellular sieves of subepithelial fibroblasts) seem to link together (Desaki and Shimizu, 2000). Basal protrusions of epithelial cells and immune cells such as eosinophils, lymphocytes, and macrophages penetrate through channels formed by these pore structures. In the lower area of the villi, flat cells with broader cell processes are close to each other and often overlap (Desaki and Shimizu, 2000), and penetration of immune cells are occasional. Based on their location, these flat cells are postmitotic cells, which have migrated from the crypts. With continuity of the villi, a rough network of flattened fibroblast-like cells surrounds individual tubular glands like a basket (Desaki and Shimizu,

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A

B

Desmin

Apoptosis

Vimentin

a-SMA

Differentiation Proliferation

Figure 4.1 Cellular network of subepithelial fibroblasts along the crypt^villus axis. (A) An SEM view of a villus after removal of the epithelium and connective tissue components.The villus istentativelydivided into upper and lower areas at a levelthat istwo-thirds from the tip (A, arrowheads). In the upper area (U), stellate cells with numerous slender branches form a cellular networklike cellular sieve. Lymphocytes and epithelial extensions traverse through these sieves between the lamina propria and the epithelial layer. In the lower area (L), flat cells with broad processes are in close proximity to one other, and often overlap. (Reproduced from Desaki and Shimizu, 2000.) (B) A schematic representation of subepithelial fibroblasts in a villus. Subepithelial fibroblasts proliferate in the crypt, and postmitotic cells migrate tothe upper partof the villus, in approximate synchrony with epithelial cells. Cells continuously express vimentin, and have decreased expression of a-SMA and increased expression of desmin toward the upper region of the villus. The immunoreactivity of desmin is confirmed by polyclonal antibody, but not by monoclonal antibody.

2000; Takahashi-Iwanaga and Fujita, 1985). These cells in the crypts are thought to have proliferative activity. 2.2.2. Transmission electron microscopic observations Beneath the basal lamina of the villous epithelium, subepithelial fibroblasts exist as a cellular network with scattered collagen fibrils. In the crypt, subepithelial fibroblasts (pericryptal fibroblasts) form a relatively tight cellular network with thick collagen fibrils, generally consisting of two or three cell layers, but subepithelial fibroblasts within the upper part of the villi are more loosely arranged and form a network of one to two cell layers with few collagen fibrils (Hosoyamada and Sakai, 2005; Joyce et al., 1987; Marsh and

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Trier, 1974a). In ultrathin sections, stellate-shaped cells in SEM appear as spindle shaped with slender cytoplasmic processes (Fig. 4.2A). The cells have a well-developed rough endoplasmic reticulum filled with filamentous materials, Golgi apparatuses, round mitochondria, oil droplets, caveolae, and lysosomes (Desaki et al., 1984; Gu¨ldner et al., 1972; Joyce et al., 1987; Komuro, 1990; Marsh and Trier, 1974a; Pitha, 1968; Toyoda et al., 1997). Bundles of microfilaments about 6 nm in diameter are accumulated in the processes and the cytoplasm beneath the cell membrane facing the epithelium (Desaki et al., 1984). In the processes of subepithelial fibroblasts, intermediate filaments and microtubules are also present. The microfilaments are abundantly filled in the processes at the crypts (Pitha, 1968), but are few at the tip of the villi (Moore et al., 1989). Gap junctions and adherence junctions can be seen at the contact region of two processes (Desaki et al., 1984; Joyce et al., 1987; Komuro and Hashimoto, 1990). Occasionally, cells are enlarged with cytoplasm occupied by well-developed rough endoplasmic reticulum and they will likely synthesize reticular fibrils more actively than spindle-shaped cells with slender cytoplasmic processes (Toyoda et al., 1997). Subepithelial fibroblasts are in close contact with nerve terminals containing clear and/or dense core vesicles (Fig. 4.2C; Desaki et al., 1984; Gu¨ldner et al., 1972; Marsh and Trier, 1974a; Nagahama et al., 2001), capillaries, and immune cells such as lymphocytes, dendritic cells, and eosinophils (Deane, 1964; Komuro, 1985; Toyoda et al., 1997). In the core region of the lamina propria, fibroblast-like cells (lamina propria fibroblast cells) with their long, slender fingerlike processes partly embrace smooth muscles, capillaries, and axon bundles or varicosities (Fig. 4.2B). Microfilaments are scarce in these processes. Unmyelinated axons and subepithelial fibroblasts form synapselike structures. Axon varicosities containing small clear vesicles and large dense core vesicles are closely apposed (15 to 20 nm) to the cell bodies and thin cell processes of subepithelial fibroblasts. Synaptic vesicles accumulate facing subepithelial fibroblasts, and presynaptic densities apparently localize (Fig. 4.2C; Desaki et al., 1984; Gu¨ldner et al., 1972; Nagahama et al., 2001). Various types of motor and sensory neurons innervate into the intestinal villi, such as motor neurons (cholinergic secretomotor neurons, cholinergic and noncholinergic secretomotor/vasodilator neurons) and sensory neurons (submucosal intrinsic primary afferent neurons, extrinsic primary afferent neurons) (Furness, 2000; Furness et al., 1999, 2004; Holzer et al., 2001). Subepithelial fibroblasts seem to receive input or output from some of these motor and sensory neurons. As shown by SEM and TEM, the basal lamina is discontinuous in the upper area of the villi, but continuous in the crypts. Lymphocytes pass through fenestrae in the upper area of the villi (Desaki and Shimizu, 2000; Hashimoto and Komuro, 1988; Komuro, 1985; Komuro and Hashimoto, 1990; Toyoda et al., 1997). Sometimes, basal protrusions of epithelial cells

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Figure 4.2 Electron micrographs of subepithelial fibroblasts in rat intestinal villi. (A, B) EM autoradiographs of intravenously injected 125I-labeled ET-1 represent localization of 125 I-labeled ET-1 binding sites (silver grains; arrows) in the thin processes of subepitlial fibroblasts (SF) beneath the epithelium (Epi) (A) and lamina propria fibroblast (LF) in the

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and enteroendocrine cells penetrate into the lamina propria through fenestrae of the basal lamina and directly contact subepithelial fibroblasts, nerve terminals, and immune cells such as eosinophils and lymphocytes (Komuro, 1985; Komuro and Hashimoto, 1990; Marsh and Trier, 1974a; Toyoda et al., 1997; Wade and Westfall, 1985). These fenestrae may provide passages within the intestinal villi for nutrients, water, and immune cells. 2.2.3. Cytoskeleton and contractile proteins of subepithelial fibroblasts in vivo Small intestine Subepithelial fibroblasts in rat intestinal villi are rich in smooth muscle tropomyosin, nonmuscle isomyosin, smooth muscle isomyosin, actin, a-SMA, and cyclic GMP-dependent protein kinase (Joyce et al., 1987). Immunostaining of contractile-associated proteins is more intense in the pericryptal fibroblasts than villous subepithelial fibroblasts, and weak in the lamina propria fibroblasts (Joyce et al., 1987). In general, intestinal myofibroblasts in the small intestine and normal colon express a-SMA, smooth muscle heavy chain myosin, vimentin, and Thy-1 but do not express smoothelin, caldesmon, or desmin (Powell et al., 2005). However, by double staining with indirect immunofluorescence, we found that stellate subepithelial fibroblasts in the upper area of the rat duodenal villi were positively immunostained by monoclonal antivimentin (Fig. 4.3A1) and polyclonal antidesmin antibodies (Fig. 4.3A2). However, they were not stained by monoclonal antidesmin antibodies (not shown). We compared the results with three antibodies such as monoclonal antidesmin antibody, polyclonal antibody from desmin purified from chick gizzards, and also polyclonal antibody from a synthetic peptide mapping near the C-terminus of human desmin. Perhaps the epitope of the monoclonal antidesmin antibody is masked in the in vivo structure, but polyclonal antidesmin antibodies can detect other epitopes. The expression of desmin in the subepithelial fibroblasts from the upper area of the intestinal villi has not been reported, because many investigations used monoclonal desmin antibodies. In the top of the duodenal villi, stellate-shaped subepithelial fibroblasts are intensely desmin positive (Fig. 4.3B2), but a-SMA immunofluorescence is weak or negative (Table 4.1 and Fig. 4.3B1), although its a-SMA immunoreactivity is positively detected by incubation of the avidin–biotin complex (ABC) followed by the DAB reaction. core region of the villus. (B).The lamina propria fibroblast has several slender fingerlike processes that partly embrace smooth muscles (SM), capillary (Cap), and lymphocyte (Lym). (A: Reproduced from Furuya et al., 1990.) (C) Neural varicosity that consists of many clear vesicles and a few dense core vesicles (Neu) in close contact with thin processes of subepithelial fibroblast (SF) under the basal lamina of the epithelial layer. (Courtesy of Dr. M. Nagahama, SuzukaUniversityof Medical Science.)

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Figure 4.3 Cytoskeletons of subepithelial fibroblasts in the upper area of the rat duodenal villi (A, B) and crypts (C, D). Cryosections were double-stained for vimentin (A1, C1) and desmin (A2, C2) and for a-SMA (B1, D1) and desmin (B2, D2). In the upper area of the villi, vimentin- (A1) and desmin- (A2) positive subepithelial fibroblasts with several slender cell processes (arrows) are located under the epithelium (Epi). Desminpositive stellate subepithelial fibroblast cells (B2, arrows) are a-SMA negative (B1) by

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Table 4.1 Cytoskeletal and contractile proteins of fibroblast-like cells in the rat small intestine and colon

Small intestine Subepithelial fibroblasts in the upper area of the villi Subepithelial fibroblasts in the lower area of the villi Pericryptal fibroblasts Colon Pericryptal fibroblasts Lamina propria fibroblasts a b,c

d

Vimentin

Desmina

a-SMA

þþþ

þþ

þb or c

þþþ

þ

þþ

þþþ

–

þþþ

þþþ þþþ

–d –

þþþþ –

Positive immunostaining for desmin was detected with polyclonal antidesmin antibody, but not with monoclonal desmin antibody. Immunostaining for a-SMA in the upper area of the intestinal villi was bpositive by ABC immunohistochemistry due to a high enhancement of immunoreactivity, but cnegative or faint by indirect immunofluorescence. Subepithelial fibroblasts in the lower area of the villi and crypts were intensely immunostained by both indirect immunofluorescence and ABC immunohistochemistry. Most of pericryptal and lamina propria fibroblasts are desmin negative, but a few desmin-positive cells with long slender cell processes are present underneath the epithelium and lamina propria of the colon.

In contrast, subepithelial fibroblasts in the lower area of the intestinal villi and the pericryptal fibroblasts exhibit intense vimentin (Fig. 4.3C1) and a-SMA immunofluorescence (Fig. 4.3D1). Desmin immunoreactivity is not detected (see Table 4.1 and Fig. 4.3C2 and D2) by indirect immunofluorescence and ABC immunohistochemistry. The decrease in a-SMA immunoreactivity of stellate cells in the upper area of the villi may be due to a decrease in a-SMA expression or to a depolymerization of a-SMA. We think that desmin immunoreactivity with polyclonal antibodies is a marker for differentiated subepithelial fibroblasts in the upper part of the duodenal villi. Subepithelial fibroblasts in the jejunum and ileum exhibited very similar immunostaining patterns (S. Furuya and K. Furuya, unpublished data). The results are summarized in Table 4.1 and a schematic model for the villus is represented in Fig. 4.1B. Colon In the colon, pericryptal (subepithelial) fibroblasts show intense immunoreactivities for a-SMA and vimentin. Lamina propria fibroblasts are vimentin positive, but a-SMA negative. Most of pericryptal and lamina propria fibroblasts are desmin negative, but a few desmin-positive cells with immunofluorescence, although a-SMA immunoreactivity is detected byABC immunohistochemistry. In the duodenal crypts, pericryptal fibroblasts are vimentin- (C1) and a-SMA (D1, arrows) positive but desmin (C2, D2) negative. SM, smooth muscle.

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long thin processes are scattered underneath the epithelium and also in the lamina propria of the colon (see Table 4.1; S. Furuya and K. Furuya, unpublished data; Zhao and Burt, 2007). In the hyperplastic and neoplastic polyps of human colon, lamina propria fibroblasts change from vimentinþ, desmin , a-SMA , SMM to myofibroblasts expressing vimentinþ, a-SMAþ, SMMþ, and spotty localization of desmin (Adegboyega et al., 2002). Thus, the expression pattern of intermediate filaments and contractile proteins changes according to the villus–crypt axis, the duodenum–colon axis, and normal–pathological conditions. 2.2.4. Differential expression of the extracellular matrix and receptors along the villus–crypt axis of the small intestine The basal lamina is located between the epithelium and subepithelial fibroblasts, and both cell types contribute to the synthesis and secretion of extracellular matrix (ECM) components. Basal lamina is composed mainly of collagen IV, laminins (laminin-1, laminin-2, laminin-5, laminin-10), and heparin sulfate proteoglycans (nidogen, perlecan), and interstitial matrix is composed of collagen I, fibronectin, and tenascin (Kedinger et al., 1998; Simon-Assmann et al., 1995; Teller and Beaulieu, 2001). Expression of these molecules, especially laminins, is tightly regulated in development and along the crypt–villus axis and is associated with functional regulations and differentiation processes of epithelial cells (Simon-Assmann et al., 1995; Teller and Beaulieu, 2001). Differential distributions of the various laminins and laminin-binding integrins have been observed along the crypt–villus axis in both the developing and the adult intestine. In adult small intestines, collagen IV and laminin-1 (a1b1g1) are distributed homogeneously both in the crypt and villus. The a2 chain of laminin 2 (a2b1g1) is restricted to the crypt region. On the other hand, the a3 chain of laminin 5 (a3b3g2) and a5 chain of laminin 10 (a5b1g1) show a clear increasing gradient from the crypt to the villus tip (Kedinger et al., 1998; Simon-Assmann et al., 1995; Teller and Beaulieu, 2001). The integrins, laminin cell receptors, are a superfamily of transmembrane ab heterodimer glycoproteins that is differentially distributed in the villus. Though the integrin b1, b4, and a6 subunits are distributed both in the crypt and the villus, the a3 and a2 subunits have an opposite expression (i.e., a3 is distributed in the villi and a2 in the crypt) (Lussier et al., 2000; SimonAssmann et al., 1995; Teller and Beaulieu, 2001). Tenascin C is predominantly localized in the upper position of the intestinal villus, but not in the crypt, indicating an importance for cell shedding at the villous tip (Aufderheide and Ekblom, 1988; Probstmeier et al., 1990). In contrast, fibronectin predominantly localizes in the crypt, with a decrease in gradient from the crypt to the top of the villus (Simon-Assmann et al., 1995).

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Cytokines, combinations of TNF-a and IFN-g, can alter some types of laminin expression in the epithelial cells (Francoeur et al., 2004), and expression patterns of laminins and tenascin differ in intestinal pathologies (Salas et al., 2003; Teller and Beaulieu, 2001). In the intestinal villi, the cell shape of subepithelial fibroblasts is flat at the lower position of the villi, which changes to a stellate structure at the upper two-thirds of the villi. As described in Section 2.3.2, shape conversions depend on intracellular cAMP concentrations in vitro. Factors that trigger stellate formation of subepithelial fibroblasts in vivo are unknown; however, it seems that the differential expression of the ECM secreted from subepithelial fibroblasts and the epithelium may be one factor that regulates cell shape and/or differentiation in a paracrine and autocrine manner.

2.3. Culture of subepithelial fibroblasts 2.3.1. Intestinal myofibroblast cell lines Several clonal myofibroblast cell lines have been established from rats and human intestinal mesenchyme. These cloned cell lines express a-SMA, vimentin, and smooth muscle myosin heavy chain, but not desmin (Powell et al., 1999a,b, 2005). 18Co cells, a myofibroblast clone derived from human colon, change cell shape by dBcAMP treatment (Valentich et al., 1997), just as primary cultured subepithelial fibroblasts of duodenal villi (Furuya and Furuya, 1993). However, 18Co cells take longer time (4 to 24 h) to change cell shape compared with primary cultured villous subepithelial fibroblasts (15 to 60 min). 18Co cells contain carbachol receptors coupled to PGE2 synthesis that elicit the stellate transformation (Kim et al., 1998; Shao et al., 2006). The proinflammatory cytokine IL-1 increases secretion of PGE2 (Kim et al., 1998; Valentich et al., 1997) and also induces a wide variety of cytokine secretions (Rogler et al., 2001). Clonal myofibroblast cell lines isolated from different levels of gut axes (rat jejunum, ileum, and colon) display regional characteristics, that is, secrete different expression patterns of growth factors such as HGF, TGF-b1, and epimorphin (Fritsch et al., 2002; Plateroti et al., 1998), and they regulate differentiation of epithelial cells in a region-specific manner (Plateroti et al., 1998; Ratineau et al., 1997). Two morphologically distinct clones isolated from the rat ileum, F1G9 and A1F1, display clear-cut differences in the morphogenesis and differentiation of the epithelial cells (Fritsch et al., 1997, 2002; Kedinger et al., 1998; Subramanian et al., 2001). The F1G9 clone expressing a-SMA responds to TGF-b and induces crypt–villus structures in vivo. In contrast, the a-SMAnegative A1F1 clone induces cryptlike structures. F1G9 and A1F1 clones alter the level of Cdx-1 and Cdx-2 homeobox genes differentially in epithelial cells (Duluc et al., 1997).

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As each clonal cell line maintains specific gene expression profiles through replicated culture passages (Powell et al., 2005), they are useful in studying epithelial–mesenchymal interactions, immune cell–mesenchymal interactions, repair, and tumorigenesis. 2.3.2. Primary culture of subepithelial fibroblasts isolated from duodenal villi To study the functions of subepithelial fibroblasts, it is necessary to keep cells in a differentiated cellular network similar to those under physiological conditions in vivo. We established a primary culture system that maintains their two- or three-dimensional networks of differentiated subepithelial fibroblasts just as observed in vivo (Furuya and Furuya, 1993). In this culture system, the proliferation of subepithelial fibroblasts is scarce, and more than 90% of cells are subepithelial fibroblasts, which show a rapid reversal change in cell shape (Furuya and Furuya, 1993). Moreover, subepithelial fibroblasts are highly sensitive to many bioactive substances (Furuya et al., 1994). Therefore, these cultures are adequate in examinations that measure cellular responses such as electrophysiology, Ca2þ measurements, and morphological analyses. In contrast to cloned myofibroblast cell lines, this primary culture system is not adequate for biochemical analyses, because several other types of cells in the lamina propria contaminate, such as capillary endothelial cells, smooth muscle cells, and immune cells. Culture method Epithelium-free villi were obtained from 10- to 12-dayold rat duodenal villi by incubation with Ca2þ- and Mg2þ-free phosphatebuffered saline followed vibration. Subepithelial fibroblasts migrate within 1 day from epithelium-free lamina propria and form monolayer cellular networks. To prevent dedifferentiation, the concentration of FCS is decreased from 10% to 1–3% after migration. Characteristics of subepithelial fibroblasts such as cell shape conversion and Ca2þ responses to various bioactive substances are well maintained for 2 to 4 days and disappear in older cultures. Most cells began to die after 1 week. In cultures isolated from 3-week-old rats, cell migration from the clusters is slow, and the threedimensional structure of the lamina propria is well maintained. They are more sensitive to endothelins and other bioactive substances than cultured cells isolated from 10- to 12-day-old rats, and contractions evoked by mechanical stimulations (touch or stretch) are intense (K. Furuya and S. Furuya, unpublished data). Cell shape conversions Migrated subepithelial fibroblasts from the epithelial-free villi are flat shaped with several broad processes. By treatment with 0.3 to 1 mM dBcAMP, 10 mM forskolin, 20 mM cholera toxin or 0.5 mM PGE2, more than 90% of migrated cells change to a stellate morphology within 15 to 60 min, which represents a round cell body with several

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slender processes (Fig. 4.4A). Stellate-shaped cells are flattened upon the addition of 10 7 to 10 9 M endothelins (ET-1, ET-3) or FCS (see Fig. 4.4A). Subepithelial fibroblasts change cell shape reversibly depending on the intracellular concentration of cAMP (Furuya and Furuya, 1993; Furuya et al., 1994). These phenomena are very similar to those observed in astrocyte cultures (Table 4.2; Moonen et al., 1976; Shapiro, 1973). In colonic 18Co cells, the coaddition of carbachol and IL-1 elicited the synthesis of PGE2 followed by a stellate transformation (Kim et al., 1998; Shao et al., 2006; Valentich et al., 1997). However, subepithelial fibroblasts isolated from duodenal villi did not respond to carbachol or acetylcholine by Ca2þ measurements (K. Furuya et al., 1994, 2005). Substance-P elicits an intense intracellular Ca2þ increase but does not induce cell shape changes (Furuya et al., 1994). Both ET-1 and ET-3 induce cell shape changes from stellate to flat. This process was blocked with a combination of an ETA antagonist, BQ123 and

Figure 4.4 Rapid reversal shape conversion of cultured subepithelial fibroblasts isolated from rat duodenal villi. (A) Subepithelial fibroblasts convert cell shape reversibly from flat to stellate upon treatment with dBcAMP and from stellate to flat shape upon the addition of FCS and ET-1. (Reproduced and modified from Furuya and Furuya, 1993.) (B) Subepithelial fibroblasts uptake vitamin A in culture, and vitamin A possessing flat-shaped cells (B1) convert to stellate shape by dBcAMP treatment, while maintaining vitamin A fluorescence (B2).Vitamin A was detected by autofluorescence excited with 360 nm using a UV laser confocal microscope.

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Table 4.2 Comparison of characteristics between subepithelial fibroblasts and astrocytes in primary culture Subepithelial fibroblasts of duodenal villia

Proliferation Actin a-SMA Intermediate filaments Gap junctions Connexins

 or ? þþþ þ Vimentin, desmin þþ CX43

Permeability cAMP-independent Not inhibited by ETs Stellate conversion With cAMP Rapid, þþþ reagents d With carbachol Contraction þþ Effect of ETs Cell shape change Contraction Transient Ca2þincrease and oscillation Mechanical stimuli, ATP Ca2þ wave triggers a b c d

Astrocytesb

þ þþþ þc Vimentin, GFAPc, desminc (partially) þþ CX43(major) CX26 and Cx30 (a few) cAMP-dependent Inhibited by ETs

Rapid, þþþ þþ þ or  Cell shape change Proliferation Transient Ca2þincrease and oscillation Mechanical stimuli, ATP, glutamate

Subepithelial fibroblasts isolated from duodenal villi summarized from Furuya and Furuya (1993), K. Furuya et al. (1994, 2005), and S. Furuya et al. (2005). Astrocytes summarized from Buniatian et al. (1999), Kettenmann and Steinha¨user (2005), Ransom and Ye (2005), Shain et al. (1992), and Shapiro (1973). Expression of a-SMA, GFAP, and desmin are variable with the regions of the brain, ages of animals, and culture periods (Buniatian et al., 1999). Villous subepithelial fibroblasts change to stellate shapes and respond to PGE2 but do not respond to carbachol. In contrast, colonic myofibroblast 18Co cells have carbachol receptors coupled to PGE2 synthesis that elicit stellate transformation.

an ETB antagonist, BQ788. In cultures isolated from ETB-mutant (sl/sl ) rats (Karaki et al., 1996; Kunieda et al., 1996), reversal changes in cell shape were induced similar to wild-type cells (S. Furuya et al., 2005). However, shape conversion from stellate to flat by ET-1 was blocked only by BQ123. Shape conversion from stellate to flat by ET-1 is attributable to both ETA and ETB receptors in wild-type cells, but only to ETA receptors in the sl/sl cells (S. Furuya et al., 2005). The signal for ET-1-induced stress-fiber formation is transmitted through a Gq/PCL- and G12-dependent pathway to the Rho/ ROCK system examined in ETA transfected CHO cells (Kawanabe et al., 2002),

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and stress-fiber formation in astrocytes is inhibited by C3 ADP-ribosyltransferase (Koyama and Baba, 1996). In culture, cells abutted to smooth muscles in the center of the clusters and change shape reversibly and more rapidly than cells migrated to the periphery. In vivo, lamina propria fibroblasts extend their long, slender fingerlike processes (see Fig. 4.2B), and immunoreactivities for contractile proteins are absent or less than those seen in subepithelial fibroblasts ( Joyce et al., 1987). Cyclic AMP concentrations in lamina propria fibroblasts may be higher than those in subepithelial fibroblasts in vivo and in vitro. Both types of cells are continuous and form a cellular network that communicates with gap junctions as described in Section 4. The aforementioned results may indicate that subepithelial fibroblasts and probably lamina propria fibroblasts have common cellular characteristics and form three-dimensional cellular networks in vivo. Compared to in vivo structures observed by SEM (see Fig. 4.1A; Desaki and Shimizu, 2000), cells cultured with medium containing 10% FCS show flattened cell bodies with broad processes, which are similar in morphology to subepithelial fibroblasts in the lower area of the villus. Subepithelial fibroblasts treated with dBcAMP, forskolin, cholera toxin, or PGE2 display features such as a round cell body with several slender processes. Subepithelial fibroblasts in the upper area of the villi in vivo have a stellate multipolar shape and seem to be a transient form between a flat shape and highly stellate shape with a round cell body and several thin processes. Uptake of vitamin A in subepithelial fibroblasts In situ, 3H-labeled vitamin A is incorporated into oil droplets of both subepithelial fibroblasts and lamina propria fibroblast-like cells in mouse intestinal villi (Hirosawa and Yamada, 1977), as observed in hepatic stellate cells (Hirosawa and Yamada, 1973). Vitamin A storing ability is one of the unique characteristics of these fibroblast-like cells. In culture, almost all subepithelial fibroblasts isolated from duodenal villi take up vitamin A (Fig. 4.4B1) and change cell shape from flat to stellate by dBcAMP treatment (Fig. 4.4B2).

2.3.3. Ultrastructure of cultured subepithelial fibroblasts In primary cultures of subepithelial fibroblasts isolated from rat duodenal villi, cells are rich in thick bundles of microfilaments, intermediate filaments, microtubules, expanded rough surface endoplasmic reticulum filled with amorphous materials, Golgi apparatuses, oil droplets, and caveolae (Fig. 4.5A; Furuya and Furuya, 1993). Subepithelial fibroblasts in primary culture show features very similar to those in vivo. In 3- to 4-day cultures with 10% FCS, collagen fibers abut the periphery of flat-shaped cells (see Fig. 4.5A). Upon 1 mM dBcAMP or 10 mM forskolin treatment for 30 min to 2 h, the cell shape changes from flat

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Figure 4.5 Electron micrographs of cultured subepithelial fibroblasts. (A) Flat cell represents thick bundles of microfilaments (mf), rough surface ER (rER) expanded with amorphous materials, electron-dense oil droplets, and collagen fibers abutting cell surface (col: arrow and arrowhead). mvb, multivesicular body. (B, C, D) Stellate cell treated with dBcAMP represents round cell body (B) and thin elongated processes (C, D) with microtubules (mt) and intermediate filaments (fil). Note the disappearance of bundles of microfilaments. (Reproduced from Furuya and Furuya,1993.)

to stellate such as a round cell body (Fig. 4.5B) with several thin processes (Fig. 4.5C and D). In the thin processes, thick bundles of microfilaments disappeared though microtubules and bundles of intermediate filaments are present (see Fig. 4.5C and D). Gap junctions are revealed between adjacent cell processes by freeze-fracture analysis and by functional dye coupling both in flat- and stellate-shaped cells (shown in Section 4).

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2.3.4. Cytoskeleton of cultured subepithelial fibroblasts Villous subepithelial fibroblasts cultured for 1 to 4 days are rich in F-actin, aSMA, vimentin, and desmin (Fig. 4.6 and Table 4.2). Although subepithelial fibroblasts of rat duodenal villi in vivo are immunoreactive only to polyclonal antidesmin antibody, cultured subepithelial fibroblasts are immunoreactive not only to polyclonal antidesmin antibody but also to monoclonal antidesmin antibody. The presence of desmin in cultured cells indicates that these cells are in a differentiated state, which corresponds to those at the upper

Figure 4.6 Cytoskeletons in cultured subepithelial fibroblasts. (A) Stellate cells treated with dBcAMP (A1, phase contrast) are vimentin (A2) and desmin (A3) positive. Intensely immunostained vimentin-positive but desmin-negative large spindle-shaped cell (arrow in A1 and A2) is not subepithelial fibroblast. (B) Fat-shaped cells express a-SMA (B1) with a ‘‘stress-fiber’’ pattern and are desmin positive (B2). (C) Treatment with dBcAMP induces depolymerization of a-SMA (C1) and rearrangement of desmin (C2) in the stellate cells.

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two-thirds of the villus (Fig. 4.6A3, B2, and C2; compare cell features with Fig. 4.3B2). By dBcAMP treatment, about 90% of cells converted to a stellate shape and these cells express vimentin (Fig. 4.6A2) and desmin (Fig. 4.6A3). A few intensely vimentin-positive but desmin-negative cells are contaminated in the culture (see Fig. 4.6A2, arrow). They do not change cell shape and are usually spindle shaped and larger than subepithelial fibroblasts (Fig. 4.6A1). Most probably, these contaminants represent other type of cells. A thick stress fiber pattern of a-SMA is present in flat-shaped cells (Fig. 4.6B1). Treatment with dBcAMP induces depolymerization of filamentous a-SMA and decreases a-SMA immunofluorescence (Fig. 4.6C1). Vimentin and desmin are reorganized from dispersed conformations to form bundles according to changes in slender processes (Fig. 4.6A2 and A3). Changing from stellate to flat by the addition of ET-1, ET-3, or FCS, many thin fibers appear within 10 min (Furuya and Furuya, 1993).

3. Receptors in Subepithelial Fibroblasts From their anatomical location in the villi, subepithelial fibroblasts are supposed to possess many receptors to transmitters released from nerve terminals, vasoactive substances from vessels, and neuropeptides secreted from endocrine cells in the epithelium and from mast cells in the lamina propria. Here we summarize the receptors obtained by immunohistochemical observations and Ca2þ measurements.

3.1. Receptors detected by immunohistochemistry and in situ hybridization 3.1.1. Endothelin receptors ET-1 binding sites revealed by electron microscopic autoradiography EM autoradiography of intravenously injected 125I-labeled ET-1 revealed the localization of endothelin receptors on the long processes of both subepithelial fibroblasts beneath the epithelium (see Fig. 4.2A) and lamina propria fibroblasts (see Fig. 4.2B). Receptor-mediated endocytosis was seen 30 to 60 min after the injection (Furuya et al., 1990, 1991). These receptors were confirmed as ETA receptors by immunoelectron microscopy (S. Furuya et al., 2005). Light microscopic immunohistochemistry and in situ hybridization ETA receptors are located in the subepithelial fibroblasts throughout the rat duodenal villi and crypts (Fig. 4.7A; S. Furuya et al., 2005) and colon. ETB receptors are located in the enteric neurons and glial cells of submucosal and myenteric plexi (Fig. 4.7B). In the normal rat colon, the mRNA of

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Figure 4.7 Localization of ETA, ETB, and P2Y1receptors in the ratduodenum and in culture. (A, B, C) In the rat duodenum, ETA receptors (A) and P2Y1 receptors (C) are located in subepithelial fibroblasts (arrows) throughout villi and crypts. ETB receptors are restricted to the ganglion cells in both submucosal and myenteric plexus (B: arrows). (D, E, F) In cultured subepithelial fibroblasts, ETA (D) and P2Y1 receptors (F) diffusely are located throughout the cells. The expression of ETB receptors is evoked by culture and they mostly accumulate to the perinuclear region, probably in the Golgi apparatus (E).

ETA is abundantly expressed in subepithelial fibroblasts, and the mRNA of ETB receptors is expressed only in a few pericryptal fibroblasts in addition to the vasculature and neurons (Egidy et al., 2000a). Under pathological condition, both ETA and ETB receptors are abundantly expressed in many colonic myofibroblasts (Egidy et al., 2000b). In culture, ETA receptors are diffusely located in almost all subepithelial fibroblasts (Fig. 4.7D) (S. Furuya et al., 2005). ETB expression is induced by culture, and receptors localize mostly to the perinuclear region, probably to Golgi apparatuses (Fig. 4.7E). 3.1.2. P2Y1 receptors P2Y1 immunoreactivity is distributed in subepithelial fibroblasts throughout rat intestinal villi and crypts (Fig. 4.7C; S. Furuya et al., 2005) and colon. In culture, P2Y1 receptors are located diffusely throughout the cells (Fig. 4.7F).

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3.1.3. NK1 receptors NK1 receptor (substance-P receptor) immunoreactivity is abundantly located in the pericryptal region and lamina propria of the human colon (Riegler et al., 1999). In mouse small intestine, immunoelectron microscopy revealed localization of intense NK1 immunoreactivity to the processes of subepithelial and lamina propria fibroblasts (Vannucchi and Faussone-Pellegrini, 2000). 3.1.4. Angiotensin II receptors Angiotensin II receptor AT1 and AT2 immunoreactivities are located in a-SMAþ myofibroblasts in human colonic mucosa (Hirasawa et al., 2002). In the intestinal villi, AT1 immunoreactivity is abundant in the lamina propria and the muscular layer but not restricted to subepithelial fibroblasts. AT2 immunoreactivity is present in the lamina propria up to the mid portion of the villi in the lamina propria (Johansson et al., 2001).

3.2. Receptors detected by Ca2þ measurements in culture Ca2þ measurement is often used to detect receptors in a certain cell, although receptors that do not elicit Ca2þ change cannot be detected. Receptors detected by Ca2þ measurements are shown in Table 4.3 and are compared to astrocytes. 3.2.1. Vasoactive and neuroactive substances In 2- to 4-day cultures of subepithelial fibroblasts isolated from rat duodenal villi, intracellular Ca2þ increases transiently upon the addition of 10 mM ATP, 10 nM bradykinin, 1 nM ET-1, 10 nM ET-3, and 100 nM substance-P, respectively (Fig. 4.8A). Almost all cells intensely respond to endothelins, substance-P, and ATP. Some subepithelial fibroblasts respond to 10 mM serotonin (40% of cells). We previously reported that some cells respond to angiotensin II (Furuya et al., 1994); however, this probably belongs to another cell type, because they did not convert cell shape upon treatment with dBcAMP (K. Furuya and S. Furuya, unpublished data). Subepithelial fibroblasts do not respond to glutamate, GABA, acetylcholine, NE, VIP, oxytocin, or vasopressin. Though C-type natriuretic peptide (CNP) is reported to relax colonic myofibroblasts (Chitapanarux et al., 2004), CNP did not elicit a Ca2þ response in cultured subepithelial fibroblasts (K. Furuya and S. Furuya, unpublished data). Colonal 18Co cells have carbachol receptors coupled to PGE2 synthesis that elicit cell shape changes (Valentich et al., 1997), and VIP elevates cAMP concentration in the human colonic myofibroblasts ( Jobson et al., 1998). These findings suggest that the cell shape of subepithelial fibroblasts of human colon is modulated by cholinergic and peptinergic inputs in vivo. However, subepithelial fibroblasts isolated from rat duodenal villi did not

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Table 4.3 Ca2þ responses to various neuroactive and vasoactive substances and subtypes of receptors between subepithelial fibroblasts of intestinal villi and astrocytes in primary culture

ET-1 ET-3 Receptors Bradykinin Receptors Angiotensin II ANP Receptors VIP Vasopressin Receptors Oxytocin Substance-P Receptors Serotonin Glutamate Receptors GABA Receptors Norepinephrin Receptors Acetylcholine Receptors Histamine Receptors ATP UTP Receptors a b

Subepithelial fibroblastsa

Astrocytesb

þþþ (oscillation þ) þþ (oscillation þ) ETA> ETB þþþ not identified

þþþ (oscillation þ) þþþ (oscillation þ) ETB>ETA þþþ B1, B2 þ þþ ANPA, ANPB, ANPC þþ þþ V1A þþ þþþ NK1, NK2, NK3 þþ þþþ mGluR,iGluR(AMPA) þþ GABA A, GABAB þ a1,b2 þ mAchR,nAchR þ H1, H2 þþ þþþ P2X,P2Y1,P2Y2

þ or  ?

? þþþ NK1 þþ ? ? ? ? ? þþþ þ P2Y1

Data from Ca2þ response experiment (K. Furuya et al., 1994, 2005, 2005, and unpublished data). Summarized from Finkbeiner (1995); Kettenmann and Steinhauser (2005). Responses vary within regions of the brain.

respond to acetylcholine or carbachol in Ca2þ measurements, or change cell shape (K. Furuya et al., 1994, 2005). 3.2.2. Subtypes of endothelin receptors in culture Applications of ETs (0.1 to 100 nM) evoke transient and sometimes oscillatory Ca2þ responses in cultured subepithelial fibroblasts isolated from duodenal villi (Fig. 4.8B). ET-1 is nearly one order more effective than ET-3. The Ca2þ

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Figure 4.8 Intracellular Ca2þ responses to various substances in cultured subepithelial fibroblasts. (A) Typical responses to serotonin (5HT, 10 mM), substance P (Sub-P, 100 nM), ATP (10 mM), bradykinin (BK,10 nM), and endothelin-1 (ET1,10 nM). Acetylcholine (Ach, 100 mM) and vasoactive intestinal peptide (VIP, 5 mM) were not effective. (B) Ca2þ responses to ET1 (10 nM) in normal Ringer (left) and in Ca2þ -depleted solution (right). (C) Time courses of morphological changes were measured simultaneously with Ca2þ measurements. Morphological changes were monitored by fluorescence changes in the cell, indicating that cell shape changes from stellate to flat are fast (about 10 min) and Ca2þ independent. (Modified from Furuya et al.,1994.)

response to 10 nM ET-1 is inhibited by an ETA antagonist, BQ123 (100 nM), but not by an ETB antagonist, BQ788 (100 nM), and the response to 10 nM ET-3 is inhibited by BQ788 but not by BQ123. In addition to these pharmacological results, RT-PCR and Western blotting analyses indicate both ETA and ETB receptors are expressed in cultured subepithelial fibroblasts, although ETA is more prominent (S. Furuya et al., 2005). As mentioned previously, ETA receptors are dominantly expressed in subepithelial fibroblasts of the small intestine in vivo and in culture (see Fig. 4.7A and D).

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3.2.3. Subtypes of ATP receptors in culture Cultured subepithelial fibroblasts respond to some nucleotides. Even in Ca2þ-free solution, a similar Ca2þ response is initially observed. The order of potency is 2MeSATP > ADP ATP  UTP > UDP; here, 2MeSATP (2-methyl thio ATP) is known to act specifically on P2Y1 from the P2Y family. MRS2179 (30 mM), a P2Y1 antagonist, blocks the Ca2þ response to 2MeSATP, but MRS2159 (100 mM), a P2X1 antagonist, has no effect. In addition to these pharmacological results, RT-PCR and Western blotting analyses indicate P2Y1 is dominantly expressed in ATP receptors in subepithelial fibroblasts (K. Furuya et al., 2005). 3.2.4. Comparison of cellular characteristics and Ca2þ response between subepithelial fibroblasts and astrocytes In diverse tissues, special types of fibroblast-like cells that are rich in actin filament similar to smooth muscle localize at specific anatomical locations in close contact to epithelial cells, parenchyma cells, neural cells, and vasculature (Komuro, 1990). They are called myofibroblasts (Gabbiani et al., 1971; Powell et al., 1999a,b) because they are activated by injury and inflammation and express a-SMA, which produce the force of wound contraction. Myofibroblasts are usually quiescent in vivo and express specific morphologies and functions in each tissue (Komuro, 1990; Powell et al., 1999a,b; Zhao and Burt, 2007). Subepithelial fibroblasts of the small intestine and colon, hepatic stellate cells, and brain astrocytes are typical myofibroblasts that are located at the blood–tissue interface and express a number of similar properties in vivo and in vitro (Buniatian et al., 1999; Cassiman et al., 2002; Knittel et al., 1999; Ramadori and Saile, 2002; Senoo, 2004). In particular, subepithelial fibroblasts of intestinal villi and astrocytes form a cellular network communicated via gap junctions composed of connexin 43 in vivo and in culture. They change cell shape rapidly and reversibly from flat to stellate by cAMP reagents (Furuya and Furuya, 1993; Goldman and Abramson, 1990; Moonen et al., 1976; Shain et al., 1992; Shapiro, 1973) and from stellate to flat by endothelins or FCS (Furuya and Furuya, 1993; Goldman et al., 1991; Koyama and Baba, 1996; Koyama et al., 1993). These cell shape conversions accompany depolymerization and polymerization of actin and rearrangement of intermediate filaments. In contrast, hepatic stellate cells do not form a syncytium in the sinusoid. Hepatic stellate cells in culture take 48 h to change their cell shape from flat to stellate after the application of dBcAMP (Kawada et al., 1996), so its process seems to be quite different from the rapid-reversal shape conversion process in subepithelial fibroblasts and astrocytes in culture. However, subepithelial fibroblasts of intestinal villi and hepatic stellate cells have common characteristics such as uptake and storage of vitamin A in the oil droplets in vivo (Hirosawa and Yamada, 1973, 1977) and contraction elicited

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by endothelins (K. Furuya et al., 2005; Kawada et al., 1996, 1999; Kernochan et al., 2002). Cell characteristics and Ca2þ responses to transmitters and hormones are compared with cultured subepithelial fibroblasts and astrocytes (see Tables 4.2 and 4.3). Both types of cells respond well to ET-1, ET-3, substance-P, bradykinin, ATP, and serotonin (Finkbeiner, 1995; Goldman et al., 1991; Kettenmann and Steinha¨user, 2005; K. Furuya et al., 1994, 2005; S. Furuya et al., 2005). In the villous sensory and motor neurons, acetylcholine, NE, substance-P, VIP, CGRP, and neuropeptide Y are revealed by immunohistochemistry (Furness, 2000). Cultured villous subepithelial fibroblasts did not respond to these transmitters except substance-P and serotonin, although colonic myofibroblasts responded to carbachol (Valentich et al., 1997). In contrast, astrocytes contain many such receptors including glutamate, acetylcholine, NE, GABA, and glycine (Finkbeiner, 1995; Kettenmann and Steinha¨user, 2005). Astrocytes are enveloping all neurons in the brain. So it is reasonable that astrocytes respond to almost all transmitters, although the responses vary within regions of the brain. Astrocytes dominantly express ETB receptors more than ETA (Hori et al., 1992; Sasaki et al., 1998), and P2Y2 subtypes are major purinergic receptors rather than P2Y1 (Zhu and Kimelberg, 2001). Subtypes of receptors in astrocytes have been examined by immunohistochemical, biochemical, pharmacological, and RT-PCR analyses (Finkbeiner, 1995; Kettenman and Steinha¨user, 2005). As described in Section 5, both types of cells are mechanosensitive. Mechanical stimuli evoke ATP release and intercellular Ca2þ wave propagations in a cell shape–dependent manner in astrocytes (Cotrina et al., 1998b, 2000) and in subepithelial fibroblasts (K. Furuya et al., 2005). Although subepithelial fibroblasts do not respond to glutamate, glutamate evokes Naþ waves in parallel with Ca2þ waves in astrocytes, which give rise to spatially correlated increases in glucose uptake (Bernardinelli et al., 2004).

4. Gap Junction Communication 4.1. Morphology of gap junctions in situ and in culture Gap junctions are present between adjacent cell processes of subepithelial fibroblasts in small intestinal villi by ultrathin sections (Desaki et al., 1984; Joyce et al., 1987; Komuro and Hashimoto, 1990). In culture, small clusters of gap junctions were revealed between adjacent cells by freeze fracturing (Fig. 4.9B and C; Furuya and Furuya, 1993) and immunohistochemistry (Fig. 4.9D and E). Gap junctions are frequently present on the cell periphery of flat cells and slender processes of stellate cells. About 64% of clusters are small (see Fig. 4.9C), and the remaining 36% are larger (see Fig. 4.9B). The number

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Figure 4.9 Gap junctions in the cellular network of cultured villous subepithelial fibroblasts. (A) Lucifer yellow injected into a stellate-shaped cell treated by dBcAMP (A1, A2, arrow) was rapidly transferred to adjacent cells (A2). (B, C) Freeze-fracture images of stellate cells show gap junctions between thin processes (B, C, arrows, arrowhead). Most gap junctions are small (C, arrow), and a few are very small (C, arrowhead). (Modified from Furuya and Furuya,1993.) (D, E) Connexin 43 immunoreactivity is localized at the periphery of the flat cells (D, arrows) and thin processes of stellate cells (E, arrows).

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and the size distribution of gap junction clusters do not change significantly during cell shape conversion between flat- and stellate-shaped cells. Connexin 43 is the main component of gap junctions revealed by RT-PCR analysis and immunohistochemistry of cultured subepithelial fibroblasts (Fig. 4.9D and E; S. Furuya et al., 2005).

4.2. Dye coupling between adjacent cells in culture In cultured subepithelial fibroblasts, Lucifer yellow (MW 522) injected into a cell spread rapidly to adjacent cells within several tens of seconds, indicating cell–cell coupling via gap junctions (Fig. 4.9A1 and A2). This dye coupling is present both in flat cells and stellate cells treated with dBcAMP, indicating that gap junctions are open independently with the concentration of intracellular cAMP.

4.3. Permeability changes measured by the FRAP method FRAP (fluorescence recovery after photobleaching) is a noninvasive method that measures the permeability of dyes through gap junctions and can measure the change within the same cell under different conditions. Using this method, the dynamics of permeability can be measured even in very thin cells such as subepithelial fibroblasts. Calcein (MW 622.5) was loaded into cells using an acetoxymethyl ester (AM) form, and several cells were photobleached by intense laser exposures. If gap junctions are open, nonbleached calcein is transferred to photobleached cells from the adjacent cells, and the fluorescence recovers. Recovering time courses were fitted exponentially with a recovery time constant. Gap junction permeability varied with each culture. Generally, gap junctions are open with recovery constants less than several hundred seconds (Fig. 4.10A). Treatment with carbenoxolone (CBX, 100 mM ), a gap junction blocker, perfectly suppressed the recovery, and the recovery was observed after washout (see Fig. 4.10A; S. Furuya et al., 2005). In some cultures, gap junctions were closed in all measured cells. The application of dBcAMP, ETs, and substance-P had no effect on opening these closed junctions. Gap junction permeability is controlled by various factors such as membrane potential, pH, intracellular Ca2þ concentration, adenyl cyclase, phospholipase C, phospholipase A2, and tyrosine kinases (Enkvist and McCarthy, 1992; Giaume and McCarthy, 1996; Paulson et al., 2000; Ransom, 1995; Ransom and Ye, 2005). In astrocytes, the gap junction permeability increases by the elevation in intracellular cAMP concentration and decreases by ETs treatment (Blomstrand and Giaume, 2006; Blomstrand et al., 1999, 2004; Ehrenreich, 1999; Giaume et al., 1992). In villous subepithelial fibroblasts, the gap junction permeability is unchanged by the increase or decrease in cAMP level (Fig. 4.10B; S. Furuya et al., 2005). To avoid ambiguity caused

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Figure 4.10 Gap junction permeability and its cell shape dependence measured by the FRAP method. Several calcein-loaded subepithelial fibroblasts were photobleached by intense laser exposure. Fluorescences of the cells gradually recover when gap junctions are open. Recovery time course is a good indicator of gap junction permeability. (A) Recovery time courses and the inhibitory effect of CBX. Changes in the normalized intensity in the eight cells are averaged. The recovery of fluorescence was completely blocked by CBX (100 mM) treatment and retrieved by washout. (B) Effects of cell shape on recovery time course. Open circles show nontreated flat cells and closed circles show dBcAMP-treated stellate cells. (C) Comparison of recovery time constants obtained from the same cells under different conditions: nontreated (control), dBcAMP treated for 20 to 60 min, and 10 nM ET-1 treated for about 10 min. Each data point is the average of five experiments. Bars represent standard errors. (Modified from S. Furuya et al., 2005.)

by variations between culture to culture, the gap junction permeabilities were measured in the same cells but under different conditions. Calculated fluorescence recovery constants did not change in the same cells under different conditions: control (flat shape), 20 to 60 min after dBcAMP treatment (stellate shape), and 10 min after ET-1 application (reflattened) (Fig. 4.10C). The gap junction permeability in subepithelial fibroblasts is independent of changes in intracellular cAMP concentration, at least in the short term (10 min to 2 h). Blomstrand et al. (2004) reported that ET inhibits astrocyte gap junctions in wild-type rat hippocampal slices and in culture, but not in ETB mutant preparations. The phosphorylation of Cx43 is inhibited by ETs in wild-type rat astrocytes, but not in ETB mutant forms. This result indicates that the ET-induced inhibition of gap junction permeability does not occur through ETA receptors alone, and that the inhibition requires the integrity of ETB receptors. Astrocytes predominantly express ETB receptors (Hori et al., 1992). On the other hand, ETA is the dominant functional receptor of subepithelial fibroblasts. Therefore, a reason must exist as to why the gap junction permeability in subepithelial fibroblasts is not significantly modified by ET-1. Recently, Bedner et al. (2006) measured the cAMP permeability of different types of connexin channels, Cx26, 32, 36, 43, and 45, using cyclic nucleotide–gated channel activity measurements. As a result,

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the Cx43 gap junction is the most permeable gap junction to cAMP with up to a thirtyfold difference in efficacy. Although, the open/close mechanism of gap junctions in subepithelial fibroblasts is uncertain, the network of subepithelial fibroblasts may maintain intercellular communication via gap junctions independent of changes in cell shape. This property may help in the rapid transduction of signals (including cAMP) evoked in a local area to spread to the whole network, even under adverse cellular conditions. This means that cell shape itself is regulated in some large areas of the villus, but not within each cell. Cell shape seems to be an essential factor for the functioning of subepithelial fibroblast networks within villi.

5. Mechanosensitive Networks via ATP Receptors ATP and other nucleotides are important and are the most ubiquitous extracellular messengers in various kinds of tissues and organs (Abbracchio and Williams, 2001; Burnstock, 2006; Burnstock and Knight, 2004). ATP is often released upon mechanical stimulation, and it activates surrounding cells via many subtypes of P2Y metabotropic and P2X ionotropic ATP receptors (Guthrie et al., 1999; Schwiebert, 2000). This process forms intercellular Ca2þ waves, propagating cell to cell in an autocrine/paracrine fashion in many types of cells, for example, mast cells (Osipchuk and Cahalan, 1992), astrocytes (Arcuino et al., 2002; Blomstrand et al., 1999; Cotrina et al., 1998a,b, 2000; Fields and Stevens, 2000; Guthrie et al., 1999; John et al., 1999; Koizumi et al., 2003; Ostrow and Sachs, 2005; Schwiebert, 2000), keratinocytes (Denda et al., 2002; Koizumi et al., 2004), liver epithelium (Frame and de Feijter, 1997), and pancreatic islet cells (Cao et al., 1997). ATP plays pivotal mechanotransduction roles in many tissues and organs. For example, in blood vessels, mechanical shear stress by blood flow induces ATP release from endothelial cells and ATP enhances mechanosensitivity (Burnstock, 1999; Yamamoto et al., 2000); in the airway, mechanical stress induced by foreign substances evokes ATP release from ciliated epithelial cells, and ATP enhances salt and water transport, ciliary beat frequency, and mucin secretion to increase defense mechanisms (Hansen et al., 1993; Homolya et al., 2000); in bone, shear stress and mechanical loading induce ATP release from osteocytes and ATP enhances bone formation (Hoebertz et al., 2003; Robling et al., 2006; Rubin et al., 2006); in mammary alveoli, mechanical stress induced by myoepithelial cell contraction evokes ATP release from secretory epithelial cells, and ATP enhances milk secretion (Furuya et al., 1997, 2004; Nakano et al., 2001); and in tubular and bladder visceral organs, such as intestine (Burnstock, 2001) and urinary bladder (Cockayne et al., 2000; Knight et al., 2002),

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the distention or distortion of these organs induced ATP release from the epithelium, and ATP mediates nociceptive mechanosensory transduction. Recently, we revealed that subepithelial fibroblasts are highly sensitive to mechanical stimulations and release ATP by touch or stretch stimulations. From these findings, we have proposed that subepithelial fibroblasts work as a mechanosensor in the intestinal villi via ATP release (K. Furuya et al., 2005, 2006).

5.1. Mechanical stimulations evoke Ca2þ responses and ATP release in cultured subepithelial fibroblasts Two types of mechanical stimulations were applied to cultured subepithelial fibroblasts isolated from rat duodenal villi. One type of stimulation involved touching a cell with a fine glass rod, and the other stimulation involved stretching of cells that were cultured on an elastic chamber made with silicone elastomers (K. Furuya et al., 2005). We measured intracellular Ca2þ changes with indo-1 fluorescence using an UV-laser scanning confocal microscope and released ATP with luciferin-luciferase bioluminescence assay and bioluminescence real-time imaging methods. 5.1.1. Touch-induced ATP release followed Ca2þ waves The touching of a subepithelial fibroblast with a fine glass rod induces an intracellular Ca2þ increase in the cell, and the Ca2þ increase propagates to surrounding cells (intercellular Ca2þ waves). Ca2þ waves propagate 150 to 200 mm in radius with a speed of 5 to 10 mm/sec. Ca2þ wave propagations are reversibly blocked by MRS2179 (100 mM ), an inhibitor of P2Y1, but not by CBX (100 mM ), a gap junction blocker (Fig. 4.11A). Ca2þ waves propagate to separate cells where no physical contact exists between cells (K. Furuya et al., 2005). The touching of cells in a Ca2þ-free solution also induces Ca2þ increases in the cell and Ca2þ waves in surrounding cells during initial stimulations. These results strongly suggest that touch induces a release of nucleotides from stimulated cells, which activates P2Y1 receptors in surrounding cells. To confirm the result, the touch-induced ATP release was measured using a real-time imaging system of luciferin-luciferase bioluminescence. By touching a cell briefly, intense luminescence occurred, and it diffused to the surroundings (Fig. 4.11B), confirming the release of ATP from the touched cells. So mechanical stimulation of subepithelial fibroblasts by touching induces ATP release and released ATP activates P2Y1 in the surrounding cells. These processes propagate intercellular Ca2þ waves in networks of subepithelial fibroblasts.

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Figure 4.11 Mechanical stimulations induced intercellular Ca2þ waves and ATP releases. (A) Touch mechanical stimulations induce intercellular Ca2þ waves. A slight touch with a blunted thin glass rod induces intercellular Ca2þ waves (control). Ca2þ waves continue for about 40 sec and propagate about 200 mm. MRS2179 (100 mM), a P2Y1 blocker, inhibited the Ca2þ waves. CBX (100 mM), a blocker of gap junctions, did not affect the initiation and propagation of the Ca2þ waves. (Modified from K. Furuya et al., 2005.) (B) Imaging of ATP released by touch mechanical stimulations. Released ATP was visualized by luciferin-luciferase bioluminescence using a real-time luminescence imaging system, where the luminescence was amplified using an image intensifier and a high-sensitivity cooled CCD camera. The luminescence images are represented by green and are superimposed on Nomarski images, which are taken simultaneously with luminescence imaging using infrared. (C) Imaging of ATP released by stretch mechanical stimulations. A stretching (16%, to vertical direction) of the cells in elastic chambers induced ATP release (green) in several cells within the colony (0.5 sec). The released ATP spread to the surroundings (2 sec, 3 sec).

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5.1.2. Stretch-induced ATP release and Ca2þ responses Another type of mechanical stimulation, the stretching (8 to 60%, 1 to 3 sec) of cells cultured on silicone elastomers, also induces Ca2þ increases (e.g., Fig. 4.12B). The response is transient and disappears after a few tens of seconds. The numbers of responsive cells increase upon an increase in stretch length but not stretch duration or stretch speed (K. Furuya et al., 2005). Stretch mechanical stimulations also induce ATP release in subepithelial fibroblasts. Released ATP causes delayed wavelike Ca2þ responses in the colony. Luciferin-luciferase bioluminescence assays confirmed the release of ATP by stretching. ATP was detected by the luminescence measurement of perfusate (e.g., Fig. 4.13A) and by the real-time luminescence imaging of cells (Fig. 4.11C). ATP is released only by 8 to 10% stretch and the amount increases with the length of stretch.

Figure 4.12 Cell shape-dependent mechanosensitivities. Ca2þ responses by mechanical stimulations, touch (A) and stretch (B) in subepithelial fibroblasts, were suppressed in dBcAMP-treated stellate-shaped cells (upper panels in both A and B). Mechanosensitivities were recovered in ET1-treated flat-shaped cells (lower panels in both A and B). (Modified from K. Furuya et al., 2005.)

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Figure 4.13 Cell shape^dependent ATP release by stretch. In dBcAMP-treated stellate-shaped cells, strong stretches (32 to 40%) induced a small amount of ATP release. ET treatment changed the cell shape to flat. At that time, a small ATP release was observed. After that, the same stretches induced larger (about seven times) ATP releases. ATP contents in the perfusate were assayed every minute by luciferin-luciferase bioluminescence using a luminometer. (Modified from K. Furuya et al., 2005.)

5.2. Changes in mechanosensitivity with cell shape As shown in Section 2.3.2, subepithelial fibroblasts change shape from flat to stellate depending on the intracellular cAMP level. Mechanosensitivities are highly cell shape dependent (Fig. 4.12; K. Furuya et al., 2005). In stellateshaped cells treated with dBcAMP, the touch mechanical stimulation does not evoke Ca2þ waves (Fig. 4.12A, upper part), and the stretch does not induce Ca2þ increases (Fig. 4.12B, upper part). ETs (1 to 10 nM treatment induces cell shape change from stellate to flat within 10 min. In reflattened cells, touch stimulation elicits Ca2þ waves similar to controls (Fig. 4.12A, lower part), and stretch stimulation evokes Ca2þ increases (Fig. 4.12B, lower part). Suppression of Ca2þ waves in stellate-shaped cells suggests suppression in ATP release from touched cells and/or suppression of ATP sensitivity in surrounding cells. First, the amount of ATP released by stretch was measured in the perfusate. The release of ATP is suppressed in dBcAMP-treated stellate-shaped cells and is recovered or further enhanced in ET-1–treated reflattened cells (Fig. 4.13A). Substance-P (Sub-P, 100 mM ), which is known to induce Ca2þ increases but not morphological changes in subepithelial fibroblasts (Furuya et al., 1994), did not recover the ATP release. On average, dBcAMP treatment (over 1 h) suppressed the ATP release to about 11% of that under control conditions, and further treatment with ET1 (10 nM, 10 to 20 min) recovered and enhanced ATP release about 3.5 times (Fig. 4.13B). Second, cell shape dependence of ATP sensitivity was checked. The Ca2þ response to ATP (10 mM) decreased to about 40% in dBcAMP-treated

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stellate-shaped cells, and the suppression was recovered by ET-1 treatment (K. Furuya et al., 2005). These findings indicate that cAMP-mediated intracellular signaling causes cell shape changes, which accompany changes in mechanosensitivities and ATP sensitivities. In many cell and tissue types, mechanically induced ATP release and the following activation of ATP receptors are observed, and ATP release is thought to be an important and ubiquitous mechanosensing process (Arcuino et al., 2002; Burnstock, 2001; Furuya et al., 2004; Schwiebert, 2000). ATP is released by several pathways: the exocytosis of vesicles (Bodin and Burnstock, 2001; Coco et al., 2003; Osipchuk and Cahalan, 1992), anion channels (Hisadome et al., 2002; Sabirov et al., 2001), hemichannels of gap junctions (Cotrina et al., 1998a; Stout et al., 2002), transporters (Bodin and Burnstock, 2001; Roman et al., 1997), and transient nonselective membrane channels (Arcuino et al., 2002). However, the details of the mechanism, the molecules included, and an overview of ATP release have not yet been clarified. In subepithelial fibroblasts, thick stress fibers depolymerize during shape conversions from flat to stellate by dBcAMP or forskolin treatment, and stress fibers repolymerize in the cells changed from stellate to flat shape by ETs treatment (Furuya and Furuya, 1993). According to these morphological changes, the sensitivity of ATP release to mechanical stimulation changes remarkably. Upon treatment with Y27632, a Rho kinase inhibitor, stress fibers in flat-shaped cells diminish and mechanically induced ATP release is inhibited, although ATP sensitivity remains unchanged (K. Furuya et al., unpublished observations). These findings suggest an actin cytoskeleton contribution to ATP release in subepithelial fibroblasts as indicated in cultured astrocytes (Cotrina et al., 1998b).

5.3. Contractility of subepithelial fibroblasts As subepithelial fibroblasts are morphologically characterized as rich in contractile-associated proteins (see Sections 2.2.3 and 2.3.4), they are thought to be contractile. Contractility is a critical feature for playing roles in intestinal functions: wound healing from epithelial injuries (Powell et al., 1999b), villous movements in normal (Gu¨ldner et al., 1972) and disease states (Moore et al., 1989), and the regulation of mechanical properties of the villi (see Section 6). However, only a few reports have ascertained the contractility of subepithelial fibroblasts (Kernochan et al., 2002). When a subepithelial fibroblast is stimulated by touch, transient cellular contractions sometimes propagate concomitantly with Ca2þ wave propagation (Fig. 4.14A; K. Furuya et al., 2005, 2006). Cell contractions reach a maximum after about a 10-sec delay (Fig. 4.14A3) following the Ca2þ peak increase (Fig. 4.14A2). In the cultures isolated from 3-week duodenal

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Figure 4.14 Released ATP conveys the wave of contraction of subepithelial fibroblasts and the signal to neurons. (A) Propagating contractions of subepithelial fibroblasts. Transient cell contractions were observed following touch-evoked Ca2þ waves (A1). Ca2þ changes (A2) and contractions (A3; relative cell length) in each cell (shown in A1, 10.2 sec) were plotted. Contractions peaked at about 10 sec following Ca2þ peaks. (B) Propagation of Ca2þ signals from subepithelial fibroblasts to neurons. Differentiated NG108^15 neural cells were cocultured on subepithelial fibroblasts (B1). Neurons (N) extend well-developed processes (N. process) on subepithelial fibroblasts. Touching of subepithelial fibroblasts evoked Ca2þ waves (white parts). Ca2þ signals spread through subepithelial fibroblasts (cell 1, 2, and 3) and also reached the neural processes (a, b) and neural cell body (c). Traces of Ca2þ changes in several subepithelial fibroblasts and in neurons (a, b, and c) are shown (B2).

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villi, which are maintaining villous structures, mechanically elicited Ca2þ waves induce intense villous contractions (K. Furuya et al., unpublished observations). The application of ATP (0.1 to 100 mM) and ETs (0.1 to 10 nM) also induces contractions following the Ca2þ increase (K. Furuya et al., 2005). The contraction elicited by ATP is only transient but that by ETs is somewhat persistent, similar to the Ca2þ response. These transient and brief cellular contractions of subepithelial fibroblast networks may affect the mechanical properties of villi and may partly cause villous motility. As networks of subepithelial fibroblasts overlay the vascular tree, contraction of subepithelial fibroblasts may regulate the permeability of the vasculature.

5.4. Propagation of Ca2þ signals from subepithelial fibroblasts to neural cells Subepithelial fibroblasts may function as a mechanosensor in the villi as discussed in Section 6.3. To confirm which Ca2þ wave in subepithelial fibroblasts propagates to adjacent neurons, differentiated NG108–15 cells were cocultured with subepithelial fibroblasts. Differentiated NG108–15 cells possess neural properties and are rich in P2Y1, P2Y2, P2Y4 and P2X4, mRNA. Ca2þ waves, evoked in the network of subepithelial fibroblasts by touch mechanical stimulation, propagated in the cell processes of NG108–15 cells (Fig. 4.14B; K. Furuya et al., 2005). This finding suggests that mechanically induced ATP release from subepithelial fibroblasts can propagate to and activate purinergic sensory neurons in the villi.

6. Roles of Subepithelial Fibroblasts in the Villi A distinctive property of subepithelial fibroblasts is the fast and reversal shape conversions that are dependent on cAMP level. The network of subepithelial fibroblasts provides mutual communication through gap junctions regardless of their cell shape. Cyclic AMP can also pass through gap junctions, so shape conversions do not simply occur in each cell but rather within a widespread area of the network. Cell shape conversions alter many properties, such as the size of cellular sieve, tension, contractility, and the mechanosensitivity of the network (Fig. 4.15). By these properties, networks of subepithelial fibroblasts become unique multifunctional apparatuses in the villi. Subepithelial fibroblasts are postulated to have the following roles: they act as (1) a barrier/sieve, (2) a regulator of mechanical properties, and (3) a mechanosensor in the villi.

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Flat Stellate cAMP concentration

Size of cellular sieve Gap junction permeability Mechano-sensitivity (ATP-release, Ca2+-waves) ATP-sensitivity tension, contractility

Figure 4.15 Summary of how cell shapes affect the properties of subepithelial fibroblast networks.

6.1. Regulation of the barrier/sieve function 6.1.1. Control of sieve size by cell shape conversion The basal lamina and underlying subepithelial reticular sheet in the upper area of intestinal villi have numerous small pores or fenestrae (called fenestration, foramula, or cellular sieve) ranging from 0.3 to 5 mm in diameter (Desaki and Shimizu, 2000; Komuro, 1985; Komuro and Hashimoto, 1990; TakahashiIwanaga and Fujita, 1985; Toyoda et al., 1997). Fenestrae work as sieves or passages for nutrients and migrating immune system cells (Desaki and Shimizu, 2000; Komuro, 1985). Fenestrae in the basal lamina are scarce and small in size at the lower part of the villi, and absent within the crypts (Desaki and Shimizu, 2000). This seems to well reflect processes such as absorption in the upper part of the villi and secretion in the crypts. Subepithelial fibroblasts change shape quickly and drastically by the application of ETs, depending on intracellular cAMP levels (Figs. 4.4 and 4.15; Furuya and Furuya, 1993). Endogenously, ETs are thought to be released from epithelial cells, neurons, and immune cells such as mast cells and macrophages under their specific physiological and pathological condition. The sieve size of subepithelial fibroblast network may therefore change on local and dynamic levels upon cell shape conversions. In addition to the physical control of the sieve size, subepithelial fibroblasts regulate epithelial permeabilities through the release of humoral factors. 6.1.2. Regulation of epithelial barrier functions In a variety of physiological and pathological conditions, bidirectional cytokine signals among epithelia, subepithelial fibroblasts, and immune cells result in the modulation of tight junctions between epithelia. Nutrients and bacterial

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toxins also affect tight junctions (Clayburgh et al., 2004; Nusrat et al., 2000; Shen and Turner, 2006). In cocultures of colonic myofibroblasts and epithelial cell lines, the secretion of TGF-b from myofibroblasts enhances barrier functions and modulates electrogenic chloride secretion in epithelial cells (Beltinger et al., 1999; Powell et al., 1999b). Subepithelial myofibroblasts secrete various growth factors and cytokines including TGF-b, HGF, and TNF-a, which regulate the assembly of tight junctions (Beltinger et al., 1999; Plateroti et al., 1998; Powell et al., 1999b; Valentich et al., 1997; Walsh et al., 2000). In contrast to TGF-b, HGF (Grisendi et al., 1998), TNF-a, and IFN-g (Clayburgh et al., 2004; Poritz et al., 2004) disrupt tight junctions through the phosphorylation of myosin light chain kinase. Thus, subepithelial fibroblasts in vivo may locally and dynamically regulate epithelial permeabilities for nutrients, ions, and H2O by releasing growth factors and cytokines (Fig. 4.16A).

6.2. Contractile mechanical frame and motility of the villi The intestinal villi move spontaneously with retraction–extension and bending modes (Fig. 4.16B; Hambleton, 1914; Lee, 1971; Nanba et al., 1970; Womack et al., 1987, 1989). Networks of subepithelial fibroblasts sheathe the lamina propria of intestinal villi just like a nylon stocking, and the networks are contractile and generate tension. Thus, they work as a mechanical frame to maintain the flexibility and contractility of the villi. Mechanical properties of networks, such as tension and contractility, are dependent on properties of individual network components, that is, of subepithelial fibroblasts. Subepithelial fibroblasts change shape temporarily and locally between flat and stellate structures, depending on their local environmental conditions. Flat cells abundantly express filamentous a-SMA, which leads to high tension and contractile cells. On the other hand, stellate cells demonstrate a lower immunoreactivity for a-SMA, probably due to depolymerization, which leads to lower tension and less contractile cells (see Figs. 4.15 and 16B). Subepithelial fibroblasts change shape along the crypt to the villus axis (e.g., a flat shape in the crypts and lower part of the villi and a stellate shape in the upper part of the villi) (see Fig. 4.1), meaning the lower part of the villi is tighter or maintains a higher tension and is more contractile than the upper part of the villi. These features determine the mechanical properties of the intestinal villi. In addition to these regional differences, subepithelial fibroblasts can rapidly change shape by ETs. ETs induce rapid shape conversion from stellate to flat. Moore et al. (1989) have reported that denudation of the villus tip by epithelial injury induced condensation of microfilaments in the cytoplasmic processes of subepithelial fibroblasts and contraction of the subepithelial fibroblast network, although microfilaments were few in controls (Moore

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Figure 4.16 Schematic modelsdescribinghowsubepithelial fibroblastswork inthe intestinal villi. (A) Barrier/sieve functions: subepithelial fibroblasts work as a barrier or a sieve for nutrients, ions, H2O, and immune cells by changing their morphology and releasing cytokines that affect the permeability of epithelia. (B) Regulation of villous movements: subepithelial fibroblasts work as a mechanical frame that determines the mechanical properties of the villi. Passive and active movements of villi may be regulated by cell shape changes and cell contractions. (C) Mechanosensor: subepithelial fibroblasts release ATP by mechanical stresstovilli. Released ATPactivates P2X2,3 onterminals insensory neurons.

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et al., 1989). This means repolymerization of actin and cell shape conversion from stellate to flat shape. It seems that direct contact between subepithelial fibroblasts and epithelial cells may be essential to maintain stellate features. Thus, the network of subepithelial fibroblasts can locally and dynamically change its mechanical properties. In addition to mechanical property changes by cell shape conversion, subepithelial fibroblasts contract when exposed to ETs and ATP. Contractions elicited by ATP are transient, which continue for a few tenths of a second, but those by ETs continue for a longer time and are sometimes oscillatory. Though the contraction by ATP was only transient, contractions evoked by ETs were somewhat persistent. In collagen gels containing colonic myofibroblasts, a plateau of contractile tension evoked by ET1 persists over 15 min (Kernochan et al., 2002). Besides smooth muscle contractions, these contractions of subepithelial fibroblasts may contribute on villus movements (see Fig. 4.16B). Intestinal villi are not simple amplifiers of the mucosal surfaces that absorb nutrient, but rather they give rise to graceful surface properties that resolve contradictory functional demands such as smooth transfers and the long-term folding of food. Dynamic networks of subepithelial fibroblasts achieve these extraordinary mechanical properties of intestinal villi.

6.3. Mechanosensors in the villi The ingestion of food and water gives rise to chemical and mechanical signals that induce villous motilities and peristaltic movements in the gut. Chemical and mechanical signals control motility, secretion, blood flow, and immunity in local and/or central neural and/or nonneural pathways (Buchan, 1999; Furness et al., 1999, 2004; Ho¨fer et al., 1999). In the mucosa, luminal stimuli release sensory mediators from the mucosal epithelium, which then activate the nerve terminals of sensory neurons (Buchan, 1999; Burnstock, 2001; Cooke et al., 2003). At present, it is thought that serotonin and ATP are mechanomodulators, and that enterochromaffin, epithelial, or neural cells in the gut wall are mechanosensitive cells that detect change in tension from touch, stretch, pressure, and shear stress (Bertrand, 2003; Buchan, 1999; Burnstock, 2001; Cooke et al., 2003). 6.3.1. Mechanosensory signals from the epithelium Serotonin in enterochromaffin cells Serotonin (5-HT) is a chemomediator and mechanomediator that is released from enteroendocrine cells in the epithelial layer by luminal stimuli (Eastwood et al., 1998; Kirchgessner et al., 1992; Wade et al., 1996). In luminal perfusions to rat duodenum by high intracellular pressures, the luminal release of serotonin correlates with the intensity of the luminal pressure and elicits peristaltic contractions (Bu¨lbring and Crema, 1959; Fujimiya et al., 1997). Recently, by the use

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of electrochemical techniques with carbon fiber electrodes, the release of serotonin was measured in real time (Bertrand, 2004, 2006). Pressure and stretch-evoked reflexes and 5-HT release from enterochromaffin cells strongly correlate with local motor reflexes in the smooth muscles (Bertrand, 2006). Human carcinoid BON cells have 5-HT immunoreactivity associated with granules and are assumed to be a model of enterochromaffin cells (Kim et al., 2001). The mechanical stimulations to BON cells directly or indirectly activate Gaq signaling pathways, mobilize intracellular calcium, and cause the release of 5-HT (Kim et al., 2001). ATP in enterochromaffin and epithelial cells ATP is also reported to activate nerve terminals of intrinsic sensory neurons via P2X2 and/or P2X3 (Bertrand and Bornstein, 2002; Bian et al., 2003; Castelucci et al., 2002; Poole et al., 2002; Ren et al., 2003), or P2Y1 (Cooke et al., 2004). In P2X2 or P2X3 knockout mice, the intraluminal pressure-induced peristalsis was inhibited (Bian et al., 2003; Ren et al., 2003). In a rat model of colitis, ATP release and mechanosensory transduction are increased (Wynn et al., 2004). In endocrine cells and neurons, a high concentration of ATP is stored in the secretory granules (Winkler and Westhead, 1980) and clear or dense-cored vesicles (White and Al-Humayyd, 1983). ATP is thought to colocalize with serotonin in secretory granules of enterochromaffin cells, and is coreleased, just as seen in adrenal chromaffin cells (Winkler and Westhead, 1980). ATP and nucleotides are ubiquitously distributed in the cells and tissues. ATP is also released from epithelial cells by mechanical stimuli (Grygorczyk and Hanrahan, 1997), by hypotonic stimuli (Dezaki et al., 2000; Hazama et al., 1999; Hisadome et al., 2002; Okada et al., 2001; Sabirov et al., 2001; van der Wijk et al., 1999, 2003), or by infection of enteropathogenic Escherichia coli (Crane et al., 2002, 2005). However, these data were obtained only from intestinal cell lines. Measurements of ATP release from intestinal cells in vivo are still to come.

6.3.2. Subepithelial fibroblasts as a mechanosensor in the villi Subepithelial fibroblasts are located just under the epithelium, and their processes sometimes closely contact with nerve endings. In addition, subepithelial fibroblasts are highly sensitive to mechanical stress and subsequently release ATP. Their anatomical location and inherent characteristics strongly suggest that subepithelial fibroblasts are a mechanosensor in the villi (Fig. 4.16C). The finding that a mechanically induced Ca2þ wave propagated through the subepithelial fibroblast network could activate neural cells cocultured with subepithelial fibroblasts (see Fig. 4.14B) supports the aforementioned idea. Thus, we propose that subepithelial fibroblasts release ATP upon stretching or distention of villi and that the released ATP acts on P2X2 and/or P2X3 receptors on intrinsic sensory enteric neurons to induce mechanotransduction (see Fig. 4.16C).

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The mechanosensitivity of subepithelial fibroblasts changes drastically with cell shape, and cell shape changes locally and temporally in the villi. Cell shapes of subepithelial fibroblasts change along the crypt–villus axis in intestinal villi. This suggests that mechanosensitivity changes along the crypt–villus axis and is higher at the lower part of the villi. Cell shapes also change temporally by ET that is released from several kinds of cells in villi. So the mechanosensitivity of the villi changes locally and dynamically depending on cellular conditions, which may be essential to achieve graceful villous functions.

6.4. Other signal transduction in the villi 6.4.1. Interactions with smooth muscle Subepithelial fibroblasts communicate with lamina propria fibroblasts, which embrace smooth muscles in the villus core in vivo (Gu¨ldner et al., 1972). By calcein-loaded FRAP analysis in culture, stellate cells abut smooth muscles in the center of the inoculated villi (probably corresponding to lamina propria fibroblasts in vivo) and communicate with subepithelial fibroblasts that migrate to the periphery via gap junctions; however, they do not communicate with smooth muscles (K. Furuya and S. Furuya, unpublished data). Smooth muscle contractions are elicited by touch stimulation to a subepithelial fibroblast distant from smooth muscle (K. Furuya and S. Furuya, unpublished data). Ca2þ waves elicited by mechanical stimuli to subepithelial fibroblasts propagated to smooth muscles, because smooth muscles possess purinergic P2Y receptors (Giaroni et al., 2002; Matsuo et al., 1997). 6.4.2. Interactions with microvasculature The enteric circulation is mediated by neural mediators, humoral mediators, paracrine and autocrine mediators, and metabolites (Matheson et al., 2000; Vanner and Surprenant, 1996). Distention, mechanical stimulation, and the uptake of nutrients stimulate secretomotor and vasodilator reflexes, involving intrinsic and extrinsic vasodilatory neurons in mucosal reflexes (Vanner and Surprenant, 1996). Blood flow increases during the absorption of nutrients (Sidery and Macdonald, 1994), and absorption of fluid takes place principally in the upper third region of the villi (Kinter and Wilson, 1965). In addition to neural regulation, the microcirculation is modified by adjacent myofibroblasts in each tissue such as pericytes (Kawamura et al., 2003; Peppiatt et al., 2006), astrocytes (Filosa et al., 2004; Koehler et al., 2006; Metea and Newman, 2006; Mulligan and Mac Vicar, 2004; Takano et al., 2006), hepatic stellate cells (Reynaert et al., 2002; Thimgan and Yee, 1999), and subepithelial fibroblasts. As the cellular network of subepithelial fibroblasts overlays the villous vascular tree, Ca2þ waves evoked by mechanical stimulation may propagate from subepithelial fibroblasts to capillaries, which express P2X1 receptors (Gro¨schel-Stewart et al., 1999).

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Moreover, contractions and cell shape conversions of subepithelial fibroblasts may regulate the absorption of nutrients or regulate blood flow. As subepithelial fibroblasts of the lower area of villi are flat and rich in filamentous a-SMA, contraction evoked by mechanical stimuli seems to propagate longer distances than in the upper portion of the villi. Cell shape conversions and persistent or oscillatory contractions of subepithelial fibroblasts are evoked by ETs, which are secreted from nerve terminals, immune cells, or blood vessels. These processes seem to regulate the microcirculation locally and dynamically in the villi. In addition to vasoconstriction, there is a possibility that subepithelial fibroblasts secrete vasodilators just as reported in astrocytes in the brain (Metea and Newman, 2006; Takano et al., 2006). 6.4.3. Interaction with immune cells In the lamina propria, various hematopoietic cells are present such as lymphocytes, macrophages, eosinophils, plasma cells, basophiles, monocytes, mast cells, and dendritic cells (Deane, 1964; Hashimoto and Komuro, 1988; Hunyady et al., 2000; Komuro, 1985; Toyoda et al., 1997). In the small intestine, there is close contact (about 20 nm) between absorptive epithelial cells and lymphocytes, eosinophils, or macrophages (Hashimoto and Komuro, 1988). Subepithelial fibroblasts also closely contact lymphocytes, and dendritic cells, which are professional antigenpresenting cells (Toyoda et al., 1997). Subepithelial fibroblasts of normal colon constitutively express MHC class II molecules, and isolated colonic myofibroblasts in culture are able to stimulate allogeneic CD4þ T cell proliferation (Saada et al., 2006). The aforementioned data indicate that colonic subepithelial fibroblasts are nonprofessional antigen-presenting cells and play critical roles in mediating tolerance to luminal antigens (Saada et al., 2004). Immune cells such as lymphocytes, macrophages, and mast cells in the lamina propria secrete a variety of cytokines. IL-1 stimulates production of prostaglandin E2 via increases in the expression of cyclooxygenases COX-1 and COX-2, in 18Co colonic myofibroblasts, which regulate epithelial cell proliferation, migration, and secretory responses (Hinterleitner et al., 1996; Mahida et al., 1997; Mifflin et al., 2002; Shao et al., 2006; Valentich et al., 1997). Moreover, colonic myofibroblasts activated by ILs, LPS, TNF, and PMA secrete a wide spectrum of cytokines (IL-6, IL-8, and granulocytemacrophage–CSF) (Rogler et al., 2001) and matrix-degrading molecules (Andoh et al., 2005). In particular, crosstalk between subepithelial fibroblasts and mast cells by the secretion of chemokines, cytokines, and growth and differentiation factors is reciprocally important in proliferation, differentiation, and activation. SCF, TGF-b, and NGF are secreted from subepithelial fibroblasts, and PDGF and SCF, TNF-a, TGF-b, and ILs (IL-1, IL-6, and IL-8) are from mast cells (Crivellato et al., 2005). The number of pericryptal fibroblasts

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correlates with the density of mast cells in the duodenal crypt, which implicates them in the homeostasis of the villous structure (Crivellato et al., 2006). Reciprocal interactions between subepithelial fibroblasts and immune cells play important roles not only in the process of inflammation but also in the homeostasis of the villus structure and function. 6.4.4. Source and functions of endothelins in the villi ET-1 is a potent vasoconstrictor peptide that was originally isolated from the supernatant of cultured porcine endothelial cells (Yanagisawa et al., 1988). Moreover, members of the endothelin family (ET-1, ET-2, ET-3) exert a wide variety of physiological functions such as neuromodulator-like activity in the central nervous system (Kuwaki et al., 1997), increased proliferation in a wide range of cells (Inoue et al., 1989; Sasaki et al., 1998), inhibition of gap junction permeability, and enhancement of Ca2þ signaling in astrocytes (Blomstrand and Giaume, 2006; Blomstrand et al., 1999; Giaume et al., 1992). ETs elicit migration, but not proliferation, in the cultured colonic myofibroblasts (Kernochan et al., 2002). In the primary culture of subepithelial fibroblasts isolated from intestinal villi, ETs induced cell shape changes by decreasing the intracellular cAMP level, transient and oscillatory intracellular Ca2þ responses, and cellular contractions but do not affect gap junction permeability (Furuya and Furuya, 1993; K. Furuya et al., 1994, 2005; S. Furuya et al., 2005). ETs and, ETA and ETB receptors are abundantly distributed in the gut. Proliferation and differentiation of enteric neurons are regulated by ET-3 through ETB receptors during embryonic development (Baynash et al., 1994; Hosoda et al., 1994). In adult small intestine and colon, immunoreactivity of ET-1 and mRNA of prepro-ETs and endothelin-converting enzyme (ECE) were detected in ganglion cells of submucosal and myenteric plexus, mast cells, neutrophils, macrophages, endothelial cells, and epithelial cells (Egidy et al., 2000a; Escrig et al., 1992; Inagaki et al., 1991; Liu et al., 1998; Massai et al., 2003; Takizawa et al., 2005). ET-2 protein and mRNA are mainly distributed in epithelial cells, and weakly in VIP neurons in the myenteric plexus (Takizawa et al., 2005). In cultured enteric neurons that contain VIP, ET-1 is released into the culture medium, independent of depolarization (Eaker et al., 1995). ET-1 biosynthesis is stimulated in endothelial cells by angiotensin II and vasopressin (Imai et al., 1992), and in cultured epithelial cells by IL-2 treatment (Shigematsu et al., 1998). The immunoreactivities or mRNA of ETs and ET receptors are increased in colon tumors, with an especially high increase of ETB receptors (Edigy et al., 2000b). In inflammation, increases of ET-1–positive stroma cells and high prepro-ET-1 mRNA expression are observed in the lamina propria of patients with ulcerative colitis and Crohn’s disease (Murch et al., 1992) and appendicitis (Massai et al., 2003). Thus, ET functions are thought to be intensely related to inflammation and malignancy, especially in the colon and appendix. However, even under

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normal physiological conditions in the colon and small intestine, ETs will be released from endothelial cells in response to vasoactive substances, epithelial cells, neurons, and immune cells in response to conventional antigens and bacteria. Upon exposure to ETs, subepithelial fibroblasts may locally and dynamically change cell shape, mechanosensitivity, and contractile ability in vivo.

7. Concluding Remarks Subepithelial fibroblasts form a contractile cellular network under the epithelium in the intestinal villi and respond to many kinds of vasoactive and neuroactive substances, including ATP and endothelins (ETs). Subepithelial fibroblast networks closely contact microvascular networks, sensory and motor neuronal networks, smooth muscles, and epithelia. The first remarkable property of subepithelial fibroblasts is their fast and reverse shape conversions that are dependent on cAMP levels, although gap junctions in the cellular network are kept open independently of cell shape. Cell shape conversions lead to alterations in the size of fenestra and mechanical properties of the network. They also contract by ATP and ETs and contribute to graceful villous motility. The second remarkable property of subepithelial fibroblasts is their high sensitivity to mechanical stress and the resulting ATP release. Liberated ATP works as autocrine and paracrine mediators. Subepithelial fibroblasts communicate among other cell systems in the villi via released ATP and other humoral factors. By these unique characteristics, subepithelial fibroblast networks serve as (1) a barrier/sieve, (2) a flexible mechanical frame, (3) a mechanosensor, and (4) a signal transduction machinery in the intestinal villi. These functions are likely regulated locally and dynamically in the villi by rapid cell shape changes, contraction, and cell shape–dependent mechanosensitivities, which may play essential roles in the intestine.

REFERENCES Abbracchio, M. P., and Williams, M. (2001). ‘‘Purinergic and Pyrimidinergic Signalling. Handbook of Experimental Pharmacology,’’ Vol. 151/I, II. Springer-Verlag, Berlin. Adegboyega, P. A., Mifflin, R. C., DiMari, J. F., Saada, J. I., and Powell, D. W. (2002). Immunohistochemical study of myofibroblasts in normal colonic mucosa, hyperplastic polyps, and adenomatous colorectal polyps. Arch. Pathol. Lab. Med. 126, 829–836. Andoh, A., Zhang, Z., Inatomi, O., Fujino, S., Deguchi, Y., Araki, Y., Tsujikawa, T., Kitoh, K., Kim-Mitsuyama, S., Takayanagi, A., Shimizu, N., and Fujiyama, Y. (2005). Interleukin-22, a member of the IL-10 subfamily, induces inflammatory responses in colonic subepithelial myofibroblasts. Gastroenterology 129, 969–984.

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Syndrome of Aluminum Toxicity and Diversity of Aluminum Resistance in Higher Plants Jian Feng Ma

Contents 1. Introduction 2. Syndrome of Aluminum Toxicity 2.1. Aluminum-inhibited root cell elongation and cell division 2.2. Mechanisms of aluminum toxicity 3. Aluminum Resistance 3.1. Exclusion mechanisms 3.2. Internal detoxification of aluminum with organic acids 4. Beneficial Effect of Aluminum on Plant Growth 5. Concluding Remarks Acknowledgments References

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Abstract Aluminum (Al) is the most abundant metal in the earth’s crust, while its soluble ionic form (Al3þ) shows phytotoxicity, which is characterized by a rapid inhibition of root elongation. Aluminum targets multiple cellular sites by binding, resulting in disrupted structure and/or functions of the cell wall, plasma membrane, signal transduction pathway, and Ca homeostasis. On the other hand, some plant species have evolved mechanisms to cope with Al toxicity both externally and internally. The well-documented mechanisms for external detoxification of Al include the release of organic acid anions from roots and alkalination of the rhizosphere. Genes encoding transporters for Al-induced secretion of organic acid anions have been identified and characterized. Recent studies show that ABC transporters are involved in Al resistance. The internal detoxification of Al in Al-accumulating plants is achieved by the formation of nontoxic Al

Research Institute for Bioresources, Okayama University, Kurashiki 710–0046, Japan International Review of Cytology, Volume 264 ISSN 0074-7696, DOI: 10.1016/S0074-7696(07)64005-4

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2007 Elsevier Inc. All rights reserved.

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complexes with organic acids or other chelators and sequestration of these complexes in the vacuoles. In some plant species, Al shows beneficial effects on plant growth under particular conditions, although the exact mechanisms for these effects are unknown. Key Words: Al toxicity, Al resistance, Beneficial effect, External detoxification, Internal detoxification, Organic acid anions, Transporter. ß 2007 Elsevier Inc.

1. Introduction Aluminum (Al) is a light metal of atomic number 13 and is widely used in industry and in our daily life. Aluminum composes 7.5 to 8.1% of the Earth’s crust, making it the most abundant metallic element and the third most abundant element after oxygen and silicon. It occurs primarily in aluminosilicate minerals, most commonly as feldspars in metamorphic and igneous rocks, and as clay minerals in well-weathered soils (Driscoll and Schecher, 1988). Aluminum has two isotopes in nature: a stable isotope 27Al and a radioisotope 26Al. 27Al has a natural abundance of 100%, while 26Al is rarely used as a tracer in biological studies due to the high production cost. The solubility of Al is very low in the neutral pH range (6.0 to 8.0); however, the solubility is enhanced under acidic (pH < 6.0) or alkaline (pH > 8.0) conditions. Aluminum is nontoxic as a metal; however, it becomes toxic to all living cells when it is present in an ionic form (either Al3þ or Al[OH]4 ). For example, ionic Al is a neurotoxin and may be a cause of Alzheimer’s disease, although it is not certain whether an accumulation of Al in the brain is a consequence or a cause of Alzheimer’s disease (Crapper et al., 1973). Ionic Al also rapidly inhibits root elongation and functions as detailed below, consequently resulting in poor plant water and nutrient uptake and increased sensitivity to various stresses, especially drought stress (Ma, 2005b). Therefore, Al toxicity has been recognized as a major factor limiting crop production in acid soils, which comprise approximately 30 to 40% of the world’s arable soils and up to 70% of potentially arable land (von Uexku¨ll and Mutert, 1995). Knowledge of the mechanisms of Al toxicity and resistance has been accumulated during the past decades, and a number of excellent reviews on these topics have been published (Barcelo and Poschenrieder, 2002; Kochian et al., 2005; Ma, 2005b; Ma et al., 2001; Rengel, 2004; Ryan et al., 2001). The present review highlights recent progress made over the past 5 years on the mechanisms of Al toxicity and resistance in higher plants.

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2. Syndrome of Aluminum Toxicity 2.1. Aluminum-inhibited root cell elongation and cell division The characteristic symptom of Al toxicity is the inhibition of root elongation. Most Al is accumulated in the root tips (Fig. 5.1A). Recent studies showed that the inhibition occurs as early as 30 to 120 min after exposure to Al (Barcelo and Poschenrieder, 2002; Doncheva et al., 2005). Root elongation consists of cell elongation and cell division. The rapid Al-induced inhibition of root elongation suggests that the initial effect is likely to be caused by the inhibition of cell elongation rather than of cell division. However, a recent study showed that Al also inhibits cell division in the proximal meristem (250 to 800 mm from the root tip) within 5 min after Al exposure in an Al-sensitive cultivar of maize (Doncheva et al., 2005). Although cell division per se does not increase root length, both the rate of cell division and the time period that mitotic cells remain active determine the supply of cells to the elongation zone and hence the elongation rate. Therefore, Al-inhibited cell division may affect root elongation at a later stage.

2.2. Mechanisms of aluminum toxicity A number of possible mechanisms responsible for Al-induced inhibition of root elongation have been proposed, as detailed below. However, the exact mechanism by which Al initially causes the inhibition of root elongation has not been determined. Aluminum may interact with the root cell wall, disrupt the plasma membrane, and inhibit transport processes on the plasma membrane. It may inhibit enzyme activity and DNA replication, disrupt signal transduction pathways, inhibit the formation of microtubules, and cause dysfunction of mitochondria. Aluminum may also interact with Ca homeostasis within the root cell and other symplasmic constituents such as calmodulin (Ma, 2005b). Mechanisms involved in Al toxicity may vary with Al concentrations and exposure time (Ma et al., 2004b). In roots exposed to a low concentration of Al for a short time, only the apoplasm of the roots such as the cell wall may be influenced by Al. However, in roots exposed to a high concentration of Al (>mM) for a long time, the plasma membrane, DNA, and enzymes may also be affected. It should be noted that the Al-induced inhibition of root elongation occurs within minutes at very low concentrations (micromolar). Moreover, Al concentration in acid soil solutions rarely

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Figure 5.1 Al accumulation and Al-induced physiological changes in barley roots exposed to 5 mM Al for different times. (A) Localization of Al accumulation in the root tip (stained with eriochrome cyanine); (B) cellular location of Al accumulation (stained by Morin); (C) callose formation (stained by aniline blue); (D) integrity of the plasma membrane (stained by FDA-PI); (E) production of reactive oxygen species (stained by 6-carboxy-20,70 -dichlorodihydrofluorescein diacetate, di(acetoxymethyl ester) 2 mm from the root tip. Bar ¼ 100 mm. (Photo by Naoki Yamaji.)

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exceeds 140 mM (Ma et al., 2004b). Some previous results derived from laboratory experiments on the plants grown in the presence of Al at millimolar concentrations may not be applied to plants under field conditions. 2.2.1. Aluminum interaction with cell wall When the roots are exposed to Al, the cell wall is the first site in contact with Al. Previous studies have shown consistently that most Al is bound to the cell wall. For example, Clarkson (1967) found that 85 to 90% of the total Al accumulated by barley roots was tightly bound to cell walls. In giant algal cells of Chara corallina, up to 99.9% of the total cellular Al accumulated in the cell wall (Rengel and Reid, 1997). Almost 90% of the total Al was associated with the cell wall in cultured tobacco cells (Chang et al., 1999). Using hypocotyls of okra as an experimental model, Ma et al. (1999) found that 95% of the total Al was associated with the cell wall of the epidermis. Furthermore, most Al was localized on the epidermis and the outer cortex (Fig. 5.1B; Jones et al., 2006; Ma et al., 2004b). Recent studies have shown that Al bound on the cell wall affects the properties of the cell wall. Exposure to Al for 6 h decreased the extensibility of root cell walls in an Al-sensitive cultivar of wheat, Scout 66, in a study using the tensile test method (Tabuchi and Matsumoto, 2001). Aluminum also caused a rapid decrease in elastic and viscous extensibility and the whole extensibility of the cell wall of the root apices of the same cultivar in a study using a creepextension analysis (Ma et al., 2004b). The cell elongation is accompanied by an increase in cell wall extensibility that is regulated by two physical parameters (i.e., the viscosity coefficient and elasticity modulus). Therefore, Al-decreased extensibility may cause the inhibition of root elongation. Furthermore, the cell wall was more easily broken in Al-treated roots (Ma et al., 2004b), suggesting that Al makes root cell walls brittle. Aluminum also causes changes in cell wall polysaccharides. In wheat roots, exposure to Al resulted in an increase of pectin and hemicellulose but not cellulose (Eticha et al., 2005; Hossain et al., 2006; Tabuchi and Matsumoto, 2001). Aluminum also increases wall-bound ferulic and diferulic acids. These changes may lead to an increase in cell wall rigidity and therefore decrease cell wall extensibility as described above. However, it is unknown whether the Al-induced increase of pectin and hemicellulose is the result or the cause of Al-induced inhibition of root elongation. Several studies have suggested that Al is bound to pectin. Pectin is mainly composed of galacturonic acid chains, which contain negatively charged binding sites of Al. There is evidence that pectin content is positively correlated with the Al-induced loss of cell viability in maize suspension cells (Schmohl and Horst, 2000), suggesting that binding of Al to the pectin matrix is an important step in the expression of Al toxicity.

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Cell elongation includes both a loosening process and cell wall synthesis. Al-induced rapid inhibition of cell elongation suggests that the loosening process of the cell wall rather than cell wall synthesis is affected by Al. During the loosening process of the cell wall, the matrix polysaccharides undergo an enzymatic breaking of bonds and/or enzymatic deformation within the wall framework, generating more binding sites of Al. Wall loosening and continued deposition of new materials into the wall are tightly integrated events. It is therefore suggested that tight binding of Al to the newly generated sites of the cell wall hampers the binding of newly synthesized cell wall material to newly generated sites of the cell wall, which are necessary for the reorganization of the cell wall during cell elongation (Ma et al., 2004b). Such an interruption of cell wall deformation may result in a decrease in both elastic and plastic extensibility and break load of the cell wall as described above. 2.2.2. Aluminum-induced callose formation One of the most rapid physiological responses to Al is the increased deposition of callose, which occurs within minutes after exposure to Al (Fig. 5.1C) ( Jones et al., 2006). Therefore, callose deposition in plants has been used as a marker for Al toxicity in many studies (Teraoka et al., 2002). Callose deposition is observed in the epidermal layers surrounding the root apex ( Jones et al., 2006). Furthermore, callose production occurs only in the actively growing root regions. Callose could cement the cell walls together, preventing cell wall loosening, blocking plasmodesmata and inhibiting the symplasmic transfer of solutes (Sivaguru et al., 2000). However, when callose production was prevented by the presence of a lipophilic antioxidant, butylated hydroxyanisole, the Al-induced inhibition of root elongation was not alleviated (Yamamoto et al., 2001). Therefore, its seems that callose production does not represent a primary mechanism of Al-induced inhibition of cell elongation ( Jones et al., 2006). Callose is synthesized by the membrane-bound, 1,3-b-D-glucan synthase that uses UDP-glucose as a substrate. This enzyme is activated by various stresses such as heavy metals and wound, as well as Al. However, Al is a powerful inhibitor of this enzyme in vitro (Bhuja et al., 2004). The Al-induced increase of Ca concentration in the cytosol may be a trigger for enhanced synthesis of callose in vivo. However, a recent study reported that an increase of Ca concentration in cytoplasm is not the only factor modulating the increase in callose synthesis and deposition in the presence of Al because a significant increase of Ca concentration by the Ca ionophore, A23187, only caused a small increase in callose deposition (Bhuja et al., 2004). 2.2.3. Aluminum-disturbed plasma membrane The plasma membrane is negatively charged and contains phospholipids. Therefore, Al binds to these sites in the plasma membrane and replaces other cations such as Ca, resulting in a disruption of the structure and function of

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the membrane (Fig. 5.1D). Aluminum alters the membrane surface potential (Kinraide, 2001), causes rigidity and lipid peroxidation of the plasma membrane, and blocks Ca and K channels and other transporters (Pineros and Kochian, 2001). A short exposure to Al rapidly causes lipid peroxidation as described below. Aluminum also inhibits Hþ-ATPase in the plasma membrane (Ahn et al., 2002), resulting in disruption of the proton gradient. Since the transmembrane proton gradient serves as the major driving force for secondary ion transport processes, Al-induced disruption of the proton gradient could indirectly alter the ionic status and ion homeostasis of root cells (Kochian et al., 2005). 2.2.4. Aluminum-disrupted cytoskeleton The effect of Al on the cytoskeleton has been reported in a number of studies. Aluminum appears to affect both microtubule and actin filament cytoskeleton (Frantzios et al., 2005), which are closely related to root cell elongation. Aluminum induces microtubule depolymerization in the root elongation zone (Sivaguru et al., 2003) and rapid disintegration of cortical microtubules in the transition zone (Horst et al., 1999). A recent study showed that a short exposure to Al resulted in the formation of additional bundles of cortical microtubules, whereas the thickness of the individual bundles decreased in tobacco cell lines (Schwarzerova et al., 2002). Prolonged exposure caused disorientation of cortical microtubules. As these changes preceded the decrease of cell viability by several hours, the microtubular cytoskeleton has been suggested as one of the early targets of Al toxicity. Aluminum also induces a significant increase in the number of actin filaments, its bundling, and its disorganization in interphase root tip cells of Triticum turgidum (Frantzios et al., 2005). These cytoskeletal responses to Al could be triggered indirectly through a continuum between the cell wall, plasma membrane, and cytoskeleton in consequence of the primary Al response in the cell wall and plasma membrane as described above (Ahad and Nick, 2007). Aluminum entering into the cytoplasm might inhibit the GTPase or ATPase functions of tubulin and actin (Frantzios et al., 2005). It also might affect the cytoskeleton through an Al-induced disturbance of Ca2þ homeostasis, as described below, and other events. 2.2.5. Aluminum-induced oxidative stress Aluminum induces peroxidation of lipids in pea roots, but it seems that this event is not the primary cause of Al-induced inhibition of root elongation (Yamamoto et al., 2001) because this was not alleviated when lipid peroxidation was reduced in the presence of a lipophilic antioxidant, butylated hydroxyanisole. Overexpression of mitochondrial manganese superoxide dismutase, an important enzyme of the antioxidant pathway, resulted in enhanced Al resistance in canola (Basu et al., 2001). Overexpression of other antioxidation enzymes such as glutathione S-transferase and peroxidase

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from tobacco also resulted in enhanced Al resistance in Arabidopsis (Ezaki et al., 2001). These findings suggest that Al-induced damage during a prolonged period could be partially alleviated by increased antioxidation enzymes. Aluminum also induces an instantaneous and sustained production of reactive oxygen species (ROS) (Fig. 5.1E) (Darko et al., 2004; Jones et al., 2006). Production of ROS might be caused by Al binding to the plasma membrane and an Al-induced increase of cytoplasm Ca. 2.2.6. Aluminum interaction with signal transduction pathways Aluminum was reported to inhibit the inositol 1,4,5-trisphosphate signal transduction pathway in wheat ( Jones and Kochian, 1995; Ramos-Diaz et al., 2007). A short exposure (1 h) to Al decreased phospholipase C (PLC) activity in wheat roots but did not affect the activity of other enzymes ( Jones and Kochian, 1995). Phospholipase C hydrolyzes phosphatidylinositol 4,5-bisphosphate (PIP2) to produce inositol 1,4,5-triphosphate (Ins [1,4,5]P3) into the cytoplasm and diacylglycerol (DAG) into the membrane. Al-induced inhibition of Ins(1,4,5)P3 formation may lead to altered spatial and temporal dynamics of cytoplasmic Ca transients, leading to subsequent breakdown in signal transduction events. In Coffea arabica suspension cells, Al also inhibits the formation of phosphatidic acid (PA), which has been implicated in various stress-signaling pathways in plants (Ramos-Diaz et al., 2007). As PA is produced from DAG by phosphorylation, Al-induced inhibition of PA formation is a result of PLC inhibition (Ramos-Diaz et al., 2007). Nitric oxide (NO) is an important signaling molecule modulating numerous physiological processes in plants. Recently, it was reported that Al inhibits the activity of nitric oxide synthase (NOS) and reduces endogenous NO concentrations in Hibiscus moscheutos (Tian et al., 2007). There is a good correlation between the Al-induced inhibition of root elongation and Al-reduced endogenous concentrations in this species. Furthermore, Alinduced reduction of endogenous NO concentration in the root apical cells occurred earlier than the Al-induced inhibition of root elongation, suggesting that the reduced endogenous NO concentration is a cause rather than a consequence of Al toxicity (Tian et al., 2007). 2.2.7. Aluminum-disrupted Ca homeostasis in cytoplasm There are numerous studies on the interaction between Al and Ca in plants (Rengel and Zhang, 2003). Although Al inhibits the hyperpolarizationactivated Ca2þ channel, Al rapidly causes an increase in cytosolic Ca2þ activity. This increase is partly from extracellular sources including the Al-resistant portion of the flux through depolarization-activated Ca2þ channels and fluxes through Ca2þ-permeable nonselective cation channels in the plasma membrane. The increase may also come from intracellular

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sources including enhanced activity of Ca2þ release channels in the tonoplast and the endoplasmic reticulum membrane. Cytosolic Ca is an important second messenger. Disruption of cytoplasmic Ca2þ homeostasis may disturb numerous biochemical and physiological processes, including those involved in root growth. For a more detailed description on Al and Ca interaction, refer to Rengel and Zhang (2003). 2.2.8. Aluminum-inhibited Mg uptake Previous physiological studies have shown that Al inhibits Mg uptake by intact roots and that application of Mg can alleviate Al toxicity in several plant species (Yang et al., 2007). The activity of a magnesium transporter from Arabidopsis (AtMGT1) was inhibited by relatively low concentrations of Al (Li et al., 2001). Magnesium is an important part of chlorophyll and serves as a cofactor with ATP in a number of enzymatic reactions (e.g., ATPases and RNA polymerases). Therefore, Al-induced inhibition of Mg uptake will cause Mg deficiency, resulting in inhibition of plant growth. This is supported by the finding that overexpression of an Arabidopsis magnesium transporter gene (AtMGT1) alleviates Al toxicity in tobacco (Deng et al., 2006). 2.2.9. Aluminum-inhibited auxin polar transport Auxin is required for cell elongation. It is transported from auxinsynthesizing shoot tissues via the phloem toward the root apical meristems, where it is unloaded from the central stele into cortical and epidermal cells and then translocated basipetally to the elongation zone (Estelle, 1998). Aluminum inhibits polar transport of auxin in maize roots (Kollmeier et al., 2000). However, the mechanism responsible for Al-induced inhibition of auxin polar transport is not understood. A recent study suggested that Al-induced inhibition of auxin transport is also involved in the Al-induced alteration of root cell patterning (Doncheva et al., 2005).

3. Aluminum Resistance Some plant species and cultivars have evolved mechanisms for detoxifying Al. Two strategies for the detoxification of Al by plants have been suggested. One is the exclusion of Al from the root tips (exclusion mechanism) and the other is tolerance to Al that enters the plant (internal tolerance mechanism).

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3.1. Exclusion mechanisms A number of physiological studies have presented several mechanisms for detoxifying Al externally, including release of chelating ligands or phosphate, increased pH in the rhizosphere, modified cell wall, redistribution of Al, and internalization of Al (Fig. 5.2). 3.1.1. Aluminum-induced secretion of organic acid anions A brief overview of the physiology of aluminum-induced organic acid anions The best documented mechanism of Al exclusion is the secretion of organic acid anions from roots (Fig. 5.2A). Since the first report on Alinduced malate secretion in wheat (Kitagawa et al., 1986), a wide range of plant species has been reported to secrete organic acid anions in response to Al, including dicots and monocots such as wheat, maize, rye, and soybean. Physiological studies have been extensively carried out to determine the nature of Al-induced secretion of organic acid anions (Kochian et al., 2005; Ma, 2005a; Ma et al., 2001; Ryan et al., 2001). Plants differ in the species of organic acid anions secreted, temporal secretion patterns, temperature sensitivity, and dosage responses to Al (Ma and Furukawa, 2003). Up to now, citrate, oxalate, and/or malate have been identified as the organic acid anions secreted by roots in response to Al. In some plant species, two Shoot A

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Figure 5.2 Mechanisms of external Al detoxification. (A) Secretion of Al-chelating substances from the roots including organic acid anions and phenolic compounds; (B) alkalination of the rhizosphere; (C) increased methylation of pectin; (D) redistribution of Al; (E) efflux of Al.

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organic acid anions are secreted in response to Al. All three organic acid anions can chelate Al, thereby preventing the binding of Al to cellular components, although their chelating ability differs, following the order citrate > oxalate > malate. The site of secretion is localized in the root apex and the secretion is highly specific to Al. Two patterns of organic acid anion release can be identified on the basis of the timing of secretion (Ma, 2005a). In Pattern-I plants, secretion occurs almost immediately following the addition of Al, suggesting that Al activates a preexisting anion channel in the plasma membrane and that the induction of genes is not required (Ma, 2005a). Recently, several studies on the process leading to the secretion of organic acid anions have been reported. Patch-clamp studies have shown that Al triggers the opening of citrate and malate channels in the plasma membrane, which facilitates the efflux of malate and citrate (Kollmeier et al., 2001; Pineros and Kochian, 2001; Zhang et al., 2001). In Pattern-I plants, the possible involvement of protein phosphorylation was suggested in Alinduced secretion of malate in wheat (Osawa and Matsumoto, 2001). In buckwheat, evidence showed that ABA is involved in the secretion of oxalate (Ma et al., 2001). ABA activates the anion channel in stomatal guard cells and may play a similar role in the roots. In contrast, in Pattern-II plants organic acid anion secretion is delayed for several hours after exposure to Al, suggesting that gene induction is required. Some inducible proteins could be involved in organic acid metabolism or in the transport of organic acid anions. However, recent molecular work suggests that the transport system rather than the metabolism of organic acids is more important in controlling organic acid anion secretion as described below. Modulation of plasma membrane Hþ-ATPase has been suggested to be involved in the Al-dependent secretion of citrate. In a mutant carrot cell line showing high citrate secretion, high plasma membrane Hþ-ATPase activity was found (Ohno et al., 2003). In an Al-tolerant cultivar of soybean, which secretes citrate in response to Al, a higher activity of plasma membrane Hþ-ATPase was found in the roots (Shen et al., 2005). Both the expression of the plasma membrane Hþ-ATPase gene and its encoded protein are upregulated by Al. Furthermore, Al activated the threonineoriented phosphorylation of plasma membrane Hþ-ATPase in a dose- and time-dependent manner (Shen et al., 2005). Secretion of citrate may be accompanied by proton exudation for charge balance (Yan et al., 2002). Therefore, Al-activated plasma membrane Hþ-ATPase is probably required for Al-enhanced secretion of citrate. Genes responsible for secretion of organic acid anions Physiological studies have shown that the secretion of organic acid anions is mediated through anion channels or transporters. Two studies with maize revealed that Al activates Cl efflux and the citrate permeable anion channel

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(Kollmeier et al., 2001; Pineros and Kochian, 2001). These studies also indicated that at least a subset of the Al-activated channels requires extracellular Al3þ to maintain channel activity and that the activation machinery is localized to the plasma membrane. Recently, a gene, ALMT1 (Al-activated malate transporter 1), which is responsible for malate release, has been identified in wheat by a subtraction approach between near-isogenic lines of wheat ET8 and ES8 (Sasaki et al., 2004). The gene consists of six exons and five introns (Raman et al., 2005) and is constitutively expressed in the root apices of the Al-resistant line (Sasaki et al., 2004). Introduction of this gene into rice, cultured tobacco cells, and barley resulted in increased Alactivated malate efflux and increased Al resistance for all except rice (Delhaize et al., 2004; Sasaki et al., 2004). Genetic analysis revealed that TaALMT1 colocalized with the Al resistance locus on chromosome 4DL in several populations of wheat (H. X. Ma et al., 2005; Raman et al., 2005). Therefore, TaALMT1 is a major Al resistance locus in wheat. The protein encoded by this gene is localized to the plasma membrane (Yamaguchi et al., 2005), which is predicted to have between six and eight putative transmembrane regions. Heterologous expression of this gene in Xenopus oocytes showed transport activity for malate, but not for citrate. There are two alleles of the TaALMT1 coding region, TaALMT1–1 and TaALMT1–2, in the near-isogenic lines of wheat differing in Al resistance (Sasaki et al., 2004). They differ in six bases and two amino acids. However, heterologous expression in Xenopus oocytes showed no difference in the transport activity of malate. Furthermore, the level of TaALMT1 expression is correlated with overall Al tolerance across 13 wheat cultivars differing in Al resistance. This evidence indicates that Al resistance is not conditioned by these alleles but by the level of TaALMT1 expression (Raman et al., 2005; Sasaki et al., 2004). Recently, analysis of genomic regions upstream and downstream of TaALMT1 in 69 wheat lines suggested that the presence of the sequence repeats upstream of TaALMT1 is related to gene expression and Al resistance in wheat lines of non-Japanese origin (Sasaki et al., 2006). Homologs of TaALMT1 have been cloned from Arabidopsis, rape, and rye (Fontecha et al., 2007; Hoekenga et al., 2006; Ligaba et al., 2006). However, the expression patterns of these genes differ among these plant species. AtALMT1 from Arabidopsis is expressed only in the roots as TaALMT1, but as opposed to TaALMT1, its expression is induced by Al treatment (Hoekenga et al., 2006). Expression of this gene in Xenopus oocytes showed transport activity for malate. This gene is localized on chromosome 1, but interestingly, it does not represent the major Al tolerance QTL, which was previously found in chromosome 1 (Hoekenga et al., 2003, 2006). Two homologs of TaALMT1 have also been cloned from the roots of rape (Brassica napus) (Ligaba et al., 2006). Rape secretes both citrate and malate in response to Al (Ma and Furukawa, 2003). Expression of these two homolog genes (BnALMT1 and BnALMT2) in Xenopus oocytes showed

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transport activity for malate, but not for citrate. BnALMT1 and BnALMT2 are 95% identical to each another, 80% identical to the AtALMT1 gene from Arabidopsis, and 40% identical to the TaALMT1 gene from wheat. Similar to AtALMT1, but different from TaALMT1, the expression of BnALMT1 and BnALMT2 is induced by Al (Ligaba et al., 2006). Therefore, Al is required for two key steps: the induction of transcript and the activation of the proteins to transport malate in rape. Rye also secretes both citrate and malate from the roots in response to Al (Li et al., 2002). A homolog of TaALMT1 in rye (ScALMT1) was recently identified (Fontecha et al., 2007), which is 86% identical to TaALMT1. This gene is mapped to the short arm of chromosome 7R, and the gene is also induced by Al. Interestingly, the expression of this gene is not confined to the root apices. However, it remains to be determined whether the protein encoded by this gene has transport activity for malate. Most recently, a gene encoding the Al-activated citrate transporter (HvAACT1) has been identified in barley (Furukawa et al., 2007). Barley is the most sensitive species to Al toxicity among cereal crops, although there is a wide genotypic variation in Al resistance in barley (Zhao et al., 2003). A physiological study showed that an Al-tolerant cultivar of barley secretes citrate from the roots in response to Al, with a good correlation between Al resistance and the amount of citrate secreted (Zhao et al., 2003). A locus controlling Al-induced secretion of citrate is localized on chromosome 4HL, where an Al-resistant gene (Alt) exists (Ma et al., 2004a). Fine mapping combined with microarray analysis led to isolation of the gene responsible for Al-induced secretion of citrate from an Al-resistant cultivar of barley, Murasakimochi (Furukawa et al., 2007). This gene belongs to the multidrug and toxin excursion (MATE) family. The expression of HvAACT1 is not induced by Al, which is in agreement with the rapid secretion of citrate upon Al exposure (Zhao et al., 2003). Interestingly, this gene is expressed not only in the root tips but also in the mature region of the roots. A positive correlation was observed between Al-activated citrate secretion and the expression of this gene in cultivars differing in Al resistance, suggesting that high expression of this gene is crucial for Al resistance. When the cRNA encoding this gene was injected into Xenopus oocytes with citrate or malate, transport activity was observed for citrate, but not for malate. Overexpression of HvAACT1 in tobacco enhanced the secretion of citrate and Al resistance. The protein encoded by this gene is localized on the plasma membrane. Furthermore, immunostaining showed that the protein was localized in the epidermal cells of the roots. Because a number of plant species secrete citrate in response to Al, identification of the homologs of HvAACT1 will help us better understand the Al-induced citrate secretion process. Al-induced secretion of oxalate has been reported in several plant species including buckwheat and taro, and physiological studies also show that a

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transporter is involved in this secretion (Zheng et al., 1998). However, genes responsible for oxalate secretion have not been isolated. Metabolism of organic acid Is not a key factor in aluminum resistance Altered organic acid metabolism is also implicated in the Al-induced secretion of organic acid anions (Kochian et al., 2005). The effect of Al on organic acid metabolism has been examined in several studies, but there has been no consensus regarding the effect of Al on internal organic acid contents or citrate synthase activity. In a study with two triticale lines differing in Al resistance, the concentrations of citrate (root apices and mature root segments) and malate (mature segments only) in roots increased during exposure to Al, but similar changes were observed in Al-sensitive and -tolerant lines (Hayes and Ma, 2003). The in vitro activities of the four enzymes involved in malate and citrate metabolism (citrate synthase, phosphoenolpyruvate carboxylase, malate dehydrogenase, and NADP-isocitrate dehydrogenase) were similar in the sensitive and resistant lines in both root apices and mature root segments. The response of these enzymes to pH did not differ between tolerant and sensitive lines or with the presence of Al. These results indicate that the Al-dependent efflux of organic acid anions from the roots of triticale is not regulated by their internal levels in the roots or by the capacity of the root cells to synthesize malate and citrate. Transgenic plants overexpressing genes that encode enzymes involved in organic acid metabolism have been made in several studies. Although enhanced secretion of organic acid anions and Al resistance has been found in some cases, the effect is quite small or not observed. For example, transgenic tobacco lines expressing a citrate synthase (CS) gene derived from Pseudomonas aeruginosa did not show a higher internal citrate concentration, enhanced citrate secretion, or greater Al resistance (Delhaize et al., 2001). In another study, Arabidopsis thaliana was transformed to overexpress CS isolated from carrot mitochondria. The transformants showed up to threefold increases in CS activity and 1.6-fold increases in citrate secretion compared with controls (Koyama et al., 2000), but the Al resistance in these plants was increased only slightly. In alfalfa, overexpression of malate dehydrogenase (MDH) resulted in enhanced organic acid synthesis and secretion and greater Al resistance (Tesfaye et al., 2001). However, overexpression of the phosphoenolpyruvate carboxylase enzyme in transgenic alfalfa did not result in increased root exudation of organic acids. Overexpression of an Arabidopsis mitochondrial CS gene in canola also resulted in increased citrate secretion and Al tolerance (Anoop et al., 2003), but the effect is small. In contrast, barley overexpressing the wheat ALMT1 gene significantly enhanced the secretion of malate and Al resistance both in solution culture and in an acid soil (Delhaize et al., 2004). These findings suggest that metabolism of organic acids is not a limiting factor for Al-induced secretion

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of organic acid. In fact, only a small percentage of internal organic acid is secreted in response to Al. 3.1.2. Aluminum-induced secretion of phenolic compounds Secretion of phenolic compounds may also be one of the mechanisms for Al resistance (see Fig. 5.2A) (Barcelo and Poschenrieder, 2002). Aluminum triggers release of catechol and flavonoid-type phenolics: catechin and quercetin from maize roots (Kidd et al., 2001). Phenolic compounds also form stable complexes with Al, thereby detoxifying Al. However, the difference in the secretion of phenolic compounds between cultivars differing in Al resistance is small. Therefore, the role of phenolic compounds in Al resistance remains to be examined. 3.1.3. Aluminum-altered pH in the rhizosphere Alkalinization of the rhizosphere by root apices has long been proposed to be an Al resistance mechanism because Al toxicity dramatically decreases with increasing pH due to decreased toxic Al3þ activity (Fig. 5.2B). Evidence supporting this hypothesis has been given in a study with A. thaliana (Degenhardt et al., 1998). In an Al-resistant mutant of Arabidopsis (alr-104), exposure to Al induced a twofold increase in the net Hþ influx localized to the root apex. Recently, a new Arabidopsis mutant (alt1–1) was isolated through mutagenesis of the Al-sensitive Arabidopsis mutant als3–1 (see later for details) (Gabrielson et al., 2006). This mutant showed increased Al resistance, which was attributed to the ability to adjust the pH of the rhizosphere. When the roots were exposed to Al in a pH-buffered solution, the increased Al resistance was lost. However, it is not clear if the alt1–1 Al resistance mechanism is directly dependent on pH adjustment or if it represents an alteration in transport of an unidentified substrate that mediates the observed Al resistance (Gabrielson et al., 2006). Future identification of this gene will help in understanding this mechanism. 3.1.4. Aluminum resistance and cell wall pectin and its methylation Immobilization of Al in the cell wall may be one of the exclusion mechanisms, but convincing evidence supporting this hypothesis has not been obtained. However, there is evidence that a larger accumulation of Al in the cell wall results in higher sensitivity to Al in maize cells (Schmohl and Horst, 2000). A recent study showed that lower pectin content and its higher degree of methylation contribute to Al resistance in maize (Fig. 5.2C) (Eticha et al., 2005). As described above, pectin in the cell wall is the major site of Al binding. Methylation of the pectin decreases the negativity of the cell wall and subsequently Al accumulation. The degree of methylation could explain the differential accumulation of Al and the resulting

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differential responses to Al in maize cultivars differing in Al resistance (Eticha et al., 2005). Aluminum may enhance the activity of methyl esterase in the Al-resistant cultivar of maize, although the mechanisms remain unknown. 3.1.5. Redistribution of aluminum Functional analysis of genes cloned from Al-sensitive Arabidopsis mutants showed that redistribution of Al is involved in Al resistance (Fig. 5.2D) (Larsen et al., 2005). One example is ALS3, which encodes a plasma membrane localized transmembrane protein. This protein has some ABC transporter features but lacks an ATPase domain. This gene is expressed in both the roots and shoots; the expression in the roots is induced by Al. ALS3 is localized to the phloem throughout the plant, leaf hydathodes, and the epidermis of the root. ALS3 may represent a component of a mechanism responsible for movement of Al away from the sensitive tissues for sequestration in more tolerant tissues (Larsen et al., 2005). Homologs of ALS3 are found in both dicots and monocots, suggesting that redistribution of Al away from the sensitive part may be a common Al resistance mechanism in plants. Recently, a novel gene (ALS1) was also cloned from an Al-sensitive Arabidopsis mutant (Larsen et al., 2007). This gene also encodes an ABC transporter, but in contrast to ALS3, the protein encoded by this gene is localized at the vacuolar membrane of root cells. The gene is expressed in the roots, leaves, stems, and flowers. The expression of this gene is not induced by Al. ALS1 may be important for facilitating vacuolar sequestration of Al in Al-sensitive tissues such as root tips (see Fig. 5.2D) (Larsen et al., 2007). 3.1.6. Uncharacterized aluminum resistance mechanisms Some plant species show very high resistance to Al toxicity, but their high resistance could not be explained by any known mechanisms as described above. For example, signalgrass (Brachiaria decumbens), a widely sown tropical forage grass, shows high Al resistance compared to the closely related ruzigrass (Brachiaria ruziziensis). However, the secretion of organic acid anions from the roots was small, and there was no significant difference in the secretion between signalgrass and ruzigrass, which differ in Al resistance (Wenzl et al., 2001). Although root apices of signalgrass alkalinized the rhizosphere more than did those of ruzigrass, the difference also cannot explain the difference in Al resistance between these two species. Rice is a highly Al-resistant species. In contrast to other cereal plants, organic acid anion secretion from the roots is not a mechanism for Al resistance in this species (Ma et al., 2002). Several QTLs for Al resistance have been identified in this plant (Ma and Furukawa, 2003), but neither the genes controlling high Al resistance nor the mechanisms responsible for high Al resistance are understood. Recently, a rice mutant (als1) that shows hypersensitivity to Al has been isolated from M3 lines derived from a

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cultivar of Koshihikari (Al-resistant) subjected to g irradiation ( J. F. Ma et al., 2005). Map-based cloning led to the identification of this gene (Huang and Ma, 2006). Als1 consists of four exons and three introns. Sequence analysis of this gene in the mutant showed a 15 bp deletion at the second exon. This gene was mainly expressed in the roots, and the expression was upregulated by a short exposure (2 h) to Al. The protein encoded by this gene is localized in the plasma membrane of all cells at the root tips. Furthermore, higher Al was detected in the cytoplasm of all root cells of the mutant, but in low concentration in the wild-type rice. These results suggest that Als1 is involved in the exclusion of Al from the cells, which represents a novel resistance mechanism (Fig. 5.2E). Further functional analysis will help us better understand the high Al resistance in rice. 3.1.7. Other mechanisms Immobilization of Al with phosphorus in the root cell wall may be an additional external detoxification mechanism of Al in buckwheat (Zheng et al., 2005). After a long exposure to Al, more Al and P were observed in the roots of an Al-resistant cultivar than in an Al-sensitive cultivar of buckwheat. Furthermore, Al was found to be localized in the cell wall, suggesting that Al is precipitated with P in the cell wall to prevent Al from entering the root cell. However, P deficiency is also a limiting factor of crop production in acidic soils; therefore, coprecipitation between Al and P in the roots will affect P acquisition. Buckwheat is an Al-accumulating species and possesses internal detoxification mechanisms as described below. Therefore, unlike most plant species, it does not seem necessary to stop Al in the cell wall. The phenomenon observed after long Al treatment may not be the cause, but the result of differential Al resistance between cultivars. Release of an Al-binding mucilage by border cells may play a role in protecting root tips from Al-induced cellular damage (Miyasaka and Hawes, 2001; Pan et al., 2004). However, a study with maize showed that although the mucilage secreted from the root cap cells has a strong binding ability with Al, the total binding capacity of the mucilage was too small to confer effective protection from Al-induced root inhibition (Li et al., 2000).

3.2. Internal detoxification of aluminum with organic acids In most plant species, Al is retained in the roots as described above, and little is translocated to the shoots. However, a small number of plant species such as tea, hydrangea, and buckwheat accumulate large concentrations of Al in the aboveground plant tissues without suffering from Al toxicity ( Jansen et al., 2004). For example, hydrangea plants accumulate more than 3000 mg kg-1 in both leaves and sepals during a several-month growth period (Ma et al., 1997). Buckwheat accumulated 12,000 mg kg-1 Al when grown in a

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very acidic soil (Shen et al., 2004). Some woody species adapted to acidic soils also accumulate high levels of Al. For example, Melastoma malabathricum accumulates more than 10,000 mg kg-1 Al in the mature leaves (Watanabe and Osaki, 2001). A high accumulation of Al in the shoots suggests that Al is transported across the plasma membrane into the symplasm in these Al-accumulating plant species. Symplasmic solutions usually have a pH above 7.0. Although the concentration of free Al is decreased to less than 10-10 M at pH 7.0 due to formation of insoluble Al(OH)3, such low concentrations are still potentially phytotoxic because of the strong affinity of Al for oxygen donor compounds. For example, Al binds almost 107 times more strongly to ATP than does Mg; therefore, less than nanomolar concentrations of Al can compete with Mg for the phosphate sites (Martin, 1988). These facts suggest that Al-accumulating plants must possess effective mechanisms to detoxify Al internally. Recent studies revealed that the internal detoxification is achieved by both complexation and sequestration. Furthermore, there are distinct transport systems for Al from roots to shoots in these Al-accumulating plants (Fig. 5.3).

D

OAA

Vacuole Al-OAA Leaf Al-OAB

A

Al3+

B OAA

C OAB

Al-OAA

Root

Al-OAB

Xylem

Figure 5.3 Mechanisms of internal Aldetoxification. Aluminum istakenup in the form of ionic Althrough anunidentified transporter (A) andthenchelated withorganic anions (OA) including oxalate and phenolic compounds (B). Ligand exchange from Al^OA to Al^citrate occurs when Al is released to the xylem and then Al^citrate is translocated to the shoots (C). Finally, Al is sequestrated in the vacuoles in the chelated form (D).

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3.2.1. Aluminum uptake and translocation Detailed studies with buckwheat and Melastoma have been conducted regarding the process leading to the accumulation of Al (Ma, 2005a; Ma et al., 2001). Because buckwheat secretes oxalate in response to Al, the roots might take up Al in the form of an Al–oxalate complex. However, uptake experiments with different forms of Al revealed that the form of uptake is not the Al–oxalate complex, but the ionic Al (Al3þ) (Fig. 5.3A) (Ma and Hiradate, 2000). Furthermore, the uptake is very fast and seems to be a passive process due to a large inwardly directed electrochemical gradient for this ion, because the uptake was not affected by a respiratory inhibitor, hydroxylamine (Ma and Hiradate, 2000). In Melastoma, Al uptake is also likely to be a passive process because the uptake was not affected by carbonyl cyanide m-chlorophenylhydrazone (CCCP), a protonophore (Watanabe et al., 2001). Rapid uptake suggests that Al3þ uptake is mediated by a transporter, which remains to be identified. The form of Al in buckwheat roots has been identified as Al–oxalate at a 1:3 molar ratio (Ma et al., 2001). This suggests that following uptake, Al3þ is immediately chelated with the internal oxalate in the root cells, forming a stable, nonphytotoxic complex of Al–oxalate (Fig. 5.3B) (Ma et al., 2001). In some plant species, phenolic compounds have been proposed as a ligand for chelating Al (Ofei-Manu et al., 2001; Tolra et al., 2005). In buckwheat and Melastoma, the synthesis of oxalate was not induced by Al (Ma and Hiradate, 2000; Watanabe et al., 1998). In Rumex acetosa, Al induced high levels of anthraquinone in the roots (Tolra et al., 2005). There was a good correlation between Al tolerance and the concentration of soluble phenolic compounds in 10 woody plants, although synthesis of these compounds was not affected by Al (Ofei-Manu et al., 2001). It remains to be examined whether Al is bound to these phenolic compounds in the root cells. It is interesting that Al in the xylem is present in the form of Al–citrate in buckwheat (Ma and Hiradate, 2000). This suggests that ligand exchange from oxalate to citrate occurs when Al is released to xylem (Fig. 5.3C). Oxalate forms insoluble precipitates with Ca2þ. Therefore, buckwheat plants use citrate as a safe carrier for Al in the xylem. In Melastoma, Al is also translocated in the form of Al–citrate (Watanabe and Osaki, 2001). However, in contrast to buckwheat in which the citrate concentration in the xylem sap was not affected by Al (Ma and Hiradate, 2000), the concentration of citrate in the xylem was increased in Melastoma (Watanabe and Osaki, 2001). 3.2.2. Internal detoxification mechanisms in leaves Aluminum is mainly found in the leaves of Al-accumulating plants. Therefore it is important to know the form and subcellular localization of Al in the cells in order to understand the internal detoxification mechanisms.

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The chemical forms of Al in the leaves have been identified in several Al-accumulating plants (Fig. 5.3D). Al was identified as the form of the catechin–Al complex in tea using 27Al nuclear magnetic resonance (NMR) (Nagata et al., 1993). In hydrangea leaves, Al is complexed with citrate (Ma et al., 1997). In buckwheat, Al is present in the form of an Al–oxalate (1:3) complex. In Melastoma leaves, Al is present in the form of monomeric, Al– oxalate, Al–(oxalate)2, and Al–(oxalate)3 (Watanabe et al., 1998). As monomeric Al and Al–oxalate (1:1) show phytotoxicity, it will be interesting to investigate where Al is located in the leaf cells of Melastoma. It has also been suggested that Al is complexed with Si in the leaves of Faramea marginata, which accumulates Al up to 23 g kg-1 (Britez et al., 2002). A positive correlation was found between the Al and Si levels, and both elements showed a similar distribution in leaf and stem tissues. Furthermore, the Al and Si elution patterns were similar to those of aluminum silicate, and the mole ratio of Si:Al was approximately 0.5. However, Si forms a weak complex with Al, compared with organic acids (e.g., citrate); therefore, the role of Si in the internal detoxification of Al needs to be examined further. In Rumex acetosa, Al induced high levels of catechol, catechin, and rutin in the shoots (Tolra et al., 2005). However, the role of these phenolic compounds in internal detoxification of Al is still unclear. The patterns of tissue, cellular, and subcellular localization of Al have been extensively investigated in buckwheat. More Al is accumulated in the old leaves than in the young leaves (Shen and Ma, 2001). In a leaf, a steep gradient in the Al concentration (central part < middle part < marginal part) was observed regardless of leaf position. These observations indicate that Al distribution in buckwheat leaves is controlled by transpiration. This may also explain why buckwheat seeds accumulate much less Al compared to the leaves because of a low transpiration from the seeds (Shen et al., 2006). In addition, Al is not mobile once it is accumulated in leaves (Shen and Ma, 2001). Direct isolation of pure protoplasts and vacuoles from buckwheat leaves showed that Al in the cells is localized in the vacuoles (see Fig. 5.3D) (Shen et al., 2002). In Melastoma, Al was located in the upper epidermal cells and also in mesophyll cells (Watanabe et al., 1998), but the subcellular localization of Al in this plant species is unknown. A recent interesting finding is that the form of Al in buckwheat leaves varies with leaf position, mainly depending on Al concentration in buckwheat (Shen et al., 2004). When the Al concentration in the leaf was high, and the ratio of oxalate to Al was lower than 3, Al was present in the form of both Al–oxalate (1:3) and Al–citrate (1:1) in the old leaves. As the biosynthesis of oxalate is not enhanced by increased Al, it seems that the conversion from Al–citrate in the xylem to Al–oxalate in the leaves does not occur at higher Al concentrations.

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4. Beneficial Effect of Aluminum on Plant Growth Several previous studies have shown that Al has a beneficial effect on plant growth in some plant species such as tea, Melastoma, and Quercus serrata Thunb. under certain growth conditions. For example, in contrast to Al toxicity described above, root elongation was enhanced 2.5 times in tea grown in a nutrient solution containing 0.5 mM Al at pH 4.3 (Ghanati et al., 2005). The root biomass of Q. serrata increased with increasing Al concentrations up to 2.5 mM in a solution with pH 3.5 (Tomioka et al., 2005). In Melastoma, Al increased the root activity and stimulated the elongation of root cells (Watanabe et al., 2005). The exact mechanisms for the Al-induced beneficial effects are unknown. Two possible mechanisms were previously suggested. The first suggests that Al probably ameliorates proton toxicity in roots because the beneficial effect of Al is usually observed at low pH. However, a recent study with Q. serrata showed that Al-induced growth enhancement is not due to the amelioration of Hþ toxicity by Al (Tomioka et al., 2005). The second suggests that Al improves P nutrition. However, results from a study with Melastoma do not lend support to this hypothesis (Watanabe and Osaki, 2001). Recently, other possible mechanisms for Al-induced beneficial effects have been reported. Aluminum increased the activities of superoxide dismutase (SOD), catalase (CAT), and ascorbate peroxidase (APX) in the roots of both intact tea plants and cultured cells (Ghanati et al., 2005). An Al-induced increase in the activities of these antioxidant enzymes may cause increased membrane integrity and delayed lignification and aging, resulting in a stimulation of growth. In Melastoma, the primary reason for the Al-induced growth enhancement may be the alleviation of Fe toxicity by Al (Watanabe et al., 2006). The growth was enhanced by Al more under an excess of Fe, and the Fe concentration was decreased by Al in both the roots and shoots. Excess Fe induces the production of reactive oxygen species, leading to the disorder of various cell functions. Plants grown in acid soils are also exposed to an excess of Fe; therefore, Al-induced alleviation of Fe toxicity is important for Melastoma species in these soils (Watanabe et al., 2006).

5. Concluding Remarks Al-induced inhibition of root elongation is a result of a number of Al-induced physiological and biochemical changes (see Fig. 5.1). Although these Al-induced changes are diverse, all of them are basically caused by the binding of Al to extracellular and intracellular substances. Aluminum has a

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strong binding affinity for the oxygen donor compounds such as inorganic phosphate, nucleotides, RNA, DNA, proteins, carboxylic acids, phospholipids, polygalacturonic acids, heteropolysaccharides, lipopolysaccharides, flavonoids, and anthocyanins (Martin, 1988). The binding of Al to the cells results in considerable damage to the structure and function of the roots. Therefore, Al targets multiple sites of the root cells simultaneously, not only one site. In fact, recent genome-wide analysis showed that Al induces more than 10 genes within 30 min after exposure to Al in rice and more than 500 genes after 6 h ( J. F. Ma et al., unpublished data). Furthermore, most genes are also induced by other general stresses such as phosphorus deficiency, wounding, and oxidative stress. Therefore, in my opinion, it is difficult to identify the primary mechanism of Al toxicity. On the other hand, great progress has been made in the identification of physiological mechanisms of Al resistance during the past decade. In particular, the Al-induced secretion of organic acid anions has been investigated and characterized in a number of plant species. However, only a few genes responsible for Al resistance have been identified so far. Aluminum resistance may be controlled by a single gene or multiple genes, depending on the plant species. For example, Al resistance is controlled by a single dominant gene in barley (Ma et al., 2004a), but by multiple genes in maize (Pineros et al., 2005). Many QTLs for Al resistance have been reported in a number of plant species. Mutants sensitive or tolerant to Al have also been isolated. With the development of molecular techniques, genes responsible for Al resistance can be identified using the QTL information and mutants; this will in turn lead to a better understanding of the molecular mechanisms of Al resistance. In Al-accumulating plant species, although great progress has been made in understanding the internal detoxification of Al at the physiological level, less is known at the molecular level. For example, for transport of different Al forms from roots to leaf vacuoles, Al must pass across the plasma and tonoplast membranes (see Fig. 5.3). However, the transport system involved remains unknown. Identification of transporters involved in the transport of different Al forms in different cells will help in better understanding the internal detoxification of Al in plants.

ACKNOWLEDGMENTS This review paper focuses on progress made over the past 5 years; therefore most of the earlier papers are not cited. I thank Naoki Yamaji for photos and figures and Fangjie Zhao and other colleagues for critical reading. Some work described in this paper is supported by a grant from the Ministry of Agriculture, Forestry and Fisheries of Japan (Green Technology QT 3001) and a grant-in-aid for General Scientific Research (Grant 18380052) from the Ministry of Education, Sports, Culture, Science and Technology of Japan.

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Index

A Actin filaments desmosome crosstalk, 120–121 subepithelial fibroblasts colon, 176–177 small intestine, 174, 176 Adherens junctions, desmosome crosstalk, 118–120 Alcoholism, diacylglycerol kinase pathophysiological studies in animal models, 42–43 ALMT1, aluminum detoxification in plants, 236–237 ALS, aluminum detoxification in plants, 240 Aluminum abundance, 226 isotopes, 226 plant growth beneficial effects, 245 resistance in plants exclusion mechanisms cell wall pectin and methylation, 239–240 organic acid anion secretion induction, 234–239 pH alteration in rhizosphere, 239 phenolic compound secretion induction, 239 redistribution, 240 internal organic acid detoxification leaves, 243–244 overview, 241–242 uptake and translocation, 243 mucilage binding, 241 phosphorous immobilization, 241 prospects for study, 246 species distribution, 240–241 toxicity in plants cell division inhibition, 227 mechanisms auxin transport inhibition, 233 calcium homeostasis disruption, 232–233 callose deposition, 230 cell wall interactions, 229–230 cytoskeleton disruptions, 231 magnesium uptake inhibition, 233 overview, 227, 229 oxidative stress, 231–232 plasma membrane disturbances, 230–231

signal transduction pathway interactions, 232 root elongation inhibition, 227 Angiotensin II, subepithelial fibroblast receptors, 187 Arm-repeat domain proteins desmosome composition, 80–84 phosphorylation, 106–107 Arrhythmogenic right ventricular cardiomyopathy, desmosomal gene mutations, 125, 126, 128 Ascorbate peroxidase, induction in plants by aluminum, 245 Astrocyte, subepithelial fibroblast comparison, 190–191 AtEMR1, vacuole sorting receptor, 9 ATP receptors, subepithelial fibroblasts mechanosensitive networks, 195–196 P2Y1 immunofluorescence microscopy, 186 subtypes, 190 Auxin, transport inhibition by aluminum, 233 B BP-80, vacuole sorting receptor, 8–9 Brain, diacylglycerol kinase expression, 30–31 pathophysiological studies in animal models cerebral infarction, 37–39 emotion and alcoholism, 42–43 hypothalamic energy balance regulation, 41–42 seizure susceptibility, 39–41 transient cerebral ischemia, 34–35, 37 C Cadherin b-catenin complex, 81, 108 desmosome classical cadherin homology, 78–79 domain structures, 76–77, 79–80 genomic organization, 74 phylogenetic tree, 74–76 types, 72–73 Calcium flux aluminum disruption in plants, 232–233 desmosome dynamics, 101–103, 105–106 subepithelial fibroblasts

253

254

Index

Calcium flux (cont.) mechanosensitive networks stretch induction, 198 touch induction, 196 receptor studies, 187–191 signal propagation to neural cells, 202 Callose, deposition in aluminum toxicity, 230 Cancer, desmosome mediation, 133–135 Catalase, induction in plants by aluminum, 245 b-Catenin cadherin complex, 81, 108 cancer mutations, 134 Central vacuole, functions, 2 Collagen IV, subepithelial fibroblasts, 177 Corneodesmosin, desmosome composition, 87 D Desmin, subepithelial fibroblasts, 176 Desmocalmin, desmosome composition, 87 Desmocollins desmosome composition, 72, 81, 94–96 knockout mouse phenotypes, 109–110, 115, 117 ectopic synthesis studies, 114, 117–118 Desmogleins bacterial toxin targeting, 132 desmosome composition, 72, 81–82, 94–96 ectopic synthesis studies, 113–114, 117–118 knockout mouse phenotypes, 109, 112, 115, 117 Desmoplakin desmosome composition, 72 knockout mouse phenotype, 110–111, 113, 115 phosphorylation, 107 Desmosome biogenesis developmental studies, 97–98 experimental analysis, 98–100 cadherins classical cadherin homology, 78–79 domain structures, 76–77, 79–80 genomic organization, 74 phylogenetic tree, 74–76 types, 72–73 crosstalk adherens junctions, 118–120 cytoskeletal filaments actin filaments, 120–121 intermediate filaments, 120 microtubules, 121 nucleus protein shuttling, 121–122 signal transduction, 122–123 diseases autoimmune disease, 129–132 bacterial toxin targets, 132 cancer, 133–135

gene mutations, 123–129 dynamics adhesion regulators, 107–108 calcium-dependent alterations, 101–103, 105–106 cell cycle, 100–101 phosphorylation-dependent alterations, 106–107 knockout mouse phenotypes, 108–117 morphology diversity, 70–72 ultrastructure, 67, 69–70 plaque components arm-repeat domain proteins, 80–84 cell type specificity, 89–97 desmocollins, 72, 81, 94–96 desmogleins, 72, 81–82, 94–96 desmoplakin, 72 miscellaneous components, 87, 89 plakins, 84–87 plakoglobin, 72, 80–82 plakophilin, 72, 80, 82–84 types, 72 tissue distribution, 66 transgenic mouse studies, 117–118 Desmoyokin desmosome composition, 87 knockout mouse phenotype, 112 DGK, see Diacylglycerol kinase Diacylglycerol kinase brain expression, 30–31 diacylglycerol features and actions, 26–27 isoforms, functions, and classification, 27–30 pathophysiological studies in animal models brain cerebral infarction, 37–39 emotion and alcoholism, 42–43 hypothalamic energy balance regulation, 41–42 seizure susceptibility, 39–41 transient cerebral ischemia, 34–35, 37 dorsal root ganglion, 43–44 heart hypertrophy, 47–49 myocardial infarction, 49–51 knockout mouse studies, 52–53 lung, 51 ovary and placenta, 51–52 T cells, 44–47 prospects for study, 54 subcellular localization, 32–34 Dorsal root ganglion, diacylglycerol kinase pathophysiological studies in animal models, 43–44

255

Index

E Ectodermal dysplasia skin fragility syndrome, desmosomal gene mutations, 124, 126 Electron microscopy, subepithelial fibroblasts cultured cell ultrastructure, 182–183 scanning electron microscopy, 170–171 transmission electron microscopy, 171–172, 174 Endothelins intestinal villi sources and functions, 210–211 subepithelial fibroblasts cell shape conversion induction, 180–181 receptors, 185–186, 188–189, 210 ENTH domain, vacuolar transport role, 9 Erbin, desmosome composition, 87 F Fibroblast, see Subepithelial fibroblast Fluorescence recovery after photobleaching, subepithelial fibroblast permeability studies, 193–195 FRAP, see Fluorescence recovery after photobleaching G Gastrointestinal tract fibroblasts, see Subepithelial fibroblast Green algae vacuoles, see Vacuoles, plants H Heart, diacylglycerol kinase pathophysiological studies in animal models hypertrophy, 47–49 myocardial infarction, 49–51 HvAACT1, aluminum detoxification in plants, 237 I Intermediate filaments, desmosome crosstalk, 120 Intestinal villi, see Subepithelial fibroblast K Keratocalmin, desmosome composition, 87 L Laminins, subepithelial fibroblasts, 177–178 Localized autosomal recessive hypotrichosis, desmosomal gene mutations, 125, 127 Lung, diacylglycerol kinase expression, 51

Microvasculature, subepithelial fibroblast interactions, 208–209 Myocardial infarction, diacylglycerol kinase pathophysiological studies in animal models, 49–51 Myofibroblast, see Subepithelial fibroblast N Naxos disease, desmosomal gene mutations, 125, 127–128 Nitric oxide synthase, aluminum inhibition, 232 NOS, see Nitric oxide synthase O Ovary, diacylglycerol kinase expression, 51–52 Oxidative stress, aluminum toxicity, 231–232 P P2Y1, see ATP receptors Pectin, methylation and aluminum resistance in plants, 239–240 Pemphigus foliaceus desmosomal gene mutations, 129–131 pathogenesis, 131–132 Pemphigus vulgaris desmosomal gene mutations, 129–131 pathogenesis, 131–132 Phosphatidic acid features and actions, 26–27 synthesis, see Diacylglycerol kinase Phosphorous, aluminum immobilization in root cell wall, 241 Pinin, desmosome composition, 87 Placenta, diacylglycerol kinase expression, 51–52 Plakins, desmosome composition, 84–87 Plakoglobin desmosome composition, 72, 80–82 knockout mouse phenotype, 111–112, 116 mutation and disease, 125 phosphorylation, 106 Plakophilin desmosome composition, 72, 80, 82–84 knockout mouse phenotype, 112, 115 phosphorylation, 106–107 Plant vacuole, see Vacuoles, plants Proton-ATPase, aluminum detoxification in plants, 235 R

M Magnesium, uptake inhibition by aluminum in plants, 233 Microtubules, desmosome crosstalk, 121

Root, aluminum toxicity elongation inhibition, 227 organic acid anion secretion induction in resistance, 234–239

256

Index

S Serotonin, enterochromaffin cell mechanosensory signaling, 206–207 Smooth muscle, subepithelial fibroblast interactions, 208 Staphylococcus aureus toxin, desmoglein targeting, 132 Striate palmoplantar keratoderma, desmosomal gene mutations, 124, 126 Stroke, diacylglycerol kinase pathophysiological studies in animal models cerebral infarction, 37–39 transient cerebral ischemia, 34–35, 37 Subepithelial fibroblast barrier/sieve function regulation in villi cell shape conversion and sieve size, 203 cytokine regulation of epithelial barrier function, 204 calcium flux propagation to neural cells, 202 cell cycle, 169 contractility, 200, 202, 204, 206 culture cell shape conversions, 179–182 cytoskeleton, 184–185 intestinal myofibroblast cell lines, 178–179 primary culture, 179–182 ultrastructure, 182–183 vitamin A uptake, 182 cytokine secretion, 204, 209 functional overview, 167–168 gap junctions dye coupling between adjacent cells, 193 fluorescence recovery after photobleaching permeability studies, 193–195 morphology, 191, 193 intestinal villi morphology, 168–169 mechanosensitive networks ATP receptors, 195–196 calcium flux and ATP release stretch induction, 198 touch induction, 196 cell shape effects, 199–200 villi mechanosensory signals, 206–211 morphology cytoskeleton and contractile proteins colon, 176–177 small intestine, 174, 176 extracellular matrix and receptor differential expression along villus–crypt axis, 177–178

scanning electron microscopy, 170–171 transmission electron microscopy, 171–172, 174 origins, 169 proliferation, 169 receptors astrocyte comparison, 190–191 calcium flux studies, 187–191 immunohistochemistry and in situ hybridization, 185–187 Substance P, subepithelial fibroblast receptors, 187 Superoxide dismutase, induction in plants by aluminum, 245 T T cell diacylglycerol kinase in function, 44–47 subepithelial fibroblast interactions, 209 Tenascin C, subepithelial fibroblasts, 177 TGF-b, see Transforming growth factor-b TNF-a, see Tumor necrosis factor-a Transforming growth factor-b, subepithelial fibroblast secretion and barrier function regulation, 204 Tumor necrosis factor-a, subepithelial fibroblast secretion and barrier function regulation, 204 V Vacuoles, plants central vacuole functions, 2 embryophyte vacuoles functions autophagy, 6–7 storage, 3–6 turgor, 3 protein targeting, 8–10 structure and development studies, 7–8 types, 3 evolution, 13, 15–17 green algae vacuoles contractile vacuole, 10–11 development, 12 granules, 11–12 prospects for study, 17–18 size, 2 Vascular sorting signals, plants, 9–10 Vimentin, subepithelial fibroblasts, 176 Vitamin A, subepithelial fibroblast uptake, 182