International Review of Cytology, A Survey of Cell Biology, Vol. 191

VOLUME 191 SERIES EDITORS Geoffrey H. Bourne James F. Danielli Kwang W. Jeon Martin Friedlander Jonathan Jarvik 1949...

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VOLUME 191

SERIES EDITORS Geoffrey H. Bourne James F. Danielli Kwang W. Jeon Martin Friedlander Jonathan Jarvik

1949-1 988 1949-1 984 19671984-1 992 1993-1 995

EDITORIAL ADVISORY BOARD Eve Ida Barak Rosa Beddington Howard A. Bern Robert A. Bloodgood Dean Bok Stanley Cohen Rene Couteaux Marie A. DiBerardino Laurence Etkin Hiroo Fukuda Elizabeth D. Hay P. Mark Hogarth Anthony P. Mahowald

M. Melkonian Keith E. Mostov Andreas Oksche Vladimir R. Pantic Jozef St. Schell Manfred Schliwa Robert A. Smith Wilfred D. Stein Ralph M. Steinman M. Tazawa Donald P. Weeks Robin Wright Alexander L. Yudin

Edited by

Kwang W. Jeon Department of Biochemistry University of Tennessee Knoxville. Tennessee

VOLUME 191

ACADEMIC PRESS San Diego London

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Front cover photograph: Electron micrograph of axon terminals bordering the perivascular space which surrounds capillaries of the primary portal plexus in the neurohypophyseal median eminence. (For more details, see Chapter 5, Figure 1.)

This book is printed on acid-free paper.

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Copyright 0 1999 by ACADEMIC PRESS 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. (222 Rosewood Drive, Danvers, Massachusetts 01923), 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. COOVfees for ore-1999 chaoters are as shown on the title oaws. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0074-7696/99 $30.00 1,

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CONTENTS

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

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Role of Activin and Other Peptide Growth Factors in Body Patterning in the Early Amphibian Embryo Makoto Asashima, Kei Kinoshita, Takashi Ariizumi, and George M. Malacinski I. II. Ill. IV. V. VI. VII. VIII.

Introduction . . . ................................... Mesoderm-Inducing Factors and Their Modifiers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Effects of Activin on the Regional Expression of Specific Axial Patterning by Activin(s) . . . . . . . . . . . . . . . . . . . . . . . Life History of Activin Signaling Mechanisms of the Embryo . . . . . . . . . . . . . . Proposed Molecular Models for Activin's Role in Signal Tr Activin Causes a Broad Array of Dlfferentiations in Vitro ........................ Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............ References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ......

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Calcium Regulation of the Actin-Myosin Interaction of Physarum polycephalum Akio Nakamura and Kazuhiro Kohama I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Calcium Inhibition of Motile Events Related to Actornyosin ...................... 111. Calcium Inhibition of the Actin-Myosin Interaction of Physarum as Detected In L/ltro.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Ca-Binding Properties of Physarum Myosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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CONTENTS

Phosphorylated State and Calcium Inhibition of Physarum Myosin . . . . . . . . . . . . . . . . Actin-Binding Proteins of Physarum That Are Involved in .............................. Calcium Inhibition . . . . . . . . . . . . . VII. Ca-Binding Proteins in Physarum .................... VIII. Ameba1 Myosin and Arnebal-Plas .................... IX. Concluding Remarks . .................... References . . . . . . . . . . . ................................. V. VI.

72 77 03 00 90 92

Characteristics of Skeletal Muscle in Mdx Mutant Mice Sabine De La Porte, Sophie Morin, and Jeanine Koenig I. II. 111. IV. V. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . Animal Models . . . . . . . . . . . . . . . . . . . . . . . . Dystrophin, Utrophin, and Associated Proteins Mdx Muscle Cells . , , , Therapeutic Projects . . .......... Concluding Remarks . . .......... References . . . . . . . . .

Regulation of Phosphate Transport and Homeostasis in Plant Cells Tetsuro Mimura

111.

Distribution of Inorganic Phosphate

V. Homeostasis and Detection of Pi Status in Plant Cells VI. Concluding Remarks

.......... .......... ............... ............... ...............

149 151 152 157 184

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Synaptic-like Microvesicles in Mammalian Pinealocytes Peter Redecker I. II. 111. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Mammalian Pineal Organ: A Mediator of Darkness . . . . . . . . . . . . . . . . . . . . . . . . . UltrastructuralObservations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Molecular Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emergence of Functional Concepts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

201 203 205 21 1 224

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CONTENTS

VI. VII.

Biogenesis of Synaptic-like Microvesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding Remarks and Future Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Invertebrate Integrins: Structure, Function, and Evolution Robert D. Burke I. 11. 111. IV.

Introduction ............................ Invertebrate lntegrins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Evolution of lntegrins . . . . . . . . . . . . . . . . . . . . Concluding Remarks ..................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

257 259 266 281 281

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Numbers in parentheses indicate the pages on which the authors' contributions begin.

Takashi Ariizumi (1)1 CREST Project, University of Tokyo, Komaba, Meguro, Tokyo 153, Japan Makoto Asashima (l), Department of Life Science and CREST Project, University of Tokyo, Komaba, Meguro, Tokyo 153, Japan Robert D. Burke (257),Department of Biology, University of Victoria, Victoria, British Columbia, Canada V8W 3N5 Sabine De La Porte (99), Laboratoire de Neurobiologie Cellulaire et Mo/eculaire, CNRS UPR 9040, 91 198 Gif sur Yvette Cedex, France Kei Kinoshita (l), CREST Project, University of Tokyo, Komaba, Meguro, Tokyo 153, Japan Jeanine Koenig (99), lnstitut de Myologie, Groupe Hospitalier Pitie-Salpetriere, 75651 Paris Cedex 13, France Kazuhiro Kohama (53),Department of Pharmacology, Gunma University School of Medicine, Maebashi, Gunma 371-8511, Japan George M. Malacinski (1), Department of Bio/ogy, Indiana University, Bloomington, Indiana 47405 Tetsuro Mimura (149),Biological Laboratory, Hitotsubashi University, Naka 2-1, Kunitachi, Tokyo 186-8601, Japan Sophie Morin (99), Laboratoire de Neurobiologie Cellulaire, Universite de Bordeaux 11, 33405 Talena Cedex, France Akio Nakamura (53) Department of Pharmaco/ogy,Gunma University School of Medicine, Maebashi, Gunma 371-8511, Japan Peter Redecker (201), Department of Anatomy 1, Hannover Medical School, 0-30625 Hannover, Germany ix

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Role of Activin and Other Peptide Growth Factors in Body Patterning in the Early Amphibian Embryo Makoto Asashima,*lt Kei Kinoshita,t Takashi Ariizumi,t and George M. Malacinski$ *Department of Life Science and ?CREST Project, University of Tokyo, Komaba, Meguro, Tokyo 153, Japan, and $Department of Biology, Indiana University, Bloomington, Indiana 47405

The amphibian body plan is established as the result of a series of inductive interactions. During early cleavage stages cells in the vegetal hemisphere induce overlying animal hemisphere cells to form mesoderm. The interaction represents the first major body-patterning event and is mediated by peptide growth factors. Various peptide growth factors have been implicated in mesoderm development, including most notably members of the transforming growth factor-p superfamily. Identification of the so-called "natural" inducer from among the several candidate peptide growth factors is being achieved by employing several experimental strategies, including the use of a tissue explant assay for testing potential inducers, cloning of marker genes as indices of early induction events, and microinjection of altered peptide growth factor receptors to disrupt normal embryonic inductions. Activin emerges as the most likely choice for assignment of the role of endogenous mesoderm inducer, because it currently best fulfills the rigorous set of criteria expected of such an important embryonic signaling molecule. Activin, however, may not act alone in mesoderm induction. Other peptide growth factors such as fibroblast growth factor might be involved, especially in the regional patterning of the mesoderm. In addition, several genes (e.g., Wnt and noggin), which are expressed after the mesoderm is initially induced, probably assist in further definition of the mesoderm pattern. Following mesoderm induction, the primary embryonic organizer tissue (first described in 1924 by Spemann) develops and contributes further to body patterning by its action as a neural inducer. Peptide growth factors such as activin may also be involved in the inductive event, either directly (by facilitating gene expression) or indirectly (by serving to constrain pathways). Inrrmurronal Review of CyruluKy. V d 191

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Copyright 0 1999 hy Academic Press. All rights of reproduction in any form reserved.

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MAKOTO ASASHIMA ETAL.

KEY WORDS: Mesoderm induction, Animal-cap explants, Activin, Fibroblast growth factors, Peptide growth factors, Embryogenesis. 0 1999 Academic Press.

1. Introduction The major aim of contemporary developmental biology research is to understand the mechanisms which guide formation of the body plan of a complex, multicellular animal from the single-celled egg. Inductive interactions, which involve signaling between groups of cells, are widely considered to play a fundamental role in the establishment of the basic body plan. A variety of such inductive interactions operating at different times and in different places in the embryo most likely combine to generate the changes in the overall form, shape, and regional specialization features which characterize morphogenesis. A research focus on the nature of inductive interactions began (for amphibian embryos) with the pioneering studies of Hans Spemann. Together with his student Hilde Mangold he described the role the so-called primary embryonic organizer tissue plays in orchestrating the formation of body plan in early embryos. For several decades the biochemical identity of the molecules which are responsible for the remarkable properties of the primary embryonic organizer remained unknown. Recently, dramatic progress has not only been made in identifying specific proteins (e.g., peptide growth factors) as probable inducers but also in uncovering other subcellular components (e.g., receptors) which play roles in establishing the body plan of the early embryo. In order to provide background information for understanding the role that peptide growth factors play in embryonic induction, and to appreciate the inherent complexity of the regulatory circuitry which comprises inductive interactions, this review will summarize some of the key early events which lead up to and follow the action of Spemann’s primary embryonic organizer tissue. A. Embryonic Axis Specification

Amphibian eggdembryos represent the experimental system of choice for most studies on polarization phenomena for obvious reasons: The eggs are large (approx 1-3 mm diameter), readily collected, easily manipulated, and conveniently pigmented [animal (upper) hemisphere = dark; vegetal (lower) hemisphere = light]. Originally, urodele embryos (e.g., newts and

ROLE OF ACTlVlN IN BODY PATTERNING

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axolotls) were researched, but recently anuran embryos (especially Xenopus) have become more popular as a ready source of large numbers of eggs. Most of the information included herein was collected from Xenopus studies, although comparisons of anuran inductive interactions to urodele embryos reveal strong similarities (Malacinski et al., 1996). Axis specification actually begins during oogenesis, the period during which, while in the ovary, the egg develops from a tiny, cytoplasm-poor cell to a large cell which consists mainly of yolk-laden cytoplasm. During oogenesis the oocyte becomes polarized along its animal (darkly pigmented)-vegetal (lightly pigmented) axis. Typically, the egg contains large, densely packed yolk platelets in the vegetal hemisphere and smaller, more loosely arranged yolk platelets in the animal hemisphere (Ubbels, 1977; Neff et al., 1984). The egg cytoplasm is organized in a radially symmetrical fashion about the animal-vegetal axis. That is, all meridia are identical, and each has the potential to form future dorsal or ventral structures (Fig. 1). It is in this state that a mature oocyte awaits fertilization. Fertilization breaks the egg’s radial symmetry. The entering sperm causes (i) the egg cortex to contract toward the side of the egg where the sperm entered, and (ii) the underlying cytoplasm to shift with it. This cortical rotation establishes the bilateral symmetry of the egg. The side opposite the sperm entrance site becomes the dorsal side and the sperm entrance side the ventral side (Gerhart et al., 1989). Right and left halves are thus simultaneously established vis-a-vis the animal-vegetal polarization mentioned previously. Experimental verification of this scenario has been achieved by disrupting the cortical rotation with ultraviolet irradiation and rescuing “depolarized” eggs by subsequently reorienting them 90” along the earth’s gravitational axis for a brief period (Chung and Malacinski, 1980; Elinson and Rowning, 1988; Gerhart et al., 1989). The molecular basis of dorsal-ventral polarization was speculated by Gerhart et al., (1991) to involve the activation of “dorsal determinants of polarity.” This activation is thought to occur as the vegetal hemisphere cortex encounters the animal hemisphere cytoplasm on the future dorsal side and the animal cortex meets the vegetal cytoplasm on the prospective ventral side. This speculation was further extended to encompass, as a subsequent effect of the activation reaction, the formation of a dorsoventral signaling center in the vegetal hemisphere (Nieuwkoop, 1969), previously known as the “Nieuwkoop organizing center.” This organizing center is generally believed to induce mesoderm formation in some of the equatorial cells of the early cleaving egg. Finally, the mesoderm is considered the source of an inductive signal to overlying cells, which causes them to acquire the features of Spermann’s primary embryonic organizer. Thus, establishment of body patterning is not a single-step process, even during its earliest phases. Rather, body patterning is the product of a series

(a) Unfertilized egg animal

(b) After fertilization

( c ) Morula

ventralwdorsal

(d) Blastula ventral ectoderm

------

]mesoderm DV marginal zone

gene is required for the cell-autonomous formation of posterior mesoderm in both mouse and zebrafish embryos. The high degree of sequence conservation represented in the Xenopus homolog (Xbra) suggests that it has the same function in amphibian embryogenesis. Smith et al. (1991) demonstrated that Xbra is expressed in the ring of involuting mesoderm during gastrulation and is also expressed in the notochord and posterior tissues at later stages. Its expression in animal caps can be induced by either FGF or activin. Xbra mRNA injection experiments demonstrated that it is able to induce formation of ventral mesodermal tissue (Cunliffe and Smith, 1992).

6. Activin Induces Expression of Dorsal Marker Genes The expression of several genes can be experimentally induced by activin, Vgl, and Wnt but not by FGF. Goosecoid (gsc) is one example. It is a homeobox gene which encodes a DNA-binding specificity similar to that of the Drosophila bicoid protein (Blumberg et al., 1991). Treatment of animal caps with high concentrations of activin generates rapid (within 30 min) expression of gsc. Its rapid appearance is not inhibited by the protein synthesis inhibitor cycloheximide, so gsc is thought of as an early response gene. Distribution of gsc mRNA in normal embryos, as revealed by in situ hybridization, closely corresponds to the primary organizer region (Cho et al., 1991b). gsc mRNA disappears shortly after gastrulation. Microinjection of gsc mRNA into ventral blastomeres leads to the formation of an additional body axis including head structures. Approximately 10% of the secondary axes are complete (Cho et al., 1991a). The progeny of goosecoid mRNA-injected cells participate in secondary axis formation. Other homologs of Drosophila homeobox genes, such as the Fork head/ HNF3-related Pintallavis/XFC-I and XFKHUXFD-I genes, are expressed rapidly upon activin treatment (Dirksen and Jamrich, 1992; Ruiz i Altaba and Jessel, 1992). In normal embryos they are localized in the dorsal blastopore lip. Pintallavis encodes a member of the HNF-3/fork head transcription factor family. It is expressed in the organizer, notochord, and in the midline neural plate cells which give rise to the floor plate. The expression of fork head genes in midline cells may contribute to the establishment of the floor plate fate.


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A transcript of the LIM class, another homeobox gene (Xfim-Z),has been discovered in the unfertilized Xenopus egg. The transcript becomes concentrated in the organizer region of gastrulae (Taira et al., 1992). Then it appears in prechordal mesoderm and notochord during gastrulation, and it slowly disappears from these tissues during the neurula stages, except for the posterior tip of the notochord, where it lingers. These observations suggest that Lim-I plays a role in early mesoderm formation and later in the specification of the differentiation pattern of the CNS. Expression of Xlim-Z mRNA is induced by either activin or retinoic acid. Treatment of animal caps with both factors gives a synergistic effect (Moriya et af., 1993; Uochi and Asashima, 1996). C. Secondary Gene Expression Responses of Dorsal Marker Genes

As verification of the gene expression cascade theme mentioned previously, several recent observations of dorsal-related gene expression sequences have been reported. For example, expression of the homeobox gene chordin begins in the organizer region subsequent to goosecoid expression (Sasai et af., 1994). The cordin gene was isolated by differential screening. Its expression can be activated by gsc and Xnot2 genes. Since its induction by activin requires de n o w protein synthesis, chordin expression is best considered to represent a secondary response. Microinjection of chordin mRNA induces a secondary axis and can completely rescue axial development in ventralized embryos. The Xenopus chordin gene has been identified as the functional homolog of Drosophifa sog, which antagonizes the effects of dpp, a homolog of the vertebrate peptide growth factor BMP-4 (Holley et al., 1995; Francois and Bier, 1995). Chordin is thought to be a potent dorsalizing factor which is expressed at the right time and in the right place to participate in organizer functions. Noggin transcripts, initially restricted to the presumptive dorsal mesoderm, reach their highest levels at the gastrula stage in the organizer region (Smith and Harland, 1992). In the neurula stage, noggin is transcribed in the notochord and prechordal mesoderm. When gastrula animal cap is placed in a high concentration of noggin protein, it acts as a neural inducer without displaying any mesoderm induction (Lamb et al., 1993; Knecht et af., 1995). Finally, we discuss one more secondarily expressed gene of interest: the Xenopus homolog of sonic hedgehog (Xhh). It is detected in both normal embryos and activin-treated animal caps (Yokota et al., 1995). It is not induced by bFGF. Sonic hedgehog is expressed in mammalian tissues with known signaling capacities, such as the notochord, floor plate of the CNS, and the zone of polarizing activity in the limb. Expression of Xhh, which

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is induced in animal caps by activin, is not inhibited by cyclohexamide. This suggests that Xhh represents an immediate early response gene. D. Later (Postgastrulation] Gene Expression Promoted by Activin

Following gastrulation several position-specific genes are expressed, probably in response to the action of an initial signal molecule such as a MIF. Activin likely initiates a cascade, which eventually includes the expression of several markers for anterodorsal and posteroventral mesoderm. XlHboxl is expressed in anterior mesoderm and neural tissues, for example, in response to activin treatment (De Robertis ef al., 1989). Likewise, Xprs-I, another homeobox gene, is induced by activin in Xenopus animal caps (Takahashi et al., 1998). Later in development additional gene transcripts can be identified. Two of the most convenient markers are the mesoderm-specific muscle actin (detected in the dorsolateral mesoderm) and blood island globin (ventral mesoderm). Activin can induce both in animal caps. Neural markers such as N-CAM are also expressed, presumably as an indirect effect of activin treatment. That is, activin probably first induces mesoderm, which in turn induces the formation of the embryonic organizer region, which then induces neural tissue in ectoderm cells. Feedback loops probably act to regulate each of the steps just mentioned. For example, Smad7, which is induced by MIFs, inhibits both activin and BMP signaling and therefore interferes with mesoderm formation and axis development. At appropriate concentrations Smad7 can act as a neural inducer (Casellas and Brivanlou, 1998). It thus appears that although researchers place their major emphasis on the discovery and study of the positive (inducing) aspects of MIF action in order to understand the molecular features of patterning in early amphibian embryogenesis, extensive parallel, concomitant negative (inhibitory) regulatory mechanisms likely play roles that are equally as important. The inhibitory Smads provide a paradigm for future research designed to elucidate the nature of what may eventually prove to be extensive networks of mechanisms which downregulate the positive aspects of MIF activities.

IV. Axial Patterning by Activin(s) Based on experiments with heterogeneous inducing factors, Toivonen and SaxCn (1955) proposed a “dual-gradient’’ model to explain how neuralizing and mesodermalizing agents act to generate axial patterning in the early


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amphibian embryo. Recently, a “three-signal’’ model has been proposed by Slack (1991). No doubt more complex versions will be developed as the regulatory circuitry becomes better understood. The models proposed to date share a common feature-concentration gradients of a limited number of signal molecules play key roles in specifying regional patterns. Experimental evidence for endorsing activin as a key player in gradient/multiple signal models has so far been compelling. Most of these models will, however, likely need at least minor modification because it has recently been discovered that superimposed on “forward-acting’’ signaling models is a set of inhibitory intermediates such as Smad proteins. These anti-Smads act to inhibit both activin and BMP signaling by binding to MIF receptors (Casellas and Brivanlou, 1998). A. Animal Cap Responses t o Various Activin Concentrations

Mesoderm patterning is widely acknowledged to be influenced by the concentration of MIF, especially in animal cap explant cultures (Green and Smith, 1990; Ariizumi et al., 1991a,b; et al., 1992). The mesoderm is more dorsal in character as concentrations of MIF are increased (Fig. 6).

FIG. 6 Spectrum of tissue types (scored by conventional histology) which are induced in animal cap explants cultured in increasing concentrations of activin (a, control, no activin; b, 0.3 ng/ml: c, 5 ng/ml: d. SO ngiml). bl, blood cells; cg. cement gland; co, coelomic epithelium; epi, epidermis: mes, mesenchyme; rnus, muscle; not. notochord.

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The studies of Ariizumi et af. (unpublished data) revealed that low concentrations of FGF (0.5-10 ng/ml) differentiate ventral mesoderm, including mesenchyme, blood, and coelomic epithelium. Higher concentrations (30-120 ng/ml) induce small pieces of muscle but no notochord. In contrast, activin induces all types of mesoderm, including muscle and notochord (Ariizumi et al., 1996). Low activin concentrations (0.1-1 ng/ml) induce ventral mesoderm tissue, whereas higher concentrations (1-10 ng/ml) induce dorsal mesoderm. Notochord, the tissue with the most dramatic dorsal character, is induced at 50 ng/ml. At this high concentration muscle formation was reduced. It therefore appears that activin can act alone, without the aid of, for example, FGF, to induce a broad range of mesodermal tissues. The three activin types (see Fig. 5) induce similar mesodermal tissues. Activin is also very likely responsible for the development of neuralinducing activity since activin does not induce neural tissue directly. Animal caps treated with activin develop organizer activity and can therefore induce neural tissue secondarily in gastrula animal caps (Ariizumi and Asashima, 1995b).

6. Toward an Understanding of Activin’s Mechanism of Action The animal cap’s responses to different concentrations of activin might depend on the proportion of the activin receptors on the surface of animal cap cells which become saturated. Longer treatments with activin can mimic the effects of shorter treatments with higher activin concentrations (Ariizumi el al., 1991b). Further insight into the mechanism of action will likely come from the use of marker genes to track activin’s effects and thereby sort out early from late responses of a tissue to activin treatment. Green et ul. (1990, 1992) obtained several insights using this approach. Animal cap cells become more sensitive to activin treatment in terms of marker gene expression when blastomeres are dissociated into single cells. Using this system they demonstrated that narrow dose ranges of activin, bounded by sharp thresholds, induce at least five different states of differentiation ranging from posterolateral mesoderm to dorsoanterior organizer mesoderm. Thus, the notion that activin acts in the normal embryo in graded concentrations is reinforced. Most likely, activin acts to specify regional mesoderm features. Following such regional specification of the mesoderm, genes which play roles in mesoderm patterning that are somewhat limited compared to the more general role just proposed for activin are expressed. Bruchyury is probably one of them. It is required for differentiation of the notochord (the archetypical dorsal mesodermal tissue) in mouse and fish embryos,


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but, even in high concentrations, injection of Xenopus brachyury mRNA does not induce notochord formation in animal caps. Xbra alone, however, is sufficient to specify ventral mesoderm in animal caps, as evidenced by the expression of Xhox3 and low levels of muscle actin (Cunliffe and Smith, 1992). It has recently been demonstrated that injection of Xbra mRNA into embryos at low doses induces animal caps to develop mesothelial smooth muscle vesicles and mesenchyme. At higher doses somitic muscle is formed (O’Reilly et al., 1995). Coexpression of Xbra with either noggin or Xwnt-8,however, does induce animal caps to develop dorsal mesodermal features. Coexpression of Xbra mRNA with noggin mRNA in animal caps yields dorsal tissues such as muscle, notochord, and neural structures. Coexpression with Xwnt-8 converted animal caps to muscle masses. Cunliffe and Smith (1994) concluded that Xbra defines a cell state in the embryo which can respond to diffusible dorsal signals such as noggin and Xwnt-8 and thereby differentiate dorsal mesodermal features. In addition, O’Reilly et al. (1995) demonstrated that coexpression of Pintallavis (but not goosecoid) with Xbra causes formation of dorsal mesoderm, including notochord. The effect of Pintallavis, like that of Xbra, is dose dependent. Thus, it appears that MIF action serves as a trigger for deployment of the complex regulatory circuits of the types just described that further refine (e.g., spatially localize) the pattern of mesoderm differentiation. The manner in which gradient components such as activin might impose pattern on early embryos is of course not well understood. It is generally assumed that diffusible morphogens activate, at various threshold concentrations, different genes. In the case of activin cell-cell contact appears to be required for a multithreshold response to occur (Green et al., 1994; Wilson and Melton, 1994). In fact, it appears that cells recognize their positions in a gradient of activin by indirect mechanisms since brief (e.g., 3 h) exposures to activin fail to show the graded response detected after a longer (e.g., 15 h) exposure. These mechanisms might involve the establishment of cell-cell contacts. There is difficulty in interpreting data derived from dissociated or cultured embryonic cells since the natural cellular adhesions and local environmental cues which are presumably so important for morphogenesis are absent. Thus, it is perhaps not surprising that a contradictory view of the activin actiodcell-cell contact issue has emerged. Gurdon and Mahony (1995) demonstrated, based on in situ analysis of blastula tissue containing activin-loaded beads, that cells respond directly to various morphogen concentrations. Their conclusion is that the response of animal cells is direct and depends on the highest concentration of MIFs to which they are exposed during their period of “competence to respond.”

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MAKOTO ASASHIMA ET AL.

Regardless of which model is correct, it is likely that eventually it will be surmised that defined concentrations of activin (e.g., see Fig. 6) induce expression of various sets of genes, a proportion of which specify mesodermal features directly and a proportion of which function to prepare the secondary signals which further refine patterning. In virtually all scenarios of the mechanism of action of activin, competence to respond and signal timing are no doubt vitally important. C. Competence [Prepatterns) and Signal (Induction] Timing

Several studies (Sokol and Melton, 1991; Bolce et al., 1992; Christian and Moon, 1993) have shown that the prospective dorsal and ventral regions of the blastula animal cap respond differently to the same concentration of activin. These observations reveal that a dorsoventral competence prepattern to activin responsiveness exists in the animal blastomeres. In order to initiate an investigation of dorsoventral prepatterning, Kinoshita et al. (1993, 1995) assayed the responsiveness to activin of Xenopus dorsal and ventral blastomeres isolated at the 8-cell stage. These embryo segments constitute a “half-animal assay” and are useful for the study of mesoderm patterning because they contain the prospective (uninduced) mesoderm region which the conventional animal cap assay lacks (Masho, 1988; Vodicka and Gerhart, 1995). The isolated half-animal explants cultured without MIF form atypical epidermis, like conventional animal caps similarly treated. This of course implies that the prospective mesoderm region has not yet received induction signals at the 8-cell stage. Even when they were exposed to activin at later times they did not respond until they reached the 32-cell stage (Kinoshita el al., 1995), which corresponds to the starting period of mesoderm induction in vivo (Nakamura and Takasaki, 1970; Jones and Woodland, 1987). When the isolated dorsal and ventral blastomeres were separately treated with activin, dorsal blastomeres differentiated both dorsal and ventral mesoderm, whereas ventral blastomeres formed only ventral mesoderm. These results indicate that a competence prepattern for responding to MIF exists as early as the eight-cell stage, Since this prepattern is absent in ultravioletirradiated embryos (Sokol and Melton, 1991), it is possible that the prepattern requires a cortical rotation (which is known to be diminished in irradiated embryos). As candidate molecules that create the prepattern, we suggest that the dorsal determinants and dorsal modifiers (described in Section II,D), which can affect mesoderm patterning but do not induce mesoderm, be considered. The prepattern, in whatever form it exists in the embryo, can be


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experimentally modified by lithium treatment in such a way that ventral blastomeres form dorsal tissues (Kinoshita et al., 1995j. This observation might prove useful for future studies directed at understanding competence prepatterning. The fact that the half-animal assay deals with uninduced presumptive mesoderm enables the examination of the early events of mesoderm patterning. Kinoshita and Asashima (1995) reported that the timing of induction affects mesoderm patterning. Half-animal embryo segments (both dorsal and ventral halves), treated with activin before the MBT, preferentially form dorsal mesoderm including notochord. However, after the MBT they differentiate mostly ventral mesoderm. Despite the low responsiveness to activin, the cleavage-stage half-animal segments preferentially expressed the dorsal marker gsc rather than the ventral marker Xwnt-8. Our supposition is that the altered response of animal cells represents not simply the loss of competence but rather the emergence of a dorsal determination process which occurs in animal blastomeres before the MBT stage. There are several reports that support this notion: Embryo dorsalization caused by lithium is lost after the MBT (Yamaguchi and Shinagawa, 1989), ventral marker Xwnt-8 mRNA acts as a dorsal determinant before the MBT (Christian et al., 1991; Smith and Harland, 1991), and induction of erythroid (ventral marker) differentiation by ventral mesoderm does not occur earlier than the MBT (Maeno et al., 1994a). An alternative view-that FGF is a competence factor for the activin signaling system-has been proposed. In studies performed by Cornell et al. (1995) various molecular markers, rather than indices of histological differentiation (as employed by Kinoshita er al., 1993,1995), were followed in vegetal hemisphere explants. Addition of FGF induced ectopic expression of the mesodermal markers Xbra and MyoD. This is due in part, at least according to these authors, to the enhancement that FGF provides to the endogenous activin levels present in vegetal cells.

V. Life History of Activin Signaling Mechanisms of the Embryo A comprehensive view of the manner in which activin functions in axis determination should include considerations of how it is delivered to the egg, how it contributes to a signaling system, and how various signaling systems are linked or networked. Beginning with its synthesis in follicle cells and accumulation in the oocyte, activin will be traced through the early embryo (Fig. 7).

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MAKOTO ASASHIMA H A L .

FIG. 7 Hypothetical scheme which describes the “life history” of activin in amphibia. It is synthesized in the ovary, testis, kidney, liver, brain, and other organs in adults (origin 2) and transported through blood vessels to oocytes, where it is complexed with the yolk protein vitellogenin (which is synthesized in the liver). Follistatin is also synthesized in reproductive organs, brain, and follicle cells. Activin is synthesized in follicle cells (origin 1) and transported into the growing oocyte. Activin is also detected in oocyte at stages 1 or 2 (origin 3). There it remains, presumably stored, until early cleavage stages, when it serves as a signal which induces animal hemisphere cells to form mesoderm. Later, neural induction, which may also involve activin (perhaps indirectly; see text) occurs. By then the body pattern is fully established.

A. Accumulation of Activin[s] as Maternal (Oocyte] Information Activin proteins as well as follistatin (its inhibitory binding protein) exist in the unfertilized egg, presumably having arrived there during oogenesis. Both activin and follistatin are synthesized in follicle cells and then accumulate in the early (stage VI) oocyte. Activin mRNAs have been detected in ovarian follicle cells (Dohrmann et al., 1993; Rebagliati and Dawid, 1993), and electron microscopic immunolabeling of gold colloidal particles (Uchiyama et al., 1994) provides evidence for the synthesis of these proteins in follicle cells. Both molecules were found to be localized in yolk platelets but not in cytoplasmic organelles. Furthermore, it appears that activin and follistatin bind preferentially to vitellogenin. Thus, it is likely that yolk platelets act as a reservoir for activin (and other MIFs?) in the early embryo. How it is released, in an active form, remains a question for further research. Most likely, a mechanism comes into play which uncouples activin from follistatin, thereby permitting activin to initiate a signaling process. Such a mechanism must be signaled since yolk platelets exist everywhere in the


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early embryo, yet the activin induction system functions in a localized (e.g., dorsal) region. Once released in an active form, activin is then conceptualized as interacting with its protein receptor.

B. Activin Receptors and Signal Transduction The prospective geographical location of MIF receptors was first examined in urodele (newt) gastrula animal caps (Asashima and Grunz, 1983). The newt gastrula ectoderm consists of two cell layers, which can be conveniently separated.The outer layer has a lower competence to form dorsal mesoderm structures compared to the inner layer. When monitored with activin AI”’, receptors were found to be localized on the surface of inner-layer cells (M. Asashima, unpublished data). Receptors are therefore nonuniformly distributed in the embryo. This finding is consistent with notions about the regionalization of competence described previously. Substantial information about the receptors for members of the TGF-@ family exists. They represent a heteromeric complex composed of two types of transmembrane serinehhreonine kinases (types I and 11) (Kingsley, 1994). Type I1 receptor is thought to be responsible for initially binding the ligand. Then a type I receptor is recruited and subsequently phosphorylated by the type I1 receptor (Wrana et al,, 1994a). The phosphorylated type I receptor in turn phosphorylates intracellular substrates and in the process initiates a signal cascade. A number of type I and I1 receptors have been shown to exist in vertebrates, including receptors for activin, TGF@, and BMP. Two types of activin receptors are known in Xenopus (XactRs): type I (ALK-2 and ALK-4; Armes and Smith, 1997) and type I1 (Kondo et al., 1991; Mathews et al., 1992; New et al., 1997). In addition, various subtypes of activin receptors are known and likely generate different downstream gene expression patterns, as suggested by New et af. (1997). The Xenopus BMP-4 receptor is type I (Graff et al., 1994), as is another TGF@-relatedreceptor (Mahony and Gurdon, 1995). Overexpression studies with activin receptors in Xenopus embryos indicate that they are indeed involved in transmission of the activin signal. XactRZZB, cloned by Mathews et al. (1992), is a homolog of mouse ActRZZB. When XactRZZB mRNA was injected into the early embryo’s ventral blastomeres, a secondary body axis was induced. X A R l , which is highly homologous to AactRZZB (Hemmati-Brivanlou and Melton, 1992), is expressed continuously in the ovary, unfertilized egg, and neurula embryo. Maternal XactRZZB mRNA is found uniformly in the early embryo but is restricted in distribution to the neural plate at the neurula stage. These observations imply that activins play roles later in neurogenesis as well as earlier in mesodermal induction.

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C. A Conflicting Opinion on Activin’s Role

An opinion which contrasts sharply with the view that activin/activin receptors play key regulatory roles in mesoderm/neural patterning has, however, recently been offered. It has been suggested that activin signaling may be unnecessary for mesoderm formation. Both Schulte-Merker et al. (1994) and Kessler and Melton (1995) have demonstrated that truncated versions of Xenopus activin receptors also interfere with the activity of mature Vgl protein. This observation presents the possibility that such modified receptors may block the function of several members of the TGF-fi family and not just activin. Thus, it is possible that the effects of truncated activin receptors on Xenopus development can be explained not by activin inhibition but rather by inhibition of Vgl signaling. However, swinging the pendulum back toward activin as the dominant PGF signal, Dyson and Gurdon (1997) prepared an activin receptor construct which selectively blocks activin but not Vgl action. They claim it inhibits mesoderm differentiation. Clearly, the use of defective receptors as inhibitors for amphibian embryonic induction research requires further study, especially in view of the fact mentioned previously that numerous closely related and cross-reacting receptors likely exist. In the mouse, however, gene targeting recently has been used to inactivate a type I activin receptor. Embryos failed to gastrulate properly, providing genetic evidence for a role for activin in early embryonic patterning (Gu et al., 1998). Whether extrapolation to amphibia is warranted is a matter of speculation. It should be noted that follistatin (an activin-binding protein) mRNA injection fails to block mesoderm induction in the whole, intact embryo (follistatin does not inhibit Vgl signaling). A lack of a follistatin effect, although it represents “negative evidence,” argues against a function for activin in mesoderm induction. Finally, it should be mentioned that if the “truncated receptor” data are to be heavily weighed, so as to discount the role of activin in normal mesoderm development, more compelling data for assigning to Vgl the role previously ascribed to activin should be generated. After all, as mentioned earlier, arguments promoting Vgl as a natural mesoderm inducer have several shortcomings.

D. Networks and Cascades of Inductive Signals Various studies have suggested that natural induction and patterning processes of mesoderm development involve a complex system involving multiple signal molecules. For example, mesoderm induction by activin (Cornell and Kimelman, 1994; LaBonne and Whitman, 1994) and processed Vgl


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(Schulte-Merker et al., 1994) is facilitated by FGF-mediated signals. In several studies it has been shown that a number of the components of the FGF signal transduction pathway are important for activin signaling in the early embryo (LaBonne and Whitman, 1994;MacNicol et al., 1993; Whitman and Melton, 1992). For example, activation of the tyrosine kinase FGF receptor appears to activate the small GTPase p2lras, which in turn activates the cytosolic kinases raf-Z and mitogen-activated protein (MAP) kinase. As with the dominant inhibitory F G F receptor, dominant inhibitory mutants of raf and ras inhibit mesoderm induction by both FGF and activin (LaBonne and Whitman, 1994; Whitman and Melton, 1992). Despite the apparent need €or a functional FGF signaling pathway for activin induction, activin does not cause a significant activation of MAP kinase or p2lras (Graves et al., 1994); LaBonne and Whitman (1994) indicated that activin does not directly stimulate this pathway. These data suggest that early signaling responses can be used to distinguish pathways in vivo and thus help define the roles played by individual factors or families of factors. Recent studies using MAP kinase-specific phosphatases (LaBonne et al., 1995; Gotoh et al., 1995; Umbhauer et al., 1995) demonstrated that MAP kinase activation is required for mesoderm induction by FGF and activin. In fact, MAP kinase is sufficient for the induction of mesodermal markers (Umbhauer et af., 1995). Presumably, MAP kinase acts downstream of the initial signaling caused by FGF or activin. As more signaling molecules are identified, mesoderm induction is increasingly being viewed as an intrinsically complex phenomenon. The following are recent additional entries in the mesoderm-induction circuit “derby.” The Xenopus homolog of the mouse nodal gene ( X n r s ) has been reported to act in mesodermal patterning (Smith et al., 1995; Jones et al., 1995). Nodal is a diverged member of the TGF-P superfamily gene and is expressed in the mouse node during gastrulation. Xnrs appears transiently during embryogenesis and is specifically expressed in the Spemann organizer at the early gastrula stage. When X n r mRNAs were injected into early embryos, they rescued the embryonic axis in ultraviolet-ventralized embryos, dorsalized ventral marginal-zone explants, and induced muscle differentiation.Although no maternal Xnr mRNAs were found, the activities of the X n r proteins suggest that a signaling pathway involving nodalrelated peptides is an essential element in mesoderm differentiation and axial patterning. It has been discovered that a TGF-P-related type I receptor, XTrR-1, (Mahony and Gurdon, 1995) is expressed in all regions of embryos throughout early development. Overexpression of this receptor does not affect ectoderm or endoderm but dorsalized the mesoderm such that muscle appears in ventral mesoderm and notochord appears in lateral mesoderm

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normally fated to become muscle. In addition, overexpression of XTrR-1 in irradiated embryos is able to cause formation of a partial dorsal axis. These results suggest that XTrR-1 encodes a receptor which responds in normal development to a TGF-like ligand so as to promote dorsalization. Its function would therefore be to direct mesodermalized tissue into muscle or notochord. There is at least one more interesting gene which is known to be expressed in the embryo’s dorsovegetal region-the homeobox-containing gene Siarnois. Its RNA is first detected shortly after the midblastula transition, earlier than mRNAs for goosegoid or Xbra, and is present most abundantly in the dorsal endoderm of early gastrulae (Lemaire et af., 1995). Embryos injected with it in a ventral-vegetal blastomere develop a complete secondary axis. The progeny of Siarnois-injected cells do not participate in secondary axis formation. It would be easy to imagine Siarnois as playing a role in the formation of the Nieuwkoop center.

E. Inhibition of Neural Induction by Activin? Although activins induce neural tissue in animal caps, they do not act directly as a neural inducer. Microinjection of low doses of activin A protein directly into the blastocoel (see Fig. 2b) generates a secondary embryo. High doses yield a headless embryo with an outgrowth of tail (Ariizumi et al., 1991b). These observations hint that activin might interfere with the regulatory circuitry which comprises normal neural induction. Indeed, expression of a dominant inhibitory form of the activin type I1 receptor, which blocks mesoderm-inducing activity of activin, induces the formation of anterior neural tissue in animal cap explants (Hemmati-Brivanlou and Melton, 1994). Also, overexpression of this truncated receptor in ultravioletirradiated embryos yielded embryos with neural structures but that lacked major axial structures. In addition, expression of follistatin, a specific inhibitor of activin, in animal caps results in expression of neural markers (Hemmati-Brivanlou et al., 1994). Since Xenopus follistatin is expressed first in the gastrula organizer and later in the notochord and anterior nervous system, these localizations coincide with the tissues which would be expected to produce neural inducing factors. Thus, activin’s role in normal embryogenesis might extend beyond early dorsal mesoderm induction to include later inhibitory effects on neural induction. A key to understanding these different roles might lie in the mechanisms which regulate the endogenous levels of activin protein at various developmental stages. Accordingly, Klein and Melton (1995) investigated the translational control of activin in Xenopus embryos. They conclude that maternal

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factors regulate translation of activin mRNA by interacting with the 3‘untranslated region of the mRNA.

VI. Proposed Molecular Models for Activin’s Role in Signal Transduction Pathways As mentioned previously, the lack of a systematic approach (e.g., genetics) to elucidate the regulatory circuits in which activin is involved generates models which are likely to be incomplete. Nevertheless, such models are usually very valuable because they provide testable features and can be used as a point of departure for further studies. A few are illustrated in Fig. 8. They should not activin Smad2

activin

$.(+)

FAST-1

actkiin signal; Mix.2 expression type I1

kinase

signal transduction expression

activin

activin signal FIG. 8 Various ways of viewing the role of activin in signal transduction pathways. (Top) Two membrane receptors (types I and 11) are required for activin action. They each have a cytoplasmic domain which acts as a serinelthreonine kinase. The type I receptor becomes phosphorylated and is subsequently responsible for propagating the signal downstream [see Chen et al. (1997) for more complex versions of this model]. (Middle) The expression of the goosecoid (gsc) gene is initiated by the action of both activin and Wnf. The product of the gsc gene then triggers further steps in the gene expression cascade, which ultimately leads to development of Spemann’s primary organizer on the embryo’s dorsal side. (Bottom) BMP is employed as a negative regulator of activin action.

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be considered as competing models. Rather, they represent models which have been developed by using different databases, and consequently they depict different views of activin’s role in signal pathways. One model, based on the data of Chen et al. (1996) and Wrana et al. (1994a,b), proposes that activin interacts with its type I1 receptor, which in turn phosphorylates the activin type I receptor. This subsequently leads to phosphorylation of a cytoplasmic protein (Smad2) (Lo et al., 1998). It is translocated to the nucleus, where it interacts with a transcription factor (FAST-1) and binds to a promoter region [the so-called activin response element (ARE), a 290-base pair sequence], leading to transcription of the Mix.2 gene. M h . 2 is a homeobox gene which presumably plays a role in initiating a gene expression cascade. Thus, in this model activin is a key component in upregulating gene expression by virtue of its ability to activate a set of “transcriptional partners.” These transcriptional partners-when present in sufficiently high concentrations-may also act in a negative feedback loop to control activin action (Nakao et aL, 1997). Also, additional coactivators (CBP and p300) have recently been discovered to be involved in a cooperative mechanism which promotes Smad2 activation (Janknecht et al., 1998). Similar models which link activin and Smad2 are reported to be active in patterning the early mammalian (mouse) embryo in ways which mimic those described previously for amphibian embryos (Nomura and Li, 1998; Zhao et al., 1998). Another model emphasizes the manner in which activin and other peptide growth factors induce gsc gene expression early in a signaling pathway (Watabe et al., 1995). It proposes that the gsc promoter has a distal element (DE) and a proximal element (PE) which respond to activin (DE) and Wnt (PE) action. Simultaneous but independent interactions at both these promoter sequences is believed to be required for the guusecuid gene expression. A third model depicted in Fig. 8 emphasizes the role activin plays in promoting the expression of the early response gene X F K H l (Kaufman et al., 1996). The expression of this gene is postulated to comprise part of the Spemann primary organizer. An A R E located in the promoter region of the X F K H l gene is depicted as responding to activin. In addition, a second promoter region [BIE (BMP-activated)] in this gene is thought to respond to BMP signaling and override the activity of the activin-ARE system. Thus, both positive and negative regulation are included in this model.

VII. Activin Causes a Broad Array of Differentiations in Vitm One of the most remarkable features of activin is its ability to generate a diverse array of tissue differentiations in animal cap explant cultures when


39

applied at various doses (see Fig. 3). Its ability to induce the differentiation of such a broad spectrum of tissue differentiations provides one of the most compelling arguments, when viewed in terms of the rigorous criteria (that it largely fulfills) discussed earlier, for considering it to be an endogenous inducer. A few examples of these differentiations are reviewed in the following sections. A. Induction of Endoderm by Activin

The group of MIFs from heterogeneous tissues (e.g., chick embryo extract) have been termed vegetalizing factors based on their ability to induce endoderm and mesoderm differentiation in newt animal cap explants (Grunz, 1979; Grunz and Tacke, 1989; Kocher-Becker and Tiedemann, 1971). At low concentrations they typically induce mesoderm, whereas at high concentrations they preferentially induce endodermal differentiations. When a pellet of vegetalizing factor is implanted into the blastocoel, a secondary axis is induced. Thus, these vegetalizing factors mimic the action of the inducing signal from the Nieuwkoop center. Do activins possess the functions expected of the Nieuwkoop center? Activin induces the expression of an endoderm marker gene (Mix. I ) and antigens recognized by endodermal-specific antibodies (Jones et al., 1993). Also, high concentrations of activin (100 ng/ml) can induce histological differentiation of endoderm in newt animal caps (Ariizumi and Asashima, 199%). For example, endodermal epithelia and, occasionally, a beating heart develop. Heart induction has been shown to require signals derived from both endoderm and organizer tissue in Xenopus ventralmesoderm explants (Nascone and Mercola, 1995), so a role for activin is very plausible. Furthermore, activin-treated newt animal caps can induce mesoderm in uninduced animal caps (Ariizumi and Asashima, 199%). We can therefore propose the following model for these in vitro results. Activins act strongly on a portion of the animal cap to induce endoderm. This endoderm in turn induces mesoderm in the rest of the (uninduced) animal cap. This model is clearly compatible with activin serving as a key component in the embryo’s natural Nieuwkoop center.

6. Activin-Induced Organizer Guides Neural Patterning The vertical signal of neural induction implies that the involuting dorsal mesoderm signals the overlying ectoderm to become neural tissue. The anteroposterior pattern in neural structures is established during this period,

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presumably as a result of patterned induction from mesoderm. The results of an organizer graft (Spemann and Mangold, 1924; see Fig. 2b) can be mimicked by the action of activin. Ariizumi and Asashima (1995b) treated newt animal caps with the high concentrations of activin (100 ng/ml) mentioned previously and then kept them in saline for different periods. They were later combined with uninduced ectoderm, and subsequent differentiation was monitored. Animal caps incubated for a brief period before being combined with ectoderm induced posterior (trunkhail) structures, whereas those precultured for a long time induced anterior (head) structures. Remarkably, these data correlate well with the original Spemann and Mangold (1924) observation that the early gastrula blastopore lip induces head structures and the late gastrula lip induces trunkhail structures. At the very least, these findings should offer an experimental model system for in vitro studies on the molecular mechanisms of primary embryonic induction. The activin-treated tissues represent a much simpler system than the whole, intact embryo.

VIII. Conclusion A variety of MIFs can be demonstrated to have profound effects on body patterning when either administered to whole embryos or added to the culture medium of explanted embryonic tissue. When the life histories of

FIG. 9 Activin’s role in early embryonic patterning events is depicted as being at the center of the regulatory circuits which regulate the development of various individual tissues and organs.


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several MIFs, including FGF, Vgl, and activin, are carefully reviewed, activin stands out as having the most profound effects and fulfilling most of the rigorous criteria expected of a natural (endogenous) inducer. It is of course unlikely that any single MIF-in the normal, intact embryocontrols all early programming events and differentiations. Nevertheless, activin is likely to play the key, early role. Figure 9 positions activin at the epicenter of the profusion of circuits and pathways which comprise early amphibian embryonic patterning. Activin has also been identified as playing a key role in various cell differentiation events which are separate from its role in embryogenesis, the subject of this review. In some regards, it might be considered an all-

FIG. 10 The life history of an amphibian is illustrated. Many of its most dramatic morphogenetic events are shown. The time periods during which activin is active are diagrammed as bellshaped curves around the life cycle circle. Activin involvement in developmental life cycle events “comes and goes” from oogenesis through adulthood. In this sense, activin can be conceptualized as serving as a “timekeeper” gene since it appears to play a regulatory role in each major morphogenetic transition.

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MAKOTO ASASHIMA

ET AL.

purpose signaling molecule. At the cellular level it is known to induce apoptosis in B lineage cells (Nishihara et al., 1995) and to facilitate wound repair (Hubner et al., 1996). At the subcellular level is can affect Kf and Ca2+ channels (Mogami et al., 1995) and promote insulin secretion (Furukawa et al., 1995). At the molecular level it is known to cause alterations in transcription factor (e.g., Pit-1) activity (Gaddy-Kurten and Vale, 1995). Figure 10 conceptualizes the life cycle of the vertebrate (e.g., amphibian) and illustrates the various stages during which activin is known to play roles in one or another pathway or regulatory circuit. It should therefore be presumed that studies on the role that activin plays in specifying embryonic patterns (e.g., via dorsal mesoderm induction) stand to benefit from the data being collected in subdisciplines which do not overlap with embryology/developmental biology. By collecting information from a broader range of activin discoveries, developmental biology researchers might be able to make better use of intuition for deducing the signaling pathways and regulatory circuits used in amphibian embryos. Thereby, more freedom from the current heavy dependence on Drosophila studies, which lead to an invertebrate bias in the development of hypothetical models, could be achieved.

Acknowledgments M. A., K. K., and T. A. acknowledge the financial assistance provided by a grant-in-aid from the Ministry of Education, Science and Culture of Japan and CREST (Core Research for Evolution Science and Technology) of the Japan Science and Technology Corporation. G. M. M. acknowledges the National Science Foundation for support of the Indiana University axolotl colony. Susan Duhon provided expert editing of the manuscript.

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Calcium Regulation of the Actin-Myosin Interaction of Physarum polycephalum Akio Nakamura and Kazuhiro Kohama Department of Pharmacology, Gunma University School of Medicine, Maebashi, Gunma 371-8511, Japan

Plasmodia of Physarum polycephalum show vigorous cytoplasmic streaming, the motive force of which is supported by the actin-myosin interaction. Calcium is not required for the interaction but inhibits it. This calcium inhibition, a regulatory mode first discovered in Physarum, is the overwhelming mode of regulation of cytoplasmic streaming of plant cells and lower eukaryotes, and it is diametrically opposite to calcium activation of the interaction found in muscle and nonmuscle cells of the animal kingdom. Myosin, myosin I I in myosin superfamily, is the most important protein for Ca2+action. Its essential light chain, called calcium-binding light chain, is the sole protein that binds Ca2'. Although phosphorylation and dephosphorylation of myosin modify its properties, regulation of physiological significance is shown to be Ca-binding to myosin. The actin-binding protein of Physarum amplifies calcium inhibition when Ca2' binds to calmodulin and other calcium-binding proteins. This review also includes characterization of this and other calcium-binding proteins of Physarum. KEY WORDS: Physarum polycephalum, Cytoplasmic streaming, Actin, Myosin, Actin-myosin interaction, Calcium ion, Phosphorylation, Inhibitory effect of Ca2'. 0 1999 Academic Press.

1. Introduction Concentration of intracellular Ca2+is usually kept as low as possible by extrusion through the cell membrane and by sequestration into the endoIiirernononol Review of Cviology. Vol. I Y I

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plasmic reticulum. When the cell is excited, the concentration is increased by the entry of extracellular Ca2’ and by the release of sequestered Ca2t. However, the increase is transient because Ca2+concentration ([Ca2+])will soon be reduced again. The cell utilizes such a transient increase in [Ca”] as a second messenger to regulate various intracellular reactions. Most examples of calcium regulation observed involve activation, as exemplified by muscle contraction (Ebashi and Endo 1968): Muscle is relaxed in low [Ca”], and the elevation of [Ca”] leads to the interaction of myosin with actin to induce contraction. With the subsequent decrease in Ca”, the interact is abolished to cause relaxation. An alternative mode of regulation is theoretically possible: The interaction proceeds without requiring Ca”. An increase in [Ca”] inhibits the interaction, which will be relieved when [Ca”] returns to low levels. Such a mode has been indicated for the actomyosin system of the lower eukaryote, Physarum polycephalum (Kohama et al., 1980). Physarum plasmodium has attracted the interests of scientists in the field of cell motility because of its vigorous cytoplasmic streaming. The streaming is induced as a result of the contraction of plasmodium (Kamiya, 1981). Therefore, quantitative evaluation of the motile event in plasmodium is possible by measuring the contraction by physiological methods. Biochemical methods to purify proteins are also applicable because Physarum can be cultured in a large quantity despite the cells being eukaryotic (Kohama et al., 1998). This feature enables actin, myosin, and actomyosin-related proteins to be purified by modifying the procedures for muscle proteins (Hatano, 1973). In this review, we will explain how the interaction between actin and myosin is inhibited by Ca”. The regulation by Ca2+ of Physarum has been reviewed in detail by Kohama (1987) and subsequent reviews have appeared (Kohama, 1990; Kohama et al., 1992, 1993; Nakamura and Kohama, 1995), but these do not include the results obtained by up-to-date methods, which will be included in this review. Actin and myosin are believed to be present in all eukaryotic cells and take part in their motile events (Citi and Kendrick-Jones, 1987). Typical examples are muscle contraction in animal cells and cytoplasmic streaming in plant cells. Muscle contraction occurs at high [Ca”] and cytoplasmic streaming at low [Ca”.]. Therefore, it is expected that the effect of Ca2+ on the actin-myosin interaction is diametrically different between animals and plants (Kohama, 1990), i.e., CaZt works as an activating signal for contraction but as an inhibition one for streaming (Table I). This review also addresses an inhibitory mode of calcium regulation: calcium inhibition of the actin-myosin interaction of plant and other cells.

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TABLE I Regulation of Actin-Myosin Interaction by Ca2+

Cells Plant Animal

Resting (Ca2+< phf)

Excited (Ca2+< phf)

+

-

-

+

Note. +, active interaction between actin and myosin; -, no active interaction. Ca2+concentrations in the cells of both kingdoms are increased transiently when they are excited. The interaction of Physarurn can be classified as plant type.

II. Calcium Inhibition of Motile Events Related to Actomyosin A. Calcium Inhibition of Cytoplasmic Streaming When plant cells are observed under a microscope, organelles move along actin cables that run beneath the cell membrane. The internodal cells of Chara and Niteffa are popular materials for the observation of this cytoplasmic streaming. When they are excited by the electrical stimulation, their cytoplasmic streaming is abolished transiently (Kishimoto and Akahori, 1959; Tazawa and Kishimoto, 19681, an observation that is associated with an increase in the intracellular concentration of Ca2+(Kikuyama and Tazawa, 1982; Williamson and Ashley, 1982). The inhibitory role of Ca2" was confirmed by perfusing artificial protoplasm containing various concentrations of Ca2+into the cells and by destroying the plasma membrane to soak the cytoplasm in various concentrations of Ca2+ (Williamson, 1975; Shimmen and Yano, 1984; Tominaga et af., 1983). A similar inhibitory effect of Ca2+is demonstrated with Vaflisneria cells; its cytoplasmic streaming is detectable when observed under red light and is abolished by the application of infrared light (Takagi and Nagai, 1985, 1986). Red light induced efflux of Ca2+to, and infrared light influx of Ca2+from, the medium outside the cell (Takagi, 1993). It has been suggested for some time that the organelles of the internodal cells are associated with myosin, which interacts with actin cable to causes cytoplasmic streaming. The myosin is expected to be subjected to calcium inhibition. However, myosin (or myosin-like protein) purified from Chara is not sensitive to Ca2+(Yamamoto et af., 1995). In the myosin superfamily, conventional myosin corresponds to myosin 11. Currently, the superfamily

'

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consists of more than 10 varieties (Mooseker and Cheney, 1995). However, biochemical characterization as a motor protein has been carried out only with myosins I, 11, V, and VI. This suggestion was recently confirmed by purifying myosin-like protein from the pollen tube of Lilies, in which cytoplasmic streaming has been demonstrated to be abolished by an increase in the cystoplasmic [Ca2’] (Kohno and Shimmen, 1988). The major component of the protein is a 170-kDa peptide whose actin-myosin interaction observed in vitro was inhibited by Cazt (Yokota and Shimmen, 1993). A myosin-like protein susceptible to calcium inhibition has also been purified from BY-2, a cell line established from tobacco (Yukawa et al., 1997).

B. Calcium Inhibition of the Cytoplasmic Streaming of Plasmodia of Physarum polycephalum Cytoplasmic streaming of plasmodia is observed as a vigorous shuttle movement of organelles. Unlike in plant cells, plasmodia1 actin does not form actin cables but rather forms a complex with myosin to build up “vessels,” in which cytoplasm streams passively due to the contraction of the actomyosin vessel wall (Komiya, 1981). The concentration of cytoplasmic Ca2’ changes within pM levels, as can be observed by injecting aequorin into plasmodia as a Ca2’ probe (Ridgway and Durham, 1976). When plasmodia are soaked in a solution containing caffeine, tiny drops of cytoplasm are released from plasmodia. Hatano (1970) observed the movement of organelles in these caffeine drops and found that an increase in [Ca2’] in the solution is associated with an increase in movement. A similar claim was also published in the 1970s (Ueda et al., 1978) in analogy to the activation of muscle contraction by Ca2’ (Ebashi and Endo, 1968). It must be noted that the activating effect of Ca2t is based on observations with models for cytoplasmic streaming that are furnished with intact cell membranes. The concentrations of Ca2+ are not necessarily identical between the outside and inside of the membrane. Yoshimoto et al. (1981a) and Yoshimoto and Kamiya (1984) permeabilized the membrane of a plasmodial strand to alter the cytoplasmic concentration of Ca2’ directly and related its contraction to the concentration of Ca2+.They concluded that the actin-myosin interaction, as measured by the contraction, is maximal in the absence of Ca2’ and reduced with an increase in the Ca” concentration. This inhibitory effect of Ca2+ is confirmed with similar cell-free models under various concentrations of cytoplasmic Ca2’ (Achenbach and Wohlfarth-Bottermann, 1986a,b; Pies and Wohlfarth-Bottermann, 1986) (Table 11).

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TABLE I1 Calcium Inhibition of Actin-Myosin Interaction of Physarum Plasmodium Actin-activated ATPase activity of myosin" Superprecipitation of actomyosin* In vitro motility assay Nire//a-bassl motility assay" Myosin-coated surface assay" Tension development of actomyosin threads" Contraction of cell-free model' Intact cellR ~

~~

Ogihara er a/. (1983). Kohama and Kendrick-Jones (1986). Kohama ef al. (1991a). and Ishikawa e t a / . (1991). " Kohama et a/. (1980) and Ogihara et a/. (1983). ' Kohama and Shinimen (1985). "Okagaki et al. (1989). '' Sugino and Matsumura (1983). 'Yoshimoto et rr/. (1981a). Yoshimoto and Kamiya (1984), Achenbach and Wohlfarth-Bottermann (1986a,b), and Pies and Wohfarth-Bottermann (1986). fi Ishigami ef al. (1996). o

It has been proposed that the cytoplasmic concentration of Ca2+might not be identical throughout the plasmodium (Kuroda et al., 1988). Recent technical advances have made it possible for this argument to be resolved. Using a small fragment of plasmodium that shows contraction-relaxation movement rather than the cytoplasmic streaming, Ishigami et al. (1996) were able to relate this movement to the changes in cytoplasmic concentration of Ca2+.They concluded initially that the increase in local [Ca2+]acts as a trigger to make the plasmodium relax (Table 11).

C. Calcium Inhibition of Motile Events in Vertebrate and Invertebrate Cells Ca" activates the interaction between thick and thin filaments of muscle cells. However, the effect of Ca2+on the cleavage furrow, which is one of the typical structures that are related to actomyosin system, is diametrically different. Yoshimoto and Hiramoto (1985) related changes in Ca2+levels of echinoderm and medaka eggs to the cleavage cycle. They found that the construction and destruction of the cleavage furrow were respectively associated with a decrease and increase of Ca" concentration and suggested that the actin-myosin interaction of the furrow is subject to calcium inhibi-

58


tion. This suggestion conforms with their further observation using demembranated cell models of the eggs that a [Ca”] higher than 0.1 p M inhibited construction of the furrow (Yoshimoto et al., 1985). The inhibitory effect of Ca2+ reported for contractile rings isolated from newt eggs was such that their ATP-dependent contraction was inhibited by p M levels of Ca2+ (Mabuchi et al., 1988). The inhibitory effect of Ca2’ has been documented (Kohama, 1993). Examples are accumulating not only for motility-related events [for sperm motility, see Hong et af. (1991); for secretion of renin and parathyroid hormone, see Churchill (1985) and Shoback et af. 1983, respectively] but also for various other phenomena [for adenylate kinase of cardiac muscle, see Colvin et al. (1991); for adenylyl cyclase, see Nakamura and Bannai (1997); for NO synthetase, see Mittal and Jadhav (1994); for rhodopsin kinase, see Kawamura et af. (1993); for thioredoxin reductase, see Schallreuter and Wood (1989); for regucalcin of liver, see Simokawa and Y amaguchi (1993)]. Recently, myosin-like proteins, which have calmodulin as their light chain, have been purified from vertebrate nonmuscle cells. Myosin I from brush border consists of a 119-kDa heavy chain and calmodulin. Ca2+ dissociates calmodulin from the heavy chain and prevents the interaction of myosin I with actin (Collins et aL, 1990; Swanljung-Collins and Collins, 1991). Myosin V, a myosin-like protein from brain, is another calmodulincontaining protein. The effect of Ca2+as monitored by the in vitro motility assay is inhibitory (Cheney et af., 1993). However, the actin-activated ATPase activity of the protein is stimulated by Ca2+,a discrepancy that must be solved in the near future. Some of vertebrate smooth muscles are known to maintain their tone even if they are immersed in a solution containing EGTA for a few hours. Upon returning to ordinary Ca-containing solution, they relaxed with an increase in cytoplasmic [Ca”] (Sasaki and Uchida, 1980). We speculate that acalmoddin-containing myosin is involved in the contraction in these conditions because myosin I is present in smooth muscles (Kohama et d.,1991c; Chacko et al., 1994;Hasegawa et af., 1996). Similar speculation is possible for the type of myosin associating with the cleavage furrow and contractile ring mentioned previously: It might be a calmodulin-containing myosin.

111. Calcium Inhibition of the Actin-Myosin Interaction of Physarum as Detected in Vitro A. Studies with a Crude Actomyosin Preparation

The effect of Ca2+ on actin-myosin interaction of Physarum could be examined in vitro if one prepared crude actomyosin from plasmodia. Early


59

examination of this preparation indicated that Ca2+stimulated the interaction in a similar way to the actomyosin prepared from vertebrate muscle (Nachmias and Asch, 1974; Kato and Tonomura, 1975). Concerning the movement of amebae, Taylor et al. (1973) prepared a cell-free model of Chaos charolinesis; the model contracted in a Ca2+-containing solution and relaxed in the presence of EGTA. Myosin, whose actin-activated ATPase activity is stimulated by Ca2+,has been purified from Acanthamoeba (Collins and Korn, 1981). On the other hand, the actin-activated ATPase activity of myosin purified from amebae of Physarum (see Section VIII) was inhibited by Ca2+ essentially in the same way as that of plasmodia1 myosin (Kohama et al., 1986a). Inhibition by Ca2+of the contraction of a cell-free model of Amoeba proteus (Sonobe et al., 1985) is in good accordance with Physarum myosins. We have also observed a marked elevation of the ATPase activity of the crude preparation (Table 111; Fig. 1). However, when the preparation was subjected to further purification, the effect of Ca2+ on the activity was eventually reversed, with the ATPase activity of purified native actomyosin decreasing with an increase in [Ca”]. Because a crude extract of plasmodia is abundant in calcium-activated soluble ATPase(s) (Table 111), and contains an ATP pyrophosphohydrolase (Kawamura and Nagano, 1975), the apparent activation by Ca2+of the crude preparation was due to contamination by the soluble ATPase(s). Accordingly, we have reached the conclusion that Ca2+controls the actin-myosin interaction by inhibiting the ATPase activity of actomyosin. This inhibition is also detected by inducing the

TABLE 111 Soluble and Actomyosin ATPase Activities of Physarum Plasmodia

Condition

ATPase activity (nmol/min/mg protein)

Supernatant (0.1 mM EGTA, 50 yM Ca2’) Native (0.1 mM EGTA) Actomyosin (50 yM Ca”)

11.2 57.2 15.7 4.7

~

~~

Total ATPase activity (nmol/min/100 g plasmodia)

3.1 x 16.0 x 0.14 x 0.04 x

104 104 104 104

~~

Nore. Fresh plasmoida of Physarum polycephalum were harvested, homogenized in high salt (0.5 M NaC1, 10 mM EGTA, 0.1 mM D’IT, and protease inhibitors) at pH 8.4, and centrifuged at 100,000g for 1 h. The supernatant, adjusted to pH 6.5, was dialyzed against 5 vol of water, and centrifuged at 10,OOOg for 20 min. The supernatant was then assayed for ATPase activity. The precipitate was dissolved in high salt and used as crude native actomyosin. From this, native actomyosin was purified by repeated washing. ATPase activities were determined by the pH-stat method in 0.5 mM Mg-ATP, 1.0 mM Mg2+,30 mM KCI, and 0.1 mM EGTA-Ca buffer at pH 7.50 and 25°C (Kohama and Ebashi, 1986).

60


v

I

I

I

I

6

5

4

3

FIG. 1 Relative NTPase activity of crude native actomyosin as a function of [CaZt] (Kohama and Kendrick-Jones, 1986). Mg-NTPase activities of crude native actomyosin were determined at various concentrations of CaZt.The activities were determined by the pH-stat method under 5 mM Na+/K+,5 mM Mg acetate, 1.5 mM NTP, and 0.1 rnM EGTA-Ca buffer at pH 7.50 and at 25°C. In the presence of EGTA (loo%), the ATPase (O), UTPase (A) and GTPase (0)activities were 7.2,15.8, and 32.3 nmol/min/mg actomyosin, respectively. Calcium inhibition of actomyosin i n crude actomyosin preparation can be monitored by UTPase and GTPase activities without complicating the Ca-activatable soluble ATPase(s) contaminating the preparation (see Table 11). The difference between the Ca2+concentration that causes half-maximal inhibition for UTPase and GTPase activities can he attributed to the actomyosin properties, because a similar difference can be detected with actomyosin reconstituted from skeletal muscle actin and Physnrurn myosin (see Fig. 6 in Kohama and Kendrick-Jones, 1986).

superprecipitation as shown in Fig. 2 (Kohama et al., 1980; Ogihara et aZ., 1983) (Table 11). It should be noted that the crude preparation has been prepared as quickly as possible to avoid possible artificial modifications (see legend to Fig. 1). Therefore, actomyosin in this crude preparation is as close as possible to in vivo actomyosin. The calcium inhibition can be detected by


61

A660

*Y . c) I

0

1

3

2

Time(min)

I

6

4

pCa2+ FIG. 2 Inhibitory effect of Ca” on superprecipitation of purified preparation of native actomyosin from Physunrrn plasmodia. The extent (relative value) of superprecipitation at various concentrations of CaZ-was plotted against &a2* (Kohama e t a / . . 1980). Inset shows the time course of superprecipitation after the addition of ATP in the presence of 0.1 mM EGTA (G) or 10 p M Ca” (Ca). and the increase (relative value) in the absorbance at 660 nm in 3 min was plotted against pCa2+.

measuring UTPase and GTPase activities of such an actomyosin (Fig. 1) by utilizing the property of the Ca-activated ATP pyrophosphohydrolase mentioned previously-that it hydrolyzes UTP and GTP very slowly.

B. Myosin Is the Site of Action of Ca2’: Myosin-Linked Nature of Calcium Inhibition Observed with Crude Actomyosin from Physarurn As the first step in identifying the site of action of Ca”, we purified myosin and actin from the actomyosin preparation that shows calcium inhibition (Kohama and Kendrick-Jones, 1986).

62


1. Purification of Myosin The actomyosin preparation was dissolved in 20 mM ATP containing DTT, mixed with concentrated Mg acetate to give 0.1 M in the final concentration, and centrifuged at 100,OOOg for 30 min. The supernatant was diluted with cold water by 2.5- to 3.0-fold and allowed to stand for 3 or 4 h on ice. Myosin is then recovered as a precipitate. Because this myosin preparation is often contaminated by a trace amount of actin, as monitored by SDSpolyacrylamide gel electrophoresis (PAGE), we normally repeat these steps once more (Kohama and Kendrick-Jones, 1986). To prepare myosin that shows calcium inhibition, it is important to avoid artificial modifications during the purification procedure. Unlike the procedure originally developed by Hatano and Tazawa (1968), our procedure avoids column chromatography and hour-long centrifugation, and this may contribute to obtaining intact, native myosin. This can be monitored, for example, by examining whether sulfhydryl (SH) groups of myosin analogous to reactive thiols (SH2/SH1) of skeletal myosin (Sekine and Kielly, 1964) are protected from oxidation. Physarum myosin prepared by our procedure has intact reactive thiolds, the modification of which by N-ethylmaleimide makes myosin insensitive to calcium (Kohama et al., 1987). 2. Purification of Actin Because Physarum contains inhibitors to actin polymerization such as fragmin (see Section VI,D), the procedures that include acetone treatment for purifying actin from muscles are not applicable. Hatano and Owaribe (1977) have overcome this by heat treatment, which we employed as follows. The actomyosin preparation in 0.1 M Mg acetate and 20 mM ATP (see Section 111,BJ) was centrifuged and the resulting precipitate was suspended in 0.1 M KCl containing ATP and DTT. The suspension was incubated at 55°C for 20 min and dialyzed against 5 mM NaHC03 containing ATP and DTT. From the supernatant obtained after centrifuging the dialyzate, actin was purified by polymerization-depolymerization procedures. 3. Calcium Inhibition of Hybrid Actomyosins

The site of action of Ca2+is contained neither in actin nor in myosin of skeletal muscle. To use such a property, we reconstituted actomyosins from actin and myosin of Physarum and those of skeletal muscle. The respective actomyosins were then tested for Ca sensitivity by measuring of ATPase activity (Kohama and Kendrick-Jones, 1986). As shown in Fig. 3, the activity of the actomyosin was inhibited by Ca2+when myosin was from Physarum.

63


100

50

‘I

I

I

I

6

5

4

I

3

FIG. 3 Effect of Ca” on the ATPase activity of hybrid actomyosin formed from skeletal muscle and Physarum plasmodia (Kohama and Kendrick-Jones, 1986). Specific activities (nmol/min/mg myosin) at 100%were 164.5 for actomyosin reconstituted from Physarum actin and Physarum myosin ( O ) ,118.4 for actomyosin reconstituted from skeletal muscle actin and Physarum myosin (0),128.6 for actomyosin reconstituted from Physarum actin and skeletal muscle myosin (A),and 232.1 for actomyosin reconstituted from skeletal muscle actin and skeletal muscle myosin (A). Activities were determined by the pH-stat method in 15 mM KCI, 2 mM NaCI, 3.5 m M Mg”, 1.5 mM Mg-ATP, and 0.1 mM EGTA-Ca buffer at 25°C at pH 7.50.

However, the ATPase activity of the actomyosin produced from skeletal muscle myosin was not affected by Ca2+ no matter whether actin was from Physarum or skeletal muscle. Thus, Ca2+is demonstrated to exert an inhibitory effect through myosin. In the absence of actin, the ATPase activity of Physarum myosin is also inhibited by Ca2+,although the effect is slight (Kohama and Kendrick-

64


Jones, 1986). This observation indicates that the site of action of Ca2+is myosin and suggests that the effect of Ca2+should be amplified by actin. The role of actin will be further discussed in relation to the phosphorylated state of myosin in Section V,B.

4. Calcium Inhibition of Actin-Activated ATPase Activity of Physarum Myosin under Physiological Conditions 31P-NMR magnetic resonance studies of living plasmodia1 cells (Kohama et al., 1984) revealed that the intracellular concentrations of Mg2+and ATP and the intracellular pH were about 1 mM, 0.1-0.5 mM, and about 6.9, respectively. By other methods, the intracellular conditions of ATP, Kt, and pH were estimated to be 10.5 mM (Yoshimoto et al., 1981b), 30 mM (Anderson, 1964), and 7.0-7.5 (Morisawa and Steinhardt, 1982), respectively. Therefore, we measured the actin-activated ATPase activity of Physarum myosin in 1 mM Mg-ATP, 1 mM Mg2+,and 30 mM KC1 at pH 7.0 and observed a clean inhibitory effect of Ca2+ at p M levels as shown in Fig. 4 (Kohama and Kohama, 1984). Thus, the results shown in Fig. 3 were confirmed under physiological conditions.

5. Calcium Inhibition as Detected by in Vitro Motility Assays To demonstrate the myosin-linked nature of calcium inhibition by a quite different method, small latex beads were coated with Physarum myosin. They were then allowed to move along actin cables of Nitellu internodal cells as described by Shimmen and Yano (1984). As expected from the inhibitory effect of Ca2+on the ATPase activity of myosin, the beads moved much faster in the absence of Ca2' than in its presence (Shimmen and Kohama, 1984; Kohama and Shimmen, 1985; Table IV). Calcium regulation of the ATPase activities for myosin from scallop muscle is well characterized as will be described in Section IV,B. In this case, the effect of CaZt is diametrically different, stimulating the activity (Vale et al., 1984). Accordingly, the beads with scallop myosin moved in the presence of Ca2+,and the movement was abolished in the absence of Ca2+ (Table IV). The different effect of Ca2' using the same actin cables in this Nitella-based motility assay indicates that myosin is the site for Ca2' inhibition of the actin-myosin interaction in Physarum. To try another in vitro motility assay (Kron and Spudich, 1986; Harada et af., 1987), we prepared coverslips coated with either Physarum myosin or scallop myosin. Fluorescent actin from skeletal muscle was mounted on the coverslips in the presence of ATP and was observed with a fluorescence microscope (Okagaki et al., 1989). As shown in Fig. 5, actin moved in the


65

2001 A

c

'8

P

L

L.

L.

.5 E

tc Y

P

'5

. I

c1

..

7

5

6

4

3

pCaa FIG. 4 Effect of [Ca"] on the Mg-ATPase activity of Physariim myosin in the presence ( 0 ) and absence (0)of skeletal muscle actin under physiological conditions (0.5 mM Mg-ATP. 1 mM Mg", 30 mM KCI, and 0.1 mM EGTA-Ca buffer at pH 6.90 and 25°C) as measured by the pH-stat method (Kohama and Kohama, 1984). The conditions were estimated from "P-NMR spectrum (inset) from living plasmodia (Kohama et a/., 1984).

TABLE IV Myosin Movement along Actin Cables (pmlsec)

Solution

Physaritrn

EGTA Ca"

1.40 2 0.08 ( n = 18) 0.40 2 0.08 ( n = 11)

Scallop 0 (n 1.22 ? 0.39 ( n

= =

3) 3)

Note. Beads coated with Physarum or scallop myosin were introduced into Nitella cells together with EGTA solution (30 mM PIPES, 5 mM EGTA, 2 mM MgCI2, 66 mM KOH, 1 mM ATP, 1 mM DTT. and 170 mM sorbitol, final pH = 7.0) or with Ca'f solution (30 mM PIPES, 5 mM EGTA, 5 mM CaCll, 3 m M MgCI2, 76 mM KOH, 1 mM ATP. 1 mM DTT, and 170 mM sorbitol, final pH = 7.0) and observed with a Nomarski microscope (Shimmen and Kohama, 1984; Koham and Shimmen, 1985).

66

AKlO NAKAMURA AND KAZUHIRO KOHAMA Physarum

Scallop

l o t a

s

Q w

C

5-

5.

I

0

0

1.o

2Io

0

0

1.o

2.0

pm 'sec-1 Ca2+ myosin C (active) 4

L

b myosin-ca (inactive)

G+

myosin (inactive) 4 6

myosin-Ca (active) 2

+

FIG. 5 Inhibitory (a, b) and stimulatory (c, d) effects of CaZ' as measured by the myosincoated surface assay (Okagaki et al., 1989). Actin was labeled with rhodamine-phalloidine and mounted on a coverslip coated with Physarurn myosin (a, b) or scallop myosin (c, d). ATP-dependent movement of actin was observed in the presence of 0.1 mM EGTA (a, c) or 0.1 mM Ca2+(b, d) with a fluorescent microscope equipped with a video camera. Ordinate, number of moving actin; abscissa, velocities (pmhec). Arrows indicate average velocity of the movement.

absence of Ca2+on the Physarum myosin coverslip, and the movement was inhibited by Ca2+.Conversely, the same actin moved on the surface coated with scallop muscle myosin in the presence of actin but not in the absence of Ca2+.These differences confirm the myosin-linked nature of calcium inhibition. The effects of Ca2+on the actin-myosin interaction of Physarum myosin as examined by these two motility assays are not necessarily identical, as can be seen when Physarum myosin is dephosphorylated (see Section V,B,3).

67


IV. Ca-Binding Properties of Physerum Myosin Physarum myosin was found to bind 45Ca2twith a high K D affinity at p M levels (Kohama and Kendrick-Jones, 1986) and with a binding capacity of 2 mol CaZt per mole myosin (Table V). These data are comparable with those of myosin from scallop muscle, a typical myosin that is in an active form when it binds in Ca-containing solution. Release of Ca2' from scallop myosin upon the withdrawal of Ca2+inactivates its activity (Kendrick-Jones et al., 1970). The Ca-binding properties of Physarum myosin suggest that its calcium switch resembles that of scallop myosin (see Section IV,B). However, the effect of Ca2' for the interaction is quite distinct; Physarum myosin is in an active form when it loses Ca2+and in an inactive form when it binds Ca2t (Fig. 5).

A. Calcium-Binding Light Chain as a Ca-Receptive Subunit of Physarum Myosin Physarum myosin is composed of a pair of heavy chains (230 kDa in SDS-PAGE) and two pairs of light chains (18 and 14 kDa in SDS-PAGE). The domain structure of heavy chain has been examined; the binding sites for ATP, actin, and light chains are all in the heavy chain (Kohama et al., 1988b). The Ca-binding site is localized in the 14-kDa light chain, which we named the calcium-binding light chain (CaLc). The primary structure of CaLc was determined by both peptide analysis and cDNA cloning as shown in Fig. 6 (Kobayashi et al., 1988), and its molecular weight is calculated to be 16,084 Da. The EF-hand structure, which is a consensus sequence for Ca-binding proteins (Kretsinger, 1980), is identified at one position at the N terminal. The highest homology of CaLc

TABLE V Calcium Binding to Pbysarum Myosin and Its Subunits ~

~

Myosin (phosphorylated) Myosin (dephosphorylated) Calcium-binding light chain Phosphorylatable light chain

1.27 % 0.16 mol/mol (n = 3) 1.21 2 0.07 mol/mol ( n = 3) 0.37 -C 0.092 mollmol ( n = 6 ) ND

Nore. The binding of CA" to myosin and its subunits was measured in 0.5 M KCI, 1 nM MgClz, 20 m M Tris-HCI (pH 7.5) and 30 pM Ca" containing 4'Ca by equilibrium dialysis (Kohama et al. 1991a,b). ND, not detectable.

68


Physarum CaM 10 20 30 40 50 60 A C - V D S L T E E O I A E F K E A F S L F D K D G ~ ~ ~ N I ~ B L ~ S ~ ~ ~ Q D M I N ~ ~

70

80

90

100

110

120

G T I D E ' P B E ' L T B B I R E A F K V F D K D G N G F I S ~ ~ ~ ~ S D B E

130

140

VDEMIRBADVDGDGWNYDEFVKMMLSK

Physarum CaLc 10 20 30 40 50 60 Ac-TASAWIOECFPIFDKD~KVSIBBLGSAtRSLGlWPTNAELNTIK~LWLKEFDLATF 70 80 90 100 110 KTWRKPIKTPTEQSKEbEDAPRALDKBGN~IO~LRQL

120

130 140 VSVSGDGAINYESFVDbEVTGYPLASA FIG. 6 Amino acid sequences of calcium-binding light chain (CaLc) (Gene Bank/EMBL accession number(s) 503499) of Physarum myosin (Kobayashi etal., 1988) and Physarum calmodulin (CaM) (DDBJ/Gene Bank/EMBLaccessionnumber(s)AB022702) (Todaetal., 1990).Molecular weights and statistical pIhfor CaLc are 16,080and 4.33, and for CaM 16,606 and 3.92.

is found in calmodulin of bovine brain (see Section IV,C for phylogenic considerations). On a biochemical basis, CaLc shows calmodulin-like activity in that it activates phosphodiesterase activity (Table VI; Fig. 7). Interestingly, Ca2' at TABLE VI Summary of Functional and Structural Properties of Calcium-Binding Light Chain of Physarum Myosin Binds Ca"" Interacts with actin calcium-dependentlyh Interacts with heavy chain of skeletal muscle myosin as substitute for essential light chain" Shows similarity with vertebrate calmodulin in the amino acid sequence and in activating phosphodiesterase" Confers calcium inhibition on the actin-myosin interaction by binding to actin" "Kohama e t a / . (1991a). " Kohama et al. (1988a). ' Kohama et al. (1991b). " Kobayashi ef al. (1988). Kohama et al. (1985).

f . POLYCEPHALUMCALCIUMREGULATION

69

Y

FIG.7 CaLc stimulates phosphodiesterase activity in the presence of Ca2+.Phosphodiesterase activities (continuous line) (Kohama er al., 1991b) were measured in the presence of CaLc under conditions that allowed direct comparison with the ATPase activity as shown by the broken line.

pM levels activates phosphodiesterase activity through CaLc in the reverse manner as Ca2+inhibition of the ATPase activity of Physarurn myosin via CaLc. It has been determined that CaLc is not the site of activation or inhibition by Ca”. Thus, CaLc appears to work merely as a Ca-receptive subunit in Physarum myosin (Kohama et al., 1991b). Other evidence for CaLc being a Ca-receptive subunit includes the following: (i) The mobility of CaLc in SDS-PAGE of Physarum myosin changes in the presence of Ca2+(Kessler et al., 1980), (ii) the CaLc band after SDS-PAGE of Physarum myosin binds 4sCa2+(Kohama et al., 1985, 1986 a), and (iii) biochemical measurement (Table V) shows that CaLc binds 0.4 mol Ca” per mole. This figure is too low to explain Ca binding (1.3 mol Ca2’ per mole) of parent myosin. We speculate that the Ca-binding activity of CaLc may increase when incorporated into the myosin molecule. As will be described in Section V,A,1, the 18-kDa myosin light chain is phosphorylatable and is called the phosphylatable light chain (PLc). The

70


isolated PLc binds to skeletal muscle heavy chain as a substitute for 5 3 ' dithiobis(2-nitrobenzonic acid) (DTNB) light chain (Kohama et al., 1991b), but it does not bind Ca2+(Table V).

B. Speculative Mode of Ca Binding of Physarum Myosin Based on the Knowledge of Scallop Myosin The crystal structure of the regulatory domain of scallop myosin, which is composed of the essential and regulatory light chains and the light chainbinding fragment of the heavy chain, shows that essential light chain sequesters Ca2+at its N-terminal EF-hand structure, where the regulatory light chain is closely associated (Xie et al., 1994). The isolated essential light chain is unable to bind Ca2+.However, the regulatory domain can bind Ca2+ as strongly as parent scallop myosin (Kwon et al., 1990). The association of regulatory light chain allows the regulatory domain to bind Ca2+by stabilizing the bond. Chimeras between the essential light chain of Ca-binding scallop myosin and that of non-Ca-binding cardiac myosin were produced in Escherichia coli as recombinant proteins, and it was demonstrated that the third EFhand structure of the scallop essential light chain was required for the myosin to bind Ca2+ (Jancso and Szent-Gyorgyi, 1994). Similar analysis was carried out with chimeras between regulatory light chains of scallop and skeletal muscle myosins. The crucial portion for Ca binding by scallop myosin is at the C terminal of scallop regulatory light chain (Fromherz and Szent-Gyorgyi, 1995). Analysis with amino acid replacement showed that Gly117 at the C terminal of the regulatory light chain is of primary importance to support Ca2+binding to the scallop essential light chain (Jancso and Szent-Gyorgyi, 1994). Physarum CaLc and PLc are classified into essential and regulatory light chains, respectively, as shown in Table VI (Kohama et al., 1991b). An EFhand structure is identified at the N terminal of CaLc of Physarum myosin as described previously (Fig. 6). Resembling scallop myosin, Ca binding to CaLc is too weak to explain the Ca-binding activity of Physarum myosin (Table V). We speculate that Physarum myosin binds Ca2+at its N terminal in a mode similar to that for scallop myosin and that the binding is stabilized by PLc. It would be intriguing to know whether PLc has a Gly residue analogous to Gly117 in its C-terminal portion.

C. Phylogenic Considerations of Ca-Binding Light Chain of Physarum Multiple alignment of amino acid sequences of essential light chains (ELc) including CaLc were performed with DNASIS (Version 3.6), and phyloge-

71

f . POLYCEPHALUMCALCIUMREGULATION

netic trees were constructed from the sequence alignment with the Neighbor-joining method in the PHYLIP package (Version 3 . 5 ~ )Physarum . ELc showed the highest identity with Dictyosteliiim ELc (65%). The identity with protista and other invertebrate is 19%, and the level of identity with vertebrate and invertebrate ELc is 16%. As shown in Fig. 8, the sequences were separated into two major clusters, with one cluster containing skeletal muscle and smooth muscle. Physarum and Dictyosteliiim ELc sequence form a subcluster with high bootstrap probability (100%) in the protista/ other invertebrate clusters. We speculated that Physarum ELc evolved by a different route to vertebrate ELc.

Acanthamoeba

FIG.8 A phylogenetic tree constructed from the alignment of 15 sequences of myosin essential light chains using the SEQBOOT. PROTDIST, NEIGHBOR, and CONSENSE prograins provided in PHYLIP 3 . 5 ~(Felsenstein. 1993). The numbers at each branching point show bootstrap values (100 replications).

72


V. Phosphorylated State and Calcium Inhibition of Physsrum Myosin A. Phosphorylation and Dephosphorylation of Physarum Myosin

1. The Sites of Phosphorylation in Physarum Myosin Myosins from vertebrate muscle and nonmuscle sources are purified in the unphosphorylated state. However, those from lower eukaryotes, such as Physarum, Dictyostelium (Kuczmarski and Spudich, 1980; Maruta et al., 1983a), and Acanthamoeba (Collins and Korn, 198l), are prepared in the phosphorylated form. In the case of Physarum myosin, the total phosphate content as determined after assignment was 4.0-6.8 mol Pi per mole myosin (Kohama and Kendrick-Jones, 1986). We cultured Physarum cells in the presence of H3[32P]04and then prepared crude myosin. Autoradiography after SDS-PAGE showed that the major site of phosphorylation is in the heavy chain, but PLc is also partly phosphorylated (Kohama and KendrickJones, 1986). 2. Dephosphorylation and Calcium Inhibition We dephosphorylated Physarum myosin with phosphatase (Ogihara et al., 1983) and compared its actin-activated ATPase activity with that of phosphorylated myosin (Kohama et al., 1991a). As shown in Fig. 9, the activity of phosphorylated myosin is high in the absence of Ca2' and decreased with an increase in the Ca2+ concentration. In contrast, the activity of dephosphorylated myosin was low irrespective of Ca2+concentration. We examined the Ca-binding activity of myosin following dephosphorylation. As shown in Table V, dephosphorylated myosin bound Ca2+ as strongly as the phosphorylated form. Thus, dephosphorylation of Physarum myosin minimizes its ATPase activity so that there is no activity to be inhibited by Ca2+(Kohama et al., 1991a). The next step is to identify the site(s) that is responsible for the effect of dephosphorylation of myosin by replacing phosphate groups.

B. The Role of Actin in Calcium Inhibition 1. Dephosphorylation Affects the Affinity of Physarum Myosin to Actin Dephosphorylation reduces the affinity of myosin for actin, whereas ATPase maximum activity (Vmax) remains unaffected (Kohama et al., 1991a).

a

b .

t o -* IlKm I

0

n.s

I/Acl

2 0 4 0

I I .Q

Actin (mp ml -1)

FIG. 9 (a) Actin-activated ATPase activities of control, phosphorylated Physarum myosin ( 0 ) and . dephosphorylated Physarum myosin (0)(Kohama et a[., 1991a). Dephosphorylation reduced phosphate to 1.6 mol P, per mole of 500 kDa myosin. The activity was measured by calorimetry in 0.08 rng/ml of phosphorylated or dephosphorylated Physarum myosin, 0.08 mgl ml skeletal muscle actin, 13 m M KCI, 1 mM ATP, 2 mM MgCI2 20 mM Tris-HCI (pH 7.5), 0.1 m M DTT, and 0.1 mM EGTA-Ca buffer. (b) Effect of actin on ATPase activities of phosphorylated ( 0 )and dephosphorylated (0)Physarum myosins (KohamaetaL, 1991a). Double-reciprocal plots (inset) showed that V,,, (the ATPase activity at infinite concentration of actin) was not significantlydifferent between phosphorylated and dephosphorylated myosins. K , (the actin concentration required to achieve one-half V,,,,,) was smaller for phosphorylated myosin than for dephosphorylated myosin. Assay conditions: 0.044 mg/ml myosin, 12 mM KCI, 1 mM ATP, 2 mM MgC12,20 mM Tris-HC1 (pH 7.5). 0.1 mM DTT, and 0.1 mM EGTA.

74


Thus, when actin concentration is increased toward the concentration that gives Vmax, the activity of dephosphorylated myosin is similar to that of phosphorylated myosin. Thus, under actin-poor conditions, the activities of the dephosphorylated myosin were reduced to the basal level, irrespective of Ca2' concentration (Fig. lob). On the other hand, the activities of phosphorylated myosin were high in the absence of CaZt and low in the presence of CaZt (Fig. 10a). When actin concentration was elevated about 10-fold over that of myosin on a weight basis (Fig. lOd), the activities of the

Actin-Rich

Actin-Poor

0

50

100

0

I

I

I

50

100

150

--

0

so

100

0

50

100

d

I50

nmolmln-hny'

FIG. 10 Effect of phosphorylation on the actin-activated ATPase activity of Physariim myosin

(Kohama et al., 1991a). Physariim myosin was prepared in the phosphorylated form and an aliquot was dephosphorylated as described in the legend to Fig. 9. Actin-activated ATPase activities of the phosphorylated (a, c) and dephosphorylated (b, d) myosins were determined by the colorimetry in the presence of 0.1 rnM EGTA (solid bars) or 0.1 mM Ca2' (open bars). The protein concentrations were 0.072 mglml myosin (a-d), 0.056 mg/ml actin (a, b), and 0.46 mg/ml actin (c, d). Numbers on the abscissa are the ATPase activities in nmol min-' myosin, and bars are SEM ( n = 3). Under actin-rich conditions, the ATPase activity of dephosphorylated myosin shows Ca inhibition.

P.POLYCEPHALUM CALCIUM REGULATlON

75

dephosphorylated myosin were similar to those of phosphorylated myosin, i.e., high in the absence of Ca” and low in its presence.

2. Ca-Binding Mode, Rather Than Phosphorylating Mode, Is of Physiological Importance Under actin-poor conditions (see Section V,B,l), the ATPase activity of Physarum myosin is modified both by binding Ca2+ to myosin and by changing its phosphorylated state (Figs. 10a and lob). Under actin-rich conditions, only the former mode regulates the activities (Figs. 1Oc and 10d). Therefore, the crucial factor in determining which of the two modes is dominant in vivo is the concentration of actin in Physarum cells. In muscular tissues, the concentration of actin is comparable to that of myosin on a weight basis. Nonmuscle tissues, however, are expected to be in an actin-rich condition because actin greatly exceeds myosin in concentration (Pollard and Weihing, 1974). This is true in Physarum cells (Ishikawa et al., 1991), and hence the mode of Ca binding should be physiological. Change in the phosphorylated state of myosin may not play a major role in vivo.

3. In Vitro Motility Assays in Relation to Actin-Rich and Actin-Poor Conditions Superprecipitation studies with a spectrophotometer detect an ATP-dependent shrinkage of the actomyosin complex (Kohama et al., 1986b). Physarum myosin superprecipitated with actin is in the untreated, phosphorylated form. The extent of superprecipitation was higher in EGTA than in Ca2+(Ogihara el al., 1983). These observations are in agreement with the results of ATPase measurement under actin-poor conditions (Fig. 2). The myosin-coated surface assay also provides actin-poor conditions because single actin filaments move on a two-dimensional surface coated with myosin (Fig. 5). When coated with Physarum myosin in the phosphorylated form, rapid ATP-dependent movement of actin can be observed especially in the absence of Ca2+.However, actin does not move on the surface coated with dephosphorylated Physarum myosin irrespective of Ca’+ concentrations (Kohama et al., 1991a). Another in vitro motility assay detects an ATP-dependent movement of myosin along the actin cables fixed inside the cell membrane of Nitella. This Nitella-based motility assay allows myosin to interact with actin under actin-rich conditions. Accordingly, Physarum myosin moves faster in EGTA than in Ca2+irrespective of whether it is in the untreated, phosphory-

76


lated form as shown in Figs. l l a and b or 11 in the dephosphorylated form as shown in Figs. l l c and l l d (Kohama et al., 1991a).

C. Dependence on Ca2+of Protein Kinases and Phosphatases That Act on Myosin Kinases specific for the myosin heavy chain and PLc have molecular weights of 76 and 55 kDa, respectively, as indicated by SDS-PAGE (Okagaki et

Phosphorylated

Dephosphorylated

a

C

1

I

loc ':LA 2o

00

20

0.2

0.4

0.6

-

o

0.8

b

0.2

0.4

0.6

0.8

d

20

10

0

o

0.2

0.4

0.6

0.a

0

I

1

F

I

0.2

0.4

0.6

0.8

pm sec-' FIG. 11 Velocities of movement of phosphorylated and dephosphorylated myosins as examined by the Nitellu-based motility assay (Kohama et al., 1991a). Actin cables in a Nitella internodal cell were exposed by internal perfusion. Small latex beads coated with phosphorylated (a, b) or dephosphorylated (c, d) myosin were introduced to the actin with EGTA solution (a, c) or with Ca2' solution (b, d). Velocities of the ATP-dependent movement of the beads were measured by observations under Nomarski microscope. Ordinate, number of moving particles; abscissa, velocities (pmlsec). Arrows indicate average velocity of the movement. With the Nitella-based assay, dephosphorylated myosin moved in the similar way to phosphorylated myosin. The movement of dephosphorylated myosin is blocked by Ca2+ almost perfectly. The data of a and b confirms those of a and b in Fig. 5 because Physarum myosin in Fig. 5 is in a phosphorylated form.

P. POLYCEPHALUMCALCIUM REGULATION

77

al., 1991a,b). The effect of Ca2+on their activities is similar to its effect on myosin, i.e., these activities are most pronounced in the absence of Ca2' and are reduced with an increase in Ca2+concentration. A sole Ca-binding protein is involved in the inhibitory effect of Ca2+(see Section VI1,A). The phosphatase activities for myosin are much lower in Physarum cells than the kinase activities. However, native actomyosin preparation contained not only the kinase activities but also the phosphatase activities. We incubated the preparation with [32P]y-ATPto phosphorylate myosin; the phosphorylation was terminated by the addition of the kinase inhibitor staurosporine (Okagaki et al., 1991a). We observed that the radioactivity incorporated into these proteins was gradually lost by the phosphatase activity. The decrease in radioactivity was abolished by the phosphatase inhibitor okadaic acid. The phosphatase activities were low in the presence of Ca2t and increased with an increase in [Ca2+]to p M levels. Furthermore, the calmodulin inhibitor trifluoperazine also inhibited the phosphatase activities, suggesting the involvement of calmodulin in these activities (Kohama et al., 1993). Figure 12 summarizes the role of Ca2' and phosphorylation in the relationship between Ca binding and phosphorylation of Physarum myosin (Kohama el al., 1990, 1993). Step 2 is mediated by phosphatase activities, which works at high Ca2+levels. Kinase reactions are expressed by step 4, which occurs at low Ca2+concentrations. Steps 1 and 3 are mediated by direct Ca binding to myosin. The Ca-binding activity remains the same regardless of the phosphorylated state of myosin (see Table V, and Section V,A,2). Under actin-poor conditions (see Section V,B,l), the sole form of active myosin is phosphorylated and free from Ca2' (Fig. 12). As discussed in Section V,B,l, dephosphorylation of myosin reduces its affinity to actin, and an actin-rich condition rescues the reduction. Therefore, under actinrich conditions (Fig. 12), as found in Physarurn cells, myosins in both phosphorylated and dephosphorylated forms are in the active form in low [Ca2+].Ca binding to both myosins inhibits their activities. In other words, the physiological switch that determines whether Physarum myosin is active or inactive is Ca binding to myosin (see Section V,B,2).

VI. Actin-Binding Proteins of Physarum That Are Involved in Calcium Inhibition

The actin-linked mode of calcium inhibition has been indicated in an earlier stage of our studies (Kohama, 1981; Kohama et al., 1985). We currently understand that the actin-binding proteins may back up the Ca inhibition due to Ca binding to myosin.

78


FIG. 12 A model of the role of Ca2’ and phosphorylation in fhysctrum myosin. Myosin is active only in the phosphorylated form at low Ca” concentrations (asterisk). Binding of Ca2+ occurs irrespective of the extent of phosphorylation when high [Ca2+](see steps 1 and 3). Myosin can be dephosphorylated at high [Ca”] (step 2) and phosphorylated at low Ca” (step 4). The Ca-binding proteins are calcium-binding light chain (CaLc) for steps 1and 3, calmodulin for step 2. and calcium-dependent inhibitory factor for step 4. The phosphorylated form of myosin is denoted by P. Under actin-poor conditions, the sole active form of myosin is shown by *, and under actin-rich condition the two active forms of myosin are shown by * and **. As described in the text, the actin-rich condition is physiological for fhysarurn plasmodium. Therefore, binding and release of Ca2+should be the physiological mode of regulation.

A. Search for a Regulatory Mode Other Than That Inhibited to Myosin When ATPase activity of native actomyosin was compared with actomyosin reconstituted from purified actin and myosin, the inhibitory effect of Ca2+ of the former was more pronounced than that of the latter (Kohama and Kohama, 1984). If the inhibition were exerted exclusively through myosin, the effect should be the same. Thus, the inhibitory factor(s) is speculated to be lost during the purification. The same has been suggested by the following experiment. When Mg2+ concentration was elevated in the assay medium, calcium inhibition of the actin-myosin interaction was obscured (Table VII), presumably by competition for the Ca-binding site of myosin. A fraction was obtained from the actomyosin preparation that shows calcium inhibition. This fraction was mixed with actin and myosin at high [Mg2’] concentration and superprecipi-

79


TABLE VII Effect of Calcium-Binding Light Chain (CaLc) of Physarum Myosin and Bovine Brain Calmodulin on CaInhibition (%) of Actin-Activated ATPase Activity of Physarum Myosin ~

ATPase activity (nmol/rnin/rng myosin) Myosin (30 pgiml)

EGTA

50 p M Ca”

%“

Untreated NEM modified NEM modified

0 0 3.5

3.5 3.5 3.5

5 5 5

96.8 43.9 66.7

33.9 27.3 33.3

65.0 37.9 50.0

Untreated Untreated Untreated

0 0 3.5

3.5 8.5 8.5

20 5 5

79.3 119.5 123.2

28.0 97.6 61 .o

64.7 18.3 50.5

Nore. (Top) Desensitization by CaLc of the myosin desensitized by NEM modification (about 2 mol NEM/mol myosin) (Kohama et 01.. 1987). (Bottom) Desensitization by CaLc of the myosin apparent by desensitization in the presence of high (8.5 m M ) Mg” (Kohama et ul.. 1985). Assay conditions: 1.5 mM ATP, 0.1 m M EGTA-Ca buffer with a pH-stat method at p H 7.50 and 25°C. Actin concentrations are 100 pg/ml. ‘‘ 100 X (ATPase in EGTA-ATPase in Ca”)/(ATPase in EGTA).

tated, and the ATPase activity of the mixture was shown to be inhibited by Ca2+(Kohama, 1981). This inhibitory fraction binds to an affinity column, suggesting that Ca inhibition is exerted via actin. We speculate that the Ca-binding components in the factor(s) could be CaLc andlor calmodulin as will be described in Seclions VI,B and V1,C.

B. CaLc as an Actin-Binding Protein When CaLc was isolated from Physarunz myosin, it was found to interact with actin (Table VI). The interaction was first demonstrated by the binding to an actin affinity column (Kohama et al,, 1985) and was further confirmed by detecting the effect of CaLc on the actin polymerization (Fig. 13). The effect, as examined by the measure of viscosity and flow birefringence of actin, was inhibited by Ca*+(Kohama et al., 1988a). Such effects have never been observed with light chains isolated from vertebrate myosins, although it has been observed that essential light chains show a subtle interaction with actin (Yamamoto and Sekine, 1983; Kohama, 1987). To determine whether CaLc confers calcium inhibition to the actinmyosin interaction of Physarum, we examined its effect on the actin-activated ATPase activity of Physaritm myosin under the conditions in which the activity was not sensitive to Ca”. As shown in Table VII, Ca2+hardly

80


0.2

1

o ! 0

I

I

I

30

60

90

min

FIG. 13 Interaction of calcium-binding light chain (CaLc) as monitored by Ca-dependent inhibition of actin polymerization (Kohama et al., 1988). Skeletal muscle monomeric actin, whose polymerization alone showed no Ca dependence, was mixed with KCI at a final concentration of 114 mM and allowed to polymerize. Viscosity was measured with an Ostowald capillary viscometer at 25°C. Conditions: 140 kg/ml skeletal muscle actin, 3.3 pg/ml CaLc, 1 mM MgC12,23 mM Tris-HCI (pH 7.5), and 0.1 mM EGTA-Ca buffer. 0 ,in 0.1 mM EGTA; 0, in 0.1 mM Ca2+.

affected the activity in the absence of CaLc. However, a lower activity was detected in the presence of Ca2+ when CaLc was mixed with actin and myosin. In Physarum cells, a significant amount of CaLc is present without associating myosin (Kohama et al., 1985), although CaLc is present in the cells as a myosin subunit. We speculate that the isolated CaLc may take part in such a role. The previous experiments were carried out using CaLc purified from P h y s a n m Now that recombinant CaLc is available, we need to confirm the results and to identify the sequences responsible by mutating CaLc.

C. Caldesmon-like Protein of Physarum We purified a 210-kDa, heat-stable protein from Physarum that reacts with an antibody against caldesmon, an actin-binding, regulatory protein of

81

f . POL YCEPHALUM CALCIUM REGULATION

smooth muscle (Ishikawa et al., 1991, 1992). This caldesmon-like protein binds to actin and stimulates the actin-activated ATPase activity of Physarurn myosin and the actin motility on a surface coated with Physarurn myosin (Table VIII). The stimulation is abolished when calmodulin is mixed with the caldesmon-like protein in the presence of Ca2+.Furthermore, calmodulin is found in Physarurn (Ishikawa et al., 1991). Therefore, caldesmon-like protein is able to produce calcium inhibition together with calmodulin. Such an actin-linked mode may back up the inhibitory effect of Ca2+ on Physarurn myosin by CaLc (Fig. 14). It must be noted that the stimulatory effect of caldesmon-like protein is distinct from the inhibitory effect reported for smooth muscle caldesmon (Sobue and Sellers, 1991). However, smooth muscle caldesmon is also shown to stimulate its actin-myosin interaction under the specified conditions. Because this stimulation is related to the myosin-binding property of caldesmon (Lin et al., 1994), we need to reexamine the regulatory mode of Physarurn caldesmon-like protein.

D. Other Actin-Binding Proteins of Physarum Fragmin binds to actin filaments at their barbed end and severs them in the presence of Ca2’ (Hasegawa et al., 1980). Furthermore, it forms a complex with monomeric actin, and the complex becomes a nucleus for polymerization of actin into filaments (Hasegawa et al., 1980; Hinssen, 1981a,b; Sugino and Hatano, 1982; Maruta et al., 1983b). Fragmin is considered to be an analog of vertebrate gelsolin (Yin and Stossel, 1980). Isoforms of fragmin were purified by Uyeda et al. (1988) and Furuhashi and Hatano (1989). Profilin was also identified in Physarurn as another protein that forms a complex with monomeric actin (Ozaki et al., 1983). TABLE Vlll Velocities of Movement of Actin Filaments on Coverslips Coated with Physarum Myosin

Actin filament

Velocity ( p d s e c , n = 30)

Control (0.1 mM EGTA) + Caldesmon-like protein (0.1 mM EGTA) Control (0.1 mM Ca2+) + Caldesrnon-like protein (0.1 mM Ca”) + Caldesmon-like protein + calmodulin (0.1 mM Ca”)

1.39 ? 0.49 1.95 t 0.66 0.99 ? 0.42 1.26 ? 0.60 1.08 t 0.50

Note. Caldemon-like protein was purified from Physarurn plasmodia as a heat-stable, actinbinding protein. Unlike smooth muscle caldesmon, caldesmon-like protein stimulated the velocity of movement. Because calmodulin in the presence of CaZ+abolishes binding to actin, caldesmon-like protein works cause calcium inhibition (Ishikawa et al., 1991).

82


FIG. 14 Relationship between Ca2+ and the actin-activated ATPase activity of Physarum myosin (Jshikawa et al., 1991, 1992). (A) Myosin is regulated both by phosphorylation and by Ca” binding. Myosin must be phosphorylated to be active, and Ca” inhibits the activity by binding to the myosin. (B) A caldesmon-like protein of Physarum enhances the activity. The enhancement is abolished by calmodulin in the presence of Ca2’ with resulting enhancement of calcium inhibition of the activity (Ishikawa et al., 1991, 1992). According to Section V,B, the assay condition is actin poor.

The actin-binding proteins that may be involved in the organization of Physarum plasmodium are high-molecular-weight actin-binding protein (Sutoh et al., 1984), a homolog of smooth muscle filamin (Wang, 1977); connectin/titin (Ozaki and Maruyama, 1980; Gassner et al., 1985), an elastic protein originally found in muscle tissue (Maruyama et al., 1976); a 52kDa protein that bundles actin filaments (Itano and Hatano, 1991); and


83

caldesmon-like protein (Ishikawa et al., 1991, 1992). lshikawa et al. (1995) purified an ATP-dependent actin-binding protein from Physarum plasmodia. The effect of actin binding has not yet been detected. As described in Section VI,C, caldesmon-like protein together with CaM takes part in the calcium inhibition of the actin-myosin interaction (Jshikawa et al., 1991). Sugino and Matsumura (1983) showed that the tension of the actomyosin thread was reduced by fragmin in the presence of Ca" (Table 11). Connectin/titin, although prepared from skeletal muscle, binds to actin in a calcium-dependent manner and inhibits the actin-myosin interaction (Kellermayer and Granzier, 1996).

VII. Ca-Binding Proteins in Physarum A. Three Ca-Binding Proteins That Support Calcium Inhibition In this review, we have described three Ca-binding proteins in Physarum as summarized in Fig. 15. The one most characterized is CaLc as shown in Table VI (Kohama et al., 1992). Not only does CaLc work as a Ca-binding subunit of Physarum myosin but also it binds to actin. Both myosin- and actin-linked properties contribute to the actin-myosin interaction as described in Sections IV,A and VI,B. The amino acid sequence of Physarum calmodulin (Fig. 7 ) is very similar to that of brain calmodulin (88% identity). Calmodulin binds to caldesmonlike protein in the presence of Ca2+to abolish the stimulatory effect of the protein on the actin-myosin interaction (see Section V1,C). The abolition allows calcium inhibition of the actin-myosin interaction. In the presence of Ca", calmodulin stimulates myosin phosphatase activity (Kohama et al., 1993). In the absence of Ca2+,myosin phosphatase remains low to keep myosin phosphorylated and thus sensitive to Ca". Therefore, calmodulin indirectly contributes to calcium inhibition of myosin. As described in Section V,C the activities of the kinases for the heavy (76 kDa in SDS-PAGE) and light (55 kDa) chains of Physarum myosin are also calcium inhibitory, i.e., at low Ca2+concentration Physarum myosin tends to be phosphorylated. Because phosphorylated myosin is active, calcium inhibition of kinase activities also contributes to the inhibition of the actin-myosin interaction. A Ca-binding protein of 38 kDa in SDS-PAGE has been shown to be involved in the calcium inhibition of kinase activities and is called calcium-dependent inhibitory factor (ClF) as shown in Fig. 15 (Okagaki et al., 1991a,b). Actin kinase of Physarum has also been shown

84


CALCIUM INHIBITION

FIG.15 Three Ca-binding proteins involved in calcium inhibition of the Physarum actomyosin system. CIF, calcium-dependent inhibitory factor for kinases (Okagaki ef al., 1991a.b); CaLc, calcium-binding light chain of Physarum myosin (Kobayashi et al., 1988; Kohama et al., 1991b); CaM, Physanrm calmodulin (Toda et al., 1990); CaD, caldesmon-like protein of Physarum (Ishikawa et al., 1991). CaLc works as a sole Ca2+-receptive protein in Physarum myosin, producing calcium inhibition. When CIF binds Ca2+,CIF interacts with kinases for Physarum myosin. The myosin in the phosphorylated form interacts with actin more effectively than when dephosphorylated. Ca2+stimulates myosin phosphatase activity (Kohama el al., 1993) to dephosphorylate myosin, which is lost calcium inhibition. CaD enhances the interaction by binding to actin. The enhancement is negated by CaM binding Ca2+.CaLc binds actin and allows Ca2+to cause inhibition (Kohama et al., 1985).

to be subject to calcium inhibition (Okagaki et al., 1991a). When actin forms a complex with fragmin, the kinase is able to phosphorylate actin (Furuhashi and Hatano, 1990, 1992; Furuhashi et al., 1992; Gettemans et al., 1992, 1993).

B. Preliminary Characterization of Recombinant 40-kDa Protein A 40-kDa Ca-binding protein was purified from Physarum (Nakamura et al., 1994). The cloning of its cDNA showed that it is identical to LAV1-2 cloned by Laroche et al. (1989) as an abundant mRNA specific to Physarum plasmodia. We expected 40-kDa protein to work as the CIF described in Sections V,C and VII,A because CIF has a similar molecular mass (38 kDa) (Okagaki et al., 1991a,b). However, the 40-kDa protein obtained as a recombinant protein in E. coli failed to exert an inhibitory effect on myosin heavy and light chain kinases. We interpreted this to mean that CIF differs from 40-kDa protein. The other interpretation is that posttran-


85

scriptional modifications such as glycosylation and phosphorylation may be required for 40-kDa protein to inhibit the kinases.

1. Structure of 40-kDa Protein As shown in Fig. 16, the 40-kDa protein consists of 355 amino acid residues (Nakamura et al., 1994). The calculated molecular mass and p l are 40,508 Da and 4.97, respectively. Four EF-hand Ca-binding consensus sequences (Kretsinger, 1980) were found in the C-terminal half, which also contained a consensus sequence for nuclear proteins as reported by Robbins et al. (1991). The sequence is 226 RKIDTNSNGTLSRKEFR 242. The N-terminal half contained an @-helical structure predicted by MacDNASIS Pro 1993. Residues 1-32 formed aggregates of the 40-kDa protein.

2. Ca-Binding Properties of the 40-kDa Protein Recombinant 40-kDa protein was obtained as a soluble form in low ionic strength. However, elevation to p M levels allowed the 40-kDa protein to form large aggregates (Nakamura et al., 1994; Nakamura and Kohama, 1995). The ability to form aggregates was abolished by producing a mutant that was deleted for residues 1-32. When this truncated protein was subjected to the Ca-binding assay with a flow dialysis apparatus (Womack and Colowic, 1973), 4 mol Ca/mol of 40-kDa protein was bound with halfmaximal binding at a concentration of 0.29 p M . The concentration is much lower than that of Physarum calmodulin (5.7 p M ) , which was also obtained in the recombinant form (Fig. 6). The C-terminal half of the 40-kDa protein is highly homologous to calmodulin, whereas the N-terminal half is devoid of calmodulin sequences. We speculate that the secret of the higher Cabinding activity of the 40-kDa protein is in its N-terminal half. It will be of interest to see whether a fusion protein of calmodulin with N-terminal half 40-kD protein has increased Ca-binding properties.

3. Other Properties of the 40-kDa Protein An antibody was raised against the 40-kDa protein. Immunofluorescent studies with the antibody showed staining of Physarum nuclei, although the cytoplasm was also partially stained (Fig. 17). When the staining between the plasmodial and amebal stages of Physarum was compared (see Section VIII), the staining was observed only in plasmodial stages. The different expression of the 40-kDa protein was confirmed by subjecting total lysates and total RNAs from both plasmodial and amebal cells to Western blotting and RT-PCR, respectively. When the genomic PCR product was compared with that from RT-PCR, they showed the same mobility

86


A~ - S Y ~ B A W N10P M S S L D E I20I S I ~ L K S K30T G A V K B I F4S0O E L M R B ~50K K ~ B N L B g60L Q I ( U D 70 80 90 100 110 120 B T S F A B K B D R D R C B A ~ I A O K B O B O ~ Y ~ ~ O N B F D ~ ~ R B R B ~ G D ~ K 130

140

150

160

170

180

190

200

210

220

230

240

~~LLKDLBDIU8GY~SKP~SBB~ILROL~SSAVSGSGKFSFODLKO~

K Y A D T I P B G P L K U ~ ~ G ~ S Y I r r V A V A V ~ ~ V ~ F ~ I ~ S N ~ S ~ 250 260 270 280 290 300 ~ B F J R L G P D K K S V O D A L F G R aYVeLGLCLLVLRILYAFADFDKSGQ Y ~ 310

320

330

340

350

~ O ~ ~ D A a I P S S ~ K ~ B O F S ~ ~ D S KVLLWEDD S L S Y O B ~

FIG. 16 Structure and function of the 40-kDa protein (Gene Bank/EMBL accession number(s) X14.502). (A) Amino acid sequence of the 40-kDa protein deduced from its nucleotide sequence. Molecular weight, 40,508 Da; statistical pl, 4.97. (B) Domain structures of the parent 40-kDa protein and the truncated form of the 40-kDa protein compared with that of calmodulin (CaM). (C) We expressed the 40-kDa protein, its truncated form, and Physarum calmodulin (CaM) in E. coli and purified them. The measurement of Ca binding to the 40-kDa protein was hampered by its Ca-dependent aggregation. Therefore, we used the truncated form as shown (0).We also measured the Ca-binding activities of CaM (0).Assay conditions; 100 mM NaCI, 20 mM MOPS (pH 7.0), and various concentrations of Cazt.


87

FIG. 17 Immunostaining of Physarum plasmodium. Physarum plasmodium (strain Ng-1) was treated for fluorescent microscopy (Uyeda and Kohama, 1987) and double stained with D A P I and the antibody against 40-kDa protein. (A) Phase-contrast micrograph; (B) immunofluorescent micrograph. Arrows and arrowheads show the staining of cytoplasm and nuclei, respectively. (C) Nuclear staining with DAPI. Scale bar = 10 pm. (D) Specificity of the antibody. Total homogenate (PCL) and recombinant 40-kDa protein (40K) were subjected to Western blots using the antibody. CBB. protein staining with Coomassie brilliant blue R-250; anti 40K, immunostaining with the antibody.

88


in the agarose gel, indicated that the 40-kDa protein had no introns; this has been confirmed by nucleotide sequencing of the products. 4. Immunostaining of Vertebrate Cells with the Antibody against the 40-kDa Protein CHO-k1 was one of the vertebrate cell lines to react with the antibody against the 40-kDa protein. In Western blots of the cell lysate, the antibody reacted with a 40-kDa band, suggesting the presence of a homolog in the CHO-k1 cell. Immunofluorescent studies showed staining of the nucleolus, as shown in Fig. 18. During mitosis, the dot-like staining of nucleoli moved to chromosomes. At the late stage of mitosis, the dot-like staining disappeared and the staining became homogeneous throughout the cells (Fig. 19). The other cell to react with the antibody was the 10T1/2 cell, which showed immunofluorescent staining at both the nucleolus and endoplasmic reticulum (Fig. 18).

VIII. Amebal Myosin and Amebal-Plasmodia1 Transition The life cycle of Physarum consists of two phases with different modes of motility: uninucleate haploid amebae showing slow ameboid movement and multinucleated diploid plasmodia showing rapid cytoplasmic streaming. Actomyosin was obtained from the amebae as described in Section III,A (Kohama and Takano-Ohmuro, 1984) and amebal myosin was purified from the actomyosin preparation as described in Section II1,B (Kohama et al,, 1986a). Amebal myosin resembles plasmodia1 myosin in its twoheaded, long-tailed shape and in its subunit composition of heavy chain, Ca-binding light chain and phosphorylatable light chain. The heavy chain and phosphorylatable light chain differ between the myosins as examined by peptide mapping, whereas the Ca-binding light chain of both myosins appears to be identical. The actin-activated ATPase activities of the amebal variety did not differ greatly from those of plasmodial myosin; the effect of Ca2' was also inhibitory. Therefore, the different motility between ameba and plasmodium cannot be ascribed to myosin but to other contractile proteins. The contractile proteins specific to plasmodium are the 40-kDa Ca-binding protein discussed in Section VI1,b and high-molecular-weight, actinbinding protein (HMWP) (Uyeda and Kohama, 1987). Immunofluorescent studies with an antibody against HMWP showed that it was detectable during the transition from ameba to plasmodium (Fig. 20). The transition


89

FIG. 18 Immunostaining of vertebrate cells with the antibody against Physarum 40-kDa protein. (A-F) CHO-K1 cells (A-C) and 10 T1/2 cells (D-F) were cultured on coverslips and double stained with the antibody and DAPI. ( A and D) Phase contrast micrographs; (B and E) immunofluorescent images-arrowheads and arrows indicate the staining of nucleoli and endoplasmic reticula, respectively; (C and F) nuclear staining with DAPI (scale bar = 10 pm. (G) Western blots of CHO-K1 cell extracts. The lysate of CHO-Kl cells was applied to the an affinity column conjugated with the antibody. After removing unbound materials, the bound proteins were eluted. PCL, lysate of Physaricm plasmodia (see the legend to Fig. 17); CHO, lysate of CHO-kl cells: AFP, elute from the affinity column; CBB, protein staining with CBB; anti 40K, immunostaining with the antibody.

90


FIG. 19 Immunofluorescent images of CHO-K1 cells during mitosis. CHO-K1 cells were double stained with D A P I and the antibody against the Physururn 40-kDa protein. A-C, interphase; D-F, metaphase; G-I, anaphase. (A, D, G) phase-contrast micrographs; (B, E, H ) DAPI-fluorescent images, and (C, F, I) immunofluorescent images. As shown by the arrows in C and F, the staining of nucleoli in interphase moved to chromosomes during metaphase. In anaphase (I), the whole cytoplasm was stained with the antibody.

was also examined using antibodies specific to either amebal or plasmodial myosin heavy chain. Stage-specific expression of both proteins was confirmed. Interestingly, intermediate cells with swollen nuclei contained both heavy chains (Anderson et al., 1976). Similar observation was reported (Uyeda et al., 1988) with amebal and plasmodial fragmins, which are homolog of vertebrate gelsolin (Yin and Stossel, 1980). These observations indicate that the proteins related to the actomyosin system were switched in a coordinated manner during the intermediate stage. The switching contributes to a radical reconstruction of the cytoskeletal architecture during amebal and plasmodia1 transition.

IX. Concluding Remarks In this review, motility of Physarum is shown to be subject to calcium inhibition by various methods (Table VI). The major path for calcium


91 Plasmodia

-

Amoeba

PMHC AMHC ? PF AF CaLC

PLC d HMWP

40K 4 FIG. 20 Changes in synthesis ofcytoskeletal proteins that are related to the actomyosin system during amebal-plasmodia1 transition of a colonial strain (schematically presented) (modified from Uyeda and Kohama. 1987). PMHC, plasmodial myosin heavy chain: AMHC, amebal myosin heavy chain: PF, plasmodial fragmin: AF, amebal fragmin; CaLc, calcium-binding light chain expressed in both plasmodiai and amebal myosins; PLC, phosphorylatable light chain of plasmodial myosin: HMWP. high-molecular-weight myosin-binding protein: 40K, 40kDa protein.

inhibition is through myosin: Myosin is in an active form when Ca2+ is absent, and it is inactivated by binding Ca7+to CaLc (Fig. 5). Another route is via actin-binding proteins such as CaLc (Table VII), fragmin (Sugino and Matsumura, 1983),and caldesmon-like protein in association with calmodulin (Table VIII). Phosphorylation and dephosphorylation also modify the properties of Physariim myosin. It is purified in a phosphorylated form. Dephosphorylation procedures reduce its affinity to actin. Although dephosphorylation does not affect its Ca-binding property (Table V), dephosphorylated myosin does not show calcium inhibition. When the actin concentration is increased to the level found in living plasmodia, the calcium inhibition lost by dephosphorylation of myosin is recovered (Fig. 12). Therefore, changes in the phosphorylated state of myosin may not play a major regulatory role. More important for regulation is the binding of Ca2’ to myosin. The kinase activities responsible for phosphorylation of myosin are detectable for both PLc and heavy chain and are inhibited in a Ca-containing solution. Calcium-dependent inhibitory factor is expected to work as a Cabinding protein (Fig. 15). It is important to note that the kinases are active

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without requiring Ca”. Such enzymes are expected to be involved in general cell maintenance and thus be exempted from the regulation by Ca”. However, the discovery of calcium inhibition indicates that exemption is an erroneous idea and suggests that some of the housekeeping enzymes may be under the control of Ca2+if they are associated with novel Ca-binding proteins that should exert an inhibitory effect. We hope that our review may inspire scientists in the search for such proteins.

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Vale, R. P., Szent-Gyorgyi. A. G . , and Sheetz, M. P. (1984). Movement of scallop myosin on Nitella actin filaments: Regulation by calcium. Proc. Natl. Acad. Sci. USA 81, 6775-6778. Wang, K. (1977). Filamin, a new high-molecular-weight protein found in smooth muscle and nonmuscle cells. Purifications and properties of chicken gizzard filamin. Biochemistry 16, 1857-1865. Williamson, R. E. (1975). Cytoplasmic streaming in C h a m A cell model activated by ATP and inhibited by cytochalasin B. .I. Cell Sci. 17, 655-688. Williamson, R. E., and Ashley, C. C. (1982). Free CaZt and cytoplasmic streaming in the alga Chara. Nature 296, 647-651. Womack, F. C., and Colowic, S. P. (1973). Rapid measurement of binding of ligands by rate of dialysis. Methods Enzymol. 27, 464-471. Xie, X., Harrison, I., Schlichting, D. H., Sweet, R. M., Kalabokis, V. N., Szent-Gyorgyi, G., and Cohen, C. (1994). Structure of the regulatory domain of scallop myosin at 2.8 A resolution. Nature 368,304-312. Yamamoto, K.. and Sekine, T. (1983). Interaction of alkali light chain 1 with actin: Effect of ionic strength on the cross-linkingof alkali light chain 1with actin. J. Biochem. 94,2075-2078. Yamamoto, K., Kikuyama, M., Sutoh-Yamamoto, N., Kamitsubo, E., and Katayama, E. (1995). Myosin from alga Chum: Unique structure revealed by electron microscopy. J . Mol. Bid. 254, 109-112. Yin, H. L., and Stossel. T. P. (1980). Purification and structural properties of gelsolin, a Ca2+activated regulatory protein of macrophages. J. B i d . Chem. 255, 9490-9493. Yokota, E., and Shimmen, T. (1993). Regulation of myosin isolated from pollen tubes of lily by calcium. Cell Ficnct. 18, 642. [Abstract] Yoshimoto, Y.. and Hiramoto, Y. (1985). Cleavage in a saponin model of the sea urchin egg. Cell Struct. Funct. 10, 29-36. Yoshimoto, Y., and Kamiya, N. (1984). ATP and calcium-controlled contraction in a saponin model of Physarum polycephalum. Cell Struct. Funct. 9, 135-141. Yoshimoto, Y., Matsumura, F., and Kamiya, N. (1981a). Simultaneous oscillations of Ca2+ efflux and tension generation in the permeabilized plasmodia1 strand of Physarum. Cell Motil. 1,432-443. Yoshimoto, Y., Sakai, T., and Kamiya, N. (1981b). ATP oscillation in Physarum plasmodium. Protoplasma 109, 159-168. Yoshimoto, Y., Iwamatsu, T., and Hiramoto. Y. (1985). Cyclic changes in intracellular free calcium levels associated with cleavage cycles in echinoderm and medaka eggs. Biomed. Res. 6, 387-394. Yukawa, C., Yokota, E.. Sonobe, S., Mutoh, T.. and Shimmen, T. (1997). Two types of myosin in tobacco cultured cell BY-2. Cell Struct. Ficnct. 22, 704. [Abstract]

4.

Characteristics of Skeletal Muscle in Mdx Mutant Mice Sabine De La Porte,* Sophie Morin,t and Jeanine Koenigt,' *Laboratoire de Neurobiologie Cellulaire et MolCculaire. CNRS UPR 9040, 91198 Gif sur Yvette Cedex, France; and tLaboratoire de Neurobiologie Cellulaire, Universite de Bordeaux 11, France

We review the extensive research conducted on the mdx mouse since 1987, when demonstration of the absence of dystrophin in mdx muscle led to X-chromosome-linked muscular dystrophy (mdx) being considered as a homolog of Duchenne muscular dystrophy. Certain results are contradictory. We consider most aspects of mdx skeletal muscle: (i) the distribution and roles of dystrophin, utrophin, and associated proteins; (ii) morphological characteristics of the skeletal muscle and hypotheses put forward to explain the regeneration characteristic of the mdx mouse; (iii) special features of the diaphragm; (iv) changes in basic fibroblast growth factor, ion flux, innervation, cytoskeleton, adhesive proteins, mastocytes, and metabolism; and (v) different lines of therapeutic research. KEY WORDS: Mdx, Skeletal muscle, Dystrophin, Utrophin, Regeneration, Basal lamina, Fibroblast, Nuclear magnetic resonance. 0 1999 Academic Press.

1. Introduction Approximately 700 neuromuscular diseases have been described in man and are dominated by those with primary muscle involvement. Of the latter, Duchenne muscular dystrophy (DMD), a recessive X chromosome-linked disease, is the most frequent (1 boy in 3500) and severe (lethal around the age of 20). DMD was first described in the 1860s by the French physiologist G. Duchenne of Boulogne. It is characterized by elevated serum levels of muscle enzymes (more than SO times higher than normal) and by progressive loss of muscular strength, perceptible from the age of 4 or 5. Death due

' Preserir address: Institut de Myologie, Groupe Hospitalier PitiC-Salpitriirre, Paris, France. Irirernurronid R ~ w i r i v(I! ('yrolo~v,Vol. lYI

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to respiratory or cardiac insufficiency generally occurs around age 20. This myopathy is sometimes associated with mental retardation. A less severe and more variable form of the disease, Becker muscular dystrophy (BMD), was described in 1955. This myopathy is 10 times less frequent than DMD. The gene responsible for these myopathies was localized on the X chromosome in the early 1980s by several research teams and was isolated in 1986 (Monaco et af., 1986). This 1800 kb gene is located on the short arm of X chromosome, at Xp21. It comprises exons of mean size 150 pb separated by introns of about 16 kb (Chelly and Kaplan, 1988). It is the largest of all the genes identified to date (Koenig et al., 1987); the second largest, which is 10 times shorter, is that of neurofibromatosis. It accounts for between 0.05% (Brown and Hoffman, 1988) and 0.1% (Hoffman and Kunkel, 1989) of the human genome and one-third of the Escherichia coli genome (Brown and Hoffman, 1988). The corresponding protein, dystrophin, was identified in 1987 (Hoffman et al., 1987a). It is produced in all types of muscle tissues (cardiac and skeletal striated muscle and smooth muscle) and in certain neurons (Hoffman et al., 1988). Dystrophin is large (Mr427kDa, i.e., six times that of hemoglobin) and shares many features with spectrin (Koenig et af., 1988), a cytoskeletal protein which contributes to the plasticity of the membranes of red blood cells. It is thought that dystrophin contributes to the stability of muscle cell membranes. The discoveries of the DMD gene and of dystrophin were followed by characterization in animal models of muscular dystrophies genetically identical to those in humans. The discovery in the mouse (Bulfield el af., 1984), dog (Cooper et al., 1988), and cat (Carpenter et al., 1989) of homologous mutants with the same genetic deficiency as DMD led to a new experimental approach to muscular dystrophies. These mutant animals, however, do present specific differences.

II. Animal Models

A. Mdx Mouse In 1984, Bulfield et af., who were seeking glycolysis metabolism mutants in mice, discovered some animals of the C57BL/10 strain that had abnormally elevated levels of serum pyruvate kinase and creatine kinase, suggesting the existence of a muscular dystrophy. Genetic observations established that the disease was linked to X chromosome, and histological and ultrastructural examinations revealed primary lesions of muscle fibers, without

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nervous system involvement. The mutant was named mdx (X-linked muscular dystrophy) and in 1987 the finding that its muscles lacked dystrophin suggested it was a homolog of DMD (Monaco et af., 1986; Brockdorff et af., 1987; Heilig et af., 1987; Hoffman et al., 1987a). The mdx mouse has a mutation in the dystrophin gene and so its muscle fibers degenerate following membrane damage, resulting in particular in calcium uptake by the cell, overcontraction of muscle fibers, and activation of intracellular proteases. Unlike the situation in DMD, in the dystrophindeficient mouse, there is minimal fibrosis and fatty tissue replacement, and cellular necrosis is long compensated by regeneration of muscle fibers. The molecular aspects of the mutation were studied by Sicinski et af. (1989): The replacement of a cytosine by a thymine at position 3185 of the sequence coding for the gene leads to the appearance of a stop codon which prematurely terminates the translation of dystrophin. The truncated protein (about 27% of the normal length) comprises the N-terminal domain and seven of the repetitive sequences of the central domain. The rest is absent, but it is difficult to know whether the residual protein product is stable and functional.

B. The GRMD Dog

In 1986 and 1988, dogs developing progressive muscular dystrophy linked to X chromosome with absence of dystrophin were described (Valentine et af.,1986; Kornegay et af., 1988). These dogs have a mutation which results in a change in reading phase and a premature stop codon in the gene (Sharp et al., 1992). The creatine kinase level is elevated from the first week onwards, but the disease, called canine X-linked muscular dystrophy or GRMD (golden retriever muscular dystrophy), is only present around the age of 8 weeks as muscular weakness, which worsens progressively over several months. As in humans, the muscular weakness of the dogs is paralleled by a progressive loss of muscle tissue and its replacement by fibrous tissue (fibrosis). The anomalies affect the entire musculature, and in particular reduce respiratory capacity. Death generally occurs around 1 year of age (Valentine et af., 1992). In GRMD, 1% of the fibers are dystrophin positive (Nudel, 1989). As in DMD muscle, basic fibroblast growth factor (bFGF) is present only in small amounts, essentially in the nuclei and sarcoplasm (Anderson et af., 1993). Likewise, levels of mastocytes involved in development of fibrosis rise sharply and pass from the perimysium to the endomysium. This change in mastocyte distribution strictly parallels the clinical stages of the disease (Gorospe et af., 1994).

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C. The FXMD Cat In 1989, Carpenter et al. described two 2-year-old cats with walking difficulties due to a myopathy, despite the absence of progressive muscular weakness. Dystrophin was lacking, but the genomic deletion could not be defined. Serum creatine kinase levels were much above normal. This disease was given the name FXMD (feline X-linked muscular dystrophy). Unlike DMD and GRMD, dystrophin deficiency in FXMD cats is characterized by muscular hypertrophy. These animals never lose muscle tissue and conserve a certain strength, Moderate fibrosis occurs but there is no fatty infiltration. To explain these between-species differences, Hoffman et al. (1987a) hypothesized that in the human and dog, development of fibrous instead of muscle tissue prevents the muscle from regenerating and remaining functional. Another hypothesis is that in affected muscles in the human and dog, constant deterioration of muscle fibers creates a great demand for myoblasts, such that the muscle gradually loses its ability to regenerate. In smaller animals (mice and cats), the muscular demands would be less and the “pressure” on the myoblasts would be lower. Nonetheless, it is clear that fibrosis plays a central role in disease progression in humans (Hoffman, 1993).

111. Dystrophin, Utrophin, and Associated Proteins

A. Dystrophin 1. Structure Dystrophin has a multidomain structure (Brown and Hoffman, 1988; Mandel, 1989). From the NH2 terminal to the COOH terminal, there is an actinbinding region, followed by repetitive spectrin-like motifs, a cysteine-rich zone suggestive of calcium-binding motifs, and a domain penetrating in the plasma membrane and interacting with glycoproteins (Fig. 1): The N-terminal domain (240 amino acids), analogous to those of aactinin (Slater, 1987; Chelly and Kaplan, 1988; Hoffman et al., 1989) and p-spectrin (Byers et al., 1989), binds actin cooperatively (Way et a/., 1992) at two binding sites (Levine et al., 1992). The central rod-shaped domains consist of 24 (or 25, depending on the authors) repetitive structural elements of variable length (about 100 amino acids), similar to repeated triple-helix elements of a-actinin and @-spectrin(Man et al., 1990), and they combine to form filaments. Four proline-rich hinges in this domain confer elasticity and flexibility on

Actin

FIG. 1 Schematic model of the dystrophin-glycoprotein complex as a transsarcolemmal linker between the subsarcolemmal cytoskeleton and the extracellular

matrix (reproduced with permission from the Association Franpise contre les Myopathies).

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dystrophin (Koenig and Kunkel, 1990). Deletions truncating this region reduce the extensibility of the molecule without affecting actin binding, which characterizes BMD (Cross et al., 1990). The cysteine-rich domain (142 amino acids) exhibits 24% homology with the C-terminal domain of a-actinin and has two calcium-binding sites. The 420-amino-acid C-terminal domain has no sequence similar to that of a known protein; it penetrates the membrane of the muscle fiber and interacts with the membrane glycoproteins (LCger et al., 1991). The monomers of dystrophin are rods 3 or 4 nm thick and 175 nm long (Pons et al., 1990). These monomers combine spontaneously in antiparallel homodimers by matching of central repetitive domains, forming a hexagonal network such as erythrocyte spectrin (Koenig, 1989). The N and C termini are therefore juxtaposed and associate with one another or with other dimers of dystrophin (Hoffman and Kunkel, 1989). The dystrophin molecules are parallel (or almost so) to the inner face of the plasma membrane. The N and C termini and the plasma membrane are each about 50 nm apart. The dystrophin molecules are linked to the most peripheral of the filaments of sarcoplasmic actin (Wakayama et al., 1993). The nucleotide sequences in humans and mice have a highly conserved size and organization and marked homology (Robertson, 1987; Beam, 1988; Chelly and Kaplan, 1988). In the chicken, the sequence encoding dystrophin is almost the same size as its human counterpart. Conservation is 80% for the N-terminal region, 75% for the spectrin-like domain, and 95% for the C-terminal domain, suggesting that this is an important region for interactions with other proteins (Lemaire et al., 1988). Dystrophin represents 0.002% of total muscle protein (Campbell and Kahl, 1989) and 5% of the membrane cytoskeleton in skeletal muscle (Ohlendieck and Campbell, 1991a). Although dystrophin is a minor muscle protein, it is a major constituent of the muscle membrane in which it plays an important structural role. 2. Expression

Dystrophin is present in fetal and adult skeletal, cardiac, and smooth muscles (Robertson, 1987). It is expressed to equal extents in skeletal and cardiac muscle cells and in the brain (Brown and Hoffman, 1988) and to a lesser degree in smooth muscle (Beam, 1988). In the normal mouse embryo, dystrophin is visible from the 13th day in skeletal muscle (Karpati, 1989). Dystrophin expression has been extensively studied in human muscle. At the embryonic stage (8 or 9 weeks), it is first localized in the sarcoplasm at the ends of the myotubes, beside the tendons. At the fetal stage, it is distributed throughout the myofiber. By the 22nd


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week, it is localized in the sarcolemma of most fibers. This suggests that dystrophin accumulates in the cytoplasm before being associated with the membrane (Van Ommen, 1989; Prelle et al., 1991; Wessels et al., 1991; Clerk et al., 1992). Some authors consider that dystrophin content depends on muscle type. It has been reported that there is more dystrophin in slowtwitch than fast-twitch fibers in the rat and mouse, although this difference is not reflected in the dystrophin mRNA (Ho-Kim and Rogers, 1992). In contrast, Koga et al. (1992) found no significant difference in the dystrophin contents of slow and fast muscles. In rat heart dystrophin is undetectable on the 15th embryonic day, the stage at which the heart is able to generate action potentials and beat spontaneously. Development of the first functions of the myocardium therefore does not require the presence of dystrophin. A small quantity is detected on the 17th embryonic day and this increases during the perinatal period (Tanaka and Ozawa, 1990a). Cardiac work increases at this stage. Because its expression rises before these changes, dystrophin may be necessary for rapid and large contractions of the myocardium. Adult levels of dystrophin are reached 2 weeks postnatally (Tanaka and Ozawa, 1990b). Dystrophin is localized at the membrane surface of the Purkinje cells (Bies et al., 1992). Dystrophin is also found in cultured human skin fibroblasts. The promoter used in the muscle tissue is responsible for the transcription observed in fibroblasts (Hugnot et al., 1993). Dystrophin is also expressed by the neurons (Hoffman and Kunkel, 1989), notably in the cerebral cortex and cerebellum (Lidov et al., 1990). Dystrophin-like immunoreactivity is uniformly expressed in several regions of the brain involved in learning (hippocampus and cerebral cortex) and in motor function (spinal medulla, cerebellum, thalamus, and substantia nigra) (Huard et al., 1992a). It is localized in the soma and dendrites, associated with the inner part of the membrane (Lidov et al., 1990; Torelli et al., 1992). Brown and Hoffman (1988) reported that dystrophin is not expressed in vivo or in vitro in tissues other than muscle and nerve, whereas Walsh et al. (1989) detected dystrophin in minute amounts in all normal tissues.

3. Localization Biochemical and electron microscopy studies established that dystrophin is abundant in membrane fractions of skeletal muscle, at the inner face of the membranes, and in T-tubules (Hoffman et al., 1987b; Watkins et al., 1988). These authors therefore hypothesized that dystrophin was associated with the triads (Hoffman etal., 1987b; Chelly and Kaplan, 1988). ZubrzyckaGaarn et al. (1988) then showed that dystrophin was associated with the sarcolemma rather than with the triad. In rat soleus muscle regenerating after damage, dystrophin-type reactivity is first apparent in the sarcolemma

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and various inner membranes and, 4 weeks after the damage, in the triads. Dystrophin therefore appears primarily cytoplasmic, and then associates with the sarcolemma and the T-tubules during maturation. In mature fibers, dystrophin levels in the T-tubules appear to be so low that they can only occasionally be seen by means of immunocytochemistry (Bornemann and Schmalbruch, 1991). It is now considered that dystrophin is essentially localized at the inner face of the sarcolemma. Dystrophin molecules are not uniformly spread over the surface of mammalian muscle fibers. Confocal microscopy reveals aligned fluorescent points or intermittent lines along the sarcolemma, with most fluorescent points corresponding to the I bands (Masuda et af., 1992; Porter et al., 1992). The dystrophin molecules form a network of dense transverse rings or costamers and fine longitudinal interconnections. The dystrophin network at the surface of the muscle fiber is organized in relation with the contractile apparatus. Likewise, vinculin, y-actin, talin, and spectrin constitute a submembrane cytoskeletal network with a costameric distribution (Porter et af., 1992; Straub et af., 1992). Dystrophin binds strongly to talin, and this binding is inhibited by vinculin because of steric hindrance. Dystrophin can therefore interact in vitro with proteins of the muscle cytoskeleton. These proteins constitute additional sites of dystrophin binding to the sarcolemma (Senter et af., 1993). This may explain why dystrophin is found at the sarcolemma even when it lacks its C terminus (Senter et al., 1993). Dystrophin is more abundant at the membrane surface of the intrafusal fibers and at the neuromuscular junction (NMJ) than at the membrane surface of skeletal and cardiac muscle fibers (Arahata and Sugita, 1989; Miyatake et of.,1989; Huard et af., 1992b). At the NMJ, dystrophin is found in the depths of junctional folds and is absent from most AChR-rich domains of the rat NMJ (Byers et af., 1991; Sealock et af.,1991; Yeadon et al., 1991; Huard et af., 1992b). Dystrophin is present in the immediately adjacent membranes, suggesting that it is not an obligatory component of the AChR domains in the muscle; its role at the NMJ may be linked to the organization of the junctional folds (Sealock et af., 1991). Dystrophin also accumulates at the myotendinous junction (Bonilla, 1989; Mandel, 1989; Arahata and Sugita, 1989; Masuda et al., 1992), where it is one of the components binding the terminal actin filaments to the cytoplasmic face of the membrane of the junctional folds of the tendon (Samitt and Bonilla, 1990). In culture, dystrophin is undetectable in myoblasts (Nudel, 1989) but is present in myotubes (Zubrzycka-Gaarn et af., 1988; Ecob-Prince et al., 1989). Dystrophin is localized at talin-positive sites, where the sarcolemma is in apposition to the substratum. The first sites at which dystrophin appears in cultured muscle cells are therefore adhesion sites, i.e., specific sites of


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interactions between the cytoskeleton and the extracellular matrix (Kramarcy and Sealock, 1990). Dystrophin is discontinuously distributed in the submembrane region of human muscle cells in culture. Such cultures do not contract spontaneously but contractions appear in cocultures with rat spinal cord. In contracting conditions, dystrophin occurs continuously along the inner face of the sarcolemma and in periodic dense aggregates. After addition of tetrodotoxin, dystrophin resumes a discontinuous distribution, as in myotubes cultured alone. The contractile activity of the muscle therefore plays an important role in the continuous distribution of dystrophin along the sarcolemma during development (Sklar et al., 1990; Park-Matsumoto et al., 1991). In mdx mice, dystrophin mRNA is of normal size (16 kb) but present in small amounts (25% of the normal level). An mRNA containing a nonsense mutation can be degraded in the nucleus (Monaco et al., 1986; Brown and Hoffman, 1988; Chamberlain et al., 1988). There may be somatic reversion or suppression of the mdx mutation, which explains the observation of some dystrophin-positive fibers by immunofluorescence (Hoffman et al., 1990). Dystrophin-like immunoreactivity is detected in the fetal muscles of mdx mice, perhaps due to alternative splicing, which is quite often specific to the different stages of development. Hence, in the fetus, the stop codon is eliminated, thereby allowing synthesis of a dystrophin-type protein, whereas in the adult the stop codon is retained, thereby blocking translation (Kahn, 1989). Other authors consider that in the mdx mouse fetus there is synthesis of an abnormal dystrophin which cannot be integrated into the membrane and is therefore degraded (Van Ommen, 1989). Mdx mice have structural defects at the myotendinous junction-notably reduced lateral associations between the membrane and the fine filaments. These anomalies are observed before the onset of necrosis and may therefore be directly correlated with the absence of dystrophin (Law and Tidball, 1993). Tissues other than skeletal muscle are also affected. The contractile properties of the myocardium are markedly altered (Sapp et al., 1996), which is consistent with the hypothesis that dystrophin deficiency affects cardiac contractile function. The total absence of dystrophin in the cerebellum is not reflected in particular clinical signs (Huard and Tremblay, 1992). Nevertheless, in vivo and in vitro H' magnetic resonance spectroscopy revealed an increase in choline compounds and myoinositol levels, indicative of gliosis or developmental abnormalities in dystrophic brain (Tracey et al., 1996b). Systemic dysfunction may occur in smooth muscles (blood vessels) and skeletal muscle fibers. This may explain certain clinical symptoms of DMD, such as respiratory and gastrointestinal disorders (Miyatake et al., 1989). The mdx mouse expresses the Dp71 isoform of the dystrophin gene in its brain and in other nonmuscular tissues (Rapaport et al., 1992).

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B. Glycoprotein Complex Associated with Dystrophin Dystrophin is associated with a complex of glycoproteins and membrane proteins respectively called dystrophin-associated glycoproteins (DAGs) and dystrophin-associated proteins (DAPs) (Fig. 1). The large oligomeric complex with dystrophin comprises three subcomplexes (Ervasti and Campbell, 1991; Ohlendieck and Campbell, 1991b; Ibraghimov-Beskrovnaya er al., 1993; Ozawa et a[., 1995):

Dystroglycan complex: a-dystroglycan (156 kDa DAG), P-dystroglycan (43 kDa DAG) Sarcoglycan complex: a-sarcoglycan (adhalin or SO kDa DAG), p-sarcoglycan (43 kDa DAG), y-sarcoglycan (35 kDa DAG), Ssarcoglycan (35 kDa DAG) (Yoshida et al., 1997), and sarcospan, a unique 25-kDa member of the complex (Crosbie et al., 1997) Syntrophin complex: a-syntrophin (59 kDa DAG), P,-syntrophin (59 kDa DAP), &-syntrophin (59 kDa DAP) Dystrophin, the syntrophin complex, a-dystroglycan, and a muscle isoform of dystrobrevin (78 kDa) (Blake et al., 1996a) are peripheral membrane proteins, whereas the sarcoglycan complex and P-dystroglycan are integral membrane proteins. The syntrophin complex links sodium channels of the membrane to the actin cytoskeleton (Gee et al., 1998). a-Dystroglycan binds to laminin, thereby forming a link between the sarcolemma and the extracellular matrix (ECM) (Ibraghimov-Beskrovnaya et al., 1992). This link is calcium dependent and inhibited by heparin. One of laminin’s heparin-binding domains is therefore involved in binding to dystroglycan. P-Dystroglycan and a syntrophin are associated with dystrophin (Jung et d., 1995). There is no interaction between the dystrophin/glycoproteins complex and fibronectin, type I and IV collagens, entactin, or heparan sulfate proteoglycan (HSPG). However, the dystrophin/glycoproteinscomplex cosediments with actin and may therefore play a part in the binding of cytoskeletal actin to the ECM (Ervasti and Campbell, 1993). a-Syntrophin is also complexed with neuronal nitric oxide synthase (nNOS) (Brenman et al., 1996) in normal muscle. NOS is not anchored to the skeletal muscle sarcolemma of mdx and DMD muscle, but it is mislocalized to the interior of the muscle fibers (Brenman et al., 1995; Chang et al., 1996). NOS has been reported as present in the sarcolemma of fast-twitch or both fast- and slow-twitch fibers (Kobzik et at., 1994; Grozdanovic et al., 1996). The hypothesis that free radical toxicity is due to the mislocalization of nNOS is now excluded: Transgenic mice devoid of both nNOS and dystrophin present the same dystrophic characteristics as mdx mice (Crosbie et al., 1998). NOS is also concentrated at the NMJ (Kusner and Kaminski,


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1996; Oliver etal., 1996), where its role is unclear, except in possible involvement in synaptic suppression (Wang et al., 1995). In the model proposed by Minetti et al. (1992), a cytoskeletal structure links a-actinin of the Z-line of the sarcomere to the plasma membrane via F-actin, dystrophin, and associated glycoproteins. Complex molecular interactions involving other cytoskeletal proteins (vinculin, talin, and integrin) maintain the structural integrity of the plasma membrane. Lastly, a dystroglycan acts as a link between the plasma membrane and the ECM (Ibraghimov-Beskrovnaya et al., 1992). In the mdx mouse, all dystrophin-associated proteins are greatly reduced (80-90%) in the muscle due to the absence of dystrophin and not to secondary effects of the degradation of muscle fibers. This loss could have secondary effects on basal lamina components (Ohlendieck and Campbell, 1991b). There is an 85% reduction in a- and 0-dystroglycan in mdx muscle (Ervasti et al., 1990), whereas their mRNA is expressed at normal levels (Ibraghimov-Beskrovnaya et a/., 1992). Similarly, a-sarcoglycan mRNA is present in mdx muscles, as in DMD muscles, but the protein is greatly reduced (Roberds et a/., 1993). Regulation of these proteins is therefore posttranslational in these muscles. DAGs are produced, but in the absence of dystrophin they are not correctly assembled and/or integrated in the sarcolemma and are degraded (Matsumura and Campbell, 1994). Following surgical or pharmacological denervation of skeletal muscles of mdx mice, the levels of adhalin and P-dystroglycan increased at the extrajunctional sarcolemma together with AChR, suggesting that their association is independent of the presence of dystrophin (Mitsui et al., 1996). C. Utrophin

A 13-kb autosomal transcript encoded by a gene of the long arm of chromosome 6 has been identified in human fetal muscle. It encodes a protein with more than 80% homology with dystrophin (395 kDa) (Love et a/., 1989); thus because of its ubiquitous localization, it is called dystrophinrelated protein (DRP) or utrophin (Fig. 2), The homology between dystrophin and utrophin extends over their whole lengths, suggesting that they derive from a common ancestral gene. Utrophin, like dystrophin, binds actin. The C-terminal domain is highly conserved (Tinsley et al., 1992). In the mouse, the utrophin locus is on chromosome 10. This gene is expressed to varying degrees in numerous tissues; the transcript is particularly abundant in several human fetal tissues (e.g., heart, placenta, and intestine). In normal human and murine muscles, utrophin is always found at the membrane surface of immature fibers (Zhao et al., 1993). It is concentrated at the NMJ of mature muscle fibers and reappears at the membrane surface

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FIG. 2 Schematic model of the utrophin-glycoprotein complex as a transsarcolemmal linker between the subsarcolemmal cytoskeleton and the basal lamina. ARIA, acetylcholine receptor-inducing activity; CGRP, calcitonin gene-related peptide; MASC, MuSK accessoryspecific component; MuSK, muscle-specific receptor tyrosine kinase; RATL, rapsyn-associated transmembrane linker (reproduced with permission from the Association FranGaise contre les Myopathies).

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of muscle fibers after denervation. Its localization is similar to that of the AChR, at the tops of the postsynaptic folds (Bewick et al., 1992; Appel and Merlie, 1995). Utrophin mRNA selectively accumulates within the postsynaptic sarcoplasm of adult muscle fibers (Gramolini et al., 1997). Utrophin may be one of the molecules of the cytoskeleton which organize and stabilize the cytoplasmic domain of the AChR (Takemitsu et al., 1991a, b). It is present at the earliest stages of the concentration of AChRs at the NMJ in the mouse embryo and is also concentrated in the large AChR clusters on myotubes of the C2 line, suggesting that it is mainly involved in the growth of AChR clusters (Phillips et al., 1993). Utrophin-deficient mice are healthy and show no signs of weakness. However, their NMJs have reduced numbers of AChR and decreased postsynaptic folding (Deconinck et al., 1997a; Grady et al., 1997a). Therefore, utrophin alone is not essential for AChR clustering at the NMJ. Utrophin is associated with laminin (Khurana et al., 1995) via a complex of sarcolemmal proteins identical to or at least antigenically similar to that of dystrophin (Fig. 2) (Matsumura et al., 1992). In the mdx mouse, various authors have shown that utrophin is expressed at the NMJ and is found in muscle extracts from this region (Anderson et al., 1990; Fardeau et al., 1990; Pons et al., 1991). Utrophin and associated proteins colocalize at the NMJ (Karpati et al., 1993a), where utrophin is overexpressed (Takemitsu et al., 1991a, b; Koga et al., 1993). Outside the NMJ, the results are discordant. Anderson et al. (1990), Fardeau et al. (1990), and Pons et al. (1991) report that utrophin is absent from extrasynaptic zones. Karpati et al. (1988) found utrophin along the whole sarcolemma of mdx muscle small-caliber fibers and of cardiac fibers. Because these muscles exhibit few pathological changes, utrophin may compensate for the absence of dystrophin by holding the DAGs at the extrajunctional sarcolemma (Matsumura et al., 1992; Tinsley et al., 1992). Takemitsu et af. (1991a, b) and Koga et al. (1993) consider that utrophin is present at the membrane surface of all mdx muscles. Contradictory results have emerged from studies of utrophin expression during development. All authors agree that utrophin is overexpressed in fetal mdx muscle (Takemitsu et al., 1991a, b; Koga et al., 1993; Zhao et al., 1993). Some authors consider that this overexpression continues after birth (Takemitsu et al., 1991a, b; Koga et al., 1993; Zhao et al., 1993) and that utrophin expression in the adult mdx animal is twice that in normal mice (Sugita et al., 1993; Law et al., 1994). Law and colleagues, however, did not find this increase in 2-week-old mice. Other authors have found that utrophin decreases in fast muscles before the third week and disappears in slow muscles between the third and fourth week. Love et al. (1991) identified utrophin in many fetal tissues, including the heart, placenta, and intestine. In the nervous system of the mdx mouse,

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utrophin is present in the vascular walls, pia mater, and choroid plexus of the brain but not in neuronal cells (Ishiura et al., 1990; Uchino et al., 1994).

D. Function of Dystrophin, Utrophin, and Associated Proteins Although dystrophin is a cytoskeletal protein located on the inner face of the plasma membrane of the muscle fiber, its function is still incompletely understood. It seems to be involved in maintenance of the morphological and functional structure of the striated muscle fiber (Petrof, 1998). Two main hypotheses concerning its role have been proposed. The first attributes a mechanical function to dystrophin, which is seen as the main element of an elastic net or sort of lattice within the membrane of the muscle fiber. The molecules “slide” over one another, thereby allowing the whole net to change shape during contraction and relaxation (Watkins, 1989; LCger et al., 1991). By connecting the cytoskeleton to the cytoplasmic face of the muscular membrane (Zubrzycka-Gaarn et al., 1988), dystrophin may help resist the stresses that develop during contraction (Beam, 1988). It may therefore play an important role in preservation of membrane stability (Park-Matsumoto et al., 1992). Indeed, dystrophin is located very close to the plasma membrane of the muscle fibers. The N terminus is linked to the cytoskeleton’s actin network and the C terminus to the plasma membrane. In terms of the glycoprotein complex, dystrophin is supposed to play an important role in transduction of the mechanical force of the contractile apparatus to the ECM (Straub et al., 1992; Petrof et al., 1993a). According to the second hypothesis, dystrophin is involved in calcium homeostasis: It binds the contractile filaments of the internal membrane system, thereby ensuring a link between the membranes which release calcium and the contractile proteins which are activated by this calcium (Slater, 1987; Hoffman et al., 1987a; Tay et af., 1989). Dystrophin may also play an important physiological and/or structural role in cell motility (Miyatake et al., 1991). At the synapse, dystrophin may participate in the formation of a submembrane cytoskeletal network which promotes AChR clustering (Jasmin et af., 1990). At the myotendinous junction, it may form lateral associations between the fine filaments and the cell membrane (Tidball and Law, 1991; Law and Tidball, 1993). Dystrophin’s localization in the central nervous system indicates a physiological function in the conduction of the nerve impulse rather than a mechanical function (Yoshioka et af., 1992). It may be involved in the structure and maintenance of the dendritic tree and neuronal survival (Huard et af., 1992a).


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Utrophin may have a function similar to that of dystrophin during development (Anderson and Kunkel, 1992). It may also play an important role in the organization of the postsynaptic membrane of the NMJ (Ohlendieck et al., 1991). a-Dystroglycan constitutes an agrin-binding site at the surface of the muscle cell. With DAG 50 kDa and utrophin, it colocalizes with agrininduced AChR clusters. It may therefore be involved in AChR clustering (Gee et al., 1994). One hypothesis is that agrin stabilizes the membrane cytoskeleton specific to the synapse, which in turn serves as a support in the concentration of synaptic molecules (Campanelli et al., 1994). The role of a-dystroglycan remains unelucidated (Sugiyama et al., 1994) since Glass et al. (1996) demonstrated that the main component of the receptor complex which mediates agrin signaling is a muscle tyrosine kinase (Musk).

IV. Mdx Muscle Cells A. Degeneration/Regeneration of t h e Muscle

1. Morphological Defects Mdx muscles exhibit early necrosis. From the first day, there is disorganization of the Z-line and hence of the contractile apparatus. Necrosis starts on Day 5 , but only in the muscles of the head, trunk, and girdle. The limbs are affected later (Torres and Duchen, 1987). In the first 10 days following birth, all the limb muscle fibers seem relatively normal. The necrotic fibers start to appear at 21 days and become numerous by Day 28. A lack of synchronization has been described in the course of the disease: a craniocaudal gradient in the appearance of necrosis followed by regeneration (Muntoni ef al., 1993). The degeneration and regeneration of the mdx muscle occurs essentially between the third and 10th week, with marked differences depending on the authors: Onset of degeneration around 2 weeks, with a peak between Weeks 3 and 8 (Karpati, 1989; Karpati et al., 1990; Nagel et al., 1990) Onset between Weeks 3 and 5 (Brown and Hoffman, 1988; Coulton et al., 1988) Peak at 6 weeks (DiMario and Strohman, 1988) Peak between Weeks 5 and 8 and complete regeneration around 10 weeks (Torres and Duchen, 1987; DiMario et al., 1989) Muscle fibers whose girth is below 20-25 p m in diameter (such as extraocular muscle fibers) are not susceptible to necrosis (Karpati et al., 1988). In

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such muscle fibers there is no significant differences in nNOS concentration and distribution compared to other mdx muscles (Wehling et al., 1998). Apoptosis precedes any detectable necrotic change in mdx muscle, and apoptotic events continue into the stage of necrosis (Tidball et al., 1995; Smith et al., 1995). Overcontracted fibers are seen from birth in mdx muscles (Torres and Duchen, 1987) and at least to the age of 18 months. They reflect an early stage of necrosis, due to the massive influx of calcium in a segment of the muscle fiber, provoking abnormal contraction of the sarcomeres. Calpains (neutral calcium-dependent thiol proteases) are autolytically cleaved during the disease process of mdx dystrophy, indicating that they may be activated and play a role in the proteolysis that occurs in muscle (Spencer et aL, 1995). The fragility of skeletal muscle fibers of mdx mice is attested by (i) a significant accumulation of Evans blue, a tracer molecule used to analyze sarcolemmal integrity (Straub et al., 1997), and (ii) a significant and transient rise in the level of creatine kinase and P-galactosidase (by using a construction with a muscle-specific promoter in trangenic mdx mice) in the serum of mice after eccentric running exercise (Vilquin et al., 1998). Fast-twitch fibers are preferentially but not exclusively involved (Carnwath and Shotton, 1987). Only 5% of the original fibers survive to 26 weeks in EDL muscle (fast). Muscle destruction is delayed in the slow-twitch soleus muscle and in the fast-twitch gastrocnemius compared to the intermediate tibialis anterior muscle (Dangain and Vrbova, 1984). The presence of intermediate fibers may result from coexpression of slow and fast myosin heavy chains, indicating a transition from fast to slow in regenerating fibers (Pastoret and SCbille, 1993b). The onset of muscular hypertrophy in the mdx mouse can be explained by three nonexclusive hypotheses: hypertrophy of the fibers (arise in fiber diameter and therefore in muscle bulk), hyperplasia (fiber neosynthesis), or an abnormal increase in connective tissue. In the adult mutant, none of these hypotheses alone can account for the increases in weight and muscle area. This hypertrophy peaks between 6 and 12 months of age and then regresses such that animals more than 18 months of age show substantial muscular atrophy (Pastoret and SCbille, 1993a). These animals exhibit marked fibrosis in some muscles, notably the largest proximal muscles (Hoffman and Kunkel, 1989). The contractile function of muscles of old mdx mice displays many similarities to that of DMD (Hayes and Williams, 1998), but the results of Bobet et al. (1998) disagree with this hypothesis. It is a moot point whether regeneration of mdx muscles is transient or continuous. Strohman’s team (Berkeley, CA) has shown that mdx muscle expresses fetal and neonatal myosin mRNA from Weeks 6 to 16, but that from Weeks 10 to 56 the frequency of the fibers expressing embryonic myosin decreases to about 1%(DiMario et al., 1991). This suggests that


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mdx muscle regeneration is transient. Conversely, other studies suggest that regeneration of mdx muscle is a continuous process: (i) Beilharz et al. (1992) have shown that the expression of MyoD and of myogenin, which are early and specific markers of regeneration, is elevated between 3 and 6 weeks. It then decreases to a constant, albeit relatively high, level; (ii) McGeachie et af. (1993) have shown that proliferation of muscle cells starts at about 3 weeks, peaks between 4 and 8 weeks, and continues at a lower level at least to Week 44; and (iii) necrotic foci are present throughout the life of the animal (MacLennan and Edwards, 1990; Pastoret and SCbille, 1995) but are compensated by regeneration up to 18-20 months. The number of fibers with central nuclei, i.e., fibers that have been necrotized and have regenerated, increases with age (Torres and Duchen, 1987; Karpati et af., 1990), and there is great variation in fiber diameter (Torres and Duchen, 1987). From 3 months, 80-90% of these fibers of the skeletal muscles of the mutant have a central nucleus (Carnwath and Shotton, 1987). However, the formation of such fibers appears to predominate during the acute phase of the disease, whereas peripheral relocalization of the nuclei becomes the major event during the chronic phase. The central localization of the nuclei in mdx muscle which has regenerated is not permanent but is more lasting than that in other studied species (McGeachie et al., 1993). The pattern of expression of various myogenic regulatory factors (MyoD, myogenin, Myf-5, and Myf-6) differs in mdx regenerating muscle fibers from that in developing muscle in embryos (Bhagwati et al., 1996). Leukemia inhibitory factor (LIF) and interleukin (IL-6) have been shown to promote the proliferation of myoblasts. Normal muscle rarely expresses mRNA for these two molecules, but mdx muscle expresses both LIF and IL-6 mRNA (Kurek et al., 1996). 2. Basal Lamina

The ECM of skeletal muscle consists of a fibrillar matrix, which essentially contains collagen I, 111, and V (Duance et al., 1977) and fibronectin (Chiquet et al., 1981), and a basal lamina, consisting particularly of type IV collagen, laminin, fibronectin, entactin, and HSPG (Kefalides et al., 1979; Hoover et al., 1980). Laminin is a heterotrimer (Vachon et al., 1996) composed of a heavy chain (a;400 kDa) and two similar but not identical light chains ( p and y ; 205-220 kDa). The classical laminin, laminin-1, is composed of an a1 chain and two p , I y I subunits. It has recently been demonstrated that laminin a1 is not found in muscle basal lamina of developing and adult mice (Patton et al., 1997; Tiger and Gullberg, 1997). The predominant laminin variants in the muscle fibers contain the a2 heavy chain (two fragments of 300 and 80 kDa) and are known as merosin. In muscle basal

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lamina, the a2chain associates predominantly with pl/yl chains as laminin2, whereas it associates with the p2/yIchains as laminin-4 at NMJs. The signal-transducing receptors for laminins, collagens, and other extracellular matrix proteins are integrins, a family of heterodimeric (ap)transmembrane proteins (Hynes, 1992; Sonnenberg, 1993). Different isoforms are specifically distributed in synaptic and extrasynaptic zones (Martin et al., 1996). The expression of laminin pI/yIchains and HSPG (by immunofluorescence staining and immunoblot quantification) is increased in mdx muscle no matter what the age of the mice and not just during the acute phase of the disease (Morin et al., 1993). This is in agreement with previous reports of a marked accumulation of collagen, reticulin, fibronectin, and laminin (Marshall et al., 1989; Goldspink et al., 1994; Prattis et al., 1994; Seixas et al., 1994; Quiricosantos et al., 1995). This increase in laminin a l l y l chains seems to be specific since Arahata et al. (1993) found that the laminin cyz chain is not affected. An increase in both these basal lamina components in mdx mice could contribute in different ways to the intense muscle regeneration observed in this mutant. Components of the extracellular matrix are thought to facilitate muscle regeneration (Vracko and Benditt, 1972;Sanes et al., 1978). The mechanisms by which the basal lamina influences regeneration may include stimulation of satellite cell division and growth (Ocalan et al., 1988) and fusion (Vachon et al., 1996). The increase in basal lamina HSPG, concomitant with the increase in bFGF in regenerating areas (DiMario et al., 1989; Anderson et al., 1991, 1993; Matsuda et al., 1992), would favor their interaction and HSPG could deliver the growth factor to high-affinity receptors that initiate the stimulation of myogenic cell proliferation and/or differentiation (Rapraeger et al., 1991;Yayon et al., 1991). This phenomenon could be amplified by the greater sensitivity of mdx satellite cells to bFGF (DiMario and Strohman, 1988). The mechanism of the persistent regeneration in mdx muscle is therefore currently unclear. It does seem reasonable that basal lamina components could be partly responsible for a successful regenerative process. The extracellular matrix protein tenascin-C (TN-C), expressed in wound healing and nerve regeneration, is undetectable in normal muscle except at the myotendinous junction but is prominent in degeneratingkegenerating areas of mdx muscle. TN-C staining declines around stable regenerated mdx myofibers (Settles et al., 1996). 3. Muscle Fibroblasts

Fibroblasts were thought by many to be structure-supporting cells with a negligible role in cellular homeostasis. However, the in vitro cultivation of these cells revealed their fundamental role and their interaction with other


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cell types. For instance, growth inhibitory factors were shown to be secreted by fibroblasts maintained in culture (Harel, 1981; Harel et al., 1978). Thirteen-kDa (Hsu and Wang, 1986) and 45-kDa proteins were purified from medium conditioned by dense cultures of 3T3 fibroblasts. The 45-kDa factor completely inhibits growth of chick embryo fibroblasts (Blat et al., 1987, 1989a) and has been shown to be the insulin-like growth factor-binding protein3 (IGF-BP-3) (Blat et al., 1989b). IGF-BP-3 also accumulated to high levels in conditioned medium (CM) of quiescent and senescent human fibroblasts (Goldstein et al., 1991). IGF-BP-4 and -5 were also shown to be synthesized and secreted by fibroblasts (Camacho-Hubner el al., 1992) and to play a role in the regulation of cell proliferation (Fowlkes and Freemark, 1992: Jones et al., 1993; Neely and Rosenfeld, 1992: Reeve et al., 1995). Macieira-Coelho and Soderberg (1993) also described the presence of a 1-kDa glycopeptide with growth inhibitory activity in extracts from normal human fibroblasts. These results suggest that changes in fibroblast functions may have a profound effect on muscle fibers, and in the case of muscle disease the fibroblast may play a role in determining the extent to which a disease phenotype is expressed, even in the presence of a dystrophic genotype. In view of the hypothesis that the regenerative process is successful at least partly because of modifications of muscle fibroblast functions in the mdx mouse, our approach has been to study the properties of mdx and DMD muscle fibroblasts by analyzing their proliferative response in vitro (Morin et al., 1995). We found that fibroblasts taken from human DMD and control muscle had a similar in vitro proliferation capacity. In mdx mice, the study was performed at various ages to determine if there were changes during the course of the disease. We observed a growth arrest of muscle fibroblasts during the acute phase of the disease: This arrest was partial at 6 and 10 weeks but total at 8 weeks (Fig. 3). This is in marked contrast to fibroblasts taken from animals at other ages, which show a normal proliferative capacity. These results suggested that 8-week-old mdx male mouse muscle fibroblasts release a factor that inhibits their own proliferation. Our hypothesis was confirmed using CM experiments. CM from 8-week-old mdx mouse muscle fibroblasts inhibited the proliferation of control fibroblast cultures. This growth inhibitory factor seems to be specific for fibroblasts and when added to control or mdx myoblast cultures it stimulated their proliferation, as did 8-week-old control fibroblast CM. It has previously been shown that proliferating bovine muscle fibroblasts produce a specific “growth factor activity,” distinct from bFGF or plateletderived growth factor, which increases proliferation of bovine myoblasts but not of bovine or 3T3 fibroblasts (Quinn et al., 1990). From our results, it seems that even quiescent 8-week-old mdx mouse muscle fibroblasts produce a myoblast growth factor. The nature of the factor remains to be

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Age (weeks) FIG. 3 Proliferation rates of control and mdx mouse muscle fibroblasts, determined as the slope of the proliferation curve, at 1, 4, 6, 8, 10, and 13 weeks of age. Data points represent the averages of triplicate determination ? SEM. Significant statistical differences are indicated as follows: ***p < 0,001, **p < 0.01, *p < 0.05. The proliferation rate of mouse muscle fibroblasts decreases during the acute phase of the disease, and inhibition is complete in fibroblasts from 8-week-old mdx mice. The proliferation capacity of control fibroblasts was very high at 1 week and decreased with the age of the mice. The reduction, however, was only significant between 1 and 4 weeks. The proliferation capacity of mdx fibroblasts was usually less than that of control fibroblasts. The ability of the fibroblasts to proliferate decreased between 6 and 10 weeks, the period corresponding to the onset of muscle regeneration in mdx mouse muscle. The 8-week-old mdx mouse muscle fibroblasts produce an inhibitor of their own proliferation and a growth factor specific for myoblasts in vitro. If these factors are secreted in vivo, they could directly and indirectly stimulate satellite cell proliferation, thus favoring muscle regeneration (reproduced with permission from Differentiation, Inhibition of proliferation in 8-week-old mouse muscle fibroblasts in vitro, Morin, S., De La Porte, S., Fiszman, M., and Koenig, J., 59, Fig. 3, copyright 0 1995 by Springer-Verlag).

determined. It could be, at least partly, responsible for the mitogenic activity detected in mdx and control muscle extracts (Chen et al., 1994). In addition to this factor, it is well known that growth factors such as FGFs are synthesized by myogenic cells and stimulate their proliferation. One hypothesis was that the growth inhibitory factor could be one of the IGF-BPs. Preliminary results indicate an exclusive expression of IGFBP-5 in muscle fibroblasts from 8-week-old mdx mice. In addition, messenger RNA is increased three-fold in mdx mice compared to controls, and


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the inhibitory effect on fibroblast proliferation is decreased by 37% when mdx fibroblast CM is previously incubated with antibodies against IGFBP-.5 (Zhang et al., 1996). Our results suggest that fibroblasts release a growth factor specific for myoblasts and that 8-week-old mdx mouse muscle fibroblasts release a growth inhibitory factor specific for fibroblasts. If these factors are released in vivo, the growth inhibitory factor may act in an autocrine way to stop fibroblast proliferation, whereas the mitogenic activity could stimulate satellite cell proliferation, thus favoring muscle regeneration. This hypothesis postulates an active role for quiescent fibroblasts in myogenesis and is supported by the three times increase of the number of cells in culture of myogenic cells accumulating around individual living fibers isolated from muscles of 8 week-old mdx mice (Bockhold et al., 1998).

6 . Diaphragm

The diaphragm of the mdx mouse is the only muscle to exhibit marked degeneration, fibrosis, and functional insufficiency similar to that seen in DMD muscles. Because there seem to be no differences in terms of utrophin expression between limb and diaphragm muscles, it is unclear why the diaphragm is severely affected and other muscles are relatively protected (Sugita et al., 1993). Gillis (1997) considers that the mdx mouse diaphragm combines three unfavorable factors: a large proportion of fast oxidative fibers with large diameter, a life-long sustained activity, and forced lengthening during each contraction. The collagen density in the diaphragm is 7 times that in the diaphragm of control mice and 10 times that in the limb muscles of the mdx mouse. These changes are rare before 2.5 days. At 30 days, there are foci of degeneration, necrosis, and regeneration, as in the limb muscles. However, unlike the limb muscles, the diaphragm exhibits progressive degeneration. At 6 months, fiber size varies greatly, necrosis is continuous, and there is substantial proliferation of the connective tissue. At 16 months, there is extensive loss of muscle fibers and marked fibrosis. These divergences between the diaphragm and limb muscles may reflect intrinsic differences in regenerative capacity. However, regeneration persists even at 16 months (embryonic myosin and central nuclei) and in vitro studies of the satellite cells of the diaphragm and different limb muscles indicate identical proliferative capacities. At 18 months, similar but less severe histological changes are seen in muscles involved in respiration (Stedman et al., 1991). As in the limb muscles, in the diaphragm of the 3or 4-month-old mdx mouse there is an increase in the number of fibers coexpressing the myosin heavy chains of slow and fast muscles. Regenerating fibers express embryonic myosin, whereas the number of fast-twitch

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fibers drops sharply. At 22-24 months, fibers with slow-type myosins are abundant, fast-twitch fibers have been eliminated, embryonic myosin is undetectable, and endurance is increased. The mdx diaphragm therefore responds to the progressive degeneration of the muscle by a transition to a slower phenotype associated with greater endurance. This preserves the contractile function and enhances survival of muscle fibers by lowering energy requirements (Petrof et al., 1993b). Some authors have hypothesized that dystrophin is a mechanical transducer which transmits growth stimuli from the skeleton to the muscle. Dystrophin controls the rate of addition of sarcomeres to the ends of the fibers by participating in the mechanism which allows the satellite cells to fuse with the muscle fibers. In the mdx mouse, the most acute phase of the disease corresponds to the stage at which, in controls, the rate of addition of sarcomeres to the ends of the muscle fibers is maximal. Likewise, in the diaphragm of elderly mdx mice, the fibers are 35% shorter, which supports the hypothesis that dystrophin plays a key role in the regulation of the longitudinal growth of the fibers (Brown and Lucy, 1993). C. bFGF

Studies in mdx mice have shown that (i) in vitro, replication of myoblasts and, to a lesser extent, of fibroblasts is stimulated by smaller amounts of bFGF than in control cells (DiMario and Strohman, 1988) and (ii) the ECM of mdx muscle is richer in bFGF (DiMario et al., 1989; Anderson et al., 1991) and in acidic F G F (Oliver et al., 1992a) than that of control muscle, principally in the regeneration zones. bFGF may participate in the degenerative and regenerative responses of the muscle. In the control muscle, bFGF is localized at the periphery and in the nuclei of the muscle fibers, in the satellite cells, and, to a lesser degree, in the cytoplasm. In mdx muscle, intact fibers are labeled in the cytoplasm and nucleus. Small regenerating fibers accumulate more bFGF than do larger adjacent fibers. The smaller the cells, the more intense the staining. Mature fibers no longer express bFGF. The decrease in the immunoreactivity of bFGF is therefore a function of the size of the regenerating cells. The intensely labeled cells are more numerous at 5 weeks than at 10 weeks. Elsewhere, injection of bFGF promotes in vivo muscle regeneration in mdx muscle by enhanced replication of muscle satellite cells (Lefaucheur and SCbille, 1995). bFGF may also be involved in the physiology of the different striated muscles: The slow muscles contain more bFGF than do the fast muscles, both in mdx and in normal muscles (Anderson et al., 1991, 1993; Matsuda et al., 1992). A recent study indicates that mdx muscle cells have elevated levels of HSPG receptors for bFGF (Crisona et al., 1998).


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D. Ionic Flux 1. Calcium

Studies of calcium content in mdx muscle fibers have yielded contradictory results. Some authors have reported that free intracellular calcium levels increase at rest and during stimulation, resulting in an increase in degradation of muscle proteins (Turner et al., 1988; MacLenna et al., 1991; Kamper and Rodemann, 1992; Hopf etal., 1996). Likewise, in vitro this concentration is twice as high in the mdx myotubes at rest as in normal myotubes (Bakker et al., 1993). The intracellular calcium concentration is unchanged in mdx smooth muscle at rest, indicating that the absence of dystrophin does not always result in perturbed calcium metabolism (Boland et al., 1993). Several hypotheses have been put forward to explain the origin of this rise in intracellular calcium in mdx muscle: In young mdx mice, mechanosensitive ion channels, i.e., open at rest and inactivated by stretching of the membrane, are highly active (Franc0 and Lansman, 1990). This activity diminishes during development but is compensated for because the density of channels remains constant in mdx fibers but decreases in normal fibers. An early stage in the dystrophic process could be an alteration in the mechanisms governing expression of functional channels (Haws and Lansman, 1991). Fong et al. (1990) observed calcium leakage at rest. In controls, channels are little active and calcium is strictly regulated, whereas in the mdx mouse channels are active, leading to poor calcium regulation. Carlson and Officer (1996) attribute part of this calcium leakage activity to unusual physical interactions between AChR and cytoskeleton in mdx mice. Proteolysis raises membrane permeability to calcium at rest, creating positive feedback which results in an additional entry of calcium and muscle fiber necrosis (Turner et al., 1993). The sarcoplasmic reticulum pump is functionally altered in dystrophic mdx muscle (Takagi et al., 1992; Kargacin and Kargacin, 1996). Contradictory observations have been reported regarding parvalbumin, a protein which links calcium and relaxing factor in fast muscles. Sano et al. (1990) have shown that the decrease in the parvalbumin content of mdx muscle may contribute to a rise in intracellular free calcium and to activation of calcium-dependent proteolysis. Gillis (1996), on the other hand, reports that levels of parvalbumin and its mRNA (Gailly et al., 1993a) are higher in fast muscles of mdx mice than in normal muscles. A deficit in the level of calmitine has been reported in young mdx mice (Lucas-Hkron et al., 1990). The calmitine level remained low in 3-,

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5, and 6-week-old mdx mice and was similar to controls in 16-weekold mdx mice. However, calmitine deficiency does not affect the main calcium-binding structures involved in the regulation of mitochondria1 calcium (Lucas-HCron et al., 1994). Reports of an increase in intracellular calcium in mdx muscle are now contested. Several authors (Head, 1993; Pressmar et al., 1994; Gillis, 1996) consider instead that the resting potential of the membrane, the resting calcium concentration, and the transient calcium currents are identical in mdx and normal muscles. Leijendekker et al. (1996) report that calcium concentration is similar in mdx and normal myotubes, but increased permeability to calcium is noted for mdx myotubes under specific stress conditions. Gailly et al. (1993b) report that the concentration of cytosolic free calcium is comparable in mdx and normal muscles, although the total calcium increases. Most studies have examined muscle from mice of only one age. Reeve et al. (1997) demonstrated large changes in muscle calcium content during the postnatal development of the mdx mouse, thereby providing one possible explanation for the contradictory data reported.

2. Sodium and Potassium There is no apparent difference between the properties of the sodium channels of mdx and control muscles (Mathes et al., 1991), but intracellular sodium concentrations are higher in mdx muscles. This may reflect a reduced flow via the Na/K-ATPase, which would lead to poor control of cell volume and cell death (Dunn et al., 1993, 1995). The absence of dystrophin in the muscle sarcolemma does not affect the main KATp(Allard and Rougier, 1997) and K' delayed rectifier (Hocherman and Bezanilla, 1996) channel properties.

E. Innervation Axonal transport is broadly normal (Yamashita et al., 1989), as are innervation and myelinization, but in degenerating zones nerve terminals fragment into subunits. Although there is no direct injury of motoneurons, the growth-associated protein B50/GAP-43, which is involved in axonal outgrowth and synaptic remodeling following neuronal injury, is increased in terminal nerve branches at motor endplates of mdx mice, particularly in the areas of degeneratinghegenerating myofibers (Verze et al., 1996). The frequency of the miniature endplate potentials, their quanta1 content, and their amplitude are normal in mdx muscle (Hollingworth et al., 1990; Lyons


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and Slater, 1991). In contrast, Nagel et al. (1990) report that the amplitude of the miniature endplate potentials decreases and the quanta1 content of the endplate potential increases. There is a decrease in the number of subneuronal folds, simplification of the postsynaptic membrane (Torres and Duchen, 1987), widening of the synaptic zone (Nagel et al., 1990), and redistribution of the postsynaptic molecules. The G4 form of AChE is deficient in certain mdx muscles, likely reflecting a secondary effect of the dystrophy (Oliver et al., 1992b). The postsynaptic membrane’s AChR population is normal, but two types of receptors are expressed in mdx muscle: the adult type, as in normal muscle, and the embryonic type, similar to that of denervated muscle, with changes in opening time and current amplitude (Koltgen and Franke, 1992). This expression of the embryonic AChR is not a characteristic of dystrophy but a consequence of muscle regeneration (Koltgen and Franke, 1994). Salpeter’s group reports that AChR in innervated muscles of mdx degrades as AChR after denervation in normal mice (t112-3-5 days) but at no time as embryonic receptors ( t I l 2-1 day) (Xu and Salpeter, 1997). Most nerve terminals are abnormally complex at the NMJs of the regenerated fibers (Kitaoka et al., 1997). The absence of dystrophin in the postsynaptic membrane therefore has little effect on the function of the neuromuscular junction, but the degeneration and regeneration of the fibers leads to remodeling of the pre- and postsynaptic components. Carbonic anhydrase activity, a marker of mouse proprioceptive neurons in dorsal root ganglia, is regulated by neuron-muscle interactions. In mdx mice, neuronal carbonic anhydrase expression stops when the period of muscular degeneration-regeneration begins, and this alteration persists during adulthood (Mayeux et al., 1996).

F. Cytoskeleton The distribution and relative abundance of vinculin, desmin, and nebulin are unchanged in mdx muscle. The absence of dystrophin therefore does not result in alterations in the structures linking the sarcolemma to the contractile apparatus (Massa et al., 1994). These results partially contradict those of Law et al. (1994), who showed that in 2-week-old mice vinculin and talin are expressed comparably in mdx and normal muscles, whereas their expression is increased approximately 200% in mdx mice aged 11 months. This increase is localized in the myotendinous junction. It is argued that the mdx mouse partly compensates for the absence of dystrophin by overexpression of molecules which also have a mechanical function. Vinculin binds to actin and to talin, which binds integrins, and this ensemble may provide the link between the cytoskeleton and the ECM.

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G. Adhesion Proteins The expression of N-CAM is comparable to that in normal muscle before the start of the degeneration and regeneration cycles, but then it increases. It seems to be related to the muscular regeneration process (Dubois et al., 1994).

H. Mastocytes In the mdx mouse, the number of mastocytes is three times the normal value by 4 weeks and then decreases beginning at week 9; however, as in normal muscles the mastocytes are localized around the major blood vessels supplying the muscle. The peak coincides with the period of massive necrodregeneration (Gorospe et al., 1994; Lefaucheur et al., 1996).

I. Metabolism Changes in the lipid composition of the skeletal muscles of mdx and control (C57BL10) mice have been studied by nuclear magnetic resonance (NMR), a powerful noninvasive technique for studying living material (Gillet et al., 1993). The COSY sequence has been used to assign larger molecules such as fatty acids (Gillet et al., 1989). All the muscle spectra of dystrophic and control mice showed scalar correlations of saturated and unsaturated fatty acids. However, an additional correlation corresponding to triunsaturated fatty acid (linolenic-like) was highly visible in the muscles of mdx mice aged between 2 and 4 weeks (Fig. 4). This signal decreased as the mice grew and was not detected once they were more than 2 months old. Because myogenesis and regeneration of muscle in vivo can be mimicked in vitro using cultures of myogenic cells, the metabolism of muscle cell cultures has been studied in order to determine the origin of this linolenic-like signal in comparison with the different steps of myogenesis. The state of fusion of C2 cells can be distinguished from that of replicating cells by the presence of high-resolution signals consistent with saturated and unsaturated lipids and especially with triunsaturated “linolenic-like” lipids. Myoblasts can fuse to form myotubes during muscle regeneration, so it is possible that the specific triunsaturated linolenic-like signal in the mdx mouse muscle comes from the fusing cells and that it is characteristic of the regeneration of muscle fibers (SCbriC et al., 1988). Brain metabolism is abnormal in mdx mice (Tracey et al., 1996a): an increase in inorganic phosphate/phosphocreatine and pH, a reduction in


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FIG. 4 hi vivo 'H COSY spectrum of the hindleg of 3-week-old mdx mice. Changes in the lipid composition of mdx and control skeletal muscles are studied by nuclear magnetic resonance (NMR). The spectrum shows characteristic correlations of saturated and unsaturated fatty acid chains. In contrast to control mice muscle, the mdx muscle shows an additional correlation corresponding to triunsaturated (linolenic-like) fatty acids: K signal. This specific signal is present in the muscles of mdx mice aged between 2 and 4 weeks, but decreases once they are over 2 months old. This K signal is also detected in the spectra of fusing C2 cells. It is possible that the specific signal in the mdx mouse muscle comes from the fusing cells and that it is characteristic of muscle fiber regeneration. If this hypothesis is confirmed, 2-D 'H NMR could be used noninvasively to follow the regeneration process (reprinted from Neuromusculur Disorders 3, B. Gillet et al., In vivo 2D 1H NMR of rndx mouse muscle and myoblast cells during fusion: Evidence for a characteristic signal of long chain fatty acids, 433-438, copyright 1993, with permission from Elsevier Science).

total creatine, and an increased extracellular and decreased intracellular volume. The mdx mutation results in a decrease in energy metabolism: Metabolic rate, food intake, and physical activity all decrease in mdx mice between

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4 and 6 weeks. This difference compared with control animals is no longer apparent in 1-year-old mice (Dupont-Versteegden et al., 1994). In vivo, protein synthesis and degradation are higher in mdx mice than in controls at all ages. This may be the specific consequence of the absence of dystrophin since in the liver, in which there is no dystrophin, protein turnover is identical in mdx and control mice (MacLennan and Edwards, 1990). The high rate of proteolysis may be due at least in part to the accumulation of damaged proteins since abnormal proteins tend to be quickly degraded. Elimination and replacement of damaged proteins may constitute an important feature of the strategy by which the mdx muscle responds to the absence of dystrophin (MacLennan et al., 1991). In mdx muscle, levels of the glucose transporter GLUT4 are raised by 55%. In the diaphragm, the levels are identical to control levels at 5 or 6 weeks and decrease by 40% at 6 or 7 months. The expression of the major glucose transporter is related to the capacity of the muscle to regenerate rather than to the absence of dystrophin (Olichon-Berthe et al., 1993).

V. Therapeutic Projects Current research on the treatment of DMD is developed and tested using the mdx mouse. Three types of treatment are envisaged: pharmacologic, myoblast transplantation, and gene therapy. Most pharmacologic studies have focused on the effects of a glucocorticoid, prednisone. In vitro, prednisone stimulates myogenesis in cultures of mdx and normal muscle: the number of myotubes and levels of AChR, dystrophin, and D R P increase (Metzinger et al., 1993; Passaquin et af., 1993). In contrast, when administered to 15- to 45-day-old mdx mice, prednisone has no effect on the course of necrosis and regeneration (Weller et af., 1991), whereas another glucocorticoid, deflazacort, has a beneficial effect on both (Anderson et al., 1996). Anabolic steroid treatment increases myofiber damage in the mdx mouse (Krahn and Anderson, 1994). NMR is a useful noninvasive method to follow the effects of treatments on skeletal muscle metabolism (McIntosh et al., 1998). Glucocorticoid therapy slows the progression of DMD (Hardiman et al., 1992). Part of the beneficial effect of prednisone in DMD patients could be attributed to a reduction in Ca2+ influx and in the size of Ca2+ pools in dystrophic muscle fibers (Metzinger et al., 1995). However, the duration of this beneficial effect and the risk of long-term use remain to be determined. In 1989, two teams showed that when myoblasts from healthy mouse muscle were injected into muscles of mdx mice, the new cells fused with the mdx muscle fibers and produced dystrophin (Karpati etaf.,1989; Partridge et


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al., 1989). Myoblast transplantation leads to fiber formation only when immunocompetent but fully histocompatible donors and recipients are used (Vilquin et al., 1995). The use of a Y-chromosome-specific probe to track the fate of donor male myoblasts injected into dystrophic muscle of female mdx mice revealed rapid and massive death of the donor myoblasts soon after myoblast injection. There is limited movement of the injected donor myoblasts and fusion into host myofiber is rare (Fan et al., 1996b). Nevertheless, a high percentage of muscle fibers of donor origin can be obtained when myoblasts grown with bFGF are injected (Kinoshita et al., 1995) or when a short-term immunosuppressive treatment of mice with FK 506 is used (Asselin et al., 1995). Acute myoblast death can be prevented by control of the inflammatory reaction (Guerette et al., 1997). The use of sliced muscle grafts as a potential alternative strategy for myoblast transfer therapy proposed by Fan et al. (1996a) seems hypothetical because of the lack of myoblast migration between transplanted and host muscles (Moens et al., 1996). The application of this technique to DMD muscle initially yielded some dystrophin-positive fibers, but it was subsequently found that the functional benefit was extremely limited or short-lived (Law et al., 1990; Gussoni et al., 1992: Huard et al., 1992c; Karpati et al., 1993b; Mendell et al., 1995; Miller et al., 1997). Gibson et al. (1995) proposed an alternative approach to myoblast transfer therapy and reported the presence of dystrophin-positive fibers after implanting cloned dermal fibroblasts from normal mice into mdx muscle. The conversion of dermal fibroblasts (not muscle fibroblasts-the two are of different embryological origin) to a myogenic lineage is induced by a soluble factor derived from myoblasts (Wise et al., 1996). It has been demonstrated that the regional expression of recombinant dystrophin in dystrophic muscle leads to regional restoration of normal muscle morphology (mice: Cox etal., 1993; Dunckley etal., 1993:Matsumura et al., 1993; Ragon et al., 1993: Inui et al., 1996; Fassati et al., 1997a; dog: Howell et al., 1998). Numerous groups, in particular that of Karpati, are developing vectors, viral or plasmid DNA, which can be injected intravenously, intramuscularly, or systemically. Mdx mouse muscle treated by injections in the neonatal period expressed more dystrophin than muscle treated during adulthood or old age (Acsadi et al., 1996). In the case of herpes simplex virus-mediated gene delivery, a possible explanation is that basal lamina would be a physical barrier to infection (Huard et al., 1996). By 2 months postinjection, there is a substantial reduction in the number of dystrophin-positive fibers. This effect appears to be due in part to the activity of CD8+ cytotoxic lymphocytes directed against the transduced cells, leading to eventual elimination (Petrof et al., 1996). This hypothesis is substantiated by two observations: (i) In immunodeficient (SCID) mice, lacking both humoral and cellular immune competence, expression of

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transfected dystrophin is maintained over a longer period (Acsadi et al., 1996) and (ii) optimization of immunosuppression permits a sustained highlevel expression of dystrophin after gene transfer of dystrophin (Lochmuller et al., 1996; Chen et al., 1997; Zhao et al., 1997; Yang et al., 1998). The dystrophin minigene seems more effective than the full-length dystrophin gene (cf. Acsadi et al., 1996; Deconinck et al., 1996; Yanagihara et al., 1996). In fact, it has been shown in transgenic mdx mice expressing dystrophin with deletions that the cysteine-rich domain of dystrophin is critical for functional activity, presumably by mediating a direct interaction with pdystroglycan, and that the COOH terminus is not required for this assembly (Rafael et al., 1996). Contractile properties of diaphragm muscle segments from old transgenic-mdx mice do not differ from those of young transgenicmdx mice or control mice (Lynch et al., 1997). Minidystrophin gene transfer in mdx diaphragm leads to rapid and significant functional improvements (Decrouy et al., 1997). This line of research is expanding considerably. Important questions regarding the safety of the vector used, the difficulties of access to certain muscles, the number of transfected muscle cells, and the reproducibility of the results remain to be solved, however, before this demonstration finds a simple application in clinical therapy. The possibility of implanting a large numbers of genetically modified primary fibroblasts massively converted to myogenesis by adenoviral delivery of MyoD ex vivo has been tested in regenerating muscle of immunodeficient mice (Lattanzi et al., 1998). Three other ways to circumvent the numerous obstacles to gene therapy have been proposed: (i) the use of engrafted macrophages as potential shuttles for delivering a therapeutic agent (Parrish et al., 1996), (ii) the transplantation of retroviral producer cells toward the long-term goal of gene therapy (Fassati et al., 1997b), and (iii) the antisense strategy to transform DMD in Becker dystrophy phenotype by inducing exon skipping (Matsuo, 1996). Mdx mice are only mildly dystrophic, and utrophin-deficient mice show only subtle neuromuscular defects (Deconinck et al., 1997a; Grady et al., 1997a). However, utrophin-dystrophin-deficient mice present skeletal and cardiac myopathy and are considered a better model for DMD that mdx mice (Deconinck et al., 1997b; Grady et al., 1997b). The sparing of extraocular muscle in mdx mice is lost in mice lacking utrophin and dystrophin (Porter et al., 1998). Another model is proposed: the mutant generated by targeted disruption of exon 52 to disrupt the expression of the four other shorter isoforms that are also expressed from the dystrophin gene (Araki et al., 1997). In this mutant, muscle degeneration is similar to that observed in DMD. An alternative approach is to compensate for dystrophin loss by utrophin. There is evidence that utrophin may be capable of performing the same cellular functions as dystrophin (Tinsley and Davies, 1993; Blake et al.,


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1996b; Campbell and Crosbie, 1996). Expression of utrophin, with localization at the sarcolemma, is increased in DMD and Becker dystrophy (Karpati et al., 1993a; Mizuno et al., 1993) and mdx mice (see Section 111, C). Nevertheless, Vainzof et al. (1995) have shown an absence of correlation between utrophin localization and quantity and the clinical severity in DMD and Becker dystrophy. By transgenic expression of high levels of utrophin in mdx mice, Tinsley et al. (1996) demonstrated that utrophin could functionally replace dystrophin: Overexpression of utrophin leads to the restoration of all the components of DAGs, and mechanical performance of muscle is improved (N. Deconinck et al., 1997). Overexpression of utrophin even rescues the deterioration of the diaphragm, the most severely affected mdx muscle. The longevity of the truncated transgene expression has to be determined. The expression of a truncated utrophin transgene in muscle of utrophin-dystrophin-deficient mice prevents death and development of any clinical phenotype (Rafael et al., 1998). Identification of molecules or drugs that could upregulate utrophin is a very important goal for therapy. Today, only muscle and neural isoforms of agrin have been found to increase utrophin expression (Gramolini et al., 1998). The main advantages of such putative agents are (i) that there would be no need for gene replacement, because patients already have a functional utrophin gene and (ii) minimization of immunological responses. These different strategies should not be viewed as exclusive but rather can be clinically complementary.

VI. Concluding Remarks The mdx mouse is the best studied and therefore best understood animal model of Duchenne myopathy. Certain results are contradictory and further work is needed. There are various characteristics in common between these two myopathies: the absence of dystrophin, a reduction in the complex of associated glycoproteins, and the presence of utrophin along the muscular membrane and not restricted to the NMJ as in healthy muscle. There is, though, a very important difference which remains partly unexplained: In the mdx muscle intense regeneration compensates for the degeneration of muscle fibers. In DMD muscle, on the other hand, the regeneration process is weakened and muscle fibers are progressively replaced by fatty and connective tissues. Our results underpin the hypothesis that the increase in the components of basal lamina in vivo and the cessation of fibroblast proliferation observed in vitro may at least in part favor the marked muscular regeneration noted in mdx muscle. Indeed, we have found that fibroblasts taken from human DMD and control muscle have similar in vitro prolifera-

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tive capacities (Morin et al., 1995), and immunocytochemical visualization with polyclonal antibodies shows that the rate of expression of two components of basal lamina (laminin and HSPG) is lower in DMD than in control human muscle (Morin et al., 1993). The observed differences in these various parameters between mdx and DMD muscles could partly explain the regenerative capabilities of these two muscles: The myoblast/fibroblast balance seems t o favor myoblasts in mdx muscle. In DMD muscle the poor capacity of regeneration of the muscle seems to be due to a poor proliferative capacity of the satellite cells: A preparation of pure muscle satellite-cell populations has shown that the potential to generate myogenic cells for myofiber growth or regeneration is severely lacking even in the youngest DMD patients (Webster and Blau, 1990). All the treatments envisaged are designed to palliate this absence of regeneration, by promoting myogenesis or by limiting degeneration through pharmacologic treatments, by compensating for the weakness of the satellite cells by injection of myoblasts, or by compensating for the absence of dystrophin directly by complementing the genome through the use of vectors or indirectly by attempting to overexpress utrophin. In parallel with attempts to alter the genetic material of the DMD muscle through gene therapy, it appears necessary to continue the approaches employing cell and pharmacologic therapies.

Acknowledgments We thank the Association Franqaise contre les Myopathies (AFM), which has supported this work through grants. Sophie Morin was a recipient of a scholarship from AFM.

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by combined microscopic display of adenoviral DNA and dystrophin. H u m Gene Ther. 13, 1565-1573. Zubrzycka-Gaarn, E. E., Bulman, D. E., Karpati, G., Burghes, A. H. M., Belfall, B., Klamut, H. J.. Talbot, J., Hodges, R. S., Ray, P. N., and Worton, R. G. (1988). The Duchenne muscular dystrophy gene product is localized in sarcolemma of human skeletal muscle. Nature (London) 33, 466-469.

Regulation of Phosphate Transport and Homeostasis in Plant Cells Tetsuro Mimura Biological Laboratory, Hitotsubashi University, Naka 2-1, Kunitachi, Tokyo 186-8601, Japan

The inorganic phosphate (Pi) status of plants was reviewed based on the current knowledge of membrane transport systems and ion homeostasis. The Pi content and Pi distribution in plant cells were first considered in relation to experimental procedures used in their measurement. In order to understand the mechanisms which contribute to the Pi status, transport systems for Pi across the plasma membrane and the tonoplast were examined in detail. Recent progress in molecular biological approaches is discussed, especially for Pi transport across the plasma membrane. The molecular basis for Pi efflux across the plasma membrane and for Pi movements across the tonoplast still remains to be resolved. The involvement of Pi transport in Pi homeostasis is discussed from both the cellular and the whole plant perspectives. KEY WORDS: Apoplast, Inorganic phosphate (Pi), Membrane transport, Pi homeostasis, Plasma membrane, Plant cell, Tonoplast, Vacuole 0 1999 Academic Press.

1. Introduction Phosphorus is one of the most essential elements for biological organisms. There are many compounds in cells which contain phosphorus. The genetic apparatus is based on phosphate groups in the polynucleotides of DNA and RNA. Phospholipids are fundamental building blocks for cellular membranes, whereas energy transduction revolves around phosphate in ATP and in numerous other metabolic compounds. Recently, it has been established that protein phosphorylation-dephosphorylation by phosphate transfer is a major reaction in the regulation of various cellular functions (Ranjeva and Boudet, 1987). lnrernnriunal Rrvrrw of Cyrologs, Vol. 191

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Copyright ri3 1999 hy Academic Press. All rights 01 reproduction in any form rrherved.

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Phosphorus was one of the first elements to be recognized, being isolated from human urine in 1669. Isolation of elements from organic material is quite rare and phosphorus may therefore be considered an element symbolic of living organisms. Of course, carbon, nitrogen, oxygen, and hydrogen may be more important elements than phosphorus, but the position of phosphorus as an essential element in living organisms is unique. First, the total phosphorus in the earth which is utilizable by living organisms is limited differently from that of other major elements. All phosphorus in organisms originates in phosphate molecules which are taken up from the soil by plants. Free phosphate concentrations in the soil, however, are very low-usually z

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day, but generally it takes 2 or 3 days to detect a distinct activation. Kinetic analysis of the activation of Pi uptake showed that K , and/or V,,, changed during Pi starvation. In experiments using Neurospora, Burns and Beever (1977a,b) showed that the K,,, of the low-affinity transport system decreased from 0.9 to 0.3 mM when the Pi levels in the culture medium changed from 0.05 to 10 mM. On the other hand, the K,, of the high-affinity transport system (around 3 p M ) did not change. In higher plants, there are two phases of changes. After Pi starvation, V,,,, first increases and then K,, decreases. Sometimes we can see only one of these responses. If we assume that the K,,, is a characteristic feature of one protein, then a change in K , for Pi implies that the molecular species has also changed. On the other hand, changes in V,,, may occur either by changes in the nature of the molecular species or by changes in the number of the same species. Although it was known that Pi deficiency increased Pi uptake, it remained uncertain whether in higher plants there were really different transporters with different affinities for Pi. Molecular biological research on Pi transporters finally concluded that Pi transporters belonged to a multiple gene family. Muchhal et al. (1996) and Leggewie et al. (1997) have shown that there are more than one gene that complements the yeast PH084 mutant. Mitsukawa et al. (1997b) isolated four different genes from Arabidopsis. Among these genes, some were expressed only in Pi-starved conditions (Table IV). Thus, there is no doubt that Pi deficiency can induce different kinds of transporters. Pi transport activities of these genes were analyzed by complementation of the yeast Pi transport mutant. K,, values found in these experiments do not always agree with affinities for Pi obtained from in vivo flux measurements in higher plants (Leggewie et al., 1997). Mitsukawa et al. (1997b) have also succeeded in introducing a Pi transporter gene from Arabidopsis into tobacco suspension-cultured cells. These transgenic tobacco cells showed higher Pi uptake activity than before, indicating that at least some of the genes for Pi transporters must have a high affinity for Pi. There is evidence from split-root experiments that Pi uptake activity is determined not by localized Pi status but at the tissue or whole plant level. Drew and Saker (1984) divided barley roots into two parts, one part remaining Pi deficient and the other part supplied with Pi. Pi uptake of the Pi-deficient part was found to be affected by the Pi-supplied part. This was confirmed at the molecular level by Liu et al. (1998), who showed that expression of the Pi transporter in the Pi-deficient part changed when Pi was supplied to the other part. Drew and Saker suggested that the internal Pi level of cells in the Pi-deficient part might change by Pi movement from the Pi-supplied part. However, there is no evidence that the cytoplasmic Pi level of the Pi-deficient part really changes when the other part is supplied with Pi. We need t o determine how the Pi level of the apoplast in the Pideficient part is influenced by Pi supply through the xylem and phloem. It

PHOSPHATE TRANSPORT AND HOMEOSTASIS IN PLANT CELLS

177

would be interesting to determine whether Pi transport activity can respond to internal signals even when there is no Pi in the extracellular medium. ii. Growth and Differentiation There have been many studies on the relationship between plant growth and Pi nutrition. Lefebvre and Glass (1982) traced growth-dependent changes in Pi influx in barley. After germination, Pi influx based on the root fresh weight gradually decreased in the presence of an adequate Pi. Under Pi starvation, it dramatically increased after 12 days and then returned to the original activity after 16 days. It is likely that Pi uptake in the root is regulated by the demand for Pi for shoot growth. Pi uptake of growing plants has been measured by various methods (Breeze et af., 1984, 1985; Ericsson and Ingestad, 198S), with the general finding that growth rate and Pi uptake rate are strongly correlated. Clarkson and Saker (1988) concluded that under low levels of external Pi (the normal condition in nature), growth and Pi uptake rates were constant between a wide range of external Pi concentrations. Plants seem to control their Pi uptake activity to keep the growth rate constant, independently of the extracellular Pi concentration. Experiments in which Pi uptake is studied using suspension-cultured cells under a continuous flow of culture medium may help to clarify the relationship between Pi uptake rate and cell division (cell cycle) at the cellular level. Molecular analyses of Pi transporter genes have demonstrated that the expression of each gene is dependent on not only the Pi status but also the type of plant (Table IV). Liu et al. (1998) showed that the expression of Pi transporters was differently regulated in the genes (LePTI and LePT2) in tomato tissues. Both were expressed in the root epidermis. Under Pi starvation, LePTI transcripts were also detected in the root central cylinder and in the leaf palisade parenchyma and phloem cells, whereas LePT2 was not detected in the leaf at all.

c. Pho Regulon and Two-Component Systems A high-affinity Pi transport system of prokaryote cells has been shown to be composed of outer membrane protein, a Pi-binding protein in the periplasm, and a phosphate transporter belonging to the ABC transporter group. Although I have not dealt with Pi transport in procaryotes in this review (see Nakata et al., 1987; Rao and Torriani, 1990), I draw attention to its regulation system. The high-affinity Pi transport system is under the control of the phosphate regulon. Induction of the Pho regulon is mediated by a two-component system. Two-component signaling systems have been identified as important receptors of environmental signals, e.g., osmolarity, chemotaxis, and nitrogen source (Wurgler-Murphy and Saito, 1997; Chang and Stewart, 1998). The level of extracellular phosphate is also an environmental signal. Pho R and Pho B proteins function to transmit information about the Pi level to the Pho regulon. The Pho R protein is known to be a sensor protein of

178

TETSURO MIMURA

the cell membrane and is believed to detect the extracellular Pi concentration and not the cytoplasmic Pi level. Blue-green algae also respond to the extracellular Pi conditions via a similar two-component system (Aiba et al., 1993). In higher plants, two-component systems have recently been reported to work as ethylene receptors and cytokinin receptors (Wurgler-Murphy and Saito, 1997; Chang and Stewart, 1998). Currently, there is no experimental evidence that they may operate for the detection of nutritional status in higher plant cells, as they do in procaryotes.

5. Measurement of Pi Efflux and Xylem Loading Pi efflux from cells through the plasma membrane is an important aspect of Pi transport. When Pi moves from root to shoot, Pi must be released from root cells into the xylem (xylem loading). Movement into the apoplast will normally require the expenditure of metabolic energy for export across the plasma membrane. There are few reports on Pi efflux compared to those on the influx measurements. In order to measure the efflux accurately, it is necessary to use tissues loaded with radioisotopes of phosphorus. The most popular method is the time-dependent efflux analysis from labeled cells. The efflux of "Pi usually shows three different time-constant curves, as is observed for many other ions. The fastest efflux is interpreted as release from cell wall, the second fastest as efflux from the cytoplasm across the plasma membrane, and the slow phase comes from the vacuole across the tonoplast (Lefebvre and Clarkson, 1984; Woodrow et al., 1984). However, smaller phases with time constants for exchange similar to those of the larger pools will not be detected by this method. McPharlin and Bieleski (1989) reported the following characteristics of Pi efflux in Spirodela or Lemna. In Pi-adequate plants, Pi efflux was about 10% of the influx at moderate Pi concentrations. In Pi-deficient plants, it decreased to approximately 1%. A t very low external concentrations, efflux can be similar to influx (i.e., no net uptake). This is known as the equilibrium concentration point. In Pi-deficient plants, this was < 0.1 p M . Pi-adequate plants showed three or four times higher values than did Pi-starved ones. This was confirmed by Bieleski and Lauchli (1992). Low temperature and metabolic inhibitors such as CCCP, NaN3, and KCN increased Pi efflux. Such experimental treatments usually cause membrane depolarization. Membrane depolarization, however, is favorable for Pi influx, not Pi efflux. Thus, we do not know the reason why metabolic inhibition increases Pi efflux. Table VI summarizes the characteristics of Pi efflux. McPharlin and Bieleski (1989) observed that both Pi efflux and Pi influx increased as the extracellular Pi concentration increased. A rapid increase

-I

*

TABLE VI Comparison of Pi Influx and Efflux

33

z

U

Material

Pi concentration of external medium ( p M )

Influx Original value 0.11 pmolldcm’

Hydrodictyon africanum

n

Efflux Estimated (nmol/min/g FW) 0.31“

Original value 0.02 pmol/s/crn’

-I

D

Estimated (nmol/minlg FW) 0.06‘

z 0

Reference Raven (1974)

I 0

5

g

--I

!%

Hordeum vulgare Zea maiz

0.2 2.0

836 nrnol/h/g FW

13.9

206 nmol/h/g FW

3.43

Lefebvre and Clarkson (1984)

0.27 nmol/min/g FW 4.91 nmol/min/g FW

0.27 4.91

0.24 nmol/min/g FW 0.62 nmollminlg FW

0.24 0.62

Elliot ef al. (1984) McPharlin and Bieleski (1989)

Lemma major

1 10

21 nmol/h/g FW 109 nmollhlg FW

0.35 1.82

15 nmollhlg FW 25 nmollhlg F W

0.25 0.42

Spirodela oligorrhiza

1 10

22 nmollhlg FW 130 nmol/h/g FW

0.37 2.17

17 nmol/h/g FW 30 nmol/h/g FW

0.28 0.5

Azolla mexicana

1 10

22 nmol/h/g FW 101 nmollh/g FW

0.37 1.68

11.7 nmol/h/g FW 13.8 nmol/hr/g FW

0.20 0.23

Flux was estimated based on 0.005 mz/g FW.

0

z

9

b z -I

0

Bieleski and Lauchli (1992)

rn r

cn

180

TETSURO MIMURA

in Pi efflux from Chara cells was also found when Pi was added to the extracellular medium during the measurement of efflux (Mimura et al., 1998). Thus, cells appear to have an ability to sense changes in the extracellular Pi supply and to respond through the Pi efflux system. It is well known that symbiotic mycorrhizal fungi play an important role in Pi uptake from soil in some plants (Schachtman el al., 1998). In phosphate transfer from mycorrhizae to plant root cells, Pi efflux from fungal cells to the plant host is an important step. Unfortunately, the factors that determine the rate of efflux are not understood. Currently there is no information on the molecular mechanisms which mediate Pi efflux across the plasma membrane. Since the electrochemical potential gradient of Pi is usually outward, it is feasible for Pi to efflux through ion channels. I have not encountered any reports of studies in which the Pi permeability of anion channels in plant plasma membrane has been examined. Some mutants of Arabidopsis relate to Pi efflux. The Phol mutant may be a Pi efflux mutant (Poirier et al., 1991; see Table IV) since it was not able to accumulate Pi in the shoot, even though uptake by the root was normal. The most likely explanation for this observation is that there is a defect in the loading of Pi into the xylem in the root. The Phol gene may code the transporter contributing Pi efflux. 6 . Tonoplast Pi taken up into the cytoplasm across the plasma membrane is partly metabolized and most of the rest is transported into the vacuole (Bieleski and Laties, 1963) which occupies more than 80% of the cell volume. Pi accumulated in the vacuole is used to buffer the cytoplasmic Pi level against fluctuations caused by variable external supply. Thus, Pi transport across the tonoplast (the vacuolar membrane) becomes an important factor in Pi metabolism of plant cells.

1. Measurement of Pi Influx There are many methods for measuring Pi transport across the tonoplast. The in vivo measurement using "P-NMR can simultaneously measure both the cytoplasmic and the vacuolar Pi contents. Time-dependent changes in the vacuolar Pi content reflect the net flux of Pi. In sycamore suspensioncultured cells, when Pi was depleted from the culture medium, the vacuolar Pi level gradually decreased over 50 h until it coincided with the cytoplasmic Pi level. When Pi was added to the Pi-deficient cells, the Pi level of the cytoplasm increased within 2 h, and then the vacuolar Pi level gradually


181

increased (Rebeille et al., 1983). More detailed analyses were obtained in Cutharunthus suspension-cultured cells which were almost Pi deficient (Sakano et al., 1995). Pi added to the culture medium induced a twofold increase in the cytoplasmic level, after which it remained steady. Afterwards, the vacuolar Pi level dramatically increased, although the rate of increase was slower than that in the cytoplasm. Pi influx has also been estimated from direct measurement of Pi concentrations in vacuolar sap. In C h r u cells, it is easy to isolate the sap because of the large size of the internodal cells. Smith (1966) showed that labeled Pi added to the medium first appeared in the cytoplasm and then in the vacuole. The level of radioactivity in the cytoplasm increased only for the first few hours, but in the vacuole it continued to increase for more than 10 h. In barley mesophyll cells, the intact vacuoles of protoplasts, were isolated every 30 min after an addition of '2Pi and the radioactivity in the vacuoles was measured (Mimura etal., 1990). "Pi was detected in the vacuole within 30 min in Pi-deficient protoplasts but it took longer than 1 h in Pi-rich protoplasts (Fig. 7a). This suggests that Pi deficiency may induce new Pi transport activity

a

T

I

0

I

I

I

20 40 60 Time (min)

80

FIG. 7 Pi uptake in vacuoles of Barley mesophyll cells. (a) I n vivo Pi uptake into vacuoles. Mesophyll protoplasts were incubated with "Pi, and then the vacuoles were isolated and 33P in the vacuole was counted. (b) Pi uptake by the isolated vacuoles. After isolation of vacuoles, they were incubated with "Pi. 0 and A. Pi-adequate barley; 0 and A, Pi-deficient barley. In b. solid lines indicate that the isolated vacuoles were incubated with A T P (broken line, without ATP) (from Planla, Phosphate transport across biomembranes and cytosolic phosphate homeostasis in barley leaves, Mimura. T.. Dietz. K.-J., Kaiser, w., Schramm, M. J.. Kaiser, G . . and Heber. U., 180, 139-146. Fig. 6, 1990, 0 Springer-Verlag).

182

TETSURO MIMURA

not only in the plasma membrane but also in the tonoplast. This was confirmed by isolating vacuoles from both Pi-rich or Pi-deficient barley protoplasts and comparing Pi uptake rates. Pi uptake rates into the vacuole of Pideficient cells were higher than those of Pi-rich cells (Fig. 7b). However, the increase in Pi uptake under Pi deficiency was only observed in the presence of ATP. Without ATP, Pi influxes in both vacuoles were the same. Thus, Pi transport into the vacuole may depend on the cellular energy status, at least in the Pi-deficient cells. This experiment also indicated that Pi influx to the vacuoles of Pi-rich cells may be very slow. Measurement of transport activities across the tonoplast can also be performed using tonoplast membrane vesicles. If a transport process is energy dependent, the membrane vesicles can be energized by addition of ATP or PPi to drive Hi pumps, p H jumps of incubation media to create pH gradients, or generation of membrane potential differences with K t and ionophore. Kaestner and Sze (1987) measured potential-dependent anion transport using tonoplast vesicles from oat. They showed that the permeability of Pi was very low compared with that of other anions. However, because Pi may also have been acting as an inhibitor of both H+ATPase and PPase (Gonzalez and Medina, 1988; Takeshige et al., 1992), their results were ambiguous. 2. Mechanism and Regulation of Transport Few reports deal with the mechanism of Pi transport across the tonoplast. Mimura et al. (1990) measured the concentration dependence of Pi influx into vacuoles isolated from Pi-rich barley mesophyll cells. Pi influx did not saturate until 20 mM Pi in the extravacuolar medium, which is at the upper end of measured cytoplasmic Pi concentrations. These results help establish the electrochemical conditions under which Pi transport to the vacuole operates and allow educated guesses regarding the likely molecular mechanism. It is difficult to estimate accurately the electrochemical potential gradient of Pi across the tonoplast because it is unknown which molecular species of Pi is transported and the vacuolar Pi level varies greatly depending on the Pi nutrient status. At pH 7.5 in the cytoplasm, most of the Pi is in the divalent HP04’- form, but in the vacuole, where the pH is approximately 5, most of the Pi is present as the monovalent H2P04-. The electrical potential difference across the tonoplast is usually slightly positive with respect to the cytoplasm. If we consider only HPO?-, both concentration and electrical potential gradients are inward into the vacuole. HP04*- can therefore be passively transported via either carrier or through ion channels. On the other hand, if we consider H2P04-,the electrochemical potential gradient changes its direction dependent on the vacuolar content and it


183

may therefore be necessary to invoke an active transport mechanism such as Pi-ATPase, Pi-PPase, or H' antiport system under some conditions. In Pi-deficient vacuoles, the presence of extravacuolar ATP accelerated Pi influx (Mimura et al., 1990).Eligny et ul. (1 997), using "P-NMR, measured the effects of anoxia on cellular Pi-containing substances. In their measurements, when cells were exposed to low-oxygen conditions, the cellular ATP level decreased and Pi moved from the vacuole to the cytoplasm. On recovering to normoxia, Pi in the cytoplasm returned to the vacuole. They suggested that ATP played an important role in Pi uptake to, and keeping Pi in, the vacuole. A possible complication with these NMR experiments arises from the fact that if anoxia causes the pH of cytoplasmic organelles to fall to a similar pH to that of the vacuole, then the apparent vacuolar pool will seem to increase (see Section 11), thereby causing an underestimation of the efflux across the tonoplast. There are three possible roles of ATP. ATP may work as a direct energy source for Pi transport. ABC transporters such as the glutation-S-conjugate transporter are known to operate in the tonoplast (Martinoia et al., 1993; Lu et al., 1998). Pi uptake into the vacuole may be mediated by one of these transporters. Second, V-type H+-ATPases may be involved in the transport of Pi. Activation of the H'-ATPase produces a vacuole-inside positive potential. If Pi transport is driven by the electrical potential gradient, addition of ATP may increase Pi influx. In preliminary experiments, specific inhibitors of H'-ATPase, bafilomycin and folimycin, partly inhibited Pi uptake into the isolated vacuole. Very recently, we have confirmed that not only ATP but PPi is also effectual to Pi uptake of Pi-deficient vacuoles isolated from Cutharunthus roseus cell (Mimura et al., unpublished data). Lastly, ATP is known to be a transport regulator of the tonoplast. Dietz et al. (1990) showed that ATP activated amino acid transport into the vacuole without hydrolysis of ATP. Such a role for ATP in Pi transport is possible but there is no evidence to support such a mechanism. Klughammer et al. (1992) measured the ion channel activities of the tonoplast integrated into planer lipid bilayers. They detected anion-dependent currents with a range of anions including Pi, although the conductance of Pi was very low compared to that of some other anions. Two reports have recently dealt with Pi transport across the tonoplast in yeast (Booth and Guidotti, 1997; Kulakovskaya and Kulaev, 1997). Booth and Guidotti suggested the existence of a bidirectional Pi transporter in the tonoplast. This transporter had a relatively high affinity for Pi (K,,, approx 0.4 mM) and had a higher activity at low pH. Kulakovskaya and Kulaev described Pi uptake through a channel-like transporter, which was independent of the electrochemical proton gradient.

184

TETSURO MIMURA

There is currently no information on the molecular structure of the tonoplast Pi transporter(s) in plants. However, as shown in Table IV, there are many Pi transporter genes that have been identified in Arabidopsis, and some of them may well be tonoplast transporters. 3. Measurement of Pi Efflux from the Vacuole The most important function of the vacuole is to store excess Pi and buffer the cytoplasmic Pi level. Mobilization of vacuolar Pi under conditions of deficiency involves controlled efflux of Pi across the tonoplast. As with influx, efflux of Pi has also been measured by "P-NMR by following the changes in the vacuolar and the cytoplasmic Pi contents. Since the cytoplasmic Pi may be immediately metabolized, the changes in vacuolar Pi are a better indication of the efflux of Pi. When sycamore suspension-cultured cells were deprived of sucrose, most of the Pi accumulated in the vacuole. Addition of sucrose immediately changed the cytoplasmic Pi level but did not affect the vacuolar pool (Roby er al., 1987). This suggests that efflux of Pi from the vacuole in sitic may be very slow. The Pi efflux from isolated vacuoles has also been investigated (Martinoia et al., 1986). Incubation of isolated barley vacuoles in Pi-free medium did not result in a Pi release compared to other anions (C1- and NO,-), which is consistent with the view that Pi movement out of the vacuole is slow. Observations with NMR of pea leaves under anoxia, however, showed that Pi rapidly moved between the vacuole and the cytoplasm (Bligny et al., 1997). In yeast, the addition of Pi to the extravacuolar medium induced rapid efflux of Pi (Booth and Guidotti, 1997). The apparent conflict between these studies remains to be resolved. Figure 8 shows a schematic diagram of membrane transport of Pi in plant cells. Molecular analyses of Pi transporters have concentrated only on Pi uptake across the plasma membrane. Studies on the other transporters and on the mechanisms of regulation of Pi transport are important subjects that should be studied in the near future.

V. Homeostasis and Detection of Pi Status in Plant Cells Homeostasis of Pi is an overall phenomenon resulting from the orchestration of many individual processes, including membrane transport, binding to the membrane, sequestration as Ca or Mg precipitate, and metabolic conversion between inorganic and organic phosphates. Homeostasis means the control of physiological status and the maintenance of a certain steady state. Feedback regulation is used to counter any perturbation. For this, the system has to be able to constantly monitor the prevailing conditions.


185

FIG. 8 Schematic diagram of Pi transport systems in plant cells. Transporters drawn with broken lines are putative molecules.

In Pi homeostasis, this may require monitoring of the Pi concentration in each compartment. Here, I focus only on the relationship between Pi homeostasis and Pi transport activities. Regarding metabolic processes, there are excellent reviews (Theodorou and Plaxton, 1993: Duff era/., 1994). A. Cytoplasm and Vacuole Bieleski and Laties (1963) made it clear with radioisotopes that there are two different pools of Pi in plant cells; one in which the Pi is actively

186

TETSURO MIMURA

metabolized and one in which Pi metabolism is static. It is evident that the former is the cytoplasm and the latter is the vacuole. In all experiments in which Pi influxes to the cytoplasm and to the vacuole have been measured separately, the transported Pi accumulated first in the cytoplasm and then in the vacuole. The cytoplasmic Pi level eventually reached a steady level, whereas the vacuolar Pi level continued to increase. These studies suggested that the cytoplasmic Pi level might be regulated at a certain value but that the vacuolar Pi level might not. 'lP-NMR measurements showed that under Pi deficiency, the vacuole was acting as a Pi reservoir to maintain the cytoplasmic Pi pool constant (Rebeille et al., 1982, 1983; Lee and Ratcliffe, 1983; Lee et al., 1990). In 1990, Mimura el al. isolated protoplasts and vacuoles from barley leaves under Pi deficiency and measured changes in subcellular distribution. Mimura et al. (1992, 1996) have analyzed changes in Pi concentrations in subcellular compartments during the growth of barley. These studies showed that Pi concentrations in the cytoplasm were always constant and were independent of the Pi supply. On the other hand, the vacuolar Pi changed in order to keep the cytoplasmic Pi concentration constant. Although the exact cytoplasmic level of Pi is still unknown (see Section 111), it is believed to be from a few to approximately 10 mM. Maintenance of this level of Pi in the cytoplasm is essential to keep the Pirelated metabolism at a normal state, i.e., Pi homeostasis in the cytoplasm. Lee and Ratcliffe (1983, 1993) found that Pi homeostasis was achieved under moderate Pi deficiency but once the vacuolar Pi pool was completely exhausted, the cytoplasmic Pi concentration also began to decrease. Cytoplasmic Pi homeostasis is mainly achieved via a combination of membrane transport and metabolic conversion from organic phosphates to inorganic phosphates. When the cell increases in volume or divides into two daughter cells, the cytoplasmic Pi content will be diluted because a part of Pi is consumed for structural materials of new cells. To compensate, Pi is taken up from the external medium and from the vacuole. Under Pistarved conditions, recovery of the cytoplasmic Pi concentration can only be achieved by transport from the vacuole. The use of the vacuole as a reservoir for cytoplasmic Pi also applies in yeast (Shirahama et al., 1996). It is known that yeast cells accumulate large amounts of phosphate as polyphosphate which is stored in the vacuole. Cytoplasmic Pi homeostasis in yeast and other polyphosphate-accumulating fungi and algae is dependent on the synthesis and the degradation of polyphosphate and may therefore be a more complicated process than that in higher plant cells. Metabolism of Pi has a greater effect on the cytoplasmic Pi level than on the total cytoplasmic phosphorus content, which is the sum of inorganic and organic phosphates. In green cells, light activates photosynthesis and some of the Pi present in the dark is esterified into organic compounds in


187

chloroplasts. Takeshige et al. (1992) measured the cytoplasmic Pi level of Chara cells between light and dark conditions and found that 20 min after the beginning of the light period, the cytoplasmic Pi level decreased to almost 50-60% of that in the dark. In suspension-cultured cells of Acer, it was found that sucrose starvation induced a large accumulation of Pi in the vacuole and a small increase in the cytoplasmic Pi concentration (Rebeille et al., 1985) as a result of the degradation of organic phosphate. Thus, although cytoplasmic Pi levels measured over long periods may appear to be constant, there may nevertheless be shorter term fluctuations resulting from variations in metabolic activity. Intracellular depletion of Pi can lead to changes in many processes, for example, activation of Pi uptake, induction of phosphatase, and RNase. How do these processes detect changes in Pi levels in the different compartments in which they operate? Unfortunately, this question cannot be answered for plant cells. Currently, there is only knowledge of the bacterial two-component systems which detect osmotic or nutritional conditions (Wurgler-Murphy and Saito, 1997). The main problem in plant cells is that many of these changes occur in the absence of, or before, measurable changes in the Pi level of the cytoplasm. It is known however, that the vacuolar Pi concentration undergoes marked changes and this might therefore be a suitable level to monitor. Mimura et al. (1998) analyzed the activation of Pi transport under Pi deficiency using mature Chara cells which do not grow or divide (k, intracellular Pi stores are not diluted by growth or division). In these cells, changes in Pi transport resulting from Pi starvation were completely independent of the vacuolar Pi level. Thus, the assumption that the vacuolar Pi level may act as the signal for Pi status is excluded, at least in these cells. The second difficulty is that the cytoplasmic Pi level, especially that of photosynthetic cells, changes greatly between light and dark within a few minutes. If the cell responds to the cytoplasmic Pi level, deficiency-induced responses should also occur in the light (the stimulation of Pi uptake may be consistent with this). Another possibility is that the plant cell may detect the extracellular Pi concentration. In prokaryotes and yeast. the extracellular Pi is likely to be monitored by the cell. It has been proposed that the two-component system membrane receptors detect extracellular Pi in E. coli and Syrzecococcus (Torriani, 1990; Aiba et al., 1993). In yeast, some of the transporter proteins may be involved in the detection of extracellular Pi (Oshima, 1997) but further evidence is needed. Recently, in an ingenious experiment using tomato suspension-cultured cells, Kock etal. (1998) showed that the coexistence of extracellular Pi and Pisequestering substances such as mannose induced RNase, which was usually induced under Pi-deficient conditions. They suggested that the application

188

TETSURO MIMURA

of these substances sequestered intracellular Pi and transiently decreased the cytoplasmic Pi level before recovery as a result of release of Pi from the vacuole. This experiment provides support for the proposal that the cell detects the cytoplasmic Pi level and not the extracellular Pi level. Moreover, since the changes in Pi were expected to be transient (they did not actually measure changes in cytoplasmic Pi), it implies that responses can be rapid. In addition to detecting intracellular Pi or extracellular Pi, it is also possible that plant cells are able to detect Pi fluxes directly, but this suggestion remains entirely speculative. There is clearly a need to find a receptor, a process, or a level that can be used to predict a plant’s response to changes in Pi status.

B. Apoplast The “symplast” is the intracellular phase of a plant. Its counterpart, the “apoplast,” comprises those parts within the plant that are extracellular. The main components of the apoplast are the cell walls and extracellular spaces and the conductive tissue of the xylem. I previously discussed how intracellular phases of plant cells are subject to homeostasis. This is also true of animals, but in animal tissues homeostasis of the extracellular fluids is of a similar importance to the maintenance of a controlled intracellular phase. In plants, there is evidence for homeostasis of the apoplast (Canny, 1995), albeit limited by the constraints imposed by widely varying environmental conditions. Measurements of Pi levels in the xylem (Bieleski, 1973; Bieleski and Ferguson, 1983; Marschner, 1995) yield values much higher than those in the external medium. There must therefore be steps between the external medium and the xylem in which Pi becomes concentrated. Except in the small region near the root tip where the endodermis is immature, there is no direct connection between the external medium and the xylem. The increase in concentration of Pi in the xylem must therefore originate in processes involving the symplast, either uptake into the root cells or unloading into the xylem. The Arubidopsis Phol mutant, which appears to be defective in xylem loading (Poirier et uf., 1991),may be useful in this respect. . measured the growth-dependent changes in xylem Mimura et ~ l (1996) Pi levels. When plants were subjected to Pi deficiency, the Pi levels in the xylem decrease to approximately 1 m M compared to 5-7 mM in the nondeficient control plants. As the plant became more P deficient and the whole plant Pi concentration gradually declined, the Pi level in the xylem remained constant at approximately 1 mM. It seems likely that the xylem Pi concentration is a function of both the supply of Pi from the soil and the demand for Pi by the shoot.

189


There is also evidence that Pi levels in the cell walls are regulated. Mimura etal. (1992,1996) measured Pi in the apoplastic fluid collected by infiltration and centrifugation of barley leaves. They found that the concentration of Pi in the apoplast paralleled that of the whole leaf, which in turn was dependent on growth and Pi nutrition. However, under Pi deficiency, in which the whole leaf Pi concentration dropped to approximately 15% of the control, the apoplastic Pi level fell to only 30% of the control (Mimura et al., 1990, 1992, 1996). In experiments in which Pi was fed to detached leaves via the transpiration stream, the level of Pi in the apoplast was independent of the supply concentration up to approximately 10 mM (Fig. 9). When the 10 mM Pi solution was replaced by pure water, the apoplastic Pi concentration remained constant. These results suggest that the apoplastic Pi concentration can be kept quasi-constant and it seems likely that this occurs via exchange with vacuolar Pi, a process which requires the participation of both the tonoplast and the plasma membrane. C. Whole Plants Pi is one of the most mobile molecules in plants (Bieleski and Ferguson, 1983). Pi is known to be retranslocated from older to younger tissues, from

I

I

I

0

10

I

20

I

30

I

40

I

50

Pi concentration fed to the leaves (mM) FIG. 9 Changes in the apoplastic Pi Concentration of barley leaves as a function of the Pi concentrations fed through the transpiration stream (modified from Planfa, Phosphate transport across hiomembranes and cytosolic phosphate homeostasis in barley leaves, Mimura, T., Dietz. K.-J.. Kaiser, W.. Schramm, M. J., Kaiser. G., and Heber. U.. 180, 139-146. Fig. 2. 1990, 0 Springer-Verlag).

190

TETSURO MIMURA

Pi-supplied to Pi-demanding tissues, or back to the root from the shoot (Clarkson et al., 1978; Mimura et al., 1996; Jeschke et d., 1997). These are well-known phenomena which reflect an apparent homeostasis of Pi at the whole plant level (Bieleski and Ferguson, 1983). Biddulph et al. (1958) first showed retranslocation of Pi from old leaves to fresh leaves using radioautograms. Furthermore, the following studies provide more detail on Pi retranslocation (Greenway and Gunn, 1966; Smith er al., 1990). The split root experiments of Drew and Saker (1984) clearly showed that Pi taken up by Pi-supplied roots moved to Pi-deleted ones. Recently, Liu et al. (1998) reinvestigated Drew and Saker's work and found that Pi transporter genes were expressed not only in the Pi-deficient roots but also in the Pi-supplied roots in the divided root system. Pi depletion in one part of roots affected the other part of roots. They suggested that the internal signal (possibly the cytoplasmic Pi level) induced an expression of Pi transporter in both parts. Mimura et al. (1996) visualized and quantified the Pi movement between barley leaves with radioisotopes (Fig. 10). Using a highly sensitive imaging plate (Fuji Film, BAS2000), Mimura et al., analyzed the redistribution of Pi in the same individual plantlet (Mimura, 1995a; Mimura et al., 1996). After pulse labeling of the Pi-deficient barley plant, 32Pmoved sequentially from the first to the second and from the second to the third leaves (Fig. lo), confirming the results of Biddulph et al. (1958). Also using the imaging

FIG. 10 An example of autoradiograms obtained using an imaging plate. Barley grown in Pideficient conditions was pulse labeled with '*Pi for 30 min and then radioactivity was chased (from Mimura, 1995, with kind permission of Japanese Society of Plant Physiologists).

191


plate, Mimura et al. could quantitatively estimate amounts of 32Pin each part (Fig. 11). Interestingly, the Pi-adequate plant (Fig. l l a ) behaved like the Pi-deficient plant (Fig. l l b ) after the plant was exposed to Pi-depleted medium. Since the Pi-adequate plant was grown in the presence of Pi for 10 days, it should have enough Pi in tissues. The imaging plate technique should prove useful for investigating the time-dependent and space-dependent changes in 32Pdistribution in whole tissue. Retranslocation of Pi between tissues is believed to occur via phloem (Bieleski, 1973). However, little is known about the mechanism of Pi movement from cells to phloem and from phloem to cells. There are two pathways from cells to phloem, symplastic and apoplastic. For symplastic movement, a concentration gradient of Pi must be formed between the cytoplasm of cells and phloem through the plasmodesmata. Such a concentration gradient of Pi between cells has not been detected. For the apoplastic pathway, the Pi status of the external phases must be known. Katou and Enomoto (1991) mathematically demonstrated the existence of concentration gradients in the apoplast in order to explain a driving force for the water transport in growing tissues. They named a small and thin space of the cell wall con-

b

a +Pi culture

4 -Pi culture

-Pi culture

40

-b

-Pi culture

14

16

h

v

4

30

h

60

j .

B

“a 4 0

2 N

a

20

v

N

PI m

m

N

0

8

10

12

14

16

18

Days after sowing

0 20

8

10

12

18

20

Days after sowing

FIG. 11 Numerical analysis of 32P distribution in autoradiograms. Diamonds. whole plants;

inverted triangles. root: circles, the first leaf; triangles. the second leaf, squares, the third leaf. In (a), radioactivity of Pi-adequate barley was chased in the absence of Pi. In (b), radioactivity of Pi-deficient barley was chased in the absence of Pi (from Mimura, T., Sakano, K., and Shimmen, T., Studies on distribution, re-translocalion, and homeostasis of inorganic phosphate in barley leaves, Planr Cell Envirnn., with kind permission from Blackwell Science).

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nected to the xylem the “apoplast canal.” In these canals, the extracellular concentrations of solutes decrease proportional to the distance from the xylem as these solutes are taken up into cells (Katou and Okamoto, 1992). A similar situation occurs in solute retranslocation from cells to the phloem through the apoplast. Pi effluxed creates a concentration gradient toward the phloem. The Pi level in the phloem is a major factor in determining and controlling Pi translocation. We have recently measured the Pi level in the phloem of rice using the aphid method (Mimura et al., unpublished data). Under Pi deficiency the Pi level of the phloem decreased to one-fifth its original value, in contrast to root or mesophyll cells. This was also confirmed by Jeschke et al. (1997). The phloem appears to lack a homeostatic mechanism for Pi, but the large fluctuation in phloem Pi content may be a necessary part of the regulation of long-distance Pi transport as suggested by Clarkson and Saker (1988). In Arabidopsis, the Pho2 mutant (Delhaize and Randall, 1995) accumulates Pi in the shoot more than the wild type. Dong et al. (1998) suggested that the Pi accumulation in shoot might result from a defect in Pi uptake to the phloem in Ph02, i.e., the Pi accumulation in the shoot was due to the depression of retranslocation from shoot to root through the phloem. They proposed an alternative explanation-that the shoot cells lacked the ability to regulate intracellular Pi concentration. The latter could arise if there was a defect in the mechanism for export of Pi from the vacuole. To accomplish Pi homeostasis, a complex system is necessary-detection of Pi levels in each compartment, changes in the distribution of Pi between organelles in the cell or between cells in a whole plant, or changes in metabolic processes. After attainment of a new Pi state, the new level of Pi might be referred back to the Pi detection system (Fig. 12)-a conventional feedback system. In a simpler alternative model, the strongest Pi sink assumes a central role. If the cytoplasm in the cell or the youngest tissues in the whole plant absorbs Pi according to demand, the other parts may respond dependent on their individual Pi status. In this system Pi levels are driven by the demand from the strongest sink and no feedback system is needed. I believe that the former system operates at the cellular level, but the latter may be sufficient between cells and tissues.

VI. Concluding Remarks Phosphorus is one of the most essential elements in not only plants but also animals. All phosphorus in living organisms is derived from Pi absorbed


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// FIG. 12 A hypothetical diagram of Pi homeostasis in a plant cell. Putative sensor seems t o respond to the following factors; 1, the cytoplasmic Pi (andlor organic-P) level; 2. the extracellular Pi level: or 3. Pi flux across the membrane. Afterwards, the sensor would influence the membrane transport systems of both plasma membrane and tonoplast. and the Pi metabolisms directly or indirectly. There are also interactions between transport systems and metabolisms, for example. an inhibition of H+-ATPase by Pi.

by plants. The total amount of phosphorus easily accessible by plants in the environment is quite limited. Because of limited phosphorus, plants have developed various adaptive systems, i.e., high-affinity Pi uptake, storage of Pi in the vacuole, retranslocation of Pi between tissues, and secretion of phosphatases or organic substances. I have undoubtedly concentrated on those aspects of phosphorus which are closest to my own research interest, perhaps at the expense of other, equally important aspects such as the metabolic processes involving phosphorus. Owing to the importance of phosphorus, there is a multitude of published papers, and it was impossible for me to refer to them all. I hope that I have not overlooked any of the important studies. There is clearly a need to unify all knowledge about Pi in plants, i.e., Pi transport and Pi-related metabolism. The recent molecular approaches of both fields are certainly making clear the molecular networks for Pi in the cell, especially mechanisms for Pi uptake across the plasma membrane. However, there is still a large gap between physiological and molecular research which needs to be bridged. Furthermore, we may need to consider more carefully the relationship between global Pi circulation

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and plant availability in order to maximize agricultural production while protecting natural environments.

Acknowledgments I express my best gratitude to Prof. M. Tazawa (Fukui Institute of Technology), Prof. T. Shimmen (Himeji Institute of Technology), Prof. U. Heber (University of Wuurzburg), Dr. K. Sakano (National Institute of Agrobiological Resources), Prof. K.-J. Dietz (University of Bielefeld), Prof. E. Martinoia (University of Neuchatel), Prof. F. A. Smith (University of Adelaid), Dr. R. J. Reid (University of Adelaide), and all my colleagues for their kind and critical discussions. Especially, 1 express my sincere appreciation to Dr. Reid and Prof. Shimmen for their kind reading and many suggestions for the manuscript. I also thank Yokogawa Analytical Systems, Inc., and Dr. Y. Inoue for their continuing support to measure phosphates.

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Synaptic-like Microvesicles in Mammalian Pinealocytes Peter Redecker Medizinische Hochschule Hannover, D-30625 Hannover, Germany

The recent deciphering of the protein composition of the synaptic vesicle membrane has led to the unexpected identification of a compartment of electron-lucent microvesicles in neuroendocrine cells which resemble neuronal synaptic vesicles in terms of molecular structure and function. These vesicles are generally referred to as synaptic-like microvesicles (SLMVs) and have been most intensively studied in pancreatic pcells, chromaffin cells of the adrenal medulla, and pinealocytes of the pineal gland. This chapter focuses on the present knowledge of SLMVs as now well-established constituents of mammalian pinealocytes. I review the results of morphological, immunocytochemical, and biochemical studies that were important for the characterization of this novel population of secretory vesicles in the pineal organ. The emerging concept that SLMVs serve as a device for intercellular communication within the pineal gland is outlined, and unanswered questions such as those pertaining to the physiological function and regulation of pineal SLMVs are discussed. KEY WORDS: Pineal gland, Secretory vesicles, Synaptic membrane proteins, Paracrine communication, Neuroactive amino acids, Neuroendocrine cells, Melatonin. 0 1999 Academic Press.

1. Introduction Many endocrine cells share common structural and functional features with neurons and have therefore been classified as members of the diffuse neuroendocrine system or of the family of paraneurons (Pearse and Takor Takor, 1979; Fujita et al., 1988). Importantly, these similarities also manifest themselves in the regulated secretory pathways of neurons and neuroendocrine cells. Thus, it has been known for some time that the peptide hormone/~ltc'r?l

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