Role in Cell Physiology (The Cytoskeleton)

ROLE IN CELL PHYSIOLOGY

THE CYTOSKELETON

A Multi-Volume Treatise, Volume 2

This Page Intentionally Left Blank

ROLE IN CELL PHYSIOLOGY Editors: JOHN E. HESKETH Rowett Research Institute Aberdeen, Scotland IAN F. PRYME Department of Biochemistry and Molecular Biology University of Bergen Bergen, Norway

VOLUME 2 •

1996

UHU) jAI PRESS INC. Greenwicli, Connecticut

London, England

Copyright © 1996 byJAI PRESS INC. 55 Old Post Road, No. 2 Greenwich, Connecticut 06836 JAI PRESS LTD. The Courtyard 28 High Street Hampton Hill Middlesex TW12 IPD England All rights reserved. No part of this publication may be reproduced, stored on a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, filming, recording, or otherwise, without prior permission in writing from the publisher. ISBN: 1-55938-688-6 Manufactured in the United States of America

CONTENTS

LIST OF CONTRIBUTORS

vii

INTRODUCTION John Hesketh and Ian Pryme

ix

O N THE ROLE OF THE CYTOSKELETON IN METABOLIC COMPARTMENTATION Colin Masters

1

ASSOCIATION OF mRNAS A N D POLYRIBOSOMES WITH THE CYTOSKELETON: POTENTIAL ROLES IN TRANSPORT A N D COMPARTMENTALIZATION OF mRNAS John E. Hesketh and Ian F. Pryme

31

SPECIALIZATIONS IN CYTOSKELETAL FUNCTION DURING EARLY DEVELOPMENT David G. Capco

59

MICROTUBULE-BASED INTRACELLULAR TRANSPORT OF ORGANELLES Howard Stebbings

113

ROLE OF THE CYTOSKELETON IN THE DEVELOPMENT OF EPITHELIAL POLARITY Detlev Drenckhahn, Thomas Jons, Bernd Puschel, and Frank Schmitz

141

FOCAL ADHESIONS A N D INTEGRIN-MEDIATED CELL SIGNALING Susanne M. Bockholt and Keith Burridge

167

vi

CONTENTS

INTERACTIONS OF MEMBRANE RECEPTORS AND CELL SIGNALING SYSTEMS WITH THE CYTOSKELETON Coralie A. Carothers Carraway and Kermit L. Carraway

207

FUNCTION OF MICROTUBULES IN PROTEIN SECRETION AND ORGANIZATION OF THE GOLGI COMPLEX Jaakko Saraste and Johan Thyberg

239

INDEX

275

LrST OF CONTRIBUTORS

Susanne M. Bockholt

Department of Biology University of Utah

Keith Burridge

David G. Capco

Department of Cell Biology and Anatomy University of North Carolina, Chapel Hill Molecular and Cellular Biology Program Arizona State University

Coralie A. Carothers Carraway

Department of Biochemistry and Molecular Biology University of Miami School of Medicine

Kermit L. Carraway

Department of Cell Biology and Anatomy University of Miami School of Medicine

Detlev

Institute of Anatomy University of WiJrzburg Wurzburg, Germany

Drencl
Jotin E. Hesl<eth

Rowett Research Institute Aberdeen, Scotland

Thomas Jons

Institute of Anatomy University of Wiirzburg Wurzburg, Germany

Colin Masters

Faculty of Science and Technology Griffith University Brisbane, Australia

VII

LIST OF CONTRIBUTORS Ian F. Pryme

Department of Biochemistry and Molecular Biology University of Bergen Bergen, Norway

Bernd Puschel

Institute of Anatomy University of Wurzburg Wurzburg, Germany

Jaakko Saraste

Department of Biochemistry and Molecular Biology University of Bergen Bergen, Norway

Frank Schmitz

Institute of Anatomy University of Wurzburg Wurzburg, Germany

Howard Stebbings

Department of Biological Sciences Washington Singer Laboratories University of Exeter

Johan Thyberg

Department of Cell and Molecular Biology Medical Nobel Institute Karolinska Institute Stockholm, Sweden

INTRODUCTION During the last 10 years it has become evident that the cytoskeleton is intimately involved in different aspects of cell physiology. The chapters in this second volume of the The Cytoskeleton, A Multi-Volume Treatise, describe a wide variety of cell functions in which the cytoskeleton has been either implicated or shown to have a role; the emphasis is on its role in general cell processes rather than specialized aspects in particular cells or tissues which will be described in volume three. A persistent theme throughout this volume is the important role that the cytoskeleton plays in compartmentation, targeting, and subcellular organization. For many years cell biologists and biochemists have speculated as to whether there is compartmentation and spatial organization of metabolic reactions within the cytoplasm. The discovery of the cytoskeleton provided a possible mechanism for such subcellular organization, but to date conclusive evidence for a role of the cytoskeleton in metabolic compartmentation has remained elusive. The possible association of glycol3^ic enzymes with actin is discussed in the context of metabolic compartmentation by Masters. As reviewed by Hesketh and Pryme in the second chapter, there is also an increasing body of evidence that a proportion of both polyribosomes and mRNAs is associated with the cytoskeleton; such interactions may have important roles in mRNA localization and the spatial organization of the protein synthetic apparatus. Compartmentation and spatial organization is particularly evident in embryonic development and, as described by Capco, the cytoskeleton plays an important role in the early stages of development. ix

X

INTRODUCTION

Cell organization involves transport of material within the cell, as particularly illustrated by axonal transport. The cytoskeleton has well defined roles in organelle transport and this is discussed in the chapter by Stebbings. As described in detail by Drenckhahn and colleagues, such transport by the cytoskeleton can be highly specific in a spatial sense and this allows the cytoskeleton to contribute to the generation of cell polarity; furthermore the interaction of membrane proteins with the cytoskeleton may be the basis of domains within the membrane. The interaction of the cytoskeleton with membrane proteins also has other important functions: as discussed by Bockholt and Burridge, the best-characterized is the interaction of the cytoskeleton with the extracellular matrix which brings about cell adhesion and movement; there is also increasing evidence that links between membrane receptors and the cytoskeleton are important in signaling processes between extracellular and intracellular environments (Carraway and Carraway); finally, Saraste and Thyberg discuss the evidence that the cytoskeleton is involved in secretion. John Hesketh and Ian Pryme Editors

ON THE ROLE OF THE CYTOSKELETON IN METABOLIC COMPARTMENTATION

Colin Masters

I. II. III. IV. V VI. VII. VIII.

Introduction 2 Evidence for the Micro-Compartmentation of Carbohydrate Metabolism . . . . 3 Enzyme Multiplicity and Interactions with Cellular Structure 9 Variation of Structure Within the Cytoskeleton 11 CovalentModificationof Enzymes and Cellular Structure 13 Energy Requirements for Signal Transduction 16 Matrical Compartmentation 18 Perturbations of Compartmentation During Cellular Differentiation and Dysfunction 21 IX. Compartmentation in Intermediary Metabolism 25 X. Concluding Comments 26 References 28

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 1—30 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 1

2

COLIN MASTERS

I. INTRODUCTION There is increasing evidence that many of the cytoplasmic enzymes exist in vivo as part of an organized structural system that provides a framework for the coordination of metabolic activities (Masters, 1981; Clegg, 1984; Srere and Ovadi, 1990). This evidence has been derived from a number of divergent techniques, but probably the most complete and systematic series of investigations into this phenomena in recent years has been those studying the interactions between glycolytic enzymes and the cytoskeleton (Humphreys and Masters, 1986; Chen et al., 1986; Chen and Masters, 1988; Shearwin et al, 1990 a, 1990b; Masters, 1991, 1992). Using this data as an appropriate base, this review seeks to draw attention to the need for modification of many of the classical views of cytoskeletal function and the regulation of intermediary metabolism. Historically, for example early research into intermediary metabolism assumed that individual metabolic pathways were controlled by single regulatory enzymes— an assumption which is no longer recognized as valid (Keleti and Ovadi, 1988). In addition, recent studies have served to emphasize other regions of potential inappropriateness in the classical biochemical approaches to this topic: many classical studies have tended to rely on investigations with purified enzymes in dilute aqueous solution, and assume that cell metabolism is merely a linear superposition of the kinetic characteristics of single enzymes established under these conditions; alternatively, many other metabolic investigations have been carried out using cytosolic fractions, prepared by the classical methods of subcellular fractionation, and viewed as closely approximating metabolism in the cytoplasmic compartment of the cell. Without wishing to decry the substantial advances achieved in the past by these methodologies, and their contribution to our understanding of normal and abnormal carbohydrate metabolism, it needs to be recognized that, in the light of present knowledge, these procedures do not provide a fully satisfactory simulation of cellular conditions in vivo. For example, the first of these methodologies does not allow for the high protein concentration in cells, nor the marked influence of this molecular crowding on the interactions and kinetic characteristics of individual enzymes, while the second approach also generally involves a dilution effect. As well, and central to the thrust of this review, both methods disregard any contribution of the cytoskeleton towards the compartmentation of carbohydrate metabolism. With regard to the cytoplasmic compartment, for example, where a major part of intermediary metabolism is located, the concept of self-organization via transient macromolecular associations has received increasing support of late (Srere and Ovadi, 1990). There is now compelling evidence that much of the intermediary metabolism in living cells is carried out within the confines of microenvironments such as that engendered by enzyme-cytomatrix assemblages, and a growing realization of the critical importance of such positional factors to our understanding of the living state (Masters, 1981, 1992; Clegg, 1984). Such assemblies of enzymes

The Cytoskeleton and Metabolism

3

offer the possibilities of increased efficiency of the overall processes due to proximal juxtaposition of active sites, reductions in substrate transit times, variable localization within the cell, and ready response to variations in metabolic status. The micro-compartmentation of glycolysis deserves especial consideration in this context as a central element of control in carbohydrate metabolism, and with this in mind, this article reviews the available data on the microcompartmentation of carbohydrate metabolism, and comments on localized enzyme associations, the heterogeneity of cytoskeletal structure, the covalent modification of enzymes and structure, energy requirements during signal transduction, the perturbations of micro-organization during cellular dysfunction, and the role of the cytoskeleton in modulating intermediary metabolism, in general.

11. EVIDENCE FOR THE MICRO-COMPARTMENTATION OF CARBOHYDRATE METABOLISM It has been clearly established by a variety of techniques that an extensive, differential binding of glycolytic enzymes to the cytoskeleton exists in most mammalian cells (Masters, 1984, 1992). Certain of the glycolytic enzymes bind more-readily than others and included in this category are the specific binding characteristics of phosphofructokinase, aldolase, glyceraldehydephosphate dehydrogenase, pyruvate kinase, and lactate dehydrogenase. Many of these enzymes have been shown to possess structurally distinct binding sites for substrate and for actin (Humphrey et al., 1986), and interactions with intracellular structures allow these enzyme activities to be positioned in the cell near regions involved in dynamic activities (e.g., the contractile units of muscle, or the cytoskeleton), and hence to contribute to rapid energy production at just those positions in the cell where it is most required. Adding to the potential metabolic advantage of these associations are two other features of the glycolytic enzyme-actin interactions which have emerged from previous studies. These are the modification of enzyme kinetics which occur concomitantly with binding, and the interrelationship between the degree of binding and the emphasis of cellular metabolism. When aldolase binds F-actin-tropomyosin-troponin, for example, the K^ value is increased by two orders of magnitude, whereas Vj^^^ ^^^^^ four-fold relative to free aldolase. Again, in conditions of elevated glycolysis (such as during muscle contraction), a markedly increased degree of binding of the glycolytic enzymes to cellular structure was observed (Masters, 1981, 1984; Figure 1). A number of workers have noted that micro-compartmentation of this type appears to be of major consequence in the regulation of carbohydrate metabolism, and intimately involved in aspects such as the balance of the aerobic versus the anaerobic glycolytic rate (Masters, 1981; Storey, 1985). While aerobic metabolism is of major importance in energy production, for example, many organisms retain anaerobic pathways of metabolism, which despite their low energy yields, can be of great value in periods of stress where demand outstrips the aerobic capacity (e.g.,

COLIN MASTERS

o

I (D

40

80 120 160 Fructose 1,6 - bisphosphate, mM

200

Figure 1. Substrate saturation curves of free and bound aldolase A4. A. Free Enzyme; B. Enzyme bound to actin-tropomyosin-troponin.

periods of intense muscular contraction). The strategy here involves a greatly increased catabolism of substrate to compensate for the relatively lov^ energy yields, and the experimental data indicates that the aerobic to anaerobic transition is accompanied by a shift in the soluble: particulate association of glycolytic enzymes which generally over-rides allosteric type controls and allov^s for unfettered, maximal expression of activities of these enzymes (Masters, 1984; Storey, 1985). There is also evidence that in the cell the sequence of glycolytic enzymes may often operate as a composite of shorter metabolic sequences, each v^ith a degree of independent function and associated v^ith the cytoskeleton, rather than as an entire and soluble sequence as described in most textbooks, or as a complex of all the glycolytic enzymes as postulated by Kurganov and coworkers (1985). It has been proposed, for example, that aldolase (ALD), glyceraldehydephosphate dehydrogenase (GAPDH), triosephosphate isomerase (TPI) and phosphoglycerokinase (PGK) form one cluster of enzymes which may function on its own to produce energy—^and indeed may possibly be the most important anaerobic energy source available to most cells (Masters, 1981; Figure 2, Masters et al., 1987). Studies with purified components have indicated that both aldolase and GAPDH bind to actin with comparatively high affinity, and the binding sites for these two enzymes are closely adjacent and periodically spaced on the actin filaments. Thus, a mechanism

The Cytoskeleton and Metabolism

,3PGA

Figure 2. Diagrammatic representation of the association between a segment of the glycolytic sequence and an actin-containing filament, with indications of the attendant possibilities of metabolic channelling.

for the anchoring of the metabolic segments to the structural elements is readily available (Masters 1981, 1984; Humphreys et al., 1986). With regard to the other two components of this segment, TPI and PGK, it has been shown that they bind to actin with far lower efficiency than aldolase and GAPDH; but interestingly, once this complex is an achored by an initial binding of ALD and GAPDH to cytoskeletal components, then TPI and GPK may add on and bind quite firmly—the aptly termed phenomenon of piggy-backing or facilitated binding. Thus the formation and stabilization of this particular cluster of four enzymes can occur, and this in turn allows the possibilities of metabolic channeling and the advantageous positioning of this cluster in appropriate locations within the cell. While the original proposal for the existence of this enzyme cluster was mainly based on binding characteristics such as those outlined above (Masters 1981), it is of interest to note that this concept has received independent kinetic confirmation recently. Han et al. (1992) have demonstrated by kinetic means that skeletal muscle contains a compartmentalized reaction sequence consisting of these four glycolytic enzymes (aldolase, triosephosphate isomerase, glyceraldehyde-3-phosphate dehydrogenase, and phosphoglycerate kinase) which in the structure-associated state (i.e., bound to actin containing filaments) is active in the synthesis of ATP in the triadic junction. They also provided evidence that fructose-1,6-bisphosphate was especially effectively channelled in this system, and that the function of this enzyme cluster was linked to some of the major cell signalling systems (e.g., the phospholipase C and protein kinase systems). Other enzyme clusters in the glycolytic sequence—^four in all—^are similarly indicated as being able to function with increased efficiency under appropriate

6

COLIN MASTERS

cellular conditions, and these clusters are illustrated in Figure 3. Several studies of micro-environmental and ontogenic variation have pointed to the independent functioning and positioning of these clusters within the cell, and to the associated consequences of an increased flexibility and appropriateness of the glycolytic processes with regard to their many interrelated physiological roles (Masters, 1992). As has been mentioned in preceding sections, there are major difficulties in defining the detail of biphasic interactions between enzymes and cell structure by means of classical procedures such as subcellular fractionation. Novel experimental

I

ATP-N ADPV

GLUCOSE

I HK

ENERGY CONSUMING (PRIMING) SEGMENT

GLUCOSE -6 - PHOSPHATE I GPI FRUCTOSE - 6 - PHOSPHATE PFK

ATP ^ ADP r\ur^^^

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n ENERGY PRODUCING SEGMENT

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ENERGY PRODUCING SEGMENT

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PHOSPHOENOUPYRUVATE 2 ADP 2 ATP^

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PK

PYRUVATE

IV ANAEROBIC SEGMENT

Figure 3.

2NADH-^ 2 NAD ^

•-irj

LDH

LACTATE

Representation of the glycolytic sequence as a series of segments.

The Cytoskeleton and Metabolism

7

approaches which allow the retention of cytoskeletal structure and the close simulation of cellular conditions in vivo are required (Masters, 1992). One particular technique which has proved particularly valuable in assessing the role of the cytoskeleton in enzyme-structure interactions is the use of digitonin-treated cells, where the cell membrane is perforated but the cytoskeletal structure preserved (Mackall et al, 1979; Humphreys and Masters, 1986). The use of this technique, for example, has allowed the extent of interactions between enzymes and cell structure under physiological conditions to be assessed. Maretski et al. (1989) based their speculation that such interactions were non-physiological on a comparison of the extent of the binding of the glycolytic enzymes under hypotonic and isotonic conditions, with the diminished binding in the latter situation leading to their negative conclusion. However, in their experiments these authors failed to give recognition to one of the major factors in establishing the characteristic status of the cellular milieu, namely protein concentration. When the digitonin-treated cell system was used to test the effect of physiological concentrations of both protein and electrolytes on the binding of glycolytic enzymes to the cytoskeleton, retention of the enzyme-structure associations was shown to be significantly increased (Shearwin et al., 1990a, 1990b), with the volume occupied by the added protein crowding the cellular protein into a smaller space, and favoring hetero-associations such as those between glycolytic enzymes and actin (Figure 4). Clearly, then, the positive influence of molecular crowding needs to be included in any assessment of the physiological significance of the binding of glycolytic enzymes to cytoskeletal structure. In a similar manner, the influence of a number of metabolic effectors (specific metabolites, hormones, second messengers) on the binding of glycolytic enzymes to the cytoskeleton have been tested by the digitonin-treated cell technique. When, for example, the effect of physiological levels of fructose-1,6-bisphosphate (F-1,6P2) on such interactions was tested in the cellular situation a significant and specific effect on the desorption of aldolase was noted without any marked perturbation on the binding of other glycolytic enzymes (Chen et al., 1986). F-l,6-P2 is, of course, the major physiological substrate of aldolase, and in the bound state, increased concentrations of F-l,6-P2 would be expected to provide an increased ability to interact with the sequentially vicinal enzyme glyceraldehydephosphate dehydrogenase, resulting in a higher flux rate. On the other hand, further increases in the concentration of F-1,6-P2 lead to a decreased binding of aldolase to the cytoskeleton and a marked alteration in the catalytic properties of this enzyme. Thus, these interactions may modulate the flux rate, and perhaps the direction of triosephosphate metabolism, depending on the concentration of F-l,6-P2 in that particular region of the cell (Keleti and Ovadi, 1988). Also, it may be noted that an alteration in the degree of binding of enzymes in response to changes in the concentration of cellular metabolites is a key element in the concept of enzyme ambiguity (Wilson, 1978)—that is, that the extent of interaction of enzymes with cellular structure varies in response to the metabolic status of the cell.

COLIN MASTERS

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GLYCOLYTIC ENZYMES Figure 4, The differential retention of glycolytic enzymes by the cytoskeleton of cultured animal cells which have been permeabilized with digitonin. The open histograms represent retention when cells were treated with protein free release buffer, and the shaded area of the histogram represents the increase in retention when the release buffer contained 0.15% saline and 20% protein (after Shearwin et al., 1990).

When the influence of pH on binding was studied in digitonin-treated cells (Shearwin et al., 1990b) it was noted that a decrease of pH within the physiological range was accompanied by an increase in binding of many of the glycolytic enzymes to the cytoskeleton, and it was noticeable that amongst the enzymes most markedly influenced were those most traditionally cited as control points in the glycolytic sequence (e.g., phosphofructokinase and pyruvate kinase). It seems likely therefore that these observed changes may well be significant in relation to the control of glycolysis. It is also of interest to note that the glycolytic enzymes which showed increased binding to the cytoskeleton under acidic conditions in these experiments in vitro are precisely the same as those which were observed to bind to actin filaments in significantly increased quantities during increased muscular activity in vivo and its associated lowering of cellular pH (Walsh et al., 1981). Two other points are worth mentioning in this general context of biphasic interactions. Firstly, in relation to associations between the cytoskeleton and

The Cytoskeleton and Metabolism

9

glycolytic enzymes, it may be noted that interactions between tubulin and glycolytic enzymes such as glyceraldehyde phosphate dehydrogenase and aldolase have been noted (Kumagai and Sakai, 1983; Balaban and Goldman, 1990). Actin and tubulin are, of course, two of the most important structural components of the cytoskeleton, so that this particular conjunction of properties not only broadens the link between glycolytic enzymes and cytoskeleton, but also increases the scope for coordination of the energetic requirements of these cellular structures. Secondly, in relation to the spatial organization of carbohydrate metabolism within the cytoplasm, it may be noted (a) that actin filaments have been identified as localized near mitochondria (Pardo et al., 1983), and (b) the enzymes needed to carry out the synthesis and degradation of glycogen have been shown to be bound to the surface of the glycogen granules in the cytoplasm (Alberta et al., 1983). Both of these observations obviously have the effect of broadening the previous observations on the micro-compartmentation of glycolysis and enzyme-structure interactions, and allow an inclusion into these considerations of the aerobic and storage elements of carbohydrate metabolism, and the integration of their activities.

III. ENZYME MULTIPLICITY AND INTERACTIONS WITH CELLULAR STRUCTURE Another factor which is directly relevant to the extent of enzyme-structure association in the physiological situation is the occurrence of enzyme multiplicity. In higher organisms, most individual tissues display characteristic distributions of activity amongst multiple enzyme forms, and the occurrence of these specific patterns of heteromorph distribution also extends to the subcellular level (Masters and Holmes, 1975). Enzyme multiplicity, of course, may arise by several mechanisms. Multiple forms of an enzyme may arise from covalent modification, from posttranslational modification, or from differences in the primary amino acid structure. In any of these cases, the individual forms of an enzyme may interact quite differently with cellular structure and with metabolism, hence it is important to consider the isoenzyme status of any enzyme activity, and the cellular distribution of the multiple forms, before attempting any specific definition of these interactions (Masters, 1984). Most glycolytic enzymes exhibit distinctive isoenzyme profiles. Aldolase, for example, has been shown to exist in at least nine different forms in vertebrate species, with these isoenzymes representing the possibilities of hybridization between three types of gene product. A-type is the classical muscle type activity, B-type predominates in liver, and C-type occurs predominantly in brain (Masters and Holmes, 1975). Not only do these multiple forms possess distinctive kinetic characteristics, but they also display markedly different actin-binding properties between the individual multiple forms (Masters, 1981). The A-type enzyme is the form which has the highest affinity for actin fibers, and is of special interest in relation to the higher Kj^ and V^^^^ ^f ^^e bound form of the enzyme, and the

10

COLIN MASTERS

Table 1. Major Glycolytic Enzymes—Subunit Types and Predominant Distribution in Adult Mammalian Tissues Enzyme

Isoenzyme

Tissue Location

Phosphofhictokinase

A4 B4 C4

Muscle, Brain Liver Brain

Aldolase

A4 B4 C4

Muscle, Brain Liver, Kidney Brain

Pyruvate Kinase

A4 B4 C4

Muscle, Brain Liver Kidney

Lactate Dehydrogenase

A4 B4 C4

Muscle, Liver Heart, Brain Sperm

Specificity in the degree of desorption of this enzyme brought about by the substrate, fructose -1,6-bisphosphate (Masters, 1981). As has been noted, the existence of an equilibrium between free and bound forms of aldolase, with each form having different catalytic properties and the equilibrium between the forms being dependent on substrate concentration, presents the ingredients of a substantial control system for glycolysis (Masters et al, 1969). Apart from aldolase, a number of other glycolytic enzymes also exhibit a considerable degree of heterogeneity (Table 1), and there are several other instances of differential binding capacities amongst the multiple forms of individual glycolytic enzymes, of modification of kinetic characteristics on binding, and functional relationships of these phenomena (Masters, 1981). Phosphofhictokinase, pyruvate kinase and lactate dehydrogenase, in particular, display a marked propensity for binding to actin-containing filaments, with the affinity of this binding varying markedly amongst the individual multiple forms. It is notable, though, that it is the muscle-type isoenzymes that associate most strongly with the cytoskeleton in a variety of tissues. The other tetrameric component of the glycolytic system which binds extensively to the cytoskeleton is glyceraldehyde phosphate dehydrogenase, but it may be noted in this case that the enzyme does not display an extensive multiplicity in mammalian tissues. It has been argued that the reason for this is that the enzyme plays a key structural role in the cell, interacting with a number of components (such as actin, tubulin and the band III protein), and hence being of particular significance in locating glycolytic enzyme sequences at appropriate points in the cytomatrix—^with the attendant necessity for maintaining all of these binding sites leading to evolu-

The Cytoskeleton and Metabolism

11

tionary constraints, a high degree of conservation and the lack of multiple forms (Reid and Masters, 1986). As discussed in a later section of this review, it may further be noted that many of the key protein components of the signaling pathways associated with glycolytic events also bind to the cytoskeleton and exist in multiple enzyme forms, and in a wider context, it may be commented that enzyme heterogeneity is a general phenomenon, which is common to most areas of metabolism in the cell. Giving regard to all these observations then, it is clear that, when investigating interactions between glycolytic (or other) enzymes and cell structure, it is important to give consideration to the established differences between the binding constants for particular isoenzymes and different types of actin fibers, and to define both the isoenzyme status of the catalytic component and the exact nature of the structural component.

IV. VARIATION OF STRUCTURE WITHIN THE CYTOSKELETON As mentioned above, one of the salient features of previous studies into the interactions between glycolytic enzymes and the cytoskeleton has been the marked diversity in binding characteristics between the different forms (isoenzymes) of the glycolytic enzymes. This heterogeneity of response of the catalytic component of any enzyme-cytoskeleton association points to a need to also examine the implications of variation in the structural component, that is, cytoskeletal heterogeneity. Many text book-accounts, perhaps constrained by a requirement for concise expression, tend to direct the great majority of their comment towards the principal structural characteristics of the cytoskeleton, and these limitations often leave an overall impression of a predominantly static and homogeneous character of cytoskeletal structure. A more detailed consideration of the functional and structural data as described in other chapters of this treatise, however, leads to a very different conclusion. At the functional level, regulated changes in cytoskeletal structure are now recognized as occurring in a large number of biological responses. Movement in cells—change of shape, the repositioning of internal organelles, and migration of the whole cell or its cytoplasm—^is acknowledged as being dependent on the cytoskeleton, and playing a prominent role in these various processes, is the major structural component, F-actin, which is assisted in its dynamic roles by a variety of actin-binding proteins (e.g., myosin, tropomyosin, filamin, a-actin, filamin, fimbrin, profilin, villin, gelsolin, and capping proteins). While the role of these binding proteins in modifying the structure of the cytoskeleton to suit the differential motility requirements of cells has been extensively documented and is well recognized (Pollard et al., 1984), less acknowledgment has been given to the established involvements of the glycolytic enzymes in modifying the functional role of the cytoskeletal action (see later sections of this review).

12

COLIN MASTERS

In line with this estabhshed diversity of both function and means of structural modulation, much information at the research level refers to an evident heterogeneity in the cytoskeletal structure, and a variation in functional involvement in different locations within the cell. Morphologically, bundles of parallel filaments, cross-linked by proteins such as fimbrin, have been observed as present in the microvilli and sterocilia, and appear to be mainly involved in a structural role, whereas other bundles of actin filaments are associated with myosin and found in regions of the cell where contractile processes occur (e.g., the contractile ring of the dividing cell, and the belt desmosomes). In addition, further networks of actin filaments are found in a cortical layer adjacent to the plasma membrane-networks, which are thought to be responsible for a variety of cell surface movements. These networks appear to be formed by flexible, actin-cross-linking proteins, and they undergo a calcium induced decrease in viscosity that is mediated by actin fragmenting proteins. By interacting with non-muscle myosin and with proteins that anchor them to the plasma membrane, these networks are thought to be responsible for a variety of cell surface movements, and to play a crucial part in the complex process of cell locomotion (Heuser and Kirschner, 1980; Porter et al., 1983). Clearly, then, a degree of variability and heterogeneity in the cytoskeletal structure of most cell types is recognizable even by the comparatively gross morphological parameters which have been in common use up to the present time. In relation to the functional-structural correlations of these variations in cytoskeletal morphology, comparisons are of course limited by the nature of the information on morphological heterogeneity which has been available. It is of interest, though, that much of the data points to the cortical layer of the cytoskeleton as most involved in the shifts of enzyme distribution which accompany physiological variations and abnormalities of carbohydrate metabolism. The transition from aerobic to anaerobic metabolism in many tissues for example, is recognized as accompanied by a marked shift in the soluble-particulate association of the glycolytic enzymes, and an increased binding in the cortical region of the cytoskeleton (Masters, 1984; Storey, 1985). Similarly, it is known that peptide hormones affect the shape of cells quite dramatically, and these morphological changes are accompanied by the appearance of numerous cytoplasmic processes attached to the substratum, and an increased binding of the glycolytic enzymes thereto (BereiterHahn, 1988; Masters, 1992). In the knowledge that the actions of peptide hormones are effected via stimulation of adenyl cyclase, and elevation of intracellular cAMP, it is also of interest to note that treatment of cultured cells with cAMP alone results in an increased binding of glycolytic enzymes to peripheral cytoskeletal structures (Masters, 1992). Growth factors, and the second messenger inositol triphosphate also produce similar responses; and changes in the extent and intracellular location of these interactions between the glycolytic enzymes and cytoskeletal structure have also been observed in many of the major disorders of carbohydrate metabolism, as discussed in later sections of this review.

The Cytoskeleton and Metabolism

13

As actin modifies enzyme structure and function, so the reciprocal action of enzymes on actin structure and function needs to be considered—^a point first made by Masters (1975) and based on the observations that under certain conditions, glycolytic enzymes can organize actin filaments into highly ordered filament bundles (Masters, 1975; Morton et al., 1976). Enzymes such as aldolase and GAPDH are large molecules which are present in high concentration in cells, and each enzyme has its own unique tertiary and quaternary structure which contributes to its specific actin binding properties. The experimental evidence indicates that the conformations and actin binding of these enzymes are under exquisite metabolic control via the highly evolved and specific binding sites for their substrates, products, coenzymes and other metabolic effectors (Table 3). Aldolase -FBP binds in a totally different fashion to aldolase, for example, and the presence of other glycolytic enzymes can also have a marked effect on the nature of the actin-aldolase assembly. The available data demonstrates the potential of specific glycolytic metabolites to regulate the physical properties of actin-enzyme complexes, and by such means, glycolytic metabolism may be rendered responsive to and integrated with the regulation of cytoskeletal structures; an apposite combination of substrate and enzyme may act to bring about an appropriate conjunction of enzyme and structural element, topographically and functionally linked so as to provide the cell with an appropriate source of energy just where required within the cell. Clearly the glycolytic enzymes deserve more consideration as actin-binding proteins which affect the nature and function of cytoskeletal structure, as well as their major catalytic role in particulate and non-particulate metabolism. It should be noted in relation to the modification of cytoskeletal properties, that the cortical regions of the cytoskeleton appear to be formed by flexible, actin cross-linking proteins, and that the glycolytic enzymes possess a demonstrated ability to form cross-links in just such a manner (Masters, 1975). Overall, then, these variations of enzyme binding within different regions of the cytoskeleton may be said to provide strong indications of the potentially broad metabolic ramifications of these interaction phenomena.

V. COVALENT MODIFICATION OF ENZYMES AND CELLULAR STRUCTURE Another important aspect of enzymic and structural heterogeneity, and the influence of the interactions between enzymes and the cytoskeleton on the integration and control of metabolic processes, is the covalent modification of protein structure. Enzyme modification by the covalent incorporation of phosphate, for example, is a wide-ranging phenomenon which has significant consequences in relation to metabolic control in vivo (Cohen, 1980), and often provides a means for considerable changes in enzyme activity over a short time scale. The process of phosphorylation is stimulated by cAMP and Ca'^Vcalmodulin, and is in turn responsive to hormonal and growth factor influences. Included among the variety of regulatory

14

COLIN MASTERS

enzymes which are subject to this type of covalent modification are phosphofructokinase, pyruvate kinase, glycogen phosphorylase, glycogen synthase, acetyl CoA carboxylase and triglyceride lipase, as well as a number of glycolytic enzymes (Cohen, 1980; Li et al., 1988; Sukhodolets et al, 1988). Covalent modification by the phosphorylation or dephosphorylation of enzymes also appears to play a major role in controlling the relative emphasis of aerobic versus anaerobic glycolysis in many tissues, and in the association of these enzymes with cellular structure (Storey, 1985). In the case of the important control enzyme, phosphofructokinase, for example, there are accumulating indications of an association of the covalent modification of this enzyme with anoxia (Storey, 1985), and an increased activity of the enzyme in association with actin filaments (Kuo et al, 1986). In general, significant kinetic differences occur between the dephosphorylated and phosphorylated forms of mammalian phosphofructokinase, with the phosphorylated form of the enzyme being more sensitive to ATP and citrate, and less sensitive to the influence of activators (Foe and Kemp, 1982). Similarly, covalent modification as a regulatory mechanism for glycolysis during anaerobiosis has also been shown to be the case with another important control enzyme in glycolysis, pyruvate kinase, with mammalian tissues using the phosphorylation of pyruvate kinase to regulate glycolytic versus gluconeogenic flux, in the main (Engstrom, 1978). More generally, the phosphorylation of amino acid residues such as serine clearly alters the charge distribution on the protein substrate, introducing a negative charge and influencing its conformation and binding characteristics (Figure 5). It is also noticeable, that the phosphorylated forms of regulatory enzymes generally lose their capacity for allosteric modification of activity; and this has led to the view that the combination of altered binding to the cytoskeleton and loss of allosteric control may combine to provide stronger and more far reaching influences on the regulation of anaerobic metabolism than minor changes in pH or allosteric effectors (Storey, 1985). Although not yet fully tested, this combination of mechanisms could obviously provide a powerful means for regulating metabolism, because unlike I allosteric controls which need to be tailored to individual enzymes, these controls could be applied to whole pathways. It is even likely that this combined mechanism may not only be involved in anaerobic glycolysis, but may also influence many other pathways. To cite an example outside of carbohydrate metabolism, it has been noted in relation to the regulation of the activity of cytidyltransferase and other enzymes, that phosphorylation of the enzymes alters both the activity and the extent of binding to cellular structure (Vance and Pelech, 1984). The association of binding of these enzymes and the over riding of allosteric controls in such cases indicates that metabolic biochemists need to look above the level of allosteric/kinetic control of enzyme activity, towards such higher tiers of metabolic control, and their potentially major roles in environmental stresses (Storey, 1985). Additionally, other types of covalent

The Cytoskeleton and Metabolism

wsm/\

mMmN

(b)

I^AAAAAAI

MAA.AAAM

1 m/jMH

ENZYME

FILAMENT

mmm

ENZYME - FILAMENT

Figures. Diagrammatic representation of the manner in which covalent modification (including charged groups) may lead to an increased association between enzyme and structure, through electrostatic attraction. (A) Introduction of charged groups to a binding site on the enzyme surface. (B) Introduction of charged groups to a binding site on a filamentous structure.

modification, for example, the methylation of specific amino acid residues, or the attachment of adenylate groups, may also be worthy of investigation in this respect. Furthermore, in addition to the alteration of enzymic characteristics, the involvement of covalent modification to the binding sites for enzymes on structural elements of the cytoskeleton should also be considered. As has been mentioned previously, the cytoskeleton is intimately involved in a number of motile processes (e.g., the movement of cells and organelles) which have an associated requirement for energy, and phosphorylation is recognized as a common effector of these processes. It is of interest in this connection then to consider an extension of the influence of covalent modification to the structural partner in these associations. Many of the enzyme-structure interactions which have been studied to date have been shown to be predominantly electrostatic in nature (Masters, 1984), and just as the phosphorylation of the amino acid residues in enzymes may alter the charge

16

COLIN MASTERS

distribution and influence the conformation and binding characteristics of the enzyme, so modification at or near a binding site on the cytoskeletal structure may also be expected to significantly influence the ability of this element to bind specific enzymes (Fig. 5). In this way, then, phosphorylation and the covalent modification of cellular structure should be considered as capable of exerting a powerful, if putative, influence on metabolic micro-compartmentation.

VI. ENERGY REQUIREMENTS FOR SIGNAL TRANSDUCTION In extending this consideration of enzyme-structure interactions to include phenomena such as signal transduction and the effects of hormones, agonists, and second messengers on the degree of association between glycolytic enzymes and the cytoskeleton, attention needs to be directed firstly to the molecular mechanisms underlying the transmission of signals across the cell membrane. While the complicated network of events which translates information from the extracellular environment into changes of cytoskeletal architecture are still incompletely understood, certain elements in the process may be discussed with some confidence. It is generally agreed, for example, that, among the host of transmitters participating in such processes, specificity is maintained by the presence of receptor proteins which present an appropriate region of their structure on the outer face of the plasma membrane (Lehninger, 1982). Interaction between an individual agonist and its particular receptor maintains the specificity in this relationship, and results in a conformational perturbation of the receptor protein, which in turn results in an altered confirmation of the receptor or an associated protein on the inner surface of the plasma membrane, and the transmission of a signal to the interior of the cell. In this context, the interaction between the agonist and its receptor, and the transmission of this signal across the plasma membrane by conformational change obviously induces a generalized energy requirement for such trans-surface processes. This energy requirement is even more in evidence when the membrane signal or transport is linked to a requirement for ATP. The sodium-potassium pump which occurs in the plasma membranes of virtually all animal cells and plays such an important role in the Na"*" and K"^ gradients, which in turn are responsible for the cells' membrane potential and for the active transport of sugars and amino acids, is dependent for its function on the hydrolysis of ATP (Proverbio and Hoffman, 1977). Similarly the calcium pumps in the plasma membranes of many eukaryotic cells are also ATP-ases (Wuytack et al., 1985), and indeed, on average, more than one third of the normal energy requirement of an animal cell is devoted to fueling these pumps. In the same context of the energy requirements inherent in trans-membrane signaling, the actions of a number of hormones and growth factors which act via cyclic AMP or inositol triphosphate and diacylglycerol as second messengers, deserve consideration. In the cyclic AMP-dependent processes, for example, the

The Cytoskeleton and Metabolism

17

hormone is known to interact with a specific receptor on the external surface of the cell membrane, causing the activation of the adenyl cyclase on the internal surface of the cell membrane, and the formation of cyclic AMP, which can then exert its widespread effects in the cell via the action of specific protein kineses. Again, in glycogen breakdown, cyclic AMP is formed from ATP and activates a protein kinase, which activates phosphorylase kinase, which in turn activates phosphorylase (Lehninger, 1982). It should be noted that this enzymatic cascade not only provides considerable scope for the amplification of the physiological effect, as has been well described in many texts, but also consumes a considerable energy input in the process. An amplification of a hundredfold at each step, for example, implies the involvement of ten thousand molecules of ATP in a cascade initiated by one mole of cyclic AMP (de Duve, 1988). Similarly, with the transmembrane signaling of growth factors and other agonists which utilize inositol triphosphate as a second messenger, it is noteworthy that a series of intracellular phosphorylations is involved (inositol to phosphatidyl inositol to phosphatidyl inositol-4-phosphate to phosphatidyl inositol-4:5-diphosphate and diacyglycerol to phosphatidic acid; Berridge, 1986). Hence, the action of both these second messengers in transmembrane signaling share certain common features—they represent the internal manifestation of cell surface interactions between agonists and receptors, they involve a series of phosphorylations and a degree of amplification; and hence generate an intense and localized demand for energy to ensure their continuing function and benefits. Also associated with these cell-signaling pathways, as well as those involving Ca"^"^, is protein kinase C (Nishizuka, 1984). Many physiological functions have been assigned to this system, including involvements in secretion, smooth muscle contraction, steroidogenesis, the regulation of gene expression and cell proliferation, and regulatory crosstalk between signaling pathways. It is relevant to note again, that in all these processes, there is a requirement for energy input which is often met from glycolysis, and often associated with significant proportions of protein kinase C in a form which is closely associated with the cytoskeleton (Carr and Scott, 1992). It is also of interest to note that there is an established connection between the action of growth factors and the activity characteristics of the glycolytic enzymes. Several growth factors initiate their biological effects by activating membrane receptors, as we have seen, and many of these receptors possess intrinsic tyrosine kinase activity. It is by means of this phosphorylation process that these agonists regulate particular biochemical pathways, so that it is noteworthy that several enzymes of the glycolytic pathway, including key regulatory enzymes such as phosphofructokinase and glyceraldehydephosphate dehydrogenase, may serve as substrates for receptor kinases (Reiss et al., 1986). As indicated in a previous section of this article, too, such protein phosphorylation may exert quite profound effects on the degree of association of the substrates with structure. There would appear to be a strong link then between the mechanism of action of these agonists and the

18

COLIN MASTERS

degree of association of the glycolytic enzymes with the cytoskeleton—the significance of which is developed in the next section.

VII. MATRICAL COMPARTMENTATION In the previous sections, it has been noted that not only does a typical cell show discernible heterogeneities and variabilities of cytoskeletal structure, but also that there exists a differential localization of energy requirement within these cells. In tying together these observations, it is instructive to consider the possible connection between the requirements for localized energy production for cellular function, and the provision of this energy by the binding of glycolytic enzymes to localized areas of cytoskeleton. In relation to the examples quoted, for instance, it is worth noting that the important role of glycolysis as a source of energy for cell membrane function has been cited previously by several research groups, as has the modification of the kinetics of bound glycolytic enzymes so as to provide for maximal increases in glycol3^ic flux. Daum et al. (1988), for instance, have observed an association of glycolytic enzymes with the cytoplasmic side of the plasma membrane and the adjacent portion of the cytoskeleton in glioma cells and have commented on the fact that phosphorylation of glucose is strictly coupled with its membrane transport in these cells with evident advantage for the metabolism of glucose in close vicinity to its transporter. Another apposite example is provided by consideration of the ubiquitous Na-K plasma membrane pump. This pump acts as an ATP-ase, with the active transport of glucose being driven by a sodium gradient that is generated and maintained by the Na-KATP-ase. Here also a compartmentation of cytoplasmic carbohydrate metabolism has been noted, with multiple sitings and separate functioning of glycolytic pathway components. In particular, an association of glycolytic enzymes with the plasma membrane in close juxtaposition with the membrane pump has been observed, and with the aerobic production of energy in this pathway acting quite independently of generalized glycogenolysis—being related to the specific requirements of the membrane pump rather than the general energy requirements of the cell (Lynch and Paul, 1986). In addition, a structure and function relationship has been documented between the non-uniform distribution of glycolytic systems and heterogeneous ATP supply, where glycolytically derived ATP is preferred over that from oxidative phosphorylation as the energy source for membrane function in various normal and transformed cells. In this regard, glycolytic activity has been shown to be coupled to Na and K transport in smooth muscle and erythrocytes, Ca"^"^ uptake in heart, the maintenance of membrane electrophysiological activities in cultured chick embryo and transformed hamster and Ehrlich ascites tumor cells, and K balance during neuronal activity in brain (Aw and Jones, 1988). It would appear that both mitochondrial and glycolytic systems are heterogeneously, but strategically, located

The Cytoskeleton and Metabolism

19

within cells, and under conditions of limited ATP supply this heterogeneous distribution can be an important determinant of ATP supply to different-ATP utilizing reactions. Additionally, as has been indicated previously, treatment of cells in culture with peptide hormones, or with second messengers such as cyclic AMP and inositol triphosphate, causes an increased binding of glycolytic enzymes to the cortical regions of the cytoskeleton (Chen and Masters, 1988). Here again then the intense energy requirement of these ATP-dependent regulatory cascades appears to require a separate identity and localization of the glycolytic pathway within the cellular structure, so that these important processes, and their requirements for energy supply and utilization, may be effectively coordinated. Overall, then, the available, albeit limited, evidence serves to indicate that an increasing number of cellular processes, and especially those associated with membrane function and signal transduction, appear to possess a spatially and functionally discrete association with elements of the glycolytic pathway. Rather than the commonly accepted view that the glycolytic enzymes are soluble and homogeneously distributed within the cytoplasm, the indications are that the glycolytic components often associate with particular regions of the cytoskeleton and display considerable functional heterogeneity. Indeed such a co-localization may not be merely an advantage; it may be a necessity in the cellular situation. As has been pointed out recently (Lynch and Paul, 1988), if all the energy requiring processes competed for a common pool of ATP, coordination would be rendered extremely vulnerable to any centrally located ATP-ase, which could effectively eliminate the access of other processes to ATP. Again, as has been pointed out previously in a more general context, considerable benefits in improved efficiency may accrue to the cell through this type of compartmentation, providing metabolic advantage by influencing the containment, efficiency and relative positioning of the enzyme components (Masters and Reid, 1987). Indeed these particular characteristics of the interactions between cytoskeletal structure and (glycolytic) enzymes are sufficiently novel and distinctive to merit a special nomenclature—namely, matrical compartmentation (Masters and Reid, 1987) underlying the fact that this form of compartmentation is quite distinct from the other major recognized forms of compartmentation (e.g., intercellular, organellar), yet possesses characteristics which are especially advantageous to that major metabolic sector of the cell—the cytoplasm. One advantage which may readily be perceived in terms of these enzyme-structure interactions is that contributed by the pervasive nature of the cytoskeleton. Viewed in metabolic terms, this means that the cell has a ready made scaffolding, which extends throughout the cytoplasm and which means that extreme precision is available in positioning energy providing sequences near to energy requiring sequences. Furthermore, this matrical form of compartmentation is readily reversible, and can be disassembled and reassembled as required by the cell. Matrical compartmentation also provides advantages in relation to the facilitation of the interaction between enzyme and substrate, too. If one considers the situation of a

20

COLIN MASTERS

Figure 6. Diagrammatic representation of the manner in which enzyme associations with the cytoskeleton may be utilized to achieve the positional coupling of complementary metabolic processes. A,B,C represent energy requiring membrane transport processes which may be linked with adjacent energy producing glycolytic segments (X). Similarly, within the interior of the cell, an apposite linkage may be achieved between other energy requiring processes (D, F, G) and energy producing sequences (X, Y, Z).

substrate molecule in the cell and its pathway to a target enzyme for example, an appreciable transit time may be required for the interaction of these two molecules to be achieved. With matrical compartmentation, the transit time may be reduced by 2—3 orders of magnitude, and offers major advantages in metabolic efficiency (Masters and Reid, 1987). Additionally, of course, advantages arise from the fact that the enzymes, when bound to the structural matrix, assume altered kinetic properties which often favor a maximal rate of reaction and increases possibilities of substrate channelling (Masters et al., 1987). In addition to the examples quoted previously in relation to membrane transport, there are clearly many other processes in the cell (e.g., the synthesis of macromolecules) which require energy and would benefit from a positioning of the means of energy supply liear to the site of energy demand. Even more generally, many other metabolic interconnections in the cell may be facilitated, or favored as one particular choice amongst many possible fates, by the apposite positioning of the constituent processes. Matrical compartmentation, in general, then affords unique features of flexibility in relation to assembly and disassembly, extreme precision in relation to subcellular positioning, significant reduction in transit times, and other kinetic advantages (Figure 6).

The Cytoskeleton and Metabolism

21

VIII. PERTURBATIONS OF COMPARTMENTATION DURING CELLULAR DIFFERENTIATION AND DYSFUNCTION If one accepts that the co-locaHzation of glycolysis and surface processes confers considerable metabolic advantage on cellular systems in the adult animal, it is logical to consider how this may affect carbohydrate metabolism during development and major clinical disorders. It is generally accepted that carbohydrate metabolism is of particular importance to the developing embryo, for example, with glycolysis being the prime source of energy in early embryogenesis. In addition, the cytoskeleton is known to assume different structural characteristics to that observed in adult tissues so as to be able to cope with the special developmental requirements of active cell division, morphogenesis and migration. At the same time, there is a requirement for energy for these processes, and as already observed, glycolysis is established as the main source of this energy (Masters and Holmes, 1975). It has also been well documented, that coincident with this high glycolytic rate which is characteristic of cells in the fetus, there occurs a degree of adsorption of the glycolytic enzymes to the cytoskeleton which is increased above that generally present in adult tissues (Masters, 1991; Murrell et al., 1992; Figure 7). Indeed, the developmental progressions of the glycolytic enzymes provide an important substantiation of the physiological significance of micro-compartmentation, and for its influence by cellular control mechanisms that can respond to ontogenic and physiological cues. The developmental data provides strong evidence that enzyme structure associations play a significant role in meeting the specialized energy needs of differentiating cell types during early ontogeny, jas well as offering the potential for increased flexibility and control of glycolysis in specialized physiological situations (Murrell et al., 1992; Table 2). Just as there is compelling evidence for a developmental variation in the degree of interaction between the cytoskeleton and glycolytic enzymes, so is there abundant data that the interaction of many proteins with cellular structure is a dynamic process which is responsive to and in turn influences cellular metabolic status. In addition to the well documented case of hexokinase, for example, and^the many indications of an ambiquitous behavior for this enzyme (Wilson, 1978), there is also much evidence that the extent of binding of other glycolytic enzymes varies with metabolic status. In skeletal muscle, for instance, an increased, binding of aldolase, phosphofructokinase and glyceraldehyde-phosphate dehydrogenase has been observed during contractile activity, with a reversion to normal distribution after the cessation of stimulation (Masters, 1981). Similarly ambiquitous redistributions have been demonstrated in other major areas of metabolism such as the biosynthesis of triglycerides and phosphatidyl choline (Vance and Pelech, 1984), chemotaxis (Koshland, 1981), and the wide range of functions irivolving protein kinase C (Nishizuka, 1984) and calmodulin (Hanbauer et al, 1979)—all of which

22

COLIN MASTERS

LIVER

80-1

KIDNEY

80-

N 10

H

1 M-m

-28 -14 +1 +14+21 Age (days)

80-1

0 l—i 1—MHI -28 -14 +1 +14+21 Age (days)

Adult

MUSCLE

H 9 Adult

BRAIN

80-1

^

10 -Mh -t-28 -14 +1 +14+21 Age (days)

9

-28-14 +1+14+21 Age (days)

Adult

Adult

HEART

lOOn

N •28

-4-M^

+14+21 Age (days)

Adult

Figure 7. T h e percentage of structure-bound aldolase and guinea pig tissues during pre- and post natal development (after Murrell et a l . , 1 9 9 2 ) .

The Cytoskeleton and Metabolism Table 2,

23

Interactions Between the Glycolytic Enzymes and Cytoskeleton— Indications of Physiological Significance

•

Evidence from a variety of techniques for the extensive occurrence of such binding in cells and tissues.

•

The marked alterations in the degree of interaction which occur during vertebrate growth and differentiation.

•

The changes observed as a result of physiological activities such as muscle contraction.

•

Alterations in the pattern of interactions during disease processes such as diabetes and cancer.

•

The metabolic correlations between interactions and the micro-organization of the glycolytic sequence.

adds strength to the general significance of the interactions of the interactions of the glycolytic enzymes, as discussed in this chapter. In a similar manner to the variations in the degree of enzyme structure association which have been demonstrated in the case of developmental and physiological changes, so there are also an increasing number of reports of variations during cellular abnormality and dysfunction. Tumor cells, for example, are well known as displaying atypical characteristics of both cytoskeletal structure and carbohydrate metabolism, and it is of interest in the present context to compare the relationships

Table 3, The Influence of Regulatory Metabolites on the Extent of Binding of Glycolytic Enzymes to the Cytoskeleton Metabolite

Enzyme

Influence on Binding

PFK

F-2, 6-P2 F-1, 6-P2

Decrease Increase

ALD

F-l,6-P2

Decrease

GAPDH

GA3P

Decrease

PGK

ATP NAD

Decrease Increase

PK

F-2, 6-P2 F-l,6-P2

Decrease Increase

LDH

NADH F-l,6-P2

Decrease Increase

Key: PFK, Phosphofructokinase; ALD, Aldolase; GAPDH, Glyceraldehyde-Phosphate Dehydrogenase; PGK, Phosphoglycerate Kinase; PK, Pyruvate Kinase; LDH, Lactate Dehydrogenase; FP2, Fructose Bisphosphate; GA3P, Glyceraldehyde-3 -Phosphate.

24

COLIN MASTERS

between these abnormalities. Normal cells in tissue culture are known to become stationary, and develop a well ordered pattern of stress fibers in the cytoskeleton. By contrast, most malignant cells remain motile, and exhibit much less in the way of stress fiber development. The tumor situation has been likened to de-differentiation by many workers, and certainly bears the joint characteristics of alterations in both cytoskeletal structure and the increased dependence on glycolysis as an energy source referred to in previous examples. There appears to be a close correlation between the organization of the cytoskeleton in tumors and the intensity of the Warburg effect (Bereiter-Hahn, 1988), and this proposal is supported by evidence of an increased association between the glycolytic enzymes and the cytoskeleton in tumor cells (Masters 1992), facilitating an increased supply of ATP from glycolysis in this situation. There is also an indication of the mechanism of this process, since transforming viruses appear to possess the ability to phosphorylate the glycolytic enzymes and so influence the degree of association of these enzymes with cellular structure (Reiss et al., 1986). The close interaction of the glycolytic pathway with filamentous actin may also affect calcium extrusion by the plasma membrane. There are, of course, persistently high cytosolic calcium levels in transformed cells, and in Ehrlich ascites tumor cells, it has been established that the transport of calcium through the membrane uses glycolysis as its source of an energy (ATP) supply. As has been indicated earlier, there is also data showing that hormones can cause perturbations of the cytoskeletal association between glycolytic enzymes and the cytoskeleton. Insulin, adrenalin, and glucagon cause a modest increase in the binding of the glycolytic enzymes, but adequate to cope with the energy requirements of conformational alteration of receptor proteins, while second messengers with their greater initial requirement for energy, cause more substantial degrees of cortical association (Chen and Masters, 1988; Masters, 1992; Table 3). In the case of diabetes, there is increasing evidence that an antithetical situation applies; coincident with decreased insulin secretion and the reduction in hormone-receptor interactions, there is a reduction in the degree of binding of glycolytic enzymes to the cortical regions of the cytoskeleton (Masters, 1992). This contributes both to a reduction in the cellular transport processes for sugars and amino acids, and as a consequence in glycolysis. Thus micro-compartmentation should be considered as a new and important factor in this condition, and other disorders of carbohydrate metabolism. The alterations to cytoskeletal structure and associated redistributions of localization of the glycolytic enzymes within the cytoplasm present a novel, characteristic aspect of disorders of carbohydrate metabolism; and these associations and the implications of the attendant shifts in matrical compartmentation would appear to deserve far more consideration and research attention in relation to the molecular definition of these important disease processes.

The Cytoskeleton and Metabolism

25

IX. COMPARTMENTATION IN INTERMEDIARY METABOLISM It is important to recognize that the concepts of micro-compartmentation need not be restricted to carbohydrate metabolism. As is well established, many of the cell's metabolic pathways and indeed most of its intermediary metabolism take place in the cytoplasm (Lehninger, 1982), and the advantages of propitious positioning of energy requiring processes which have been observed in the case of glycolysis may apply with equal relevance in these broader metabolic areas as well. It would be both intriguing and illuminating for cell biologists to determine whether the cytoskeleton not only formed a general framework for the modulation of carbohydrate metabolism, but also acted in a broader perspective as a self-regulatory skeleton which integrated many of the specific energy-requiring and -developing processes of intermediary metabolism. In relation to the positioning of these energy related processes, for example, it is well established experimentally that substantial ATP gradients exist in cells under ATP-limiting conditions (Aw and Jones, 1988), and many examples of the potential advantages of coupling between energy-driving and -utilizing reactions may be cited. In relation to carbohydrate metabolism, evidence has already been listed which indicated a conjunction of glycolysis with membrane transport processes, and the accompanying metabolic rationale (Lynch and Paul, 1986; Wuytack et al., 1985). In recognition of the variety of these transport processes (Na-, K-, Ca-, glucosetransport, for example), and our previous comment on multiplicity, it is relevant to point out that this situation may well be an example of a number of different positional assemblages of glycolytic components, each paired with separate transport functions. The established positional separation of glycolysis from glycogenolysis has also been cited, and a simple example of positional advantage has been provided by the readily recognizable occurrence of glycogen granules in the cytoplasm, and the fact that the enzymes which are needed to carry out the synthesis and degradation of this storage form of carbohydrate are bound to the surface of these glycogen particles (Alberts et al., 1983). Furthermore there is an evident advantage accruing in an appropriate positional coupling of aerobic glycolysis to glycogen breakdown, in the linkage of LDH with glycolysis under anaerobic conditions, in the usage of glycolytic segments to provide localized sources of energy for cell motility, and in the appropriate positioning of glycolysis in relation to other interacting pathways such as the metabolism of pentose phosphates, amino acids and lipids. Clearly in all these cases there is considerable potential advantage in matrical compartmentation. Among the hundreds of other energy requiring reactions in cellular metabolism, there is also ample scope for the coordination of metabolic functions by clustering of sequential activities within individual pathways, and the synergistic linking of appropriate pathways with consequent recognition of the separate and distinctive requirements of these elements. Indeed as has been pointed out previously (Masters

26

COLIN MASTERS

et al, 1969), the structural elements of the cell may be considered to act in the same manner as a potential allosteric effector. Just as the classical (small molecule) effectors act by modifying the kinetic properties of enzymes, so binding of an enzyme to the structural elements of the cell may influence its kinetic properties significantly, in a similar manner; and in this manner act to modulate intermediary metabolism. Consideration of the metabolic maze in the cytoplasm, where one compound may have more than a dozen metabolic fates, points to the potential scope and advantages of positional associations and their inherent ability to remove such linkages of metabolism from the tyrannies of random walks and undue allosteric restrictions on flux. Moreover, there are many particulate analogies which point to the advantages of such associations. Just as the respiratory chain contains a number of large enzyme complexes, which are embedded in mitochondrial membranes with defined orientation and so act to coordinate the efficiency of respiration with energy production (Lehninger, 1982), so the cytoskeleton may well provide the positional framework which allows an efficient coupling of metabolic processes in the cytoplasm. Again, in transmembrane signaling, it is well recognized that receptor and adenylate cyclase molecules are separate proteins that interact functionally in the plasma membrane. In just such a manner, separate processes may be linked in the cytoplasm by a utilization of the cytoskeletal framework. Just as in the organellar portion of the cell, there is a heavy dependence on particulate assemblages for the purposes of coupling, so the cytoplasm should also be expected to make beneficial use of any available opportunities for similar structural advantage.

X. CONCLUDING COMMENTS An important message emerges from these considerations on the role of the cytoskeleton and enzyme-structure interactions in intermediary metabolism. Given that there is now compelling evidence for native interactions between the enzymes of glycolysis and the cytoskeleton, and extensive indications of a considerable involvement of these interactions in many biological processes, it must be concluded that biphasic interactions between the enzymes of the cytoplasm and the cytoskeleton are deserving of wider recognition in future investigations of the balance and inter-relationship of cellular metabolism in higher organisms. As has been pointed out in the previous discussion, many of the conditions commonly employed in enzymological investigations of intermediary metabolism and its control are biologically abnormal and bear little relation to the conditions under which enzymes act in vivo. While analytical accuracy may be improved by the use of purified components in dilute aqueous solution, for example, such approaches often proceed at the cost of physiological credibility. The realities of the cellular microenvironment require that account be taken as well of factors such as the pervasivjg influence of the cytoskeleton within the cytoplasm, the level of concentration of enzymes and metabolites in tissues and the associated molecular

The Cytoskeleton and Metabolism

27

crowding, interactions between enzymes and structure and the kinetic differences induced thereby, signaHng by median metaboHtes, and matrical compartmentation. Certainly, as discussed in this article, there are many indications that the biphasic interactions between the cytoskeleton and the enzymes of the cytoplasm form a major element of metabolic regulation, and offer a unique potential for control by the appropriate coupling of reactions and the facilitation of cross-talk between signaling systems. It now seems evident that the cytoskeleton should be viewed not merely as filling a passive, structural role, but also as constituting a dynamic component of the cell, which forms a major intracellular communication system and holds considerable significance in metabolism. The cytoskeletal framework has ^parently evolved over time as an essential agent in the control of cytoplasmic processes in higher organisms, and offers unique advantages in relation to the advantageous positioning of the reactions of intermediary metabolism in relation to one another and to cellular structure. Again, just as the recognition and understanding of enzyme multiplicity has opened new vistas in the comprehension of many aspects of cell biology (Masters and Holmes, 1975), so isofiinctional studies of micro-compartmentation would appear to have much to offer towards an increasingly satisfying description of the cellular processes. As has been established, the cytoskeleton is capable of being modified in many ways and by several agents. Hormones, growth factors, second messengers, actin binding proteins, metabolic effectors, and various combinations of enzymes and substrates have all been shown to exert individual influences on cytoskeletal structure, and provide substantial scope for regional modifications and micro-heterogeneities within the cytoskeletal framework which have the capability of designating particular areas for specific groups of activities. Many of these modulations of cytoskeletal structure have also been shown to be implicated in wide ranging and important biological processes such as growth and differentiation, adaptations to physiological stress and disease, and cellular transport processes. Surely, then, an improved definition of the nature and variation of cytoskeletal heterogeneity, and the contribution of the different types of structure and their interactions with cytoplasmic enzymes to cellular metabolism, provides the promise of a richly rewarding area of biological investigation. Obviously, new techniques which differ fi'om those traditionally used in metabolic studies, will feature in the major research trends of this area. In order to make appropriate allowance for the influence of the cytoskeleton on metabolic processes, a greater emphasis will need to be placed on techniques which allow the measurement of metabolic flux without drastic perturbations or removal of cytoskeletal structure. Membrane permeabilization has been mentioned extensively in this review as one such possibility, and undoubtedly many variations on this theme will be developed in coming years. There will also be a need to improve procedures for visualizing and identifying changes in the skeletal structure and the spatial localization of enzyme-cytoskeletal interactions, which occur concurrently with physiological modulations. To give an example, procedures such as the detection of

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imaging fluorescent molecules in cells during their performance of biological functions, may be of value in this regard (Backsai et al., 1993). In any case, and by whatever ingenuity of technique, cellular biologists would be well advised to consider moving their investigations of metabolic control in the cytoplasm away from structureless uniphasic simulations, and toward a more complete definition of the role of biphasic cytoskeletal interactions in modifying the emphasis of metabolic processes. REFERENCES Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K., & Watson, J.D. (1983). Molecular Biology of the Cell Garland Publishing Inc., New York. Aw, T.Y., & Jones, D.R (1988). In: Microcompartmentation (Jones, D.R, Ed.), CRC Press, Boca Raton, FL, pp. 191-207. Backsai, B.J., Hochner, B., Mahautsmith, M., Adams, S.R., Kaang, B.K., Kandel, E.R., & Tsien, R.Y. (1993). Spatially resolved dynamics of cAMP and protein kinase A subunits in aplysia sensory neurons. Science 260, 222-226. Bereiter-Hahn, J. (1988). In: Microcompartmentation (Jones, D.R, Ed.), CRC Press, Boca Raton, FL, pp. 55-69. Berridge, M.J. (1986). Intracellular signalling through inositol trisphosphate and diacylglycerol. Biological Chemistry, Hoppe-Seyler, 367, 447-456. Carr, D.W., & Scott, J.D. (1992). Blotting and band-shifting: Techniques for studying protein: Protein interactions. Trends Biochem. Sci. 17, 246-249. Chen, N., Nancarrow, D., & Masters, C.J. (1986). The influence of fructose-l:6-bisphosphate on the release of glycolytic enzymes and cellular structure, Biochem. Int. 13, 539—547, Chen, N., & Masters, C. (1988). The influence of insulin and glycogen on the interactions between glycolytic enzymes and cellular structure. Biochem Int. 16(5), 903—912. Clegg, J.S. (1984). Properties and metabolism of the aqueous cytoplasm and its boundaries. Am. J, Physiol. 246, R133-R151, Cohen, P. (1980). Recently Discovered Systems Of Enzyme Regulation by Reversible Phosphorylation, Elsevier/North Holland Biochemical Press, Amsterdam. Daum, G., Keller, K., & Lange, K. (1988). Association of glycolytic enzymes with the cytoplasmic side of the plasma membrane of glioma cells. Biochem. Biophys. Acta 939, 277—281, de Duve, C. (1988), The Living Cell Scientific American Books Inc., New York. Engstrom, L. (1978). The regulation of liver pyruvate kinase by phosphorylation-dephosphorylation. Curr. Topics Cell Reg. 13,29-51, Foe, L.G., & Kemp, R.G, (1982). Properties of phospho- and dephospho- forms of muscle phosphofructokinase. J. Biol, Chem, 257, 6368-6372, Han, J-W., Thieleczek, R., Varsanyi, M., & Heilmeyer, L.M.G. (1992). Compartmentalized ATP synthesis in skeletal muscle triads. Biochemistry 31, 377-384. Hanbauer, I., Gimble, J., & Lovenberg, W. (1979). Changes in soluble calmodulin following activation of dopamine receptors in rat striatal slices. Neuropharmacology 18, 851-857. Heuser, J,E,, & Kirschner, M,W. (1980), Filament organization revealed in platinum replicas of freeze-dried cytoskeletons. J. Cell Biol. 86,212-234. Humphreys, L., & Masters, C. (1986). On the differential release of glycolytic enzymes from cellular structure. Biochem. Int. 13, 71-77. Humphreys, L., Reid, S., & Masters, C. (1986). Evidence for the spatial separation of the binding sites for substrate and for cj^oskeletal proteins on the enzyme aldolase. Int. J. Biochem. 18, 7—13.

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Keleti, T., & Ovadi, J. (1988). Control of metabolism by dynamic macromolecular interactions. Curr. Topics Cell Reg. 29, 133. Koshland, D.E. Jr. (1981). Biochemistry of sensing and adaptation in a simple bacterial system. Annu. Rev. Biochem. 50, 765-782. Knull, H.R., Taylor, W.F., & Wells, W.W. (1974). Insulin effects on brain energy metabolism and the related hexokinase distribution. J. Biol. Chem. 249, 6930-6935. Kumagai, H., & Sakai, H. (1983). A porcine brain protein (35K protein) which bundles microtubules and its identification as glyceraldehyde 3-phosphate dehydrogenase. J. Biochem. 93,1259-1269. Kuo, H., Malencik, D., Liou, R., & Anderson, S.R. (1986). Factors affecting the activation of rabbit muscle phosphofhictokinase by actin. Biochemistry 25, 1278-1286. Kurganov, B.I., Sugrobova, N.R, & Millman, L.S. (1985). Supramolecular organization of glycolytic enzymes. J. Theor. Biol. 116, 509-516. Lehninger, A. (1982). Principles of Biochemistry. Worth Publishers Inc., New York. Li, S.S., Pan, Y.E., Sharief, F.S., Evans, M.J., Lin, M.F., Clinton, G.M., & Holbrook, J.J. (1988). Cancer-associated lactate dehydrogenase is a tyrosylphosphorylated form of human LDH-A, skeletal muscle isoenzyme. Cancer Investigation 6, 93-101. Lynch, R.M., & Paul, R.J. (1986). Compartmentation of carbohydrate metabolism in vascular smooth muscle: evidence for at least two functionally independent pools of glucose 6-phosphate. Biochim. Biophys. Acta 887, 315-318. Mackall, J., Meredith, M., & Lane, M.D. (1979). A mild procedure for the rapid release of cytoplasmic enzymes from cultured animal cells. Anal. Biochem. 95, 270-274. Maretzki, D., Reimann, B., & Rapoport, S.M. (1989). A reappraisal of the binding of cytosolic enzymes to erythrocyte membranes. Trends Biochem. Sciences 14,93—96. Masters, C.J. (1975). Isozyme realization and ontogeny. In: Isozymes, Vol. 3 (Markert, C.L., Ed.). Academic Press, New York, pp. 281-296. Masters, C. J. (1981). Interactions between soluble enzymes and subcellular structure. CRC Critical Rev. in Biochem. 11, 105-143. Masters, C.J. (1984). Interactions between glycolytic enzymes and components of the cytomatrix. J. Cell Biol. 99, 229S^225S. Masters, C. (1991). Cellular differentiation and the microcompartmentation of glycolysis. Mech. Aging Devel. 61, 1991, 11-22. Masters, C.J. (1992). Microenvironmental factors and the binding of clycolytic enzymes to contractile filaments. Int. J. Biochem. 24, 405-410. Masters, C.J., & Holmes, R.S. (1975). Haemoglobin, Isoenzymes and Tissue Differentiation. North Holland Publishing Company, Amsterdam. Masters, C.J., & Reid, S. (1987). Isozymes and the micro-organization of the glycolytic sequence. Isozymes Curr. Top. Biol. Res. 14, 1987, 45-58. Masters, C.J., Reid, S., & Don, M. (1987). Glycolysis—New concepts in an old pathway. Mol. Cell. Biochem. 76, 3-14 Masters, C.J., Sheedy, R.J., & Winzor, D.J. (1969). Reversible absorption of enzymes as a possible allosteric control mechanism. Biochem. J. 112, 806-808. Morton, D.J., Clarke, F.M., & Masters, C.J. (1976). An electron microscope study of the interaction between fructose diphosphate aldolase and actin-containing filaments. J. Cell Biol. 74,1016-1023. Murrell, W, Crane, D., & Masters, C. (1992). Ontogenic characteristics of cavian aldolase. Mech. Aging Devel. 65, 35-50. Nishizuka, Y. (1984). The role of protein kinase C in cell surface signal transduction and tumor promotion. Nature 308, 693-698. Pardo, J.v., Pittenger, M.F., & Craig, S.W. (1983). Subcellular sorting of isoactins: Selective association of gamma actin with skeletal muscle mitochondria. Cell 32, 1093-1103. Pollard, T.D., Selden, S.C, & Maupin, P. (1984). Interaction of actin filaments with microtubules. J. Cell Biol. 99(1 Pt 2), 33^37s.

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Porter, K.R., Beckerle, M., & McNiven, A. (1983). The cytoplasmic matrix. Mod. Cell Biol. 2,259-302. Proverbio, R, & Hoffman, J.F. (1977). Membrane compartmentalized ATP and its preferential use by the Na,K-ATPase of human red cell ghosts. J. Gen. Physiol. 69, 605-632. Reid, S., & Masters, C. (1986). On the ontogeny and interactions of glyceraldehyde-3-phosphate dehydrogenase. Mech. Aging Devel. 35, 209-219. Reiss, N., Kanety, H., & Schlessinger, J. (1986). Five enzymes of the glycolytic pathway serve as substrates for purified epidermal-growth-factor-receptor kinase. Biochem. J. 239, 691-697. Shearwin, K., Chen, N., & Masters, C.J. (1990a). Interactions between glycolytic enzymes and cytoskeletal structure—^The influence of ionic strength and molecular crowding. Biochem. Int. 21,53-60. Shearwin, K., Chen, N., & Masters, C.J. (1990b). The binding of glycolytic enzymes to the cytoskeleton—Influence of pH. Biochem. Int. 22, 735-740. Srere, P.A., & Ovadi, J. (1990). Enzyme-enzyme interactions and their metabolic role. FEBS Letters 268,360-364. Storey, K.B. (1985). A reevaluation of the Pasteur effect: New mechanisms in anaerobic metabolism. Mol. Physiol. 8,439-461. Sukhodolets, M.V., Muronetz, V.I., Tsuprun, V.L., Kaftanova, A.S., & Nagradova, N.K. (1988). Association of rabbit muscle glyceraldehyde-3-phosphate dehydrogenase and 3-phosphoglycerate kinase: the biochemical and electron-microscope evidence. FEBS Letters 238, 161—166. Vance, D.E., & Pelech, S.L. (1984). Enzyme translocation in the regulation of phosphatidylcholine biosynthesis. Trends in Biochemical Sciences 9,17-20. Walsh, T.R, Masters, C.J., Morton, D.J., & Clarke, F.M. (1981). The reversible binding of glycolytic enzymes in ovine skeletal muscle in response to tetanic stimulation. Biochem. Biophysics. Acta 675,29-39. Wilson, J.E. (1978). Ambiquitous enzymes: Variation in intracellular distribution as a regulatory mechanism. Trends Biochem. Sci. 3, 124—125. Wuytack, R, Raeymaekers, L., & Casteels, R. (1985). The Ca^^-transport ATPases in smooth muscle. Experentia 41, 900-909.

ASSOCIATION OF mRNAS AND POLYRIBOSOMES WITH THE CYTOSKELETON: POTENTIAL ROLES IN TRANSPORT AND COMPARTMENTALIZATION OF mRNAS

John E. Hesketh and Ian F. Pryme

I. Introduction 32 II. Evidence that the Cytoskeleton Participates in Protein Synthesis 32 III. Which Cytoskeletal Components are Associated with the Protein Synthetic Apparatus? 35 IV. Association of Polysomes with the Cytoskeleton is Affected by Physiological Conditions 38 V. What is the Function of Cytoskeletal-Bound Polysomes? 39 VI. The Cytoskeleton and mRNA Localization 42 VII. Nature of the Interaction between Cytoskeleton and mRNAs and Polysomes . .46 A. Role of the 3'Untranslated Region of the mRNA 46 B. mRNA/Ribosome Binding Proteins 47 VIII. Future Perspectives 52 Acknowledgments 52 References 52 The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 31-58 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 31

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I. INTRODUCTION Normal cell growth and activity requires that not only are the correct proteins synthesized in the correct amounts, but that they are also delivered to their site of function. Thus, protein targeting is crucial for cell organization. There are approximately 15,000 mRNA species in a mammalian cell, of which maybe 3,000 are targeted to the endoplasmic reticulum for synthesis on a distinct class of membranebound polyribosomes (polysomes). It is a major question of cell biology whether the remaining mRNAs are all translated in one compartment of cytosolic, "free" polysomes or if there is some spatial organization of protein synthesis. Until recently, it was assumed that proteins which were not synthesized on the endoplasmic reticulum were synthesized in a spatially unorganized fashion on "free" polysomes. However, the recent observations that polysomes are found associated with the cytoskeleton have suggested that the situation may be more complicated because interactions between the cytoskeleton and protein synthetic apparatus have the potential to provide a mechanism for the transport and compartmentalization of mRNAs and polysomes. Thus, over the past seventeen years there has been increasing interest in investigation of the association of mRNAs and ribosomes with the cytoskeleton (for reviews see Nielsen et al., 1983; Hesketh and Pryme, 1991; Kirkeeide et al., 1993; Suprenant, 1993; Hesketh, 1994; Pryme et al., 1995).

IL EVIDENCE THAT THE CYTOSKELETON PARTICIPATES IN PROTEIN SYNTHESIS The development of high voltage electron microscopy (HVEM) techniques made it possible for Wolosewick and Porter (1976,1979) to make a detailed study of the cytoplasmic ground substance of eukaryotic cells. From their work it became apparent that in addition to being associated with endoplasmic reticulum (ER) membranes, ribosomes were also detected in close proximity to fine filamentous elements now recognized as components of the cytoskeleton. Aggregates of ribosomes, judged to be polysomal structures, were also detected in pockets at junctions where filaments were arranged in a cross-like network. These electron microscopical observations provided thefirstevidence for the existence of a class of polysomes found in association with cytoskeletal filaments. Based on such morphological characteristics these have been termed cytoskeletal-bound polysomes (CBP). About the same time as Wolosewick and Porter published their observations on whole animal cells, Osbom and Weber (1977) demonstrated that treatment of cultured cells with low concentrations of non-ionic detergent resulted in solubilization of the plasma membrane causing a release of integral membrane proteins and cytoplasmic constituents and leaving the cytoskeleton as an insoluble matrix (referred to in this review as the cytomatrix). Using electron microscopy Lenk et al. (1977) observed that ribosomes were present in the cytomatrix and the results

Association of mRNAs and Polyribosomes

33

were taken as evidence that these ribosomes were associated with the cytoskeleton. However, the presence of remnants of ER have been demonstrated in the cytomatrix (Dang et al., 1983; Mirande et al., 1985; Ramaekers et al., 1983), indicating that this non-ionic detergent residue would also contain ribosomes associated with the rough ER (RER). There are indeed a series of articles (e.g., Birckbichler and Pryme, 1973; Pryme et al., 1973; Pryme, 1974; Pryme and Svardal 1978; Svardal et al., 1981) where it has been shown that an ionic detergent (sodium deoxycholate) is required in order to completely solubilize the RER and release membrane-bound polysomes (MBP). One can conclude, therefore, that the cytomatrix, prepared as an insoluble residue following non-ionic detergent treatment of cells will contain both CBP and MBP Following these initial observations which indicated polysome-cytoskeleton association, later experiments have used both morphological and biochemical techniques, either alone or in combination, to investigate the interaction of mRNA and polysomes with the cytomatrix. For example, Heuser and Kirschner (1980) in their studies using quick-freeze deep-etch techniques in combination with electron microscopy found clusters of ribosomes which were apparently attached to filaments in the cytomatrix. Further work using electron microscopy showed that ribosomes were closely associated with cytoskeletal filaments in both lens cells (Ramaekers et al., 1983) and ascidian eggs (Moon et al, 1983) respectively. Based on size measurements Ramaekers et al. (1983) were able to tentatively identify the filaments as microfilaments. Toh et al. (1980), in their studies using auto-immune serum obtained from lupus erythromatosus patients which contains large amounts of anti-ribosomal antibodies, were able to identify by immunocytochemistry an aligned punctate labeling in the cytomatrix, which indicated an association of ribosomes with filamentous structures. Hesketh et al. (1991a) raised antibodies against ribosomal subunit proteins and used these to stain non-ionic detergent treated 3T3 fibroblasts; the results showed a pattern of linear structural arrays either beaded or punctate in nature, which was suggestive of an association between ribosomes and filamentous structures of the cytoskeleton. Home and Hesketh, (1990a, 1990b), again using anti-ribosomal subunit antibodies, obtained similar data from experiments on skeletal muscle; they showed an organization of ribosomes in a systematic, repetitive manner along myofibrils (see below). Other workers have used antibodies against other components of the translatory system; thus Zumbe et al. (1982) who used specific antibodies which recognize the 5'mRNA cap binding protein, and Heuijerjans et al. (1989) using antibodies against initiation factor eIF-2, were able to label filamentous structures in the cytomatrix. Using indirect immunofluorescent microscopy, Shestakova et al. (1991) have utilized antibodies against elongation factor EF-2 to study the location of the factor in mouse embryo fibroblasts. In a double-staining technique actin filaments were visualized with rhodamine-phalloidin, and the results showed that cables which were positively stained for EF-2 were identified as microfilaments. The interesting

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JOHN E. HESKETH and IAN F. PRYME

observation was made, however, that not all actin-containing filaments were positive for EF-2, suggesting that not all microfilaments are associated with ribosomes that are actively involved in protein synthesis. Other studies utilizing an antibody against a nuclear envelope antigen which binds to mRNA, produced a staining pattern indicative of an association with microfilaments (Schroder et al., 1988). Detailed studies on mRNA-cytoskeletal interaction were made possible by the introduction of the combination of electron microscopy with in situ hybridization; this allowed the simultaneous study of individual components of the cytoskeletal filament system and the location of specific mRNA species (see Bassell, 1992). Using this approach Singer et al. (1989) were able to provide important evidence that the mRNAs coding for actin, tubulin, and vimentin were clustered around filamentous elements present in the cytomatrix. The above-mentioned electron microscopy, immunohistochemistry, and in situ hybridization studies have thus provided a strong body of evidence indicating that components of the protein synthetic apparatus in cells are distributed in a manner suggesting an association with cytoplasmic filamentous structures. Polysomes appear to be bound to the cytoskeleton through mRNA and not ribosomes since a removal of ribosomes and initiation factors from the cytomatrix has been demonstrated after heat shock (Welch and Feramisco, 1985) or incubation of cells with fluoride or pactamycin (Howe and Hershey, 1984). The possibility that components of the translatory apparatus are not merely trapped within the cytomatrix as a result of an artefact occurring during the perturbation of cells has been ruled out by several studies. Adams et al. (1983) investigated whether or not non-specific polysome-protein interactions occurred in the presence of Triton XI00 utilized during cytomatrix preparation. Using gel filtration they found no evidence which would suggest that such occurs. The possibility that an artefactual trapping of mRNA in the cytomatrix took place during preparation of cytomatrix from Xenopus eggs was tested by Pondel and King (1988). No evidence which suggested a trapping of either endogenous mitochondrial mRNAs or exogenous globin mRNA was found. These results corroborated well with the earlier results of Jeffrey (1984), who, using in situ hybridization studies with ascidian eggs, showed that the overall distribution of either total or specific mRNAs in the cell was not affected by non-ionic detergent treatment. Other evidence suggesting that it is unlikely that ribosomes are retained in the cytomatrix by some form of non-specific trapping is the fact that more than 70 percent of cell protein and virtually 99 percent of lactic dehydrogenase activity is released by non-ionic detergent treatment (Vedeler et al., 1991a). The fact that identical amounts of (free) polysomes were released from identical portions of cells either by treatment with non-ionic detergent or by disrupting cells using nitrogen cavitation (Pryme, 1974) also suggests that the observed presence of polysomes and associated factors in the cytomatrix reflects the situation in the cell.

Association of mRNAs and Polyribosomes

35

III. WHICH CYTOSKELETAL COMPONENTS ARE ASSOCIATED WITH THE PROTEIN SYNTHETIC APPARATUS? It has been observed that although polysomes can be extracted by salt from the cytomatrix of ascites cells vimentin is not affected in the same manner (Traub and Nelson, 1982; Vedeler et al, 1991a). Furthermore, eIF2 has been shown to be associated with filaments in extracts prepared from three different cell lines all lacking vimentin (Heuijerjans et al, 1989). In addition in these three cell lines both polysomes and specific mRNAs for actin and histone H2 are distributed between soluble and non-ionic detergent insoluble fractions as in normal vimentin-containing HeLa cells. It seems from these three lines of evidence that cytoskeletal-bound polysomes are not associated with intermediate filaments (IF). However there is evidence that nucleic acids can associate with IF (see Traub, Volume 1 of this treatise) and the Vgl mRNA was found associated with a cytokeratin-enriched fraction in Xenopus oocytes (Pondel and King, 1988). It is possible that it is mRNAs which are not being translated which are associated with IF (see Figure 1) and support for such a view has come from the observations that ribonucleoprotein particles called prosomes are found to be co-localized with the IF network (Grossi deSaetal., 1988). Early results obtained by Lenk et al. (1977) indicated that CBP are not associated with microtubules since colchicine had no effect on polysome distribution in cultured cells. The fact that colchicine, which causes microtubule disruption, can also promote collapse of the intermediate filament system (Hynes and Destree, 1978) would suggest that the two filament systems are linked with one another. Thus the finding that colchicine does not release polysomes from the cytomatrix (Vedeler et al., 1991a) would indicate that there is no major interaction between CBP and either the microtubule or the intermediate filament systems. It must be taken into account, however, that the majority of cellular microtubules are very susceptible to depolymerization at low temperature so that cell fractionation on ice would inevitably lead to a major disruption of microtubules and possible release of polysomes or mRNAs associated with them. Vedeler et al. (1991a) have in fact shown that the bulk of tubulin in Krebs II ascites cells is recovered in the non-ionic detergent soluble fraction. One should not rule out the possibility, therefore, that microtubules may play a more important role in protein synthesis than is at present understood. Indeed, there is evidence in vitro studies to suggest that ribosomes can associate with microtubules (Suprenant,1993). In ooc3^es there is evidence that mRNA transport depends upon microtubules (Raff et al., 1990; Yisraeli et al., 1990; Pokrywka and Stephenson, 1991) and in insect nutritive tubes, where there is extensive ribosome transport, electron microscopy has provided evidence to suggest that ribosomes are associated with microtubules (MacGregor and Stebbings, 1970). It may be, therefore, that microtubules are involved in mRNA, ribosome and perhaps polysome transport (see Figure 1).

36

Nucleus

JOHN E. HESKETH and IAN F. PRYME

U - ^

mRNAs

•

mRNAs sorted according to site of translation

Signal sequence direcU polysome complexes to ER

Untranslated mRNAs sequested in RNPs (intermediate filaments ?)

(^

C^

y ^^^^

CytOSkeletal-bound polysomes

A' s' ^ , ^ ^

membrane-bound polysomes

ER

Figure 1. Overview of the compartmentatlon of the protein synthesis illustrating possible roles of the cytoskeleton.

Recent studies of both oocytes and neurones have provided further evidence for an interaction between mRNAs, ribosomes, and microtubules: in Xenopus, microtubule preparations from oocytes contain ribosomes and those from embryos, where translation is more active, contain polysomes (Hamill et al., 1995); in Drosophila, a fusion protein containing the microtubule motor kinesin is localized in oocytes, as is the protein staufen which binds to localized mRNAs, and the localization of these proteins is destroyed by colchicine or mutations which cause mRNA mislocalization (Ferrandon et al., 1994; Clark et al., 1994); in neurones, in situ hybridization has shown 55% of mRNAs to be found in close apposition to microtubules. Overall there is increasing evidence that microtubules have a function in transporting and localizing mRNAs and other components of the translational complex. Both cytochalasin B and D, which cause a perturbation of microfilaments (Cooper, 1987), have been shown to cause a release of polysomes or mRNAs from the insoluble cytomatrix in a variety of cell lines (Lenk et al., 1977; Ramaekers et al, 1983; Bird and Sells, 1986; Omelles et al., 1986; Bagchi et al., 1987; Vedeler et al., 1991a,b). Seely and Aggeler (1991), studying the modulation of milk protein synthesis in mammary epithelial cells, showed that treatment of cells with cytochalasin D resulted in the reduced synthesis of most proteins, P-casein being first affected following disruption of the microfilament system. Using HeLa cells Omelles et al. (1986) showed that cytochalasin D, which caused a simultaneous release of both mRNA and polysomes, resulted in a reversible inhibition of protein

Association of mRNAs and Polyribosomes

37

synthesis. In myoblasts cytochalasin D causes the release of nascent myosin heavy chains (Isaacs and Fulton, 1987). DNAse I, which also causes actin depolymerization, has been shown to induce the release of polysomes (Adams et al., 1983). Vedeler et al. (1991a) demonstrated that phalloidin treatment, which stabilizes microfilaments (Dancker et al., 1975), resulted in a 30% increase in the amount of actin present in the cytoskeletal fraction of Krebs II ascites cells, only a small increase in CBP, however, was observed (12%). These results could be explained by the repolymerization of actin at the ends of filaments devoid of polysomes and are compatible with immunohistochemistry data suggesting that initiation factors are associated with some, but not all, microfilaments (Shestakova et al., 1993a). In experiments where 3T3 fibroblasts were extracted with non-ionic detergent in buffer containing either 25 or 130 mM KCl it was shown that the presence of the higher salt concentration led to an increased release of both polysomes (+50%) and actin (+76%)) from the cytomatrix to the non-ionic detergent soluble fraction (Hesketh and Pryme, 1988). In cytochalasin B treated cells the amount of actin released by extraction with detergent at a low salt concentration was equivalent to that measured following extraction with 130 mM KCl, indicating that extraction at the latter salt concentration caused a breakdown of actin filaments similar in degree to that produced by the microfilament perturbing agent (Hesketh and Pryme, 1988). These results again favored the suggestion that polysomes are associated with microfilaments. Earlier results had clearly demonstrated that salt concentrations of 100-150 mM destabilize actin filaments (Kasai, 1969) but not intermediate filaments (Steinert et al., 1982). Further supporting evidence for an association between polysomes and microfilaments has come from studies following cytomegalovirus infection (Jones and Kilpatrick, 1987) where a rapid depolymerization of microfilaments was observed early after virus infection followed later by a repolymerization. These changes were accompanied by first a decrease and then an increase in the proportion of polysomes associated with the cytomatrix. A direct interaction between components of the translatory apparatus and microfilaments is supported by the observation that EF-1 a possesses actin-binding properties (Yang et al., 1990). Considerable evidence has therefore accumulated suggesting that a population of polysomes is associated with microfilaments. Esfimates of the percentage of total cellular polysomes in the cytoskeletal-bound population varies between 25 and 40 percent in different cell lines (Ramaekers et al, 1983; Hesketh and Pryme, 1988; Vedeler et al., 1991a). When NIH 3T3 cells grown in monolayer were treated with non-ionic detergent in situ in order to avoid an initial perturbation of the cytoskeleton then 50 percent of the total polyribosome population was not extracted by 1.0 M KCl, suggesting a strong form of interaction between polysomes and structures in the cytomatrix (Biegel and Pachter, 1992). Shestakova et al. (1993a) have recently shown that when protein synthesis was inhibited either by the inactivation of eEF-2 with diphtheria toxin or by inactivating ribosomes with ricin the distribution of eEF-2 along the microfilament bundles was

38

JOHN E. HESKETH and IAN F. PRYME

not abolished. Furthermore, the disassembly of actin microfilaments by treatment of cells with cytochalasin B resulted in the disappearance of eEF-2 -bearing filaments, indicating that the organization of eEF-2 along a filament system is dependent on the integrity of the actin cytoskeleton. In further studies using indirect immunofluorescent microscopy it has been demonstrated that transition of cells from the proliferating phase into the G^ phase of the cell cycle leads to the distribution of eEF-2 mainly along intermediate filaments and/or microtubules, although a portion of eEF-2 was also co-localized on microfilaments (Shestakova et al., 1993b). Progression of the cells, however, from G^ into the proliferative phase, was shown to be accompanied by a rearrangement of the actin cytoskeleton, and a simultaneous reversal to the original pattern of distribution of eEF-2. These results suggest that in eukaryotic cells all three types of cytoskeletal filaments may well be involved in the organization of the protein-synthesizing machinery. In summary, at present the majority of evidence indicates that the major interaction of mRNAs and polysomes is with the actin-containing microfilaments but there is also increasing evidence for some interactions with the other cytoskeletal components.

IV. ASSOCIATION OF POLYSOMES WITH THE CYTOSKELETON IS AFFECTED BY PHYSIOLOGICAL CONDITIONS An increased association of polysomes with the cytoskeleton has been demonstrated in 3T3 fibroblasts under conditions of increased protein synthesis in response to the addifion of insulin (Hesketh and Pryme, 1988). Similarly Vedeler et al. (1990), using Krebs II ascites cells, have shown that insulin during short term incubation (1 h) causes a redistribution of ribosomes between free, cytoskeletalbound and membrane-bound polysome fractions, such that more polysomes were recovered bound to the cytoskeleton. It has thus been demonstrated that the proportions of polysomes in the three different fractions can be modulated by a physiological stimulus such as insulin. In the case of the ascites cells, insulin causes a series of changes over 1—3 h involving first an increase and then a decrease in the proportion of CBP recovered; this suggests that increased polysome-cytoskeleton interaction is not part of a mechanism involving activation of synthesis but plays a role in the continual change in the pattern of proteins being synthesized. This implies that a distinct set of proteins are synthesized on CBP. During a 24h incubation of Krebs II ascites cells in stationary culture in serumfree medium containing insulin as the only growth factor, about 80% of the cells were shown to attach to the substratum (Pryme and Hesketh, 1990). A large proportion of the attached cells showed morphological changes associated with normal adhesion and movement. These changes were shown to be a result of a major alteration in the degree of polymerization of actin. Kirkeeide et al. (1991; 1992) analyzed polysomes in Krebs II ascites cells attached to the substratum and in non-attached cells and found that a shift of ribosomes had occurred from CBP to

Association ofmRNAs and Polyribosomes

39

MBP (50% increase) in attached cells while the amount of FP remained unaltered. The level of protein synthesis was apparently very high in the MBP fraction since more than 70% of ribosomes were in polysomes. Morphological changes observed following insulin treatment were accompanied by a shift of certain proteins among subcellular fractions (e.g., actin and p35). The fibronectin content was about 20 percent higher in attached compared to non-attached cells. The results suggest that morphological changes induced by stimulation with insulin are associated with an increased activity of protein synthesis in the MBP fraction, presumably reflecting a requirement for an increased synthesis of membrane proteins. These results show that ribosomes can be shifted from the polysome compartment associated with the cytoskeleton into polysomes bound to the ER, and this presumably reflects changes in gene expression and in the nature of the mRNA species present in the cells under different conditions. It also appears that stimulation of protein synthesis by translational control mechanisms can also be associated with altered polysome distribution since the change in polysome distribution that occurs in fibroblasts in response to insulin was observed under conditions where it is known that the stimulation of protein synthesis is not inhibited by actinomycin D (Hesketh et al., 1986) and thus occurs in the absence of mRNA synthesis. Again, the mechanism behind the redistribution is not known but it would appear most likely that insulin changes the pattern of existing mRNAs being translated and that these are translated in different compartments; alternatively, it may be that insulin, by some unknown mechanism, redirects polysome complexes from one subcellular compartment to another.

V. WHAT IS THE FUNCTION OF CYTOSKELETAL-BOUND POLYSOMES? The failure in early experiments to extract polysomes from HeLa cells using non-ionic detergent led to the suggestion that "free polysomes" do not exist as such in the cytoplasm and that an association of the ribosome-mRNA complex with the cytoskeleton is essential for translation to occur (Lenk et al, 1977; Cervera et al., 1981; van Venroiij et al., 1981). However, further experiments on a variety of other cell lines has shown that non-ionic detergent treatment releases soluble components such as lactic dehydrogenase together with some 20 to 40 percent of the cell polysomes (Ramaekersetal., 1983; Bird and Sells, 1986;Bagchietal., 1987;Katze et al., 1989; Lequang and Gauthier, 1989; Vedeler et al., 1991 a; Hesketh and Pryme, 1991). Furthermore, these polysomes have a different profile from those retained in the cell matrix and in vitro translation data suggests that they synthesize a different pattern of proteins from the polysomes released by salt treatment (Vedeler et al., 1991a). More recently it has been found that polysomes released from transfected fibroblasts by non-ionic detergent treatment are enriched in the mRNA for P-globin, suggesting that in these cells at least FP are involved in the translation of specific mRNA species (Hesketh et al., 1994). Thus the bulk of the accumulated

40

JOHN E. HESKETH and IAN F. PRYME

data suggests that a proportion of polysomes are recovered in the cytosolic fraction and that these are distinct from those retained in the cytomatrix; therefore the association of the translational complex with the cytoskeleton does not appear to be a prerequisite for translation. Of course it is possible that the "free polysomes" released by non-ionic detergent at low salt are not originally derived from a "free" cytosolic compartment but represent polysomes which are only loosely attached to the cytoskeleton. Neither does it appear that association of mRNAs with the cytoskeleton results in their obligatory translation. Thus, during infection of cells with influenza or vaccinia viruses some host cell mRNAs are associated with the cytoskeleton although they are not translated. It thus appears that association of mRNAs with the cytoskeleton and their subsequent translation are independent events. Indeed, since it has been suggested that ribonucleoprotein particles called prosomes are associated with the intermediate filaments (Grossi de Sa et al., 1988) it is possible that some mRNAs may be associated with the cytoskeleton in a non-translated compartment. Although the association of mRNAs/polysomes with the cytoskeleton does not seem obligatory for translation, it does seem to be of physiological significance because the extent of interaction, as judged by the proportion of polysomes, or mRNAs, associated with the cytoskeleton, varies with physiological conditions. For example, in ascidian oocytes fertilization is associated with an increase in the proportion of polysomes/mRNA recovered in the cytomatrix (Moon et al., 1983) and in fibroblasts the rapid stimulation of protein synthesis by insulin is accompanied by a 20% increase in the proportion of polysomes which are co-extracted with actin (Hesketh and Pryme, 1988). Similarly, in Krebs 11 ascites cells insulin, within 1—3 h, both stimulates protein synthesis and causes a redistribution of ribosomes between the FP, CBP and MBP populations (Vedeler et al., 1990). After 24 hours the cells attach to the substratum and there is both a reorganization of the cytoskeleton and a further alteration in the proportion of polysomes recovered in the CBP fraction (Kirkeeide et al., 1992). In contrast, in virus infected cells where host protein synthesis is inhibited, viral but not the host mRNAs are associated with cytochalasin B releasable, that is, microfilament-associated, polyribosomes (Lenk and Penman, 1979). Data from in vitro translation experiments provided some preliminary evidence that indeed CBP contained a pattern of mRNAs which was different from that of free or membrane-bound polysomes (Vedeler et al., 1991a). In addition. Northern hybridization techniques, together with the use of salt or cytochalasins to separate CBP from MBP, has provided a direct approach to addressing the question of whether CBPs contain different mRNAs. Results from these types of experiments suggest that polysomes isolated from cytoskeletal fractions are enriched in certain specific mRNAs and are thus involved in the synthesis of a distinct set of proteins (see Figure 1).

Association ofmRNAs and Polyribosomes

41

Table 1, Association of Different mRNAs with Free (FP), Cytoskeletal-Bound (CBP) and Membrane-Bound (MBP) Polysomes in Different Cell Lines FP p-globin p-actin hi stone H3

transfected LTK- fibroblasts 3T3 fibroblasts Krebs II ascites

CBP c-myc cyclin A glyceraldehyde-3-phosphate dehydrogenase p-actin

3T3 fibroblasts/Krebs II ascites/HepG2 3T3 fibroblasts/Krebs II ascites/HepG2 3T3 fibroblasts/Krebs II ascites/myoblasts Krebs II ascites

histone H4 histone H3

Krebs II ascites HeLa HepG2

c-fos ribosomal protein L4 ribosomal protein 56

HeLa HepG2

MBP p2-microglobulin insulin receptor glucose transporter 1

3T3 fibroblasts/ Krebs II ascites 3T3 fibroblasts HepG2

Source: Compiled from data in Zambetti et al. (1985), Bird and Sells (1986), Hesketh et al. (1991, 1994), Houland et al. (1995), and Vedeler et al. (1991b).

Using sequential extraction with non-ionic detergent at low salt concentration (FP), 130mM KCl (CBP) and deoxycholate (MBP) the c-myc mRNA was found to be present at greatest enrichment in the CBP (Hesketh et al., 199 lb). More recent data show that the c-myc mRNA is associated with CBP not only in 3T3 fibroblasts but also in Krebs II ascites cells and HepG2 cell lines (Campbell, Houland, Vedeler, Pryme, and Hesketh, unpublished observations) and also in L9 fibroblasts transfected with the c-myc gene (Hesketh et al., 1994). It appears that c-myc mRNA is a useful marker for CBP in a variety of cell lines. As shown in Table 1 other mRNAs are also found to be recovered largely in CBP. Histone mRNA has been found in the cytomatrix and to be released by C3^ochalasins (Bird and Sells, 1986; Bagchi et al., 1987; Heujijerjans et al., 1989; Zambetti et al., 1990) and it is perhaps of significance that mRNAs for other nuclear proteins such as fos (Zambetti et al., 1985) and cyclinA (Houland et al., 1995) also appear to be associated with CBP. More recently, using salt treatment to extract CBP, we have found histone H4 mRNA to be recovered in CBP, but histone H3 mRNA to be present in both FP and CBP (Johannessen et al., 1995). The presence ofmRNAs coding for several nuclear proteins, as well as those coding for ribosomal proteins L4 and 56 (Houland et al.,

42

JOHN E. HESKETH and IAN F. PRYME

1995), on CBP may be related to a need to retain these mRNAs in the perinuclear cytoplasm to more efficient transport of the protein to the nucleus. Such a hypothesis is supported by the recent observation in transfected cells that the c-myc mRNA is not only recovered in CBP but is found in the perinuclear cytoplasm; furthermore, loss from CBP is accompanied by mRNA redistribution; it thus appears that in the case of this particular mRNA association with CBP is associated with a perinuclear distribution. The perinuclear cytoplasm contains a highly developed cytoskeletal network and CBP may partly represent polysomes retained on this filament matrix. However, the mRNAs retained on CBP are not only those for nuclear proteins since glyceraldehyde 3' phosphate dehydrogenase and ribosomal protein LI mRNAs are also found present on CBP (see Table 1). In addition, studies of myoblasts and fibroblasts have shown that the majority of actin mRNA is recovered largely on CBP (Bird and Sells, 1986; Meadus et al., 1990; Hesketh et al, 1991b), and in several of these studies the CBP also show an enrichment in actin mRNA. However, in 3T3 fibroblasts and ascites cells this was not the case (Hesketh et al., 1991b; Campbell, Vedeler, Pryme, and Hesketh, unpublished data) and the actin mRNA appeared to be equally distributed between FP and CBP. The reason for this somewhat unexpected observation is not clear, but two possible explanations present themselves: first, it is known that actin mRNA is present in the cell periphery (Sundell and Singer, 1990; Hill and Gunning, 1993) and so the recovery of actin mRNA in both fractions may reflect the loss of actin mRNA from the fragile cytoskeleton in the cell periphery during the initial detergent treatment; alternatively it may be due to the presence of (3- and y-actin mRNAs in different fractions, a possibility which appears likely in view of recent data showing the mRNAs for the two isoforms to be localized in different areas of the cytoplasm (Hill and Gunning, 1993). Analysis of actin isoform mRNAs in FP and CBP may allow progress to be made in defining the relationship between recovery of an mRNA in CBP and its spatial localization (see below). In summary, there is now good evidence that CBP are enriched in certain specific mRNAs and thus that they are involved in the translation of specific mRNAs. This requires some mechanism to segregate such mRNAs from those translated on the ER (i.e., the mRNAs for membrane proteins such as P-2-microglobulin and glucose transporter 1) or FP (see Figure 1). Since, as described below, there is evidence for spatial segregation of mRNAs within the cell it is possible that there is some functional link between mRNA localization and cytoskeleton-polysome interaction.

VI. THE CYTOSKELETON AND mRNA LOCALIZATION The majority of evidence for an association of mRNAs or polysomes with the cytoskeleton has come from studies of cells grown in culture, particularly HeLa cells and fibroblasts. However, there is data from a number of different experimental systems which indicates that such interactions are not restricted to cultured cells

Association ofmRNAs and Polyribosomes

43

but also occur under normal physiological conditions. Furthermore, it is evident from these studies that such interactions are highly relevant to the specialized structural/organizational features of these cells and that the interaction plays an important physiological role in cell function. Oocytes from amphibians and insects exhibit localization of specific maternal mRNA species to the different poles of the egg (Jeffrey, 1984; Rebagliati et al., 1985; Capco, this volume): for example, in situ hybridization studies have shown that the Vgl mRNA is localized in the vegetal half of the Xenopus oocyte (Weeks and Melton, 1987); in the Drosophila oocyte the transcripts of bicoid and nanos genes are found localized to the anterior and posterior poles respectively (Berleth et al., 1988; Wharton and Struhl, 1989; Gavis and Lehman, 1992) and the cyclin B mRNA is also localized to the posterior pole (Raff et al., 1990). Inhibitor studies suggest that mRNA localization is dependent on association with the cytoskeleton. For example, cytochalasin B has been shown to destroy the normal localized distribution of the Vgl mRNA whereas colchicine was found to prevent translocation of the mRNA to the egg cortex at an earlier developmental stage (Yisraeli et al., 1990). On the basis of such data it was proposed that microtubules were involved in transport of the mRNA and microfilaments involved in attachment and anchoring the message in the egg cortex. Similarly, the translocation of mRNA coding for cyclin B is prevented by colchicine (Raff et al., 1990), although maintenance of localization of this mRNA at the posterior pole appears to be dependent neither on microtubules nor microfilaments. In the case of bicoid mRNA perturbation of the microtubule network with colchicine or taxol leads to disruption of mRNA localization (Pokrywka and Stephenson, 1991). The failure of colchicine or cytochalasin to affect the localization of cyclin B mRNA at the posterior pole (Raff et al., 1990) suggests that either the intermediate filaments are involved in the anchoring of the mRNA or that some non-cytoskeletal factor is involved. Biochemical studies have shown that Vg 1 mRNA is recovered in a cytokeratin-containing cytoskeletal extract of Xenopus oocytes (Pondel and King, 1988), supporting the case for an association of this localized mRNA with the cytoskeleton and suggesting that the intermediate filaments may be involved in its localization. Critical evidence for a role of the cytoskeleton in mRNA distribution has come from genetic studies which have shown that genes necessary for correct localization of bicoid and nanos transcripts code for cytoskeletal or myosin-like proteins (Berleth et al., 1988; MacDonald and Struhl, 1988; Wharton and Struhl, 1989). So, although the exact role of the different filament systems in oocyte mRNA distribution is unclear, in some way the cytoskeleton is vital for maintenance of mRNA localization in various oocyte systems. There is evidence for mRNA localization and RNA-cytoskeleton interactions in other specialized situations (Table 2) where either there is a need for precise targeting of protein within a highly organized cell structure (the muscle fiber) or where the cells possess extremely long cell processes (neurones and oligodendroglia). In skeletal muscle myosin heavy chain mRNA has been reported to be present not only in the perinuclear subsarcolemmal cytoplasm but also in the myofibrillar

JOHN E. HESKETH and IAN F. PRYME

44

Table 2. Examples of mRNAs-which Exhibit Subcellular Localization Tissue Fibroblasts

skeletal muscle

intestinal cells neurones oligodendroglia Xenopus oocytes Drosophila oocytes

mRNA p-actin c-myc a-actin p-actin y-actin vimentin myosin

P-actin MAP2 myelin basic protein Vgl bicoid adducin-like nanos cyclin B

Location peripheral cytoplasm perinuclear cytoplasm perinuclear cytoplasm peripheral cytoplasm perinuclear cytoplasm costameres heavy chain myofibrils/myotendinous junction terminal web dendrites cell processes vegetal pole anterior pole 56D anterior pole posterior pole posterior pole

Reference Sundell and Singer, 1991 Hesketh et al, 1994 Kislauskis et al, 1993 Hill and Gunning, 1993 Hill and Gunning, 1993 Cripeetal., 1993 Aigner and Pette, 1991; Hesketh et al., 1991 Cheng and Bjerknes, 1989 Gamer etal., 1988 Aingeretal., 1993 Rebagliati et al, 1985 MacDonald and Struhl, 1988 Ding etal, 1993 Gavis and Lehman, 1992 Raffetal., 1990

cytoplasm (Hesketh et al., 1991a); approximate calculations suggest that 50 to 70 percent of the mRNA is in the myofibrillar cytoplasm. These experiments, using S^^-labeled riboprobes for in situ hybridization studies of normal muscle, failed to show a distinct banding pattern with respect to mRNA distribution, although a faint banding could sometimes be discerned. However, using digoxigenin-labeled probes and electrically stimulated muscle Aigner and Pette (1990) were able to demonstrate that not only was myosin heavy chain mRNA present in the myofibrillar cytoplasm but that it was also present in a banded distribution, suggesting an association with the myofibrils, albeit the actin-containing I bands. The presence of myosin heavy chain mRNA close to the myofibrils would suggest that the protein is synthesized close to its site of assembly into the sarcomere. Although there has been some debate over ribosome localization in skeletal muscle, recent immunocytochemistry using ant-60S ribosomal subunit antibodies has shown ribosomal material not only to be present in the subsarcolemmal cytoplasm but also to be associated with the myofibrils (Home and Hesketh, 1990a). The staining showed a distinct banding pattern consistent with an association of ribosomes with the myosin-containing A band and recent electron microscope studies have confirmed the presence of polysomes within the A-band regions of the myofibril (Gauthier and Mason-Savos, 1993). Myofibril-associated polysomes may represent a specialized case of CBP (Home and Hesketh, 1990a). At present no explanation has been given for the apparent discrepancy between myosin heavy chain mRNA association with the I-band and the presence of ribosomes/polysomes in the A-band.

Association ofmRNAs and Polyribosomes

45

During periods of growth or hypertrophy when there is an increase in actomyosin synthesis, there was an increase in the proportion of ribosomes associated with the myofibrils, as assessed by the ratio of myofibrillar: subsarcolemmal staining (Home and Hesketh, 1990a, 1990b). This observation suggests that indeed myofibril-associated ribosomes are involved in synthesis of sarcomeric proteins and this is supported by observations that nascent myosin heavy chains are present in polysomes present in the cell matrix of cultured myoblasts (Isaacs and Fulton, 1987). Similarly, vimentin mRNA has been found to be localized close to the costameres in skeletal muscle (Cripe et al., 1993) and such a localization is also close to the site of vimentin accumulation. The synthesis of myosin and other sarcomeric proteins close to their site of insertion into the myofibril provides a mechanism for precise targeting of the proteins during myofibrillar turnover or muscle growth. Neurones also show spatial organization of the protein synthetic apparatus in that ribosomes are found in the cell body and in dendrites but not in axons (Steward and Levy, 1982). Dendrites, but not axons, are capable of transporting RNA (Davis et al., 1987) and, although the precise nature of the transported RNA was not investigated, further work has shown that specific mRNAs are present in dendrites but not in axons (Gamer etal., 1988;Bmckensteinetal., 1990;Kleimanetal., 1990; Tiedge et al., 1991). It would thus appear that neurones show specific localization of certain mRNAs in the dendrites and, at least in the case of MAP2 this mRNA distribution is reflected in the protein localization (Gamer et al., 1988). The transport of dendritic RNA seems to involve the cytoskeleton since the radiolabeled RNA was present in a detergent-insoluble cytoskeletal fraction (Davis et al., 1987) and other neuronal transport processes involve cytoskeletal systems. Oligodendroglia also possess long cell processes and recent work has shown that these cells exhibit localization of a specific mRNA; in this case the mRNA for myelin basic protein has been found to be present in the extremities of these processes in cultured glial cells (Ainger et al.,1993). Furthermore micro-injection of fluorescently labeled mRNA into these cells shows that the mRNA is transported along the cell processes and inhibitor studies suggest that this transport is dependent on the cytoskeleton. Intestinal epithelial cells are highly polarized and in situ hybridization studies have shown that actin mRNA is localized in the apical region of such cells (Cheng and Bjerknes, 1989). Even in a spatially less complicated cell such as a fibroblast there is mRNA localization (see Table 2). Thus it was shown some years ago that in spreading fibroblasts p-actin mRNA is located in the cell periphery close to the lamellipodia where there is a high actin protein concentration (Sundell and Singer, 1990). This localization is not dependent on nascent polypeptide chains but is dismpted by cytochalasin D suggesting that microfilaments are required either for the transport or the anchoring of the mRNA (Sundell and Singer, 1991). More recently it has been shown that in myoblasts the mRNAs coding for p- and y-actin

46

JOHN E. HESKETH and IAN F. PRYME

isoforms are localized in different cell domains with the P-form in the cell periphery and the y-form in the perinuclear cytoplasm (Hill and Gunning, 1993). Thus, there is now a considerable body of evidence showing that in a variety of cells and tissues there is subcellular localization and transport of specific mRNAs and of ribosomes. In many cases localization of the mRNA corresponds closely to the protein localization, suggesting that such mRNA sorting/targeting provides a mechanism for the synthesis of certain proteins close to their site of function, that is, for a targeting of the protein synthetic apparatus. Furthermore it appears in many cases that the cytoskeleton is involved in this mechanism, either as a transport mechanism and/or as a mechanism for anchoring or compartmentalizing the mRNAs (Figure 1).

VII. NATURE OF THE INTERACTION BETWEEN CYTOSKELETON AND mRNAS AND POLYSOMES A.

Role of the 3' Untranslated Region of the mRNA

Inhibition of translation by pactamycin or fluoride fails to release mRNAs from the cytomatrix (van Venrooij et al., 1981; Howe and Hershey, 1984; Bag et al., 1987). Similarly, EDTA treatment has no effect on mRNA release. Since these compounds cause an arrest of translation and a release of ribosomes from mRNA, these observations suggest that mRNAs are retained both on the cytoskeleton and on the ER when not being translated and in the absence of intact polysomes. Such data suggest that the major polysome-cytoskeleton interaction is through the mRNA. This concept is compatible with the available data concerning mRNA localization: P-actin mRNA localization in the cell periphery is not dependent on nascent polypeptide chains (Sundell and Singer, 1990) and microinjection of exogenous mRNA into oocytes has shown that mRNA localization is determined by information within the mRNA (Yisraeli and Melton, 1988). Furthermore, work on oocytes together with data from recent experiments with cells transfected with chimaeric gene constructs has shown that the cytoskeleton-polysome or cytoskeleton-mRNA interaction depends on sequences within the 3' UTR of certain mRNAs. In developing oocytes of amphibians and Drosophila the 3' UTR of certain mRNAs is essential for correct localization of a set of mRNAs (MacDonald and Struhl, 1988; Yisraeli and Melton, 1988; Davis and Ish-Horowicz, 1991). Deletion analysis has shown that for Vgl and bicoid mRNAs a large (340-625bp) region of the 3' UTR is required (MacDonald and Struhl, 1988; Mowry and Melton, 1992), although for other localized mRNAs it has been suggested that there is a consensus sequence which is involved in targeting of a set of mRNAs to a particular location (Gottlieb, 1992). Recent experiments on transfected cells have shown that such a 3' UTR-based localization mechanism is not restricted to amphibian and insect oocytes but also

Association ofmRNAs and Polyribosomes

47

occurs in mammalian somatic cells. When cells were transfected with chimaeric constructs in which either the p-globin coding sequences were linked to the c-myc 3' UTR or the c-myc coding sequences were linked to the P-globin 3' UTR the distribution of the transcripts was shown to depend on the 3' UTR present (Hesketh et al, 1994); the results showed that the c-myc 3' UTR is sufficient to target the P-globin coding sequences to CBP and that the c-myc 3' UTR is required both for association of the c-myc mRNA with CBP and for a perinuclear localization. Similarly, transfection of reporter sequences linked to the a- or P-actin isoform 3' UTRs has shown that the 3' UTR regions of these mRNAs can direct the reporter sequence to different cytoplasmic locations (Kislauskis et al., 1993). In the case of cells transfected with c-myc constructs it was possible to assess both association with CBP and localization of the mRNA (Hesketh et al., 1994), and the data indicate that not only does the cytoskeleton play a role in localization of the c-myc mRNA but that the localized mRNA is probably also being translated. Furthermore, since relocalization from perinuclear to peripheral cytoplasm was paralleled by a shift in the c-myc mRNA from CBP to FP it appears that misdirected mRNAs are still incorporated into polysomes; thus correct localization may not be obligatory for translation to occur. Since these studies indicate that the 3' UTR affects both targeting of the mRNA to CBP and cytoplasmic localization, the picture emerging is that polysome-cytoskeleton association plays an important role in the compartmentation of translation (see Figure 1) by targeting ofmRNAs to specific intracellular locations so that translation occurs in particular sites (Hesketh and Pryme, 1991; Hesketh, 1994). Thus, part of the functional significance of CBP appears to produce a spatial organization of the translational apparatus so that localized synthesis of proteins can occur in discrete cytoplasmic compartments. The extent to which such a mechanism is employed by the cell remains to be fully elucidated, but such mRNA sorting and targeting to provide local synthesis may have particular advantages. Firstly, localized polysomes can produce a steady, continual amount of protein in one location and thus it may be more efficient to produce a greater concentration of the desired protein in such a manner rather than by using protein-based targeting mechanisms. Secondly, it provides a way of targeting protein isoforms. Thirdly, protein production is restricted and so any disadvantageous or unwanted interactions of the synthesized protein with components in other parts of the cell are minimized; this may be particularly important for proteins which are relatively unstable and have a short half-life. The nature of the 3' UTR-cytoskeleton interaction is not understood at present but it seems likely that it will either involve specific sequences or secondary structures which are recognized by specific proteins which bind to the cytoskeleton. Gottlieb (1992) has suggested that there is a conserved 9 nucleotide sequence in the 3' UTRs of some of those mRNAs which are localized in oocytes; interestingly, this sequence would be predicted to form a stem-loop structure. However, when one considers the UTRs of all the mRNAs localized in oocytes there appears to be

48

JOHN E. HESKETH and IAN F. PRYME Table 3. Comparison of Localization, Association with CBP and Targeting via 3'UTR for Four Different mRNAS which have been Studied in Fibroblasts or Myoblasts

mRNA

Localization

Polysome Population

Targeting

c-myc p-actin y-actin P-globin

perinuclear peripheral perinuclear none

CBP CBP ?? FP

3'UTR 3'UTR ??

—

little sequence homology suggesting both that in other cases different sequences are involved and that secondary structure of the 3' UTR region is important. Indeed, in a series of different Dro5o/?/z//a species the 3' UTR of the different fe/co/JmRNAs show little sequence homology but similar predicted secondary structure (MacDonald, 1990), although all the mRNAs are localized similarly. It would thus appear therefore that specific secondary structures are important in mRNA localization. B. mRNA/Ribosome Binding Proteins

In oocytes a number of 3' UTR-binding proteins have been identified (e.g., Schwartz et al., 1992) and, as expected, these appear to play some role in the localization of the mRNA. One would also anticipate that the 3' UTR-cytoskeleton interaction would involve proteins which would bind both to the cytoskeleton and to specific 3' UTR signals. Interestingly, recent studies have shown that the cytomatrix contains proteins which bind to the P-actin mRNA 3' UTR (Sharpless et al, 1993) and such proteins are clearly prime candidates for playing a targeting role. The binding of different proteins to different sequences in mRNAs could provide the basis of the spatial segregation and specific localization of mRNAs. The ability to generate a variety of localization signals is important because mRNAs associated with CBP or targeted via the 3' UTR may be targeted to different domains of the cytoskeleton and to different parts of the cell (see Table 3). P-actin mRNA is located in the cell periphery (Sundell and Singer, 1990) and c-myc to the perinuclear cytoplasm (Hesketh et al., 1994) and yet targeting of both mRNAs appears to involve microfilaments (Table 3); in other cases there may be targeting to specific regions of the cell periphery so as to achieve a polarized concentration of actin mRNA, for example in intestinal epithelial cells (Cheng and Bjerknes, 1989). A mechanism is therefore required which will produce differential mRNA-cytoskeleton interactions and thus specific localization. Such mRNA-cytoskeleton interactions and targeting mechanisms would appear to require a multitude of proteins that interact with the mRNA, cytoskeleton and perhaps also the ribosomes. In order to begin to analyze the differences in polysome-associated proteins in those polysomes associated with the cytoskeleton

Association ofmRNAs and Polyribosomes

49

Moss et al. (1991, 1993, 1994) have studied the protein composition of extracts prepared from the three polysome-containing fractions; high salt washes of FP, CBP, and MBP from MPC-11 and Krebs II ascites cells were analyzed by SDSPAGE. The protein profiles in the individual polysome fractions were quite different in the two cell lines and changes promoted by the stimulation of protein synthesis following incubation of cells with insulin were also different in nature. It is too early at present to assign specific properties to the individual proteins, however, the differences in proteins observed in the high-salt washes of the three polysome fractions in the two cell lines may suggest a role of specific polysomeassociated proteins in the regulation of protein synthesis. Altered protein patterns seen to occur in the three fractions after insulin treatment may possibly reflect modifications of the control mechanism(s) which result in the synthesis of new proteins. Insulin stimulation may result in the production or activation of proteins which ftinction as signals promoting the entry of an inactive mRNA into an active polysomal complex. It is likely that such proteins would be ultimately present in mRNA-ribosome aggregates and thus be identified in high salt wash extracts of polysomes. Hesketh and Pryme (1991) and Kirkeeide et al. (1993) have envisaged that following transport from the nucleus into the cytoplasm mRNAs may fall into two categories: firstly, species which are translocated in the cytoplasm and then sorted into the correct polysome compartment for immediate translation, and secondly, mRNA molecules which are not initially translated but first sequestered into inactive ribonucleoprotein particles for a period of storage prior to active translation. It is thus anticipated that a range of proteins play important roles in mRNA transport/storage/sorting phenomena. There are a number of instances which indicate that alterations/modifications in proteins are important in regulating the process of translation. It has been shown, for example, that phosphorylation and dephosphorylation respectively, of the a-subunit of eIF-2 and the 25 kDa (eIF-4e) subunit of eIF-4f occurs following the deprivation of nutrients or serum in cells grown in culture (Duncan et al., 1987), and fiarthermore, it appears that an activation of at least three initiation factors occurs upon maturafion (Clemens et al., 1987). Moss et al. (1994) have shown that the FP, CBP, and MBP populations isolated from Krebs II ascites and MPC-11 cells have their own individual complements of poly(A) binding and non-poly(A) binding proteins. It is likely that this may reflect the different growth characteristics of the two cell lines; Krebs II ascites cells are propagated in the peritoneal cavity of mice, large quantities of cells (>10^) accumulafing in the ascitic peritoneal fluid while MPC-11 cells grow in single cell suspension in roller or spinner culture. Krebs II cells when removed from the host and put into suspension culture do not divide and thus they appear to be completely dependent on some undefined growth factor(s) provided by the host environment. It is therefore tempting to suggest that the observations concerning the differences in content of poly(A) binding proteins associated with the three polysome fractions may reflect differences in the types of mRNAs being translated under the very different growth conditions exhibited by

50

JOHN E. HESKETH and IAN F. PRYME

the two cell lines. This possibility is supported by the findings of Olsen et al. (1988) who showed that the pattern of proteins in high-salt washes of total polysomes isolated from synchronized populations of L-929 cells varied during the cell cycle; some proteins were present throughout the whole cell cycle, while others exhibited various degrees of cyclical behavior. The levels of initiation, elongation; and termination factors required for protein synthesis during the cell cycle would not be expected to show any high degree of variation. It is possible, therefore, that the cell cycle related differences in the pattern of ribosome-associated proteins may indicate a role for such proteins in regulating the translation of specific mRNA species. Although our knowledge concerning the exact nature of the function of poly(A) binding proteins is relatively poor some information has emerged. Schroder et al. (1988) using a monoclonal antibody against the poly(A) binding nuclear envelope fraction isolated from rat liver observed reaction with two poly(A)+ binding proteins of 65 and 83 kDa. In addition the antibody stained the 83 kDa polypeptide in rat liver polysomes, and also a 112 kDa protein band. Using immunofluorescence microscopy they found that in CV-1 cells the immunoreactive polypeptides were associated not only with the envelope, but also with a cytoplasmic fibrous network; this was partially colocalized with the microfilament bundles. These observations would indicate that a common protein is involved both in the translocation of mRNA and then in its attachment to the cytoskeleton. Two polysome-associated proteins of 31 and 58 kDa, purified from rat liver, have been shown to possess high affinity for poly(A) (Schroder et al., 1986). Both proteins appear to exert their effects in the mRNA-translocation system in the nuclear envelope. It is likely that the 31 kDa protein corresponds to the mRNA-transport protein characterized by Moffett and Webb (1983). The affinity of both stimulatory proteins for poly(A) suggest that they might be associated with adenylated messenger precursors in the nucleus and remain associated with mRNA in the cytoplasm. Poly(A) binding proteins may thus be involved in binding mRNA to cytoskeletal elements and could therefore function in the translocation of newly synthesized mRNAs to their respective subcellular compartment(s) for translation in the appropriate polysome fraction. It is possible that such proteins may be one of a large number of as yet unknown proteins which act as sorting signals which ultimately direct mRNAs to their correct polysome compartment, perhaps following a period of storage in a non-translated state (Hesketh and Pryme, 1991). Although the available data indicates that mRNA localization depends upon both the cytoskeleton and 3' UTR sequences, the details of the transport and localization mechanisms, the sequences in the mRNA molecules, and the cytoskeleton-ribosome interactions remain to be defined. It is not clear for example if the mRNA is transported in a RNP particle or in a mRNA-ribosome or polysome complex. At what stage of the transport pathway does polysome formation occur? One can envisage several possible sequences of events (see Figure 2): first, the mRNA associates with a ribosome before any interaction with the cytoskeleton; second.

Association ofmRNAs and Polyribosomes (T) Binding of protein to 3'UTR

51

(2) Association with cytoskeleton: transport

(3) Localization : anchoring to cytoskeleton

MODEL I : mRNA TRANSPORT Translation cannot occur until after Localization

I (4) Localized translation complex

Ribosone binding

^ U<:?>iA ""

•-&er&—"—

MODEL I I : POLYSOME TRANSPORT

t

is translation repressed during transport or does translation occur prior to localization ?

Anchoring

31 (T) Binding of protein to 3UTR

(2) Polysome formation

(3) Association with cytoskeleton: transport

Figure 2. Two alternative models for the transport and localization of mRNA and polysomes. In model 1 the mRNA is transported itself or in a RNP and translation cannot occur until after the mRNA is localized. In model 2 the polysome complex is transported. For simplicity, (a) the localization signal in the 3'UTR is shown as a stem-loop structure (the precise nature of such signals is not known), and (b) the models show only one localization motif and one mRNA-binding protein linking the mRNA to different cytoskleletal structures whereas in reality different RNA structures and/or proteins may be involved in transport and localization.

ribosome binding occurs after association of the mRNA with the cytoskeleton and after transport of the translation complex (i.e., the mRNA is transported alone or as a ribonucleoprotein complex); third, the mRNA binds to the cytoskeleton, then binds one or more ribosomes and the translation complex is then transported. In oligodendroglia myelin basic protein mRNA has been reported to be transported via the cytoskeleton and in some form of particle which appears to contain components of the translation complex (Ainger et al., 1993; Barbarese et al., 1995). However, we do not know if translation is arrested during transport. Whatever the sequence of events which leads to the formation of the polysomecytoskeleton complex it is now clear that in addition to the primary, and we assume initial, interaction between mRNA and cytoskeleton there are also interactions between the cytoskeleton and other components of the translation complex. EDTA and ribonuclease have been found not to release ribosomes from the cytomatrix

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although polysomes were dissociated (Howe and Hershey, 1984) and this indicates that ribosome association with both the membrane and cytoskeletal elements in the deoxycholate fraction did not require the continued presence of intact mRNA. It seems likely, therefore, that after the initial interaction of the mRNA itself with the cytoskeleton there is subsequent binding of the ribosomes, perhaps to a "receptor" on the cytoskeleton, in a similar manner to that which occurs in the attachment of MBP to ER membranes (Savitz and Meyer, 1990). Loss of mRNA from the cell matrix after puromycin treatment (Taneja et al., 1992) has also been interpreted as evidence for ribosome or nascent peptide chains stabilizing the mRNA-cytoskeleton interaction. The recent observation that elongation factor la may be an actinbinding protein (Yang et al.,1990) ftirther suggests that polysome-microfilament links may partly occur through interactions which do not directly involve the mRNA. Mechanisms involving such interactions may be part of the as yet unknown mechanisms which retain or anchor the mRNA or translation complex at its correct destination.

VIIL FUTURE PERSPECTIVES Our knowledge of the interaction of ribosomes and mRNAs with the cytoskeleton has reached a critical stage. It is now clear that a proportion of polysomes are associated with the cytoskeleton and that this appears to be due to the ability of specific mRNAs to interact with the cytoskeleton. However, little is known about the functions and detailed mechanisms of the interaction. It is in these areas that future research will be directed. Initially it will be important to determine which mRNAs are associated with the cytoskeleton, and what properties these mRNAs share, so as to clarify the significance of the interaction, particularly to elucidate the role of the cytoskeleton in transport and compartmentalization of mRNAs and polysomes. Using in situ hybridization techniques it should prove possible to clarify both the relationship between the subcellular location of a mRNA and its presence on CBP and whether CBP represent one or more distinct compartments. In order to better understand the mRNA-cytoskeleton interaction it will be necessary to identify both the nature of the RNA structure/sequences required for localization and the proteins involved in mRNA-cytoskeleton interactions. At present it appears that the major cytoskeletal-polysome interaction is through the microfilaments whereas the roles of the intermediate filaments and microtubules are less well defined; future work will describe the different roles of three filament systems in mRNA transport and targeting. Knowledge of these processes will give greater understanding both of the ways in which newly synthesized proteins are targeted to the location in the cell where they are required and the relationship between mRNA localization, stability, and translational control.

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ACKNOWLEDGMENTS This work was supported by the Scottish Office Agriculture and Fisheries Department (JEH) and the Norwegian Research Council for Science and Humanities (IFP).

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SPECIALIZATIONS IN CYTOSKELETAL FUNCTION DURING EARLY DEVELOPMENT

David G. Capco

I. Introduction II. Special Developmental Problems Faced by Eggs and Embryos III. The Cytoskeleton in Early Development of Nonchordates A. Insects B. Echinoderms IV. The Cytoskeleton in Early Development of Chordates A. Ascidians B. Mammals C. Amphibians V. Summary Acknowledgment References

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I. INTRODUCTION Investigations employing somatic cells have resulted in an extensive body of knowledge concerning the organization and function of the cytoskeleton. In conThe Cytoskeleton, Volume 2 Role in Cell Physiology, pages 59-112 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 59

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trast, most studies examining the organization and function of the cytoskeleton in gametes or embryos have only involved searches for structures comparable to those identified in studies of somatic cells. Well known systems that have provided much information for such analyses in somatic cells include brush border cells from the intestines of a variety of organisms, the giant axon of the squid, the red blood cell, the giant algal cell, Nitella, and a plethora of cell types maintained by cell culture (Kreis, 1990; Vallee and Shpetner, 1990; Bretscher, 1991; Nickerson and Penman, 1991; Skalli and Goldman, 1991; Heintzelman and Mooseker, 1992). These studies have contributed greatly to the understanding of assembly mechanisms of cytoskeletal systems, the intricate and highly variable three-dimensional organization of the cytoskeleton in different types of somatic cells, and mechanisms which regulate the cytoskeleton. Studies investigating the cytoskeleton within somatic cells fall into two categories. (1) Some studies that analyze cytoskeletal organization and function have employed cells immediately removed from the adult organisms, as is the case for red blood cells or intestinal cells. These types of cells are highly specialized and represent the mature, highly differentiated form of the cell type in the organism. (2) Other studies, representing the large majority of studies directed at analysis of the cytoskeleton, have been conducted using cells maintained in vitro by cell culture techniques. Together, these analyses have characterized distinct roles for the cytoskeleton in control of cell shape, intracellular transport, protein and mRNA localization and cell mobility. 1. Cell Shape Control. Cells contain a three-dimensional organization of cytoskeletal elements which reflect the particular role they perform in the organism. This three-dimensional organization is established by the three main filament systems of the cytoskeleton (actin filaments, microtubules, and intermediate filaments) and maintains not only the intracellular organelle distribution characteristic of the cell, but also the three-dimensional configuration of the cell exterior (Brinkley et al., 1980; Fey et al, 1984; Felice et al., 1990; Matsudaira, 1991; Bretscher, 1991; Nickerson and Penman, 1991; Skalli and Goldman, 1991). This is complicated by the fact that cells undergo extensive internal and external reorganizations during cell cycle progression, locomotion, and other processes characteristic of life. Thus, the cytoskeleton acts both to support the cell infrastructure, and to effect dynamic cell rearrangements. 2. Molecular Motors and Positioning of Intracellular Components. Cells exhibit non-random patterns of organelle distribution and trafficking of vesicles that can establish polarity in cells. Maintenance of this polarity requires that organelles and vesicles must be actively transported to and from different regions of the cell. The cytoskeleton attaches to, and at least in some cases moves, organelles and other components of the cell using both microtubule- and actin filament-based systems of transport (Kreis, 1990; Theriot and Mitchison, 1992). For example, microtubules in squid axons transport organelles between the cell axon and the cell body (Vale et al, 1985 a, 1985b; see reviews by Schroer and Kelley, 1985; Vale et al., 1986;

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Vallee and Shpetner, 1990). Movement away from the cell body is associated with the protein kinesin, and movement toward the cell body appears to be associated with cytoplasmic dynein (Vale et al., 1986). Evidence suggests that microtubules play a similar role in other cell types as well, such as the amoeba (Koonce and Schliwa, 1986), and other, more widely-used model systems. For example, in cultured mammalian cells, microtubules appear to move endosomes toward the cell center where they contact lysosomes, which are also associated with microtubules (Matteoni and Kreis, 1987; Scheel and Kreis, 1991). Actin-based organelle transport has been demonstrated in the plant, Nitella, where it was shown that beads coated with heavy meromyosin move along the length of actin cables in the cytoplasm of ruptured M/e//fl cells (Sheetz and Spudich, 1983). Myosin-coated beads also move along isolated actin bundles in vitro, and organelles move along actin filaments in permeabilized amoebae (Koonce and Schliwa, 1986) providing further support for the existence of actin-based organelle movement. 3. Positioning of mRNA and Protein. As in the case of organelles, cells also exhibit non-random distributions of mRNA and protein (for reviews see Hesketh and Pryme, 1991 and Merketh and Pryme, this volume). For example, specific cytoskeletal mRNAs are localized near the site where their cognate protein is needed: Actin mRNAs can be found at the cell periphery, a region enriched in actin filaments, and tubulin mRNA is concentrated near the nucleus, a region enriched in microtubules (Lawrence and Singer, 1986). The cytoskeleton controls at least part of the heterogenous protein distribution as well as a number of different mRNAs (Lenketal., 1977; Fulton etal, 1980;Ben-Ze'evetal., 1981; Van Venrooij et al., 1981; Bonneau et al, 1985; Singer et al., 1989; Hoock et al, 1991; Vedeler et al., 1991). Evidence also suggests that the protein synthetic machinery is associated with the cytoskeleton (Dang et al., 1983; Howe and Hershey, 1984) and protein synthesis is known to occur on cytoskeleton-associated mRNA (Lenk and Penman, 1979; Van Venrooij et al., 1981; Fulton and Wan, 1983; Hesketh and Pryme, 1991). A variety of enzymes are also associated with the cytoskeleton, including tRNA-synthase (Dang et al., 1983), creatine phosphokinase (Eckert et al., 1980), glycolytic enzymes (Liou and Anderson, 1980; Hand and Somero, 1984; Shearwin et al., 1990), and aldolase (Pagliaro and Taylor, 1988). The functional significance of these interactions is not clear, but in some cases it appears that up or down regulation of the protein or mRNA may result from its association with the cytoskeleton. It has also become evident that the cytoskeleton may modulate intracellular signaling activities by controlling signal transduction across the plasma membrane. For example, tyrosyl kinase (Cinton and Finley-Whelan, 1984; Kellie et al., 1991), growth factor receptor kinase (Landreth et al., 1985; Payrastre et al., 1991), and GTP-binding proteins (Jesaitis et al., 1988) associate with the cytoskeleton, and in some cases the association is correlated with changes in the activity of these proteins (Jesaitis et al., 1988).

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4. Mediation of Cell Motility. Many cell types undergo some form of movement. In somatic cells, movement is usually of the amoeboid type, which is mediated by contracting stress fibers (bundles of actin filaments) coordinated with adhesion plaque formation and membrane flow (for review see Schliwa, 1986; Singer and Kupfer, 1986; Conrad et al., 1989; Theriot and Mitchison, 1991; Bretscher, 1991). In some single-celled organisms, movement is accomplished by the beating of numerous cilia, actin filament-containing structures which extend from the cell surface or by the rotation of one or several flagella, microtubule-containing structures which also extendfi*omthe cell surface. There are limitations as to what extent the functions of the cytoskeleton described above can be generalized to all types of cells in various organisms. Knowledge acquired from analysis of cells maintained by in vitro culture necessarily has the limitations of the cell culture method, that is (1) all cell types will not survive in culture and (2) cells in such a system lose their histotype and many lose their differentiated phenotype. Similarly removal of cells fi-om an adult organism for analysis may allow examination of terminally differentiated cells, but the physical removal of cells may induce a damage or wound response that changes the activities of the cells. Consequently, caution must be applied when generalizing functions for all cells.

II. SPECIAL DEVELOPMENTAL PROBLEMS FACED BY EGGS AND EMBRYOS Gametes are unlike any other cells of the organism in that the fusion of the male and female gametes produce the zygote, a cell with the developmental potential to produce every cell type and tissue type of the organism. Moreover, gametes represent unusually structured cells whose architectural specializations are designed to promote the formation of the zygote and allow rapid progression through early embryogenesis. The sperm of most species is a motile, highly streamlined cell whose function is to penetrate various extracellular matrices surrounding the eggs and subsequently penetrate the egg plasma membrane and activate the program of early development while delivering the paternal genetic information. In contrast, the egg is an immotile cell of much greater cytoplasmic mass. Its function is to respond to the penetrating sperm by initiating the program of early development, and also to contain the maternal genetic information. Once the sperm and egg fuse, a cascade of intracellular signals (Bement and Capco, 1990a, 1990b, 1990c; Bement, 1992; Berridge, 1993) progressively restructure the egg cytoplasm and nuclear material into that of the zygote. Though the zygote and the egg are separated in developmental time by only a few minutes, structurally and functionally the zygote becomes a very different cell from the egg (Capco and Bement, 1991). The sperm, egg, and zygote clearly are cells, however all have very different developmental roles. As cells, they must perform some activities in common with all cells.

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but their developmental role is a property not shared with all cells and it can be anticipated that they exhibit some highly specialized activities. The cytoskeleton, which is involved with many important cellular functions, is likely to have a role in the functioning of these cells. Moreover, it can be anticipated that cytoskeletal specialization will exist to mediate both the specialized cellular structure of the gametes and zygote, and perhaps just as important, the specialized developmental roles of these unique cells. This review will focus on the developmental specializations of the cytoskeleton in the egg, zygote, and early embryo. Eggs and embryos are challenged with special developmental problems which are different from other cells. First, as indicated above, the zygote must be developmentally totipotent, and thus must be able to express the developmental information to produce every cell type in the organism; not simply to make daughter cells as is the case for somatic cells grown in vitro. Second, fertilization requires the restoration of diploidy, consequently specializations must exist to promote and allow pronuclear fusion. Moreover, mechanisms must exist to allow only one paternal pronucleus to be active within the egg, either by allowing only one sperm to penetrate the egg or by inactivating other paternal pronuclei. Third, eggs and embryos exhibit unusual regulation of the cell cycle and cytokinesis. Eggs, awaiting sperm penetration are arrested in the cell cycle, with eggs of different species arrested at different points (e.g., metaphase, interphase, etc.). Sperm penetration releases the cell cycle arrest, but without fusion with sperm, eggs appear to undergo a process resembling apoptosis. Sperm penetration initiates progression through the cell cycle of the zygote and early embryo, and this cell cycle progression is quite different from that within somatic cells of the same species, usually exhibiting rapid cycling by reduction or omission of the length of Gap 1 or Gap 2 of the cell cycle. Fourth, accompanying their special developmental roles eggs, zygotes, and cells of the early embryo (i.e., referred to as blastomeres) encounter special challenges as cells. In most species these cells are large with a very small nuclear to cytoplasmic ratio. The cytoplasm of these cells exhibits extremely intricate positioning of intracellular components that results in a cellular polarity for at least the egg and zygote of most species. While cellular polarity, and the positioning of intracellular components is a problem faced by all cells, in eggs, zygotes, and embryos the scale of the problem is larger because the size of the cell is much greater. Fifth, the large size of the egg makes responding to an event occurring at the plasma membrane of a cell, be it binding of a ligand to a receptor or binding of the sperm to the egg plasma membrane, an event which requires transduction of the signal across large distances to reach the cell interior. The developmental challenges described in the previous paragraph are met by the egg and embryo in a variety of ways which are discussed in this chapter. Comparison of the approaches employed in eggs and embryos reveals that at least four cytoskeletal specializations are conserved among eggs of different species. (1) There is a highly developed cortical cytoskeletal domain, usually associated with developmental information, which undergoes reorganization at key developmental

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transitions, (2) The entire cytoskeleton is reorganized at key developmental transitions, (3) Elements of the cytoskeleton, usually microtubules, are involved in pronuclear transport, and (4) Regulation of the cytoskeletal reorganizations is mediated through signal transduction mechanisms. Investigations will be considered that demonstrate each of these conserved features of the cytoskeleton for eggs and embryos of chordates and nonchordates. These same conserved features can be observed in eggs and embryos of nematodes, annelids, and oligochaetes; however, they will not be considered here as very few reports concerning the cytoskeleton in the early development of these organisms have arisen since the area was previously reviewed (Bement et al., 1992).

III. THE CYTOSKELETON IN EARLY DEVELOPMENT OF NONCHORDATES A.

Insects

Investigations of the cytoskeleton within eggs and embryos of Drosophila serve to demonstrate many major cytoskeletal specializations unique to development that are unanticipated from the results of studies employing somatic cells. A number of recent investigations using the Drosophila system have demonstrated a highly dynamic cytoskeleton uniquely organized to meet the challenges of this developmental system. Although many of these recent studies have not made full use of the genetic manipulations possible in this organism, the approaches used in many of the recent studies, when coupled with genetic analyses, hold the potential to reveal may details of the functional significance and mechanisms of regulation of the cytoskeleton during development. As a developmental system, Drosophila has several unique features related to the formation of oocytes and embryos which are accompanied by specializations of the cytoskeleton. These unique developmental features will be outlined first and then used to illustrate the cytoskeletal structures accompanying the developmental specializations. During oogenesis the Drosophila oocyte develops as one of 16 cells linked to one another by intercellular bridges. The oocyte prepares for its developmental fate by acquiring a large amount of nutrients and developmental information from these linked cells, which are referred to as nurse cells. After fertilization, nuclear division (i.e., karyokinesis) ensues in the interior of the egg in cytoplasmic islands, without a corresponding cytokinesis. Thus the multinucleate embryo forms a syncytium (or more accurately termed a Plasmodium) bounded by one expanse of plasma membrane and cell cortex. Nuclei undergo two classes of migration to the surface. First at nuclear division 9, a few nuclei migrate to the posterior pole of the cell, divide two more times, and subsequently cellularization occurs around these nuclei only. This first migration of nuclei establishes the pole cells which are progenitors of the germ cells and essential for the continuance of the species. The second migration of nuclei occurs at nuclear division 10 when most of the remaining

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nuclei in the cell interior migrate to the cell cortex, divide four more times at the cortex, and then cellularization occurs around these nuclei. The developmental events outlined above are accompanied by several specialized cytoskeletal structures. During oogenesis these specializations include the existence of cytoskeletal elements that maintain and transport nurse cell components through the intercellular bridges and allow for mixing of the oocyte cytoplasm with the cytoplasm imported from the nurse cells. Moreover, cytoskeletal elements also precisely position developmental information within the oocyte. After fertilization, this multinucleated syncytium contains one extremely large cortical cytoskeleton which undergoes extensive reorganization into hundreds of individual pockets as nuclei, and their perinuclear cytoskeletal components, arrive at the cortex from the cell interior during one of the nuclear migrations. Subsequently, thousands of individual cytoskeletal pockets form in the cortex as nuclear divisions continue. The nuclei, after migration, face two challenges. They must remain at the cortex (i.e., not return to the cell interior) and they must continue karyokinesis in an area increasingly crowded with nuclei without the interchange of chromosomes from different mitotic nuclei that could result from mistargeting of spindle microtubules to adjacent chromosomes (i.e., without the protection of a delimiting plasma membrane isolating each nucleus). Later in development, the plasma membrane of the syncytium invaginates around each nucleus resulting in cellularization and isolating each within an island of cytoplasm bounded by membrane. These cell cycle-related events occur synchronously throughout this very large sync3l:ium suggesting there exists an elaborate regulatory system which may involve signal transduction. Late during the process of oogenesis in each egg chamber (e.g., 15 nurse cells and one oocyte) the nurse cell cytoplasm is transferred through cytoplasmic bridges into the oocyte by physical contraction of the nurse cells. Several investigations have provided evidence indicating that actin filaments are involved in this contraction (Gutzeit, 1986a, 1986b; Cooley et al., 1992, Theurkauf et al., 1992). The product of the chickadee gene, which bears structural similarity to the actin binding protein, profilin, appears responsible for the establishment of one of the two actin networks in Drosophila ovaries (Cooley et al., 1992). One actin filament network exists in the subcortical region of the nurse cells throughout oogenesis, while the second network forms just prior to the transfer of nurse cell cytoplasm to the oocyte, and appears to have a structural role in maintaining the nurse cell nuclei position. Chickadee mutations result in disruption of the second actin network allowing the nuclei to drift, and consequently clog the cytoplasmic bridges to the oocyte, thus preventing transfer of cytoplasm from the nurse cells to the oocyte (Cooley et al., 1992). Based on these results Cooley et al. (1992) speculated that the subcortical actin network in nurse cells is involved in generation of contractile force. As the nurse cell cytoplasm enters the oocyte it is mixed with the oocyte cytoplasm by cytoplasmic streaming within the oocyte. The cytoplasmic streaming appears to be a microtubule-dependent process (Gutzeit, 1986; Theurkauf et al., 1992) generated

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by a spiral array of microtubules around the subcortex of the ooc5^e (Theurkauf et al., 1992). Although there is transfer of the mass of cytoplasm from nurse cells to the oocyte late during oogenesis, a more specific form of transport exists that translocates developmental information, without the bulk of the cytoplasm, earlier during oogenesis and localizes it at specific sites in the egg (St. Johnston et al., 1989; Ephrussi et al., 1991; Kim-Ha et al., 1991; Pokrywka and Stephenson, 1991; Cheung et al., 1992). Such specific transport may be mediated through the microtubule array extending from the oocyte to the nurse cells at early stages of oogenesis in Drosophila (Theurkauf et al., 1992) and other insects (Valdimarsson and Huebner, 1989). In support of this, the translocation of at least one developmental component, the bicoid mRNA, appears to require this intact microtubule network (Pokrywka and Stephenson, 1991). These studies demonstrate that oogenesis in Drosophila occurs concurrently with a continuous reorganization of the cytoskeleton in both nurse cells and the oocyte of each egg chamber. At fertilization the Drosophila egg contains a relatively uniform cortical cytoskeleton approximately 3 |im deep and composed of actin, myosin, and microtubules (Warn et al., 1980, 1984; Warn and Warn, 1986; Karr and Alberts, 1986). At this early stage spectrin, a protein known for its ability to link actin to the plasma membrane (Nelson and Veshnock, 1986; Del Buono et al., 1988), as well as actin filaments and microtubules are enriched in the cortical cytoskeleton, and are also present throughout the interior cytoplasm (Kellogg et al., 1988; Pesacreta et al., 1989). When the nuclei migrate to the periphery during nuclear divisions 8-10 these nuclei cause extensive reorganization of this cortical cytoskeleton such that it divides into hundreds of cytoskeletal pockets in which the actin, tubulin, spectrin, and myosin are organized around each of the hundreds of nuclei which entered the cortex (Warn et al., 1980: Karr and Alberts, 1986). Nuclear migration in other developmental systems is a problem faced by the egg and/or sperm pronuclei within the cytoplasm of the fertilized egg. In these other systems, as in Drosophila, the genetic information within the sperm and egg must migrate together so that syngamy can occur resulting in the formation of the zygote nucleus. However, inDrosophila embryos the problem of nuclear migration is faced again later in development when several hundred nuclei must migrate from the interior of this syncytium to the periphery between nuclear divisions 8 and 10. The positioning of nuclei in the interior of the embryo as well as migration of these nuclei to the periphery appears to be under the control of three independent maternal genes N26, N441, and paralog mapped to loci in which genes for cytoskeletal proteins have not been identified (Hatanaka and Okada, 1991a, 1991b). Aphenocopy of these mutants can be induced with cytochalasin B (Hatanaka and Okada, 1991b), suggesting that nuclear migration involves actin filaments. However, cytochalasin B does not appear to inhibit nuclear migration in all insect embryos (Wolf, 1978) and a role for actin in nuclear migration appears in conflict with reports by Zalokar and Erk (1976) and Raff and Glover (1989) who showed that drugs which disrupt microtubules (i.e., colchicine) inhibit nuclear migration. These

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observations are not as disparate as they might first seem, however, since recent studies have suggested a parallel role for actin and microtubules in cells (Clark and Meyer, 1992; Lees-Miller et al., 1992); thus, disruption of one cytoskeletal element could affect the other and block nuclear migration. In the future it is likely that genetic approaches using Drosophila will allow the role of actin filaments and/or microtubules in nuclear migration to be defined; for example, by attempts to rescue mutants lacking, or depleted in, actin filaments, microtubules, or actin binding proteins by injection of specific cytoskeletal proteins (or the gene linked to an inducible promoter) and monitoring of the ability of nuclei to translocate. An increasing amount of evidence suggests that it is not the migration of nuclei to the cortex but rather the centrosomes which accompany the nuclei, which induce reorganization of the cortical cytoskeleton. A pharmacological agent, aphidicolin, injected into early Drosophila embryos prevents nuclear migration, although the drug has no effect on the migration of the centrosomes to the cortex; upon arrival of these centrosomes, the actin and tubulin within the cortex are reorganized as they would be in untreated embryos (Raff and Glover, 1989). At first evaluation it might be considered that this effect was an artefact of treatment with the pharmological agent. However, this is unlikely for two reasons. Firstly, there is a normal separation of the centrosome from the nucleus, accompanied by centrosomal migration, in the yolk-filled embryo interior (Callaini and Dallai, 1991), suggesting that the results induced by the drug are not that unusual iox Drosophila embryos. Secondly, results from other investigators have supported a role for the centrosome in reorganization of the cortical cytoskeleton; in injection of exogenous DNA or irradiation of embryos with ultraviolet light has shown that exogenous DNA induced assembly of the nuclear lamina, nuclear membrane and nuclear pores, while centrosomes were capable of independent migration to the cortex and upon arrival the centrosomes induce reorganization of the cortical cytoskeleton Yasuda et al. (1991). Once the centrosome has migrated to the cortex a variety of components of the cortical cytoskeleton reorganize around the centrosome, including microtubules, microtubule-associated proteins, actin filaments, actin-binding proteins, spectrin, and myosin (Warn et al., 1980: Karr and Alberts, 1986; Kellogg et al., 1989; Miller et al., 1989; Pesacreta et al., 1989; Young et al., 1991). How does the centrosome serve as a center which organizes this diverse array of proteins? Clearly, for microtubules, the centrosome can serve as an anchoring or nucleation site, but its role for organizing the other cytoskeletal proteins is less clear. Disruption of centrosomal microtubules alters the cap of actin filaments (Callaini et al., 1991) while disruption of actin filaments alters the centrosomal organization (Callaini and Riparbelli, 1992) suggesting that there is an interaction between microtubules and actin filaments. This result is not surprising in light of recent reports suggesting that a form of actin is present in the centrosome and that actin-related proteins are necessary for microtubule-dependent transport (Goldstein and Vale, 1992; LeesMiller et al., 1992; Clark and Meyer, 1992). If the centrosome can affect the organization of both microtubules and actinfilamentsthen other proteins associated

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with microtubules and actin filaments may become reorganized because of their binding affinities for actin and microtubules. Clearly further studies are needed to resolve how the different components interact during reorganization. The unique organization of cytoskeletal elements around the centrosome and nucleus at the time when nuclei migrate to the cell cortex permits an assessment of potential developmental function(s) of the reorganized cytoskeleton. One role for the individual islands of cytoskeletal elements organized around each of the nuclei in the cell cortex spectrin in preparation for their role in the formation of cleavage furrows at the time of cellularization, that is to organize actin, myosin, and has been addressed by several investigators, (Pesacreta et al, 1989; Warn and Robert-Nicoud, 1990;Planquesetal., 1991; Young etal., 1991). In addition, these cytoskeletal islands (in particular the dome-shaped actin caps) may function to delimit the c)^oplasmic territory occupied by each nucleus in the cortex. During mitosis the actin cap reorganizes to form an actin shell surrounding each mitotic spindle, and thus maintaining a territorial delimitation. Such a territorial delimitation may not be essential when the nuclei first migrate to the cortex because there is ample space between nuclei, but as the cortical region becomes more crowded with nuclei such a barrier may be important to prevent mismatching of mitotic spindles and inappropriate chromosomal disjunction. Kellogg et al. (1988) first suggested that the actin shell may be important to restrict the movement of the mitotic spindle in the cortex as a result of their observations on living Drosophila embryos where the spindle can make oscillatory movements of approximately 3 |Lim. Additional support for this suggestion comes from two different lines of evidence. Firstly, Callaini et al. (1992) showed that treatment of embryos with cytochalasin, which disrupts actin filaments, caused spindle fiision between adjacent spindles. In these experiments Callaini et al. (1992) showed that cytochalasin treatment disrupted the actin cytoskeleton in all cases, however when there were fewer nuclei in the cortex (i.e., mitotic divisions 8, 9, and 10) mitotic spindles did not fuse. When cytochalasin-treated embryos were allowed to proceed to later developmental stages (i.e., mitotic divisions 11,12, and 13), with more nuclei in the cortex, the mitotic spindles fuse and karyokinesis was disrupted. The second line of evidence comes from genetic studies. Sponge is a maternal effects gene whose deletion alters some, but not all of the actin-dependent structures in the cortex of the Drosophila embryo. Embryos deficient in sponge do not exhibit the actin cap or the shell of actin that surrounds the mitotic spindle, although later they are able to organize actin filaments into the hexagonal arrays that are believed to mediate cellularization (Postner et al., 1992). Karyokinesis occurs relatively normally in sponge deficient embryos up to mitotic division 10. At subsequent divisions, however, as the cortex becomes more crowded with nuclei, spindle fusion occurs and karyokinesis is disrupted (Postner et al, 1992). It is significant that this is the same stage at which Callaini et al. (1992) observed a comparable result by treatment of embryos with cytochalasin. This suggests an essential role for the actin cap and mitotic shell between divisions 10 and 11 in maintaining the cytoplasmic space or territory.

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Elements of the cytoskeleton also have a central role in the cellularization process in Drosophila embryos. Cellularization is a two phase process in which first actin, myosin and spectrin accumulate in the cortex and form a hexagonal array along which plasma membrane invaginates, and second as the membrane furrow extends beyond the nucleus a contractile ring forms to pinch off the forming cell from the yolk mass (Figure 1; FuUilove and Jacobson, 1971; Warn and Magrath, 1983; Pesacreta et al, 1989; Schweishuth et al., 1990; Warn and Robert-Nicoud, 1990; Simpson and Wieschaus, 1990; Young et al, 1991). These changes require an extensive and relatively rapid set of alterations in the cytoskeleton that appear to be under the control of at least two zygotic genes, nullo and serendipity alpha, whose function may be partially redundant. Deficiencies in either locus result in defective cellularization and production of multinucleated cells, probably by disrupting the actin-myosin organization of the cleavage furrows (Schweisguth et al., 1990; Simpson and Wieschaus, 1990). What mechanism(s) regulate the orderly, and relatively synchronous changes in cytoskeletal organization that occur during embryogenesis in Drosophila! Reports described above indicate a series of genes that could act in succession to regulate the organization of actin filaments and accompanying developmental events. Maternal genes N26, N441, 3nd paralog appear to act early in development to regulate migration of nuclei and centrosomes to the embryo cortex (Hatanaka and Okada, 1991b). Once nuclei have migrated to the cortex another maternal gene sponge appears to regulate the actin caps and shells which delimit the cytoplasmic territory around the nucleus and mitotic apparatus, respectively. Sponge, however, does not appear to regulate earlier developmental stages where actin filaments control nuclear migration (Postner et al., 1992). Later in development the zygotic genes nullo and serendipity alpha become active and regulate actin organization in the cellularization process, but have no role at earlier developmental stages (Simpson and Wieschaus, 1990; Schweisguth et al., 1990). Thus differential gene expression could play an important role in regulating the changes in cytoskeletal organization throughout development of the embryo. How are the changes in cytoskeletal organization performed in concert throughout such a large cell? Since the early embryo is delimited by only one plasma membrane it is possible that signal transduction may have a significant role in regulating the synchrony of these events. However, with one exception, investigations have not been directed at mechanisms regulating the synchrony of these events. The exception concerns the synchronous cellularization which occurs across the embryo and which is accompanied by major changes in cell cycle control (Edgar et al., 1986; Edgar and Schubiger, 1986; O'Farrell et al, 1989). It was proposed that membrane furrows that mediate cellularization would extend by a simultaneous contraction all over the embryo (Fullilove and Jacobson, 1971; Warn and Magrath, 1983). However, this notion seems unlikely as local inactivation of the contractile network by microinjection of antimyosin antibodies (Lutz and Keihart, 1987) or phalloidin (Planques et al., 1991; Warn and Robert-Nicoud, 1990), which should disrupt the synchrony of contraction in the embryo, has no

'J&«S«IILIK--

Figure 1. (1) Myosin in the furrow canals during cellularization of Drosophila, as viewed by confoca! microscopy after labelling with anti-myosin antibodies. Earlier to later stages of cellularization are shown from top to bottom of the panel. Left column shows optical cross section of myosin pattern (arrowheads indicate major alterations in myosin pattern). Middle column shows optical en face sections at the plane of the forming furrow canal corresponding to developmental stage in left column. Right column shows corresponding furrow canal region at higher magnification. Scale bar for two left columns is 10 ^m and for the right column is 5 \xm. Reproduced from Young et al., 1991, with permission of Company of Biologists, Ltd. (2) Microtubules in the cortex of a starfish oocyte labeled by immunofluorescence. Scale bar is 10 |im reproduced from Otto and Schroeder, 1984, with permission of Academic Press. (3) Cortical granules linked to isolated cortex of sea urchin egg interconnected with 6 nm filaments. Scale bar is 2 ^im. Reproduced from Chandler, 1984, with permission of Academic Press. (4) Immunogold localization of spectrin in sea urchin egg cortex. Anti-spectrin antibody coats vesicles (arrow in a) and granules and actin in microvilli (b). Scale bars are 0.5 jim. Reproduced from Fishkind et al., 1990, with permission of Academic Press. 70

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effect on the cellularization over the rest of the embryo. Clearly more studies are needed to determine the mechanisms of cytoskeletal regulation. Few studies of the Drosophila system have described the spatial organization of intermediate filaments during embryogenesis, and some investigators have expressed skepticism that intermediate filaments exist in Drosophila (Fyrberg and Goldstein, 1990; but see Biessmann and Walter, 1989). However, intermediate filaments have been identified both in embryos and cvXtm^d Drosophila cells based on biochemical and immunological criteria. The intermediate filament network appears to be involved in the heat shock response of the embryo (Walter and Biessmann, 1984; Walter et al., 1990). Heat shock causes the intermediate filament network to collapse around the nucleus or chromosomes. The embryo is capable of recovering from this state once it reaches the developmental time when heat shock proteins can be synthesized, however prior to this time heat shock is lethal (Walter et al., 1990). Walter et al. (1990) propose that the heat shock-induced collapse of the intermediate filament network may be responsible for altering the distribution and proper translation of mRNAs responsible for the establishment of segment identity, and thus be responsible for the segmental abnormalities induced by heat shock (Eberlein, 1986). Investigations using Drosophila have the major advantage of allowing the use of mutants with different genetic backgrounds to investigate the functional significance of changes in cytoskeletal organization. It would have been anticipated that genetic screening of Drosophila eggs and embryos would have identified many mutants containing defective, or totally lacking cytoskeletal proteins. However, until recently few such mutants have been identified. This may be because such mutants are so disruptive that the embryo dies at an early stage and goes unnoticed in the screening, and/or because such mutants are not detectable as a pattern in the cuticle. Some genetic defects are now recognized as due to cytoskeletal elements or their associated proteins. First, for example, the shibire gene codes for a protein highly similar to the molecular motor protein dynamin (van der Bliek and Meyerowitz, 1991). A fly carrying a temperature sensitive shibire mutation undergoes a temperature sensitive, reversible paralysis which is thought to be due to depletions of synaptic vesicles, an event which results because membrane recycling is inhibited at the nonpermissive temperature. The identification of the shibire protein as a dynamin-like protein suggests a model wherein the dynamin-like protein is associated with the synaptic vesicles and acts as a molecular motor to mediate vesicle transport at the permissive temperature, but is inhibited at the nonpermissive temperature. Such a model allows the design of a series of experiments to determine the precise cellular mechanism(s) resulting from this genetic defect (van der Bliek and Meyerowitz, 1991). Second, the nod mutations which cause nondisjunction and chromosomal loss appear to result from a dysfunction in another molecular motor protein, kinesin (Zhang et al., 1990). Subsequent analyses have shown that there is a multi-member kinesin gene family in Drosophila (Endow and Hatsumi, 1991; Stewart et al., 1991) which may be responsible for several different devel-

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opmental abnormalities in embryos. Analysis of deletion mutants for each of these kinesin genes as well as mutants for each of the actin- and microtubule-associated proteins (after the respective gene has been identified) reported by Alberts and coworkers (Kellogg et al., 1989; Miller et al, 1989) is likely to allow determination of specific roles for a variety of cytoskeletal-associated proteins. Evidence exists which suggests a role for the cytoskeleton in the localization of developmental information which specifies the body plan in Drosophila eggs and embryos. It is well known that oocytes, eggs, and embryos of insects exhibit dynamic, nonrandom distributions of mRNAs (Capco and Jeffery, 1978, 1979; Edgar et al, 1987; Berleth et al, 1988; Wang and Lehmann, 1991). Molecular studies analyzing the structure of gene products for some of the genes involved with pattern formation such as BicaudalD show homology with cytoskeletal proteins, in this case myosin heavy chain (Wharton and Struhl, 1989). Support for the assertion that cytoskeletal elements are involved in positioning developmental information comes from studies in which pharmacological agents that act on microtubules are applied to oocytes. These studies reveal that agents which disassemble microtubules disrupt the anterior localization of bicoid mRNA (Pokrywka and Stephenson, 1991). Pharmacological agents that disrupt microtubules also disrupt the perinuclear localization of cyclin B mRNA (which participates in regulation of the embryonic cell cycle) in the embryos, however they have no effect once the posterior localization of cyclin B mRNA has been established (Raff et al, 1990). The posterior localization of cyclin B is due to the nucleotide sequence in the 3' nontranslated region of the message. Early in development (i.e., prior to stage 9 of oogenesis) a shorter cyclin B is transcribed lacking nucleotides at the 3' nontranslated region and this message is not localized. Later in development, however, a longer form of cyclin B is transcribed and sequencing reveals the additional nucleotides in the 3' nontranslated region which alters the secondary structure of the mRNA; as a result of this additional sequence the mRNA is localized to the posterior pole of the egg (Dalby and Glover, 1992). Dalby and Glover (1992) have proposed that the localization phenomena is due to the secondary structure of the cyclin B mRNA acting directly on microtubules. Oogenesis and embryogenesis in Drosophila are accompanied by many unique developmental transitions that are absent in non-insect species. Yet for each of these developmental transitions the cytoskeleton is integrally involved, for example the transfer of developmental information and cytoplasm from nurse cells to oocyte, the positioning of pattern formation information within the egg, translocation of nuclei within the embryos and cellularization, to mention only a few. A cortical cytoskeleton clearly exists within the egg and is extensively reorganized during embryogenesis. Developmental events within Drosophila embryos occur with a great deal of synchrony suggesting a cytoplasmic regulatory mechanism, but few studies have yet investigated the mechanism(s) regulating this synchrony during embryogenesis.

Cytoskeleton in Early Development B.

73 Echinoderms

Echinoderms, particularly sea urchins, have been a popular system for investigation of events related to fertilization and embryogenesis. Moreovier, the extensive cytoskeletal networks in these eggs (Figure 1) have been demonstrated to be involved in overcoming a iiumber of the problems posed by fertilization. For example, these organisms rely on the cytoskeleton for the prevention of polyspermy; the sea urchin egg undergoes a massive exocytosis reaction, releasing the contents of cortical granules to modify the extracellular matrix and form a barrier impenetrable to sperm, the fertilization envelope. As discussed below, not only does the cytoskeleton anchor the cortical granules immediately beneath the plasma membrane, but, in addition it also mitigates potentially unfavorable consequences of precocious exocytosis. An extensive cortical cytoskeletal domain containing actin filaments is present in the cortex of unfertilized star fish eggs (Otto and Schroeder, 1984) and sea urchin eggs (Spudich and Spudich, 1979; Kidd aiid Mazia, 1980; Schatten et al., 1986b; Bonder et al, 1989). In the sea urchin egg, filamentous actin in the cortex is concentrated predominantly in the numerous microvilli, as demonstrated by phalloidin staining which reveals a punctate pattern on the egg surface (Bonder et al., 1989). Fodrin (also known as spectrin), a peripheral membrane protein which associates with actin, also exhibits a punctate pattern of staining in the unfertilized egg (Schatten et al., 1986b; Henson and Begg, 1988; Bonder et al., 1989) and is associated with cortical granules, pigment granules and yolk platelets (Fishkind et al., 1990). in addition to microvilli, ultrastmctural analysis of the egg cortex reveals that short, actin-sized filaments connect the cortical granules to the plasma membrane (Figure 1; Chandler, 1984; Henson and Begg, 1988). Actin filaments also run adjacent to the plasma membrane (Henson and Begg, 1988) and the cortical granules are underlain by a layer of filamentous actin, as demonstrated by immunofluorescence (Bonder et al., 1989). The cortical granules may be immobilized within this actin network by linking with the spectrin coating the surface of the granules (Figure 1; Fishkind et al., 1990). In addition to the filamentous actin described above, the sea urchin egg cortex contains a second, non-filamentous pool of actin (Spudich and Spudich, 1979; Henson and Begg, 1988; Spudich et al., 1988; Bonder et al., 1989). Spudich ahd Spudich (1979) initially noted that although Isolated sea urchin egg cortices contained a large amount of actin, very few actinfilamentscould be seen by electron microscopy, and they proposed that much of the cortical actin was in a non-filamentoiis form. Using immunofluorescence and immunoelectron microscopy, Spudich et al. (1988) demonstrated that although actin immunolocalizes in the microvilli, the subplasma membrane region, and around the cortical granules, the actin surrounding the cortical granules is non-filamentous. Bonder et al. (1989) confirmed and extended these findings, and proposed a precise spatial map of filamentous and non-filamentous actin pools in the cortex, wherein filamentous

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Figure 2. (1) Schematic diagram illustrating the distribution of filamentous actin (A, cross-hatch), non-filamentous actin (A, closed circles) and spectrin {B, thin strands) in the cortex of sea urchin eggs. CG denotes cortical granule, AG denotes acidic vesicle. Reproduced from Bonder et al., 1989, with permission of Academic Press. (2) Immunofluorescence images of intermediate filament network in cortex of starfish oocytes, referred to as snoods. The network lies in a single plane (a) and exhibit a beaded structure (b,c). At higher magnification the network is seen to have a periodicity. Scale bars are 10 |Lim. Reproduced from Schroeder and Otto, 1991, with permission of Academic Press. (3) Schematic diagram illustrating presumed sequence of signal transduction events within sea urchin eggs at the time of fertilization. 74

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actin is located in the microvilli and in a layer beneath the cortical granules, whereas non-filamentous actin is localized around the cortical granules (Figure 2). What is the purpose of the unpolymerized pool of actin? The timing of the disappearance of the unpolymerized actin pool suggests it is needed immediately upon fertilization: Fertilization triggers a wave of exocytosis, and hence the insertion of large amounts of membrane into the plasma membrane. As a result, the membrane surface area of the egg increases dramatically (Schroeder, 1979). The egg responds to this increased amount of membrane by following the wave of exocytosis with a wave of microvillar elongation (Begg and Rebhun, 1979; Schroeder, 1979). As exocytosis proceeds, the unpolymerized pool of actin is rapidly recruited into the microvilli, which lengthen nearly 3-fold (Schroeder, 1979). The microvilli remain elongated until endocytosis retrieves sufficient membrane to return the plasma membrane surface area to normal (Schroeder, 1979), thereby mitigating the potentially unfavorable consequences of massive insertion of cortical granule membranes into the egg plasma membrane. Fertilization also triggers an increase in the amount of filamentous actin in the cortex below the plasma membrane (Spudich and Spudich, 1979). In addition, detergent extraction of sea urchin eggs and embryos has demonstrated an increased binding of poly(A)"^RNA to the cytoskeletal fraction that is associated with: (1) an increase in the number of filaments in the cortex and (2) an increase in the amount of protein synthesis in the embryo (Moon et al., 1983). Based on work conducted using somatic cells. Moon et al. (1983) suggested that association of embryonic mRNA with the cytoskeleton promotes protein synthesis. What regulated the extensive changes in actin organization? One possibility is that the actin-associated protein, villin, drives these changes based on the altered intracellular free calcium ([Ca^"^]|) that results at fertilization. Villin is present in sea urchin eggs and may be responsible for the establishment of the brush borderlike microvilli which are present on the egg at fertilization (Wang and Bonder, 1991). Wang and Bonder (1991) proposed that the fertilization-dependent rise in [Ca^"^]- induces rapid polymerization of short actin filaments from the non-filamentous actin at the cortex. At the same time villin could disrupt the plasma membraneassociated actin filaments which are thought to prevent cortical granules from contacting the plasma membrane, thus permitting cortical granule exoc5^osis. As the [Ca^"^]. level decreases, villin could then promote the assembly of actin filaments

Figure 2. (Continued) Steps indicated with "V are speculative. Reproduced from Chandler, 1991, with permission of Wiley-Liss, Inc. (4) Cortex of ascidian egg viewed as a replica after deep-etching and rotary shadowing. An extensive anastomosing network of membrane and cytoskeletal elements can be seen. PM is plasma membrane, mt is microtubule, T is tubule of endoplasmic reticulum. Scale bar is 0.3 ^m. Reproduced from Sardet et al., 1992 with permission of Company of Biologists, Ltd.

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to stabilize the long microvilli which form from addition of the cortical granule membrane to the plasma membrane (Wang and Bonder, 1991). Cortical microtubules are found in both sea urchin and starfish eggs (Schroeder and Otto, 1984; Boyle and Ernst, 1989). In sea urchin eggs, reorganization of the microtubule network serve to mediate the pronuclear migration required for syngamy. Microtubules elongate from the sperm centrosome and extend to the female pronucleus (Bestor and Schatten, 1981) and the male and female pronuclei appear to migrate along these microtubules until pronuclear fusion. Pronuclear migration is blocked by agents which disrupt microtubules (Schatten et al, 1989), further supporting the idea that a microtubule-based system of motility mediates pronuclear migration. Oocytes and eggs of starfish and sea urchins contain intermediate filaments (Boyle and Ernst, 1989; St. Pierre and Dufresne, 1990; Schroeder and Otto, 1991). In starfish an extensive meshwork of intermediate filaments is present in the cortex of the oocyte (Figure 2). This network is unusual in that it is composed of parallel bundles of intermediate filaments that fill the entire cortex, except in the animal hemisphere. In the animal hemisphere unbranched bundles of intermediate filament appear to radiate out from the region of the centrosomes and become continuous with the meshwork of intermediate filaments in the cortex. In starfish oocytes this network disappears during resumption of meiosis and does not reappear (Schroeder and Otto, 1991). The intermediate filament network also disappears as the sea urchin oocyte resumes meiotic maturation (Boyle and Ernst, 1989). Although the function of this network in the oocytes is unclear, its presence in oocytes and its disappearance during the resumption of meiosis parallel the observations reported for intermediate filaments in oocytes of the 3m.phihmn,Xenopus laevis (see below) and these filaments may serve to stabilize the cortex during the long period of oogenesis. The intense study of oocyte fertilization in sea urchins has led to many insights into the signal transduction mechanisms which govern the cytoskeletal transitions accompanying fertilization. The key signaling event in sea urchin fertilization, as in fertilization of other organisms (Jaffe, 1983), is a transient increase in ([Ca^"*"]}) (Steinhardt et al., 1977). It has been proposed that sperm penetration results in the cleavage of a membrane lipid phosphoinositol (PIP2) by phospholipase C resulting in the production of inositol trisphosphate and diacylglycerol. The former reaction product induces a rise in [Ca^'*"]^ while the latter activates protein kinase C (PKC) and these two intracellular signals initiate two different cascades of changes within the egg that mediate the fertilization events (Figure 2; for review see Chandler, 1991). The [Ca^"^]i rise propagatesfi"omthe site of sperm entry across the egg in a wave-like manner, acting as the initial stimulus for numerous morphological and biochemical events. The [C2?'^\ wave is followed rapidly by exocytosis, microvillar elongation, and an increase in intracellular pH, and each of these events can be triggered in the absence of fertilization by artificially inducing an elevation [Ca^"^]^. The precise relationship between the [Ca^"^]} wave, the rise in pH, and microvillar

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elongation has been demonstrated by studies wherein calcium was increased under conditions which prevented the pH increase, or pH was increased under conditions which did not result in a [Ca^'^jj rise (Begg and Rebhun, 1979; Begg et al, 1982; Carron and Longo, 1982). From these results it appears that the [Ca^'^]^ rise and the pH increase each play a separate role in microvillar elongation, probably regulated by the calcium- and pH-dependent protein, villin (Wang and Bonder, 1991). It is likely that the rise in [Ca^^]j also participates in the activation of the calcium- and phospholipid-dependent enzyme, PKC, which appears necessary for a variety of events including normal formation of the first mitotic apparatus and subsequent cleavage furrow; in the presence of inhibitors of PKC the mitotic apparatus and furrow are abnormal (Mabuchi and Takano-Ohmuro, 1990). Thus, the sea urchin embryo utilizes specific intracellular signals to execute particular cytoskeletal transformations. A clear correlation between signal tranducers and developmental transitions can be identified in the echinderm system. As with Drosophila, there is a highly developed cortical cytoskeletal domain which undergoes extensive reorganization as progression through the meiotic cell cycle is initiated and again at the time of fertilization. Thus again, extensive cytoskeletal reorganization occurs during key developmental transitions. Moreover, in this system it is possible to link pronuclear migration to the microtubules of the sperm aster.

IV. THE CYTOSKELETON IN EARLY DEVELOPMENT OF CHORDATES A.

Ascidians

Developmental specializations of the cytoskeleton seen in lower organisms are also apparent in chordates particularly with regard to the cortical cytoskeleton, whose role appears to be conserved between chordates and nonchordates. One of the most extensively studied cortical cytoskeletal domains is that of the ascidian, Styela. This organism was first used for investigation of cytoskeletal rearrangement and localization of developmental information, as expressed by mRNA localizations, by William Jeffery and coworkers. This system has features which make it ideal for such studies. First, the eggs are transparent and have differentially colored components associated with different egg regions, making it possible to follow the fate of the different egg regions in living embryos by observing the redistribution of the colored components during embryogenesis. Second, Styela has three obvious egg cytoplasmic regions, each of which has a specific developmental fate: Ectoplasm, a clear region originating from the oocyte germinal vesicle, accumulates in ectodermal cells. Myoplasm, a region with yellow pigment granules, accumulates in muscle and mesenchyme cells. Endoplasm, a white region containing yolk, accumulates in endodermal cells.

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Each region of Styela oocytes and embryos exhibit characteristic patterns of mRNA distribution, as might be expected given their different developmental fates. This was demonstrated by in situ hybridization of nucleic acid probes to histological sections. In the full grown oocyte, poly(A)"^RNA is enriched in the germinal vesicle (Jeflfery and Capco, 1978). In contrast, actin mRNA is found in the yellow cj^oplasm of the egg cortex and the germinal vesicle, whereas histone mRNA is apportioned equally to all regions of the cytoplasm (Jeflfery et al., 1983). Fertilization initiates a dramatic redistribution of these cytoplasmic regions and their constituent RNAs. The myoplasm and its associated actin mRNA redistributes to the vegetal hemisphere cortex. Both then move to the subequatorial region of the zygote and form the yellow crescent, which later gives rise to the posterior pole. The ectoplasm becomes enriched in poly(A)"^RNA and, subsequently, the poly(A)"^RNA(like the ectoplasm) is predominantly distributed to ectodermal cells (Jeflfery et al., 1983). Histone mRNA, in contrast, remains uniformly distributed and is equally partitioned into the blastomeres. What mechanism is responsible for the localization and subsequent segregation of the different regions of the cytoplasm and their associated RNAs? Several lines of evidence indicate that the cytoskeleton controls both localization and redistribution of Styela cytoplasm and RNAs. First, detergent-extracted Styela eggs retain the bulk of their mRNA on the cytoskeleton (Jeflfery, 1984). Second, the spatial distribution of mRNA in detergent-extracted eggs is virtually the same as in unextracted eggs (Jeflfery, 1984). Third, stratification of eggs by centrifugation results in distinct bands corresponding to the ectoplasm, endoplasm and myoplasm, and in situ hybridization reveals that mRNA stays in the same region it was located in before stratification (Jeflfery, 1984). Fourth, electron microscopy and biochemical analysis of detergent-extracted eggs and embryos reveals the presence of a filamentous network in the cortex which contains actin and intermediate filament proteins and which co-localizes with the yellow pigment and mRNA of the myoplasm (Jeflfery and Meier, 1983, 1984; Jeffery, 1985). Fifth, as a response to fertilization, the cortical cytoskeletal domain undergoes contraction and migration to the vegetal cortex, and subsequently to the region of the yellow crescent, following exactly the pattern of reorganization undergone by the myoplasm in normal development (Jeffery and Meier, 1983, 1984). The above studies demonstrate that: (1) RNA and protein components of the different plasms are tightly associated with elements in the cell, (2) these elements remain behind after detergent extraction and, (3) RNA and other cytoplasmic components move in temporal and spatial concert with embryonic cytoskeletal domains. Taken together, these investigations clearly demonstrate the role of the cytoskeleton in localization and segregation of cytoplasmic components during Styela development. Moreover, the fact that the different domains maintained and transported by the cytoskeleton have distinct developmentalflatesfurther illustrates how embryonic patterning events are dependent on the cytoskeleton.

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The myoplasm which becomes localized by the cortical cytoskeletal domain as a response to fertilization becomes concentrated in the larval tail muscle cells during embryonic development. Using immunochemical methods Swalla et al. (1991) have identified a protein localized in the myoplasm which binds to the cytoskeleton and accumulates in the larval tail muscle cells. Surprisingly, when the distribution of this protein was examined in ascidians which do not form tails it was found not to localize to the myoplasm and not to accumulate in cells of muscle lineage (Swalla et al., 1991). Subsequent studies have shown that this protein, as well as the paternal genome, are both essential for a partial rescue of the larval tail when eggs from the tailless species are fertilized (Jeffery and Swalla, 1992). Thus, this cytoskeletonassociated protein, which was part of the cortical cytoskeleton of the egg, plays an essential role in the developmental processes which regulate tail formation in ascidians. In addition to their cortical actin filament network, an elaborate network of microtubules are present in ascidian eggs as part of the meiotic asters and as a microtubule array filling the egg interior. Using immunofluorescence, Sawada and Schatten (1988) demonstrated a network of microtubules which extended throughout the cytoplasm. As a response to fertilization this network is reorganized, resulting in a microtubule array projecting from a centrosome located in the myoplasm at the vegetal pole. The precise role of this network is unknown, but it is thought to be required for migration and syngamy of the male and female pronuclei. The cortical cytoskeleton of unfertilized eggs exhibits an animal-vegetal polarity with an enrichment of actin filaments and possibly intermediate filaments in the vegetal cortex (Figure 2; Sardet et al., 1992). However microtubules are excluded from the cortex on the vegetal side of the egg until after fertilization (Sawada and Schatten, 1988; Sardet et al., 1992). The large centrosome which forms at the vegetal pole as a response to fertilization (Sawada and Schatten, 1988) may serve the same function of the centrosomes which arrive in the cortex of the Drosophila embryo, that is they may serve to reorganize the cortical cytoskeleton. In support of this contention, immediately after the centrosome forms, the cortical cytoskeletal domain of the myoplasm moves to the vegetal pole. Subsequently, the centrosome is translocated to the equatorial region and again the myoplasm with its associated cytoskeletal components, mRNAs, and developmental determinants accumulates at the new location for the centrosome. From the available evidence it is not possible to determine whether the centrosome is responsible for the repositioning of the myoplasm or is simply carried along with the myoplasm by some other force. Additional studies which employ microtubule-stabilizing or destabilizing agents could help to define the active force, as could microinjection of additional centrosomes which might serve to subdivide the myoplasm into additional cytoplasmic domains. The means by which ascidians regulate these cytoskeletal transitions are not clear. However, preliminary evidence suggests that a rise in [Ca^"^]-, triggered by fertilization (Brownlee and Dale, 1990; Speksnijder et al., 1990), may be an important

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signal. For example, Jeffery (1982) demonstrated that translocation of myoplasm to the vegetal pole of Styela could be perturbed by placing eggs next to a rod treated with a calcium ionophore; under these conditions the myoplasm always migrated toward the side of the egg closest to the rod, without respect to the animal-vegetal axis (Jeffery, 1982). Support for the notion that elevation of [Ca^'^Jj directs the migration of the myoplasm comes from Speksnijder (1992) who showed that ascidian eggs produce repetitive calcium transients after fertilization and that the majority of transients are initiated at the vegetal pole, the same site where the myoplasm initially accumulates during unperturbed development. The ability of the vegetal pole to generate these calcium transients may be due to the extensive network of endoplasmic reticulum (which contains a calsequestrin-like protein) at the vegetal pole (Sardet et al., 1992). Moreover, Bates and Jeffery (1988) showed that treatment of egg fragments with calcium ionophore disrupts normal patterns of myoplasm migration, suggesting that (1) the normal trigger for myoplasm movement was a rise in [Ca^"^] j and, (2) in unperturbed development, an endogenous gradient of [Ca^"^]! regulates migration of the myoplasm to the vegetal pole. In summary, in response to fertilization signal transduction events appear to be temporally and spatially associated with extensive reorganizations of the cytoskeleton. These cytoskeletal reorganizations, while using components which can be found in somatic cells, are quite different from those typically found to occur in somatic cells, and in many cases are associated with developmental events such as the localization and positioning of the developmental determinants for the larval tail. As will be described later for mammals and amphibians, sperm penetration initiates a cascade of intracellular signals in the egg involving changes in [Ca^"^]j (Speksnijder, 1992) and the enzyme, protein kinase C (Yokosawa et al, 1989), and it is likely that these signals, in contrast to differential gene expression, act in concert to induce rapid reorganization of the cytoskeleton. B. Mammals

Early development in mammalian embryos serves as an excellent example of a system that contains cytoskeletal specializations which apparently exist to complement developmental specializations. As with other systems described in this chapter, fertilization of mammalian eggs initiates the program of early development. However, unlike the other developmental systems, mammalian embryos undergo two unique developmental transitions which do not have counterparts in nonmammalian development, namely embryonic compaction and blastocyst formation. Embryonic compaction occurs between the 8 and 32 cell stage and is a time when the embryo acquires two types of polarity. First, the entire embryo acquires an inside-outside developmental polarity with the outside blastomeres fated to form the trophectoderm and the inside blastomeres fated to form the embryo proper. In addition, the outside blastomeres acquire a cellular polarity mediated by the formation of tight junctions, repositioning of cadherins and accompanied by

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differential distribution of ion channels, microvilli, and nuclear position along the apical to basolateral axis. The second unique developmental transition is the formation of the blastocyst, at which time, prior to gastrulation, a differentiated epithelium forms as part of the embryo, that is the trophectoderm. Although all three cytoskeletal networks are present in mammalian systems their eggs also contain unique cytoskeletal elements, which have been termed sheets, that undergo dramatic reorganizations simultaneous with fertilization, embryonic compaction and blastocyst formation in ways which suggest that the sheets are integrally involved with these transitions. The sheets form an extensive cytoskeletal network in eggs of all mammals which have been examined to date (e.g., eggs of cows, dogs, hamsters, humans, mice, pigs, rats and rhesus monkeys; Hope, 1965; Gallicano et al, 1992; Capco, unpublished data for the dog). These cytoskeletal sheets can be observed using conventionally fixed material embedded in the standard plastic resins used for electron microscopy (Hadek, 1966; Weakly, 1966; Schlafke and Enders, 1967; Koehler et al., 1985), but are easier to detect after extraction in a medium containing a non-ionic detergent which removes the soluble components and leaves behind insoluble cytoskeletal elements (Capco et al., 1982; Capco and McGaughey, 1986). When such specimens are viewed as relatively thick, embedment-free sections (Capco and McGaughey, 1986; McGaughey and Capco, 1989; GaUicano et al, 1991), they provide many spatial cues which allow observation of changes in spatial organization. In fact, it was the serendipitous use of embedment-free sections prepared in a removable embedding medium such as Pentament (Capco, 1993) which allowed detection of the extensive changes in spatial organization exhibited by the sheets. Other investigators who observe the sheets in intact eggs and in plastic-embedded thin sections required for conventional transmission electron microscopy (Hadek, 1966; Weakly, 1966; Schlafke and Enders, 1967; Koehler et al., 1985) could not detect the extensive changes in spatial organization during development and consequently no importance was assigned to these structures and they were largely ignored until recently. The changes that occur in the spatial organization of the cytoskeletal sheets during early development in the mouse (Gallicano et al., 1991) are illustrated in Figure 3. In unfertilized mouse eggs the sheets are excluded from the cortex of the egg by a thin filament network (Figure 3). By the time of embryonic compaction the sheets form long concourses and contact the plasma membrane in outer blastomeres (Figure 3). At the blastocyst stage the sheets splay apart into individual filaments with a diameter of 10 nm in the outer blastomeres which form the first epithelium, the trophectoderm. However, the sheets remain present in the pluripotent inner cell mass cells which will form the embryo (Figure 3). The changes in spatial organization of the sheets during embryogenesis of the hamster are similar to those in the mouse except that the sheets in hamster eggs are in a whorled configuration in the unfertilized egg and exhibit a more linear configuration as a response to fertilization (Capco and McGaughey, 1986). In addition, the sheets in

MOUSE SHEET 82

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hamster eggs are large, solid-looking planar structures instead of the bundles of fibers which are observed in mouse eggs. We have compared the cytoskeletal sheets in eggs from different species and have found that the sheets exist in two basic forms; the sheets in eggs of mice, humans, cows and pigs are bundles of fibers (Figure 3), whereas the sheets in hamster and rat eggs are the large solid-looking planar structures (Figure 3; Gallicano et al., 1992). While observation of the sheet surface topography from these different species suggests that they represent very different structures, three lines of evidence suggest that the sheets in different species are comparable. First, both forms of the sheets undergo virtually identical changes in spatial organization at fertilization, at compaction, and again as the blastocyst forms. In addition, both forms of the sheets splay into 10 nm filaments in blastomeres of compacted embryos (Capco et al., 1993; Gallicano et al., 1993), and completely disassemble into 10 nm filaments in the trophectoderm cells of the blastocyst (Capco and McGaughey, 1986; Gallicano etal., 1991). Second, and more significantly, examination of the substructural organization of the sheets demonstrates a high degree of similarity between hamster and mouse (Figure 3). In both forms of the sheets, 10 nm filaments are linked with crossbridges every 20 nm. However, in the mouse egg sheets contain only one layer of crosslinked filaments and can roll into a cylinder (Gallicano et al., 1993), whereas in hamster eggs sheets contain two layers of crosslinked filament immobilized by an additional type of crossbridge (i.e., vertical crossbridges), and consequently are planar structures (Capco et al., 1993). The 10 nm filaments in both forms of the sheets are coated with a particulate protein which can be removed by differential extraction with detergents or salts and characterized with SDS-polyacrylamide gel electrophoresis (McGaughey and Capco, 1989; Gallicano et al., 1991, 1993; Capco et al., 1993). The coating with particulate protein masks the 10 nm filaments giving the sheets a solid appearance in hamster eggs and a fibrous appearance in mouse eggs, and

Figure 3. Cytoskeletal sheets in mouse eggs and embryos viewed as embedment-free sections after detergent extraction. (1) Unfertilized egg; arrow points to sheet, Z is zona pellucida. Scale bar is 1 fim. (2) Four blastomeres of a compaction stage embryo are shown. At this magnification sheets appear as rod-like structures present in the cytoplasm. Scale bar is 10 jam. (3) Blastocyst stage embryo showing blastomere of trophectoderm (T) and inner cell mass (I). Sheets are present in inner cell mass cells (arrow), but sheets disassemble into intermediate filaments in trophectoderm cells (arrowheads). Scale bar is 1 jim. Figure 3-1 to 3-3 are reproduced from Gallicano et al., 1991, with permission of Wiley-Liss. (4) View of fibrous sheet typical of mouse eggs. Scale bar is 0.1 ^m. Gallicano and Capco, unpublished. (5) View of solid planar sheet typical of hamster egg. Scale bar is 0.5 fim. Gallicano and Capco, unpublished. (6) Model depicting structural differences between the solid planar sheet in hamster eggs (A) and the fibrous cylindrical sheet of mouse eggs (B). Reproduced from Capco et al., 1993, with permission of Wiley-Liss.

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masks the antigenic sites on the 10 nm filaments. Third, after displacement of the particulate protein coating the 10 nm filaments, the filaments bind antibodies to keratin, but not antibodies to tubulin or vimentin (Capco et al., 1993; Gallicano et al., 1993). These results support the model that the sheets are composed of cytokeratin intermediate filaments and the major difference in surface topography is caused by the absence of one type of crossbridging element in the form of sheets present in mouse eggs (i.e., Figure 3, vertical crossbridges). In the absence of vertical crossbridges, the sheets in mouse eggs are composed of only one layer of intermediate filaments which can roll into a cylinder. The presence of vertical crossbridge in sheets of hamster eggs links two layers of intermediate filaments together resulting a more planar structure. Recognition that assembled intermediate filaments are highly organized within sheets and are also masked with protein should put to rest the controversy as to when intermediate filaments appear during mammalian embryogenesis. Different researchers have reported that (in the mouse) intermediate filaments first appear at times during early development rangingfi-omthe oocyte (Lehtonen et al., 1983) to post-compaction (Jackson et al., 1980; Paulin et al., 1980). The investigators who observed intermediate filaments early in embryogenesis employed more stringent permeabilization approaches (Lehtonen et al., 1983) to allow antibody penetration and are likely to have disrupted the protein coating the sheets, thereby exposing the intermediate filaments. Indeed, we have found that, in mouse eggs and embryos, the use of Triton X-100 as a permeabilizing agent disrupts much of the cytoskeleton and it is necessary to carry out the extraction using Tween-20 in order to maintain the integrity of the cytoskeleton (Mutchler et al., 1988; Gallicano et al, 1991). In mouse embryos we have observed that the sheets begin to splay apart into intermediate filaments at compaction (Gallicano et al., 1991), the stage at which some investigators first observed cytokeratin filaments using immunofluorescent techniques. This observation has implications for the study by Emerson (1988) which tested whether intermediate filaments were required for events in early embryogenesis. Emerson (1988) injected anti-cytokeratin antibodies into early embryos and demonstrated that the embryos could continue to develop normally through the blastocyst stage, suggesting that intermediate filaments were not necessary for the developmental events required for blastocyst formation. However, since antibodies to intermediate filaments will not bind filaments within the sheets, this study does not rule out that, as a part of the sheets, intermediate filaments have an essential role during development. A structure comparable to the sheets exists during M-phase (mitosis) of the cell cycle in some somatic cell types. During M-phase the intermediate filament network forms lamellar and globular structures (Franke et al., 1982; Chou et al, 1989) driven, perhaps, by increased phosphorylation of intermediate filament proteins which is known to occur at this time (Evans, 1989). The oocyte remains at the interphase/meiosis boundary for an extended period of time, thus, it is possible that this delay results in extensive intermediate filament phosphorylation.

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causing reorganization of intermediate filaments into lamellar structures (i.e., the sheets). However, in contrast to the case for M-phase cells the sheets do not disappear as the blastomeres enter interphase, therefore their presence is not simply due to the action of M-phase kinases. Webster and McGaughey (1988, 1990), have investigated reorganization in the cortical cytoskeleton of hamster eggs by preparing isolated cortical lawns from eggs during fertilization. Ultrastructural examination of the thin filament network in the egg periphery not only revealed an extensive thin filament network excluding the cytoskeletal sheets, but also demonstrated that the cortical thin filament network possesses two domains (Webster and McGaughey, 1988,1990). The first is a loose filament network (LN) consisting largely of actin filaments, while the second is a localized dense network composed largely of non-actin filaments (Figure 4). Upon sperm penetration the loose network forms physical associations with the sperm head and appears to mediate sperm entry. This work confirms and extends previous studies which demonstrated an involvement of actin-associated structures (e.g., microvilli and the fertilization cone) at the time of sperm penetration (Longo and Chen, 1985; Maro et al., 1984). Although actin filaments appear to be involved in sperm ingression they are not essential for sperm penetration as it will occur even in the presence of actin filament inhibitors, which reduce or inhibit the formation of the fertilization cone (Maro et al, 1984). At compacfion the actin cytoskeleton undergoes further modification to form caps in the apical region of blastomeres as they polarize (Ducibella et al., 1977). An additional reorganization, revealed by immunofluorescent analysis with anti-acfin antibodies, occurs at the blastocyst stage. A diffuse fluorescent pattern exists in the inner cell mass cells, whereas the trophectoderm shows a concentration of actin in the peripheral region of each blastomere (Lehtonen and Badley, 1980). As the mitotic apparatus forms, the amount of actin in the cortex decreases and, upon entry into interphase, actin again becomes concentrated in the cortex. This pattern of actin reorganization suggests a dependence on the cell cycle (Ducibella et al., 1977). Myosin also undergoes changes in distribution during this actin reorganization. At compaction, myosin moves from the areas of contact between blastomeres to regions occupied by actin (Sobel 1983a, 1983b, 1984). Vinculin, an acfin-binding protein, also accumulates near the termination sites of actin and appears first in precompacted embryos. Vinculin staining disappears at compaction and then reappears in the trophectoderm at blastomere contact sites (Lehtonen et al, 1988). Other acfin-associated, submembrane skeleton proteins such as fodrin and alpha-actinin also alter their spatial organizafion during early development (Lehtonen and Badley, 1980; Lehtonen et al, 1988; Sobel and Alliegro, 1985; 1986; Damjanov et al., 1986). Sobel and Szczesny (1991) have examined isolated cortices in hamster embryos before and after compaction. They have shown that before embryonic compaction the submembrane skeleton in hamster embryos resembles a lacy filamentous network, but that after compaction the organization of the submembrane skeleton changes into a thin mat of densely woven filaments. Moreover, they demonstrate that the

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cytoskeletal sheets, which they refer to as striated filament bundles, also change their organization between the two and eight-cell stages of embryogenesis. In the 2 cell embryo the sheets are found in association with globular membrane components, while this association was not detected in the eight-cell embryo (Sobel and Szczesny, 1991). In mouse eggs, at the time of fertilization, microtubules appear to regulate pronuclear migration as they do in the oocytes of other species described in this review. The microtubules (Figure 4) extend from microtubule organizing centers (MTOC) and contact the pronuclei (Schatten et al., 1985,1986a; Albertini, 1992). In other developmental systems, MTOCs appear to be derived from the sperm; however, in the mouse the MTOCs are maternally derived (Schatten et al., 1985, 1986a, 1991). In interphase of the pre-compacted mouse embryo, tubulin is

Figure 4, (1) Replica of isolated cortical cytoskeleton in unfertilized hamster egg composed of actin filaments and short crosslinking filannents (arrowheads). The localized dense network at the center of the field of view is surrounded in the loose network. Scale bar is 0.5 jim. Reproduced from Webster and McGaughey 1990. with permission of Academic Press. (2) Microtubule and actin filament distribution in rodent oocytes and eggs during meiotic maturation. (A) Microtubules in preantral follicle stage oocyte, (B) oocyte in large antral follicle, and (C) late prometaphase I oocyte. (D) shows a taxol treated metaphase I oocyte showing taxol-inducible microtubule organizing centers. (E) is same oocyte as in "D" except stained to show actin filaments. (F) shows the tubulin distribution in a metaphase II egg while (G) and (H) show the taxol-inducible microtubule organizing centers and actin filaments, respectively in the same metaphase II egg. Scale bar is 10 ^im. Reproduced from Albertini, 1992, with permission of the ISCU Press.

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uniformly distributed throughout each blastomere; however, during mitosis, tubulin assembles into the mitotic apparatus. As the cleaving blastomeres re-enter interphase, microtubules reassemble near areas of cell contact (Ducibella et al., 1977; Lehtonen and Badley, 1980; Maro et al., 1991). The microtubule reorganization is accompanied by redistribution of mitochondria and lipid droplets (Ducibella et al., 1977; Wiley and Eglitis, 1980), suggesting that microtubules may be responsible for positioning these and other organelles in the blastomeres. This is plausible as investigations employing somatic cells have shown that organelles, even nuclei in fungi and yeast, can be translocated along microtubules with the aid of kinesin, a molecular motor protein (Enos and Morris, 1990; Meluh and Rose, 1990). Microtubule distribution in the blastocyst has been described, using immunofluorescence, by Lehtonen and Badley (1980) who showed that tubulin was uniformly distributed throughout each blastomere in the inner cell mass and localized in the perinuclear region in trophectoderm cells. Interestingly, a discrepancy has arisen as to the exact distribution of microtubules prior to blastulation. Lehtonen and Badley (1980) reported an enrichment of microtubules in the areas of cell contact while Houliston et al. (1987) found a uniform distribution of microtubules at the same stage of development. This discrepancy may have occurred because of the two very different techniques used to visualize microtubules. Houliston et al. (1987) extracted isolated blastomeres with Triton X-100 whereas Lehtonen and Badley (1980) did not use detergent extraction, nor did the latter investigators employ isolated blastomeres. We have found that the presence of either Triton X-100 or NP-40 in extraction buffers disrupts the spatial integrity of the cytoskeleton of mouse eggs (Mutchler et al., 1988; Gallicano et al., 1991) whereas another detergent, Tween-20, allows extraction of soluble components of the cell while maintaining cytoskeletal integrity. In addition, the disassociation of embryos into isolated blastomeres or blastomere aggregates may have an affect on microtubule organization. Although much less work has been conducted concerning the role of intracellular signals in the mediation of these mammalian developmental transitions compared to sea urchin and amphibian systems, evidence indicates a role for calcium and PKC at fertilization and compaction. Treatments that increase intracellular free calcium in unfertilized eggs have been shown to induce egg activation (Steinhardt et al., 1974; Cuthbertson and Cobbold, 1985; Tombes et al., 1992), as have PKC agonists (Miyazaki et al., 1986; Endo et al, 1987; Colonna et al., 1989). More recently we have demonstrated that PKC acts downstream of the fertilization-induced rise in [Ca^"*']i to mediate formation of the second polar body and the transit to first mitotic interphase, while calcium is necessary for chromosomal disjunction at the release from the arrest at meiotic metaphase II (Gallicano et al., 1993). In addition, compaction stage embryos have been shown to have a requirement for extracellular fi-ee calcium (Pratt et al., 1982; Goodall, 1986), whereas increased PKC activity has been shown to induce premature compaction while inhibition of this activity induces decompaction (Winkle et al, 1990). Similarly, PKC activity has been

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shown to induce cytoskeletal reorganization (Bloom, 1989) and evidence suggests that PKC may regulate mammalian embryonic cytokinesis (Mystkowska and Sawicki, 1987). Thus it is possible that PKC mediates many of the changes in cytoskeletal organization during early development including alterations in the spatial organization of the cytoskeletal sheets which occur during the time that PKC is reported to be active (see above). There is also evidence to suggest that phosphorylation and cell cycle regulatory molecules affect the ability of the centrosome to nucleate microtubules and influence the distribution of microtubules during early mammalian embryogenesis (Mattson and Albertini, 1990; Messinger and Albertini, 1991; Plancha and Albertini, 1992). In support of this it has been shown that microtubule assembly in activated mouse eggs is essential for progression from meiosis to first mitotic interphase (Schatten et al, 1989). The cytoskeletal reorganizations which occur in mammalian embryos provide further support for the notion that cytoskeletal domains exist in embryos and that these are reorganized at key developmental transitions in early development. Even the cytoskeletal sheet domain unique to mammalian eggs is subject to these reorganizations. Moreover, evidence further suggests that intracellular signals, in particular, PKC, regulate these global changes in cytoskeletal organization. Future studies linking intracellular signals to cytoskeletal reorganizations may provide insight into mechanisms regulating these changes in other classes of organisms. C.

Amphibians

Amphibian oocytes, eggs, and zygotes, particularly those of the frog Xenopus laevis, have been extremely useful for study of many developmental and cellular phenomena, including cytoskeletal dynamics. In this developmental system actin filaments, intermediate filaments, and microtubules undergo extensive reorganizations at key transitions both during Xenopus oogenesis and early development (Ryabova, 1988; Bement and Capco, 1990a,c). Some of these cytoskeletal reorganizations are crucial for the processes of fertilization, cytokinesis, and axis determination, while others result from specializations to allow the egg to cope with its large size. It is likely that many, if not all, of these cytoskeletal reorganizations are controlled by intracellular signaling pathways, allowing rapid, transient regulation of complex cytoskeletal events (Bement and Capco, 1990c; Bement, 1992). Comparable to that described for other organisms, amphibian oocytes and eggs have a cortical cytoskeletal domain which undergoes extensive changes in structure and composition during oogenesis and early development. Larabell (1993) has recently reported a technique for gentle isolation of the cortex and cortical cytoskeleton ofXenopus eggs. This manipulation can be conducted under physiological conditions and allows for rapid exchange of the ionic milieu without damage or distortion to the cortex. Using this technique Larabell (1993) has demonstrated an intricate submembrane skeleton linked to a more internal network by long filaments (Figure 5). These internal cytoskeletal filaments are connected to cortical

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Figure 5, Cortical lawn prepared from Xenopus egg. Gentle preparation of the cortex reveals both the submembrane cytoskeleton (SMC) and a more internal network {arrows) interlinking cortical granules (C), pigment granules (P), and membrane cisternae (M). Scale bar is 0.5 |Lim. Larabell, unpublished.

granules, endoplasmic reticulum, and pigment granules and probably have a role in the positioning of these components. This view^ of the cortex bears similarity to the two-component cytoskeletal system proposed by Ryabova (1982, 1988; see below). It is anticipated that this method of preparing and manipulating the cortex will be of great use in evaluating the regulatory mechanisms which affect the cortical cytoskeleton. Actin filaments have been demonstrated in the cortex of the amphibian oocytes and eggs by electron microscopy (Franke et al, 1976; Ryabova, 1982, 1990a), immunofluorescence (Colombo et al., 1981) and immunoelectron microscopy

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(Gall et al., 1983). Although both oocytes and eggs have cortical actin filaments, in oocytes many of the filaments appear to be within microvilli, and the actin filament network underlying the plasma membrane is relatively sparse. In contrast, in eggs the microvilli are greatly reduced in size (Charbonneau and Grey, 1984; Bement and Capco, 1989a) but the filament network beneath the plasma membrane is much more dense (Ryabova, 1982). Ryabova (1982, 1988, 1990a, 1990b) has suggested that a two-component cytoskeletal system composed largely of actin filaments exists in the cortex of oocytes and eggs. The first component of this system is a sparse actin filament network immediately beneath the plasma membrane and is devoid of organelles. The second cytoskeletal component is also composed of actin filaments and is immediately adjacent in the subcortex. This second network appears to contain both pigment and cortical granules. The structural differences in these networks have been detected by electron microscopy and assays have shown a differential sensitivity to calcium ions and cytochalasin B as well as changes in contractile activity during resumption of meiosis (Ryabova, 1982, 1988, 1990a, 1990b). The reorganization of actin filaments during the resumption of meiosis is accompanied by the reorganization of the actin-binding protein vinculin (Evans et al., 1990). In the oocyte vinculin is present only in the interior, but in the fertilization-competent egg vinculin is present in the cortex where it may associate with, and bind, actin filaments into a network capable of contractile activity (see below). Fertilization initiates a wave of cortical granule exocytosis (Grey et al., 1974), microvillar enlargement (Ezzell et al., 1985), and contraction of the egg cortex (i.e., cortical contraction), such that the pigmented surface of the animal hemisphere appears reduced in size with respect to the vegetal hemisphere surface (Wolf, 1974). Microvillar elongation allows the egg plasma membrane to cope with the insertion of cortical granule membrane, as it does in sea urchins. Cortical contraction is followed by relaxation of the cortex, and subsequently, in synchrony with each cleavage cycle, the egg cortex contracts and relaxes again (Hara et al., 1980). The initial contraction may be necessary for emission of the second polar body and evidence suggests that it helps position the sperm nucleus near to the site where the female pronucleus will form (Elinson, 1977). The subsequent contractions are thought to be required for cytokinesis (Hara et al, 1980), a presumption confirmed by our observation that eggs which fail to contract also fail to form cleavage furrows (Bement and Capco, 1990b). Microvillar enlargement and cortical contraction are apparently mediated by interaction of the cortical actin filament network with myosin since (1) they are sensitive to N-ethylmelamide-modified heavy meromyosin (Ezzell et al., 1985) and (2) depletion of myosin from eggs sliced in half prevents contraction, whereas readdition of myosin restores contractility (Christensen et al, 1984). Intermediate filaments are also present in amphibian oocytes, and, similar to other cytoskeletal networks, undergo reorganization as the oocyte becomes the egg, and again as the egg becomes the zygote. Cytokeratin filaments were first identified

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inXenopus oocytes by immunofluorescence and immunoelectron microscopy (Gall et al., 1983; Franz et al, 1983; Godsave et al, 1984a). The clearest picture of the oocyte cytokeratin network has been provided by Klymkowsky et al. (1987) by wholemount immunofluorescence. These studies demonstrated the presence of an asymmetric, highly organized "geodesic" cytokeratin network which extends throughout the vegetal hemisphere cortex. Another intermediate filament, vimentin, is also present in the Xenopus oocyte. In the oocyte animal hemisphere, immunofluorescence reveals that vimentin is present in mitochondria-filled corridors of cytoplasm, while in the vegetal hemisphere it exhibits a punctate pattern (Godsave et al., 1984b; Tang et al., 1988; Torpey et al, 1992; but see Herrmann, 1989, Dent et al., 1992). The cytokeratin and vimentin networks are not colocalized, instead cytokeratin filaments occupy a more cortical position, while vimentin filaments lie more internally (Torpey et al, 1992). This lends further support to the notion that a two-domain cytoskeletal system exists within Xenopus oocytes as was proposed by Ryabova (1982, 1988, 1990a). The oocyte is converted into the fertilization-competent egg during the meiotic resumption. During this time virtually all of the cytokeratin network disappears (Godsave et al., 1984b; Gall and Karsenti, 1987; Klymkowsky et al., 1987), leaving a few residual cytokeratin filaments in the egg cortex (Klymkowsky et al., 1987). The disruption of the cytokeratin network appears to be regulated by intracellular signals (see below) and is caused by a phosphorylation-induced fragmentation of the cytokeratin network into oligomers (Klymkowsky et al., 1991). Vimentin distribution also changes, becoming dispersed uniformly throughout the egg (Godsave et al, 1984b), however, phosphorylation does not appear to regulate this change in organization and instead evidence suggests that the change in organization may be due to the inactivation of an inhibitory facto (Dent et al., 1992). Fertilization triggers a further reorganization of the cytokeratin network. A cytokeratin network reforms, but it is again asymmetrically organized between the animal and vegetal hemispheres. In the animal hemisphere a sparse network of filaments form, whereas in the vegetal hemisphere the distribution of cytokeratin is initially punctate followed by the appearance of first bundles which then form an interlinked system of filaments. The cytokeratin networks maintain this asymmetry until the early blastula stage. Unlike cytokeratin, vimentin remains randomly distributed after fertilization (Godsave et al., 1984b; Tang et al., 1988). The animal and vegetal hemispheres differ both in the form (i.e., filamentous versus nonfilamentous) and amount of the two intermediate filament proteins they contain, and it is reasonable to assume that the different blastomeres derived from each hemisphere will likewise differ in their intermediate filament composition. This difference may well contribute to the different developmental fates of the blastomeres derived from the animal and vegetal hemispheres. Study of microtubules in amphibian oocytes, eggs, and zygotes has revealed similarities to somatic microtubules, as well as a number of specializations of this cytoskeletal component that are entirely unanticipated from somatic cell studies.

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In the Xenopus oocyte, immunofluorescence and electron microscopy have shown that microtubules are found both in the cortex (Dumont and Wallace, 1972; Huchon et al., 1988; Houhston and Elinson, 1992) and in association with the germinal vesicle (Jessus et al., 1986), a distribution similar to that seen in other cell types (Brinkley, 1985). Oocyte microtubules may help position the germinal vesicle within the cytoplasm, since treatment of oocytes with drugs inducing microtubule disassembly induces displacement of the germinal vesicle toward the oocyte cortex (Lessman, 1987; although see Lessman and Kessel, 1992 for a possible role for intermediate filaments in nuclear positioning). Meiotic maturation initiates an extensive reorganizations of the oocyte microtubule network. While the nuclear envelope of the oocyte germinal vesicle disrupts, a prominent microtubule network becomes visible (by light microscopy and immunofluorescence) at the vegetal side of the disintegrating germinal vesicle (Huchon et al., 1981; Jessus et al., 1986; Gard, 1992). As meiotic maturation progresses, the network moves to the animal pole, and upon assembly of the first meiotic spindle, the cytoplasmic network disappears (Huchon et al., 1981) perhaps positioned by actin filaments (Ryabova et al., 1986). Curiously, in the meiotically mature egg, which is arrested at metaphase II, microtubules are present both in the cytoplasm (Elinson, 1983; Huchon et al., 1988; Elinson and Houliston, 1990) and in the spindle, a situation quite different from somatic cells (Brinkley, 1985). Fertilization results in an initial decrease in the level of polymerized tubulin (Elinson, 1985) followed by an explosive increase (Stewart-Savage and Grey, 1982; Gard and Kirschner, 1987a, 1987b; Elinson and Rowning, 1988). The microtubule network that forms at this time has two distinct tasks (Houliston and Elinson, 1991a; Schroeder and Gard, 1992). First, microtubules from the sperm aster elongate dramatically, ultimately contacting the female pronucleus and accomplishing pronuclear migration (Stewart-Savage and Grey, 1982). The sperm aster microtubules elongate at 12 |Lim per minute (Stewart-Savage and Grey, 1982), a rate 10-fold faster than somatic cell microtubules (Gard and Kirschner, 1987a). This rapid elongation is required to overcome the long distances travelled by the male and female pronuclei through the relatively vast cytoplasm of the egg; if the microtubules elongated at the rate of somatic cell microtubules, pronuclear migration would take 5 hours instead of 30 minutes. Second, as the microtubules extend from the sperm aster and contact the cortex they bend and from a subcortical array of microtubules in both vegetal and animal hemispheres (Houliston and Elinson, 1991a; Schroeder and Gard, 1992). The cortical microtubule network persists until the end of the first cell cycle and is disrupted in part by the activation of Maturation Promoting Factor MPF; (Schroeder and Gard, 1992). The cortical network is composed of numerous parallel microtubules (Figure 7-1; Elinson and Rowning, 1988), of similar polarity (Houliston and Elinson, 1991b,) and the formation of this network occurs at the same time as the zygote cortex rotates with respect to the zygote interior. The colocalization of kinesin with this microtubule network suggests that molecular motors may be involved in the cortical rotation (Houliston and Elinson, 1991b).

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This rotation establishes the dorsal/ventral axis of the embryo (Gerhart et al., 1986), and several lines of evidence indicate that it is mediated by the parallel microtubule network: (1) The network is in a "shear zone" between the egg cortex and the interior cytoplasm, which is the appropriate position to rotate or facilitate rotation of the cortical cytoplasm (Elinson and Rowning, 1988). (2) Microtubule poisons, which disrupt the network, also prevent rotation and axis formation (Manes et al., 1978; Elinson, 1983; Ubbels et al., 1983). (3) Irradiation of the zygote at the time the network is forming (but not before or after) disrupts the microtubule network and prevents rotation (Elinson and Rowning, 1988). As in the species discussed in previous sections, Xenopus oocytes, eggs, and zygotes exhibit dynamic, non-random RNA distributions. The periphery of the vegetal hemisphere of the oocyte contains a striking localization of poly(A)"^RNA (Capco and Jeffery, 1982; Larabell and Capco, 1988). In addition, the vegetal periphery is also enriched in specific mRNAs, including Vgl (which encodes a protein related to transforming growth factor beta; Weeks and Melton, 1987; Melton, 1987) and tubulin mRNA (Larabell and Capco, 1988). Interestingly, the pattern of poly(A)'*"RNA localization throughout oogenesis (Figure 6; Capco and Jeffery, 1982) precisely parallels that exhibited by the Vgl mRNA (see Melton, 1987). The similarity of the spatial patterns suggests that there may be a common mechanism involved in the localization of mRNAs to the vegetal cortex. Biochemical analysis has revealed that other regions of the oocyte contain specific mRNAs. By separating oocytes into animal and vegetal hemispheres, and each hemisphere into peripheral and central regions (Figure 6; "spatial fractionation"; Capco and Mecca, 1988), we were able to purify and analyze, by blotting and probing with radiolabeled nucleic acid probes, mRNA from each of these different regions (Perry and Capco, 1988). Using this approach, we showed that actin and tubulin mRNA are concentrated in the oocyte periphery, whereas histone mRNA is distributed uniformly throughout the egg (Perry and Capco, 1988). Meiotic maturation is accompanied by RNA redistribution just as elements of the cytoskeletal are reorganized during the meiotic resumption. In eggs, tubulin mRNA is concentrated in the egg interior rather than the periphery, and a slight increase in the concentration of actin mRNA in the egg center is also detectable (Figure 7). The distribution of histone mRNA, on the other hand, does not change. In situ hybridization provides further confirmation that meiotic maturation triggers mRNA redistribution. For example, in the egg, poly(A)^RNA is no longer concentrated in the vegetal periphery (Capco and Jeffery, 1982) nor is mRNA for Vgl or tubulin (Melton, 1987; Larabell and Capco, 1988). Using timecourse in situ analysis of the poly(A)'^RNA distribution, we have determined that the localization in the vegetal periphery disappears at about the time the germinal vesicle breaks down and apparently is triggered by inhibition of a calcium-dependent chloride efflux which normally exists in the oocyte, but not the egg (Larabell and Capco, 1988). Fertilization re-establishes the concentration of tubulin and actin mRNAs in the zygote periphery, whereas histone mRNA is concentrated in the animal hemisphere

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Figure 6, (1) Microtubules in the Xenopus egg at the time of cortical rotation viewed with a confocal nnicroscope. (A) Microtubule bundles in the vegetal cortex. (B) Cross sectional view of vegetal hemisphere showing microtubules extending into the vegetal cortex. (C) Microtubules in the animal cortex after cortical rotation. (D) Microtubules in the animal cortex before cortical rotation. Scale bars are 25 |im. Reproduced from Schroeder and Card, 1992, with permission of Company of Biologists, Ltd. (2) Schematic showing the distribution of polyadenylated messenger RN A during oogenesis in Xenopus. Stippled areas show regions were RNA is localized. Reproduced from Capco and Jeffery, 1982, with permission of Academic Press. (3) Illustration of steps involved in "spatial fractionation" of Xenopus oocytes, eggs, or zygotes. (A) intact oocyte, (B) bisection of oocyte, (C) removal of internal cytoplasm of a hemisphere, (D) removal of peripheral cytoplasm from coelomic envelope. Reproduced from Perry and Capco, 1988, with permission of Elsevier Scientific Pub., Ltd.

(Figure 7; Perry and Capco, 1988). While it is unclear whether these RNA redistributions reflect active translocation or compartmentalized synthesis and degradation, two reports suggest the translocation can account for at least part of the observed patterns of RNA distribution. In the first, labeled poly(A)'^RNA isolated from the vegetal hemisphere and then injected into the animal pole of the activated egg was found to redistribute back to the vegetal pole (Capco and Jeffery, 1981). In the second, labeled Vgl mRNA injected into oocytes was found to translocate to the vegetal periphery, the normal location for endogenous Vgl

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¥\gure 7. (1) Schematic shows the spatial distribution of four different mRNAs as the Xenopus oocyte is converted to the fertilization-competent egg and subsequently fertilized. Stipples indicate the relative concentrations of the mRNAs. The pattern of tubulin mRNA undergoes extensive reorganization, whereas the pattern of actin mRNA remains relatively constant. Histone mRNA was initially randomly distributed, but becomes enriched in the periphery of the animal hemisphere after fertilization. Based on the results of Perry and Capco, 1988. (2) Schematic diagram illustrating pathways of signal transduction that act in Xenopus to convert the fertilization-competent egg into a zygote. Phorbol esters (PMA and PDD) or diacylglycerols (OAG and DiC8) activate protein kinase C (PKC) downstream of the sperm-induced rise in calcium. Reproduced from Bement and Capco, 1990, with permission of the American Society for Cell Biology.

mRNA (Yisraeli and Melton, 1988). In both of these studies, the kinetics of translocation and the relative stability of the RNA employed suggested that translocation, rather than compartmentalized synthesis and degradation, was responsible for the observed RNA redistribution. Sincepoly(A)'"RNA(Capco and Jeflfery, 1982), Vgl mRNA(Melton, 1987), and tubulin mRNA (Larabell and Capco, 1988) colocalize in the oocyte vegetal periph-

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ery with the cytokeratin network (Klymkowsky et al., 1987), while actin mRNA (Perry and Capco, 1988) colocalizes in the periphery with the actin network (Franke et al., 1976; Gall et al., 1983), and in view of the association of RNA andcytoskeletal elements in other systems, it seems likely that the Xenopus oocyte and egg cytoskeleton might be associated with localized populations of RNA. Indeed, although much work remains to be done, this seems to be the case. For example, it has been shown that Vgl mRNA is associated with the detergent-resistant cytoskeleton of the oocyte, but in the egg is associated with soluble components (Pondel and King, 1988). Cytokeratin filaments are not believed to be the component which immobilizes Vgl since experimental manipulations which disrupt the cytokeratin network do not solubilize a large portion of the Vgl mRNA and conversely conditions which solubilize the Vgl mRNA can leave the cytokeratin network intact. (Klymkowsky et al., 1991). It has also been demonstrated that the normal patterns of Vgl mRNA localization are disrupted by treatment of oocytes with other cytoskeletal poisons (Yisraeli et al., 1990). Using a detergent-extraction protocol originally designed for large cells or segments of tissue (Capco et al., 1987), we have examined associations between actin and tubulin mRNA and the cytoskeleton of Xenopus oocytes and eggs (Hauptman et al., 1989). In both the oocyte and the egg, 80 percent of the actin and tubulin mRNA are associated with the detergent-resistant cytoskeleton (Hauptman et al. 1989). Since actin and tubulin mRNAs have characteristic patterns of localization in both the oocyte and the egg (Perry and Capco, 1988), a simple interpretation of the above reports is that actin and tubulin mRNA retain their characteristic patterns of localization by binding to the cytoskeleton in the region where they are localized. In contrast, Vgl mRNA and other RNAs localized in the vegetal periphery bind to a detergent-resistant network only in the oocyte, and upon meiotic maturation either the mRNA detaches from the networit, or the network itself becomes soluble. In no other developmental system, except perhaps sea urchins, has the relationship between intracellular signaling events and observed patterns of cellular reorganization been as closely scrutinized as in Xenopus (Bement and Capco, 1989a, 1990c; Bement, 1992). As with sea urchins and ascidians, it has been demonstrated that amphibian eggs and embryos employ intracellular signaling pathways to control cytoskeletal transitions (Bement and Capco, 1990b, 1990c). A fertilizationinduced rise in [Ca^"^]} acts as the initial stimulus for many of the observed changes, including cortical granule exocytosis and cortical contraction (Schroeder and Strickland, 1974; Busa andNuccitelH, 1985; Kubota et al., 1987). The rise in [Ca^"']^ is coupled to cortical granule exocytosis, microvillar enlargement, and cortical contraction by protein kinase C (PKC; Figure 7; Bement and Capco, 1989b, 1990b; Capco et al., 1992; Bement and Capco, unpublished results). Evidence also indicates that PKC regulates formation and closing of the contractile actomyosin ring which mQdiaXQS Xenopus embryo cytokinesis (Bement and Capco, 1989b, 1990a, 1990c; Bement and Capco 1991; Brice and Capco, unpublished results). Moreover, PKC activity is also responsible for transit into the first mitotic interphase. The

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fertilization-induced elevation of [Ca^'^]j induces activation of PKC and also inactivates MPF thus driving the zygote into interphase (Bement and Capco, 1991). In fact, chronic activation of PKC using pharmacological agents can override the MPF signal and drive the cell into interphase even when MPF is active (Bement and Capco, 1991). PKC is a calcium- and phospholipid-dependent kinase, and presumably therefore exerts its effects by way of phosphorylation. What are the relevant PKC substrates underlying cortical contraction and cytokinesis? All of the available evidence suggests that phosphorylation of myosin light chain may represent the key trigger for PKC-mediated contraction and cytokinesis: First, myosin is required for cortical contraction both in vitro and in vivo (see above). Second, phosphorylation of myosin light chain is correlated with actin gel contraction in Xenopus egg extracts (Ezzell et al., 1983). Third, phosphorylation of myosin light chain in vivo is correlated with cortical contraction, and this phosphorylation appears to be mediated by PKC (Bement and Capco, unpublished data). Finally, phosphorylation of embryonic myosin light chain is known to increase the actin-activated ATPase activity of myosin (de Lanerolle and Nishikawa, 1988). These data, and the fact that PKC antagonists prevent cytokinesis in Xenopus embryos (Brice and Capco, unpublished results), lead us to suggest the following hypothesis for amphibian embryonic cytokinesis: The rise in [Ca^"^]j which accompanies each meiotic and mitotic division (Whitaker and Patel, 1990) transiently activates PKC. Upon activation, PKC phosphorylates myosin light chain, thereby imparting an increase in the actin-binding affinity and actin-activated ATPase activity of myosin (de Lanerolle and Nishikawa, 1988). This results in an increase in actomyosin-based force production and consequently, cell contraction and cytokinesis. Intracellular signals have been demonstrated to play a central role in the reorganization of the cortical cytoskeleton as the oocyte is converted to the fertilization-competent egg during the meiotic resumption (Capco et al., 1992). It is well known that the cortex of amphibian oocytes acquires contractile ability during the meiotic resumption (Gingell, 1970; Schroeder and Strickland, 1974; Merrian and Sauterer, 1983; Ryabova, 1983; Ezzell et al., 1985) and evidence suggests that this is due to an interaction between actin and myosin (Meeusen and Cande, 1979; Christensen et al., 1984; Ezzell et al., 1985). By bisecting oocytes, as originally described by Christensen et al. (1984), at increasing time intervals after the resumption of meiosis we showed that the oocyte acquires this contractile ability in two phases. An oocyte cortex has virtually no contractile ability and when bisected the hemisphere undergoes no change. However, when bisected immediately after meiosis has been reinitiated the bisected surface forms a contractile ring capable of closing and sealing off the hemisphere (Figure 8). When bisected between 70-80% through the meiotic resumption the oocyte's contractile ability changes again as the contraction becomes oriented from the bisected surface to the apex of the hemisphere causing the cortex to pinch off of the interior cytoplasm and a cortical cap forms (Figure 8). The ability to form a contractile ring and

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* TIME TO PKC •SNDuCtD C O N T R A C - O N

(conf/nL/ecO

Figure 5. (1) Time course of contracti le ring formation i n progesterone treated oocytes bisected immediately after progesterone addition. Oocytes were cytochemically fixed to arrest changes. (A) intact oocyte, (B) oocyte immediately after bisection with cut surface lying on petri plate, (C) same as in "B" except bisected surface is standing on edge, (D) contractile ring forming (arrowheads), and (E) further closure of the contractile ring. (2) Time course of cortical cap formation as a result of bisection late after progesterone addition. The same animal hemisphere is shown in time sequence of 30 minutes to demonstrate cap formation. Pigment is black; internal yolk is white.

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subsequently a cortical cap can be prevented by pharmacological agents which inhibit PKC. Moreover, the entire sequence of contractile events (e.g., contractile ring formation and subsequently cortical cap formation) can be induced by agents which activate PKC (Figure 8; Capco et al., 1992), suggesting that PKC plays a pivotal role in the reorganization of the cortical cytoskeleton. Intracellular signals also appear to regulate microtubule dynamics during meiotic resumption and egg activation. Maturation promoting factor (MPF), a complex of phosphoproteins which acts at the top of a cascade of kinase/phosphatase reactions, converts the cytoplasmic microtubule array of the oocyte into the spindle array of the egg in vitro (Lohka and Mailer, 1985) and in vivo (Karsenti et al., 1984). For example, a microtubule-associated protein with molecular weight of 220 kD has been isolated from M-phase egg and interphase cells (i.e., activated eggs); the protein from interphase cells bound to microtubules, promoted tubulin polymerization and was dephosphorylated whereas the protein from M-phase eggs, which may be a downstream effector of MPF, exhibited neither activity but was phosphorylated while in interphase (Shiina et al., 1992). Similarly, phosphorylation of other microtubule associated proteins (MAPs) may regulate sperm aster formation and possibly formation of the vegetal microtubule array. For example, Gard and Kirschner (1987) identified "XMAP", a 215 kD MAP found in the cytoplasm of Xenopus eggs. XMAP promotes extremely rapid microtubule polymerization, and high XMAP activity is correlated with dephosphorylation of XMAP. Since formation of the sperm aster and the subcortical microtubule array coincide with high XMAP activity, it is likely that XMAP contributes to the regulation of these structures. Further evidence for cell cycle regulators acting on the cytoskeleton comes from the work of Buendia et al. (1992) who showed that one of the cyclins, cyclin A, appears to act directly on centrosomes to increase their microtubule nucleating activity. As with the other systems considered in this c\va^\,QX Xenopus oocytes exhibit an elaborate cortical C3^oskeleton which is reorganized during progression through meiosis. Fertilization again alters the organization of the cytoskeleton and reveals that this cytoskeleton has acquired contractile ability. Pronuclear movement is

Figure 8. (Continued) As cortex undergoes actln-myosin contraction pigment becomes restricted to a small cap at the center of the hemisphere progressively revealing more yolk. (3) The effects of the two PKC simulators a phorbol ester (A) and the synthetic diacylglycerol, DiC8 (B) is shown. The ability to form a contractile ring or cortical cap was measured against the time required for PKC to induce cortical contraction. When the times monitored were expressed as a percentage of the time for PKC-induced contraction, the kinetics of these changes exactly paralleled the effects of progesterone on oocytes, suggesting that PKC is sufficient to remodel the actin-myosin cytoskeleton in the oocyte. Reproduced from Capco et al., 1992, with permission of Wiley-Liss.

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mediated in part by this contractile event and by the elaboration of microtubules from the sperm centrosome. This system clearly demonstrates cytoskeletal regulatidn by signal transducers.

V. SUMMARY A useful starting point for investigating the role of the cytoskeleton in early development has been the results obtained from studies examining the cytoskeleton in somatic cells. However, the developmental challenges faced by eggs, zygotes, and early embryos appear to require specializations of cytoskeletal function, as described in this chapter. The cytoskeletal specializations outlined in this article, are not meant to be an exclusive list but rather a starting point for the recognition of these special roles. Indeed, future studies are likely to lengthen the list of specialized cytoskeletal functions in early development. Conversely, it is also useful to remember that somatic cells originate from the development of an embryo, and there is no reason to assume that somatic cells lose all of the specialized functions present in their progenitor cells (i.e., the zygote and blastomeres of the embryo). But in somatic cells some or all of these specialized roles may be greatly reduced in scope, making them difficult to detect. Thus characterization of the specialized functions of the cytoskeleton during early development may aid the identification of more subtle cytoskeletal functions in somatic cells.

ACKNOWLEDGMENT The author thanks Deitmar Schmucker for critical reading of the section on Dwsophila. work was supported in part by NIH grant HD27151.

This

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MICROTUBULE-BASED INTRACELLULAR TRANSPORT OF ORGANELLES

Howard Stebbings

I. Introduction II. Microtubules as Motility Substrates A. Microtubule Disposition in Cells B. Microtubule Polarity C. Microtubules and Structural MAPs III. Which Organelles are Translocated Along Microtubules? IV. Model Systems Extensively Used to Study Microtubule-Based Intracellular Movement A. Movements within Cultured Cells B. Translocation Along Nerve Axons C. Pigment Granule Movements in Chromatophores D. Movement Between Nurse Cells and Oocytes in Insect Ovaries E. Movement within Reticulopodia of Protists V. The Interaction Between Microtubule and Microfilamentous Systems in Intracellular Transport

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 113-140 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 113

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VI. Video-Microscopy: The Breakthrough in the Characterization of Microtubule-Based Movement VII. The Identification and Isolation of Microtubule Motors VIII. Occurrence of Kinesin and Cytoplasmic Dynein— Distribution and Cellular Localization IX. The Characterization of Microtubule Motors A. The Structure and Molecular Structure of Kinesin and Dynein B. Force Generation and the Mechanochemistry of Microtubule Motors . . C. Nucleotide Specificity and Drug Sensitivity of Kinesin and Cytoplasmic Dynein X. Families of Microtubule Motors A. Genetic Approaches to Identifying and Characterizing Molecular Motors XI. The Regulation of Microtubule Motors and the Control of Directionality of Organelle Transport A. cAMP and Calcium in the Regulation of Microtubule-Based Motility . . B. The Association of Motors with Vesicles C. The Influence of Structural Maps on Microtubule Motor Activity XII. Perspectives References

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1. INTRODUCTION Since the days of early microscopes it has been clear that components of all varieties move around and are transported within cells. This transport is more pronounced in some cells than others and is particularly obvious w^here cells have adopted an asymmetric shape. This they may do to extreme degrees, as is the case with nerve axons, and in such instances transport is readily observed in the cell processes. The advent of electron microscopy and its application to the study of cell ultrastructure demonstrated that the cell processes along which intracellular transport had been seen to occur, invariably contained elements of the cytoskeleton and notably microtubules arranged parallel to the long axis of the process and to the direction of movement. Such findings encouraged the belief that microtubules played a role in intracellular transport, and this view was confirmed indirectly in experiments using antimitotic agents which disrupted the microtubules, bringing about a collapse of the cell processes and a cessation of movement along them. Carefiil ultrastructural studies were suggestive of associations, in the form of cross-links, between microtubules and translocating organelles of a variety of descriptions but the equation of microtubules and other cytoskeletal structures with intracellular movement was for many years essentially circumstantial. A highly significant breakthrough came in the early 1980s with the advent of video microscopy. In conjunction with image processing it immediately became possible to visualize individual "living" microtubules in vivo and in vitro and as a consequence it was confirmed that organelles are indeed transported along microtubules. Subsequent to this, conditions began to be developed which "trapped" certain polypeptides onto

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microtubules, and these proved to be motor proteins which are capable of driving artificial beads or organelles along microtubules in one or other direction. Since the initial identification and characterization of two microtubule motor proteins which propel components in opposite directions along microtubules, it has become apparent that they are only part of the story and it is emerging that they represent members of families of motor proteins. The significance of the multiplicity of motor proteins in different cellular situations is receiving a great deal of attention as is the way these might be regulated in cells. All these topics will be covered in this review of the role of the cytoskeleton in the intracellular transport of organelles. Whereas the studies outlined above have demonstrated organelle transport to be microtubule-based, cytoplasmic streaming, a phenomenon more typical of plant cells, has in parallel studies been shown to be based on actin filaments. Both microtubules and actin filaments are integrated components of the cytoskeleton, and it is becoming increasingly clear that it is artificial to separately compartmentalize the two as more examples demonstrate movements of specific components by both systems.

II. MICROTUBULES AS MOTILITY SUBSTRATES A. Microtubule Disposition in Cells

If, as already outlined, microtubules are involved in the intracellular translocation of organelles, a consideration of their disposition in cells is clearly pertinent. In cells generally, microtubules arise from organizing centers, which in animal cells are usually positioned close to the nuclear envelope. Microtubules radiate outwards from these microtubule organizing centers toward the cell periphery, and where the cell has differentiated and shows any degree of asymmetry, the microtubules can usually be seen to extend along the axis of asymmetry. This is true not only of differentiated nerve cells with their long axonal processes, but also many cell types, and a number of these like nerve axons, have become extensively-researched models for the study of microtubule-based organelle translocation. These include a range of chromatophores offish and amphibia which adopt stellate patterns with microtubules extending along the cell extensions. Some protozoans such as the heliozoans also show dramatic asymmetry, with very fine axopodial extensions along which run geometrically elaborate aggregates of microtubules. While the above arrangement is true for many cells, in polarized epithelial cells the microtubules do not radiate outwards from a single nucleating center, but run vertically through the cell from the basal to the apical surface (Gorbsky and Borisy, 1985). These extreme examples illustrate the importance of microtubules as skeletal organelles, as well as organelles implicated with motility. In addition to microtubules occurring in cell extensions there are also examples of microtubules being the main components of intercellular connections. Such an example is seen in the

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ovaries of certain insects, where the developing oocytes and nurse cells which supply them with various components throughout oogenesis, are linked by extensive cytoplasmic channels which can often extend several millimeters in length and which are packed with an astonishing number of parallel microtubules. These are not the only arrangements of microtubules in cells. Elsewhere in these volumes is described how microtubules from pairs of organizing centers assemble into mitotic spindles, and how chromosomes move in various ways in conjunction with these microtubule arrangements. Fascinatingly, the spindle also appears to be involved in the rearrangement of membranous organelles during the different mitotic stages in this way may perform what might be regarded as a double function (Waterman-Storer et al., 1993). In some cells microtubules appear to be organized from multiple centers, and in plant cells ultrastructural and immunocytochemical studies have shown that microtubules adopt a helical pattern around the cell periphery. B. Microtubule Polarity Because of the way in which tubulin dimers assemble into the walls of microtubules, the organelles have an intrinsic polarity. This polarity was, from the time of its elucidation, regarded as likely to be significant in a component of the cytoskeleton involved in the directional transport of organelles in cells. Microtubule polarity is readily assessed in cellular situations where it is clear that the microtubules have assembled from a distinct organizing center. In other cases the origin of microtubules is less obvious, and there are many instances where there has been considerable debate as to the polarity of a particular set of microtubules. There are also examples where microtubules detach from their origin of assembly, and some where the polarity of microtubules is the opposite to what might have been expected. To clarify these situations ingenious techniques have been devised so that the polarity of a particular set of microtubules can usually be determined in situ. Certain of these methods are based on the way in which particular markers, such as dynein but probably more effectively tubulin itself, bind to the microtubule lattice which can then be seen ultrastructurally as directional projections. Other procedures depend on growing labeled tubulin onto existing microtubules and then detecting microtubule polarity from the different growth characteristics of the two ends of a microtubule. In general these techniques have shown that the minus ends of microtubules are associated with the centrosomes, while the plus, fast-growing ends are peripherally located. C. Microtubules and Structural MAPs Since microtubule proteins were first isolated from tissues, most notably initially from mammalian brain, it has become clear that while the tubulins which comprise

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the walls of the microtubules are the most prominent proteins, other minor proteins also exist, and these have engendered a great deal of study. These microtubuleassociated proteins (MAPs), for reasons already mentioned, have been most extensively studied from mammalian brain tissue, but comparable MAPs have subsequently been identified and characterized from non-neuronal mammalian tissue, from other vertebrate sources, from invertebrates and also a variety of plants. Many of the large number of MAPs now characterized have properties similar to those described for the better known brain MAPs. Importantly, there are indications that the MAPs isolated by current techniques may represent only a small fraction of the proteins that actually associate with microtubules in situ and new methods are being devised to assess the extent of existence of these molecules. With regards function, MAPs have been shown to promote microtubule assembly, to conserve microtubule stability, and of importance for this discussion they form side arms from microtubules which are thought to mediate associations and interactions between neighboring microtubules; or between microtubules and different cellular organelles. Those MAPs whose structure has received attention have been shown to have a portion of the polypeptide bound to the microtubule lattice, the binding domain, with the larger portion extending from the microtubule surface, the projection domain, which may or may not be easily seen ultrastucturally. Not only do MAPs function in assembling and maintaining microtubule substrate bundles, but their binding sites obviously occupy significant but clearly variable amounts of the microtubule surfaces and therefore have the potential for influencing the way in which any other components might interact with a particular microtubule—a, point which will be returned to later. There is also the possibility that MAPs might anchor organelles to microtubules in between periods of translocation.

III. WHICH ORGANELLES ARE TRANSLOCATED ALONG MICROTUBULES? Evidence has accumulated that the organization and arrangement of most, if not all, of the membrane compartments of eukaryotic cells, particularly those of the exocytic and endocytic pathways and the membrane trafficking between them, depend on the existence of an intact microtubule system within the cell and microtubule-based movement (Figure 1). Nuclei may be positioned within the cytoplasm by movement along microtubules, and this certainly appears to be the case with nuclear migration in yeast (Jacobs et al., 1988). The nuclei in viral-induced syncytia have also been described to aggregate along tracts of microtubules. In the cytoplasm cells tend to concentrate their Golgi membranes at the cell center and to disperse their endoplasmic reticulum (ER) throughout the cytoplasm. The reliance of the actual positioning of the Golgi on microtubules has been demonstrated by the experimental disruption of the cell's microtubules, which results in

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Figure 1. Microtubule-based translocation in one or other direction has been implicated in the organization of, and trafficking between, the different nnembrane compartments of eukaryotic cells.

the dispersement of Golgi elements. On the other hand, when the microtubules reform Golgi membranes have been shown to recluster around the centrosome by migrating along those microtubules (Ho et al., 1989). Moreover, another approach has served to confirm this, since Golgi complexes applied to semi-intact cells in culture have been shown to be actively transported to the minus ends of microtubules and to accumulate at the centrosome region (Corthesy-Theulaz et al.,1992). The behavior of distending along microtubules is also seen in other organelle networks such as the ER. Concerning the ER the opposite to that seen in the case of the Golgi occurs with the ER membranes retracting towards the cell center on microtubule depolymerization, and reforming to a typical extended distribution on microtubule repolymerization (Terasaki et al., 1986). This would appear to be confirmed from in vitro experiments using membranous extracts believed to be from the ER which form tubular extensions during an active migration relative to microtubule substrates (Vale and Hotani, 1988). Lysosomes, like the Golgi, interact with microtubules and are maintained in the perinuclear region (Matteoni and Kreis, 1987); and in macrophages, where lysosomes have been readily visualized due to the uptake of the dye lucifer yellow, they also adopt a tubular shape which is dependent on their association with microtubules within the cell.

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In relation to the anabolic and catabolic membrane compartments of the cell there has been increasing interest in membrane trafficking within these systems. Studies have shown that trafficking from the ER to the Golgi apparatus, the brefeldin-induced rapid redistribution of Golgi back into ER in the opposite direction (Lippincott-Schwartz et al., 1990), and also movements within the Golgi apparatus (Cooper et al., 1990), all occur along microtubules. Furthermore, secretory granules in exocytosis and endosomes in endocytosis, labelled with the fluorescent probes acridine orange and lucifer yellow respectively, have been seen to migrate along microtubules between the cell center and the cell periphery (Kreis et al., 1989; Hollenbeck, 1993a). Mitochondria are also translocated along microtubules and one prominent example of this is seen with insect ovaries where mitochondria migrate along microtubules to accumulate in large numbers in the developing oocytes (Stebbings and Hunt, 1987). Microtubules may also form associations, and indeed interact, with non-membrane bound organelles, such as ribosomes (Suprenant et al., 1989), and of course other elements of the cytoskeleton—^including other microtubules. With regards cytoskeletal components, there is a great deal of evidence for microtubules interacting with microfilaments via MAP interconnections. Intermediate filaments appear to be positioned in cells by microtubules. Destruction of the latter by colchicine has been shown to result in a collapse of the intermediate filament network. When the microtubules are allowed to reform, the intermediate filaments redistribute themselves to the cell periphery, and are thought to do so by being actively transported along the microtubules (Gyoeva and Gelfand, 1991). Microtubule translocation by microtubules, resulting in their relative sliding has been well documented. The relative sliding of parallel microtubules within the axonemes of cilia and flagella forms the basis of their bending movements, and a similar sliding of anti-parallel microtubules occurs during anaphase movements of chromosomes at cell division. It is believed too that this phenomenon of the relative sliding between microtubules within cells might be a general one.

IV. MODEL SYSTEMS EXTENSIVELY USED TO STUDY MICROTUBULE-BASED INTRACELLULAR MOVEMENT A. Movements within Cultured Cells

Cultured fibroblasts have been used to study the movement of organelles on microtubules, and are useful for such studies as they have thin processes (Hayden and Allen, 1984) through which the phenomenon can be observed. This has allowed the study of the behavior of such particles, the movements of which have been described as saltatory, with frequent stops, reversals in direction and changes in velocity.

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Comparable intracellular organelle motility to that seen in living fibroblasts can be reproduced in extracts of fibroblasts (Dabora and Sheetz, 1988), and in such cases the microtubule-associated movements are even easier to observe. In general, however, the movements of cellular organelles in relation to microtubules becomes particularly pronounced in some differentiated cells, and hence it is these cases which have received the most experimental attention. The most accentuated and obvious examples of the phenomenon occur where a cell has become highly asymmetric, and in such instances translocation invariably occurs in association with microtubules along the axis of asymmetry. B. Translocation Along Nerve Axons

An excellent example of microtubule-based intracellular translocation over long distances is seen in nerve cell axonal processes (Figure 2). These extensions are often enormous on a cellular scale reaching, for example, more than one meter in length in human motor neurones. They contain a parallel arrangement of microtubules, most oriented with their plus ends toward the synapse, as well as other cytoskeletal elements such as neurofilaments and microfilaments. Fast movement of neurosecretory vesicles towards the nerve terminal and synapse can be seen directly using light microscopy, and has been estimated at rates of 50-500 mm/day. This is superimposed on a much slower movement of cytoskeletal proteins which occurs at 0.2-0.8 mm/day and which can only be detected by means of radiolabeling. Even here, careful studies have resolved the slow movement into different component groups with the slowest (known as SCa), comprising neurofilament and microtubule protein which moves en bloc—probably as neurofilaments and microtubules, and a slightly faster fraction (known as SCb) which includes actin, myosin, clathrin, and spectrin and the so-called carrier proteins. As well as these anterograde movements towards the axon terminal there are simultaneous retrograde movements which again have different characteristics, in the opposite direction, toward the cell body. In addition to observing axonal translocation in intact nerve axons, movements have, as in the case of fibroblasts, been seen to great advantage in permeabilized axons and also in axoplasmic extracts. Regarding the latter, the giant axon of the squid (Allen et al., 1985) has proved to be an immensely valuable and rewarding material for study. C. Pignnent Granule Movements in Chromatophores

The dermal chromatophores, or pigment cells, involved in color changes in many lower vertebrates show a striking example of microtubule-based intracellular organelle transport (Figure 2). In these cells microtubules radiate from the cell center outwards to the cell periphery The redistribution of pigment granules within the cells occurs either outwards towards the plus ends, or inwards toward the minus ends, of this microtubule framework. Chromatophores, which may be termed

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cell body (C)

Figure2, Diagram illustrating microtubule-based translocation (a) along nerve axons, (b) along cell processes of chromatophores and (c) from nurse cells to the developing oocytes in insect ovaries.

xanthophores, iridophores, erythrophores, or melanophores depending on the type of pigment concerned, differ from axons in terms of their movement in that anterograde and retrograde movements do not occur simultaneously but as two distinct events. This has made them favorite material for investigating the control of intracellular transport. The movements of pigment granules in chromatophores, which is related to the organism's behavioral patterns, is rapid, in the order of 1-6 jiim/sec. It has different characteristics in the two directions. In vitro models of the system have been extensively studied and these have been produced by isolating chromatophores which can be maintained in culture, and then either lightly permeabilized (Rozdzial and Haimo, 1986a) or stripped of most of their membrane (McNiven and Ward, 1988). The beauty of these preparations is

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that aggregation or dispersion of the pigment granules can then be attained by applying the appropriate stimulus. This has shown that pigment dispersion is cAMP-dependent and requires the phosphorylation of certain proteins, while conversely pigment aggregation involves the activity of the Ca^"*" and calmodulindependent phosphatase, calcineurin (Rozdzial and Haimo, 1986b), a point which will be returned to in a later section. D. Movement Between Nurse Cells and Oocytes in Insect Ovaries The developing oocytes in the ovaries of many insects are in cytoplasmic continuity with nurse cells, and the latter have been shown to supply the oocytes with various components, at least during the previtellogenic stages of oogenesis (Fig. 2). In most orders of insects where this occurs the two cell types are in close proximity, but in the hemipterans, the bugs, the nurse cells remain at the anterior end of the ovarioles and are then connected to the oocytes by nutritive tubes which may lengthen to become a few millimeters long in some cases. A range of components, predominantly ribosomes, and in some species mitochondria, have been shown to pass unidirectionally to the oocytes where they accumulate. The overall translocation of materials down the nutritive tubes is slow, and can only be monitored using autoradiography. Using this technique a faster movement superimposed on the slow movement has been detected much as has been shown for axonal transport (Stebbings, 1986). The most obvious feature, ultrastructurally is that the nutritive tubes are packed with tens of thousands of parallel microtubules all arranged with their plus ends towards the nurse cells and their minus ends towards the oocytes (Stebbings and Hunt, 1983). The microtubules are interspersed with ribosomes and mitochondria, and transport towards the oocytes is therefore retrograde. The numbers of microtubules within the nutritive tubes is so great that the tubes are very strongly birefringent and this feature has enabled them to be dissected from ovarioles by manual microdissection using tungsten needles. The addition of ATP to isolated nutritive tubes has been shown to result in the reactivation of translocation of certain components along their length, suggesting that their transport along this system is an active process (Stebbings and Hunt, 1987). E. Movennent within Reticulopodia of Protists The marine foraminiferan, Allogromia, and the giant freshwater amoeba, Reticulomyxa, both have extensive reticulopodial networks which contain microtubules and have been valuable for the study of microtubule-based movements. Investigations of AUogromia have shown that microtubules predominate within the reticulopodia and that the microtubules are themselves motile; and this motility has been regarded as propelling pseudopod extension. In addition, the organism shows continuous bidirectional transport of organelles of all varieties along the micro-

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tubules (Travis and Bowser, 1986), and its dependence on microtubules has been demonstrated using antimitotic agents. The giant multinucleated syncytial freshwater amoeba, Reticulomyxa, is very similar to Allogromia, existing as a highly dynamic extensively-branched system of membrane-bound filamentous strands (Koonce and Schliwa, 1986). Again bidirectional streaming of mitochondria, various sorts of vesicles and food vacuoles occurs within the strands at fast rates, up to 20 |Lim/sec. Some strands possess many microtubules as well as microfilaments, but even thefineststrands containing single microtubules show bidirectional movements. Additionally, membrane-bound organelles can clearly be seen to move relative to the microtubules, which are also believed to move relative to each other. A further advantage of these protists is that, as with other systems, lysed cell models yield extensive cytoskeletalfi-ameworkswhich still support the movement of endogenous organelles. Moreover, even after severe extraction the movements can be reactivated and so provide a valuable model for studying the molecular basis of the phenomenon.

V. THE INTERACTION BETWEEN MICROTUBULE AND MICROFILAMENTOUS SYSTEMS IN INTRACELLULAR TRANSPORT While there are examples, such as in the insect ovary, where translocation occurs solely in association with microtubules, in most cellular situations microtubules intermingle with other cytoskeletal elements. However, where microtubules and microfilamentous systems are found together little is known about the association and possible interaction between these two systems in intracellular transport. Evidence is emerging that it may be unwise to treat these totally separately. Studies have suggested that the movement of chloroplasts in the alga Bryopsis requires organized arrays of both microtubules and actin filaments. For some time it has been realized that fast axonal transport of neurons requires actin filaments (Brady et al., 1984) as a result of the gelsolin-inhibition of transport. Indeed more recent observations of axoplasmfi*omsquid giant axons has shown that there are two types of translocation in such preparations. Bidirectional movement of vesicles occurs, as has already been discussed, along the parallel microtubules within axonal preparations, but in addition other directed movements have also now been described which occur independently of microtubules, often at right angles to them, unidirectionally, and in conjunction with what has proved to be an intermingling network of actin filaments (Kuznetsov et al., 1992). These observations are fascinating as they illustrate an association, and perhaps a cooperation between microtubule-based and microfilament-based intracellular movement.

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VI, VIDEO-MICROSCOPY: THE BREAKTHROUGH IN THE CHARACTERIZATION OF MICROTUBULE-BASED MOVEMENT Until about 10 years ago the equation of intracellular transport with microtubule components of the cytoskeleton was essentially circumstantial. Using light microscopy a variety of cell organelles, as has already been described, had been seen to move in cellular locations which electron microscopy revealed to contain aggregates of parallel microtubules. The frustrating fact was that it was not possible to observe both the intracellular movements and the cytoskeleton with the same instrument so as to be able to assess the implied interrelationship. Development of the technique known as video microscopy allowed this and provided the essential breakthrough in understanding microtubule-based intracellular movement. Briefly, the basic equipment for studying such movement is a research compound microscope fitted with differential-contrast optics of high numerical aperture, to which is attached a high resolution video camera. The signal from the camera passes to an image processor which enhances the raw image by increasing contrast, and at the same time allows one to visualize structures 5—20 times smaller than one could see or photograph using a light microscope in the standard fashion. Image processing can also be used amongst other things, to "clean" images by subtracting background noise. These techniques, pioneered by S. Inoue and by R.D. Allen, were first used for visualizing microtubules (24 nm) in living cells and extracts, and then in reconstructed systems; since membrane-bound vesicles could also be seen with the method it provided the means to observe and investigate the movements of one relative to the other. One of the main caveats with the procedure (not always appreciated) is the fact that sub-resolution structures such as microtubules are not portrayed as their real dimension, but are increased in size to the limit of resolution. Hence, images obtained require a certain amount of interpretation. This is not a problem when, for example, one might wish to observe the movement of a particle or vesicle relative to a microtubule. It does create difficulties if one wishes to observe the movement of a microtubule relative to a neighboring microtubule since video microscopy does not distinguish between a single microtubule and pairs and aggregates of closely adjacent parallel microtubules; and the relative sliding of microtubules can only be implied from the technique when a certain amount of splaying has resulted. These difficulties have been overcome to an extent by binding beads to microtubules, after which the monitoring of the bead movements has permitted an indirect analysis of microtubule movements (Kamimura and Kamiya, 1989). With the advent of video microscopy a range of preparations known to exhibit microtubule-associated movement began to be subjected to close scrutiny. One of the earliest and most vigorously examined of these was the axoplasm extruded from the giant axons of squid. Such preparations were already known to contain large

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numbers of microtubules as well as other cytoskeletal organelles and to continue the transport of organelles and vesicles for several hours after extrusion. Observation of such preparations by video microscopy showed that particles exhibited Brownian motion in suspension, but on coming into contact with a microtubule adhered to it and were transported along the microtubule with characteristics seen in the intact axons, remaining attached to a microtubule until they reached its end. Again, as with whole axons, bidirectional movement of particles could be observed, even along a single microtubule. Surprisingly, as well as showing particles riding along microtubules, the axoplasm preparations also revealed short lengths of microtubules gliding over the glass substrates, and larger microtubules tended to produce writhing serpentine shapes. This was interpreted as being a different manifestation of the activity of the same mechanochemical enzymes that were believed to produce transport along the microtubules. The observations outlined from viewing axoplasm preparations have since been confirmed in a multitude of systems from intact cells, particularly protozoa, where bidirectional transport has been shown to occur along cell processes which contain only single microtubules, to permeabilized cells where cytoskeletal functions are retained, a range of other cytoplasmic extracts for example from cultured cells (Dabora and Sheetz, 1988), and even instances where microtubule transport systems can be microdissected by hand for subsequent observation.

VII. THE IDENTIFICATION AND ISOLATION OF MICROTUBULE MOTORS Closely following the observation of membranous organelles translocating along microtubules in axoplasmic extracts came the identification of the first brain (neuronal) ATPase with properties expected for the fast axonal transport motor. The key to its discovery proved to be the use of the non-hydrolyzable analogue of ATP, adenylyl imidodiphosphate (AMP PNP) which was known to inhibit axoplasmic translocation, and was found to form stable interactions between membranous organelles and microtubules. Microtubule pellets incubated with AMP PNP and soluble factors from brain tissue were shown to be enriched with a 130 kD polypeptide and an associated 70 kD species, the former having ATPase activity and interacting with microtubules in an ATP-dependent manner (Brady, 1985). This ATPase had many properties suspected for an enzyme involved in fast axonal transport, and this was in part confirmed when it was shown to produce movement of plastic beads, to which it had been added, along microtubules—3, demonstration which resulted in it being named kinesin. Kinesin has in fact been demonstrated to generate several forms of ATP-dependent microtubule-based movements in vitro. Bead movements driven by kinesin along microtubules were seen to be smooth, continuous and unidirectional and to occur at velocities of 0.3-0.5 |Lim/sec (Vale et al, 1985). Kinesin also

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supported the movement of organelles along microtubules at a velocity comparable to that seen in dissociated axoplasm and the movement of microtubules themselves over glass substrates to which it had been applied. In order to assess directionality, kinesin-coated beads were then applied to microtubules grown onto isolated microtubule organizing centers (MTOCs) which are then known to have their plus ends distal, and their minus ends proximal to the MTOCs. In such instances all the beads moved outwards towards the plus ends of the microtubules, indicating that kinesin appeared to be a candidate for the anterograde motor. Since organelles from different systems had clearly been shown to translocate in two directions even along a single microtubule, and following the discovery of a plus-end directed microtubule motor protein, kinesin, attention began to focus on the search for the minus-end directed motor. Mammalian brain microtubules had already for some time been known to possess a number of high molecular weight (HMW) MAPs, and one of these named MAP IC was shown to have nucleotidedependent binding to microtubules—^indicating it to be a possible motor (Paschal et al.,1987). It had previously been regarded as a minor MAP, but was found to be enriched in microtubules assembled in the absence of nucleotides, something which had become possible with the advent of the drug taxol which readily promotes microtubule assembly. The confirmation of MAP 1C as a microtubule motor protein came from the finding that it possessed ATPase activity, which like kinesin was enhanced in the presence of microtubules; and when applied to glass substrates it was shown by video microscopy to support microtubule gliding in the presence of ATP. MAP IC was then shown to share some properties with axonemal dynein, in particular being cleaved by exposure to ultraviolet in the presence of vanadate and ATP. These similarities indicated MAP IC to be a cytoplasmic form of dynein and therefore a possible retrograde motor. The actual demonstration of MAP IC as a retrograde motor came from a microtubule gliding assay, using axonemes from the biflagellate unicellular alga, Chlamydomonas. The microtubules making up the axoneme are all of the same polarity and have a built in polarity marker because under certain conditions their plus ends distal to the cell body have a distinct tendency to fray. MAP IC caused axonemes to glide in the opposite direction to kinesin from the compact (minus) towards the frayed (plus) end (Paschal and Vallee, 1987). Like kinesin, cytoplasmic dynein has been shown to be capable of driving beads along microtubules, and the retrograde movement of dynein-coated beads is slightly faster and more jerky than is seen with kinesin. They also show some lateral displacement suggesting that dynein may move from one microtubule protofilament to another. Hence to be designated microtubule motor proteins, kinesin and dynein are enzymes which have nucleotide and microtubule binding sites. They demonstrate microtubule-activated nucleotidase activity and they use energy from nucleotide hydrolysis to generate movement relative to microtubule substrates (see Figure 3).

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Figure 3. Nutritive tube translocation channels isolated from ovarioles of the hennipteran insect, Notonecta. The nutritive tubes are comprised of many thousands of microtubules interspersed with ribosomes and mitochondria (electron micrograph, top right), and translocation of the latter along with the microtubules can be reactivated and viewed in dark-field, following the application of ATP {top left) (Stebbings and Hunt, 1987). Isolated nutritive tubes can be extracted with Triton X-100 so that only a parallel bundle of microtubules remains (center). Motor-coated beads applied to microtubule substrates from isolated and extracted nutritive tubes (bottom), translocate on reactivation with ATP (Anastasi, Hunt and Stebbings, 1990). (Left hand panels xl,000; right hand panels approx. xl 15,000.)

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VIIL OCCURRENCE OF KINESIN AND CYTOPLASMIC DYNEIN—DISTRIBUTION AND CELLULAR LOCALIZATION Kinesin is in all likelihood a ubiquitous protein, and has been demonstrated to be present in a variety of tissues and organisms. These include systems which possess large numbers of microtubules and examples where microtubule-based translocation has been well-documented, such as neuronal material from cow (Bloom et al., 1988) as well as from other mammals and vertebrates, squid axons, sea urchin eggs (Scholey et al, 1985), Drosophila (Saxton et al, 1988), and the ovaries of other insects (Anastasi et al., 1990), and Acanthamoeba (Kachar et al., 1987). Kinesin has been shown by immunotechniques to be largely soluble, and in fibroblasts, for example, about one third of it is organelle-bound (Hollenbeck, 1989). It is present in widely different amounts in different cell types, and possesses a distribution which is consistent with a role in organelle transport and organelle positioning. This has been illustrated by a survey of chick tissues by Hollenbeck (1989) where he found that neuronal tissue had the highest levels of kinesin while red blood cells, which show no organelle translocation, had no detectable kinesin. Dynein-like enzymes had been identified in the C3^oplasm of sea urchin eggs at a much earlier stage (Weisenberg and Taylor, 1968) but there had been considerable uncertainty as to whether these were cytoplasmic forms of dynein involved in a range of cellular motor activities including mitotic spindle function and organelle translocation, or whether they were ciliary precursors merely stored within the egg (see Stebbings, 1988). This dilemma was resolved with the already-discussed equation of HMW MAPIC with dynein, and bolstered at about the same time by the discovery of a dynein-like motor in the nematode worm, Caenorhabditis elegans, (Lye et al., 1987) which was significant as this organism forms no motile cilia or flagella and possesses no axonemal dynein. Subsequently, as well as being purified from mammalian brain (Paschal et al., 1987) cytoplasmic dynein has also been obtained from other mammalian and vertebrate organs including liver and testis (Collins and Vallee, 1989). Also, in addition to sea urchins, it has been isolated from a range of invertebrate organisms including amoeboid protozoa (Euteneuer et al., 1988), squid, Dictyostelium, and hemipteran insects (Anastasi et al., 1990), and like kinesin the amounts of dynein in different tissues show considerable variation, but the degree of phylogenetic conservation remains to be fully assessed.

IX. THE CHARACTERIZATION OF MICROTUBULE MOTORS A. The Structure and Molecular Structure of Kinesin and Dynein Fundamental to an understanding of microtubule-based organelle transport is the requirement for an appreciation of the structure and function of microtubule motor proteins.

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Kinesin is a tetramer consisting of two heavy chains (120kD) and two light chains (62kD). Rotary shadowing prior to electron microscopy has shown kinesin to be an elongated rod-shaped molecule about 80nm in length (Hirokawa et al., 1989; Scholey et al., 1989). It has two globular heads at one end which have been seen to associate with microtubules, and a fan shaped tail at the other. The shaft connecting the two appears to possess a "kink" some 35nm from the head end. Using electron microscopy and monoclonal antibodies to kinesin heavy and light chains the submolecular domains have been identified, the heavy chains being mainly but not exclusively located to the globular heads and the light chains to the fan shaped tail (Hirokawa et al., 1989). Determination of the primary sequence of the heavy chain of kinesin, first of all from Drosophila (Yang et al, 1989) and then from squid (Kosik et al., 1990) as well as from other organisms including man (Navone et al., 1992), has shown there to be high evolutionary conservation. It has also confirmed the tripartite structure of the molecule consisting first of the amino terminal head domain, with its associated ATP-binding and microtubule-binding sites, which has been shown to be the minimum portion of the molecule which will produce movement in an in vitro assay (Yang et al., 1990), second a heptad repeat pattern characteristic of a rod-like a-helical coiled coil which probably affects the association between the heavy chains, and third a carboxyl-terminal region which is globular to an extent and which with part of the rod region has been shown to interact with the light chains. The kinesin light chains, the primary structure of which has also been investigated for a number of organisms (Cyr et al, 1991; Wedaman et al., 1993) have been shown to illustrate inter and intraspecies differences and such heterogeneity is thought to be important as the light chains are believed to represent the part of the molecule which possesses organelle interaction sites and a domain which may target and bind to specific organelles within a cell. Cytoplasmic dynein has more subunits than kinesin (at least 8) with two similar if not identical heavy chains (>400 kD) and also intermediate-sized accessory polypeptides in the 75—80 kD region. Electron microscopy has shown dynein to be composed of two globular heads connected by stalks to a common base, but the actual location of the different chains within this structure is not clear. As with kinesin, the globular head domains are thought to be the mechanochemical transducers which interact with the microtubules to produce motility, while the base domains which are unique to cytoplasmic forms of dynein (Mikami et al., 1993) are believed to attach to the translocating cargo. B. Force Generation and the Mechanochemistry of Microtubule Motors It is not clear as yet how motor proteins generate force, but one aim toward the understanding of this has been the accurate monitoring of movements generated by motor molecules, and in this regard a number of studies using in vitro assays have been very telling.

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First, the nature and velocity of the movements generated by kinesin and by dynein have been found to be quite different. With kinesin the monitoring of beads with great precision (Gelles et al., 1988) has shown them to maintain position above a particular protofilament as they are translocated along a microtubule, an observation confirmed on the one hand by the use of zinc-induced sheets of protofilaments in which adjacent protofilaments are arranged with alternate rather than common polarity but which support unhindered unidirectional translocation with kinesin (Kamimura and Mandelkow, 1992), and also by the fact that microtubules comprised of twisted protofilaments rotate during gliding on kinesin-coated slides, while those with parallel protofilaments do not (Ray et al, 1993). Dynein-driven retrograde translocation is quite different in both velocity and also in nature. In general dynein propels beads at a faster rate along microtubules than does kinesin. The movement tends to be intermittent rather than continuous, and it appears unlikely that the dynein motor confines itself to traversing a single microtubule protofilament. To obtain better temporal resolution than is possible with video, with its limited number of frames per second, Kamimura and Kamiya (1989) have imaged beads complexed to flagella microtubules onto photodiodes and so recorded nanometer displacements of approximately 300Hz thought to be produced by individual dynein strokes, the precise frequency being dependent on the concentration of ATP applied. Regarding the number of motors required to move a microtubule in an in vitro assay, it has been shown that the number of microtubules moving over a surface increases as the density of kinesin on the slide's surface is increased (Howard et al., 1989). Calculations from these observations have shown that a single kinesin molecule will move a microtubule, and attempts to measure the force generated have been made using "optical tweezers" (Kuo and Sheetz, 1993) and also the centrifugal microscope (Hall et al., 1993). These findings have allowed the conclusion that the kinesin molecule must be able to maintain its association with a microtubule during repeated attachment and detachment during nucleotide hydrolysis. Such studies are also of great significance to the movement of organelles within cells as they demonstrate that only a few motor molecules are required for translocation. They also facilitate studies of how motor proteins use energy from nucleotide hydrolysis to generate force. Central to the dilemma is what actually powers the cross-bridge cycle—conformational change within the motor protein, or thermal energy in which ATP hydrolysis rectifies the random fluctuations of the cross bridge allowing it to act as a "thermal ratchef (see Cordova et al., 1992) to produce force in a single direction. It is possible that different cytoskeletal motors are involved differently in the force-generating step and certainly the mechanochemical cycles of kinesin and dynein differ.

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Nucleotide Specificity and Drug Sensitivity of Kinesin and Cytoplasmic Dynein

Kinesin and cytoplasmic dynein are different, as we have seen, not only in structure and biochemistry but also in enzymatic properties, and their substrate specificities are quite different. Whereas kinesin has been shown to have broad substrate specificity, being capable of using all nucleotide triphosphates to a degree (Kuznetsov and Gelfand, 1986), although dynein hydrolyzes a range of nucleotide triphophosphates at high rates only ATP acts as a substrate for in vitro motility (Gibbons, 1986). Still further, in a comprehensive study Shimizu and coworkers (1991) tested the ability of a wide range of ATP analogues to be turned over by, and to support movement by kinesin and dynein. As a result they found that the two motors did indeed have distinct "nucleotide fingerprints" which, they suggest, could be used in conjunction with inhibitor studies as a means of identifying motors involved in particular forms of motility. An example in which this approach has been particularly useful is seen with the bidirectional movement in the giant syncytial freshwater amoeba, Reticulomyxa, where surprisingly the two directions of organelle transport which occur have identical usages of ATP analogues (Schliwa et al., 1991). Hence they appear to be driven by the same motor—^in which case the differences in direction would have to be affected by some modification of the motor. Many studies have been conducted of agents which inhibit microtubule-based motility, but most that inhibit kinesin, such as the ATPase inhibitors vanadate and EHNA, the nonhy drolyzable analogue of ATP, AMP PNP, and the alkylating reagent NEM also inhibit dynein—^albeit generally at lower concentrations.

X. FAMILIES OF MICROTUBULE MOTORS In the recent past it has become clear that the naive picture of there being two microtubule motors—one driving organelles in the anterograde direction and the other driving them in the retrograde direction, as one might have anticipated, is far too simplistic, and it is now clear that families of microtubule motors exist and this may indeed account for some of the exquisite selectivity of translocation which clearly occurs in cells. A. Genetic Approaches to Identifying and Characterizing Molecular Motors

Information obtained by the cloning and characterization of genes encoding known and putative motor proteins has shown that families of kinesins and probably dyneins exist. Following its purification, the kinesin heavy chain was cloned by screening a Drosophila cDNA library with the appropriate antibody, and the gene sequenced

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(Yang et al, 1989). This has subsequently led to the identification of an ever-increasing family of related proteins by molecular genetic analysis of mutants in a range of organisnis amenable to this approach (see Endow and Titus, 1992). Mutations resulting in defects in microtubule-based motility in these organisms were found to reside in genes encoding proteins similar to kinesin heavy chains, and specifically in its amino-terminal mechanochemical head domain. Indeed, the non-motor tail domains are quite different, as might perhaps be expected for the cargo end of the molecules. Knowledge of the motor domain sequence has led to the production of antibodies which in turn have been used to identify further, and indeed large numbers of kinesin-like proteins in organisms such as Drosophila. Many of the kinesin family have been found to be required for cell division and have been implicated in mechanisms of mitosis or meiosis. Certain members of the kinesin family, however, such as that specified by unc-104 in the nematode Caenorhabditis elegans have been shown to be required for axonal transport (Hall and Hedgecock, 1991). Moreover, while apparently being involved in the transport of synaptic vesicles, it is not involved in the translocation of other membrane-bound organelles. Similarly, five members of the kinesin family (KI 1—5) have been identified in the mouse central nervous system (Aizawa et al, 1992), and some if not all of them are thought to be involved in membrane organelle transport. Regarding function and functional comparisons, whereas kinesin has been shown to drive movement towards the + or the fast-growing ends of microtubules, motor activity has only actually been demonstrated so far with a few kinesin-related proteins. Certain examples, however, which have been expressed in bacteria have astonishingly been shown to drive movement towards the minus ends of microtubules, so that possession of the kinesin motor motif must clearly not be taken to indicate motility and in particular motility in a specified direction. Similar approaches to those for kinesin may result in the identification of novel dyneins—now that sea urchin dynein heavy chain (Gibbons et al., 1991; Ogawa, 1991) and also rat brain cytoplasmic dynein (Mikami et al., 1993) have been sequenced, and it will be interesting to discover how heterogeneous cytoplasmic dynein is.

XI. THE REGULATION OF MICROTUBULE MOTORS AND THE CONTROL OF DIRECTIONALITY OF ORGANELLE TRANSPORT Examples of the translocation of specific organelles to particular locations in cells have already been introduced and discussed, and such movements are clearly precisely regulated—probably at a number of levels. A number of questions are crucial to understanding how microtubule-based organelle translocation is regulated in vivo, including what determines the association of motor proteins with their vesicular cargo, what factors influence the activities of the different translocation

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motors, and to what extent the microtubule substrate modulates the activities of the motors. A. cAMP and Calcium in the Regulation of Microtubule-Based Motility Cases where intracellular movement occurs in distinct temporal events, as in fish chromatophores where a stimulus-response coupling results either in the retrograde transport and therefore the aggregation of pigment, or the anterograde movement to disperse it, lend themselves particularly well to the study of the regulation of translocation. The pigment granule movements in chromatophores are believed to be under the control of the sympathetic nervous system, and isolated melanophores have been shown to disperse pigment in response to (3 adrenergic agonists and to aggregate pigment on exposure to a adrenergic agonists. Moreover, a2 rather than a I adrenoreceptors are required for pigment aggregation which is significant since they act quite differently—a^ stimulation resulting in the elevation of intracellular Ca^"^ and the a2 response being a decrease in cAMP (see Exton, 1985). Experiments using the melanophores and indeed the chromatophores of various fish, with different degrees of demembranation have shown that pigment dispersion is c AMP dependent and requires phosphorylation of particular proteins by a protein kinase (Rozdzial and Haimo, 1986a), while pigment aggregation requires the activity of a Ca^"^ and calmodulin dependent phosphatase (Thaler and Haimo, 1990). Certain evidence has suggested therefore that both second messengers, cAMP and Ca^"^, influenced by different a adrenergic receptors are involved in the regulation of pigment transport, the direction of which, it has been suggested, may be controlled by the balance between cAMP-stimulated kinase activity and Ca^"*"stimulated phosphatase activity (Thaler and Haimo, 1992). The position is not totally clear, however, since other studies using melanophores of different species have shown that while there are changes in both Ca^^ and cAMP during experimentally-induced pigment aggregation, the dynamics of the former are not required while the dynamics of cAMP are closely linked to bidirectional intracellular movement. This shows that even in chromataphores second messenger regulation might differ. The question arises as to what substrate is regulated by the cAMP-dependent kinase and how this might be related to known motor proteins and those involved in pigment migration. In one study, of Tilapia melanophores, a 57 kD polypeptide has been shown to be phosphorylated during pigment dispersion, and ciliary axonemal dynein was seen to be activated by phosphorylation of a similarly sized associated polypeptide (Tash, 1989) suggesting that the same may be true of its cytoplasmic dynein counterpart. The extent to which phosphorylation of motor or motor-associated proteins might regulate motility remains to be determined, but such regulation could conceivably be affected by it influencing their binding to vesicular cargo and possibly modulat-

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ing their nucleotidase activity. Moreover, experiments have indicated that cell cycle control of microtubule-based membrane transport results from changes in the association of motors with membranes (Allen and Vale, 1991). In this regard, recent studies (HoUenbeck, 1993b) have shown that the kinesin heavy and light chains, as well as the kinesin-associated protein kinectin, are phosphorylated, although surprisingly perhaps not on the mechanochemical head; and this has lead to the suggestion that kinesin's phosphorylation state may influence its binding to organelles rather than acting directly on motor activity. B. The Association of Motors with Vesicles

Studies have shown that large amounts of microtubule motors exist in a soluble pool in cells (HoUenbeck, 1989), but what factors recruit particular motors to bind to specific organelles remains to be determined. Evidence that kinesin and dynein bind to membranes comes from studies of subcellular localization, immunofluorescence and in vitro assays of vesicle motility. Using antibodies specific for kinesin heavy and light chains and immunofluorescence microscopy kinesin has been localized to Triton X-100 soluble punctate structures in a range of neuronal and non-neuronal cells (Pfister et al., 1989) and a similar distribution has been seen on punctate structures in squid axoplasm (Brady etal., 1990). To localize the kinesin specifically to anterograde or retrograde-moving organelles, antibodies were applied to ligated peripheral nerves, a procedure which helps to distinguish organelles moving in the two directions. In such studies, kinesin was found to accumulate on the proximal sides of the ligatures much more than on the distal sides, suggesting that kinesin is mainly associated with organelles moving in the anterograde direction, and supporting the hypothesis that kinesin is an anterograde motor. The distribution was not, however, exclusive to organelles moving in the anterograde direction, and indeed cytoplasmic dynein also localizes to both anterogradely and retrogradely moving organelles (Hirokawa et al., 1990). In an effort to determine more precisely the nature of the membrane-bounded organelles to which kinesin binds, bovine brain homogenates were fractionated prior to immunoblotting and immunoelectron microscopy. In these studies kinesin was found to copurify with some fractions and in particular those containing synaptic vesicles, mitochondria and coated vesicles, but was not found to be associated with Golgi membranes or nuclei—suggesting that kinesin exhibits organelle-specific binding (Leopold et al., 1992). If kinesin is bound with high aflTinity to anterogradely moving vesicles the converse would appear to be true for cytoplasmic dynein which has been immunolocalized to lysosomes in the perinuclear region of fibroblasts—^an observation consistent with these organelles associated with the later stages of the endocytic pathway being regulated by a retrograde motor such as dynein (Lin and Collins, 1992).

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For a vesicle to move in a particular direction it requires to be associated with the correct motor, and therefore interaction between a motor and the organelle surface is a crucial factor in the control of directionality of translocation. Using brain microsomes Yu and coworkers (1992) have investigated the binding of kinesin and cytoplasmic dynein. They discovered that competition occurred between the motors for binding to vesicles in the absence of ATP, indicating a shared site, but that when ATP was present there appeared to be two distinguishable sites on the organelles. They then went on to consider three possible schemes which might account for motor-vesicle interaction. All early in vitro assays involved applying motor-coated beads to microtubule substrates. The adsorption of such motor proteins to beads is clearly non-specific, and individual beads in cytosol containing both kinesin and cytoplasmic dynein show bidirectional movement. This contrasts with the movement of membranebounded organelles which move in only one direction in similar cytosols—emphasizing the importance and specificity of the motor—organelle association. Some studies have demonstrated the importance of cytosolic factors (Schroer et al., 1988) and have suggested that soluble kinesin and dynein exchangefi-eelywith organellebound motors. Such ephemeral associations between motors and organelles are an attractive possibility as they suggest a simple means by which organelle directionality might be controlled. More recently, however, some doubt has been expressed as to the importance of cytosolic factors since in in vitro assays it appears that the cytosolic factors might simply block adsorption onto glass substrates of organelles which would otherwise translocate along the microtubules . Schnapp and coworkers (1992) have shown that such adsorption can be prevented by treating glass substrates with casein, and thereafter the addition of cytosol did not increase organelle movement. They also went on to investigate the nature of the associations between motors and organelles, confirming that kinesin and dynein are probably differently attached to organelles. Whereas dynein is extracted with 300mM KI, implying a possible ionic interaction with the organelles, kinesin remains bound even when exposed to 900mM KI, suggesting that other than ionic interactions are involved. It remains possible, therefore, that the binding of motors to organelles may be mediated by integral membrane proteins, a possible candidate in the case of kinesin being the recently identified 160kD protein extracted from detergentstabilized microsomes which has been named kinectin (Toyoshima et al., 1992). On the other hand, they may interact directly with membrane lipids, as happens with myosin I (Adams and Pollard, 1989). In addition, the finding of irreversible attachment of motors to organelles and the many observations of constant switching between plus-end and minus-end directed transport in cultured cells (Herman and Albertini, 1984), a phenomenon also mimicked in vitro (Vale et al., 1992) means that further mechanisms will have to be proposed to account for control of directionality of microtubule motor-driven movement.

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HOWARD STEBBINGS C. The Influence of Structural MAPs on Microtubule Motor Activity

While in vitro studies have concentrated on the interaction of motor proteins with MAP-free microtubule substrates a plethora of MAPs associate in a selective way with microtubules in nature, and could of course be important modulators of microtubule-based translocation in vivo. It is probable in any cell, and certainly the case in neurons which have been the subject of considerable attention, that different microtubules have distinctly different MAP complements, and this may influence the ability of a microtubule to act as a substrate for motor protein activity. It is known, for example, that axonal microtubules have quite different MAPs from dendritic microtubules (Matus, 1990) and the concept that there are preferred, perhaps MAP-free sections of microtubules as substrates for translocation is not new (Miller et al., 1987). More specifically, experiments have shown that MAP2 inhibits microtubule gliding on dynein-coated coverslips and it has been suggested that this is due to the fact that MAP2 and the motor protein both interact with the C-termini of a and (3 tubulin (Paschal et al, 1989). Recent experiments, however, have repeated similar studies using both MAP2 and tau, which have similar microtubule-binding domains but very different lengths of projection side arms. These have shown that while MAP2 inhibits kinesin and dynein-driven motility, tau does not, suggesting that it is the side arm, rather than the binding domains that are inhibitory (Lopez and Sheetz, 1993). These authors go on to suggest that since the MAP2 side arm is longer (100-200nm) than either dynein or kinesin molecules simple steric hindrance could account for the inhibition observed. In vivo the rigidity of the MAP side arm could also be an important factor, perhaps controlled in turn by MAP phosphorylation (Hagestedt et al., 1989).

XII. PERSPECTIVES The significance of the possession by a cell of multiple microtubule motors, beyond the requirement for components to travel in one or other direction along the microtubules remains to be elucidated (see Goldstein, 1991; Endow, 1991; Vale, 1992). Whether the answer relates to the variety of cellular systems in which the motor activities occur, the multitude of components which are driven by such motors, control in the form of "fine-tuning" of the translocation event or some related parameters of the cell physiological process encompassing the microtubulebased translocation, are questions being addressed. Such understandings are of course a prerequisite to the focused manipulation of cell function—not least in the area of medical science.

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Lin, S.X.H,, & Collins, C.A. (1992). Immunolocalization of cytoplasmic dyneinto lysosomes in cultured cells. J. Cell Sci. 101,125-137. Lippincott-Schwartz, J., Donaldson, J.G., Schweizer, A., Berger, E.G, Hauri, H-R, Yuan, L.C., & Klausner, R.D. (1990). Microtubule-dependent retrograde transport of proteins into the ER in the presence of brefeldin A suggests an ER recycling pathway. Cell 60, 821-836. Lopez, L.A., & Sheetz, M.R (1993). Steric inhibition of cytoplasmic dynein and kinesin motility by MAP2. Cell Motil. Cytoskel. 24, 1-16. Lye, R.J., Porter, M.E., Scholey, J.M., & Mcintosh, J.R. (1987). Identification of a microtubule-based cytoplasmic motor in the nematode C. elegans. Cell 51, 309-318. Matteoni, R., & Kreis, T.E. (1987). Translocation and clustering of endosomes and lysosomes depends on microtubules. J. Cell Biol. 105, 1253-1265. Matus, A. (1990). Microtubule-associated proteins. Curr. Opin. Cell Biol. 2,10-14. McNiven, M.A., & Ward, J.B. (1988). Calcium regulation of pigment transport in vitro. J. Cell Biol. 106, 111-125. Mikami, A., Paschal, B.M., Mazumdar, M., & Vallee, R.B. (1993). Molecular cloning of the retrograde transport motor cytoplasmic dynein (MAP IC). Neuron 10, 787-796. Miller, R.H., Lasek, R.J., & Katz, M.J. (1987). Preferred microtubules for vesicle transport in lobster axons. Science 235, 220-222. Navone, R, Niclas, J., Hom-Booher, N., Sparks, L., Bernstein, H.D., McCaffrey, G., & Vale, R.D. (1992). Cloning and expression of a human kinesin heavy chain gene: Interaction of the COOH-terminal domain with cytoplasmic microtubules in transfected CV-1 cells. J. Cell Biol. 117, 1263—1275. Ogawa, K. (1991). Four ATP-binding sites in the midregion of the p heavy chain of dynein. Nature 352, 643-645. Paschal, B.M., Obar, R.A., & Vallee, R.B. (1989). Interaction of brain cytoplasmic dynein and MAP2 with a common sequence at the C terminus of tubulin. Nature 342, 569-572. Paschal, B.M., Shpetner, H.S., & Vallee, R.B. (1987). MAP IC is a microtubule-activated ATPase which translocates microtubules in vitro and has dynein-like properties. J. Cell Biol. 105, 1273-1282. Paschal, B.M., & Vallee, R.B. (1987). Retrograde transport by the microtubule-associated protein MAP IC. Nature 330, 181-183. Pfister, K.K., Wagner, M.C., Stenoien, D.L., Brady, S.T., & Bloom, G.S. (1989). Monoclonal antibodies to kinesin heavy and light chains stain vesicle-like structures, but not microtubules, in cultured cells. J. Cell Biol. 108, 1453-1463. Ray, S., Meyhofer, E., Milligan, R.A., & Howard, J. (1993). Kinesin follows the microtubule's protofilament axis. J. Cell Biol. 121, 1083-1093. Rozdzial, M.M., & Haimo, L.T. (1986a). Reactivated melanophore motility: Differential regulation and nucleotide requirements of bidirectional pigment granule transport. J. Cell Biol. 103,2755-2764. Rozdzial, M.M., & Haimo, L.T. (1986b). Bidirectional pigment granule movements of melanophores are regulated by protein phosphorylation and dephosphorylation. Cell 47, 1061—1070. Saxton, W.M., Porter, M.E., Cohn, S.A., Scholey, J.M., Raff, E.C., & Mcintosh, J.R. (1988). Drosophila kinesin: Characterization of microtubule motility and ATPase. Proc. Natl. Acad. Sci. USA 85, 1109-1113. Schliwa, M., Shimizu, T., Vale, R.D., & Euteneuer, U. (1991). Nucleotide specificities of anterograde and retrograde organelle transport in Reticulomyxa are indistinguishable. J. Cell Biol. 112, 1199-1203. Schnapp, B.J., Reese, T.S., & Bechtold, R. (1992). Kinesin is bound with high affinity to squid axon organelles that move to the plus-end of microtubules. J. Cell Biol. 119, 389-399. Scholey, J.M., Heuser, J., Yang, J.T., & Goldstein, L.S.B. (1989). Identification of globular mechanochemical heads of kinesin. Nature 338, 335-357. Scholey, J.M., Porter, M.E., Grissom, P.M., & Mcintosh, J.R. (1985). Identification of kinesin in sea urchin eggs, and evidence for its localization in the mitotic spindle. Nature 318,483-486.

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Schroer, T.A., Schnapp, B.J., Reese, T.S., & Sheetz, M.P. (1988). The role of kinesin and other soluble factors in organelle movement along microtubules. J. Cell Biol. 107, 1785—1792. Shimizu, T., Furusawa, K., Ohashi, S., Toyoshima, Y.Y, Okuno, M., Malik, R, & Vale, R.D. (1991). Nucleotide specificity of the enzymatic and motile activities of dyneip, kinesin, and heavy meromyosin. J. Cell Biol. 112, 1189-1197. Stebbings, H. (1986). Cytoplasmic transport and microtubules in telotrophic ovarioles of hemipteran insects. Int. Rev. Cytol. 101, 101-123. Stebbings, H. (1988). Cytoplasmic dynein graduates. Nature 336, 14—15. Stebbings, H., & Hunt, C. (1983). Microtubule polarity in the nutritive tubes of insect ovarioles. Cell Tissue Res. 233, 133-141. Stebbings, H., & Hunt, C. (1987). The translocation of mitochondria along insect ovarian microtubules from isolated nutritive tubes: A simple reactivated model. J. Cell Sci. 88,641-648. Suprenant, K.A., Tempero, L.B., & Hammer, L.E. (1989). Association of ribosomes with in vitro assembled microtubules. Cell Motil. Cytoskel. 14,401—415. Tash, J.S. (1989). Protein phosphorylation: The second messenger signal transducer of flagellar motility. Cell Motil. Cytoskel. 14, 332-339. Terasaki, M., Chen, L.B., & Fujiwara, K. (1986). Microtubules and the endoplasmic reticulum are highly interdependent structures. J. Cell Biol. 103, 1557-1568. Thaler, CD., & Haimo, L.T. (1990). Regulation of organelle transport in melanophores by calcineurin. J. Cell Biol. Ill, 193S^1948. Thaler, CD., & Haimo, L.T. (1992). Control of organelle transport in melanophores: Regulation of Ca ^ apd cAMP levels. Cell Motil. Cytoskel. 22, 175-184. Toyoshima, I., Yu, H., Steuer, E.R., & Sheetz, M.P. (1992). Kinectin, a major kinesin-binding protein on ER. J. Cell Biol. 118, 1121-1131. Travis, J.L., & Bowser, S.S. (1986). Anew model of reticulopodial motility and shape: Evidence for a microtubule-based motor and an actin skeleton. Cell Motil. Cytoskel. 6, 2-14. Vale, R.D. (1992). Microtubule motors: Many new models off the assembly line. Trends Biochem. Sci. 17,300-304. Vale, R.D., & Hotani, H. (1988). Formation of membrane networks in vitro by kinesin-driven microtubule movement. J. Cell Biol. 107, 2233-2241. Vale, R.D., Malik, F., & Brown, D. (1992). Directional instability of microtubule transport in the presence of kinesin and dynein, two opposite polarity motor proteins. J. Cell Biol. 119,1589-1596. Vale, R.D., Reese, T.S., & Sheetz, M.P. (1985). Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility. Cell 42, 39-50. Wedaman, K.P., Knight, A.E., Kendrick-Jones, J., & Scholey, J.M. (1993). Sequences of sea urchin kinesin Ught chain isoforms. J. Mol. Biol. 231, 155-158. Waterman-Storer, CM., Sanger, J.W., & Sanger, J.M. (1993). Dynamics of organelles in the mitotic spindle of living cells: Membrane and microtubule interactions. Cell Motil. Cytoskel. 26,19-39. Wedaman, K.P., Knight, A.E., Kendrick-Jones, J., & Scholey, J.M. (1993). Sequences of sea urchin kinesin light chain isoforms. J. Mol. Biol. 231, 155-158. Weisenberg, R., & Taylor, E.W. (1968). Studies on ATPase activity of sea urchin eggs and the isolated mitotic apparatus. Exp. Cell Res. 53, 372—384. Yang, J.T., Laymon, R.A., & Goldstein, L.S.B. (1989). A three-domain structure of kinesin heavy chain revealed by DNA sequence and microtubule binding analyses. Cell 56, 879-889. Yang, J.T., Saxton, W.M., Stewart, R.J., Raff, E.C, & Goldstein, L.S.B. (1990). Evidence that the head of kinesin is sufficient for force generation and motility in vitro. Science 249,42-47. Yu, H., Toyoshima, I., Steuer, E.R., & Sheetz, M.P. (1992). Kinesin and cytoplasmic dynein binding to brain microsomes. J. Biol. Chem. 267, 20457-20464.

ROLE OF THE CYTOSKELETON IN THE DEVELOPMENT OF EPITHELIAL POLARITY

Detlev Drenckhahn, Thomas Jons, Bernd Puschel, and Frank Schmitz

I. Introduction 11. Role of the Membrane Cytoskeleton in Epithelial Polarity ! . . A. Polarized Distribution ofAnkyrin and Protein 4.1 B. Associationof Ankyrin withNa , K"^-ATPase C. Associationof Ankyrin with Adhesion Proteins D. PartialReversalof Polarity in Certain Endoepithelial Cells III. Binding Sites between Integral Membrane Proteins and the Membrane Cytoskeleton A. Ankyrin B. Protein 4.1 IV. Roleof Microtubules in Cell Polarity A. Intracellular Transport of Apical and Basolateral Membrane Proteins . .

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 141-165 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 141

142 144 144 147 150 151 151 151 152 154 154

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B. Uniform Alignment and Polarity of Microtubules as Structural Basis for Vectorial Intracellular Transport 154 C. Alteration of Epithelial Polarity by Microtubule-Disrupting Drugs . . . .155 D. Transport Barrier Hypothesis 158 E. Associationof Apical Carrier Vesicles with Dynein 159 V. Role of the Cytoskeleton in Development of Epithelial Polarity; A Unifying Hypothesis 160 References 161

I. INTRODUCTION The plasma membrane is a complex, dynamic assembly of a large variety of lipids and proteins that allovv^s the cells of the body to generate and maintain a specific internal chemical milieu different from their environment. In addition, cells can physically interact with each other and with extracellular substrates thereby reaching and maintaining a specific position within the body's tissues. Many cells in situ and in tissue culture {in vitro) become functionally and morphologically polarized. Typical examples of polarized cell types are the simple exoepithelial cells lining the lumen of the gut (and its appending exocrine glands), the urogenital system (including kidney tubules), the respiratory surface and the skin with its exocrine glands. The plasma membrane surface of these polarized epithelial cells can be divided in apical and basolateral domains (Figure 1). The apical membrane domain faces the body's exterior and differs in many important aspects from the basolateral domain that faces the body's interior. The basolateral domain can be further separated in a lateral and basal surface. The lateral surface is located at the interface between adjacent epithelial cells, whereas the basal surface is attached to the underlying extracellular matrix (basal lamina). Epithelial cells lining internal fluid-filled compartments (endoepithelium) such as blood vessels, ventricles of the brain, the inner surface of the eye, and follicles of the thyroid gland are also often polarized cells with the apical surface facing the fluid compartment and the basolateral domain bordering on the intercellular space and on the connective tissue proper. The apical surface of many polarized epithelia is enlarged several-fold by numerous finger-like projections termed microvilli. These projections are stabilized by an internal axial supportive skeleton of actin filaments that is stabilized by different bundling proteins such as villin and fimbrin (Burgess, 1987; Bement and Mooseker, this treatise. Volume 3). This core bundle is connected to the plasma membrane by linking proteins, one of which is myosin I. Enlargement of the basolateral cell surface mainly occurs via lateral folds (microplicae) and basal invaginations. The most striking functional feature of polarized epithelial cells is their capability of directing the transport of ions and small organic molecules across the epithelial layer thereby generating electrochemical gradients. The vectorial transcellular

Cytoskeleton and Epithelial

Polarity

143

apical HgO

HCO3Na*,Glucose - Cotransporter

lateral Glucose Transporter

Glucose basal Figure 1. Examples of molecular heterogeneity of the apical and basolateral plasma membrane in acid-secreting intercalated cells (a) of kidney collecting duct and in the intestinal epithelium (b).

transport of molecules depends largely on an asymmetric distribution of pumps, channels, and carriers to either the apical or the basolateral pole of the transporting epithelial cells. Two typical examples of cooperation between apical and basolateral membrane proteins in the vectorial transport of ions and organic solutes across the epithelial layer are depicted in Figure 1. In the acid-secreting epithelial cells of the stomach (parietal cells) and the secretory epithelial cells of the kidney collecting duct and turtle urinary bladder (intercalated cells, dark cells), basolateral anion exchangers (AE) allow the removal of HCOj-ions that are left behind in the cytosol due to the activity of a proton pump that translocates H"^ ions across the apical plasma membrane (Steinmetz, 1985; Forte and Wolosin, 1987; Brown, 1989). In most epithelial cells (except certain epithelia of the nervous system) the sodium pump (Na"*", K"^-ATPase) is located at the basolateral membrane surface where it provides the major driving force for resorption of various ions and solutes via sodium-coupled cotransporters (Pedersen and Carafoli, 1987; Mercer 1993). These cotransporters are mainly located at the apical cell surface as shown in Figure 1 for the sodium-glucose cotransporter. This striking asymmetric distribution of certain membrane proteins to either the apical or basolateral membrane domain raises the question of how this molecular polarity is generated and how it is maintained (for reviews see Rodriguez-Boulan andSalas, 1989; Rodriguez-Boulan and Nelson, 1989; Hubbard etal, 1989;Caplan and Matlin, 1989; Simons and Wandinger-Ness, 1990; Mostov et al., 1992; Mays etal., 1994).

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Tight junctions (zonulae occludentes), which separate the apical from the basolateral membrane, are thought to serve as a diffusion barrier within the lipid bilayer, thereby preventing apical membrane proteins from diffusing laterally into the basolateral membrane domain, and vice versa (Stevenson et al, 1988; Citi, 1993). Thus, tight junctions appear to be of particular importance for maintaining cellular polarity. However, tight junctions alone cannot explain why certain membrane proteins show a rather uneven distribution along the basolateral membrane surface. The basolateral anion exchanger in the acid secreting intercalated cells of the kidney collecting duct (kidney AEl), for example, is mainly concentrated at pleated areas of the basolateral membrane and does not occur in significant amounts in the plain areas of the plasma membrane (Drenckhahn and Merte, 1987). Another example for restriction of an integral membrane protein to subdomains of the basolateral surface is the Na"^,K"^-ATPase which is confined to the lateral membrane surface of all exoepithelial cells and does not occur along the basal membrane domain that abuts onto the basal lamina (Koob et al, 1987; Morrow et al., 1989). Finally, tight junctions are absent from several polarized cell types such as neurons, photoreceptors, certain vascular endothelial cells (Stolz et al., 1992), and the amniotic epithelium (King, 1982). Thus, further mechanisms must exist that are responsible for the polarized and uneven distribution of certain membrane proteins along the cell surface. In the present review we will summarize experiments and observations suggesting that connection of certain integral membrane proteins to components of the cytoskeleton may play a role in polarized retention or targeting of these proteins in epithelial cells.

II. ROLE OF THE MEMBRANE CYTOSKELETON IN EPITHELIAL POLARITY A.

Polarized Distribution of Ankyrin and Protein 4.1

One possible mechanism how particular membrane proteins could be placed and immobilized at specialized sites of the cell surface could be by linkage of these proteins to components of the membrane cytoskeleton. The erythrocyte membrane provides an example of how an integral membrane protein, namely the anion exchanger (termed AEl or band 3), is restricted in its lateral mobility by linkage to the membrane cytoskeleton (for reviews see Bennett, 1989; Alper, 1991; Niggli, this treatise. Vol. 1). AEl is anchored to the subplasmalemmal fibrous meshwork of actin and spectrin mainly by ankyrin. Ankyrin contains two main binding sites, one for (3-spectrin and the other for the cytoplasmic domain of AEl. A further site of linkage between the spectrin/actin meshwork and AEl is mediated by protein 4.1 which additionally serves to increase the affinity between spectrin and actin. The first indication that the membrane cytoskeleton may be involved in epithelial polarity came from immunocytochemical and immunoblotting studies on various transporting epithelial cells, such as the epithelium of the intestine, exocrine glands

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145

and the kidney. In these epithehal cells immunoreactive forms of ankyrin were found to be confined to the basolateral plasma membrane and to be absent from the apical membrane (Figures 2 and 3) (Drenckhahn et al, 1985; Drenckhahn and Bennett, 1987). Actin and isoforms of spectrin displayed a nonpolarized distribu-

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Figure 2. (a) Peripheral membrane proteins associated with parotid Na"^, K'^-ATPase of the pig parotid gland. Coomassie Blue-stained SDS-PAGE (lanes 1-2) and corresponding Western Blots (lanes 3-5). Lane 1: hunnan erythrocyte membrane proteins; lane 2: purified Na"^, K"^-ATPase vesicles; lanes 3-5: corresponding to lane 2 blotted on nitrocellulose and probed with antibodies raised against brain sprectrin (3), erythrocyte ankyrin (4), and actin (5). (b) Localization of ankyrin along the lateral plasma membrane and basal infoldings of a striated duct in the pig parotid gland. Bar: 10 |Lim. (c-e) Morphology of peripheral membrane proteins stripped off Na"^, K"^-ATPase vesicles. Note spectrin-like tetramers, one of which (e) is associated with an ankyrin-like particle. (Compiled from Koob et al., 1990; slightly modified.)

146

Cytoskeleton and Epithelial Polarity

147

tion in these cells and were detected at both the apical and basolateral membrane domain. In the acid secreting intercalated cells of kidney collecting duct and in the dark cells of the turtle urinary bladder ankyrin was found to colocalize with spectrin andAEl at the basolateral cell surface (DrenckhahnetaL, 1987,1993). This striking codistribution between an ankyrin-binding integral membrane protein (i.e., AEl) and ankyrin/spectrin strongly suggested lateral association of epithelial AEl via ankyrin to the spectrin-based membrane cytoskeleton at the basolateral surface of these transporting epithelial cells. In certain endoepithelial cells of the nervous system, such as the retinal pigment epithelium and the epithelium lining the choroid plexus of the brain ventricles the situation appears to be different in that ankyrin is found apically and not detected basolaterally (Gundersen et al., 1991;Marrsetal., 1993; Alperetal., 1994). As will be outlined later, this reversal of the distribution of ankyrin is accompanied by reversal of the localization of certain "basolateral" membrane proteins and secretory proteins that are expressed apically in these endoepithelial cell types. Vascular endothelial cells also form a polarized monolayer with functionally and biochemically different apical and basolateral membrane domains. These cells express significant amounts of protein 4.1 that it is concentrated at the lateral cell surface of the endothelial monolayer (Leto et al., 1986), again indicating a possible mechanism for linkage of certain membrane proteins to only one of the two major domains of the cell surface. Antibodies to protein 4.1 have also been observed to bind to the lateral cell surface of Madin-Darby canine kidney (MDCK) cells (Marchesi, 1993) raising the possibility of an additional, ankyrin-independent way of linkage of the spectrin-based membrane cytoskeleton to the lateral cell surface. The membrane attachment sites for ankyrin and protein 4.1 in epithelial cells lacking AEl (or closely related other anion exchangers) are largely unknown. However, during recent years two epithelial membrane proteins have been characterized that serve as direct or indirect binding sites for ankyrin. These are Na"^, K'^-ATPase, and E-cadherin. B. Association of Ankyrin with Na"^, K'^-ATPase

One of the most abundant and widely distributed integral membrane proteins restricted to the basolateral cell surface of virtually all exoepithelial cells is the Na^, K'^-ATPase. Na"^, K'^-ATPase is a transmembrane heterodimeric protein complex that consists of two subunits, the catalytic a-subunit and the non-catalytic PFigure 3. Immunoelectron microscopy showing localization of Na"*", K'^-ATPase (a) and ankyrin (b) in epithelial cells of the thick ascending limb of the rat kidney. Note, that both proteins colocalize along the lateral cell membrane and the infoldings of the basal cell surface. No label is seen in association with the zonula adherens (asterisks) and the basal plasma membrane abutting on the basal lamina (Bl). M, mitochondrion. Bar (a, b): 0.5 |im (part of Figure 3b is taken from Koob et al., 1987).

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DRENCKHAHN, JONS, PUSCHEL, and SCHMITZ

subunit. Na"^, K"^-ATPase is capable of generating a steep transmembrane gradient for sodium and potassium ions that is important for many basic cell functions such as generation of membrane potential, regulation of cell volume and sodium-coupled transport of various ions and organic solutes (Pedersen and Carafoli, 1987; Mercer, 1993). By light- and immunoelectron microscopy, it has been shown that Na"^, K'^-ATPase and ankyrin are precisely colocalized in kidney epithelial cells, where both proteins are present in identical microdomains of the basolateral plasma membrane (Koob et al., 1987, 1990; Morrow et al., 1989). These microdomains include both basal invaginations and the entire lateral cell surface; in contrast the basal surface and intercellular junctions were not labeled by antibodies to Na"^, K'^-ATPase and ankyrin (Figures 3 and 4). If the Na"*", K'^-ATPase could freely diffuse in the plane of the lipid bilayer one would expect an even distribution of the pump along the entire basolateral domain rather than a restriction to specialized areas of the plasma membrane. In view of the precise colocalization of Na"^, K'^-ATPase with ankyrin a possible explanation for this restricted distribution of Na"*", K"^ATPase would be its linkage, via ankyrin, to the subplasmalemmal spectrin lattice. Further support for this view comes from studies on two endoepithelial cells of the nervous system, the retinal pigment epithelium and the epithelium lining the choroid plexus (Figure 4). As mentioned above, ankyrin is restricted to the apical domain in these cells. Importantly, this reversal of the distribution of ankyrin is accompanied by reversal of the Na"^, K"^-ATPase which is absent from the basolateral domain in these cells and codistributes with ankyrin at the apical cell surface (Gundersen et al., 1991; Marrs et al., 1993; Alper et al., 1994). Direct evidence for binding of ankyrin to Na"^, K"^-ATPase was obtained by immunoprecipitation and sedimentation experiments in which Na"^, K'^-ATPase (purified from kidney outer medulla) was shown to bind erythrocyte ankyrin (Nelson and Veshnock, 1987; Koob et al., 1987; Morrow et al., 1989). Most likely, the binding site for ankyrin is located on the a-subunit. Binding of ankyrin to Na"^, K'^-ATPase could be inhibited by addition of the isolated cytoplasmic domain of erythrocyte AEl. A cluster of four amino acids (ALLK) on the large cytoplasmic loop of Na"^, K"^-ATPase appears to be essential for binding (Jordan et al., 1995) Further proof for the direct association of both proteins in vivo was provided by the following three observations: 1. The complex of Na^, K"^-ATPase, and ankyrin has been recovered from whole detergent extracts of the polarized MDCK cell line that have been fractionated in sucrose gradients and then coelectrophoresed in non-denaturing polyacrylamide gels (Nelson and Hammerton, 1989). 2. A native complex between Na"^, K"^-ATPase, ankyrin, spectrin, and actin has been purified from pig kidney and pig parotid gland, two tissues particularly rich in Na"^, K"^-ATPase (Koob et al., 1990). Examination of the peripheral membrane proteins stripped off the isolated Na"^, K'^-ATPase by high ionic strength showed that the "pump" is associated with typical spectrin-like

Cytoskeleton and Epithelial Polarity

149 Retinal pigment epithelium

Choroid plexus epithelium

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Figure 4, Schematic drawing illustrating the organization of cytoskeleton-membrane complexes in (a) polarized exoepithelial cells (e.g., epithelium of the kidney and intestine) and (b) in two examples of endoepithelial cells of the nervous system (retinal pigment epithelium, choroid plexus). Note codistribution of ankyrin and Na"^, K'^-ATPase along the basolateral cell surface in (a) and the apical location of both proteins in (b). Besides Na"^, K'^-ATPase, E-cadherin appears to be directly or indirectly associated with ankyrin and may play a role in stabilizing and assembling the membrane cytoskeleton and the Na"^, K'^-ATPase at the basolateral cell surface of exoepithelial cells. The absence of cadherins capable of assembling Na"*", K'^-ATPase-cytoskeleton complexes at the basolateral cell surface could contribute to accumulation of ankyrin and Na^, K'*'-ATPase at the apical cell surface of the choroid plexus and retinal pigment epithelium. Whether spectrin extends into the apical microvilli of the choroid plexus epithelium is not clear. Bl, basal lamina; ZA, zonula adherens; Z O , zonula occludens. For details, see text.

tetramers (100-200 )im in length), some of which were associated with ankyrin-like globules attached to the central region of the tetramer (Figures 2c, 2d, and 2e). Tetramers are generally believed to represent the functional subunit of the spectrin-based membrane cytoskeleton. 3. A complex between Na"^, K'*'-ATPase, ankyrin, and spectrin was obtained from cultured retinal pigment epithelium by chemical crosslinking and subsequent immunoprecipitation with antibodies to the a-subunit of Na"^, K"^-ATPase, ankyrin, and spectrin (fodrin; Gundersen et al., 1991).

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DRENCKHAHN, JONS, PUSCHEU and SCHMITZ C. Association of Ankyrin with Adhesion Proteins

In most polarized exoepithelial cells (e.g., epithelia of the kidney, intestine, and liver) the intercellular adhesion molecule E-cadherin (uvomorulin) is restricted to the lateral membrane domain. E-cadherin is a major adhesion molecule that mediates Ca^'^-dependent homophilic adhesion between the neighboring cells (Kemler, 1992). Ankyrin is also restricted to the lateral cell surface in these cells (see Section IIA). In detergent-solubilized membranes of MDCK cells, a fraction of about 30% of E-cadherin was found to cosediment with spectrin (fodrin), ankyrin, and Na"*", K'^-ATPase (Nelson et al., 1990). Ankyrin and spectrin, but not Na"^, K'^-ATPase, coimmunoprecipitated with antibodies to E-cadherin, indicating that separate complexes may exist formed between ankyrin/spectrin and either E-cadherin or Na"^, K^-ATPase (Nelson et al., 1990). The observation that only a fraction of E-cadherin appears to be associated with ankyrin/spectrin correlates with the ultrastructural distribution of these proteins: the bulk of E-cadherin is concentrated at the belt desmosomes (zonula adherens) (Boiler et al., 1985) where E-cadherin appears to be clustered by linkage to the actin-based cytoskeleton via a different set of linker proteins including plakoglobin and a- and p-catenin (Ozawa and Kemler, 1992; Kemler, 1992;Tsukitaetal., 1992). However, unlike E-cadherin, ankyrin appears to be absent from the zonula adherens (Figures 3 and 4; Koob et al., 1987). Neurons of the central and peripheral nervous system are also highly polarized cell types with three different main membrane domains (cell body, dendrite, axon). Different isoforms of ankyrin are restricted to the cell body and axon (Lambert and Bennett, 1993). Within the axon a distinct ankyrin isoform is confined to the nodes of Ranvier and an isoform of brain ankyrin to the intemodal section of the axonal plasma membrane (Kordeli and Bennett, 1991). This distribution of ankyrin may be partly explained by linkage of ankyrin to particular integral membrane proteins (ankyrin-binding glycoproteins, AGPs), one of which has been sequenced (AGP 186) and identified as a member of the axon-bundling neuronal adhesion protein neurofascin (Davis et al., 1993). Other axonal ankyrin-binding proteins appear to be the voltage-gated sodium channel (Srinivasan et al., 1992) and the Na"^, K'^-ATPase; both proteins are concentrated at the nodes of Ranvier. Association of the spectrin-based membrane cytoskeleton with intercellular adhesion proteins may play a critical role in early steps of the generation of cell polarity: Transfection of a mouse fibroblastic cell line (L-cell) with E-cadherin, not only resulted in intercellular contact formation between neighboring cells, but also caused significant assembly and concentration of spectrin and Na"*", K"*"-ATPase at this novel lateral membrane surface in this otherwise nonpolarized cell type. In cells transfected with truncated E-cadherin lacking half of the cytoplasmic domain, no intercellular adhesion occurred and no concentration of spectrin and Na"*", K'^-ATPase was observed along the lateral cell surface (McNeill et al, 1990).

Cytoskeleton and Epithelial Polarity D.

151

Partial Reversal of Polarity in Certain Endoepithelial Cells

As mentioned above, the retinal pigment epithelium and the epithelium lining the choroid plexus differ from all exoepithelial cells in that ankyrin is associated with the apical rather than the basolateral membrane surface and that these cell types lack E-cadherin. The reversal of the polarized distribution of ankyrin is accompanied by reversal of the distribution of the Na"^, K'^-ATPase* which, like ankyrin, is also restricted to the apical membrane domain and absent from the lateral domain (Figure 4). Interestingly, the choroid plexus also secretes transthyretin, ceruloplasmin, and cystatin C apically into the cerebrospinal fluid and all three proteins are secreted basolaterally by hepatocytes (Cole et al, 1987; Aldred et al., 1987a, 1987b). In the retinal pigment epithelium the situation is somewhat more complicated than in the choroid plexus since the apical domain of the pigment epithelium is an adhesive surface that contains the neuronal adhesion protein N-CAM and forms intercellular contacts with the outer segments of photoreceptors (Gundersen et al., 1991). However, in cultured pigment epithelial cells the apical polarity of Na"^, K'^-ATPase and ankyrin is retained. When these cells are transfected with E-cadherin both ankyrin and Na"^, K'^-ATPase become redistributed to the lateral cell surface (Andersson-Fisone et al., 1993). In the choroid plexus intercellular adhesion appears to be mainly mediated by B-cadherin, a Ca^"*"-dependent adhesion molecule, that, unlike E-cadherin, does not cause redistribution of spectrin and Na"^, K"^ATPase to the lateral cell surface when expressed in transfected L-cell fibroblasts (Marrs et al., 1993).

III. BINDING SITES BETWEEN INTEGRAL MEMBRANE PROTEINS AND THE MEMBRANE CYTOSKELETON A.

Ankyrin

Ankyrins are encoded by different genes and occur as different splice variants (Bennett, 1992; Lambert and Bennett, 1993). Ankyrins possess a -90 kD N-terminal portion that contains 24 copies of a 33 amino acid motif (ankyrin repeats). At least 12 of these motifs are required for high affinity binding of ankyrin to the cytoplamic domain of the erythrocyte anion exchanger (AEl). A cluster of four amino acids (ALLK) is essential for binding of ankyrin Na"^, K'^-ATPase. A motif related to the ALLK cluster is also present in the cytoplasmic domain of AEl where this sequence (ALLLK) turned out to be also important for ankyrin binding (Jordan etal., 1995).

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DRENCKHAHN, JONS, PUSCHEU and SCHMITZ B. Protein 4.1

/.

Binding Interface between Protein 4.1 and Anion Exchanger 1

Erythroid protein 4.1 contains in its 30 kD N-terminal domain the motif "LEEDY" that is duplicated in nonerythroid isoforms of protein 4.1. This motif binds to a complementary motif located on the cytoplasmic domain of all mammalian and avian anion exchangers so far sequenced (Figure 5; IRRRY, LRRRY, VRRRY, LKKRY; Jons and Drenckhahn, 1992). Glycophorin C, the other protein 4.1-binding membrane protein of erythrocytes also contains a related motif in its cytoplasmic domain and this consists of a triplet of three basic amino acids flanked by one hydrophobic residue and by glycine (YRHKG; Colin et al., 1986). Point mutation of the LEEDY motif on protein 4.1 (LREDY instead of LEEDY) significantly reduced the binding affinity of protein 4.1 to stripped erythrocyte inside-out membranes (Jons, Fenner, and Drenckhahn, unpublished observations).

Bands (AE1)

rvBJ,

Figure 5. Hypothetical model for protein 4.1 -mediated linkage of the actin-spectrin scaffold to the cytoplasmic domain of AE1 (band 3). In binding assays using the chymotryptic fragment of the cytoplasmic domain of AE1 only the binding site in position A is accessible (from Jons and Drenckhahn, 1992).

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2. Occurrence of the LEEDY/IRRRY Motif on other Cytoskeletal and Integral Membrane Proteins

The LEEDY motif (triplet of E/D flanked by two strongly hydrophobic residues, such as F, L, I, Y, V) occurs on several membrane-associated structural proteins (the number of such motifs per protein are shown in parentheses). Proteins were identified in the protein sequence database, SWISSPROT (Release 25): protein 4.1 / human, Xenopus vinculin, metavinculin / chicken annexin 4 / human, rat annexin 5 / human annexin 8 / human

(1-2) (1) (1) (2) (2)

a-spectrin / human

(1)

p-spectrin / human

(1)

a-actinin / human etc.

(1) (2)

laminbi /human lamin 1 / Xenopus peripherin / rat, mouse desmoplakins / human, bovine plectin / rat kinesin heavy chain / mouse

(2) (1) (8) (5) (4)

The IRRRY-motif (triplet of R/K flanked by two strongly hydrophobic residue, such as F, L, I, Y, V) occurs on the cytoplasmic domains of several different integral membrane proteins. Some examples of more than 30 membrane proteins found in the data base are: anion exchanger 1, 2, 3 / human etc. Na^, K"^-ATPase / human V-cadherin / human sodium channel / human ryanodine receptor / human etc. synaptobrevin / human. Torpedo platelet membrane glycoprotein 1 a / human

(2) (1) (1) (2) (3) (1) (1)

Although, it is uncertain whether apart from protein 4.1 and AE1 the LEEDY/IRRRY motifs on these cytoskeletal and integral membrane proteins play any role in cytoskeleton-membrane associations one should at least consider the possibility of such interactions. It should be mentioned that these motifs are also found on some proteins not related to possible cytoskeleton-membrane complexes, such as proteins of the extracellular matrix or exoplasmic domains of membrane proteins.

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IV. ROLE OF MICROTUBULES IN CELL POLARITY A.

Intracellular Transport of Apical and Basolateral Membrane Proteins

The Golgi apparatus plays a crucial role in sorting membrane proteins into different populations of carrier vesicles that are either targeted to cytoplasmic membrane systems (e.g., lysosomes) or are delivered to the apical or basolateral membrane surface, respectively. While several membrane proteins are vectorially delivered from the Golgi apparatus to either the apical or the basolateral domain, other proteins may reach their final site of destination by random delivery to both domains and subsequent transcytosis to the definite domain (Hubbard et al., 1989; Wessels et al., 1990; Wandinger-Ness et al., 1990; Mostov et al., 1992). Sorting by transcytosis appears to be the predominant sorting mechanism occurring in hepatocytes in which apical (biliary) proteins are first inserted into the basolateral membrane and subsequently undergo endocytosis and transport to the apical membrane (Bartles et al., 1987). A certain degree of missorting of apical membrane proteins to the basolateral surface (followed by transcytotic delivery to the apical surface) has also been described for some apical proteins in intestinal epithelial cell lines derived from colonic carcinoma (Matter et al., 1990; Le Bivic et al., 1990). B. Uniform Alignment and Polarity of Microtubules as Structural Basis for Vectorial Intracellular Transport

The molecular basis for vectorial transport of carrier vesicles within polarized epithelial cells appears to be provided by microtubules (Mts) in the intestinal epithelium and probably in all other polarized epithelial cells Mts are uniformly oriented towards a narrow space underneath the apical plasma membrane (more precisely underneath the apical terminal web). This "subterminal space" contains Mt-organizing properties and controls uniform orientation and polarity of Mts in these cells (Achler et al., 1989). After experimental depolymerization of Mts with nocodazole or colchicine some drug-stable short Mts are always retained within this apical space (Figure 6). As soon as the drugs are eliminated new Mts grow out from this apical zone and project towards the basal cell portions. Since the Mt-organizing center is located apically in these cells, Mts must contain their slowly growing ends (minus end) located apically and their fast growing ends (plus end) pointing basally. Ultrastructural determination of Mt polarity in MDCK cells confirmed that the bulk of Mts in the cellular midportion appear to have their plus ends pointing basally (Bacallao et al., 1989). However, the most important question of Mt polarity in these cells still needs to be thoroughly addressed; that is, what is the polarity of Mts in the space between the Golgi apparatus and the apical plasma membrane?

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Figure 6. Visualization of the orientation and nucleatlon site of microtubules (Mts) in the rat intestinal epithelium by immunofluorescence, (a) Control, (b) 1 h after application of colchicine, (c) 6 h after application of colchicine. Note apicobasal orientation of Mts and drug-Induced depolymerlzation beginning In the basal part of the cells. Drug-resistent Mts remain in the subterminal space suggesting that this narrow zone serves as Mt-organizing center. From the subterminal space Mts will grow out In a basal direction after removal (elimination) of the drug (from Achler et al., 1989; slightly modified). C. Alteration of Epithelial Polarity by MIcrotubule-DisruptIng Drugs

Experimental depolymerization of Mts (induced by colchicine, nocodazole, or vinblastine), both in cultured polarized epithelial cell lines and in vivo, indicates that Mts are particularly important for the vectorial transport of carrier vesicles from the Golgi apparatus toward the apical plasma membrane (Achler et al., 1989; Eilers et al, 1989). In contrast, the transport towards the basolateral cell surface does probably not depend on the Mt-system. Drug-induced depolymerization of Mts in cultured MDCK cells infected with the apically budding influenza virus resulted in random virus budding at both the apical and basolateral plasma membrane, whereas selective basolateral budding of VSV-virus remained unaffected (Rindler et al., 1987). In the intestinal epithelium of living mice and rats, depolymerization of Mts (induced by gavage of colchicine, vinblastine, or nocodazole) caused delivery of several newly synthesized apical membrane proteins to the basolateral cell surface. In contrast, basolateral targeting of the Na"^, K'^-ATPase appeared to be unaffected by disruption of Mts (Achler et al., 1989). Similar observations were made with an enterocyte cell line derived from colonic carcinoma (Gilbert et al., 1991). Mistargeting of apical membrane proteins was followed by the formation of typical brush borders at the basolateral cell surface (Figure 7) that contained all components of the microvillus cytoskeleton (i.e., villin.

-' *''.'''-*ttW ' ''.'

''/' . f^'-:" •

>

'

•

''*^',,

^v :.^^./ ^-r^

figure 7, Intestinal epithelium of the mouse 6 h after application of colchicine by stomach tube, (a) Electron micrograph showing an ectopic brush border located in an invagination of the basal cell surface (arrow), (b) Fluorescence micrograph of a corresponding 1-jam-thick plastic section of the epithelium stained with an antibody to villin, an actin-crosslinking protein of microvilli. Note the numerous ectopic brush borders, one of which is indicated by an arrowhead. Bars: (a) 1 ^im; (b) 10 |nm (from Achler et al., 1989; slightly modified). 156

apical

lateral

bTV-f

^2h

BB-vacuole

24h

— apical membrane proteins — basolateral membrane proteins Figure 8. Schematic drawing illustrating changes of the microtubule (Mt) system and of cellular polarity in the intestinal epithelium (rat, mouse) after experimental Mt-depolymerization induced by application of colchicine (vinblastine) by stomach tube. Stages at 0 h, 6 h, 12 h, and 24 h after application of the drugs are shown. In untreated animals, Mts are uniformly oriented towards a narrow space underneath the apical terminal web (subterminal space) that contains Mt-organizing properties and controls polarized alignment of Mts. It is most likely that Golgi-derived transport vesicles containing apical membrane proteins (aTV) are vectorially guided from the Golgi apparatus (GA) to the apical cell surface by a dynein-mediated transport to the minus ends of the Mts in the subterminal space. Transport through the apical terminal web may be mediated by the actin-filament dependent motor protein myosin I. The cell nucleus (N) and the endoplasmic reticulum (ER) may be kept in a basal position by a plus-end motor protein (binding of kinesin to the ER has been well established; Toyoshima et al., 1992). Certain basolateral transport vesicles (bTV) may also use a plus-end motor. After depolymerization of Mts apical membrane proteins become mistargeted to the basal cell surface, where ectopic microvilli and brush borders will subsequently arise. Reconstitution of cell polarity occurs by endocytosis of ectopic brush borders (BB) and subsequent transport of the BB-vacuoles to the apical cell surface (transcytotic pathway of sorting). (Drawn according to data published in Achler et al., 1989.) 157

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DRENCKHAHN, JONS, PUSCHEL, and SCHMITZ

fimbrin, myosin I) normally only found in association with apical microvilli (Achleretal., 1989). This dramatic change in cell polarity was fully reversible after elimination of the drugs: basolateral brush borders were subsequently endocytosed into large vacuoles, which were finally incorporated into the apical cell surface. These brush border-containing vacuoles behaved like intracellular islands of the apical plasma membrane in that they contained Mt-organizing properties on their cytoplasmic surface and served as recipient organelles for newly synthesized apical membrane proteins. Reconstitution of cell polarity depended on the reappearance and apicobasal reorganization of the Mt system (Figure 8). Similar observations were also obtained with a polarized growing intestinal epithelial carcinoma cell line (Gilbert etal., 1991). The most likely explanation for these observations is that apically destined carrier vesicles contain a receptor for a dynein-like motor protein that is responsible for Mt-dependent transport of these vesicles to the microtubular minus end located underneath the apical plasma membrane. Since the distance between the Golgi apparatus and the apical plasma membrane is considerably larger than that to the basolateral membrane a selective transport of carrier vesicles to the apical cell pole may help to prevent these carrier vesicles from accidental fiision with the lateral membrane. That exocytotic carrier vesicles (obtained from virus-infected MDCK cells) contain binding sites for microtubules has been shown for vesicles containing apical and basolateral membrane proteins (Van der Sluijs et al., 1990). As soon as Mts become depolymerized by colchicine or vinblastine, the apical carrier vesicles will lose their guiding structures and thus will become prone to random diffusion throughout the cell. A fraction of these carrier vesicles (40-60% according to [•^H]-fiicose autoradiography; Bennett et al., 1984; Blok et al., 1981; Ellinger et al., 1983) may still reach the apical membrane whereas the rest will fuse with the basolateral membrane and, thus, deliver apically destined proteins to the basolateral cell surface. Such mistargeted apical proteins probably include proteins that nucleate polymerization of actin filaments thereby initiating paradox microvillus formation at the lateral cell surface (Achler et al., 1989). A central conclusion drawn from these experiments is that apical carrier vesicles canfiisewith both the apical and basolateral plasma membrane whereas basolateral carrier vesicles are only able to fiise with the basolateral and not the apical membrane domain. This conclusion is fiirther supported by recent identification of both Rab-like (Rab 8) and trimeric GTP-binding proteins that appear to be involved in domain-specific targeting and exocytosis (Huber et al., 1993; Pimplikar and Simons, 1993; Zerial and Stenmark, 1993; Stow and de Almeida, 1993). D. Transport Barrier Hypothesis

Targeting of apical membrane proteins appears to occur in two steps. The first step is probably mediated by Mts that guide carrier vesicles to the apical subterminal

Cytoskeleton and Epithelial Polarity

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space. Since Mts rarely project into the terminal web (which underlies the apical but not the basolateral plasma membrane) the bulk of apical carrier vesicles must be able to pass through this specialized submembranous cytoskeleton ("barrier") consisting of bundles of actin filaments (rootlets of microvilli) interconnected by spectrin and myosin. The transport through this barrier may be mediated by myosin I which has been observed by immunoelectron microscopy to be associated with the surface of vesicles within the terminal web (Drenckhahn and Dermietzel, 1988). Thus, it is tempting to speculate that myosin I binds only to apical carrier vesicles, thereby facilitating selective access of apical carrier vesicles to the lipid bilayer, whereas basolateral carrier vesicles might be excluded by such a mechanism. Recent evidence suggests that such an association between myosin I and carrier vesicles may already occur at the level of the Golgi apparatus (Path and Burgess, 1993; Path et al., 1993; Kroschewski et al., 1994). E. Association of Apical Carrier Vesicles with Dynein

Our hypothesis that apical carrier vesicles are transported from the Golgi apparatus towards the apical plasma membrane by a minus-end directed transport along Mts has been further substantiated by the following unpublished observations (Schmitz, Kraemer, and Drenckhahn, in preparation):

TOkO

Figure 9. Association of dynein with purified secretory granules (zymogen granules) from rat pancreas, (a) Electron microscopy of the zymogen granule fraction used in (b) for immunoblotting with an antibody against the 70 kD dynein intermediate chain (Steuer et al., 1990). Kinesin is not detected in this fraction. The bulk of kinesin was found in the zymogen granule-depleted supernatant (not shown). Bar: 1 jiim.

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(1) pancreatic acinar cells deliver their secretory granules (zymogen granules) vectorially from the Golgi apparatus towards the apical plasma membrane which contains the specific sites for exocytosis. Immunoblot analysis of purified secretory granules of the rat pancreas revealed the presence of dynein and the absence of kinesin (Figure 9). (2) Polarized MDCK cells infected with the apically budding influenza virus were homogenized, and a membrane fraction was purified in sucrose gradients that was enriched in apical carrier vesicles. Immunoblotting of vesicles containing cleaved influenza virus hemagglutinin (proteolytic cleavage of hemagglutinin into two fragments occurs after budding of the carrier vesicles from the Golgi apparatus) revealed cytoplasmic dynein as major motor protein associated with the hemagglutinin-containing carrier vesicles and, by double immunolabeling, dynein, but not kinesin, was observed to colocalize with hemagglutinin-containing vesicles in the apical cytoplasm of MDCK cells infected with influenza virus.

V. ROLE OF THE CYTOSKELETON IN DEVELOPMENT OF EPITHELIAL POLARITY; A UNIFYING HYPOTHESIS Development of epithelial polarity can be divided into three main stages, all of which are accompanied by the formation of specific assemblies of the cytoskeleton. First stage: Aprimary prerequisite for any kind of cellular differentiation (beyond the morula stage) is the formation of cellular contacts with the extracellular matrix (ECM). These cell-to-ECM contacts (focal contacts) typically contain clusters of ECM receptors of the integrin type which, on their cytoplasmic side, are tethered and stabilized by various components of the actin filament cytoskeleton. This complex interface formed between the actin filament system and the cytoplasmic domains of clustered ECM receptors contains several associated kinases that are involved in signal transduction, e.g., the a-isoform of protein kinase C, pp 60^^^, focal adhesion kinase 125 (pp 125^"^^), vasodilator-stimulated phosphoprotein (VASP) (for reviews see Burridge et al., 1988; Luna and Hitt, 1992; Zachary and Rozengurt, 1992). Second stage: The second messenger pathways induced by contact formation with the ECM may stimulate transcription of various proteins that are important for fiirther steps of the development of cellular polarity, such as the intercellular adhesion protein E-cadherin (uvomorulin). E-cadherin mediates tight intercellular contact formation and induces assembly of the spectrin-based membrane cytoskeleton along the new lateral (intercellular) membrane surface. The ankyrin/spectrin scaffold, in turn, may serve to capture and immobilize Na"*", K^-ATPase along the lateral cell surface. Third stage: Na"^, K'^-ATPase immobilized at the lateral plasma membrane might create an electrochemical gradient across the epithelial layer that could stimulate (by a still unknown mechanism) the final stages of polarity formation, such as the

Cytoskeleton and Epithelial Polarity

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formation of tight junctions and the translocation of the perinuclear Mt organizing center towards the apical cell pole. That application of external electric voltage gradients can cause reorientation of the Mt-system has been shown in cultured fibroblasts (Harris et al., 1990). As soon as Mts have obtained their apicobasal orientation apical membrane proteins will be selectively transported to the apical cell surface, thereby completing the polarized differentiation of the epithelial layer. This three-stage hypothesis for the development of polarity in exoepithelial cells may also hold true for hepatocytes except that Mts are not uniformly aligned in these cells (Feldm^nn, 1989). However, a mechanism has developed in hepatocytes that controls transcytotic delivery of apical membrane proteins to the apical (biliary) plasma membrane (Bartles et al., 1987). However, certain steps involved in transcytotic targeting appear to be also dependent on the Mt-system since experimental disruption of Mts in hepatocytes has been shown to result in disturbance of the polarized distribution of certain apical membrane proteins (Durand-Schneider et al., 1991; Sawadaetal., 1992). In the retinal pigment epithelium and the epithelium lining the choroid plexus stage 2 probably differs from that of exoepithelial cells in that the cells express B-cadherin or N-CAM instead of E-cadherin (Gundersen et al., 1991; Marrs et al., 1993). Thus, the absence of cadherins capable of assembling Na"*", K"^-ATPase-cytoskeleton complexes at the basolateral cell surface could contribute to the accumulation of ankyrin at the apical surface. However, the possibility remains that apical membrane binding sites for ankyrin occur in addition to those for the Na"^, K"*'-ATPase itself; these could include one of the ankyrin-binding glycoproteins. The partial reversal of the polarity of the choroid and pigment epithelium does not involve the Mt-system which appears to be also nucleated from the apical subterminal space in these cells (Rizzolo and Joshi, 1993; Alper et al., 1994). REFERENCES Achler, C , Filmer, D., Merte, C , & Drenckhahn, D. (1989). Role of microtubules in polarized delivery of apical membrane proteins to the brush border of the intestinal epithelium. J. Cell Biol. 109, 179-189. Aldred, A.R., Grimes, A., Schreiber, G., & Mercer, J.F. (1987a). Rat ceruloplasmin. Molecular cloning and gene expression in liver, choroid plexus, yolk sac, placenta, and testis. J. Biol. Chem. 262, 2875-2878. Aldred, A.R., Dickson, RW., Mariey, RD., & Schreiber, G. (1987b). Distribution of transferrin synthesis in brain and other tissues in the rat. J. Biol. Chem. 262, 5293-5297. Alper, S.L. (1991). The band 3-related anion exchanger family. Annu. Rev. Physiol. 53, 54^564. Alper, S.L., Stuart-Tilley, A., Simmons, C.R, Brown, D., & Drenckhahn D. (1994). The fodrin-ankyrin cytoskeleton of choroid plexus preferentially colocalizes with apical Na"^, K'^-ATPase rather than with basolateral anion exchanger AE2. J. Clin. Invest. 93, 1430-1438. Andersson-Fisone, C , Nabi, I., Mathews, A.R, Zurzolo, C , & Rodriguez-Boulan, E. (1993). Lateral localization of Na , K -ATPase in E-cadherin (L-CAM) transfected rat retinal pigment epithelial cells (RPE). J. Cell Biochem. Suppl. 17B, 270a.

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Bacallao, R., Antony, C, Karsenti, E., Stelzer, E., & Simons, K. (1989). The subcellular organization of Madin-Darby canine kidney cells during the formation of a polarized epithelium. J. Cell Biol. 109,2817-2832. Bartles, R.T., Feracci, H.M., Steiger, B., & Hubbard, A.L. (1987). Biogenesis of the rat hepatocyte plasma membrane in vivo: Comparison of the pathways taken by apical and basolateral proteins using subcellularfractionation.J. Cell Biol. 105, 1241-1251. Bement, W.M., & Mooseker, M.S. (Forthcoming). The Cytoskeleton of the Intestinal Epithelium. The Cytoskeleton, A Multi-Volume Treatise, Vol. 3, Cytoskeleton in Specialized Tissues (Hesketh, J.E., & Pryme, I.F., Eds.). JAI Press, Greenwich, CT. Bennett, V. (1989). The spectrin-actin junction of erythrocyte membrane skeletons. Biochim. Biophys. Acta 988, 107-121. Bennett, V. (1992). Ankyrins. Adaptors between diverse plasma membrane proteins and the cytoplasm. J. Biol. Chem. 267, 8703-8706. Bennett, G., Carlet, E., Wild, G., & Parsons, S. (1984). Influence of colchicine and vinblastine on the intracellular migration of secretory and membrane glycoproteins. III. Inhibition of intracellular migration of membrane glycoproteins in rat intestinal columnar cells and hepatocytes as visualized by light and electron microscope radioautography after H-fticose injection. Am. J. Anat. 170, 545-566. Blok, J., Ginsel, L.A., Molder-Stapel, A.A., Onderwater, J.J.M., & Daems, W.T. (1981). The effect of colchicine on the intracellular transport of H-fticose-labelled glycoproteins in the absorptive cells of cultured human small-intestinal tissue. An autoradiographical and biochemical study. Cell Tissue Res. 215, 1-12. Boiler, K.D., Vestweber, D., & Kemler, K. (1985). Cell-adhesion molecule uvomorulin is localized in the intermediate junctions of adult intestinal epithelial cells. J. Cell Biol. 100, 327-332. Brown, D. (1989). Vesicle recycling and cell-specific ftinction in kidney epithelial cells. Ann. Rev. Physiol. 51, 771-784. Burgess, D.R. (1987). In: Advances in Cell Biology. Vol. 1 (Miller, K. R., ed.), pp. 31-58. JAI Press Inc., Greenwich, CT. Burridge, K., Fath, K., Kelly, T, Nuckolls, G., & Turner, C. (1988). Focal adhesion: Transmembrane junctions between the extracellular matrix and the cytoskeleton. Annu. Rev. Cell Biol. 4,487—525. Caplan, M., & Matlin, K.S. (1989). In: Functional Epithelial Cells in Culttire (Matlin, K. S., & Valentich, J. D., eds.), pp. 71-127. Liss, New York. Citi, S. (1993). The molecular organization of tight junctions. J. Cell Biol. 121,485-489. Cole, T, Dickson, RW., Esnard, F., Averill, S., Risbridger, G.R, Gauthier, F., & Schreiber, G. (1987). The cDNA structure and expression analyses of the genes for the cysteine proteinase inhibitor cystatin C and for beta 2-microglobulin in rat brain. Eur. J. Biochem. 186, 35-42. Colin, Y., Rahuel, C, London, J., Romeo, R-H., d'Auriol, L., Galibert, R, & Cartron, J.-R (1986). Isolation of cDNA clones and complete amino acid sequence of human erythrocyte glycophorin C. J. Biol. Chem. 261, 229-233. Davis, J.Q., McLaughlin, T., & Bennett, V. (1993). Ankyrin-binding proteins related to nervous system cell adhesion molecules: Candidates to provide transmembrane and intercellular connections in adult brain. J. Cell Biol. 121, 121-133. Drenckhahn D., & Bennett, V. (1987). Polarized distribution of Mr = 210,000 and 190,000 analogs of erythrocyte ankyrin along the plasma membrane of transporting epithelia, neurons and photoreceptors. Eur. J. Cell Biol. 43, 479-486. Drenckhahn, D., & Dermietzel, R. (1988). Organization of the actin filament cytoskeleton in the intestinal brush border: A quantitative and qualitative immunoelectron microscope study. J. Cell Biol. 107,1037-1048. Drenckhahn, D., & Merte, C. (1987). Restriction of the human kidney band 3-like anion exchanger to specialized subdomains of the basolateral plasma membrane of intercalated cells. Eur. J. Cell Biol. 45,107-115.

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Drenckhahn, D., Schluter, K., Allen, D.P., & Bennett, V. (1985). Colocalization of band 3 with ankyrin and spectrin at the basal membrane of intercalated cells in the rat kidney. Science 230,1281-1289. Drenckhahn, D., Oelmann, M., Schaaf, P., Wagner, M., & Wagner, S. (1987). Band 3 is the basolateral anion exchanger of the dark epithelial cells of the turtle urinary bladder. Am. J. Physiol. 252, C570-C574. Drenckhahn, D., Jons, T., Kollert-Jons, A., Koob, R., Kraemer, D., & Wagner, S. (1993). Cytoskeleton and epithelial polarity. Renal Physiol. Biochem. 16, 6-14. Durand-Schneider, A.-M., Bouanga, J.-C, Feldmann, G., & Maurice, M. (1991). Microtubule disruption interferes with the structural andfiinctionalintegrity of the apical pole in primary cultures of rat hepatocytes. Eur. J. Cell Biol. 56, 260-268. Eilers, U., Klumperman, J., & Hauri, H.-P. (1989). Nocodazole, a microtubule-active drug, interferes with apical protein delivery in cultured intestinal epithelial cells (Caco-2). J. Cell Biol. 108,13—22. Ellinger, A., Pavelka, M., & Gangl, A. (1983). Effect of colchicine on rat small intestinal absorptive cells. II. Distribution of label after incorporation of ( H) fucose into plasma membrane glycoproteins. J. Ultrastruct. Res. 85, 260-271. Path, K.R., & Burgess, D.R. (1993). Golgi-derived vesicles from developing epithelial cells bind actin filaments and possess myosin-I as a cytoplasmically oriented peripheral membrane protein. J. Cell Biol. 120,117-127. Path, K.R., Mamajiwalla, S.N., & Burgess D.R. (1993). The cytoskeleton in development of epithelial cell polarity. J. Cell Science, Suppl. 17, 63-73. Feldmann, G. (1989). The cytoskeleton of the hepatocyte. Structure and functions. J. Hepatol. 8, 380-386. Forte, J.G., & Wolosin, J.M. (1987). In: Physiology ofthe Gastrointestinal Tract, 2nd ed. (Johnson, L.R., Ed.). Raven Press, New York, pp. 853-863. Gilbert, T., Le Bivic, A., Quaroni, A., & Rodriguez-Boulan, E. (1991). Microtubular organization and its involvement in the biogenetic pathways of plasma membrane proteins in Caco-2 intestinal epithelial cells. J. Cell Biol. 113, 275-288. Gundersen, D., Orlowski, J., & Rodriguez-Boulan, E. (1991). Apical polarity of Na,K-ATPase in retinal pigment epithelium is linked to a reversal of ankyrin-fodrin submembrane cyroskeleton. J. Cell Biol. 112,86^^72. Harris, A.K., Pryer, N.K., & Paydarfar, D. (1990). Effects of electric fields on fibroblast contractility and cytoskeleton. J. Exp. Zool. 253, 163—176. Hubbard, A.L., Stieger, B., & Bartles, J.R. (1989). Biogenesis of endogenous plasma membrane proteins in epitheUal cells. Ann. Rev. Physiol. 51, 755-770. Huber, L. A., Pimplikar, S. W, Parton, R. G., Virta, H., Zerial, M., & Simons, K. (1993). Rab8, a small GTPase involved in vesicular traffic between the TGN and the basolateral plasma membrane. J. Cell Biol. 123,35-45. Jons, T., & Drenckhahn, D. (1992). Identification of the binding interface involved in linkage of cytoskeletal protein 4.1 to the erythrocyte anion exchanger. EMBO J. 11, 2863-2867. Jordan, Ch., Puschel, B., Koob, R., & Drenckhahn, D. (1995). Identification of a binding motif for ankyrin on the a-subunit of Na'*",K''-ATPase. J. Biol. Chem. 270, 29971-29975. Kemler, R. (1992). Classical cadherins. Semin Cell Biol. 3, 149-155. King, B.F. (1982). Cell surface specializations and intercellular junctions in human amniotic epithelium: An electron microscopic andfreeze-fracturestudy. Anat. Rec. 203, 73-82. Koob, R., Zimmermann, M., Schoner, W, & Drenckhahn, D. (1987). Colocalization and coprecipitation of ankyrin and Na ,K -ATPase in kidney epithelial cells. Eur. J. Cell Biol. 45, 230-237. Koob, R., Kraemer, D., Trippe, G., Aebi, U., & Drenckhahn, D. (1990). Association ofkidney and parotid Na ,K -ATPase microsomes, with actin and analogs of spectrin and ankyrin. Eur. J. Cell Biol. 53, 93-100. Kordeli, E., & Bennett, V. (1991). Distinct ankyrin isoform at neuron cell bodies and nodes of Ranvier resolved using erythrocyte ankyrin-deficient mice. J. Cell Biol. 114, 1243—1259.

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Kroschewski, R., Ruppert, C , & Bahler, M. (1994). Myr 1, a mammalian myosin-I molecule implicated in membrane traificking. Eur. J. Cell Biol. Suppl. 40, 91. Lambert, S., & Bennett, V. (1993). From anemia to cerebellar dysfunction. A review of the ankyrin gene family. Eur. J. Biochem. 211, 1-6. Le Bivic, A., Quaroni, A., Nichols, B., & Rodriguez-Boulan, E. (1990). Biogenetic pathways of plasma membrane proteins in Caco-2, a human intestinal epithelial cell line. J. Cell Biol. 111,1351—1361. Leto, T.L., Pratt, B.M., & Madri, J.A. (1986). Mechanisms of cytoskeletal regulation: Modulation of aortic endothelial cell protein band 4,1 by the extracellular matrix. J. Cell Physiol. 127,423-431. Luna, E.J., & Hitt, A.L. (1992). Cytoskeleton—^Plasma membrane interactions. Science, 258,955-963. Marchesi, V.T. (1993). In: Guidebook to the cytoskeletal and motor proteins (Kreis, T., & Vale. R., eds.), pp. 68-69. Sambrook & Tooze Publications, Oxford. Marrs, J.A., Napolitano, E.W., Murphy-Erdosh, C , Mays, R.W., Reicherdt, L.F., & Nelson, W.J. (1993). Distinguishing roles of the membrane-cytoskeleton and cadherin mediated cell-cell adhesion in generating different Na/K-ATPase distribution in polarized epithelia. J. Cell Biol. 123, 149-164. Matter, K., Brauchbar, M., Bucher, K., & Hauri, H.-P. (1990). Sorting of endogenous plasma membrane proteins occurs from two sites in cultured human intestinal epithelial cells (CacQ-2). Cell 60, 429-437. Mays, R.W., Beck, K.A., & Nelson, W.J. (1994). Organization and function of the cytoskeleton in polarized epithelial cells: A component of the protein sorting machinery. Curr. Opin. Cell Biol. 6, 16-24. McNeill, H., Ozawa, M., Kemler, R., & Nelson, W.J. (1990). Novel function of the cell adhesion molecule uvomorulin as an inducer of cell polarity. Cell 62, 309-316. Mercer, R.W (1993). Structure of the Na,K-ATPase. Int. Rev. Cytol. 137C, 139-168. Morrow, J.S., Cianci, CD., Ardito, T., Mann, A.S., & Kashgarian, M. (1989). Ankyrin links fodrin to the alpha subunit of Na , K -ATPase in Madin-Darby canine kidney cells and in intact renal tubule cells. J. Cell Biol. 108, 455-^65. Mostov, K., Apodaca, G., Aroeti, B., & Okamoto, C. (1992). Plasma membrane protein sorting in polarized epithelial cells. J. Cell Biol. 116, 577-583. Nelson, W.J., & Hammerton, R.W. (1989). A membrane-cytoskeletal complex containing Na ,K ATPase, ankyrin and fodrin in Madin-Darby canine kidney (MDCK) cells: Implications for the biogenesis of epithelial cell polarity. J. Cell Biol. 108, 893-902. Nelson, W.J., Shore, E.M., Wang, A.Z., & Hammerton, R.W. (1990). Identification of a membrane-cytoskeletal complex containing the cell adhesion molecule uvomorulin (E-cadherin), ankyrin, and fodrin in Madin-Darby canine kidney epithelial cells. J. Cell Biol. 110, 349-357. Nelson, W.J., & Veshnock, P.J. (1987). Ankyrin binding to (Na^ + K^) ATPase and implications for the organization of membrane domains in polarized cells. Nature 328, 533—536. Ozawa, M., & Kemler, R. (1992). Molecular organization of the uvomorulin-catenin complex. J. Cell Biol. 116,989-996. Pedersen, P.L., & Carafoli, E. (1987). Ion motive ATPases. I. Ubiquity, properties, and significance to cell function. Trends in Biochem. Sci. 12, 146-150. Pimplikar, S.W, & Simons, K. (1993). Regulation of apical transport in epithelial cells by a Gs class of heterotrimeric G protein. Nature 362, 456-458. Rindler, M.J., Ivanov, I.E., «fe Sabatini, D. (1987). Microtubule-acting drugs lead to the nonpolarized delivery of the influenza hemagglutinin to the cell surface of polarized Madin-Darby canine kidney cells. J. Cell Biol. 104,231-241. Rizzolo, L.J., & Joshi, H.C. (1993). Apical orientation of the microtubule organizing center and associated gamma-tubulin during the polarization of the retinal pigmented epithelium in vivo. Dev. Biol. 157, 147-156. Rodriguez-Boulan, E., & Salas, P.J.I. (1989). External and internal signals for epithelial cell surface polarization. Annu. Rev. Physiol. 51, 741-754.

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Rodriguez-Boulan, E., & Nelson, W.J. (1989). Moq)hogenesis of the polarized epithelial cell phenotype. Science 245, 718-725. Sawada, T., Itai, H., Fujikura, Y, Kuniki, H., Tamechika, M., & Fukumoto, T. (1992). Distribution of the surface antigen HAM-4 and cytoskeleton during reformation of bile-canalicular structures in rat primary cultured hepatocytes. Exp. Cell Res. 199, 50-55. Simons, K., & Wandinger-Ness, A. (1990). Polarized sorting in epithelia. Cell 62, 207-210. Srinivasan, Y, Lewallen, M., & Angelides, K.J. (1992). Mapping the binding site on ankyrin for the voltage-dependent sodium channel from brain. J. Biol. Chem. 267, 7483—7489. Steinmetz, RR. (1985). In: The Kidney. Physiology and Pathophysiology (Selding, D.W., & Giebish, G., Eds.). Raven Press, New York, pp. 1441-1458. Steuer, E.R., Wordeman, L., Schroer, T.A., & Sheetz, M.P. (1990). Localization of cytoplasmic dynein to mitotic spindles and kinetochores. Nature 345, 266-268. Stevenson, B.R., Anderson, J.M., & BuUivant, S. (1988). The epithelial tight junction: Structure, function and preliminary biochemical characterization. Mol. Cell. Biochem. 83, 129-145. Stolz, D.B., Bannish, G., & Jacobson, B.S. (1992). The role of the cytoskeleton and intercellular junctions in the transcellular membrane protein polarity of bovine aortic endothelial cells in vitro. J. Cell Sci. 103, 53-68. Stow, J.L., & de Almeida J.B. (1993). Distribution and role of heterotrimeric G proteins in the secretory pathway of polarized epithelial cells. J. Cell Science, Suppl. 17, 33—39. Toyoshima, I., Yu, H., Steuer, E.R., & Sheetz, M.P. (1992). Kinectin, a major kinfesin-binding protein on ER. J. Cell Biol. 118, 1121-1131. Tsukita, Sh., Tsukita, S., Nagafuchi, A., & Yonemura, S. (1992). Molecular linkage between cadherins and actin filaments in cell-cell adherens junctions. Curr. Opin. Cell Biol. 4, 834—839. Van der Sluijs, P., Bennett, M., Antony, C , Simons, K., & Kreis, T. E. (1990). Binding of exocytic vesicles from MDCK cells microtubules in vitro. J. Cell Sci. 95, 545—553. Wandinger-Ness, A., Bennett, M.K., Antony, C, & Simons, K. (1990). Distinct transport vesicles mediate the delivery of plasma membrane proteins to the apical and basolateral domains of MDCK cells. J. Cell Biol. I l l , 987-1000. Wessels, H.R, Hansen, G.H., Fuhrer, C , Look, A.T., Sjostrom, H., Noren, O., & Spiess, M. (1990). Aminopeptidase N is directly sorted to the apical domain in MDCK cells. J. Cell Biol. I l l , 2923-2930. Zachary, L, & Rozengurt, E. (1992). Focal adhesion kinase (pi25 ): A point of convergence in the action of neuropeptides, integrins, and oncogenes. Cell 71, 891—894. Zerial, M., & Stenmark, H. (1993). Rab GTPases in vesicular transport. Curr. Opin. Cell Biol. 5, 613-620.

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FOCAL ADHESIONS AND INTEGRIN-MEDIATED CELL SIGNALING

Susanne M. Bockholt and Keith Burridge

I. Introduction II. Focal Adhesions A. Homologous Structures/« Vivo B. Dynamics III. Integrins A. Structure B. ECM Interactions C. Targeting to Focal Adhesions IV. Cytoplasmic Face A. Focal Adhesion Proteins that Link Integrins to Actin Filaments B. Other Focal Adhesion Protein Interactions C. Isolation of Focal Adhesions V. Regulation of Focal Adhesions A. Protein expression B. Activation of PKC C. Tyrosine Phosphorylation D. Rho

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 167-206 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 167

168 169 169 171 172 172 172 173 176 176 180 182 183 183 184 186 188

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VI. Integrin-Mediated Signal Transduction A. Tyrosine Phosphorylation B. [Ca""^]! C. pHi D. Lipid Metabolism and PKC VII. Conclusion Acknowledgments References Note Added in Proof

189 189 189 191 192 193 193 193 205

I. INTRODUCTION Cells secrete, adhere to and model arrays of extracellular matrix (ECM). In turn, the ECM affects many aspects of cell behavior, including their migratory properties, their growth and differentiation. Much of what is known about how ECM affects cell behavior has been learned from studies of cells in culture. For example, it has been known for some time that addition of the ECM protein, fibronectin, to many transformed cells in culture will restore a normal phenotype. This includes a more spread morphology, increased adhesions and the presence of large bundles of actin filaments (stress fibers; Yamada et al., 1976; Ali et al, 1977; Hynes et al., 1977; Chen et al., 1986). These effects of fibronectin are due largely to its action of promoting or stabilizing focal adhesions (also known as focal contacts or adhesion plaques). Focal adhesions are discrete regions of the cell surface involved in tight adhesion to the underlying substratum (visualized in Figure 1). It should be noted

Figure 1, Visualization of focal adhesions and stress fibers. Rat embryo fibroblasts were plated overnight, in the absence of serum, on coverslips coated with the ECM protein, fibronectin. Focal adhesions were revealed with a monoclonal antibody against tensin, followed by a rhodamine-conjugated second antibody (A). The distribution of actin filaments was visualized in the same cell with fluorescein-phalloidin (B). Many bundles of actin filaments (stress fibers) can be seen to terminate in focal adhesions (arrowheads). Scale bar =10 jam.

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Cell Signaling

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that under most conditions of cell culture, the surface to which cells adhere is a layer of ECM protein, usually fibronectin or vitronectin derived from serum, adsorbed to the plastic culture dish. At their cytoplasmic face, focal adhesions serve to anchor stress fibers (Figure 1). Members of the integrin family of ECM receptors are concentrated at focal adhesions and form a transmembrane bridge between the ECM on the outside and the cytoskeleton on the inside. Recent evidence indicates that focal adhesions are also regions of signal transduction between the ECM and the cytoplasm. In this review, we will discuss ECM-cytoskeletal interactions, concentrating on integrins and focal adhesions. In addition, we will discuss the signaling events triggered by integrin-mediated cell adhesion.

II. FOCAL ADHESIONS A.

Homologous Structures in vivo

Most cells in vivo do not display focal adhesions, and this has led to the suggestion that focal adhesions are artifacts of tissue culture. The prominence of focal adhesions in many cultured cells probably reflects the abnormal conditions in which they are grown, in particular their growth on a rigid planar surface. Although focal adhesions may be rare in vivo, many cell-ECM junctions share structural and functional similarities with focal adhesions. In general, cell-ECM junctions are sites where there is a linkage across the membrane between the extracellular matrix on the outside and actin filaments on the inside. Tension generated in the microfilament system is transmitted across the membrane in these regions to the extracellular matrix. Focal adhesions closely resemble the myotendinous junctions of skeletal muscle, and the dense plaques of smooth muscle. In skeletal and cardiac muscle, many of the focal adhesion components are also localized in costameres (Table 1), sites of lateral attachment of the myofibrils to the sarcolemma at the level of the Z-discs (Pardo et al., 1983a, 1983b). In cultured myocytes, tension can be transmitted to the underlying substratum via costameres (Danowski et al., 1992). Many focal adhesion proteins are also concentrated in the postsynaptic region of neuromuscular junctions (Table 1), where the muscle plasma membrane interacts with a specialized extracellular matrix. It is not clear, however, whether tension is transmitted across the membrane at this site, leaving the function of focal adhesion proteins in this location uncertain. Focal adhesion components have also been detected at the basal surface of epithelial cells where they interact with the underlying basement membrane (Drenckhahn et al., 1988). During blood clot formation, platelets form adhesions to the ECM protein fibrinogen. Platelets contain most focal adhesion proteins in large amounts (for references see Burridge et al., 1988). Here they may function to link the contractile platelet cytoskeleton to the insoluble extracellular matrix generated during formation of a blood clot. Although focal adhesions are diminished in many transformed cells, cells transformed by Rous sarcoma virus (RS V) exhibit structures related to focal adhesions

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Table 1. In Vivo Localization and Other Characteristics of Focal Adhesion Proteins Many focal adhesion proteins are found at in vivo sites of either cell-cell or cell-substrate adhesion. In this table the presence of proteins in these various locations is indicated by (+), absence by (-), discrepancies by (+/-), and where not determined by (?). Tyrosine phosphorylation and other characteristics of these focal adhesion proteins are also indicated. The references from which these observations were taken are listed below: Talin^ Vinculirfi aactinin^ Paxillin^ Zyxin^

Tensin

FAK^

Fibroblasts (Focal Adhesions)

+

+

+/-

+

+

+

+

Epithelial Cells (Zonula Adherens)

-

+

+

-

+

+

?

Intercalated Disc (Fascia Adherens)

+/-

+

+

9

?

+

?

Z-line

-

+

+

?

?

*I-band

?

Costameres

+

+

-

?

9

+

?

Smooth Muscle (Dense Plaques)

+

+

+/-

+

+

+

?

Neuromuscular Junction

+

+

+

+

9

+

-

Myofibrils

Myotendinous Junction

+

+

+

+

9

+

+

Tyrosine phosphorylated in RSV-transformed cells Tyrosine phosphorylated in response to adhesion Kinase activity

+

+

-

+

9

+

+

-

-

-

+

9

+

+

-

-

-

-

-

-

+

+

+

-

-

SH2 Domain LIM Domain

9

+

Notes: ^ Burridge and Connell (1983), Geiger et al. (1985), Belkin et al. (1986), Tidball et al. (1986), DeClue and Martin (1987), Drenckhahn et al. (1988), Bockholt and Burridge (1993). ''Geiger (1979), Burridge and Feramisco (1980), Geiger et al. (1980),Geiger et al. (1981), Sefton and Hunter (1981), Bloch and Hall (1983), Pardo et al. (1983a, 1983b), Shear and Bloch (1985), Small (1985), Terracioetal. (1990). *= Stromer and Goll (1972), Lazarides and Burridge, (1975), Geiger et al. (1979), Wehland et al. (1979), Geiger et al. (1981), Sefton and Hunter (1981), Tokuyasu et al. (1981), Chen and Singer (1982), Bloch and Hall (1983), Small (1985), Tidball (1987), Samuelsson et al. (1993). ^ Glenney and Zokas (1989), Turner et al. (1990), Turner (1991), Turner et al. (1991), Burridge et al. (1992), Turner and Miller (1994). ^Beckerle (1986), Crawford and Beckerle (1991), Sadler et al. (1992). f Wilkins and Lin (1986), Glenney and Zokas (1989), Davis et al. (1991), Bockholt et al. (1992), Bockhoh and Burridge (1993). 8 Burridge et al. (1992), Guan and Shalloway (1992), Hanks et al. (1992), Komberg et al. (1992), Schaller et al. (1992), Baker et al. (1994). * Tensin was originally identified as a component of Z-lines (Wilkins and Lin, 1986). Additional studies with a mAb revealed localization to the I-band (Bockholt et al., 1992). The reason for this different localization may be due to distinct epitopes recognized by the different antibodies used in these studies.

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on the ventral surface. These have been referred to as "rosette adhesions" or "podosomes" (David-Pfeuty and Singer, 1980; Carley et al., 1981; Carley and Webb, 1983; Marchisio et al, 1984,1987; Tarone et al., 1985; Nermut et al., 1991) and have more of a dot-like appearance when visualized by immunofluorescence microscopy. These structures contain many of the focal adhesion components, but appear to be more dynamic, forming and disassembling rapidly (Stickel and Wang, 1987). These structures are probably the same as "invadopodia," invasive protrusions of the cell surface that have been studied in RSV-transformed cells grown on fibronectin-coated gelatin beads (Mueller et al., 1989, 1992; Mueller and Chen, 1991). Again, many focal adhesion components are prominent in invadopodia, but these protrusions are distinguished from focal adhesions by increased proteolytic activity at their extracellular surface (Chen and Chen, 1987). It is interestmg that rosette adhesions or podosomes have been detected in normal cells that are invasive, such as cells of the monocyte/macrophage lineage (Trotter, 1981; Lehto et al., 1982; Marchisio et al., 1987). The most elaborate rosettes or podosomes are found in osteoclasts and are associated with bone resorption (Marchisio et al., 1984, 1987; Zambonin-Zallone et al., 1989; Kanehisa et al., 1990). B.

Dynamics

Focal adhesions typically form within the leading lamella of a cell in culture. Assembly of focal adhesions has been most carefully analyzed by Izzard and his colleagues (Izzard and Lochner, 1976, 1980; DePasquale and Izzard, 1987, 1991) who have combined interference reflection microscopy (IRM) with immunofluorescence microscopy to study the formation of these structures. Before a focal adhesion is detectable by IRM (which reveals the closeness of the membrane to the substratum), an actin-rich structural precursor can be seen. This contains the focal adhesion protein talin, but not vinculin (Izzard, 1988). The assembly of a focal adhesion is undoubtedly a complex process which requires a number of coordinated events, including the activation of a tyrosine kinase (Burridge et al., 1992) and of protein kinase C (PKC) (Woods and Couchman, 1992), as will be discussed later. The assembly of focal adhesions has been treated mathematically (Ward and Hammer, 1993). In this model, initial integrin-mediated adhesion is followed by incorporation of a talin-containing cytoskeletal precursor, but the resulting structure is considered to have little mechanical rigidity. It is suggested that the strengthening of the adhesion derives from recruitment of additional receptors and development of crosslinks, involving some of the cytoskeletal proteins that are concentrated at the cytoplasmic face of focal adhesions. Many of the focal adhesions that form near the leading edge of the cell are transient. Only a few become stabilized, and associate with stress fibers. Although mature focal adhesions appear to be relatively stable areas of the plasma membrane and cell cortex, exchange of constituent proteins has been demonstrated (Kreis et al., 1982;Geigeretal., 1984; Meigs and Wang, 1986). Davies and coworkers (1993)

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have observed focal adhesions in real time with tandem scanning confocal image analysis. In stationary cells they found extensive remodeling of focal adhesions, although cell adhesion overall was not significantly altered. In addition, intact focal adhesions have been observed to change in shape and size, and move relative to each other, although how this occurs has not been established (Sanger et al., 1987; Stickel and Wang, 1988; Hock et al., 1989). Evidence for actin polymerization at focal adhesions supports the hypothesis that these areas are important in regulating stress fiber assembly (Kreis et al, 1982; Wang, 1984).

III. INTEGRINS A.

Structure

Prominent components of focal adhesions are integrins, cell surface receptors that mediate adhesion to ECM molecules. They consist of a and P heterodimers. Approximately 15 a chains and 8 (3 chains have been identified so far (Albelda and Buck, 1990;Hemler, 1990;Hynes, 1992). Each subunit has a single transmembrane domain, a large extracellular domain, and a short cytoplasmic domain. Many, but not all, of the a chains are proteolytically cleaved at a site just exterior to the transmembrane domain, and the two portions of the a subunit are linked together by a disulfide bond. The P subunits contain extensive intrachain disulfide bonds. The cytoplasmic domains of P integrin subunits are highly conserved. In general, the a subunit cytoplasmic domains are much more variable in sequence (Sastry and Horwitz, 1993). The structural features of integrin subunits have been reviewed in detail elsewhere (Hemler, 1990; Hynes, 1992). B. ECM Interactions

A single ECM protein may bind to more than one type of integrin. The sequences within ECM proteins involved in binding to particular integrins have received much attention. The first sequence to be identified was in the major cell binding ft-agment of fibronectin. Within this fragment of fibronectin, the sequence GRGDSP was found to be responsible for cell attachment and it was shown that the critical residues were RGD (Pierschbacher and Ruoslahti, 1984). Interestingly, RGD sequences have been found in many ECM proteins (Yamada and Kleinman, 1992) and are involved in the binding to several integrins (Ruoslahti and Pierschbacher, 1987; Hemler, 1990; Hynes, 1992). Some integrins recognize this sequence only within a single ECM protein. The integrin, a^^^ (the fibronectin receptor), for example, binds only to the RGD sequence in fibronectin. Other integrins may bind to the RGD sequence within many ECM proteins; for example, a^^^ (^^^ vitronectin receptor) binds to this sequence in vitronectin, fibronectin, von Willebrand's factor, thrombospondin, fibrinogen, and collagen (Hynes, 1992). Sequences flanking RGD must confer integrin specificity and restrict ligand interaction, as in the case

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of a5Pi. Other sequences within ECM proteins have also been identified as sites of interaction with integrins. For example, the sequence EILDV, found in an alternatively spliced region of fibronectin, has been shown to bind to the integrin a^^^ (Komoriya et al., 1991). Identification of the ECM ligand-binding sites on integrins has been less precise than the reciprocal studies of binding sequences in ECM proteins. Crosslinking studies and analysis of mutations have indicated that the N-terminal regions of both the a and P subunits contribute to ECM binding (for references see Hynes, 1992). Several integrins require "activation" before they will bind to their ECM ligands. This is illustrated well with the major integrin on platelets, OL^^^^^ that is responsible for binding fibrinogen. Since circulating platelets are continually bathed in fibrinogen, it would be a disaster if their interaction with fibrinogen was not regulated. The stimulus for clot formation involves a cascade of regulatory events occurring in platelets that culminate in the binding of ajj^Pj to fibrinogen. How this integrin becomes activated is not fully understood. Lipids may contribute to activation. Phosphatidic acid and lysophosphatidic acid enhance fibrinogen binding to a^i^f>^ in vitro (Smyth et al., 1992), but whether these are the physiological activators of ajj^P3 has not been determined. Conformational changes in the ajj^Pj following activation have been detected both immunologically and structurally (reviewed in Hynes, 1992). The cytoplasmic domains have a critical role in regulating the activation state as evidenced by transfection experiments in which the cytoplasmic domains are deleted. Deletion of the a^^^ cytoplasmic domain leads to a constitutively activated complex, implying that the ajj^^ cytoplasmic domain normally exerts an inhibitory effect on the heterodimer that must be relieved by activation (O'Toole et al., 1991). On the other hand, deletion of the P3 cytoplasmic domain eliminates cellular functions attributed to the aii^P3 integrin, such as spreading on fibrinogen (Ylanne et al., 1993). C. Targeting to Focal Adhesions

Most cells express several types of integrins on their cell surfaces, permitting them to interact with multiple ECM components. Cells adhering to a particular ECM protein, adsorbed to the substratum, will cluster the relevant integrin that interacts with this component into focal adhesions on the ventral surface. Other integrins will remain diffusely distributed over the surface of the cell. For example, the vitronectin receptor (a^P3) was found to concentrate in the focal adhesions of cells plated on vitronectin, but the fibronectin receptor (a5pj) was absent from these focal adhesions and was dispersed over the surface (Dejana et al., 1988; Singer et al., 1988; Fath et al., 1989). Conversely, when cells were plated on fibronectin, the fibronectin receptor concentrated in focal adhesions, whereas the vitronectin receptor was diffuse. However, when cells were plated on fibronectin, but in the presence of serum, initially a^^^ was found in the focal adhesions, but over the course of a few hours, a^P3 became the predominant integrin in focal adhesions (Fath et al..

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1989). In the presence of serum, fibronectin is cleared from the substratum (Avnur and Geiger, 1981a; Grinnell, 1986) and is replaced by vitronectin from the serum (Path and Burridge, unpublished results) accounting for the change in the integrins concentrated in focal adhesions. From these results, it was concluded that the type of integrin found in focal adhesions was dictated by the nature of the ECM protein adsorbed to the substratum. The targeting of integrins to focal adhesions, however, has turned out to be more complex. This was revealed by a series of transfection experiments in which integrin cytoplasmic domains were either deleted or expressed as parts of chimeric molecules. In some of these experiments, the cytoplasmic domain of the (3 subunit was shown to target to focal adhesions in the absence of an extracellular ligand binding to the protein (Geiger et al., 1992; LaFlamme et al., 1992; Briesewitz et al., 1993; Ylanneetal., 1993). These and other experiments examining the functions of integrin a and (3 cytoplasmic domains will be considered in the next two sections. /. fi Subunit To begin to understand the relationship between integrin structure and function, hybrid integrins were constructed. When the avian Pj integrin subunit was expressed in mouse NIH3T3 cells, it was shown to associate with endogenous murine a subunits (Solowska et al., 1989). The hybrid integrins with intact Pj cytoplasmic domains localized to focal adhesions, but integrins with truncated Pj cytoplasmic domains did not. This experiment provided some of the first evidence that an interaction, besides binding to ECM proteins, is required to retain integrins at focal adhesions (Solowska et al., 1989). Targeting of integrins to focal adhesions in a ligand-independent fashion was observed with transfected chimeric integrins. Chimeric integrins composed of the Pj cytoplasmic domain and the extracellular domain of the interleukin-2 receptor (LaFlamme et al., 1992) or N-cadherin (Geiger et al., 1992), both localized to focal adhesions. These chimeras demonstrate that the Pj cytoplasmic domain, without the extracellular domain or any a subunit, is sufficient to target the chimeric protein to focal adhesions. This result was surprising in light of the previous work, described above, indicating that integrins localize to focal adhesions based upon their interactions with ECM ligands present on the substratum (Dejana et al., 1988; Singer et al., 1988; Fath et al., 1989). To account for these results, it was hypothesized that, under normal conditions, binding of ECM ligands causes a conformational change in the integrin that is transmitted to the Pj subunit cytoplasmic domain, increasing its affinity for focal adhesion components. It was suggested that the cytoplasmic domains of these chimeric molecules might exhibit this "ligandbound" conformation constitutively. To test whether ligand binding to integrins increases their affinity for focal adhesions, cells adhering to laminin were treated with soluble ligands for either the fibronectin or vitronectin receptors (the 75 kDa cell-binding fragment of fibronectin and the GRGDS peptide, respectively). In a striking result, these soluble ligands drove their corresponding integrins into focal

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adhesions (LaFlamme et al, 1992). Thus, under these conditions, the targeting of integrins to focal adhesions does appear to be dictated by the interaction of integrins with ECM components. This targeting occurs, however, because of a conformational change in the integrin in response to the ECM binding, increasing the affinity of the cytoplasmic domain for cytoskeletal components in focal adhesions. To determine which portion of the p^ cytoplasmic domain is important for localization, site directed mutagenesis and various insertions and truncations were designed in the p^ cytoplasmic domain (Hayashi et al, 1990; Marcantonio et al, 1990; Reszka et al., 1992). The data obtained by these groups confirm that the pj cytoplasmic domain is important for targeting integrins to focal adhesions. However, no simple sequence alone accounts for the targeting and several regions appear to be involved (Hayashi et al., 1990; Marcantonio et al., 1990; Reszka et al., 1992). Some mutations and deletions may affect targeting because they alter the three-dimensional folding of the pj cytoplasmic tail. 2. a Subunit

What then is the role of the a subunit? The ability of these P^ cytoplasmic domain chimeras, which lack a chains, to target focal adhesions, without ligand binding, might be due to an active conformation of the P^ cytoplasmic domain. Alternatively, in the absence of an a subunit, the Pj cytoplasmic domain may not be masked by an a cytoplasmic sequence (LaFlamme et al., 1992), enabling the p cytoplasmic domain to associate with cytoskeletal proteins such as talin (Horwitz et al., 1986) or a-actinin (Otey et al., 1990), as will be discussed later. Recent studies suggest that the a cytoplasmic domain has an inhibitory role, preventing the localization of integrins to focal adhesions except when ligand is bound (Briesewitz et al., 1993; Ylanne et al., 1993). Mouse NIH3T3 cells containing the fibronectin receptor, a5P j , were transfected with either the complete wild type human aj subunit or a truncated form in which the C-terminal eight amino acids had been deleted. This deletion would remove most or all of the cytoplasmic domain, depending upon where the protein emerges from the membrane. (There is disagreement in the literature as to where the transmembrane domain ends and the cytoplasmic domain begins.) These transfected a^ subunits paired with endogenous P^ subunits to form the a^Pj integrin. This is a collagen receptor normally found only in focal adhesions of cells plated on collagen. Unlike the wild type a^, the truncated a^ subunit was colocalized with the endogenous agPj in focal adhesions when the transfected cells were plated on fibronectin (Briesewitz et al, 1993). Thus, the transfected wild type aj subunit is targeted to focal adhesions in a ligand-dependent fashion and the truncated a^ in a ligand-independeht manner. This experiment indicates that the short cytoplasmic domain of the wild type aj is able to limit recruitment of an integrin to focal adhesions. Essentially the same conclusions were drawn from experiments in which the wild type a^i^^^ (the fibrinogen receptor) and truncated versions of the ajj^, subunit were transfected into Chinese hamster ovary (CHO) or fibrosarcoma cells that were plated on fibronectin (Ylanne et al, 1993). Truncation

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of the a cytoplasmic domain resulted in the integrin targeting to focal adhesions in the absence of the appropriate ECM ligand, but the intact a cytoplasmic domain restricted targeting to only those receptors which bound appropriate ligands. These various integrin constructs, expressing altered cytoplasmic domains, permit other functions to be tested. For example, cells transfected with intact and various cytoplasmic domain mutants of OL^I^^^^ revealed that the P3 cytoplasmic domain was critical for cell spreading on a substrate of fibrinogen and for fibrin clot retraction (Ylanne et al., 1993). To investigate the roles of the a cytoplasmic domain, chimeric integrins were designed so that they consisted of the extracellular domain of the a2 integrin and the cytoplasmic domains of either a2, a^, or a^ (Chan et al., 1992). The cytoplasmic domains were swapped between the transmembrane domain and the highly conserved GFFKR region of these chimeras, and were transfected into rhabdomyosarcoma cells. The chimera containing the a^ cytoplasmic domain, but not the a2 or a5, was found to promote migration upon collagen and laminin. Chimeras expressing the a2 and a5 cytoplasmic domains were able to contract collagen gels, whereas the a^ did not (Chan et al., 1992). These results suggest distinct roles for the a cytoplasmic domains, either reflecting interactions of the a cytoplasmic domain with different cytoskeletal proteins or that the a cytoplasmic domains affect the cytoskeletal interactions of the p subunit. In a different set of experiments, various deletion mutants in the cytoplasmic domain of the human a5 integrin were expressed in a clonal line of CHO cells that had very low levels of endogenous a5. Motility, adhesion and spreading assays indicated that the region adjacent to the membrane of the a5 cytoplasmic domain is required for efficient spreading and motility on fibronectin (Bauer et al., 1993).

IV. CYTOPLASMIC FACE A.

Focal Adhesion Proteins that Link Integrins to Actin Filaments

The localization of chimeric integrins lacking extracellular domains to focal adhesions implies that cytoskeletal proteins have an important role in anchoring integrins at these sites. The first cytoskeletal protein found to bind integrins was talin. This interaction was detected using equilibrium gel filtration, and a K^ of »7 was determined (Horwitz et al., 1986). Additionally, the integrin binding site was mapped to the large 200 kDa fragment of talin. The interaction of talin with Pj and P3 integrins has been confirmed using purified integrins and synthetic peptides that correspond to full length or partial sequences of the cytoplasmic domains of these integrins (Simon and Burridge, unpublished results). Confirming the interactions of two proteins in cells is often difficult, particularly if their binding is of relatively low affinity. Under a variety of circumstances talin and integrins co-distribute, consistent with their interacting in cells. Not only are they found to co-distribute in focal adhesions, but also in numerous other sites of cell-ECM interaction, both in culture and in vivo. For example, unlike several other

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focal adhesion proteins, talin is not found at epithelial cell-cell junctions where integrins typically are also absent. However, talin can localize to these sites in cells transfected with chimeric integrins in which the pj cytoplasmic domain is driven to intercellular junctions by an N-cadherin extracellular domain (Geiger et al., 1992). In some situations, the co-distribution appears to be a regulated phenomenon and may depend on the activity of protein kinase C (PKC). For instance, when integrins were capped in chicken lymphocytes, talin did not co-distribute with the cap, except when the cells were stimulated with phorbol esters to activate PKC (Bum et al., 1988). Co-capping of talin with aggregated integrins was notably absent from phorbol ester-stimulated lymphocytes that lacked a functional PKC (Kupfer et al., 1990). Talin is a good substrate for PKC in vitro (Litchfield and Ball, 1986; Beckerle, 1990) and also becomes phosphorylated in cells treated with phorbol esters to activate PKC (Turner et al., 1989). It will be important to determine whether phosphorylation of talin increases the affinity of talin for integrin cytoplasmic sequences. In addition, talin co-distributed with integrins at the site of cell-cell interaction when cytotoxic T lymphocytes adhered to their target cells (Kupfer and Singer, 1989). This co-distribution may again involve activation of PKC. Under these conditions where talin was induced to co-cap with aggregated integrins, it was striking that other focal adhesion proteins, such as vinculin and a-actinin, were often not detected at these sites (Bum et al., 1988; Kupfer et al., 1990). Redistribution of talin from the cytoplasm to the plasma membrane, presumably to an association with integrins, occurs following thrombin activation of platelets. Interestingly, platelet activation also results in a stimulation of PKC (Beckerle et al., 1989). Protein interactions with the 47 kDa fragment of talin have not been identified. This fragment shares homology with the cytoskeletal, membrane-associated proteins, band 4.1 and ezrin (Rees et al., 1990). Both the large and small fragment of talin have been demonstrated to target to focal adhesions following microinjection, implying that this small fragment must bind some other focal adhesion component (Nuckolls et al., 1990). There are data which suggest that talin associates with membranes (Heise et al., 1991; Goldmann et al., 1992), and that at the membrane talin can nucleate actin polymerization (Kaufmann et al., 1992). As yet, it has not been determined whether the membrane association involves the 47 kDa fragment. Focal adhesion formation can be prevented by microinjecting antibodies against talin into cells, indicating that talin is necessary for the assembly of these stmctures (Nuckolls et al., 1992). The talin associations dismpted in this study were not determined. To identify other cytoskeletal proteins capable of interacting with integrins, lysates of chick embryo fibroblasts were passed over a Pj cytoplasmic domain peptide affinity column, a-actinin and vinculin bound to the column and were eluted in high salt (Otey et al., 1990). In subsequent solid phase binding assays, it was determined that vinculin did not interact specifically with the peptide, whereas a-actinin did with an affinity of 1.6 x 10"^M (Otey et al., 1990). Somewhat

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surprisingly, talin was not detected binding to this column, possibly reflecting the relatively low affinity of its interaction with the integrin cytoplasmic domain sequence. The integrin binding site on a-actinin was found to be contained within the 53 kDa fragment rod domain of a-actinin (Otey et al, 1990). Recent data indicates that a-actinin can also bind to the cytoplasmic domain of P2 in solid phase binding assays and in co-immunoprecipitations of the P2 integrin from neutrophils stimulated with chemotactic peptides (Pavalko and LaRoche, 1993). a-actinin is an actin filament-crosslinking protein (Podlubnaya et al., 1975) originally isolated from muscle (Maruyama and Ebashi, 1965). Subsequently, it was localized at the ends of stress fibers and along stress fibers in a sarcomeric-like periodic distribution (Lazarides and Burridge, 1975). Using IRM, a-actinin was confirmed to be in focal adhesions (Wehland et al., 1979). a-actinin had previously been found in Z-lines of skeletal muscle (Stromer and Goll, 1972) and at dense bodies and plaques in smooth muscle (Geiger et al., 1981). The presence of a-actinin in dense plaques has been disputed as a result of other evidence suggesting its absence from these regions, although the reasons for this discrepancy are not clear (Small, 1985). Its presence in focal adhesions has also been controversial because immunoelectron microscopy studies place this protein further away from the membrane (Chen and Singer, 1982; Samuelsson et al., 1993; Pavalko et al., 1995; Meijne and Ross, personal communication). However, the expression of chicken muscle-type a-actinin in non-muscle monkey kidney COS cells and mouse fibroblasts revealed that the transfected a-actinin distributed along stress fibers and colocalized with vinculin in focal adhesions (Jackson et al., 1989; Tokuue et al., 1991). When microinjected into fibroblasts, the 53 kDa fragment of a-actinin was found to incorporate rapidly into focal adhesions, but then caused disruption of stress fibers and focal adhesions (Pavalko and Burridge, 1991). Interestingly, cells expressing high levels of a-actinin or a truncated form of a-actinin, in which the EF-hand domain had been removed, were also found to have few microfilament bundles and a-actinin was diffusely distributed throughout the cells (Tokuue et al., 1991). It is possible that the microinjected and overexpressed fragments of a-actinin compete with endogenous a-actinin at the sites of integrin interaction which leads to displacement of the microfilaments anchored at focal adhesions and their subsequent disassembly. Several proteins may be involved in linking actin filaments to integrins (Figure 2). A direct bridge between integrins and actin filaments is possible because both a-actinin and the large fragment of talin can bind to actin (Muguruma et al., 1990, 1992; Goldmann and Isenberg, 1991, 1992). A more indirect linkage may be established by other protein-protein interactions. For example, the 200 kDa integrin-binding fragment of talin also contains a binding site for the globular domain of vinculin (Jones et al, 1989; Gilmore et al., 1992; Lee et al., 1992). In turn, vinculin can bind to a-actinin (Belkin and Koteliansky, 1987; Wachsstock et al., 1987) and thereby establish a chain of attachment to actin filaments. It is likely that several other proteins are also involved in linking actin filaments to the plasma

Figure 2. Focal adhesion model. This simplified model depicts a focal adhesion in a cell adhering to a glass substrate coated with fibronectin (FN) or vitronectin (VN). It i s at the focal adhesion that integrins bring the cell into closest contact with the underlying ECM (FN or VN). The a and p integrin subunits span the plasma membrane (PM), and the p cytoplasmic domain is shown binding to talin (TAL) and a-actinin (a-A), both of which, in turn, bind actin. Vinculin (V) binds to talin, a-actinin, tensin (TEN)and paxillin (Px).The focal adhesion kinase (FAK)becomes tyrosine phosphorylated and activated upon integrin-mediated adhesion. We suspect that other proteins, indicated by (?), regulate FAK activity and possibly its interactions with integrins. Both tensin and paxillin become tyrosine phosphorylated upon cell adhesion and are probable substrates of FAK. Tensin also is able to bind to the barbed end of actin filaments where it may nucleate actin polymerization (arrows indicate exchange of actin monomers). The focal adhesion protein, zyxin (Z), binds to a-actinin and to the chicken homologue of the human cysteine rich protein (cCRP) (C).Both zyxin and cCRP contain LIM domains, indicating a possible role in signal transduction pathways.

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membrane via integrins. One such candidate would be the actin-binding protein, tensin. As will be discussed later, the properties of tensin are such that if it also binds to integrins, it could allow actin filament growth while permitting simultaneously the transmission of tension to the membrane. The involvement of different proteins in anchoring actin filaments to the membrane may reflect differences in focal adhesions. The organization of focal adhesions may depend on their state of maturation, and whether they are present in motile or stationary cells. Adhesion to different ECM substrates involves distinct integrins and this may alter the composition of the resulting focal adhesions. B. Other Focal Adhesion Protein Interactions

Several focal adhesion proteins have been identified as a result of generating monoclonal antibodies a-gainst phosphotyrosine containing proteins. Chick embryo fibroblasts transformed by Rous sarcoma virus (RSV) exhibit a large number of tyrosine phosphorylated proteins. In two separate studies, monoclonal antibodies were generated against phosphotyrosine proteins isolated on anti-phosphotyrosine columns from the extracts of RSV-transformed cells (Glenney and Zokas, 1989; Kanner et al., 1990). Glenney and Zokas (1989) showed that both a 6^-76 kDa and a 215 kDa tyrosine phosphorylated protein recognized by distinct antibodies were concentrated in focal adhesions (Glenney and Zokas, 1989). The 68-76 kDa protein was later shown to bind to the rod domain of vinculin and has been named paxillin (Turner et al, 1990). During embryonic development, paxillin is one of the major tyrosine phosphorylated proteins detected in western blots of whole tissues (Turner, 1991). It was recently shown that the tyrosine phosphorylated form of paxillin binds to v-Crk, an oncoprotein containing both Src homology 2 (SH2) and Src homology 3 (SH3) domains (Birge et al., 1993). In addition, paxillin binds to the SH3 domain of pp60^"^^^ (Weng et al, 1993). The interaction of paxillin with these oncoproteins suggests that paxillin may have a role in signal transduction in focal adhesions. The cloning and sequencing of this protein is in progress and may reveal more about its function. The 215 kDa tyrosine phosphorylated protein identified by Glenney and Zokas (1989) was shown to be tensin (Bockholt et al., 1992). Kanner et al. (1990) generated antibodies in a similar manner against phosphotyrosine-containing proteins from RSV-transformed cells, some of which were also found to recognize tensin (Schaller et al., 1993b). Tensin was originally characterized by antibodies against low-molecular-weight contaminants from vinculin preparations, termed HAl (Wilkins et al., 1986). These antibodies were found to cross-react with two high molecular weight proteins of about 150 and 200 kDa. The 150 kDa protein and the HAl proteins were suggested to be breakdown products of the larger 200 kDa protein, now known as tensin (Wilkins et al., 1986, 1987). The 150 kDa form of tensin was found to bind actin and vinculin (Wilkins et al., 1987). Other results suggest that tensin can cap and bundle actin filaments through two domains located

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in the N-terminal region (Lo et al., 1994), as well as an actin capping region more centrally located (Loe et al., 1994; Chuang et al., 1995). Tensin also appears to be the same as insertin (Weigt et al., 1992), a name given to the actin-capping activity that contaminated vinculin preparations (Ruhnau et al., 1989). Unlike many other barbed end actin-capping proteins, tensin (insertin) allows actin monomer addition and polymerization at the capped end (Weigt et al., 1992). This property of tensin would enable continued attachment of the actin filament to the plasma membrane, while at the same time allowing actin filament elongation to occur. Although it is not clear how many actin binding sites exist in tensin, the evidence that it is able to nucleate actin polymerization, as well as cap and bundle actin filaments, certainly suggests that tensin must play an important role, both in anchoring stress fibers and nucleating their growth at focal adhesions. Sequence analysis of tensin revealed that it contains an SH2 domain (Davis et al., 1991). SH2 domains bind to phosphotyrosine residues of other proteins. As yet no other structural components of focal adhesions have been shown to contain this domain, which is typically found in proteins involved in signal transduction. The proteins that interact with the SH2 domain of tensin remain to be identified. Additionally, tensin contains numerous potential phosphorylation sites on serine, threonine, and tyrosine (Lo et al., 1992). Indeed, as will be discussed later, tensin becomes tyrosine phosphorylated during integrin-mediated cell adhesion (Bockholt and Burridge, 1993), further indicating that tensin is likely to be a critical regulatory component of focal adhesions. A 125 kDa focal adhesion protein was identified by monoclonal antibodies (mAbs) generated against phosphotyrosine-containing proteins, as described above (Kanner et al., 1990). Cloning of this protein revealed that it was a novel tyrosine kinase that localized to focal adhesions, and hence it was named pp 125^^^ or FAK (focal adhesion kinase; Schaller et al., 1992). The discovery of a tyrosine kinase specifically concentrated in focal adhesions has had major implications for signal transduction in these regions and this will be discussed in a later section. The mouse homologue of FAK was independently identified by a PCR cloning approach to identify new protein tyrosine kinases, based upon homology to the kinase domain (Hanks et al., 1992). Notably absent from FAK are domains commonly found in other tyrosine kinases, such as acylation sites for membrane attachment and SH2 and SH3 domains (Hanks et al., 1992; Schaller et al., 1992). The human form of FAK (hFAK) was found to have 95 percent amino acid homology with chicken FAK and to be highly expressed in brain, but low in heart and skeletal muscle (Andre and Becker, 1993). An N-terminally truncated form of hFAK was observed to be expressed in brain alone. A FAK-related nonkinase (FRNK) cDNA has been identified that encodes a 41/43 kDa protein (Schaller et al., 1993a). FRNK is essentially identical to the portion of FAK that is C-terminal to the kinase domain, and like FAK localizes to focal adhesions. Unlike FAK, FRNK is not a substrate for tyrosine phosphorylation, but is phosphorylated on serine and threonine residues (for a review on FAK and

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FRNK see Schaller and Parsons, 1993). The existence of this truncated form of FAK was unexpected, and its function has not been determined. During screening of preimmune sera, an antiserum was found that stained focal adhesions and recognized an 82 kDa protein (Beckerle, 1986). This protein, zyxin, was subsequently purified from chicken gizzard (Crawford and Beckerle, 1991) and found to bind to the actin-binding domain of a-actinin (Crawford et al., 1992). Sequence analysis has revealed that zyxin contains three LIM domains (Sadler et al., 1992). LIM domains typically coordinate metal ions and have been identified in several proteins involved in transcriptional regulation. Although the LIM domains of zyxin may signify a possible role for zyxin in signal transduction pathways, these domains may simply facilitate protein-protein interactions (Sadler et al., 1992; Michelsen et al., 1993). Interestingly, zyxin has been found to bind another LIM domain-containing protein, the chicken homologue of the human cysteine rich protein (cCRP). This has also been shown to concentrate in focal adhesions (Sadler et al, 1992; Michelsen et al., 1993). Several other proteins have been localized to focal adhesions, but their roles and interactions await further characterization. For example, antibodies against tenuin, a 400 kDa protein isolated from adherens junctions of rat liver, were shown to stain focal adhesions of fibroblasts (Tsukita et al., 1989). A46/50 kDa vasodilator-stimulated phosphoprotein (VASP), was found to be phosphorylated by cGMP-dependent and c AMP-dependent protein kinases in platelets and localized in fibroblasts to focal adhesions and along microfilaments (Reinhard et al., 1992). Although many focal adhesion proteins have not been fully characterized, a working model can be proposed based on the interactions that have been established in vitro (Figure 2). Some of the properties of these focal adhesion components and their localization in cell-ECM sites are summarized in Table 1. Several components have not been included and we are confident that more components will be identified in the near future. C. Isolation of Focal Adhesions

To understand and gather more information on focal adhesions, there have been attempts to isolate these structures (Avnur and Geiger, 1981b; Nicol and Nermut, 1987; Nermut et al., 1991,1993; Mueller et al., 1992; Gates et al., 1993; Samuelsson et al., 1993). Many of these have been adapted or modified from a procedure developed by Avnur and Geiger (1981b), in which a stream of hypotonic buffer containing ZnCl2 is used to shear the cell bodies from the ventral membranes and focal adhesions of cells adhering to a substrate. A different approach allows cells to adhere to ECM-coated magnetic beads. Following detergent extraction and homogenization, the magnetic beads with their associated adhesions are then separated from other cell components (Plopper and Ingber, 1993). Wet-cleavage is another method that has been used primarily to expose ventral membranes. In this technique the dorsal surface of cells is removed by allowing attachment to moist nitrocellulose

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(Brands and Feltkamp, 1988; Feltkamp et al., 1991; Meijne and Ross, personal communication). Immunofluorescence and immunoelectron microscopy have been applied to these various procedures in order to visualize and quantitate focal adhesion components. Although these approaches have not been utilized for identifying new focal adhesion components, it is a possible avenue for future studies.

V. REGULATION OF FOCAL ADHESIONS A. /.

Protein Expression

ECM

Changes in the levels of ECM proteins such as fibronectin can have profound effects on cell adhesion. This was first observed with cells transformed by tumor viruses, in which fibronectin was shown to be decreased (Gahmberg and Hakomori, 1973; Hynes, 1973). Addition of fibronectin to some of these transformed cells increased their adhesion and the cells assumed a more normal phenotype, but it did not restore normal growth control (Yamada et al., 1976; Ali et al., 1977; Hynes et al., 1977; Chen et al., 1986). This restoration of a normal morphology by the addition of fibronectin is not exhibited equally by all transformed cells (Yamada et al., 1976; Kopelovich et al, 1985; Kellie et al., 1986). Several factors may contribute to the loss of fibronectin from the surface of transformed cells. These include decreased synthesis, increased proteolysis and decreased binding of fibronectin to the cell surface (Olden and Yamada, 1977). 2. Alterations in Integrins

The decreased binding of fibronectin to some transformed cells suggested that there may be changes in the expression of integrins on the cell surface. This possibility was explored in cells transformed by several viruses and it was noted that, indeed, there was a reduction in the level of some a subunits (Plantefaber and Hynes, 1989). In all of the transformed cells, the a^ subunit was greatly reduced when compared with its level in the untransformed parental cells. In contrast, the a3 subunit showed little change in its level of expression. It was concluded that the diminished adhesion of these transformed cells to fibronectin was due to the altered levels of integrins and, in particular, to the reduction in a high affinity fibronectin receptor {a^^^; Plantefaber and Hynes, 1989). The effects of manipulating integrin levels have been investigated directly by transfecting the a^ subunit into CHO cells. These cells normally have very low levels of a5, but relatively high levels of a3. Clones with high levels of a5 expression demonstrated increased adhesion, along with decreased migration and decreased tumorigenicity (Giancotti and Ruoslahti, 1990). These experiments suggest that a3pi and a5Pj, although both receptors for fibronectin, may have very different functions. In general, the high affinity receptor,

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a5pj, appears to promote adhesion and inhibit migration, whereas a3pj, the low affinity receptor, appears to provide just sufficient adhesion to permit migration. Consistent with this idea, a^^^ is frequently concentrated in focal adhesions of cells on fibronectin, whereas a3P^ is often absent or not as prominent (Carter et al., 1990; Elices et al., 1991). Experiments by Akiyama and coworkers (1989) also support the different functions of a5 and a3 integrins. These investigators found that an anti-pj mAb inhibited adhesion to, and migration on, fibronectin, whereas an anti-a5 mAb actually promoted migration. This result could be explained if a5p} is responsible for the tight adhesion to fibronectin that restricts migration. When this integrin-ECM interaction was blocked with the anti-a5 mAb, adhesion was reduced, but not abolished, presumably due to the presence of low affinity receptors such as a3 which would allow migration to occur (Akiyama et al., 1989). 3,

Change in Vinculin Expression

Although much is known about how alterations in the ECM can affect the organization of the cytoskeleton, and there is increasing information about the effects of different integrins, rather little is known about the occurrence or consequences of altered levels of intracellular focal adhesion components. One component, vinculin, has received extensive study with regard to the regulation of mRNA and protein expression levels. Regulation of vinculin expression has been observed in several cell types as they differentiate (for references see Rodriguez Fernandez et al., 1993). Levels of vinculin mRNA and synthesis were elevated with increasing cell density, cell-cell contacts, and cell-substrate contacts (Ungar et al., 1986; Bendori et al., 1987). Furthermore, the tumorigenicity of transformed mouse cells that either lack vinculin or express it at subnormal levels could be suppressed by transfecting chicken vinculin into these cells (Rodriguez Fernandez et al., 1992a). This effect was suggested to be due to vinculin incorporation into cell-cell and cell-substratum adhesions (Rodriguez Fernandez et al., 1992a). Overexpressing vinculin in BALB/c 3T3 fibroblasts increased cell adhesion and reduced cell migration (Rodriguez Fernandez et al., 1992b). Conversely, when vinculin expression was reduced by antisense RNA, decreased adhesion and increased motility were observed. In addition, these cells had increased anchorage independent growth as evidenced by increased growth of colonies in soft agar (Rodriguez Fernandez et al., 1993). The mechanism for these changes remains to be defined. In the future, it will be interesting to determine whether modulating the levels of other focal adhesion components, such as talin, will have comparable effects as to those found with vinculin. B. Activation of PKC

In addition to the level of expression of various proteins found at focal adhesions, cell adhesion can also be regulated by protein kinase C (PKC). Seven different subspecies of PKC have been identified, many of which exhibit distinct expression

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and localization in both tissues and cells (Nishizuka, 1988). Isozymes of PKC have been shown to associate with cytoskeletal structures. The PKC a-isozyme (type 3) was immunolocalized to focal adhesions in rat embryo fibroblasts (Jaken et al., 1989). In other cell types such as baby hamster kidney cells, SW13 (human adrenal carcinoma cell line), CEM (human T cell line), and U937 (histiocytic myeloid line), the P isozyme was found to associate preferentially with intermediate filaments, although some association with stress fibers was also noted (Murti et al., 1992). Numerous studies have used tumor promoters such as 12-0 tetradecanoylphorbol-13-acetate (TPA or PMA) to activate PKC. This agent permits cells to adhere and spread on surfaces that are hydrophobic and normally nonadherent (Danowski and Harris, 1988). It also enhances the spreading of cells on fibronectin (Brown, 1988; Danilov and Juliano, 1989), but this is not due to phosphorylation of integrins by PKC (Danilov and Juliano, 1989). The enhanced adhesion of cells treated with TPA may be a result of reduced contractility, as judged by their diminished ability to wrinkle a flexible rubber substrate (Danowski and Harris, 1988). It has been known for some time that myosin activity can be inhibited by PKC, both as a result of direct phosphorylation of a myosin light chain and indirectly as a result of inhibition of the myosin light chain kinase which is a substrate for PKC (Nishikawa et al., 1985; Kawamoto et al., 1989). The spreading ability of cells appears to be opposed by the contractile activity of the cortex, with this contractile activity presumably being generated by myosin interacting with actin. Inhibiting myosin activity in fibroblasts by microinjection of anti-myosin antibodies (Honer et al, 1988) generates a phenotype that is very similar to that of cells treated with TPA (Danowski and Harris, 1988; Lyass et al, 1988; Bershadsky et al., 1990). In both cases, the cells demonstrate an increase in the spreading of their lamellae. Some of these lamellae appear to migrate away from the body of the cell, as if the contractile activity in the cortex of the cell that normally acts against the tendency of the leading edge to extend, has been inhibited. Some of the most dramatic morphological changes induced by PKC activation are found in epithelial cells. In response to TPA, these cells show a marked increase in membrane extension and ruffling which is accompanied by a loss of stress fibers and focal adhesions (Schliwa et al., 1984; Kellie et al., 1985; Meigs and Wang, 1986) PKC may act directly on focal adhesion components and a small increase in talin phosphorylation has been detected (Turner et al., 1989). However, the disassembly of focal adhesions could be indirect, for example, a consequence of inhibiting myosin activity, with this leading to stress fiber disassembly. It should be noted that microinjecting antibodies against myosin into cells resulted in the disassembly of stress fibers, but focal adhesions were not examined in this study (Honer etal., 1988). In contrast to the dramatic morphological changes induced by PKC activation in epithelial cells, mammalian fibroblasts exhibit little or no effects on their stress fibers and focal adhesions when these cells are treated with TPA (Jaken et al., 1989; Turner et al., 1989; Beckerle, 1990). Indeed, evidence has been presented that the

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converse situation exists, that PKC activation is required for focal adhesion formation (Woods and Couchman, 1992). Previously, these and other investigators had demonstrated that cells failed to develop focal adhesions when plated on the cell-binding fragment of fibronectin. If a heparin-binding domain was also present, focal adhesion formation was induced (Izzard et al., 1986; Woods et al., 1986). The requirement for the heparin-binding domain could be alleviated if the cells were treated with TPA (Woods and Couchman, 1992). Further evidence that PKC activity may normally be required for focal adhesion formation, was obtained using inhibitors of PKC. Under conditions where these reagents inhibited PKC, but not the cyclic AMP-dependent or cyclic GMP-dependent kinases, focal adhesions, as judged by the staining of talin and vinculin, were inhibited from forming in cells that had been allowed to spread on fibronectin (Woods and Couchman, 1992). Interestingly, the aggregation of Pj integrins into focal adhesions was not prevented by inhibiting PKC. It was as if PKC activity is required for linking integrins to the cytoskeletal proteins. It will be interesting to pursue this system further and to determine what proteins require phosphorylation by PKC in order for the complete assembly of a focal adhesion. Osteoclasts are another system in which TPA enhances the formation of adhesions, and in this case induces prominent bundles of microfilaments (Teti et al., 1992). As mentioned above, osteoclasts contain arrays of podosomes. These adhesions to the bone surface generate a sealed chamber which becomes the site of bone resorption (Kanehisa et al., 1990; Teti et al., 1991). Stimulating PKC with TPA results in enlarged podosomes and the appearance of prominent stress fibers (Teti et al., 1992). Presumably, the targets for PKC action in this cell type are not the same as those in epithelial cells which exhibit essentially the opposite response, i.e., a loss of focal adhesions and stress fibers. The variety of effects induced by agents that stimulate PKC, and which sometimes appear to be opposite in different cell types, may be accounted for by the existence of multiple PKC isoforms, with diverse cellular and subcellular distributions and different substrates. C. Tyrosine Phosphorylation

Elevated phosphotyrosine is found in the focal adhesions of normal and transformed cells (Marchisio et al, 1984; Maher et al., 1985). Tyrosine phosphorylation of specific proteins was originally studied in the context of transformation by RSV. Because transformation by this virus results in a loss of focal adhesions, work on RSV-transformed cells led to the idea that tyrosine phosphorylation of focal adhesion proteins might contribute to focal adhesion disassembly (Sefton and Hunter, 1981; Pasquale et al., 1986). Recent data, however, have indicated that elevated phosphotyrosine in normal cells is associated with focal adhesion assembly rather than disassembly (Burridge et al., 1992; Romer et al., 1992). The first focal adhesion proteins found to contain phosphotyrosine were vinculin, integrins, and talin, again in cells transformed by RSV (Sefton and Hunter, 1981; Hirst et al., 1986; Pasquale et al, 1986). Subsequent investigations, however, with nontrans-

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forming RSV mutants revealed that the phosphotyrosine content of vinculin and talin did not correlate with the transformed phenotype (for references see Burridge et al., 1988), and it is uncertain whether the tyrosine phosphorylation of these proteins has any physiological significance. An increase in tyrosine phosphorylation of a number of proteins occurs in response to clustering of integrins on cells (Komberg et al, 1991), and as cells spread on ECM proteins (Guan et al., 1991; Burridge et al., 1992; Hanks et al., 1992; Komberg et al., 1992). One of these has been identified as FAK (Burridge et al., 1992; Guan and Shalloway, 1992; Hanks et al, 1992; Komberg et al., 1992) and this tyrosine phosphorylation is associated with stimulation of FAK activity (Guan and Shalloway, 1992). Recent in vitro binding experiments indicate that there may be a direct association between FAK and the p^ integrin cytoplasmic domain (Schaller et al., 1995), but how integrin clustering activates FAK has not been resolved. Additional proteins may also participate in FAK activation as several signaling pathways appear to converge on FAK and lead to its elevated tyrosine phosphorylation and activation. For instance, FAK tyrosine phosphorylation is stimulated in src transformed cells (Kanner et al., 1990; Guan and Shalloway, 1992), in cells treated with the peptides bombesin, vasopressin, and endothelin (Zachary and Rozengurt, 1992), and in response to clustering of the high affinity IgE receptor in cells adhering to fibronectin, but not bovine semm albumin (BSA; HamawyetaL, 1993). Several other focal adhesion proteins also become tyrosine phosphorylated in response to integrin-mediated adhesion, including paxillin (Burridge et al., 1992), tensin (Bockholt and Burridge, 1993), and a 130 kDa protein (Fetch et al., 1995). In contrast, talin and vinculin, do not become tyrosine phosphorylated during cell adhesion to fibronectin, indicating that tyrosine phosphorylation of focal adhesion components is selective (Bockholt and Burridge, 1993). It seems likely that FAK is the kinase largely responsible for the tyrosine phosphorylation of focal adhesion proteins. In support of this, paxillin has been shown to be a substrate for FAK in vitro (Turner et al, 1993). The tyrosine phosphorylation of focal adhesion proteins can be inhibited by kinase inhibitors, such as herbimycin A and members of the tyrphostin family of compounds (Burridge et al., 1992; Romer et al, 1992). These inhibitors prevent the formation of focal adhesions and stress fibers as cells spread on fibronectin-coated surfaces, implicating tyrosine phosphorylation as a necessary step in the assembly of these cytoskeletal stmctures. The tyrosine phosphorylation of specific focal adhesion proteins may promote their interaction. The presence of an SH2 domain in tensin indicates that it may be a critical component involved in binding other tyrosine-phosphorylated proteins during focal adhesion formation. It would not be surprising if additional stmctural proteins were identified containing SH2 domains that contribute to the early events in focal adhesion assembly. Many cells adhere to and spread well on polylysine. Unlike adhesion to ECM components, adhesion to polylysine does not induce tyrosine phosphorylation (Guan et al., 1991; Burridge et al., 1992; Guan and Shalloway, 1992; Hanks et al., 1992; Komberg et al, 1992; Bockholt and Burridge, 1993). Cell spreading on

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polylysine is likely to be mediated through negatively charged surface components, such as proteoglycans. Proteoglycans have been shown to have a role in the spreading of cells on surfaces coated with arginine, lysine and other diamines, and this spreading could be reduced by treatment of the cells with chondroitinase ABC (Massia and Hubbell, 1992). Cells that spread on polylysine do not form focal adhesions, as long as synthesis and secretion of endogenous ECM proteins is prevented with agents such as cycloheximide. The situation in platelets is complex with respect to tyrosine phosphorylation and adhesion. In response to platelet agonists, an increase in tyrosine phosphorylation of several proteins was found to be dependent on the integrin a^^^^^ (F^rrell and Martin, 1988; 1989; Golden and Brugge, 1989; Golden et al., 1990). Although one integrin-dependent tyrosine phosphorylated protein has been identified as FAK (Lipfert et al., 1992), other proteins of 50-68 kDa and 140 kDa have been found to be tyrosine phosphorylated prior to FAK (Huang et al., 1993). The kinase that phosphorylates these substrates remains to be identified, but it is unlikely to be FAK since activation of FAK is associated with autophosphorylation and would be expected to precede the phosphorylation of these other proteins. D. Rho Some cells in culture (e.g., Swiss 3T3 fibroblasts) disassemble their stress fibers and focal adhesions when the cells are serum-starved. Addition of serum back to these quiescent cultures, leads to a rapid reappearance of both focal adhesions and stress fibers (Ridley and Hall, 1992). The factor responsible for this effect has been shown to be lysophosphatidic acid, and the response has been shown to involve the low molecular weight GTP-binding protein rho (Ridley and Hall, 1992). The reassembly of focal adhesions and stress fibers can be blocked by the C3 exotransferase of Clostridium botulinum, which ADP-ribosylates and inactivates rho. Moreover, direct microinjection of an activated form of rho into serum-starved, quiescent cells leads to the reappearance of stress fibers and focal adhesions (Paterson et al., 1990; Ridley and Hall, 1992). Somewhat surprisingly, rho is not concentrated along stress fibers, nor is it found in focal adhesions (Adamson et al., 1992). How rho regulates the assembly of these structures is currently unclear. An interesting possibility is that rho regulates the activity of FAK. Support for this idea comes from studies with the neuropeptide bombesin. This peptide stimulates the formation of focal adhesions and stress fibers in serum-starved cells in a rho-dependent manner (Ridley and Hall, 1992). In a different study, bombesin was also found to stimulate the tyrosine phosphorylation of FAK (Zachary and Rozengurt, 1992). Taken together, these two studies suggest that rho may promote focal adhesion assembly by stimulating FAK activity. However, the action of rho on focal adhesions could be indirect. There is evidence that activated rho enhances the calcium sensitivity of smooth muscle contraction (Hirata et al., 1992). A similar situation could exist in nonmuscle cells with rho increasing the contractility of actin and myosin. It has been argued previously that

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increased contractility leads to stress fiber assembly (Burridge, 1981). Because stress fibers are anchored at one or both ends by focal adhesions, the induction of stress fibers could result, in turn, in the assembly of focal adhesions. Elucidating the mechanism of action of rho will be important, as this protein may regulate the assembly and disassembly of stress fibers and focal adhesions under a variety of conditions.

VI. INTEGRIN-MEDIATED SIGNAL TRANSDUCTION Signals generated as cells adhere to ECM molecules include tyrosine phosphorylation, activation of PKC, changes in intracellular calcium and pH. It is becoming apparent that integrins and other focal adhesion proteins are involved in these signaling events which regulate focal adhesion formation, cell migration, and anchorage dependent growth. A. Tyrosine Phosphorylation

Earlier in this review (section VC) we discussed the role of tyrosine phosphorylation in focal adhesion assembly. However, integrin-mediated tyrosine phosphorylation may be involved in signal transduction beyond its role in cytoskeletal organization. Currently, little is known about the other consequences of integrinmediated tyrosine phosphorylation. The use of kinase inhibitors or the inactivation of FAK by molecular techniques or by microinjection of antibodies may reveal signaling cascades that are initiated by integrin-mediated tyrosine phosphorylation. Proteins, such as paxillin and pi30, that become tyrosine phosphorylated in focal adhesions have been regarded as cytoskeletal proteins because of their location in these regions. However, it is possible that they are not structural proteins, but function primarily in signal transduction. Another novel tyrosine phosphorylated protein of 190 kDa has been found to associate specifically with the vitronectin receptor, a^^^^ in Swiss 3T3 fibroblasts following stimulation with platelet derived growth factor (PDGF) when these cells were plated on vitronectin (Bartfeld et al., 1993). The localization and function of this protein has not been determined. B.

[Ca^^li

Changes in intracellular calcium ([Ca"^^]-) have also been found in integrin-mediated adhesion events and in response to integrin clustering. It was noted that antibody crosslinking of either subunit of CD lib/CD 18 (aj^P2)' i^ nonadherent neutrophils, generated a rise in cytosolic free calcium (Ng-Sikorski et al, 1991). Preventing neutrophil adhesion with oi^^2 antibodies, inhibited the transient elevations in [Ca"^^]j (Jaconi et al., 1991). Oscillations in [Ca"^^]j were observed in cells transfected with the platelet integrin, ajj^P3, when adhered to fibrinogen or stimulated with antisera against aii|,P3 (Pelletier et al., 1992). An increase in tyrosine phosphorylation of a 125 kDa protein (most probably FAK) accompanied the rise in [Ca'^^lj. Treating these transfected cells with BAPTA, an intracellular calcium chelator, prevented cell spreading and tyrosine phosphorylation of FAK, but not

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attachment to fibrinogen (Pelletier et al, 1992). These experiments are consistent with the rise in [Ca"^^]. being upstream of tyrosine phosphorylation of FAK. An alternative explanation is that elevation in [Ca'^^Jj may be required for cell spreading and that this is necessary for FAK tyrosine phosphorylation and activation. Elevation of intracellular calcium was also observed in human umbilical vein endothelial cells (HUVECS) plated on immobilized anti-integrin antibodies or on ECM proteins such as vitronectin and fibronectin (Schwartz, 1993). These integrinmediated increases in [Ca"*'^]j were determined to be regulated by voltage-independent calcium channels. Crosslinking of integrins in HUVECS with soluble antibodies failed to stimulate an increase in [CdL^\ (Schwartz, 1993), unlike the situation observed in neutrophils (Ng-Sikorski et al, 1991). This discrepancy may be explained by the rapid internalization of the immune complexes before calcium mobilization in endothelial cells, or that these are distinct phenomena for leukocytes and endothelial cells, reflecting different roles for the cells in vivo (Schwartz, 1993). Recent evidence indicates that distinct intracellular signals can be transmitted through different integrins, even within the same cell type. HUVECS plated on vitronectin or on anti-a^Pj mAbs exhibited an increase in [Ca'^^Jj, but this was not detected when the same cells were plated on collagen or on anti-a2Pj mAbs (Leavesley et al, 1993). In addition, there was a requirement for extracellular calcium for cell migration on vitronectin, but not on cpUagen (Leavesley et al., 1993). Neutrophil migration on vitronectin and fibronectin was also found to be dependent on calcium transients. When these calcium transients were blocked, the neutrophils were unable to detach from the substratum (Marks and Maxfield, 1990; Jaconi et al., 1991; Marks et al., 1991). Inhibition of calcineurin, a calcium-calmodulin dependent serine-threonine phosphatase, decreased motility on vitronectin, but not on fibronectin (Hendey et al., 1992). The authors suggest that neutrophil migration on vitronectin requires activation of calcineurin, but that on fibronectin these same cells require different calcium-dependent processes. Although the details of elevated [Ca"^^]^ signaling have not been completely resolved, it is possible to envisage some consequences due to increased [Ca"*"^]j. The interactions of actin-binding proteins (Stossel, 1993), the activity of phosphatases and kinases (Wilson et al., 1991; Hendey et al., 1992), and the activity of proteases, such as calpain, may all be affected by elevation of [Ca"^^]j. It is noteworthy that one isoform of calpain has been localized to focal adhesions (Beckerle et al., 1987). The focal adhesion protein talin, but neither vinculin nor a-actinin, was found to be a good substrate for calpain in vitro (Beckerle et al., 1987). The calcium concentrations required for calpain activation in vitro exceed that found in the cytoplasm of most cells, suggesting that there are additional ways to activate calpain. Consistent with this idea, calpain becomes activated in platelets in response to platelet agonists such as thrombin, and this depends on the integrin a^^^^ interacting with its adhesive ligands (Fox et al., 1993). Activation of calpain is diminished in thrombasthenic platelets that have reduced levels of aii^P3. Agents such as antibodies and RGDS peptides, which block the binding of OL^^^^ ^^ adhesive ligands, inhibit calpain activation but not the Ca"^^ influx induced by

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platelet agonists like thrombin. This indicates that although elevated [Ca'^^Jj may be essential for calpain activation, some additional signal is required that is generated by the interaction of ajj|jP3 with its adhesive ligand. C.

pHi

Most normal cells require anchorage to a substratum in order to grow and complete the cell cycle. A hallmark of malignancy is that the cells lose anchorage dependence for growth, that is they become capable of growing in soft agar or in suspension. The basis for anchorage dependent growth in normal cells and its frequent loss upon transformation has not been elucidated. Intracellular pH (pHj) has been implicated in growth control and suggested to be a factor in anchorage dependence (Schwartz et al., 1990). Normal fibroblasts growing on a substrate reveal an elevated pHj compared with the same cells kept in suspension. Significantly, many transformed cells that are anchorage independent do not exhibit this change in pHj, but rather have an elevated pHj both when adherent or kept in suspension (Schwartz et al., 1990). Artificially elevating the pH- of mammalian fibroblasts by transfecting in a yeast H"*"-ATPase pump stimulated the growth and tumorigenicity of these cells (Perona and Serrano, 1988). Schwartz and coworkers have fiirther explored the relationship of anchorage-dependent growth and pHj. Adhesion to a variety of ECM proteins, which support cell growth, such as fibrinogen, collagen types III, IV, V, laminin, and vitronectin were all found to increase pHj (Schwartz et al., 1991a). Substrates that support cell attachment, but not cell spreading and growth, such as basic fibroblastic growth factor, thrombin, and concanavalin A, did not elicit an elevation in pH-. These findings suggest that the elevation in pHj is an integrin-mediated response and, furthermore, that many integrins can trigger this response. The direct involvement of integrins was demonstrated by clustering a^^^ on the surface of cells with fibronectin- or antibodycoated beads (Schwartz et al., 1991b). Interestingly, in this same study, Schwartz and coworkers found using GRGDSP peptides that integrin occupancy was not sufficient to stimulate pH^. However, using a different cell type, Galkina et al. (1992) reported that ROD peptides could cause an increase in pHj within minutes of application to spread cells. Integrin-mediated [Ca'*"^]^ and pH- signaling appear to be independent as demonstrated with HUVECS. Although an increase in [Ca"^^]j was observed in HUVECS adhering to vitronectin but not collagen, pHj was elevated in cells on both substrates (Leavesley et al., 1993). The integrin-mediated elevation in pHj has been linked to the activation of the Na"*'/H"*" antiporter (Schwartz et al., 1991b). It will be very interesting to determine whether there is a physical association between this ion channel and integrins, and whether it is concentrated within focal adhesions. D. Lipid Metabolism and PKC Inositol phosphates, diacylglycerol (DAG) and arachidonic acid (AA) were all found to be significantly increased in rat glomerular epithelial cells (GEC) plated

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on collagen compared with the same cells plated on uncoated plastic (Cybulsky et al., 1993). Maximizing the integrin interactions by crosslinking integrins with anti-Pj integrin antibody, augmented the increase in DAG and AA in comparison with control antibodies or Fab fragments (Cybulsky et al, 1993). Similarly, HeLa cells adhering to gelatin showed elevations in inositol phosphates, DAG and AA (Chun and Jacobson, 1992, 1993). AA is produced by the action of phospholipase A2 on phospholipids and AA is itself a precursor of other signaling molecules, such as eicosanoids, that may have effects on adhesion (Chun and Jacobson, 1993). DAG and inositol phosphates are generated from phosphatidylinositol bisphosphate (PIP2) by the action of phospholipase C (PLC). DAG is a second messenger involved in the activation of PKC. Studies by Woods and Couchman (1992), mentioned above, indicate that activation of PKC is required for focal adhesion formation by cells adhering to fibronectin. Cells adhering to the cell-binding fragment of fibronectin were unable to develop focal adhesions unless either PKC was activated through the addition of TPA or a heparin-binding fragment of fibronectin was added. This result implies that the heparin-binding fragment of fibronectin stimulates PKC activation. The heparin-binding fragment of fibronectin interacts with cells via a transmembrane proteoglycan that has recently been identified (Woods and Couchman, 1993) and it will be interesting to determine how this activates PKC. Normal fibroblasts maintained in suspension fail to respond to various growth factors, including one critical for most fibroblasts, platelet derived growth factor (PDGF). McNamee et al. (1993) have investigated the basis for the decreased responsiveness to PDGF of fibroblasts in suspension. PLC is stimulated following PDGF binding to its receptor. Although PDGF was able to bind to its receptor activating PLC in suspended cells, the products of PLC action, inositol phosphates, were less abundant than in cells adhering to fibronectin. Further investigation revealed that this was because the substrate for PLC, PIP2, was decreased in cells in suspension compared with adherent cells. McNamee and coworkers (1993) documented that adhesion to fibronectin stimulated a significant increase in PIP2 synthesis. If the level of PIP2 is a rate limiting step in these signaling pathways, including the activation of PKC, then the regulation of PIP2 synthesis may be a critical step in anchorage-dependent growth control. PIP2 is an important precursor for signaling systems involving inositol phosphates and DAG, but PIP2 has also been shown to have an important role in regulating the interaction of many actin-binding proteins with actin. Many of the proteins that bind monomeric actin, such as profilin, or cap the ends of actin filaments, such as gelsolin, are dissociated from actin by PIP2 (Hartwig and Kwiatkowski, 1991). The increase in PIP2 levels that arise from integrin-mediated adhesion would be expected to stimulate increased actin filament polymerization, both from the release of actin monomers from the monomer-binding proteins and the uncapping of actin filaments, exposing sites for the addition of these monomers. The localized increase in PIP2 at sites of adhesion to ECM may thus contribute to the nucleation of actin polymerization.

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VII. CONCLUSION During the last few years there has been a convergence of several areas of cell biology that has stimulated interest in focal adhesions. Focal adhesions are not only sites where the actin cytoskeleton links to the plasma membrane, but they are also important regions of signal transduction. Several signaling pathways appear to be triggered by integrin-mediated adhesion and to originate in focal adhesions. At the moment little is known about how these signaling pathways are stimulated by integrin binding to ECM ligands. Undoubtedly, this will become clearer in the near future, but we anticipate that signaling in focal adhesions will also become more complex as additional pathways are identified. A key area for future study is how different integrins mediate distinct responses to ECM substrata. In some cases, integrins promote adhesion, whereas in other situations integrins promote migration. The integrin a subunits, and particularly their cytoplasmic domains, appear to be responsible for these differences. It will be important to identify the proteins with which these cytoplasmic domains interact and how these cytoplasmic domains regulate the interactions and functions of common p subunits.

ACKNOWLEDGMENTS We thank our colleagues for valuable discussions. The authors have been supported by NIH grants GM29860 and HL45100.

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Schaller, M.D., Borgman, C.A., Cobb, B.S., Vines, R.R., Reynolds, A.B., & Parsons, J.T. (1992). ppl25FAK a structurally distinctive protein-tyrosine kinase associated with focal adhesions. Proc. Natl. Acad. Sci. USA 89, 5192-5196. Schaller, M.D., Borgman, C.A., & Parsons, J.T. (1993a). Autonomous expression of a noncatalytic domain of the focal adhesion-associated protein tyrosine kinase ppl25 . Mol. Cell. Biol. 13, 785-791. Schaller, M.D., Bouton, A.H., Flynn, D.C., & Parsons, J.T. (1993b). Identification and characterization of novel substrates for protein tyrosine kinases. Prog. Nucleic Acid Res. Mol. Biol. 44,205-227. Schaller, M.D., & Parsons, J.T. (1993). Focal adhesion kinase: an integrin-linked protein tyrosine kinase. Trends Cell Biol. 3, 258-262. Schaller, M.D., Otey, C.A., Hildebrand, J.D., & Parsons, J.T. (1995). Focal adhesion kinase and paxillin bind to peptides mimicking p-integrin cytoplasmic domains. J. Cell Biol. 130, 1181—1187. Schliwa, M., Nakamura, T, Porter, K.R., & Euteneuer, U. (1984). A tumor promoter induces rapid and coordinated reorganization of actin and vinculin in cultured cells. J. Cell Biol. 99, 1045-1059. Schwartz, M. A. (1993). Spreading of human endothelial cells on fibronectin or vitronectin triggers elevation of intracellular free calcium. J. Cell Biol. 120, 1003-1010. Schwartz, M.A., Ingber, D.E., Lawrence, M., Springer, T.A., & Lechene, C. (1991a). Multiple integrins share the ability to induce elevation of intracellular pH. Exp. Cell Res. 195, 533-535. Schwartz, M.A., Lechene, C, & Ingber, D.E. (1991b). Insoluble fibronectin activates the Na/H antiporter by clustering and immobilizing integrin alpha 5 beta 1, independent of cell shape. Proc. Natl. Acad. Sci. USA 88, 7849-7853. Schwartz, M.A., Rupp, E.E., Frangioni, J.V., & Lechene, C.P. (1990). Cytoplasmic pH and anchorageindependent growth induced by v-Ki-ras, v-src or polyoma middle T. Oncogene 5, 55—58. Sefton, B.M., & Hunter, T. (1981). Vinculin: A cytoskeletal target of the transforming protein of Rous sarcoma virus. Cell 24, 165-174. Shear, C.R., & Bloch, R.J., (1985). VincuHn in subsarcolemmal densities in chicken skeletal muscle: Localization and relationship to intracellular and extracellular structures. J. Cell Biol. 101, 240-256. Singer, I.I., Scott, S., Kawka, D.W., Kazazis, D.M., Gailit, J., & Ruoslahti, E. (1988). Cell surface distribution of fibronectin and vitronectin receptors depends on substrate composition and extracellular matrix accumulation. J. Cell Biol. 106, 2171-2182. Small, J.V. (1985). Geometry of actin-membrane attachments in the smooth muscle cell: The localisations of vinculin and a-actinin. EMBO J. 4,45—49. Smyth, S.S., Hillery, C.A., & Parise, L.V. (1992). Fibrinogen binding to purified platelet glycoprotein Ilb-IIIa (integrin anbPs) is modulated by lipids. J. Biol. Chem. 267, 15568-15577. Solowska, J., Guan, J.L., Marcantonio, E.E., Trevithick, J.E., Buck, C.A., & Hynes, R.O. (1989). Expression of normal and mutant avian integrin subunits in rodent cells [published erratum appears in J. Cell Biol. 1989; 109: 1187]. J. Cell Biol. 109, 853-861. Stickel, S.K., & Wang, Y.-L. (1987). Alpha-actinin-containing aggregates in transformed cells are highly dynamic structures. J. Cell Biol. 104,1521-1526. Stickel, S.K., & Wang, Y.-L. (1988). Synthetic peptide GRGDS induces dissociation of alpha-actinin and vinculinfromthe sites of focal contacts. J. Cell Biol. 107,1231-1239. Stossel, T.R (1993). On the crawling of animal cells. Science 260, 1086-1094. Stromer, M., & Goll, D. (1972). Studies on purified actin: II. Electron microscopic studies on the competitive binding of a-actinin and tropomyosin to Z-line extracted myofibrils. J. Mol. Biol. 67, 489-494. Tarone, G., Cirillo, D., Giancotti, F.G., Comoglio, RM., & Marchisio, RC. (1985). Rous sarcoma virus-transformed fibroblasts adhere primarily at discrete protrusions of the ventral membrane call podosomes. Exp. Cell Res. 159,179-186.

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Terracio, L., Simpson, D.G., Hilenski, L., Carver, W., Decker, R. S., Vinson, N., & Borg, T.K. (1990). Distribution of vinculin in the Z-disk of striated muscle: Analysis by laser scanning confocal microscopy. J. Cell Physiol. 145, 78-87. Teti, A., Colucci, S., Grano, M., Argentino, L., & Zambonin, Z.A. (1992). Protein kinase C affects microfilaments, bone resorption, and [Ca2+]o sensing in cultured osteoclasts. Am. J. Physiol. 263, C130-<:i39. Teti, A., Marchisio, P.C, & Zallone, A.Z. (1991). Clear zone in osteoclast function: Role of podosomes in regulation of bone-resorbing activity. Am. J. Physiol. 261, C1-C7. Tidball, J.G. (1987). a-actinin is absent from the terminal segments of myofibrils and from subsarcolemmal densities in frog skeletal muscle. Exp. Cell Res. 170,469-482. Tidball, J.G., O'Halloran, T, & Burridge, K. (1986). Talin at myotendinous junctions. J. Cell Biol. 103, 1465-1472. Tokuue, Y., Goto, S., Imamura, M., Obinata, T, Masaki, T., & Endo, T. (1991). Transfection of chicken skeletal muscle a-actinin cDNA into nonmuscle and myogenic cells: Dimerization is not essential for a-actinin to bind to microfilaments. Exp. Cell Res. 197, 158-167. Tokuyasu, K.T., Dutton, A.H., Geiger, B., & Singer, S. J. (1981). Ultrastructure of chicken cardiac muscle as studied by double immunolabeling in electron microscopy. Proc. Natl. Acad. Sci. USA 78, 7619-7623. Trotter, J.A. (1981). The organization of actin in spreading macrophages. Exp. Cell Res. 132, 235-248. Tsukita, S., Itoh, M., & Tsukita, S. (1989). A new 400-kD protein from isolated adherens junctions: Its localization at the undercoat of adherens junctions and at microfilament bundles such as stress fibers and circumferential bundles. J. Cell Biol. 2905-2915. Turner, C.E. (1991). Paxillin is a major phosphotyrosine-containing protein during embryonic development. J. Cell Biol. 115, 201-207. Turner, C.E., Glenney, J.R.J., & Burridge, K. (1990). Paxillin: Anew vinculin-binding protein present in focal adhesions. J. Cell Biol. I l l , 1059-1068. Turner, C.E., Kramarcy, N., Sealock, R., & Burridge, K. (1991). Localization of paxillin, a focal adhesion protein, to smooth muscle dense plaques, and the myotendinous and neuromuscular junctions of skeletal muscle. Exp. Cell Res. 192, 651-655. Turner, C.E., Pavalko, P.M., & Burridge, K. (1989). The role of phosphorylation and limited proteolytic cleavage of talin and vinculin in the disruption of focal adhesion integrity. J. Biol. Chem. 264, 11938-11944. Turner, C.E., Schaller, M.D., & Parsons, J.T. (1993). Tyrosine phosphorylation of the focal adhesion kinase ppl25 during development: Relation to paxillin. J. Cell Sci. 105, 637—645. Ungar, F., Geiger, B., & Ben-Ze'ev, A. (1986). Cell contact- and shape-dependent regulation of vinculin synthesis in cultured fibroblasts. Nature 319, 787-791. Wachsstock, D.H., Wilkins, J.A., & Lin, S. (1987). Specific interaction of vinculin with a-actinin. Biochem. Biophys. Res. Commun. 146, 554—560. Wang, Y.L. (1984). Reorganization of actin filament bundles in living fibroblasts. J. Cell Biol. 99, 1478-1485. Ward, M.D., & Hammer, D.A. (1993). A theoretical analysis for the effect of focal contact formation on cell-substrate attachment strength. Biophys. J. 64, 936-959. Wehland, J., Osbom, M., & Weber, K. (1979). Cell to substratum contact in living cells: A direct correlation between interference reflection and indirect immunofluorescence microscopy using antibodies against actin and a-actinin. J. Cell Sci. 37, 257-273. Weigt, C , Gaertner, A., Wegner, A., Korte, H., & Meyer, H.E. (1992). Occurrence of an actin-inserting domain in tensin. J. Mol. Biol. 227, 593-595. Weng, Z., Taylor, J. A., Turner, C.E., Brugge, J.S., & Seidel-Dugan, C. (1993). Detection of src homology 3-binding proteins, including paxillin, in normal and v-src-transformed Balb/c 3T3 cells. J. Biol. Chem. 268, 14956-14963.

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Wilkins, J.A., & Lin, S. (1986). A re-examination of the interaction of vinculin with actin. J. Cell Biol. 102, 1085-1092. Wilkins, J.A., Risinger, M.A., Coffey, E., & Lin, S. (1987). Purification of a vinculin binding protein from smooth muscle. J. Cell Biol. 105, 103a. Wilkins, J.A., Risinger, M.A., & Lin, S. (1986). Studies on proteins that co-purify with smooth muscle vinculin: Identification of immunologically related species in focal adhesions of nonmuscle and Z-lines of muscle cells. J. Cell Biol. 103, 1483-1494. Wilson, A.K., Gorgas, G., Claypool, W.D., & de LaneroUe, R (1991). An increase or decrease in myosin II phosphorylation inhibits macrophage motility, J. Cell Biol. 114, 277-283. Woods, A., & Couchman, J.R. (1992). Protein kinase C involvement in focal adhesion formation. J. Cell Sci. 277-290. Woods, A., & Couchman, J.R. (1993). Syndecan 4-A widespread transmembrane heparan sulfate proteoglycan present in focal adhesions. Mol. Biol. Cell 4, 406a. Woods, A., Couchman, J. R., Johansson, S., & Hook, M. (1986). Adhesion and cytoskeletal organization of fibroblasts in response to fibronectin fragments. EMBO J. 5, 665-670. Yamada, K.M., Yamada, S.S., & Pastan, I. (1976). Cell surface protein partially restores morphology, adhesiveness, and contact inhibition of movement to transformed fibroblasts. Proc. Natl. Acad. Sci. USA 73, 1217-1221. Yamada, Y, & Kleinman, H. (1992). Functional domains of cell adhesion molecules. Curr. Opin. in Cell Biol. 4, 819-823. Ylanne, J., Chen, Y, O'Toole, T.E., Loftus, J.C, Takada, Y, & Ginsberg, M.H. (1993). Distinct functions of integrin alpha and beta subunit cytoplasmic domains in cell spreading and formation of focal adhesions. J. Cell Biol. 122, 223-233. Zachary, I., & Rozengurt, E. (1992). Focal adhesion kinase (pl25FAK): A point of convergence in the action of neuropeptides, integrins, and oncogenes. Cell 71, 891-894. Zambonin-Zallone, A., Teti, A., Grano, M., Rubinacci, A., Abbadini, M., Gaboli, M., & Marchisio, PC. (1989). Immunocytochemical distribution of extracellular matrix receptors in human osteoclasts: A beta 3 integrin is colocalized with vinculin and talin in the podosomes of osteoclastoma giant cells. Exp. Cell Res. 182, 645-652.

NOTE ADDED IN PROOF Since writing this review the growth in the field of focal adhesions and signal transduction has been substantial. It is impossible to adequately update all of the sections of this review, but nonetheless, we would like to highlight recent research, some of which is discussed in greater detail in other reviews (Clarke and Brugge, 1995; Jockusch et al, 1995; Schwartz et al., 1995). We will first update a few components of focal adhesions and then briefly discuss signaling pathways. Further investigation of talin in thrombin-activated platelets has demonstrated that the relocalization of talin from the cytoplasm to the plasma membrane is dependent upon phosphorylation by serine/threonine kinases other than PKC, and does not require calcium dependent cleavage of talin (BertagnoUi et al., 1993). Given that talin can bind to the cytoplasmic domain of integrins, it is surprising that the redistribution of talin occurs even in the absence of the major platelet integrin, GPIIb-IIIa, suggesting that talin's redistribution is either dependent upon another cytoskeletal protein or a less abundant integrin. Alternatively, phosphorylation of

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talin may enable it to associate directly with phospholipids in the plasma membrane (Bertagnolli et al., 1993). The ability of vinculin to bind to F-actin has been a controversial subject over the past several years (see Jockusch et al, 1995). However, it has now been shown that vinculin contains a cryptic F-actin binding site within the tail domain that is masked by its head domain (Menkel et al., 1994; Johnson and Craig, 1995). Furthermore, the phospholipid PIP2 has been found to dissociate vinculin's headtail interaction, exposing both its actin-binding and talin-binding sites (Gilmore and Burridge, unpublished observations). A relatively new family of cytoskeletal proteins that provides a number of candidates in signal transduction is that which consists of protein which contain LIM domains, a motif comprised of two zinc fingers. The first of zyxin's three LIM domains was demonstrated to be necessary and sufficient to facilitate a protein-protein interacfion with cCRP (CRPl; Schmeichel and Beckerle, 1994). Thus, like SH2 and SH3 domains, LIM domains can funcfion as modular protein-binding interfaces. In addition to binding to CRPl, one of three highly conserved members of the CRP family (Weiskirchen et al., 1995), zyxin has also been shown to bind to VASP (Reinhard et al., 1995) and the protooncogene product Vav (Hobert et al, 1996). Based upon the fact that CRP family members are very closely related, zyxin may also bind to the muscle LIM protein (MLP or CRP3) which has been shown to potentiate myogenic differentiation (Arber et al., 1994). The LIM domain family of focal adhesion proteins also now includes paxillin. Chicken paxillin cDNA was shown to encode a proline-rich region at the N-terminus and four LIM domains at the C-terminus, similar to zyxin (Turner and Miller, 1994). Collectively, these findings indicate that LIM domain proteins are important components of signal transduction pathways. Integrin-dependent signaling pathways have been expanded to include the activation of Ras (Kapron-Bras et al, 1993) and MAP kinase (Chen et al., 1994; Schlaepfer et al., 1994) which together may lead to activation of transcription factors (see Figure 2 in Clarke and Brugge, 1995). In addition, evidence now suggests that protein tyrosine kinases such as FAK lie downstream of Rho (Barry and Critchley, 1994; Chrzanowska-Wodnicka and Burridge, 1994; Ridley and Hall, 1994; and Figure 3 in Schwartz et al., 1995). It has become apparent that FAK is centrally located in a web of signaling pathways (see Figure 6 in Schwartz et al., 1995). Various FAK-activating pathways, including DAG and PKC, Rho and PIP 5-kinase, integrins, and oncogenes have several downstream effectors that regulate the formafion of focal adhesions and stress fibers, and cell growth (Schwartz et al., 1995). Several studies have shown that integrin-mediated adhesion is also important in the suppression of programmed cell death known as apoptosis (see Schwartz etal., 1995).

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Arber, S., Haider, G., & Caroni, P. (1994). Muscle LIM protein, a novel essential regulator of myogenesis, promotes myogenic differentiation. Cell 79,221—231. Barry, S.T., & Critchley, D.R. (1994). The RhoA-dependent assembly of focal adhesions in Swiss 3T3 cells is associated with increased tyrosine phosphorylation and the recruitment of both ppl25 and protein kinase C-delta to focal adhesions. J. Cell Sci. 107,2033-2045. Bertagnolli, M.E., Locke, S.J., Hensler, M.E., Bray, P.F., & Beckerle, M.C. (1993). Talin distribution and phosphorylation in thrombin-activated platelets. J. Cell Sci. 106,1189-1199. Chen, Q., Kinch, M.S., Lin, T.H., Burridge, K., & Juliano, R.L. (1994). Integrin-mediated cell adhesion activates mitogen-activated protein kinases. J. Biol. Chem. 269,26602—26605. Chrzanowska-Wodnicka, M., & Burridge, K. (1994). Tyrosine phosphorylation is involved in reorganization of the actin cytoskeleton in response to serum or LPA stimulation, J. Cell Sci. 107, 3643-3654. Clarke, E. A., & Brugge, J.S. (1995). Integrins and signal transduction pathways: The road taken. Science 268, 233-239. Robert, O., Schilling, J.W., Beckerle, M.C, Ullrich, A., & Jallal, B. (1996). SH3 domain-dependent interaction of Vav with the LIM-domain protein zyxin. Oncogene. In press. Jockusch, B.M., Bubeck, P., Giehl, K., Kroemker, M., Moschner, J., Rothkegel, M., Riidiger, M., Schliiter, K., Stanke, G., & Winkler, J. (1995). The molecular architecture of focal adhesions. Annu. Rev. Cell Dev. Biol. 11, 379-416. Johnson, R.P., & Craig, S.W. (1995). F-actin binding site masked by the intramolecular association of vinculin head and tail domains. Nature 373, 261-264. Kapron-Bras, C, Fitz-Gibbon, L., Jeevaratnam, J., Wilkins, J., & Dedhar, S. (1993). Stimulation of tyrosine phosphorylation and accumulation of GTP-bound p21ras upon antibody-mediated a2pi integrin activation in T-lymphoblastic cells. J. Biol. Chem. 268,20701-20704. Menkel, A.R., Kroemker, M., Bubeck, R, Ronsiek, M., Nikolai, G., & Jockusch, B.M. (1994). Characterization of an F-actin-binding domain in the cytoskeletal protein vinculin. J. Cell Biol. 126, 1231-1240. Reinhard, M., Jouvenal, K., Tripier, D., & Walter, U. (1995). Identification, purification, and characterization of a zyxin-related protein that binds the focal adhesion and microfilament protein VASP (vasodialator-stimulated phosphoprotein). Proc. Natl. Acad. Sci. 92, 7956-7960. Ridley, A.J., & Hall, A. (1994). Signal transduction pathways regulating Rho-mediated stress fiber formation: Requirement for a tyrosine kinase. EMBO J. 13,2600-2610. Schlaepfer, D.D., Hanks, S.K., Hunter, T., & van der Geer, R (1994). Integrin-mediated signal transduction linked to ras pathway by GRB2 binding to focal adhesion kinase. Nature 372, 786-791. Schmeichel, K.L., & Beckerle, M.C. (1994). The LIM domain is a modular protein-binding interface. Cell 79,211-219. Schwartz, M.A„ Schaller, M.D., & Ginsberg, M.H. (1995). Integrins: Emerging paradigms of signal transduction. Annu. Rev. Cell Dev. Biol. 11, 549-599. Turner, C.E., & Miller, J.T. (1994). Primary sequence of paxiUin contains putative SH2 and SH3 domain FAK

binding motifs and multiple LIM domains: Identification of a vinculin and ppl25 -binding region. J. Cell Science 107, 1583-1591. Weiskirchen, R., Pino, J.D., Malcama, T., Bister, K., & Beckerle, M.C. (1995). The cysteine-rich protein (CRP) family of highly related LIM-domain proteins. J. Biol. Chem. 270,28946-28954.

INTERACTIONS OF MEMBRANE RECEPTORS AND CELL SIGNALING SYSTEMS WITH THE CYTOSKELETON

Coral ie A. Carothers Carraway and Kermit L. Carraway

I. Introduction: Membrane Signaling and the Cytoskeleton II. Ion Channels III. Nonreceptor Kinases and the Cytoskeleton A. Src Family B. Abl Family IV. Receptor Kinases A. EGF-Induced Morphology Changes and EGF Receptor Association with Microfilaments B. p 18 5^^^ B2/neu Association with Microfilaments and a Signal Transduction Particle V. Conclusions and Prospects Acknowledgments References

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 207-238 Copyright © 1996 by JAI Press Inc. All rights of reproduction in any form reserved. ISBN: 1-55938-688-6 207

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I. INTRODUCTION: MEMBRANE SIGNALING AND THE CYTOSKELETON A major question in contemporary cell biology is how extracellular information triggers specific cellular responses by binding to plasma membrane receptors. Signal transduction mechanisms have evolved to satisfy the requirement to produce a wide variety of responses, including such phenomena as changes in cell metabolism, secretion, endocytosis, cell motility, and mitogenesis. The role of the cytoskeleton in particular cellular responses is quite varied, depending on the cell type, the signal involved and the magnitude of the response. For example, changes in cell metabolism, such as those invoked by insulin in adipocytes, can occur with little apparent involvement of the cytoskeleton. Endocytosis and exocytosis, in contrast, must necessarily involve a substantial remodeling of parts of the cortical cytoskeleton. Motility necessitates plasticity of the cytoskeleton and major remodeling of specific cytoskeletal structures, particularly the cortical actin network. Cell-cell and cell-matrix interactions result in morphological alterations requiring major reorganization of cytoskeletal structures. Cytoskeletal reorganizations occurring as a result of these interactions play a vital role in normal differentiation and development. Mitogenesis, which culminates in cell division, represents an extreme in cytoskeletal reorganization, requiring global changes in all of the cellular cytoskeletal systems. Many of these same types of morphological and cytoskeletal changes occur during neoplastic transformation, in which the cell loses its normal growth regulation. Functionally, transformation involves mutations of the genes (proto-oncogenes) coding for components of normal cellular signal transduction pathways to their oncogenic forms, interrupting the tight regulation of cell growth and behavior imposed by normal signal transduction mechanisms. These signal transduction components and their oncogenic analogs can be grouped into four classes of proteins, based on their localization and function. (1) Extracellular ligands of various types provide the primary signal. (2) Membrane receptors, which are usually highly specific, bind ligand and become activated. The activated receptors are coupled, often via the production of second messengers, to the activation of intracellular enzymes and pathways. (3) Cytoplasmic transducers are enzymes and other proteins involved in transduction of signals through the cytoplasm to various sites of action, including the nucleus. (4) Transcription factors, proteins which regulate gene expression, are modulated by the transduced signal as part of the mitogenic response. Signaling via hydrophilic ligands occurs by binding to cell surface receptors. Four general classes of cell surface receptors based on both structural and functional characteristics have been identified. Structurally, the most complex receptors are the ligand-gated ion channel receptors, multi-subunit receptors forming pores which are opened and closed by means of ligand-induced conformational changes.

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The signaling ion then regulates intracellular metabolic pathways or secretory functions by modulation of intracellular transducers. Serpentine receptors, which have several transmembrane domains (frequently seven) and no intrinsic enzymatic activity, are targets for several of different types of ligands. They respond by modulation of membrane-associated enzymes such as adenylate cyclase and phospholipase Cg^^^^^^ (^LCg^^^j^J to increase the cytoplasmic concentration of second messengers, including cyclic AMP, specific types of membrane lipids and Ca^"^, which activate specific intracellular kinases. The modulation of the membrane enzymes is accomplished via trimeric G-proteins (GTPase proteins) which transmit the signal from the activated receptor to the enzymes. Structurally, the simplest receptors are those possessing single transmembrane domains and intrinsic enzymatic activities (kinases, phosphatases, one class of guanylate cyclase). The best characterized of these are the receptors for growth factors and polypeptide hormones, which are tyrosine-specific kinases. Ligand binding to the receptor itself results in dimerization, autophosphorylation and subsequent binding and phosphorylation of cytoplasmic proteins. The overall mechanism of signal transduction via receptor kinases is a cascade of phosphorylations and dephosphorylations of appropriate substrate proteins, which are frequently themselves kinases. Activated kinases modulate the function of target proteins, often through a complex sequence of events. Signaling events are transient and reversible; the cell returns to an "unactivated" state by a number of mechanisms, notably, the action of protein phosphatases and ligand-receptor internalization. A special class of receptors which does not fit into any of the categories described above are those which do not themselves have catalytic activity but which bind to and signal through nonreceptor cytoplasmic tyrosine kinases. Two well-characterized examples of this type of receptor are integrins and the T cell receptor. These will be described in more detail in the section on non-receptor kinases. The signal is transferred through the cytoplasm via intracellular transducers, 3. heterogeneous group of both enzymes and non-enzymatic proteins. These include cytoplasmic tyrosine kinases, serine/threonine-specific kinases of the MAP kinase pathway, protein kinase C and the cyclic nucleotide-dependent kinases. Information flow through the kinase pathways is often controlled by small G-proteins related to the ras family of (proto)oncogenes, which act as molecular switches, modulated by their GTPase activities. Lipid-converting enzymes such as phospholipase C and phosphoinositide-3-kinase (PI3K), are also important in the generation of second messengers and in regulating cell behavior, particularly via modulation of cytoskeleton plasticity. Transduction elements are often associated in multimeric complexes, linked together by adaptor proteins, which have no apparent enzymatic activity. These contain specific phosphorylation sites and specific domains for binding phosphorylated tyrosine residues (SH2 domains) and/or for binding proline-rich regions of some cytoskeleton-associated proteins (SH3 domains), which will be described further. These proteins are thought to serve a structural role

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in the organization of signal transduction pathways, particularly as they link to the cytoskeleton and morphological changes elicited by growth factors. The last class of signal transduction components, the transcription factors, encompass a group of proteins which ultimately control cell division by regulating transcription. Numerous recent studies have emphasized the importance of phosphorylation-dephosphorylation in the function of transcription factors, and cell cycle-specific kinases and phosphatases are being identified and described (Solomon, 1993). Whether the cytoskeleton is involved in any of these events is unclear, but the nonreceptor tyrosine kinase Abl may play an integrative role in cytoskeletal modulation of transcription. This multifunctional protein has been shown to phosphorylate transcription factors, to bind microfilaments and to partition between the cytoplasm and nucleus (Wang, 1993), as discussed later. Both the temporal and the spatial organization of the components of signal transduction pathways are critical to the ultimate cellular response to ligand. Transduction of signal requires that the appropriate substrate and/or binding proteins be localized proximal to the activated receptor or subsequent enzymes in the pathway. These components form at least transient associations into complexes, called signal transduction particles (Ullrich and Schlessinger, 1990), which are being characterized in several cell systems. Specific molecular mechanisms participating in this critical localization of substrate proteins have been identified. These include acylation in membrane targeting of proteins whose function requires membrane association, and SH2 domains in recruitment of substrate proteins, often enzymes, to the membrane for interaction with their membrane-localized substrates. Mechanisms for linking these signaling complexes to the cytoskeleton include SH3 domains in linking activated receptors to cytoskeletal elements via membrane skeletal proteins; other known actin-binding domains involved in binding membrane-associated proteins to actin; and specific lipids in the modulation of actin organization. Association with the cell cytoskeleton has been postulated for numerous signal transduction components. However, research in this area is still in an embryonic stage of development and many of these studies are fragmentary. In only a few cases have associations of any of these proteins with specific cytoskeletal proteins been demonstrated. Of necessity, this review will focus on the best-studied examples of the involvement of the cytoskeleton in signal transduction processes rather than attempt a global survey. Three different approaches have been important in defining the systems and processes described in this review. The first is the study of normal physiological processes in specialized systems, exemplified by investigations of the ligand-gated ion channel nicotinic acetylcholine receptor at the neuromuscular junction and Torpedo electroplax membrane. The second is the study of neoplastic transformation, from which arose a multitude of viral and cellular oncogenes that provided insights into many key cell regulation pathways. The third is the use of genetics to probe pathways for differentiation and development in simpler organisms and

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developmental systems. The complementarity of these approaches provides not only a rich experimental background for diverse investigations, but also a reminder of the unity of biology and the importance of evolution in understanding biological phenomena.

II. ION CHANNELS Conceptually, the simplest signal transduction mechanism involves moving the primary signal from the outside to the inside of the cell. This process is best exemplified by ligand-gated ion channels, which permit ion movements into cells in response to an extracellular signal. Localization of these channels in cells plays an important role in their function, and cytoskeletal elements have been implicated in the structures important to that localization in a number of cell types (Srinivasan et al., 1988; Froehner, 1991). The best studied example of a localized ion-gated channel is the nicotinic acetylcholine receptor from skeletal muscle, which permits influx of sodium and potassium ions in response to the binding of acetylcholine (Salpeter, 1987; Bloch and Pumplin, 1988). The acetylcholine receptors are localized to and function at the neuromuscular junction in the postsynaptic membrane (Salpeter, 1987). In an adult animal the synapses occupy only about 0.1% of the muscle cell membrane (Fraser and Poo, 1982). Extrasynaptic receptors appear to be randomly dispersed in embryonic or deinnervated muscle (Phillips and Merlie, 1992). Moreover, the acetylcholine receptor is a pentameric complex of four subunits (Stroud and Finer-Moore, 1985) which is readily diffusible in membranes (Salpeter, 1987). Thus, receptors must be anchored at the junction site. The components and mechanisms involved in this anchorage have been a major focus of research in neuromuscular synaptogenesis. A large number of cytoskeletal proteins have been identified in Torpedo acetylcholine receptor clusters or mammalian neuromuscular postsynaptic membranes, including a novel isoform of P-spectrin (Bloch and Morrow, 1989), actin, vinculin, talin, paxillin, filamin, a-actinin, tropomyosin 2, dystrophin, and ankyrin (Froehner, 1991). However, only a subset of these are likely to be involved in receptor clustering: actin, (3-spectrin, and tropomyosin. In addition, a dystrophinrelated protein and proteins of 43, 58 and 87 kDa have been implicated. Ultrastructurally, the receptor cluster resembles the membrane skeleton of the erythrocyte membrane, containing a network of filaments about 6 nm in diameter (erythrocyte, 5.3—5.9) and 29 nm in length (erythrocyte, 28) into which is inserted actin filaments (Pumplin, 1989). Since P-spectrin is the major protein in the synaptic membrane skeleton (4—7 spectrin molecules per acetylcholine receptor), the clusters appear to be stabilized by a network of these specialized spectrin molecules attached to F-actin. The mechanism of association of this network with the receptor is of particular interest. Extraction studies indicate that both the 43 and 58 kDa proteins are present in approximately equimolar ratio with the receptor and are more tightly associated

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with the membrane than spectrin (Pumplin and Bloch, 1993). Extraction and localization studies suggest that the 43 kDa protein is directly associated with the receptor (Froehner, 1991). Moreover, two different recombinant DNA approaches show the involvement of the 43 kDa protein in receptor clustering. First, injection of receptor subunit mRNAs into Xenopus oocytes resulted in a diffuse distribution of receptor on the oocyte surface. The receptors became clustered upon injection of the mRNA for the 43 kDa protein (Froehner et al, 1990). Similarly, transfection of quail fibroblasts with cDNAs for the receptor subunits led to a diffuse distribution of receptors, which became clustered upon transfection with the cDNA for the 43 kDa protein (Phillips et al., 1991). In addition, introduction of recombinant 43 kDa protein into cells in the absence of receptor led to the formation of domains enriched in the 43 kDa protein. Thus, this protein is capable of forming specialized domains without an interaction with receptor. These domains can then provide association sites for receptors, either by trapping mobile plasma membrane receptors or by acting as a site for insertion of newly synthesized receptors being incorporated into the plasma membrane. The connecting link from the receptor-43K complex to spectrin is undefined. Since actin overlay studies show binding to the 43 kDa protein (Walker et al, 1984), one possibility is that the 43 kDa protein binds directly to actin oligomers associated with p-spectrin, serving a frmction similar to that of ankyrin of the erythrocyte membrane. Alternative or additional mechanisms should also be considered, perhaps involving the 58 kDa and/or 87 kDa proteins. Immunoaffinity analyses suggest an association of the 58 and 87 kDa proteins; both of these can be immunoprecipitated with anti-dystrophin from detergent-solubilized Torpedo membranes (Butler et al., 1992). Dystrophin is a large spectrin-like protein with an a-actinin-like domain and an actin binding domain (Dubreuil, 1991), which may allow dystrophin to serve as a link to microfilaments. Moreover, p-spectrin, but not dystrophin, is found in neuromuscular junction receptor clusters, and dystrophin, but not p-spectrin, is found in Torpedo membranes. Thus, the two proteins may be serving the same function in the two systems (Sealock et al., 1991). Sequence comparisons of 43 kDa proteins from different species show four conserved domains, including an N-terminal myristylafion site, a leucine zipper domain and a C-terminal zinc finger domain. A deletion mutant protein containing only the N-terminal domain and C-terminal third of the sequence formed membrane-associated aggregates but did not cluster receptors. The C-terminal third of the molecule was also required for membrane association. Thus, myristylation may be necessary for binding to the membrane, but it is not sufficient. These results suggest that the receptor binding site is in the central domain of the molecule. The mechanism of the involvement of the 43 kDa protein in synapse formation is still uncertain. Both the acetylcholine receptors and the 43 kDa protein can aggregate independently in membranes. How they form complexes and aggregate is under investigation. Experiments with the receptor-clustering factor agrin (Campanelli et al., 1992) have suggested a role for phosphorylation of the receptor

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subunits (Wallace, 1991). Receptor clustering is inhibited by expression of the tyrosine kinase pp60^'^'^ in myotubes. Moreover, treatment of myotubes with fibroblast growth factor-coated latex beads induced acetylcholine receptor clustering, which was inhibited by the tyrosine kinase inhibitor tyrphostin (Peng et al., 1991). These results suggest the involvement of tyrosine phosphorylation in the clustering phenomenon. Other protein kinases have also been localized to postsynaptic densities, including protein kinase C, calmodulin-dependent kinase and cAMP-dependent kinase (PKA; Hubbard and Cohen, 1993). The PKA is linked via specific anchoring proteins, which bind to its regulatory subunit (Carr et al., 1992). How the anchoring proteins are associated with the membrane and/or cytoskeletal components is unknown. However, these results indicate that the postsynaptic densities contain a multifunctional regulatory complex which undoubtedly plays an important role in its formation, stabilization and function.

III. NONRECEPTOR KINASES AND THE CYTOSKELETON Protein kinases can be divided into two structural classes: receptor kinases, which are integral membrane proteins, and nonreceptor kinases. Non-receptor kinases may be either cytosolic or associated with membranes by mechanisms other than transmembrane domains. Kinase specificity is of particular importance in signal transduction, and kinases are also classified into two groups based on specificity: tyrosine and serine/threonine kinases. Much of the early work in signal transduction has emphasized the role of tyrosine kinases in the early events at the membrane and of serine/threonine kinases in later events, for example, phosphorylation of transcription factors. However, receptor serine/threonine kinases have recently been described (Massague, 1992), and the tyrosine kinase Abl is believed to act in the nucleus (Wang, 1993). Tyrosine kinases catalyze both autophosphorylation and phosphorylation of other protein substrates. Phosphorylation of protein tyrosines has two important consequences: (1) it can alter the structure and activity of the protein modified and (2) the resulting phosphotyrosine residue can act as a specific binding site for other proteins via SH2 domains, which bind tyrosine-phosphate. Conceptually, the mechanism of action of receptor kinases is simple, e.g., conformational activation of the kinase by binding of ligand. The activation mechanism is less well understood for membrane-associated non-receptor kinases, some of which appear to be activated after ligand binding to cell surface receptors. Perhaps the best studied example is the T cell receptor of lymphoid cells (Perlmutter et al., 1993). Although no subunit of the receptor complex has tyrosine kinase activity, ligand binding by the receptor rapidly triggers tyrosine phosphorylation of cellular proteins. Several kinases have been implicated, including two lymphoid cell-specific members of the Src family, Lck and Fyn (Samelson and Klausner, 1992). These kinases have been shown to associate with components of the T cell receptor complex (Shaw and Thomas, 1991; Izquierdo and Cantrell, 1992). How the association of kinases with receptors is linked to the triggering of phosphorylation

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in T cells is unclear. Interestingly, detergent extraction studies suggest that Lck is cytoskeleton-associated (Louie et al., 1988), but the significance of the observation is presently unknown. A. Src Family

The src gene family of tyrosine kinases occupies a central role in the history and understanding of signal transduction. Identification of src as the first known retroviral transforming gene (Brugge and Erikson, 1977) provided a linkage between oncogenic transformation and protein function which led to the recognition of the importance of tyrosine kinases in signal transduction and cell regulation (Hunter and Cooper, 1985). The src oncogene encodes pp60^"^'"^ (v-Src), the prototype of a family of phosphorylated tyrosine-specific protein kinases (Hunter and Cooper, 1985). Investigations of Src in non-transformed cells demonstrated a proto-oncogenic form c-Src, which is present in unstimulated cells in an inactive form. v-Src differs from c-Src by the deletion of the C-terminal segment containing Tyr 527, which, when phosphorylated, causes inactivation of the enzyme (Cooper and Howell, 1993). This deletion results in activation of v-Src kinase and oncogenic activities. Membrane localization is also critical to Src activity. The mechanism for at least the initial association with the membranes is via myristylation of the N-terminal glycine of Src. Mutants that are non-myristylated are non-transforming (Cross etal., 1984). Investigations of the structure of Src showed that it contains three domains common to a number of components of signal transduction pathways. These are designated Src homology domains SHI (tyrosine kinase domain), SH2 and SH3. All protein kinases, regardless of localization or specificity, show significant homologies in their SHI or kinase domains. SH2 domains are conserved protein structural motifs containing approximately 100 amino acids, which function by binding phosphorylated tyrosine residues of proteins, including the phosphorylation sites on tyrosine kinases (Koch et al., 1991; Pawson and Gish, 1992; Mayer and Baltimore, 1993). They are found in a number of proteins which are important in signal transduction (Table 1). SH2 domains exhibit both high affinity binding and a high degree of specificity for particular amino acid sequences surrounding the phosphorylated tyrosine residues. Thus, SH2 domains can serve as specific links between components of signal transduction pathways (Cantley et al., 1991). An interesting situation arises when a protein has both a tyrosine phosphate and an SH2 domain. If the specificity and geometry permit, these may form an intramolecular interaction which can alter the activity of the protein and block associations with other proteins. For example, phosphorylation of Tyr 527 in the C-terminal tail of Src forms a binding site for the Src SH2 domain. This intramolecular interaction has been proposed to provide an autoinhibitory function for c-Src which is absent in v-Src because of the deletion of the C-terminal region (Cooper and Howell, 1993).

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Table 1. Classes of Proteins Containing SH2 and SH3 Domains SH2

SH3

Other Domains or Activities

Non-receptor kinases and other enzymes Src family Abl family CSK Tec Fps/Fes 2 Syk 2 PTPIC 2 PLCgamma ISGF3a 1

1 1 1 1 0 0 0 1

Dig

0

1

p47 and p67 phox

0

2

tyrosine kinase tyrosine kinase, actin binding, DNA binding tyrosine kinase tyrosine kinase tyrosine kinase tyrosine kinase tyrosine phosphatase phospholipid hydrolase HLH , leucine zipper; transcriptional activator guanylate kinase (Drosophila tumor suppressor) activate neutrophil cytochrome oxidase

GTPase-modulating proteins Ras GAP CDC25, ste6

2 0

1 1

Ras GTPase activator GNEF^

Cytoskeletal proteins Tensin Nonmuscle myosin fodrin/spectrin cortactin p80/p85;HSl ABP-1 BEMl FUSl

1 0 0 0 0 0 0 0

0 1 1 1 1 1 1 1

cytoskeleton-membrane cytoskeleton-membrane cytoskeleton-membrane at focal adhesions at focal adhesions yeast actin binding protein cytoskeleton/yeast cytoskeleton/yeast

Adaptor proteins PI3Kp85 GRB2 (Drk/Sem5)

2 1

1 2

rhoGAP; links PI3K to activated receptor human (Drosophila, yeast); links through SH2 to activated receptors and through SH3 to Sos, a Ras-activating GNEF^ dbl domain (GNEF?); DAG/PE-binding domain function unknown function unknown function unknown

Vav Crk She Nek

1

Source: Information summarized from Koch et al. (1991), Mayer and Baltimore (1993), and Pawson and Gish (1992). Notes: ^helix-loop-helix DNA binding domain ^guanine nucleotide exchange factor •'diacylglycerol/phorbol ester

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SH3 domains are conserved motifs of about 60 amino acids, which bind to proline-rich regions of specific proteins (Ren et al., 1993). These domains may also act as regulatory elements, since deletions or mutations can increase catalytic activity and oncogenic potential of tyrosine kinases in which they are present (Koch et al, 1991; Pawson and Gish, 1992; Mayer and Baltimore, 1993). However, they are most often considered to be cytoskeleton binding sites (Drubin et al, 1990), because they occur in a number of proteins which are associated with the cytoskeleton (Table 1). Several proteins have both SH2 and SH3 domains (Table 1). Some of these have little or no structure or identifiable functional domains outside the SH domains and are thought to be adaptor proteins which serve as linking elements of signal transduction pathways. One of the consequences of neoplastic transformation of cells by oncogenes such as src is an alteration of cell morphology (Burridge, 1986). Since morphology is dependent on the organization of cytoskeletal elements in the cell, there has been a great deal of interest in the cellular localization of Src and the role of its localization in regulating the cytoskeleton (Kellie, 1988). Numerous studies have shown that v-Src is a membrane-associated protein which localizes to cytoskeletal structures in cells, particularly to adhesion plaques (Krueger et al., 1983). In contrast, c-Src is soluble in nonionic detergent, and not strongly associated with the cytoskeleton (Loeb et al., 1987). This localization to the membrane-cytoskeleton interface apparently plays an important role in neoplastic transformation (Hamaguchi and Hanafusa, 1987). Targeting of v-Src to adhesion plaques, but not the nucleus or perinuclear membranes, will transform chick fibroblasts (Liebl and Martin, 1992), which reduces their adhesivity and releases them from cell cycle control. Although the mechanism for the association of Src with the cytoskeleton is unclear, the binding site on v-Src has been mapped to the SH2 domain (Fukui et al., 1991). However, v-Src and c-Src have identical SH2 domains. One explanation for the differential localization is that the SH2 domain of c-Src is blocked by an intramolecular interaction with phosphorylated tyrosine 527 of the C-terminal region, which is missing from v-Src. Phosphorylation of this residue is also proposed to provide an autoinhibitory function for c-Src. Thus, the phosphorylation-dephosphorylation cycle of Tyr 527 is implicated in the regulation of both kinase activity and cytoskeleton association through changes in its intramolecular interaction with the SH2 domain. A similar mechanism has been observed in platelets, which contain a high content of c-Src (Golden et al., 1986). Thrombin activation of platelets results in an activation of their Src kinase, followed by a redistribution of the kinase to the cytoskeleton (Clark and Brugge, 1993). Microfilament association and activation of Src require dephosphorylation of Tyr 527. This dephosphorylation is proposed to disrupt the association of phosphorylated Tyr 527 with the Src SH2 domain and permit an interaction with some unknown tyrosine-phosphorylated cytoskeleton component (Clark and Brugge, 1993).

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The mechanism described above does not exclude the possibility of other modes of association of Src with the cytoskeleton through its other structural elements. The Src SH3 domain might also contribute to an association with the cytoskeleton. We have recently shown that biotinylated Src SH3 domain will bind several microfilament core-associated proteins from ascites tumor cell microvilli. One of these is a membrane- and micro filament-binding 58 kDa protein implicated in cell surface stabilization (Liu et al, 1989). This protein contains a proline-rich sequence (Juang et al., 1994) typical of those described for SH3 binding peptides (Ren et al., 1993). Furthermore, in vzYro-translated 58 kDa protein co-immunoprecipitates with Src from platelet extracts (Huang et al., submitted). The 58 kDa protein also contains phosphorylated tyrosine(s) and can potentially form linkages with both SH2 and SH3 domains. We propose that such molecules might be called "retroadaptors" and that they might serve to link Src or other proteins to the cytoskeleton or other cellular elements and to modulate the activities of Src-like enzymes. These results indicate the possibility of different Src association mechanisms in different contexts or of multiple, redundant association mechanisms. How Src association with adhesion sites contributes to the transformed phenotype is still unclear, since the Src signal transduction pathway is not yet understood. However, recent studies on the dynamics of adhesion sites provide some interesting insights which should lead to a better understanding. Investigations of the early stages in the formation of focal adhesions have demonstrated a concomitant increase in the tyrosine phosphorylation of a limited set of proteins, particularly those at about 70, 115, and 130 kDa (Romer et al., 1992). A similar tyrosine phosphorylation of 115—130 kDa components can be induced by antibody clustering of integrins (Komberg et al., 1991; Juliano and Haskill, 1993). One of the components phosphorylated in the presence of integrin crosslinking has been identified as a cytoplasmic tyrosine kinase which is associated with focal adhesions, ppl25^^^ (Schwartz, 1992; Schaller and Parsons, 1993). Other cell stimulating agents, including phorbol esters and bombesin, will also induce phosphorylation of ppl25^^^ in quiescent fibroblasts (Sinnett-Smith et al., 1993). The phorbol ester but not the bombesin effects are dependent on protein kinase C, and both types of effects are inhibited by cytochalasin disruption of cellular microfilaments. Thus, pp 125^"^^ appears to be involved in a complex network of regulatory events, located at least partially at the focal contact membrane-microfilament interface. Transformation of cells by v-src increases both cellular phosphotyrosine content (Kanner et al., 1990) andppl25^^^ enzymatic activity (Guan and Shalloway, 1992). Moreover, in the transformed cells a substantial fraction of the ppl25^'^^ is present in a stable complex with pp60^^^, mediated by the SH2 domain of pp60^^^ (Shaller and Parsons, 1993). These results suggest that a complex of Src and ppl25^'^^ may be important in the regulation of focal contacts, and that their interaction involves the same domain implicated in Src kinase activity and cytoskeleton association. The mechanism by which tyrosine phosphorylation is linked to changes in the cytoskeleton and focal contacts is unknown. The focal adhesion proteins tensin and

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paxillin can be phosphorylated by ppllS^"^^, but the consequences of those modifications are unknown (Schaller and Parsons, 1993). Tensin is a multidomain protein with an SH2 domain (Davis et al., 1991), three different actin binding domains and a domain related to catenin-a (Lo et al., 1994), a vinculin-related protein associated with the cadherins of cell-cell interaction sites (Nagafuchi et al., 1991). Thus, tensin has multiple sites for potential interactions with both the cytoskeleton and with other signal transduction components. However, there is little information currently available about the proteins to which tensin binds. Other signal transduction components may also be involved in cytoskeletal changes at focal contacts. Protein kinase C activation will induce focal adhesion formation (Woods and Couchman, 1992), and an isoform of kinase C is partially localized in focal contacts of rat embryo fibroblasts (Jaken et al, 1989). Moreover, recent studies have demonstrated the involvement of a small G-protein in the regulation of cell morphology, actin organization and focal contacts (Chrzanowska-Wodnicka and Burridge, 1992; Hall, 1993). Injection of p21rho into quiescent fibroblasts resulted in the formation of stress fibers and focal adhesions (Ridley and Hall, 1992). One explanation for these results is that p21rho is essential to a pathway involved in the formation or stabilization of focal adhesions. Whether p21rho is associated with the adhesion sites is uncertain. Such a possibility appears more attractive in light of the observation that a small G-protein Rap IB has been shown to become associated with the platelet cytoskeleton during activation (Fischer et al., 1990). These combined studies suggest the presence of a multimeric signal transduction complex(es) in focal contacts which could not only regulate cytoskeletal structure, but also couple changes in that structure to other cellular events, such as control of the cell cycle and cell division. Members of the Src family of kinases have also been implicated in the dynamics of cell-cell interactions involving cadherins. Cadherins are a family of calcium-dependent cell surface adhesion receptors (Takeichi, 1991; Magee and Buxton, 1991) which play a critical role in tissue morphogenesis (Kintner, 1992). As is the case for the integrins in focal adhesions, the cadherins provide a link between the associations at the cell surface and the cytoskeleton of the cell cytoplasm. The cytoplasmic surface of the plasma membrane at cadherin-containing cell-cell interaction sites has an associated cytoplasmic plaque which contains vinculin, a-actinin and actin filaments. Cadherins are linked to microfilaments via complexes with cytoplasmic proteins, called catenins (Ozawa and Kemler, 1992), one of which (catenin-a) has strong similarities to vinculin (Nagaftichi et al., 1991). Cell-cell junctions are dynamic structures, which change during processes such as cell migration, mitosis and neoplastic transformation (Takeichi, 1991; Volberg et al., 1992). Several types of evidence have implicated tyrosine kinases in these changes: (I) phosphotyrosine-containing proteins are concentrated in the intercellular junctions of epithelial cells (Maher et al, 1985); (2) treatment of several rat tissues with orthovanadate, a protein tyrosine phosphatase inhibitor, increased the amounts of

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tyrosine phosphophorylation in plaques of cell-cell junctions (Tsukita et al., 1991); (3) similar treatments of MDCK cells led to an increase in intercellular junction phosphotyrosine, followed by a concomitant reduction in intercellular junctions and a marked increase in focal adhesions (Volberg et al., 1992); (4) three members of the Src family (Yes, Src, and Lyn) were found associated with isolated liver cell-cell junctions (Tsukita et al., 1991); (5) transformation of lens epithelial cells with a temperature-sensitive mutant of Rous sarcoma virus resulted in an altered morphology and reduced cadherin-associated cell-cell adhesions at the permissive, but not the restrictive, temperature (Volberg et al., 1991), accompanied by increased tyrosine phosphorylation of cell-cell junctions; (6) treatment of these transformed cells with tyrosine kinase inhibitors resulted in reformation of junctions (Volberg et al., 1992); and (7) transformation of chick embryo fibroblasts with transforming, but not non-transforming, mutants of v-^rc suppresses specific cadherin-mediated adhesion without inhibiting cadherin expression (Hamaguchi et al., 1993). Both cadherin and catenins are tyrosine-phosphorylated in the transformed cells. The level of phosphorylation of the cytoskeleton linker catenin (approx 25%) is much higher than that found for other Src-phosphorylated proteins at membrane-microfilament interfaces, such as vinculin, talin, integrin and calpactin. These combined results suggest that tyrosine kinases located at the intercellular junction plaque play an important role in the dynamics of cell-cell interactions via a phosphorylationdephosphorylation cycle. B. Abl Family Abl was originally discovered as the product of the v-abl oncogene (Wang, 1993). Homologs of c-abl have been described from humans, mice, Drosophila, and C. elegans; an analogous gene arg is also present in humans. The c-abl gene encodes a protein which has similarities to Src, but is about twice the size. It contains the kinase, SH2 and SH3 domains and N-myristylation site similar to Src in the N-terminal half of the molecule. Furthermore, it also has a nuclear translocation sequence, a DNA binding domain and an actin binding domain similar to domains in a-actinin, spectrin and fimbrin (Wang, 1993). Constitutive activation of the kinase occurs in three oncogenic Abl variants. In mouse and cat oncogenic viruses the SH3 and C-terminal regions, respectively, have been deleted and the remainder of the protein fused to viral protein sequences. In the human oncogene the c-abl gene has been fused to the bcr (breakpoint cluster region) gene. Interestingly, the bcr-abl gene products have not only activated kinase, but also an increased microfilament binding ability (McWhirter and Wang, 1991). These effects are not interdependent, as they are localized to different regions of the bcr gene. These results suggest that both kinase and actin binding functions are auto-inhibited in c-Abl, as they are in c-Src. Removal of the SH3 domain of Abl activates its kinase and transforming potential (Franz et al., 1989; Jackson and Baltimore, 1989). Thus, Abl functional activity

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CORALIE A. CAROTHERS CARRAWAY and KERMIT L. CARRAWAY

could potentially be regulated by an intramolecular SH3-binding mechanism or by binding of other cellular proteins to the SH3 domain (Pendergast et al., 1991). Use of the biotinylated Abl SH3 domain as a probe allowed isolation of a cDNA from a lymphoid cell line which encodes a protein which binds SH3 domains selectively (Cicchetti et al., 1992). The sequence of this protein, called 3BP-1, was similar to that of the C-terminal segment of Bcr and to a GTPase-activating protem (GAP-rho) related to the GAP proteins which inactivate Ras. Thus, this SH3-binding protem is capable of linking Abl, as well as other selected SH3-containing proteins, to G-protein switches in signal transduction pathways. Moreover, the relationship to Rho suggests a potential involvement in the regulation of the cytoskeleton through the effects of Rho on stress fibers and focal adhesions (Ridley and Hall, 1992). Bcr-Abl is generally less oncogenic than Gag-v-Abl, a phenomenon which has been attributed to the differential localization of the proteins (McWhirter and Wang, 1993). In fibroblasts which overexpress the protein, a large fraction of the c-abl product is localized in the nucleus (van Etten et al., 1989). Deletion of an N-terminal segment from this protein activates its transforming potential and shifts its primary localization to the cytoplasm and plasma membrane. Bcr-Abl, but not Gag-v-Abl, has been shown to be associated with cellular stress fibers (McWhirter and Wang, 1993). Moreover, the isolated Abl C-terminal domain will bind F-actin in vitro. Deletion and mutation studies have shown that transformation by Bcr-Abl in fibroblasts is correlated with actin association. Thus, the behavior of Abl and the consequences of its activation are complex. One explanation of these results is that Bcr-Abl acts at multiple sites (nucleus, microfilaments, plasma membrane) in order to exert its effects. An alternative is that the kinase acts at a single site, and that modifications of the protein alter its distribution among various locations. Further information on the pathway(s) as well as the target(s) for the kinase is important for a better understanding of its role(s) in both normal and transformed cells.

IV. RECEPTOR KINASES Receptor kinases provide the cell surface transduction mechanisms for a variety of growth factors, hormones and cytokines which regulate proliferation and differentiation of cells. Since morphological transformations are an important aspect of these effects, the pathways initiated through these receptors must play a key role in integrating the response of the cytoskeleton with the responses of other cellular activities. Until recently the mechanisms for the transduction of signal through receptors to the cytoskeleton and the nucleus were largely unknown. Although many aspects, especially the more downstream steps, of most signal transduction pathways are still uncertain, two important features have emerged as critical early steps (Fantl et al., 1993). First, when stimulated by ligand binding to the extracellular domain, the cytoplasmic tyrosine kinase domains catalyze both autophosphorylation and phosphorylation of other cellular proteins. Second, the cytoplasmic domain autophosphorylated sites act as binding sites for proteins which bind

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tyrosine phosphate via SH2 domains. Thus, these kinases act not only as enzymes but also as organizing centers for signal transduction pathway components. Because of the specificity of the SH2 binding for sequences containing phosphotyrosine residues, each receptor forms different complexes. Indeed, different phosphorylated tyrosine residues in a single receptor can bind different substrate proteins. For example, autophosphorylated PDGF receptor binds GTPase activating protein (GAP), the modulator of Ras-like G proteins (Downward, 1992; Lowy and Willumsen, 1993; Polakis and McCormick, 1993), PI3K and PLCg^^^^ at different phosphorylated tyrosine residues of its cytoplasmic domain (Fantl et al, 1993). Mutational analyses of these sites have shown that PI3K and PLCg^^^^^^ are involved in independent pathways which are required for mitogenesis. Activated EGF receptor also binds specific SH2-containing signaling proteins, including PLC ^^^ and PI3K. These lipid-metabolizing enzymes are of particular interest because of the involvement of PLC j^^ in the production of phosphoinositides which regulate actin organization and of PI3K in receptor trafficking involving microtubules (described below). Interestingly, the insulin receptor appears not to form direct complexes with known signal transduction components. Instead, the receptor phosphorylates an intermediate, a 185 kDa protein, which binds certain SH2-containing proteins (Folli et al., 1992). The general role of these SH2 interactions with phosphorylated tyrosine residues appears to be the organization of signal transduction complexes, particularly the first components of the pathway. Indeed, this relocalization of SH2-containing proteins to the activated receptor and their consequent activation has become a paradigm in signaling through growth factor receptors (Cantley et al., 1991). Interactions via these specific binding sites accomplish several objectives: (1) allow localization of critical enzyme activities to the membrane, where their substrates are located, (2) enhance the phosphorylation of certain protein substrates on tyrosine residues by forming stable complexes with the receptor tyrosine kinase, (3) act as links in the formation of complexes with other components of signal transduction pathways, and (4) induce conformational changes in other regulatory components to enhance their activities (Mayer and Baltimore, 1993). These possibilities are not mutually exclusive, particularly since the coordinate actions of a number of factors are involved in pathways such as mitogenesis. The roles of SH2 and SH3 domains in signal transduction via receptor tyrosine kinases are exemplified by the mitogenic pathway recently described for EGF, which couples the growth factor-activated receptor via an unbroken series of steps to mitogen-activated protein kinases (MAP kinases) which activate transcription factors (Hunter and Karin, 1992). There are three key elements in this pathway (Carraway and Carraway, 1995): the transmembrane receptor, which receives the growth factor signal and transmits it via the activated tyrosine kinase; the small G protein Ras, which acts as a molecular switch and modulator via its GTP binding and GTPase activity (Downward, 1992; Lowy and Willumsen, 1993; Polakis and McCormick, 1993); and the MAP kinase cascade, which transmits the signal to the

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CORALIE A. CAROTHERS CARRAWAY and KERMIT L. CARRAWAY

nucleus (Blenis, 1993; Davis, 1993). Ras, like Src, requires fatty acylation and membrane association for activity. Ras is inactivated by binding of GAP, a GTPaseactivating protein. Inactive Ras can be activated by nucleotide exchange proteins which accelerate exchange of GTP for the GDP produced by the GTPase. In the EGF-stimulated signal transduction pathway, activation of Ras is coupled to EGF binding and autophosphorylation of the EGF receptor via an SH2/SH3 adaptor protein GRB2 (human; Drk, Drosophila; Sem5, yeast). GRB2 binds to a phosphorylated tyrosine residue of the receptor via its SH2 domain and to the nucleotide exchange protein SOS via its SH3 domain. Sos association with this complex promotes its ability to associate with and activate Ras (Egan et al., 1993). Activated Ras can associate with another proto-oncogene product, the Ser/Thr kinase Raf (Moodie et al., 1993), which is the first component of the MAP kinase (also called Erk) cascade. Raf phosphorylates and activates MAP kinase kinase (also called MEK), which activates the MAP kinases (Leevers and Marshall, 1992; Ruderman, 1993). Although this scheme describes a pathway by which the growth factor signal can be coupled to modulation of transcription factors, many of the details of the activation steps have not been elucidated. Also unclear is how the signal is transmitted from the plasma membrane to the nucleus and integrated with other cellular functions, such as remodeling of the cytoskeleton. Nonetheless, this pathway will undoubtedly serve as a prototype for investigating the organization of other signal transduction pathways. A.

EGF-lnduced Morphology Changes and EGF Receptor Association with Microfilaments

Growth factors regulate cell proliferation and differentiation through their actions on mitogenic pathways (Yarden and Ullrich, 1988). Thus, they play critical roles in processes leading to both normal and abnormal development, including oncogenesis (Cross and Dexter, 1991). Both growth factors and growth factor receptors have been identified as oncogenes. Growth factors have been observed to evoke a number of biological responses in vivo, in organ cultures and in cell cultures (Carpenter and Cohen, 1979; Yarden and Ullrich, 1988). Among the most important cellular effects are membrane ruffling, pinocytosis, increased transport, activation of glycolysis, stimulation of macromolecular synthesis, induction of nuclear (proto)oncogenes and enhanced cell proliferation. The perturbation of cell morphology (Chinkers et al., 1979, 1981;Kadowakietal., 1986;Lietal., 1993)isbest exemplified by studies of A431 cells, which overexpress the epidermal growth factor (EGF) receptor. Treatment of these cells with EGF resuhs in rapid formation of microvillus-like structures (30 sec), which then yield membrane ruffles (2—5 min) and cell rounding (10-20 min). These changes are accompanied by phosphorylation of two submembrane cytoskeletal proteins: ezrin, which was originally discovered in intestinal brush border microvillar cytoskeletons, and spectrin (fo-

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drin), the membrane skeleton component of erythrocytes and other cells (Bretscher, 1989). The exact mechanisms whereby these cytoskeletally mediated events are coupled to the binding of EGF to its receptor are uncertain. However, the interaction of the EGF receptor with submembrane microfilaments provides some interesting prospects. Biochemical, immunological and ultrastructural studies have indicated that the EGF receptor is associated with A431 cellular microfilaments (van Bergen en Henegouwen et al., 1989; Rijken et al., 1991). Although A431 cells contain both high and low affinity forms of the EGF receptor (King and Cuatrecasas, 1982), activation of the signal transduction cascade occurs through exclusive binding to the high affinity form (Defize et al., 1989). EGF receptors associated with microfilaments exhibit predominantly high affinity EGF binding (Wiegant et al, 1986), but association of low affinity sites can be promoted by EGF treatments (van Bergen en Henegouwen et al., 1989). The association of the receptor with microfilaments may occur via a direct interaction; recent studies indicate that the EGF receptor itself is an actin binding protein (den Hartigh et al, 1992), binding to actin via a sequence that resembles a sequence in profilin, the actin monomer-binding protein. Interestingly, deletion of this actin binding region does not prevent autophosphorylation or phosphorylation of PLC ^^^^^ or PI3K, but does reduce EGF-induced cell division (P. van Bergen en Henegouwen, personal communication). The mechanism by which EGF binding alters microfilament organization in cells is probably complex and likely involves a cascade of events. One interesting possibility for an immediate effect is that the autophosphorylation of the EGF receptor triggered by ligand binding changes the affinity of the receptor for actin. However, the more global changes in actin organization are likely to involve phosphoinositides, which have been shown to affect several actin binding proteins (Stossel, 1989; Carraway, 1990). One of the earliest events in EGF stimulation is the hydrolysis of phosphatidylinositol-4,5-bisphosphate (PIP2) to yield 1,4,5-IP3 and diacylglycerol. These phosphoinositides are important in cytoskeletal regulation because of their ability to alter the association of actin with specific actin binding proteins, such as gelsolin and profilin. Gelsolin is a filament severing protein also capable of forming complexes with actin which nucleate polymerization (Yin, 1987). Profilin is an actin monomer binding protein which blocks the ends of fast-growing filaments and accelerates nucleotide exchange (GoldschmidtClermont and Janmey, 1991). Profilin can also bind to phosphatidylinositides and prevent their hydrolysis by PLCg^j^j^^ (Goldschmidt-Clermont et al., 1990). However, when EGF-activated receptor binds and phosphorylates PLCg^^j^^ (Goldschmidt-Clermont et al., 1991), the inhibitory effect of profilin is released. This reciprocal modulation pathway is described in greater detail by Goldschmidt in this volume. The molecular details of the changes which occur in actin organization and their effects on cell structure and motility are still unresolved, but a number of the steps which may contribute have been described (Stossel, 1993).

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Hydrolysis of PIP2 triggers other events which may be important in cytoskeletal regulation. IP3 releases calcium from internal storage compartments. Calcium and the other released product diacylglycerol activate protein kinase C, a serine/threonine kinase which acts at the plasma membrane. One of the major substrates of protein kinase C is the MARCKS (myristylated, alanine-rich, C kinase substrate) protein (Aderem, 1992; Blackshear, 1993), a myristylated membrane protein which binds and crosslinks F-actin when dephosphorylated (Hartwig et al., 1992). Since the phosphorylated protein does not crosslink filaments, MARCKS has been proposed to serve as a regulator of filament organization. MARCKS is located at sites of membrane-microfilament interactions; thus, its phosphorylation-dephosphorylation cycle may be involved in the calcium-mediated control of F-actin association with the membrane (Aderem, 1992). This localization at the membranemicrofilament interface would allow it to participate in a number of important cellular functions, including motility, secretion and mitogenesis (Aderem, 1992). Because of the importance of phosphatidylinositide hydrolysis in the regulation of actin organization, there is considerable interest in the localization of PLC ^^^ and the role of that localization in actin organization. Immunocytochemical localization of PLC j^g in fibroblasts indicated that it is primarily associated with stress fibers (McBride et al., 1991). Moreover, the recombinant SH3 domain of PLCg^j^^^^ when microinjected into fibroblasts was also localized to stress fibers, while its SH2 domain exhibited a more diffuse cytoplasmic distribution (Bar-Sagi et al., 1993). These results suggest that localization of the enzyme to microfilaments is determined primarily by its SH3 domain, at least in unactivated cells. Activation and autophosphorylation of the EOF receptor creates a binding site for the PLCg^j^^^ SH2 domain which promotes its association with the receptor (Wahl and Carpenter, 1991). This membrane association would facilitate action of the phospholipase on its membrane-associated inositide substrate. This localization is also necessary for a role of phosphoinositides in regulating actin organization at the plasma membrane (Stossel, 1993). Thus, the SH2 and SH3 domains of PLCg^^^^^^ appear to be involved reciprocally in regulating the localization of the enzyme, depending on the state of activation of the EOF receptor. It is interesting to note that two related enzymes of inositide metabolism, phosphoinositide kinase (PIK) and diacylglycerol kinase (DGK), have also been found to be activated and associated with microfilaments when A431 cells are stimulated by EOF (Payrastre et al, 1991). However, localization of these enzymes has not been reported. Little is known of the role of PIK in cell function, but recent studies suggest that it plays an important role in signaling (McNamee et al., 1993). DGK is of considerable interest because it can reduce the levels of cellular diacylglycerol (DAG), which is not only a modulator of protein kinase C, but has also been implicated in microfilament nucleation at plasma membranes (Shariff and Luna, 1992). Although DAG produced immediately after cell activation is primarily from phosphatidylinositides, most of the DAG produced at later times appears to come from hydrolysis of phosphatidylcholine, (Liscovitch, 1992). Recent studies have shown that EGF can stimulate tyrosine phosphorylation

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as well as activation of DGK co-transfected into COS cells with the EGF receptor (Schaap et al., 1993). Interestingly, DGK is also phosphorylated by protein kinase A or C, depending on the method of cell stimulation. Thus, this enzyme may serve as an integrative factor by which different signaling pathways can regulate microfilament organization. These studies indicate that localization of these enzymes may be a critical factor in determining how signal transduction events regulate organization of the cytoskeleton. One interesting possibility is that one or more of these lipid-metabolizing enzymes (PIK, DGK, and PLCg^^^^ J are part of a larger complex which can shuttle between stress fibers and the EGF receptor at the membrane-microfilament interface. Although the role of the 1,4,5-IP3 is becoming clearer, the 3-substituted phosphoinositides are much less well understood, though equally intriguing. Growth factor stimulation and transformation increase the levels of PI(3,4)P2 and PI(3,4,5)P3, which are products of PI3K, but which are not hydrolyzed by characterized phospholipase C isoforms (Panayotou and Waterfield, 1992; Soltoff et al., 1992). PI3K is a heterodimeric enzyme, composed of a 110 kDa catalytic subunit and a 85 kDa regulatory subunit. The latter has one SH3 and two SH2 domains. Autophosphorylation of PDGF receptors results in binding of PI3K to the receptor via SH2-tyrosine phosphate interactions (Cantley et al., 1991). Activation of PI3K has been implicated in mitogenesis in some cells; in other cells it is involved in differentiation (Soltoff et al., 1992). The role of PI3K in these events has not been defined, but several observations provide interesting possibilities for involvement in both microfilament- and microtubule-mediated cellular processes. First, stimulation of PI3K occurs with activation of platelets and neutrophils, which involves heterotrimeric G-proteins rather than growth factor receptors. Furthermore, a fraction of the PI3K in platelets becomes associated with the actin cytoskeleton upon thrombin activation (Zhang et al., 1992). Further fractionation showed association of the PI3K with the platelet membrane skeleton along with vinculin, talin, protein kinase C and pp60^'"'^. Second, cloning and sequencing of the catalytic subunit showed a strong similarity to VPS34, a yeast protein implicated in targeting to vacuoles (Panayotou and Waterfield, 1992). Moreover, activated PI3K in insulin-treated rat adipocytes is found primarily in a low density microsomal fraction which includes endocytic vesicles. Stimulation of 3T3 cells with PDGF results in rapid endocytosis of PDGF receptors and PI3K into clathrin-coated vesicles and movement to the perinuclear region of the cell (Kapeller et al., 1993). This movement was inhibited by the microtubule-depolymerizing drug nocodazole, which also caused redistribution from the perinuclear region to the cell periphery. Optical sectioning of 3T3 cells indicated partial co-alignment of the PI3K 85 kDa subunit with microtubules. These results suggest that a complex of PI3K and PDGF receptor transits to the perinuclear region along microtubules. One mechanism facilitating this transport might be through the association of its the p85 SH3 domain with dynamin (Booker et al., 1993), a microtubule-binding mechanochemical protein related to G-proteins. Dynamin has been shown to bind the PI3K

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but not the Src or PLC ^^^ SH3 domain recombinant peptide. Whether PI3K affects microtubule-based transport is unknown, as is the role of the inositide-phosphorylating activity of PI3K. An obvious area of interest for future study of the mechanism of transit will be the effect of PI3K and its products on the role of dynamin in microtubule-based vesicle movements. Additional insights into the EGF-induced changes in cell morphology and the cytoskeleton have been provided by experiments with small G proteins (Chrzanowska-Wodnicka and Burridge, 1992; Hall, 1993). Microinjection of the Ras analog Racl into rat fibroblasts increased membrane ruffling and the accumulation of actin in the ruffles (Ridley et al., 1992). The ruffling activity is independent of a slower formation of stress fibers induced by Racl, suggesting that there are at least two signaling pathways which regulate actin organization, one regulating cortical actin and another regulating actin bundling. Analogously to addition of growth factors, oncogenic Ras can also induce ruffling activity (Bar-Sagi and Feramisco, 1986). Significantly, injection of a dominant inhibitory form of Racl into Ras-activated cells will inhibit ruffling (Ridley et al., 1992). These results indicate that Racl acts downstream of Ras and begin to provide the framework for a pathway which controls one aspect of actin organization. Ras modulation is complex, however, and these studies also raise the question of how the various pathways in which Ras is involved are organized and integrated. B. p1 SS^*"^^^^"^^ Association with Microfilaments and a Signal Transduction Particle

The neu oncogene (rat) encodes an analog of the EOF receptor and was first identified by transfection from chemically-induced neuroblastomas (Padhy et al., 1982). Its human counterpart c-erbB-2/HER2 was isolated using as a probe the w-erbB gene of avian erythroblastosis virus (Coussens et al., 1985). The structure of the c-erbB-2/neu protein pi 85®'"^^^^"®" shows significant sequence similarities to the EGF receptor. The 185 kDa protein has 40% amino acid identities in the extracellular domain, 82% in the tyrosine kinase domain and similar autophosphorylation sites in the C-terminal region (Gullick, 1990). The most significant difference between the EGF receptor and pi 85^'^^^^"^'' is that EGF does not bind to p 185^'^^^^""". Although activators for p 185'^^^2/"'" have been identified, the mechanism of activation may be more complex than that for the EGF receptor (Peles et al., 1993; Carraway and Cantley, 1994). Moreover, since pi85^'^^^^"^" is related to the EGF receptor, the question arises whether it is associated with microfilaments and plays a role in regulating the cytoskeleton. Microvilli isolated from 13762 rat mammary adenocarcinoma cells provide a highly purified, minimally perturbed, plasma membrane preparation for investigating membrane-microfilament interactions and their regulation (Carraway and Carraway, 1989). Immunoblot analyses of these microvilli indicate that they contain pl85®'^^^^"^'', but not the EGF receptor. Furthermore, the 13762 microvilli

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contain a major integral membrane protein with EGF-like sequences, called ASGP2 (Sheng et al. 1992), which can bind and activate pi 85^'^^^^''^''. Immunoprecipitation analyses of detergent-solubilized fractions of microvilli and membranes prepared from microvilli indicated that ASGP-2 and plSS^'^^^^""^" are present in a complex in the membranes (K. Carraway et al., 1993). Further, the ligand and receptor re-associate after treatment under dissociating conditions. Based on these results, we have proposed that ASGP-2 is an integral membrane activator for plg^erbB2/neu ^j^j^jj maintains the receptor in a constitutively activated state in the 13762 cells. Fractionation of microvilli by detergent extraction and differential centrifiigation showed that much of the p 185^^"^^^^"^^ and a fraction of the membrane ASGP-2 are associated with the microvillar microfilament core. These results were verified by density gradient phalloidin shift analysis (C. Carraway et al., 1993). In previous studies we have shown that the 13762 microvillar microfilaments are linked to the membrane via a high molecular weight glycoprotein complex (Carraway et al., 1991). This complex can be isolated from microvilli in two forms. Under relatively non-dissociating conditions, the complex remains associated with actin and other cytoskeletal proteins, such as ezrin, spectrin and a-actinin. This larger complex is called the transmembrane complex (TMC; Carraway et al, 1983), because it contains both an integral membrane glycoprotein core complex and cytoplasmic components. Under more stringent dissociating conditions used for isolation of the glycoprotein complex, these cytoskeletal components are lost. However, pi85erbB2/neu remains associated with the complex in both instances, indicating that it is strongly associated with the glycoprotein core components (C. Carraway et al, 1993). How this complex is associated with actin is unclear. Since p jg^erbB2/neu contains a scqucncc similar to that found in the EGF receptor to bind actin (den Hartigh et al, 1992), it may be bound to the microfilaments directly. Alternatively, pi85®^^^^^°^" may be associated with the microfilaments via a component of the glycoprotein complex which binds actin (Carraway et al., 1991). Since redundant interactions are found at other membrane-cytoskeleton interaction sites, both of the mechanisms may be operative. Previous studies have shown that activated growth factor receptors associate via SH2/SH3 adaptor proteins with Ras, which can further associate with Raf of the MAP kinase transduction cascade. These results suggest that a constitutively activated receptor, such as pi 85erbB2/neu ^f^^ microvilli, may be present in a stable complex with other components of this mitogenic pathway. Moreover, the association of pi85®^^^^^"®^ with such a large complex presents the possibility that this complex could be a site for integration of multiple signal transduction components. To test this hypothesis, we have analyzed microvilli, microvillar microfilament cores, TMC fractions and Triton-soluble fractions by immunoblotting with antibodies against a panel of signal transduction components. All of the components of the previously described mitogenic pathway tested were found associated with each of the particulate fractions. These include Ras, GRB2, Raf, MAP kinase kinase and MAP kinase. In addition a number of other signal transduction components

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were found to be associated with these microvillar fractions, including Src, protein kinase C, PI3K, GAP, Crk, and PLCg^^^^^. These observations suggest that the TMC is a large signal transduction particle which provides a site for interaction of different pathways, possibly permitting crosstalk and integration of the signals, as well as linking pi85erbB2/neu signaling to the microfilaments.

V. CONCLUSIONS AND PROSPECTS Intensive investigation in recent years has yielded significant progress in providing a basic understanding of the fundamentals of transmembrane signaling. This initial phase of research in signal transduction has described many of the components involved and has developed a few fundamental concepts. Foremost among these concepts is the importance of spatial and temporal localization of components m this complex process. A fundamental principle which has emerged as one of the most important themes is that signaling components are organized into multi-component signal transduction complexes. The molecular interactions involved in this organization are thus of critical importance. Advances in understanding molecular mechanisms involved in protein-protein interactions, such as the specificity of interactions of SH2 and SH3 domains, contribute significantly to our understanding of how some of these components may associate and provide a powerful tool in our attempts to construct pathways. An important aspect of the cellular organization of these pathways is their association with the cytoskeleton. The molecular interactions involved in this association have been analyzed in only a few cases. These molecular mechanisms will undoubtedly be a crucial aspect of the cellular remodeling events which occur in response to many external signals. In this review we have stressed the role of signal transduction complexes at membrane-microfilament interfaces, in part because less is known about the involvement of other cytoskeletal systems and elements in signal transduction and in part because this interface is a likely key site for integration of multiple signals. Other associations of signal transduction components with cytoskeletal elements or membrane skeletal proteins have also been described (Table 2). Whether these are associated with multimeric complexes remains to be seen. Contributions by SH3 and some actin-binding domains have been described, but other mechanisms remain to be delineated. This list will undoubtedly grow as the interest in the role of the cytoskeleton in signaling increases. The cortical localization of signal transduction components afforded by these supramolecular organizations of proteins after a signaling event provides two important advantages: (1) the complexes are present at the plasma membrane, the origin of most signals, and (2) the localization to the cytoskeleton facilitates integration of changes in the cytoskeleton with changes in other cellular responses. A key element of most recent models of signal transduction is the recruitment of cytoplasmic components to signal transduction complexes. One obvious purpose

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Table 2. Some Receptors and Signal Transduction Components Associated with Other Cytoskeletal Elements Receptor or Signal Transduction Component

Cytoskeletal Element

IP3 receptor

Ankyrin

Na"^-Ca exchanger Na"*"-K'^ATPase Glutamate receptor Tyrosine phosphatases

Ankyrin Ankyrin Fodrin ? Ezrin-Hke domains

Ig Fc receptor Chemoattractant receptor cAMP kinase G-proteins

Actin Binding Protein 7

A kinase anchor protein TubuHn

Reference Joseph & Samanta, 1993 Bourguignon et al., 1993 L i e t a l , 1993 Nelson & Veshnock, 1987 Simanetal, 1985 G u e t a l , 1991; Yang&Tonks, 1991 Ohtaetal, 1991 Jesaitis et al, 1986 Glantzetal., 1993 Roychowdhury et al., 1993

of this relocalization is the formation of an enzyme-substrate complex. Another equally important consequence of complex formation is localization or sequestration of a regulatory component. For example, recruitment of a phosphatase to a site on the cytoskeleton could sequester it from action on signal transduction components. It could then be released at a particular phase of the signal transduction pathway, for example, by phosphorylation, to act on components of a membrane signal transduction complex. This mechanism would restrict the functional activity of the regulatory process in both time and space and may actually operate in the control of CD45, which has been shown to bind the cytoskeletal protein ankyrin (Lokeshwar and Bourguignon, 1992). However, complex formation does not always result in phosphorylation of the recruited protein; localization may occur in the absence of phosphorylation. By such mechanisms localization functions can exert either positive or negative regulation. Another intriguing question in signal transduction is how apparently identical basic pathways in different cell types can lead to essentially opposite cellular responses. For example, activation of the MAP kinase pathway in fibroblasts leads to a mitogenic response, with an extensive breakdown and reorganization of cytoskeletal structure during cell division, but in PC 12 neuronal cells this activation results in cell differentiation (Kremer et al., 1991; Leevers and Marshall, 1992), involving polymerization and a very different reorganization of cytoskeletal components. One potential answer to this dilemma is the involvement of different isoforms of the MAP kinases in cell type-dependent responses. Another is that the enzymes in the basic signal transduction pathway are modulated in cell type-specific manners through different receptors and ligands, as well as by other signal transduction components such as the lipid-converting enzymes and protein kinase C. Other kinases such as Src, whose specific ftmctions are largely uncharacterized

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for any cell type, may well play cell type-dependent roles in organization and modulation of signal transduction pathways. It will be of particular interest to determine how specific components of the MAP kinase cascade, such as Ras, are modulated in different cells in response to external cellular signals acting through different receptors. Another potentially important aspect of cellular response is crosstalk between pathways linked to different receptors. Elucidating mechanisms of interactions between pathways will be crucial to these studies. Finally, superimposed upon these variables is the manner of linkage of signal transduction pathways with elements of the cytoskeleton. We suggest that cell type-specific proteins involved in these linkages must be a key element in determination of cell-specific responses to ligands acting through similar receptors. How cytoskeletal changes are coupled to the signaling event in different cell types through these specific linkage proteins is a subject of intense interest. Since these events usually involve tyrosine phosphorylation, considerable effort has been expended in identifying tyrosine-phosphorylated cytoskeletal components. Included among these are vinculin, talin, ezrin, paxillin, erythrocyte band 4.1, integrin pi subunit, cortactin, and calpactins (Glenney, 1992; Wu and Parsons, 1993). In spite of this effort we still have only a rudimentary understanding of the role of phosphorylation in cj^oskeletal changes. One impediment to our understanding is that phosphorylation of many proteins appears to be irrelevant to their functions. A second problem arises from the possibility that a significant fraction of the molecules of a specific protein need not be modified to elicit a cellular response. As an illustration, only a small percentage of cellular receptors are required to bind ligand for cell activation. Little is known about the level of phosphorylation of particular proteins required for cellular changes. Any attempts to implicate phosphorylation of specific proteins with cellular alterations will have to take this stoichiometry into consideration. One of the most poorly understood areas of signal transduction and the cytoskeleton involves the extensive reorganization of the entire cytoskeleton on cell division. The far-downstream events responsible for the global alterations in skeletal structure and the concomitant events responsible for initiating reassembly are obviously complex, since all of the organized cytoskeletal components are involved. This complexity and the likelihood of involvement of numerous interacting pathways will likely make this area the final frontier of research in the role of the cytoskeleton in the transduction of signal from the cell surface to the nucleus. A better understanding of the interactions which drive the mechanisms of those pathways and, particularly, of the organizations which determine the integration of individual pathways, are the logical next steps. Further research on interactions of these pathways with the cytoskeleton will ultimately play a critical role in the delineation of the cellular remodeling events which occur in response to many external signals.

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ACKNOWLEDGMENTS We thank Drs. Dafna Bar-Sagi, Johannes Boonstra, Lew Cantley, Lan Bo Chen, Thomas H. Fischer, Michinari Hamaguchi, Tony Hunter, Paul Janmey, Noboru Kuzumaki, Matthew Mescher, VerenaNiggli, J. Thomas Parsons, Jean Wang, and Gilbert C. White II for providing preprints or reprints. Dr. Paul van Bergen en Henegouwen for providing unpublished results and Dr. Richard Assoian for reading the manuscript. Original work was supported by NIH grant GM33795.

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FUNCTION OF MICROTUBULES IN PROTEIN SECRETION AND ORGANIZATION OF THE GOLGI COMPLEX

Jaakko Saraste and Johan Thyberg

I. II. III. IV.

Introduction 240 Membrane Compartments of the Secretory Pathway 240 The Cytoplasmic Microtubule System 243 Role ofMicrotubules in the Secretory Process 244 A. Transport from ER to Golgi 245 B. Membrane Recycling at the ER-Golgi Interface 248 C. Transport from Golgi to the Plasma Membrane 249 D. Transport to Lysosomes 252 E. In H/ro Studies of Protein Transport 252 V. Microtubule-Based Movements of Secretory Organelles 253 VI. Role ofMicrotubules in the Organization of the Golgi Complex 254 A. Spatial Relationship between Microtubules and the Golgi Complex . . . 255 B. Reorganization of the Golgi Complex in Cells Treated with Microtubule-Disruptive Drugs 256

The Cytoskeleton, Volume 2 Role in Cell Physiology, pages 23^273 Copyright © 1996 by JAI Press Inc. AH rights of reproduction in any form reserved. ISBN: 1-55938-688-6 239

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C. In Vitro Studies on the Interaction between Microtubules and the Golgi Complex VII. Reorganizationof the Golgi Complex During Cell Division A. Fragmentationof the Golgi Complex in Prophase-Metaphase B. Reformation of the Golgi Complex in Telophase-Cytokinesis VIII. Summary and Perspectives Acknowledgments References

260 261 261 262 265 265 265

I. INTRODUCTION The first descriptions of the subcellular organization of the secretory pathway already made it obvious that the secretory products, synthesized in the rough endoplasmic reticulum (ER), must be transported across considerable intracellular distances to reach the outside of the cell (Palade, 1975). Subsequent studies have further revealed that this transport process is highly efficient and many soluble and membrane-bound proteins reach the cell surface at a remarkably rapid rate. These findings clearly demonstrate that the cells possess intricate machineries that both accomplish the efficient sorting of their secretory products and ensure the rapid and targeted movement of carrier elements between secretory organelles located in spatially separated regions of the cytoplasm. The early electron microscopic studies identified the membranous organelles of the secretory pathway and also demonstrated their frequent association with the newly discovered cytoskeletal structures, microtubules (MTs) and microfilaments. With the introduction of drugs that cause the selective breakdown of MTs, a number of studies focused on their possible role in the secretory process (Dustin, 1984). However, although these drugs induce morphological changes of the secretory organelles, in particular the Golgi complex, their effects on the transport processes have been highly variable and conflicting results have been obtained even with the same cell types. Within the last decade, the application of video microscopy to analyze membrane movements occurring in living cells and in vitro systems, has yielded a wealth of new information which has reemphasized the role of MT-based transport in intracellular membrane traffic pathways (Schroer and Sheetz, 1991). In the present review we discuss the function of MTs in the different transport steps of the secretory pathway and summarize the evidence on their role in the organization of the Golgi complex, the organelle that represents the major traffic and sorting center of this route.

II. MEMBRANE COMPARTMENTS OF THE SECRETORY PATHWAY The general picture of the compartmental organization of the secretory pathway has changed considerably during recent years. For example, the organization of

Function of

Microtubules

241

Figure 1. Diagram of the membrane compartments of the secretory pathway and different steps of protein transport between these compartments. Abbreviations: RER, rough endoplasmic reticulum; GC, Golgi complex; PM, plasma membrane; L, lysosome; N, nucleus; CGN and TGN, cis- and trans-Golgi network; PG, pre-Golgi intermediate; SG, secretory granule; SV, constitutive post-Golgi vesicle; MT, microtubule. In this model cell, MTs radiate from the organizing center (minus ends) towards the cell periphery (plus ends).The numbers (1-8) refer to transport steps or organelle movements In which MTs have been suggested to participate (1) Formation of the ER network by tubule growth. (2) Function of the intermediate compartment (PG) in protein transport from RER to Golgi. (3) Membrane recycling at the ER-Golgi interface. (4) Juxtanuclear positioning of the Golgi complex. (5) Formation of tubular connections between different Golgi stacks. (6) Movement of SGs to the PM in regulated secretory cells. (7) Directed transport of SVs to different domains of the plasma membrane in polarized cells. (8) Juxtanuclear positioning of lysosomes.

subcompartments have evolved (Klausner, 1989; Farquhar, 1991; Mellman and Simons, 1992; Saraste and Kuismanen, 1992). The appreciation of the interaction of secretory organelles with different cytoskeletal systems, including MTs, has contributed to this development. A schematic view of the secretory pathway is

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illustrated in Figure 1. This scheme also depicts the transport steps in which MTs are presently thought to participate, as discussed in detail below. After their synthesis and insertion into the membrane or lumen of the rough ER, the transported proteins exit this compartment in budding vesicles or tubules. Recent results suggest that this occurs at widespread locations within the rough ER network. However, it is not clear whether vesicle budding takes place throughout the rough ER or is limited to transitional regions representing a ftinctionally specialized ER subcompartment. It has been suggested that protein transport from the rough ER to the Golgi is a bulk flow process (Pfeffer and Rothman, 1987). However, recent observations have shown that the concentration of some membrane and secretory proteins can occur already in the ER or in the intermediate compartment (Mizuno and Singer, 1993; Balch et al., 1994), indicating that other mechanisms than simple bulk flow are involved in the early transport steps. After leaving the rough ER, the proteins reach the intermediate compartment which functions as a way station in forward transport (Saraste and Kuismanen, 1984; Schweizer et al., 1990). This compartment has also been suggested to fiilfill important fimctions in membrane recycling and retrieval of resident ER components and was therefore termed the salvage compartment (Pelham, 1989). Partly different views on its organization and function in transport at the ER-Golgi interface have been presented. First, it could represent a smooth-surfaced subcompartment which extends the rough ER toward the Golgi complex (Hauri and Schweizer, 1992). In this case the rough ER and the Golgi would be separated by a single membrane discontinuity. Second, it has been proposed that the intermediate compartment is a distinct post-ER structure which communicates with the rough ER and the Golgi via two different vesicular pathways (Pelham, 1989; Lotti et al., 1992). Third, it has been suggested that the intermediate compartment is made up of mobile, endosome-like structures which themselves transport the proteins towards the Golgi complex (Saraste and Svensson, 1991). In any case, there is increasing evidence that these structures interact with MTs, which thus could influence protein transport and recycling at the ER-Golgi interface (Figure 1). Although it has been known for a long time that the Golgi stack is a compositionally and ftinctionally polarized structure, its exact compartmental organization is still in debate. It has also become clear that the Golgi complex, in spite of its ordered morphology, is a highly dynamic organelle, and the localization of specific marker enzymes within the stack varies considerably between different cell types (Roth et al., 1986; Velasco et al., 1993). Moreover, the view that the stacks are divided into multiple, compositionally distinct subcompartments is challenged by findings indicating that the enzymes which ftinction in the terminal glycosylation of N-linked oligosaccharides display overlapping distributions (Nilsson et al., 1993). The most recent models of the Golgi complex have given increasing attention to subcompartments located at its entry and exit faces, often referred to as cis- and trans-Golgi networks (CGN and TGN). Accordingly, the Golgi complex could be

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composed of three relatively stable subcompartments between which transport occurs by means of vesicular or tubular carriers (Mellman and Simons, 1992). Alternatively, the cistemae may be transient, maturing structures which themselves move in cis to trans direction and transport proteins through the stacks (Saraste and Kuismanen, 1992). During the last few years, an important role of cytoplasmic coat proteins in maintaining the structural organization of the Golgi complex has been recognized (Duden et al., 1991a; Klausner et al., 1992). If the assembly of these proteins on the surface of the membranes is prevented, for example by exposing cells to the drug brefeldin A (BFA), the Golgi stacks disintegrate and their constituents redistribute either to the ER (cis-, middle-, and trans-Golgi markers) or to peripheral endosomes (TGN markers). In vitro studies have pointed out that the coat proteins belong to the molecular machinery that operates in vesicle budding during intraGolgi transport (Rothman and Orci, 1992). However, these proteins also associate with the membranes of the intermediate compartment and may have a more general role in transport of proteins along the early part of the secretory pathway (Duden et al., 1991a, 1991b; Klausner et al, 1992; Saraste and Kuismanen, 1992). The sorting of proteins within the Golgi complex to different destinations has been suggested to occur in the trans-Golgi network (TGN), a reticular structure located at the trans-side of the stack (Griffiths and Simons, 1986; Geuze and Morre, 1991). If the stack is instead viewed as a maturing structure (Saraste and Kuismanen, 1992), this sorting process would be expected to begin in more proximal parts of the Golgi. From the TGN soluble lysosomal enzymes (bound to mannose-6-phosphate receptors) are transferred to lysosomes via a prelysosomal (late endosomal) pathway, involving clathrin-coated buds and vesicles (Komfeld and Mellman, 1989). In regulated secretory cells a further sorting process in the TGN involves the segregation of proteins to the constitutive and regulated pathways for delivery to different parts of the cell surface (Burgess and Kelly, 1987; Farquhar, 1991).

III. THE CYTOPLASMIC MICROTUBULE SYSTEM MTs represent a major component in the cytoskeleton of eukaryotic cells and fulfill important functions in the regulation of cell structure and function (Dustin, 1984). They are made up of protein subunits called tubulin (heterodimers of a- and p-tubulin) as well as smaller amounts of several other proteins (microtubule-associated proteins or MAPs). Polymerization of tubulin normally gives rise to tubules with an outer diameter of 25 nm and a 5 nm thick wall in which 13 protofilaments can be distinguished. The subunits are all oriented in the same direction within the protofilaments and this constitutes the basis for the polarity of MTs with a plus (fast-growing) end and a minus (slow-growing) end. The assembly and spatial distribution of MTs in the cell are regulated by the centrosome (typically a pair of centrioles surrounded by satellites of fibrogranular material), the main microtubuleorganizing center (MTOC) both during interphase and mitosis (Karsenti and Maro,

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1986). When new MTs are formed, the minus end is linked to the centrosome and the plus end grows towards the periphery. Recently, y-tubulin was identified as a new and highly conserved member of the tubulin family and found to be associated with the centrosome (Steams et al., 1991; Zheng et al., 1991). It is believed that this protein takes part in the nucleation of MTs and the establishment of MT polarity (Oakley, 1992; Joshi, 1993; Kalt and Schliwa, 1993). MTs vary in stability with a large, rapidly turning over population and a small, slowly turning over population. Moreover, growing and shrinking phases of MTs occur simultaneously (a phenomenon referred to as dynamic instability) and the shift between them has been related to hydrolysis of GTP reversibly bound to tubulin (Kirschner and Mitchison, 1986; Gelfand and Bershadsky, 1991). There also exist MT populations in which the tubulin subunits have been post-translationally modified (Greer and Rosenbaum, 1989; Bulinski and Gundersen, 1991). Thus, detyrosinated a-tubulin is concentrated in a limited subset of interphase MTs and a similar situation exists with regard to acetylated a-tubulin. MTs enriched in these isoforms of tubulin have a longer half-life and are more resistant to drug-induced depolymerization than the majority of cytoplasmic MTs and actually make up the slowly turning over population of MTs mentioned above. However, detyrosination or acetylation of tubulin is not sufficient to make MTs stable and it is likely that these modifications are accompanied by or occur as a result of other as yet unknown modifications. Drugs which interfere with the polymerization of tubulin have played a fundamental role in MT research (Dustin, 1984). As a matter of fact, tubulin was originally identified as the protein subunit of MTs on the basis of its ability to bind strongly and almost irreversibly to the plant alkaloid colchicine (Wilson et al., 1974). If cells are exposed to colchicine, assembly of new MTs is inhibited and preexisting MTs break down into free tubulin subunits. A similar effect is obtained with the synthetic compound nocodazole (De Brabander et al., 1976). However, nocodazole binds reversibly to tubulin (Hoebeke et al., 1976) and a rapid recovery follows upon its removal (De Brabander et al., 1981). The vinca alkaloids vinblastine and vincristine likewise destroy preexisting MTs, but in the presence of these drugs tubulin is precipitated into paracrystals rather than adding to the free pool of subunits (Wilson et al., 1974). In contrast to the substances referred to above, taxol is a plant derivative that promotes MT assembly (Horwitz et al., 1982). Nevertheless, the biological effects of taxol resemble those of the MT-disruptive agents, indicating that the dynamic behavior of MTs is essential for their functions in the cell.

IV. ROLE OF MICROTUBULES IN THE SECRETORY PROCESS Typically, the depolymerization of MTs with various drugs results at most in a partial inhibition of protein secretion (Dustin, 1984; Thyberg and Moskalewski,

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1985; Burgess and Kelly, 1987). This has led to the view that MTs facilitate the movement of transport carriers between the different compartments of the secretory pathway, but are not strictly required in this process. However, the use of drugs represents an indirect way of looking at the role of MTs in protein transport and is subject to limitations. It is presently known that different types of MTs in a single cell have different stabilities and can be differentially affected by MT-depolymerizing agents (Greer and Rosenbaum, 1989; Bulinski and Gundersen, 1991). Moreover, the drug-induced disassembly of MTs is a slow process and complete disassembly requires long incubation times. A faster and more extensive disruption of MTs can be obtained by combining the drug treatment with incubation of cells at low temperature (Turner and Tartakoff, 1989; Bomsel et al., 1990; Parton et al., 1991; Saraste and Svensson, 1991). Since MTs have a multitude of functions in cellular physiology, it is also possible that the observed effects are secondary and result from the inhibition of other processes caused by MT depolymerization. Finally, and specifically regarding the secretory pathway, the spatial organization of the responsible organelles is dramatically changed in cells exposed to MT-active agents (see below), further complicating the interpretation of the results. In the following, we summarize the evidence which supports a role of MTs in different transport events along the secretory pathway (Figure 1). In addition to the use of drugs, much of the data derives from more direct approaches, including localization studies with recently identified marker proteins, in vitro studies of the interaction of specific membranes with MTs, and experiments with living cells. A. Transport from ER to Golgl

With some exceptions (Busson-Mabillot et al., 1982; Pavelka and EUinger, 1983), studies with MT inhibitors have in general not indicated a role of MTs in transport between the rough ER and the Golgi complex. Since the drugs do not interfere with the terminal glycosylation of newly synthesized proteins, the latter are evidently transferred to the compartments where the modifying Golgi enzymes reside also in the absence of intact MTs (Stults et al., 1989). Moreover, morphological studies of actively secreting cells have suggested a close spatial relationship between transitional elements of the ER and the Golgi stacks (Palade, 1975). Together, these results have supported the view that the transfer of material from the rough ER to the entry face of the Golgi complex is a short-range transport step in which MTs are not required (Kelly, 1990). The identification of an intermediate compartment between the rough ER and the Golgi complex (Hauri and Schweizer, 1992; Saraste and Kuismanen, 1992; Bonatti and Torrisi, 1993), and the observed effects of brefeldin A (BFA) on membrane dynamics at the ER-Golgi interface (Klausner et al., 1992), have demonstrated the unexpectedly complicated organization of the early part of the secretory pathway. These studies have also revealed that the membranes of the

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intermediate compartment interact with the MT system and suggest that MTs play a role both in forward transport and membrane recycling at the ER-Golgi interface. The localization of recently identified marker proteins has shown that, in contrast to the Golgi complex, the elements of the intermediate compartment are widely distributed in cells. Under normal conditions they are clustered in the Golgi region, but are also present in the peripheral cytoplasm (Figure 2). This indicates that the exit of proteins takes place throughout the rough ER and is not restricted to sites close to the Golgi complex. Accordingly, ER to Golgi transport could not occur by simple diffusion of vesicles, but must be a directed process, involving MTs and possibly also other cytoskeletal structures (Saraste and Svensson, 1991). This conclusion is also supported by studies of secretory protein transport between heterologous organelles in fused cells (Valtersson et al., 1990). Moreover, it is evident that this transport step involves movement across long intracellular distances and at least two ways in which this may be accomplished can be envisaged (Lippincott-Schwartz, 1993). If the intermediate compartment consists of stable structures with a fixed position in the cytoplasm, it could be connected to the rough ER and the cis-Golgi by pathways involving vesicular or tubular carriers. Alternatively, mobile elements of the intermediate compartment could themselves act in ER to Golgi transport (Saraste and Kuismanen, 1992; see Figure 1). Protein transport between the rough ER and the Golgi complex is inhibited if the temperature is reduced to 15—16 °C, resulting in the arrest of proteins in the intermediate compartment (Saraste and Kuismanen, 1984). In parallel, incubation at low temperature results in accumulation of these pleiomorphic pre-Golgi intermediates throughout the cytoplasm, as determined by the localization of the 58 kD marker protein, p58 (Saraste and Svensson, 1991). The transport block is readily reversible and when cells are briefly warmed to 37 °C, protein transport is resumed and a rapid relocation of the pre-Golgi elements to the central Golgi region is observed. Depolymerization of MTs effectively inhibits this reclustering, suggesting that ER to Golgi transport depends on the integrity of MTs (Saraste and Svensson, 1991). The low temperature-induced transport arrest is not absolute and during longer incubation at 15 °C newly synthesized proteins are transported towards the Golgi complex, albeit slowly (Saraste and Kuismanen, 1984; Bonatti et al., 1989; Kuismanen and Saraste, 1989). Interestingly, recent experiments have demonstrated that elements of the intermediate compartment codistribute with cold-resistant MTs, suggesting that ER to Golgi transport involves a stable subset of cytoplasmic MTs (J. Saraste, unpublished results). The Golgi apparatus is associated with MTs enriched in detyrosinated tubulin (Skoufias et al., 1990; see further below). Recently, Mizuno and Singer (1994) obtained evidence suggesting that MTs belonging to this stable subset may participate in the transfer of proteins between the rough ER and the Golgi complex. These authors used a cycloheximide wash-out protocol to synchronize the export of albumin from the rough ER in human hepatoma (HepG2) cells, double immu-

1

^

1

•*

Figure 2, Double immunofluorescence localization of the intermediate compartment and the Golgi complex in control and nocodazole-treated BHK-21 cells. The cells were stained with antibodies against the intermediate compartment/cls-Golgi marker p58 (A and C) and the Golgi stack marker mannosldase II (B and D). In the control cells the p58-posltive elements are found both In the Golgi region and in the cell periphery (A), whereas the Golgi complex displays a typical juxtanuclear location (B). Following disassembly of MTs with nocodazole, p58 (C) and mannosldase II (D) colocalize in punctate structures scattered throughout the cytoplasm. Bars mark 10 jim. 247

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noelectron microscopy to localize albumin and detyrosinated tubulin in EM cryosections, and obtained evidence for the association of both of these proteins in vesicular structures located in the Golgi region. In conclusion, specific subclasses of MTs may be involved in the early part of the secretory pathway. If these MTs are in part resistant to depolymerizing drugs, this could explain why protein transport between the rough ER and the Golgi continues in the drug-treated cells. On the other hand, it has been shown that the elements of the Golgi complex maintain a close association with the intermediate compartment during and after their redistribution in cells exposed to nocodazole (Saraste and Svensson, 1991; see Figure 2). Although normally occurring as a MT-mediated, long distance transport step between peripheral ER sites and the central Golgi complex, ER to Golgi transport in the drug-treated cells could thus continue as a short-range transport between the rough ER and the dispersed Golgi elements. It is not known whether cytoskeletal structures other than MTs are required in this process. B. Membrane Recycling at the ER-Golgi Interface

As proteins exit the ER, large amounts of lipid also leave this compartment in the limiting membrane of the vesicular carriers which are transported towards the Golgi complex and must somehow be replenished to maintain the functional balance of the ER membranes. Recent evidence suggests that this is accomplished by a process which continuously recycles not only lipids (Hoffmann and Pagano, 1993) but also proteins from post-ER compartments (Pelham, 1991; LippincottSchwartz, 1993). The first evidence for the existence of this recycling pathway was derived from the study of resident ER proteins containing the carboxyterminal KDEL/HDEL-sequence. Such proteins can leave the ER, but subsequently bind to receptors which specifically recognize this tetrapeptide motif As a consequence of this, the receptor-bound proteins are retrieved back to the ER, providing a mechanism for retaining them in this compartment (Pelham, 1991). It is possible that the recycling of proteins to the ER occurs from multiple post-ER locations, including the intermediate (salvage) compartment (Pelham, 1989) and different parts of the Golgi complex (Hsu et al., 1991; Peter et al., 1992). The cytoplasmic tails of a number of type I ER membrane proteins contain a carboxyterminal peptide sequence (double-lysine motif) which functions in the retention of these proteins in the ER (Jackson et al., 1990). As in the case of KDEL-proteins, the retention involves retrieval from post-ER compartments (Jackson et al., 1993), but it is still unknown how this recycling takes place. Interestingly, short peptides containing the retention signal of one of these proteins—the adenovirus E3/19K protein—^have been shown to bind to taxol-polymerized MTs in vitro and to induce polymerization of tubulin (Dahllof et al., 1991). Thus, binding to cytoplasmic MTs could be one mechanism for retaining membrane proteins in the ER.

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Recently, it has been demonstrated that the cytoplasmic, carboxyterminal tail of the two homologous membrane proteins of the intermediate compartment, human p53 and rat p58, also contains a double-lysine motif (Schindler et al., 1993; Lahtinen et al, 1996). This finding suggests that membrane protein retention in the early part of the secretory pathway can be regulated by several different mechanisms. For example, the sequence context of the double-lysine motif could affect the affinity of the proteins to cytoplasmic effector molecules, resulting in the concentration of different proteins in different compartments along the pathway (Jackson et al., 1993). Since the function of MTs has been implicated in both anterograde and retrograde pathways at the ER-Golgi interface, the effect of nocodazole on the transport and recycling of different protein constructs containing the double-lysine motif was examined (Jackson et al., 1993). The authors concluded that although MTs may facilitate transport in both directions, neither pathway showed an absolute MT requirement. Studies on the effect of brefeldin A (BFA) on the secretory pathway have provided morphological evidence for a membrane recycling pathway at the ER-Golgi interface. As discussed above, BFA causes disassembly of the coat structures that normally associate with the membranes of the intermediate compartment and the Golgi complex. As a result, ER to Golgi transport is inhibited and Golgi membranes redistribute to peripheral locations (Klausner et al., 1992). Immunocytochemical detection of middle- and trans-Golgi marker proteins in BFA-treated cells has indicated that long tubular projections function as intermediates in this redistribution event, which leads to the mixing of the Golgi membranes with the ER-intermediate compartment system (Lippincott-Schwartz et al., 1990). Shortly after the addition of BFA, these projections can be seen to extend from the central Golgi complex towards the cell periphery, often close to and in parallel with MTs. In further support for a role of MTs in this process, nocodazole was found to inhibit the BFA-induced redistribution of Golgi antigens to the ER (Lippincott-Schwartz et al., 1990; Strous et al., 1991). Based on the above studies, it has been proposed that BFA treatment reveals a constitutive, MT-dependent recycling pathway operating at the ER-Golgi interface (Klausner et al., 1992; Lippincott-Schwartz, 1993). Tubulation of Golgi membranes has also been observed to occur in the absence of BFA (Cooper et al., 1990; Lippincott-Schwartz et al., 1990; Cluett et al., 1993), suggesting that the drug treatment amplifies an already existing pathway. In vitro studies with isolated Golgi membranes further indicate that MTs are not required for tubule formation itself. Therefore, they may simply act as tracks in the retrograde transport pathway in intact cells (Cluett et al, 1993). C. Transport from Golgi to the Plasma Membrane The general conclusion of a number of inhibitor studies has been that MT-active drugs exert their effect on protein secretion at a post-Golgi stage. The exocytosis

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of preformed secretory granules in regulated secretory cells is not inhibited by the depolymerization of MTs, whereas the extracellular discharge of newly synthesized products is impaired, suggesting that the movement of vesicles between the Golgi complex and the plasma membrane is inhibited in the drug-treated cells (BussonMabillot et al, 1982; Burgess and Kelly, 1987). The close spatial relationship between the Golgi complex and the MTOC is also suggestive of a role of MTs in transport of secretory vesicles to the cell surface. In many cell types, MTs radiate from the MTOC (minus end) toward the cell periphery (plus end) and, thus, are well suited to provide tracks for the movement of Golgi-derived vesicles toward different plasma membrane locations (Figure 1). Morphological studies of polarized secretory cells have revealed that the Golgi complex is oriented towards the apical pole of the cell, where the secretory granules fuse with the plasma membrane. Moreover, in several situations associated with changes in intracellular membrane traffic, a coordinated reorientation of the MTOC and the Golgi complex has been observed to take place (Singer and Kupfer, 1986). Endothelial cells (Gotlieb et al., 1981; Ettenson and Gotlieb, 1992) and fibroblasts migrating into an experimental wound in culture (Kupfer et al., 1982; Bergmann et al., 1983) are two examples of this phenomenon and macrophages subjected to a chemotactic gradient a third (Nemere et al, 1985). A similar shift in the localization of the MTOC and the Golgi complex occurs in natural killer cells (Carpen et al., 1982; Kupfer et al., 1983) and cytotoxic T lymphocytes (Geiger et al., 1982), directing their secretory activity towards the plasma membrane domain which interacts with the target cell. The binding and movement of lytic granules from cytotoxic T cells along MTs has also been reconstituted in vitro (Burkhardt et al., 1993). In the simultaneous presence of both plus and minus end-directed motors, the granules were preferentially translocated towards the plus end of the MTs. Inactivation of dynein (UV-induced cleavage) did not affect binding and movement of the granules, whereas removal of kinesin from the motor preparation (immunodepletion) inhibited binding by about half These results are in accordance with the orientation of MTs in intact cytotoxic T cells and suggest that the signal-induced transport of secretory granules from the Golgi complex to the plasma membrane utilizes MTs as tracks and is mediated by kinesin, possibly in cooperation with other factors (Burkhardt et al., 1993). In nerve cell axons the MTs form extensive arrays and are oriented with their minus ends facing the cell body (where the Golgi complex resides) and the plus ends pointing outwards. They support organelle movements and vesicle transport both in the anterograde and retrograde directions (Schroer and Sheetz, 1991). Some cell lines, such as pituitary tumor cells (AtT20) and pheochromocytoma cells (PC 12), maintain neuronal properties in culture and can be induced to extend long neurite-like processes (Tooze et al, 1989). In these cells Golgi-derived, dense-core granules of the regulated secretory pathway are found in the cell body, but also accumulate at the tips of the processes (Tooze and Burke, 1987). In dividing AtT20 cells, after the loss of interphase MTs, the secretory granules first become randomly

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distributed in the cytoplasm. During telophase they regather in the region of the forming intercellular bridge, which contains a bundle of MTs representing the remains of the mitotic spindle (Tooze and Burke, 1987). Together, these observations on interphase and mitotic cells suggest that the distribution of dense-core granules is determined by anterograde movements along MTs. Experiments with transfected cells have suggested that the distribution of constitutive post-Golgi vesicles involves a similar mechanism (Matsuuchi et al., 1988). The biogenesis of secretory granules and the mechanism of their transport to the cell surface has also been studied in the neuroendocrine PC 12 cells (Tooze et al., 1991). These workers identified immature secretory granules as an intermediate in the formation of TGN-derived secretory granules. Interestingly, both immature and mature granules could be induced to fuse with the plasma membrane, indicating that both are transport competent and can reach the periphery of the cells. Disruption of MTs by nocodazole efficiently reduced extemalization of the marker secretogranin II both from the immature and mature granules. These results suggest that immature granules are able to bind to and translocate along MTs. In polarized epithelial cells, constitutive transport pathways link the Golgi complex to compositionally different domains of the plasma membrane (Burgess and Kelly, 1987; Caplan and Matlin, 1989; Simons and Wandinger-Ness, 1990; see also the review by Drenckhahn in this volume). Studies on cultured epithelial cells, such as Madin-Darby canine kidney (MDCK) cells and human intestinal (Caco-2) cells, have indicated that MT-active drugs (colchicine, nocodazole and taxol) preferentially inhibit the transport of secretory and membrane proteins to the apical domain, whereas transport to the basolateral cell surface remains largely unaffected (Rindler et al., 1987; Achler et al., 1989; Eilers et al., 1989; Parczyk et al., 1989; Breitfeld et al., 1990; Matter et al., 1990; van Zeijl and Matlin, 1990; Gilbert et al., 1991; Ojakian and Schwimmer, 1992). The transport block appears to be kinetic in nature and leads to a variable degree of missorting of apical markers to the basolateral surface (Salas et al., 1986; Rindler et al., 1987; Parczyk et al., 1989; van Zeijl and Matlin, 1990). In Caco-2 cells and hepatocytes, apical proteins can first be targeted to the basolateral membrane and subsequently transported to the apical surface by transcytosis (Bartles and Hubbard, 1988; Nelson, 1992), a process which also has been found to depend on the integrity of MTs (Breitfeld et al., 1990; Hunziker et al., 1990). The above studies suggest that MTs do not function in the transport of newly synthesized proteins from the Golgi to the basolateral surface of polarized epithelial cells, which may represent a single default pathway (Simons and Wandinger-Ness, 1990). However, there is also evidence for the existence of multiple basolateral pathways (Nelson, 1992), possibly with different MT dependencies (Boll et al., 1991; De Almeida and Stow, 1991). Interestingly, when exocj^ic carrier elements containing apical and basolateral markers were isolated from perforated MDCK cells, both bound to taxol-polymerized brain MTs in vitro (van der Sluijs et al., 1990). It is, therefore, possible that in some epithelial cells both apical and

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basolateral post-Golgi pathways utilize different subclasses of MTs with different sensitivities to MT-depolymerizing agents. D. Transport to Lysosomes

Most lysosomal enzymes are soluble proteins which during their synthesis are inserted into the lumen of the rough ER and cotranslationally modified by the addition of N-linked, high-mannose carbohydrate side chains. The transport of these molecules to and across the Golgi complex is thought to involve the same pathway which is utilized by secretory and plasma membrane proteins. However, during the transport the lysosomal enzymes receive a mannose-6-phosphate recognition marker which, by binding to specific receptors, facilitates their sorting from secretory material and subsequent transfer from trans-Golgi to lysosomes via a prelysosomal (late endosomal) compartment (Figure 1). The two enzymes which are responsible for the synthesis of the phosphomannosyl residues are thought to be localized in the intermediate compartment/cis-Golgi region of the cell (Komfeld and Mellman, 1989). When MTs are depolymerized by nocodazole, the prelysosomal compartment and the lysosomes, like the Golgi elements, lose their juxtanuclear localization and become scattered throughout the cytoplasm. Pulse-chase analysis has shown that in the absence of MTs the transport of newly synthesized cathepsin D to lysosomes is efficiently inhibited (Scheel et al., 1990). However, as in the case of secretory and plasma membrane proteins, the transport from the rough ER to the Golgi complex did not appear to be affected by nocodazole treatment. Moreover, the synthesis of the mannose-6-phosphate marker occurred normally in the drugtreated cells, indicating that the basic sorting mechanism was not impaired. Therefore, the integrity of MTs is important for the transport of lysosomal hydrolases from the Golgi complex to the lysosomes. It is not clear whether MTs are part of the transport machinery as such or efficient transport in the trans-Golgi-prelysosome-lysosome pathway requires a close grouping of these structures in the juxtanuclear area, a process known to be affected by retrograde movements along MTs (Pastan and Willingham, 1981; Herman and Albertini, 1984; Matteoni and Kreis, 1987; Swanson et al, 1987; Heuser, 1989; Hollenbeck and Swanson, 1990). E. In Vitro Studies of Protein Transport

A major advance in the study of membrane traffic has been the development of in vitro assays which allow the reconstitution of certain steps of protein transport between different membrane compartments of the secretory and endocytic pathways (Rothman, 1992). However, although controlled fusion between distinct intracellular membranes can be achieved after cell disruption, the cell-free assays lack the functional cytoskeletal architecture which is necessary for or facilitates these transport events in intact cells. The preparation and use of semi-intact or perforated cells also requires conditions which lead to at least partial disassembly

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of the different filament structures. It has been noted that in vitro incubation of semi-intact CHO cells (Corthesy-Theulaz et al, 1992) and digitonin-permeabilized NRK cells (Plutner et al, 1992) causes a gradual depolymerization of MTs. Therefore, the in vitro conditions can induce organelle redistributions similar to those observed in intact cells treated with MT-depolymerizing drugs. These could affect the normal spatial relationship of the donor and acceptor compartments and lead to the utilization of abnormal pathways in the observed transport (usually recorded as an acceptor-specific modification of the marker protein). The MT-dependent interaction of membranes of a certain intracellular pathway has also been reconstituted in vitro. In polarized MDCK cells endocytosis occurs from both the apical and basolateral plasma membranes and the two pathways meet at the level of late endosomes in the perinuclear region. Using isolated apical and basolateral early endosomes, Bomsel et al. (1990) showed that the meeting of the two pathways in vitro required the presence of MTs and MT-based motors (dynein and kinesin). Stimulation of membrane fusion could be achieved by either polymerizing and stabilizing the endogenous MTs with taxol or adding exogenous taxol-polymerized MTs to the in vitro reaction. These results encourage the use of similar in vitro approaches in future studies of the function of MTs in transport along the secretory pathway.

V. MICROTUBULE-BASED MOVEMENTS OF SECRETORY ORGANELLES The introduction of specific markers for vital staining of the ER and the Golgi complex has made it possible to visualize these major organelles of the secretory pathway by fluorescence microscopy in living cells (Lipsky and Pagano, 1985; Terasaki et al., 1984). These studies have demonstrated that, as in the case of other cytoplasmic organelles (see the review by Stebbings in this volume), the organization and dynamics of the ER and the Golgi complex also depend on MT-membrane interactions and MT-based motility. The ER-specific cyanine dye DiOC^(3) has facilitated the observation of the ER as a continuous network of branching tubules in the periphery of cultured cells (Terasaki et al., 1984, 1986). Using drugs that depolymerize MTs, it was shown that the arrangement of this network is dependent on the integrity of the MTs, whereas other cytoskeletal elements do not seem to have a role. Studies of living cells further revealed unexpected dynamics of the ER system and suggested that the generation of ER organization depends on continuous membrane movements including tubule formation and fusion (Lee and Chen, 1988). Similar results were obtained in studies using in vitro systems, which also demonstrated that tubulation and network formation involve MT-based membrane motility (Dabora and Sheetz, 1988; Vale and Hotani, 1988). It appears that MTs provide the structural framework for the binding and movement of the ER membranes (Lee and Chen, 1993).

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As discussed further below, recording of Golgi membrane movements in living cells recovering from nocodazole treatment has suggested that retrograde transport along MTs could provide the mechanism for the positioning of the Golgi complex in the cell center (Ho et al., 1989; see Figure 1). Studies with permeabilized cells have further indicated that such a centralization process involves the motor protein dynein (Corthesy-Theulaz et al, 1992). In addition, tubulation of trans-Golgi membranes, labelled with NBD-ceramide in living cells, has been shown to require intact MTs (Cooper et al., 1990). The relationships of the above described movements of the ER and Golgi membranes to protein transport along the secretory pathway remain open. It has been suggested that the MT-mediated centralization of Golgi membranes and protein transport from the widespread ER network to the central Golgi complex could share similar mechanisms (Saraste and Svensson, 1991). Moreover, tubular transport has been put forward as an alternative to vesicular transport for movement of proteins across the Golgi stack (Klausner et al, 1992; Mellman and Simons, 1992). The use of acridine orange as a vital stain, combined with inmiunolocalization of a marker secretory protein, has demonstrated that MTs provide tracks for movement of secretory granules in AtT20 cells (Kreis et al., 1989). The granules in these cells have an acidic internal pH and, like the endocytic compartments, readily incorporate this dye. Video-enhanced fluorescence microscopy showed that the secretory granules are transported along cell processes by saltatory movements, predominantly in the anterograde direction. Interestingly, although nocodazoleresistant MTs, containing detyrosinated tubulin, are the predominant filament type in these neurite-like projections, they did not appear to be essential for the observed movements. Instead, it was suggested that MTs enriched in tyrosinated tubulin provide the tracks for granule movement (Kreis et al., 1989). There is also evidence for the movement of constitutive secretory vesicles by rapid, saltatory movements (Amheiter et al., 1984). In this study fluorochromecoupled antibodies, prepared against the cytoplasmic tail of the vesicular stomatitis virus G-protein, were microinjected into virus mutant-infected BHK-21 fibroblasts, facilitating the study of the post-Golgi pathway of the viral membrane proteins in living cells. The movement of the vesicles was observed to occur by random jumps, with an overall direction towards the plasma membrane.

VL ROLE OF MICROTUBULES IN THE ORGANIZATION OF THE GOLGI COMPLEX The Golgi complex of mammalian cells is made up of stacks of flattened cistemae surrounded by vesicles and tubules (Figure 1). Material enters the stacks via the cis-Golgi network (CGN) and leaves them via the trans-Golgi network (TGN). Recent studies of isolated Golgi fractions have suggested that proteinaceous bridges are responsible for holding the cistemae together in the stacked configuration

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(Cluett and Brown, 1992). Electron microscopic observations farther indicate that the stacks are linked together by tubular extensions and thus form a large, continuous system (Roth et al., 1985; Rambourg and Clermont, 1990; Tanaka and Fukudome, 1991). Nevertheless, the Golgi complex is far from static in character and its structural integrity depends on a balance in the bidirectional transport of membrane between the ER and the cis-side of the stacks, across the stacks, and between the trans-side of the stacks and the cell surface (Figure 1). This dynamic state is revealed by brefeldin A (BFA), which causes an almost complete atrophy of the Golgi complex by blocking vesicular transport from the ER to the cis-side of the stacks, whereas transport in the opposite direction via long tubular extensions continues (Klausner et al., 1992). An extensive disorganization of the Golgi complex has also been observed in cells treated with ilimaquinone, a new drug which, like BFA, prevents binding of coat proteins to the Golgi membranes (Takizawa et al., 1993), and okadaic acid, an inhibitor of type 1 and 2A protein phosphatases (Lucocq et al., 1991; Thyberg and Moskalewski, 1992b). The idea that MTs take part in the organization of the Golgi complex is largely based on observations dealing with the spatial relationship between these organelles and the effects of MT-disruptive drugs on the morphology of the Golgi complex. A few initial attempts have also been made to study the interaction between MTs and isolated Golgi elements in the test tube. A. Spatial Relationship between Microtubules and the Golgi Complex The Golgi stacks are usually grouped together in a circumscribed region near the nucleus and around the centrosome, the organizing center from which MTs radiate into the surrounding cytoplasm (Farquhar and Palade, 1981). A close spatial relationship thus exists between MTs and the Golgi elements (Thyberg and Moskalewski, 1985; Duden et al., 1990). In recent double staining experiments a particularly high degree of overlap was found between MTs enriched in detyrosinated or acetylated tubulin and the Golgi elements (Skoufias et al., 1990; Burgess et al., 1991; Thyberg and Moskalewski, 1993). Because of their enhanced stability, these MTs may be especially well suited to take part in establishing and maintaining a normal organization of the Golgi complex. A possible mechanism of action is that specific cross-links are created between them and the Golgi elements. The responsible factors could be derived from the cytosol or be integral components of the Golgi membranes themselves. The dynamic character of the connection between the cytoplasmic MT system and the Golgi complex has been demonstrated repeatedly. As discussed above, the MTOC and the Golgi elements both change position to the front of the nucleus during cell migration and this evidently serves the function of directing transport of newly synthesized membrane components to the leading lamella (Singer and Kupfer, 1986). Likewise, a coordinated relocation of the MTOC and the Golgi elements takes place during myogenesis (Tassin et al, 1985) and during the

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establishment of epithelial cell polarity (Bacallao et al, 1989). These findings could to a large extent be explained by a rebuilding and reorientation of the MT system, rather than an active movement of the MTOC and the Golgi elements within the cell. It has previously been suggested that environmental signals are able to activate cell surface structures which protect the ends of MTs against disassembly (Kirschner and Mitchison, 1986). This will lead to a selective stabilization of MTs growing out from the MTOC in the direction of the external signal (for example, a chemotactic or a morphogenetic stimulus) and give rise to new cell shape and polarity. Conceivably, the Golgi complex remains closely associated with the MTOC during this process. At the same time, the nucleus and other organelles may have to move into new positions and thereby change the internal topography of the cell. B. Reorganization of the Golgi Complex in Cells Treated with Microtubule-Disruptive Drugs

Thirty years ago it was reported that the Golgi complex is fragmented in HeLa cells treated with colchicine and vinblastine (Robbins and Gonatas, 1964a). Similar observations have subsequently been made in a large variety of other cell types exposed to MT-disruptive drugs (Thyberg and Moskalewski, 1985; Duden et al., 1990). Briefly, these studies demonstrate that the Golgi stacks are detached from each other and spread throughout the cytoplasm in the absence of MTs (Figures 2 and 3). Although the individual stacks tend to become partly distorted during this process, they are retained as distinct entities. Following removal of the drugs, MTs reassemble and the Golgi stacks recollect in the region around the centrosome (see further below). Notably, the Golgi complex is also broken up in cells exposed to the MT-stabilizing drug taxol, but in this case the stacks associate with drug-induced MT bundles accumulating close to the cell surface (Wehland et al., 1983). Taken together, these findings indicate that normal MTs are essential for holding the Golgi stacks together in an interconnected system in the central part of the cell. On the other hand, MTs are apparently not required for holding the Golgi cistemae together in stacks (Figure 4). However, in a recent study of yeast cells, the loss of MTs was found to cause unstacking of the Golgi cistemae (Ayscough et al, 1993). Further insight into the mechanisms by which the Golgi complex is broken up in cells treated with MT-disruptive drugs has been obtained in the last few years. In living cells labelled with NBD-ceramide, tubular processes were observed to emerge from trans-Golgi cistemae, extend along MTs, and link adjacent trans-Golgi elements into a continuous reticulum. Nocodazole inhibited formation of such processes and caused an extensive fragmentation of the reticulum (Cooper et al., 1990). In parallel, it was observed that the nocodazole-induced dispersal of the Golgi complex is energy dependent and suppressed by dmgs which slow down transport through the stacks without depleting ATP (Tumer and Tartakoff, 1989). These findings suggest that the breakup of the lateral connections between the Golgi

Figure 3, Double Immunofluorescence localization of MTs and the Golgi complex In rat embryo fibroblasts. Normal cells (A and B) and cells treated with nocodazole (C and D) were stained with antibodies against tyroslnated (A) or acetylated (C) tubulin and the Golgi stack marker mannosldase II (B and D). In the normal cells (A and B) the Golgi elements are organized in a defined area around the center of the MT network. In the nocodazole-treated cells (C and D) both the remaining short fragments of acetylated MTs and the Golgi elements are widely dispersed in the cytoplasm. Bar marks 20 |im.

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MT disassembly > < MT assembly

Figure 4. Schematic model showing the organization of the Golgi stacks in the region around the center of the MT system in a normal interphase cell (left) and the dispersal of the stacks after drug-induced disassembly of cytoplasmic MTs (right).

stacks depends on the loss of functional MTs but also requires ongoing membrane traffic. Only if both of these requirements are fulfilled, the stacks will become disconnected from each other and be able to move apart. However, little is still known about how the Golgi fragments move apart. They may be dispersed in the cytoplasm as a result of Brownian movements or actin- and myosin-dependent contractions (cytoplasmic streaming). They could also interact more directly with the actin filaments and be translocated along them with help of motor proteins like myosin-I (Adams and Pollard, 1989). Experimental data supporting such a possibility have recently been presented. Golgi vesicles isolated from developing chicken epithelial cells were thus found to bind actin filaments and to include myosin-I as a cytoplasmically oriented membrane component (Path and Burgess, 1993). Moreover, it was reported that the actin-binding protein comitin (p24) is a component of the Golgi complex in slime molds as well as mouse fibroblasts (Weiner et al., 1993). On the other hand, cytochalasins (drugs which interfere with actin polymerization) have generally not been found to cause distinct changes in the morphology of the Golgi complex. Therefore, actin filaments are apparently less important than MTs in the normal organization of the Golgi complex. However, their ability to interact with and influence the distribution of the Golgi elements could be increased in the absence of MTs. Additional information about the role of MTs in the spatial arrangement of the Golgi complex has been obtained by analysis of cells recovering from exposure to nocodazole or other MT-disruptive drugs. Using NBD-ceramide as a vital stain, scattered Golgi elements were observed to move along reassembled MTs back to the region around the centrosome (Ho et al., 1989). Based on this finding, it was proposed that minus end-directed motor proteins like cytoplasmic dynein bind to the membranes of the Golgi elements and move them along MTs towards the centrosomal region. On the other hand, plus end-directed motor proteins like kinesin bind to secretory vesicles budding from the trans-Golgi network and move them along MTs to the cell surface (Duden et al, 1990; Kreis, 1990). In support of

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this model, recent studies on semi-intact cells have provided experimental evidence for the function of cytoplasmic dynein in the movement of isolated Golgi membranes towards the minus end of MTs (Corthesy-Theulaz et al., 1992). Once brought together in the region around the centrosome, the Golgi elements could be organized in a more stable manner by formation of tubular connections between the stacks (see above) and by anchoring to MTs by factors other than the motor proteins. In this connection, posttranslationally modified MTs could be of particular importance (Thyberg and Moskalewski, 1989). Immunocytochemical studies have indicated that tyrosinated (nonmodified) MTs are the first to be formed after removal of nocodazole and that they are responsible for the initial recollection of the Golgi elements in the region around the centrosome (Ho et al., 1989). Subsequently, MTs enriched in detyrosinated and acetylated tubulin accumulate in this region, supposedly by modification of tubulin subunits within the tyrosinated MTs (Skoufias et al., 1990; Burgess et al, 1991; Thyberg and Moskalewski, 1993). Based on the above observations, and results concerning the functions of MTs and MT-associated motors in the cell (Walker and Sheetz, 1993), the following model can be proposed (Figure 5). The Golgi stacks are normally organized together in the region around the centrosome and are linked to MTs enriched in posttranslationally modified tubulins (modified MTs). Golgi elements which for some reason have left this region and vesicles carrying material from the ER are transported towards the Golgi complex along MTs made up of tyrosinated tubulin (normal MTs) and with help of minus end-directed motor proteins like cytoplasmic dynein. Endosomes about to receive acid hydrolases packaged in the Golgi complex (and thereby transforming into lysosomes) are moved in the same direction along these MTs. On the other hand, secretory vesicles budding from the Golgi stacks are carried towards the cell surface along the same set of MTs with help of plus end-directed motor proteins like kinesin. centrosome

modified MTs

modified MTs

norma MTs dispersed Golgi elements, ER-derived vesicles, endosomes Figure 5. Schematic model of the role of cytoplasmic MTs in the organization of the Golgi complex. For further details, see text.

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To understand the function of MTs in the organization of the Golgi complex, it is essential to identify the molecules which mediate the interaction between these organelles. In one of the first investigations along this line, a monoclonal antibody raised against MAP-2 was found to recognize a 110 kD protein associated with the cytoplasmic face of rat liver Golgi membranes and able to interact with taxolstabilized MTs in the test tube (Allan and Kreis, 1986). Later on, it was shown that the association of the 110 kD protein with the membranes of the Golgi complex is blocked by BFA (Donaldson et al., 1990; Orci et al., 1991). In addition, the 110 kD protein was recognized as a major coat component of the non-clathrin-coated Golgi vesicles and named p-COP because of its homology to P-adaptin found in the coat of clathrin-coated vesicles (Duden et al., 1991b; Serafmi et al., 1991). These and other related studies have established p-COP as an integral component in the molecular machinery that operates in vesicular transport along the secretory pathway (Rothman and Orci, 1992). On the other hand, its possible role in linking the elements of the Golgi complex to the MT network remains to be clarified. In another report, a 58 kD protein exposed on the cytoplasmic surface of rat liver Golgi membranes and able to bind to MTs in vitro was described (Bloom and Brashear, 1989). Also in this case, additional work is required to confirm that the protein acts as a bridge between the Golgi complex and MTs in the intact cell. As a complement to the above investigations, a few initial reports on the interaction of isolated Golgi fractions with MTs have been published. First, it was shown that Golgi vesicles isolated from rat parotid glands bind to taxol-induced brain MTs in a saturable manner and that the reaction is inhibited by brain MAPs (Coffe and Raymond, 1990). Next, the binding of Golgi elements isolated from Vero cells to taxol-polymerized brain MTs was demonstrated to depend on proteins associated with the Golgi membranes as well as cytosolic factors sensitive to the sulfhydryl reagent N-ethylmaleimide (NEM) and distinct from cytoplasmic dynein (Karecla and Kreis, 1992). Also in this case, coating of the MTs with brain MAPs reduced the binding of the Golgi elements. Partly similar findings were made using Golgi elements isolated from rabbit liver (Murata et al., 1992). However, in this case cytosolic factors were not found to be required and the interaction was not affected by NEM or BFA, suggesting that the 110 kD protein (p-COP) was not involved. These studies do not allow any definitive conclusions to be made regarding the identity of the molecules which take part in the interaction between the Golgi complex and MTs. However, MAPs of the type which normally decorate MTs along their entire length and the minus end-directed motor protein cytoplasmic dynein do not appear to have a primary role in this process.

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Vll. REORGANIZATION OF THE GOLGI COMPLEX DURING CELL DIVISION During cell division (M-phase), the Golgi complex should be partitioned between the daughter cells. To serve this purpose, it is fragmented into small vesicles and dispersed in the cytoplasm during prophase and metaphase, and then does not resume its normal structure until late in telophase or cytokinesis. At the same time, vesicular transport along the exocytic and endocytic pathways ceases (Warren, 1985, 1993). In parallel, the radiating array of interphase MTs is broken down and temporarily replaced by the mitotic spindle (Mcintosh and Koonce, 1989). It seems likely that these events are related, but little is so far known about the underlying mechanisms. Interestingly, exposure of interphase cells to okadaic acid, an inhibitor of type 1 and 2 A protein phosphatases, has been found to induce both fragmentation of the Golgi complex and arrest of intracellular membrane traffic similar to that observed during prophase and metaphase (Lucocq et al., 1991; Thyberg and Moskalewski, 1992b). In addition, okadaic acid causes a partial loss of cytoplasmic MTs (Eriksson et al., 1992; Thyberg and Moskalewski, 1992b), primarily affecting the stable subset enriched in detyrosinated tubulin (Gurland and Gundersen, 1993). In a study of extracts of interphase sea urchin eggs, okadaic acid was also found to induce a mitosis-like decrease in MT length and lifetime (Gliksman et al., 1992). These findings direct attention to the possible role of protein phosphorylation in the structural reorganization of the cell at the onset of mitosis. M-phase promoting factor (MPF), a complex of cdc2 kinase and cyclin, has been identified as a common cell cycle regulator in eukaryotic cells, acting via specific prqtein phosphorylation (Clarke and Karsenti, 1991; Sherr, 1993). However, it still remains to be clarified whether or not okadaic acid exerts its effects via this system (directly or indirectly). In any case, okadaic acid has emerged as a new pharmacological tool to explore the mechanisms behind the breakup of the Golgi complex during the first part of mitosis (Lucocq, 1992). A.

Fragmentation of the Golgi Complex in Prophase-Metaphase

Many years ago, electron microscopic and cytochemical studies of mitotic HeLa cells indicated that the Golgi complex disappears during prophase and reappears during telophase (Robbins and Gonatas, 1964b). Since then, these observations have been confirmed and extended in several cell types (Thyberg and Moskalewski, 1985; Warren, 1985,1993). A series of studies on HeLa cells revealed that the Golgi stacks disintegrate into groups of small vesicles (Golgi clusters) as well as numerous free vesicles during the first part of mitosis and that these distribute equally between the daughter cells (Lucocq et al., 1987, 1989; Lucocq and Warren, 1987). The identification of the clustered vesicles as Golgi elements was based on labeling with antibodies against galactosyltransferase, a marker of trans-Golgi cistemae. It was later reported that endocytic tracers like horseradish peroxidase and transferrin

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appear in tubulovesicular structures having similar characteristics to the clusters mentioned above (Tooze and HoUinshead, 1992). The apparent contradiction between these results has not yet been fully explained. One possibility is that there exists two populations of clusters in mitotic HeLa cells, one representing fragmented Golgi stacks and another representing tubular endosomes (Pypaert et al, 1993). In studies on L929 and CHO cells, the fragmentation of the Golgi complex at the onset of mitosis could be resolved into at least two steps (Moskalewski and Thyberg, 1990; Thyberg and Moskalewski, 1992a). During prophase, the Golgi stacks were first separated from each other and dispersed throughout the cytoplasm in a similar way as in interphase cells treated with MT-disruptive drugs (see above). During metaphase, the stacks were further disintegrated and replaced by Golgi clusters of the type mentioned above and similar in appearance to those found in cells treated with okadaic acid (Thyberg and Moskalewski, 1992b). In this work, mannosidase II was used as a Golgi marker. In interphase cells, the most intense staining was obtained in the medial cistemae of the stacks. In mitotic cells, a positive staining was seen only in a few of the clustered vesicles. On the other hand, a distinct staining was seen in the ER, which was negative in interphase cells (Thyberg and Moskalewski, 1992a). Based on these findings, it is suggested that the disorganization of the Golgi complex at the onset of mitosis takes place in the following way (Figure 6): in prophase, the radiating system of interphase MTs disassembles and, as a result, the lateral connections between the Golgi stacks are broken and they are dispersed in the cytoplasm (Cooper et al., 1990). Next, the individual stacks disintegrate and, during this process, the more proximal parts fuse with the ER, whereas the more distal parts are transformed into clusters of vesicles. These changes resemble those seen in cells treated with BFA (Klausner et al., 1992), ilimaquinone (Takizawa et al., 1993), and okadaic acid (Lucocq et al., 1991; Thyberg and Moskalewski, 1992b). They could be due to an interference with membrane transport through the Golgi stacks, possibly regulated via coat proteins and specific protein phosphorylation. As a result, the Golgi complex is broken down into small dispersed units and equally partitioned between the daughter cells in connection with cytokinesis. B. Reformation of the Golgi Complex in Telophase-Cytokinesis Studies on HeLa cells have indicated that the reassembly of the Golgi complex during telophase and cytokinesis involves at least two steps (Lucocq et al., 1989). First, free vesicles and small clusters of vesicles gather close to the ER at several sites and the vesicles fuse into cistemae. Next, the stacks formed in this way are brought together in the juxtanuclear region and interconnected by tubular extensions to form a united Golgi complex of interphase type. At least the initial steps in the reassembly of the Golgi elements appear to take place before newly synthesized proteins arrive from the ER (Souter et al., 1993).

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Figure 6. Schematic model showing how the MT system and the Golgi complex are reorganized during cell division. For further details, see text.

Additional studies on other cell types have pointed out that a close relationship between the Golgi complex and the MT system is re-established in the postmitotic phase (Moskalewski and Thyberg, 1992;ThybergandMoskalewski, 1992a, 1993). At the end of telophase, the mitotic spindle disassembles and new interphase MTs grow out from the centrosomes. In each daughter cell, the Golgi stacks are recollected in the region around the centrosome, supposedly by transport along reforming MTs in a similar way as in interphase cells recovering from treatment with nocodazole (Ho et al., 1989). In analogy with the situation in the latter cells, tyrosinated MTs are the first to be formed after disappearance of the mitotic spindle and are evidently responsible for the initial regathering of the dispersed Golgi elements. Later on, acetylated and detyrosinated MTs appear in close association with the Golgi complex and thus could contribute to the maintenance of its normal interphase structure (Thyberg and Moskalewski, 1993). Studies of synchronized cells have further revealed that the centrosome and the Golgi complex are both most frequently located in the region close to the intercellular bridge one hour after the release of the mitotic block and on the opposite side of the nucleus two hours after the release of the block (Moskalewski and Thyberg, 1992; Thyberg and Moskalewski, 1992a). These observations suggest that the centrosomes, which at the end of telophase appear at the poles, first are relocated to the region close to the intercellular bridge in each daughter cell (Mack and

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Figure 7. Double immunofluorescence localization of MTs and the Golgi complex in rat embryo fibroblasts in the terminal phase of cytokinesis. Synchronized cells were fixed for one (A and B) or two (C and D) hours after the release of a mitotic block and stained with antibodies against tyrosinated tubulin (A and C) and the Golgi stack marker mannosidase II (B and D). The coordinated shift in location of the Golgi complex and the MTOC from the proximal to the distal side of the nucleus (as related to the intercellular bridge) is demonstrated. Bar marks 20 i^m.

Rattner, 1993), and that the Golgi complex is reorganized here. Later on, in the terminal phase of cytokinesis, the centrosome and the Golgi complex together shift position to the other side of the nucleus (Figures 6 and 7). This relocation of the Golgi complex at the end of M phase could serve the function to direct transport of membrane constituents and secretory products first to the elongating intercellular bridge and thereafter to the leading edge as the daughter cells are about to separate and move away from each other (Moskalewski et al, 1994).

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V m . SUMMARY AND PERSPECTIVES A considerable amount of evidence has accumulated supporting the function of MTs in the transport of proteins between the different compartments of the secretory pathway. The role of MTs as tracks in the targeting of Golgi-derived vesicles to the cell surface in many cell types has been considered already for some time and is documented by recent studies of polarized epithelial and neuronal cells. In contrast, the suggested participation of MTs in the early part of the secretory pathway represents a more unexpected development. In light of the present evidence, it can be suggested that MTs have analogous roles in pre- and post-Golgi transport as tracks in the long distance movement of transport intermediates. It has generally been assumed that these intermediates in most cases take the form of small carrier vesicles. However, recent results have revealed that intercompartmental transport can also be accomplished by tubular intermediates. Whether the MT-based tubular transport represents a special case or a general mechanism of intercompartmental communication is the subject of future studies. Recent data indicates that MT-based transport is also involved in the positioning of the major secretory organelles, the ER and the Golgi complex, within the cytoplasmic space. However, it remains to be seen whether the mechanisms of these membrane movements are different from those involved in intercompartmental protein transport or whether the two processes are in fact tightly coupled in the living cell. The present distinction may largely result from the limited and nonoverlapping methodology which is available to study these events. Finally, it should be emphasized that although we have limited the present discussion to the role of MTs in the secretory process, these represent only one of the components in the interacting network of cytoskeletal filament structures. In addition to the much studied role of actin filaments in the exocytic process, there is increasing evidence suggesting that actin and associated motors (myosin I) may also function at other locations of the secretory pathway. It can be speculated that the cytoplasm in fact contains multiple motility systems with overlapping functions in intracellular traffic pathways. This possibility remains one of the unresolved questions and a major challenge for future studies of the secretory process.

ACKNOWLEDGMENTS J.S. is a Norwegian Cancer Society Fellow. J.T. was supported by grants from the Swedish Medical Research Council, the Swedish Heart Lung Foundation, and the King Gustaf V 80th Birthday Fund.

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INDEX

AA, 192 Abl, 210, 213, 219-220 Acetylcholine receptor, 211 Acridine orange, 254 Actin, 11-13, 67-68, 73-75, 85-90, 211, 223-224, 227, 258 (5ee also "Focal adhesions...") -based organelle transport, 61 integrins, actin filaments and, 176180 mRNA,42 localization of, 45, 78 polymerization of, 38-39 Actin-binding proteins, 11-12 Adenylyl imidodiphosphate (AMP PNP), 125 AEl, 144-147, 152-153 Agrin, 212-213 Aldolase, 4-5, 9-10, 13 Alpha-actinin, 85, 177-178, 182, 190 AMP PNP, 125 Amphibians, cytoskeleton in early development of, 88-100 actin filament network, 90 actin network, 93-96, 97-99 [Ca^l, 96-97 cortical cytoskeletal domain, 88 cytokeratin network, 90-91 intracellular signals, 97-99 pathways, 96

Maturation Promoting Factor (MPF),92, 99 meiotic maturation, 92, 93 myosin, 97-99 PKC activity, 95-99 poly(A)"RNA, 93-96 "spatial fractionation," 93, 94 tubulin mRNAs, 93-96 vimentin, 91 vinculin, 90 XMAP, 99 Anion exchanger, 144-147, 152-153 Ankyrin, 151-161,229 adhesion proteins, association of with, 150 and Na^ K^-ATPase, 147-149 polarized distribution of, 144-147 Aphidicolin, 67 Arachidonic acid (AA), 192 Arg, 219 Ascidians, cytoskeleton in early development of, 77-80 ATP, 16-17, 18 Basic fibroblastic growth factor, 191 BFA, 243, 245, 249, 255, 260, 262 Blastomeres, 63 Bombesin, 188, 217 Brefeldin A (BFA), 243, 245, 249, 255, 260, 262

275

276

Cadherins, 218-219 Calcineurin, 122, 190 Calmodulin, 21 Calpain, 190-191 Catenins, 218-219 Cell signaling systems and membrane receptors, interactions of with cytoskeleton, 207238 {see also "Membrane receptors...") CGN, 242-243, 254 Chondroitinase ABC, 188 Chromatophores, pigment granule movements in, 120-122 cis-Golgi network, 242-243, 254 Colchicine, 35, 43, 119, 154-157, 244, 251,256 Collagen, 172, 190-191 Comitin, 258 Concanavalin A, 191 iS-COP, 260 Costameres, 169-170 Cyclic AMP, 16-17 Cytochalasins, 258 B, 43, 66, 68 D,45 Cytoplasmic transducers, 208 Cytoplasmic streaming, 115, 258 Cytoskeletal-bound polysomes (CBP), 32, 40 c-myc mRNA, 40-41 insulin, effect of, 38 Cytoskeletal function, specializations in during early development, 59-112 {see also "Specializations...") Cytoskeleton in metabolic compartmentation, role of, 1-30 concluding comments, 26-28 covalent modification of enzymes and cellular structure, 13-16 aerobic versus anaerobic glycolysis, 14

INDEX

binding sites, 15-16 cAMP, 13-14 phosphorylation, 13-14 pyruvate kinase, 14 energy requirements for signal transduction, 16-18 ATP, 16-17, 18 cyclic AMP, 16-17 glycolysis, 16-18 protein kinase C, 17 receptor proteins, 16-18 enzyme multiplicity and interactions with cellular structure, 9-11 A-type enzyme, 9-10 aldolase, 4-5, 9-10, 13 glyceraldehyde phosphate dehydrogenase, 10 intermediary metabolism, compartmentation in, 25-26 introduction, 2-3 carbohydrate metabolism, microcompartmentation of, 3 glycolysis, microcompartmentation of, 3 history, 2 self-organization, concept of, 2 in vivo, insufficiency of procedures for, 2-3, 6-7 matrical compartmentation, 18-20 advantages, 19-20 Na-K plasma membrane pump, 18 micro-compartmentation of carbohydrate metabolism, 3-9 actin, 9 aerobic versus anaerobic glycolytic rate, 3-4, 12 ALD, 4-5 binding, degree of, 3-5 difficulties in defining, 2-3, 6-7 digitonin-treated cells, use of, 78

Index

enzyme ambiguity, concept of, 7 enzyme kinetics, modification of, 3 enzymes, glycolytic, 3-10 facilitated binding, 5 GADPH, 4-5, 13 glycolytic sequence as series of segments, 6 PGK, 4-5 protein concentration, 7 TPI, 4-5 tubulin, 9 perturbations of compartmentation during cellular differentiation and dysfunction, 2124 in diabetes, 24 during early ontogeny, 21 Ehrlich ascites tumor cells, 24 hexokinase, 21 hormones, 24 tumor cells, 23-24 Warburg effect, 24 structure within, variation of, 1113 actin-binding proteins, 11-12 cAMP, 12 F-actin, 11 fimbrin, 12 DAG, 192, 224 Development, early, specializations in cytoskeletal function in, 59-112 {see also "Specializations...") DiOC6(3), 253 Diabetes, 24 Diacylglycerol (DAG), 192, 224 Drosophila, cytoskeletal development in, 64-72 Dynamin, 225-226

277 Dynein, 126-136, 159-160, 254, 259260 (see also "Microtubulebased...") cytoplasmic, occurrence of, 61, 128 nucleotide specificity and drug sensitivity of, 131 structure and molecular structure of, 129 Dystrophin, 212 E-cadherin, 147, 149-151, 161 Echinoderms, cytoskeletal development in, 73-77 ECM, 160 -coated magnetic beads, 182 Eggs, special development problems of, 62-64 Ehrlich ascites tumor cells, 24 Embryos, special development problems of, 62-64 Endoepithelium, 142 Epithelial polarity, cytoskeleton and development of, 141-165 binding sites between integral membrane proteins and membrane cytoskeleton, 153 ankyrin, 151 LEEDY/IRRRY motifs, 152153 protein 4.1, 152-153 cytoskeleton, role of in development of, 160-161 electrochemical gradient, creation of, 161 extracellular matrix (ECM), formation of focal contacts with, 160 hypothesis, three-stage, 160-161 proteins, stimulating transcription of, 160-161 introduction, 142-144 electrochemical gradients, generating, 142-143

278

membrane cytoskeleton, role of, 144-151 AEl, 144 anion exchanger, 144 ankyrin, association of with adhesion proteins, 150 ankyrin and protein 4.1, polarized distribution of, 144-147 E-cadherin, 147, 149, 150 Na^, K^-ATPase, ankyrin and, 147-149 polarity, partial reversal of in endoepithelial cells, 151 schematic, 149 uvomorulin, 150 microtubules, role of in cell polarity, 154-160 apical and basolateral membrane proteins, intracellular transport of, 154 dynein, apical carrier vesicles and, 159-160 Golgi apparatus, 154, 157-158 microtubule-disrupting drugs, alteration by, 155-158 transport barrier hypothesis, 158-159 vectorial intracellular transport, as structural basis for, 154 plasma membrane, 142 apical domain, 142, 143 basolateral domain, 142, 143 endoepithelium, 142 microvilli, 142 tight junctions, 144 zonulae occludentes, 144 Erk, 222 Extracellular ligands, 208 Extracellular matrix (ECM), 160, 168-169 and integrins, 172-173 Ezrin, 222, 227

INDEX

F-actin, 11 FAK, 181, 187 {see also "Focal adhesions...") bombesin, 188 paxillin, 187 rho, 188-189 tyrosine phosphorylation of focal adhesion proteins, 187 Fibrinogen, 172-176, 191 Fibronectin, 168, 172-176, 183, 186, 190-193 Fimbrin, 12, 142 Focal adhesions and integrinmediated cell signaling, 167205 conclusion, 193 cytoplasmic face, 176-183 a-actinin, 177-178, 182 cCRP, 179 ECM-coated magnetic beads, 182 FAK, 181 FRNK, 181 insertin, 181 isolation of focal adhesions, 182-183 LIM domains, 182 paxiUin, 180 ppl25^''^ 181 protein interactions, other, 180182 protein kinase C, 177 proteins linking integrin to actin filaments, 176-180 Rous sarcoma virus (RSV), 180 SH2 domains, 181 talin, 176-178 tensin, 178-181 tenuin, 182 VASP, 182 vinculin, 177-181 wet-cleavage, 182 ZnCl2, 182 zyxin, 179, 182

Index

focal adhesions, 169-172 in blood clot formations, 169 costameres, 169, 170 dynamics, 171-172 homologous structures in vivo, 169-171 invadopodia, 171 model, 179 neuromuscular junctions, 169 "podosomes," 169-171 protein kinase C, 171 "rosette adhesions," 169-171 Rous sarcoma virus (RSV), 169-171 stress fiber assembly, 172 talin, 171,175 targeting to, 173-176 transient, 171-172 vinculin, 171 vitronectin, 172-174 integrin-mediated signal transduction, 189-193 arachidonic acid (AA), 192 BAPTA, 189-190 ([Ca^']i),189-190 DAG, 192 fibronectin, 190-193 gelsolin, 192 GRGDSP peptides, 191 HUVECS, 190 inositol phosphates, 192 lipid metabolism and PKC, 192193 pl30, 189 paxillin, 189 pHi, 191 PIP2, 192-193 platelet derived growth factor (PDGF), 192 profilin, 192 RGD peptides, 191 tyrosine phosphorylation, 189

279 integrins, 172-176 a and j8 cytoplasmic domains, 174-176 chimeric, 174-176 ECM interactions, 172-173 ECM ligand-binding sites, 173176 fibrinogen, 172-176 fibronectin, 172-176 a and P heterodimers, 172 on platelets, 173 RGD sequences, 172 structure, 172 vitronectin receptor, 172-174 introduction, 168-169 fibronectin, 168 regulation of, 183-189 bombesin, 188 chondroitinase ABC, 188 cycloheximide, 188 ECM, 181 FAK, 187 in epitheUal cells, 185 fibronectin, 183 heparin-binding domain, 186 herbimycin A, 187 integrins, alterations in, 183-184 lysophosphatidic acid, 188 osteoclasts, 186 paxillin, 187 phosphotyrosine, 186-188 PKC, activation of, 184-186 in platelets, 188 PMA, 185 podosomes, 186 polylysine, 187-188 protein expression, 183-184 rho, 188-189 tensin, 187 TPA, 185 tyrosine phosphorylation, 186-188 vinculin expression, change in, 184

280

Fodrin, 66, 68, 73, 85, 150, 222 FRNK, 181-182 GADPH, 4-5, 13 Gametes, 62-63 Gelsolin, 192, 223 Golgi complex, function of microtubules in protein secretion and organization of, 239273 {see also "Microtubules...") Growth factor receptor kinase, 61 GTP-binding proteins, 61 Herbimycin A, 187 Hexokinase, 21 High voltage electron microscopy (HVEM), 32 Horse-radish peroxidase, 261-262 HVEM, 32 Ilimaquinone, 255,262 Inositol phosphates, 192 Insects, cytoskeletal development in, 64-72 Insertin, 181 Insulin effect of on association of polysomes with cytoskeleton, 3839 and polysome-containing fractions, 48-49 Integrin-mediated cell signaling, 167205 {see also "Focal adhesions and...") Integrins, 209, 217 Interference reflection microscopy (IRM), 171 Intracellular transducers, 209 "Invadopodia," 171 Ion channel receptors, 208-213 IRM, 171

INDEX

Karyokinesis, 64-65,68 Kinectin, 135 Kinesin, 61, 71-72, 87, 92, 125-136, 159, 250, 258-259 {see also "Microtubule-based...") light chains, 129 nucleotide specificity and drug sensitivity of, 131 occurrence, 128 structure and molecular structure of, 128-129 Krebs II ascites cells, 38-40, 48-49 Laminin, 191 Lysophosphatidic acid, 188 M-phase promoting factor (MPF), 261 Madin-Darby canine kidney (MDCK) cells, 147, 150, 158-159,251,253 Malignancy, cell characteristic in, 23-24, 191 Mammals, cytoskeleton in early development of, 80-88 blastocyst formation, 80-88 embryonic compaction, 80-81 MAP kinase cascade (Erk), 221-222 MAPs, 116-1X1 {see also "Microtubule-based...") MARCKS, 224 Maturation Promoting Factor (MPF), 92, 99 MBP, 33 MDCK cells, 147, 150, 158-159, 251, 253 MEK, 222 Membrane-bound polysomes (MBP), 33 Membrane receptors and cell signaling systems, interactions of with cytoskeleton, 207-238

Index

conclusions and prospects, 228230 ankyrin, 229 crosstalk, 230 cytoskeleton, 228 phosphorylation, role of, 230 reorganization of cytoskeleton, 230 signal transduction complexes, multi-component, 228 spatial and temporal localization, importance of, 228 introduction, 208-211 Abl, 210 cytoplasmic transducers, 208 extracellular ligands, 208 genetics, use of, 210-211 integrins, 209 intracellular transducers, 209 ion channel receptors, 208-210 membrane receptors, 208 membrane signaling and cytoskeleton, 208-211 mitogenesis, 208 neoplastic transformation, 208, 210 nonreceptor kinases, 209, 213220 proteins, four classes of, 208 receptor kinases, 209 serpentine receptors, 209 signal transduction particles, 210 T cell receptor, 209 transcription factors, 208, 210 ion channels, 211-213 acetylcholine receptor clustering, 211-213 actin, 211 agrin, 212-213 dystrophin, 212 ligand-gated, 211 localization, 211

281

nicotinic acetylcholine receptor, 211 receptor clustering, 211-213 )8-spectrin, 211 Torpedo acetylcholine receptor clusters, 211-232 tyrosine phosphorylation, 213 tyrphostin, 213 nonreceptor kinases and cytoskeleton, 213-220 Abl, 213, 219-220 arg, 219 activation mechanism not well understood, 213 cadherins, 218-219 catenins, 218-219 dephosphorylation, 216 Fyn, 213-214 integrins, 217 Lck, 213-214 localization, 216 morphology of cell, 216 neoplastic transformation, 208, 210 orthovanadate, 218-219 p21 rho, 218 phosphorylation, 213, 217 RaplB, 218 retroadaptors, 217 serine/threonine kinases, 213 SHI, SH2, and SH3, 214-219 specificity, 213 Src family, 213-219 T cell receptor, 213 tensin, 218 3BP-1, 220 tyrosine kinases, 213 receptor kinases, 220-228 A431 cells, 222-224 actin, 223, 227 ASGP-2, 226-227 autophosphorylated sites, 220221

282

DGK, 224, 225 diacylglycerol (DAG), 224 dynamin, 225-226 EGF-induced morphology changes, 222-226 EGF receptor association with microfilaments, 222-226 Erk, 222 ezrin, 222, 227 fodrin,222 gelsolin, 223 growth factors, 222 MAP kinase cascade, 221-222 MARCKS, 224 MEK, 222 mitogenic pathway, 221-222 neu oncogene, 226 nocodazole, 225 13762 cells, 226-227 phosphoinositides, 223 P13K, 225-226 pl85"^'/"'" association with microfilaments and signal transduction particle, 226228 PIK, 224, 225 PIP2, 223-224 PLCgamma, localization of, 224225 profilin, 223 protein kinase C, 224 Racl, 226 Ras, 221-222, 226-227 SH2 and SH3 interactions, 221226 spectrin, 222, 227 transmembrane complex, 227 transmembrane receptor, 221222 tyrosine kinase domains, 220221 Micro-compartmentation, 3-30 (see also "Cytoskeleton...")

INDEX

Microtubule-based intracellular transport of organelles, 113140 introduction, 114-115 cross-links, 114 cytoplasmic streaming, 115 motor proteins, 114-115 video microscopy, 114, 124-125 kinesin and cytoplasmic dynein, occurrence of, 128 microtubule and microfilamentous systems, interaction between, 123 Bryopsis, 123 model systems used to study, 119123 Allogromia, 122-123 calcineurin, 122 chromatophores, pigment granule movements in, 120-122 movements within cultured cells, 119-120 nerve axons, translocation along, 120-121 nurse cells and oocytes in insect ovaries, movement between, 121-122 Reticulomyxa, 122-123 reticulopodia of protists, movement within, 122-123 SCa and SCb, 120 motility substrates, microtubules as, 115-117 calcium, role in regulation, 133134 cAMP, role in regulation, 133134 dynein, 116 MAPS, 116-117 microtubule disposition in cells, 115-116 microtubule polarity, 116 as skeletal organelles, 115

Index

in spindles, mitotic, 116 and structural MAPs, 116-117 motors, identification and isolation of, 125-127 adenylyl imidodiphosphate (AMP PNP), 125 calcium, 133-134 cAMP, 133-134 characterization of, 128-131 directionality of organelle transport, control of, 132-136 dynein, axonemal, 126, 134 {see also "Dynein") families of, 131-132 force generation and mechanochemistry of, 129-130 genetic approaches to, 131-132 kinectin, 135 kinesin, 125-134 (see also "Dynein") MAP IC, 126 microtubule organizing centers (MTOCs), 126 microtubule motor proteins, 126 myosin 1,135 regulation of, 132-136 structural MAPs, influence of on activity, 136 tau, 136 taxol, 126 Triton X-100 soluble punctate structures, 134 vesicles, association with, 134136 organelles translocated, 117-119 colchicine, 119 endoplasmic reticulum (ER), 117-118 eukaryotic cells, 117 Golgi complexes, 117-119 lysosomes, 118-119 microfilaments, 119

283

ribosomes, 119 tubulin, 116 perspectives, 136 video-microscopy, 124-125 axoplasm, scrutiny of, 124-125 disadvantages, 124 Microtubules, function of in protein secretion and organization of Golgi complex, 239-273 cytoplasmic microtubule system, 243-244 colchicine, 244 drugs, 244 dynamic instability, 244 nocodazole, 244 taxol, 244 tubulin, 243-244 vinblastine, 244 vincristine, 244 Golgi complex, reorganization of during cell division, 261-264 BFA, 262 fragmentation of in prophasemetaphase, 261-262 ilimaquinone, 262 M-phase promoting factor (MPF), 261, 264 mannosidase II, 262 okadaic acid, 261-262 reformation in telophasecytokinesis, 262-264 schematic, 263 Golgi complex, role of in organization of, 254-260 jS-COP, 260 brefeldin A, 255 cis-Golgi network (CGN), 254 comitin, 258 cytochalasins, 258 cytoplasmic streaming, 258 dynein, 259-260 ilimaquinone, 255 kinesin, 258-259

284

and microtubules, spatial relationship between, 255-256 myosin I, 258 NBD-ceramide, 256, 258 nocodazole, 256-257 okadaic acid, 255 reorganization of, microtubuledisruptive drugs and, 256259 schematic, 258-259 taxol, 256 trans-Golgi network (TGN), 254 in vitro studies, 260 introduction, 240 movements of secretory organelles, 253-254 acridine orange, 254 DiOC(5(3), 253 fluorescence microscopy, 253 role of in secretory process, 244253 secretory pathway, membrane components of, 243 brefeldin A (BFA), 243, 255 cis-Golgi network (CON), 242243 salvage compartment, 242 schematic, 241 TGN markers, 243 trans-Golgi network (TGN), 242-243 secretory process, role of in, 244253 brefeldin A, 245, 249 CHO cells, 253 drugs, use of, 245 ER to Golgi, transport from, 245-248 Golgi to plasma membrane, transport from, 249-252 KDEL/HDEL sequence, 248 living cells, experiments with, 245

INDEX

localization studies, 245-246 lysosomes, transport to, 252 mannose-6-phosphate recognition marker, 252 MDCK cells, 251, 253 membrane recycling at ERGolgi interface, 248-249 nocodazole, 248-252, 254 NRK cells, 253 p53, 249 p58,249 PC12 cells, 251 protein transport, in vitro studies of, 252-253 taxol, 251,253 in vitro studies, 245, 249, 252253 summary and perspectives, 265 Mitogenesis, 208 Motor proteins, 114-115 mRNAs and polyribosomes, association of with cytoskeleton, 31-57 cytoskeletal components, protein synthetic apparatus and, 3538 colchicine, 35 cytochalasin B and D, 36-37 intermediate filaments (IF), 35 polysomes and microfilaments, 37 prosomes, 35 vimentin, 35 cytoskeleton, participation of in protein synthesis, 32-34 cytomatrix, 32-33 cytoskeletal-bound polysomes (CBP),32 high voltage electron microscopy (HVEM), development of, 32 immunocytochemistry, use of, 33

285

Index

membrane-bound polysomes (MBP), 33 ribosomes, 32-34 cytoskeletal-bound polysomes (CBPs), function of, 39-42 c-myc mRNA, 40-41 "free polysomes," 39 MBPs, 40-41 prosomes, 40 interaction of with cytoskeleton and polysomes, nature of, 46-51 in compartmentation of translation, 47, 49 FP, CBP, and MBP, 48-49 insulin stimulation, 48-49 mRNA/ ribosome binding proteins, 47-51 poly (A) binding proteins, 49-50 polysome-containing fractions, three, study of, 48-49 3'UTR of mRNA, 46-48 introduction, 32 protein targeting crucial, 32 localization of, cytoskeleton and, 42-45 actin mRNA, 45 myofibrils, 44-45 neurones, 45 oligodendroglia, 45, 50 and protein localization, 45 subcellular, mRNAs exhibiting, 44-45 vimentin mRNA, 44-45 perspectives, future, 52 physiological conditions, effect of on association of polysomes with cytoskeleton, 38-39 insulin, 38-39 Krebs II ascites cells, 38 polysomes, cytoskeletal-bound, function of, 39-42 (see also "... cytoskeletal-bound...")

Myosin, 68-70, 72, 97-99, 157, 159, 185, 258, 265 Na^ K^-ATPase, 147-149 NBD-ceramide, 256, 258 Neoplastic transformation, 208, 210 Nicotinic acetylcholine receptor, 211 Nocodazole, 155, 225, 244, 248-249, 251-252,254,256-257,259, 263 Nodes of Ranvier, 150 Nonchordates, cytoskeletal development of, 64-77 echinoderms, 73-77 insects, 64-72 Non-receptor kinases, 209 NP-40, 87 Nurse cells, 64-65 Okadaic acid, 255, 261-262 Organelles, microtubule-based intracellular transport of, 113140 {see also "Microtubulebased...") Orthovanadate, 218-219 Osteoclasts, 186 pl3K, 225-226 p21 rho, 218 Paxillin, 180, 187 PDGF, 192 Pentament, 81 Phorbol esters, 217 PIP2, 192-193 Plasma membrane, 142 (see also "Epithelial polarity...") Plasmodium, 64 Platelet derived growth factor (PDGF), 192 "Podosomes," 169-171,186

286

Polyribosomes, association of with cytoskeleton, 31-57 {see also "mRNAs...") Polysomes "free," 39 Profilin, 192, 223 Prosomes, 35, 40 Protein 4.1, polarized distribution of, 144-147 Protein kinases nonreceptor, 213-220 receptor, 220-228 Protein kinase C, 17, 21, 171,^177, 184-185 Proteins classes of, four, 208 SHI, SH2, and SH3 domains, 214-219 Pyruvate kinase, 14 Rap IB, 218 Ras, 221-222, 226-227 Receptor kinases, 209 Retroadaptors, 217 Rho, 188-189 "Rosette adhesions," 169-171 Rous sarcoma virus (RSV), 169-171, 180, 186-188, 219 invadopodia, 171 "podosomes," 169-171, 186 "rosette adhesions," 169-171 RSV, 169-171, 180, 186-188 SDS-PAGE, 48, 145 Sea urchin, cytoskeletal development in, 72-77 Ser/Thr kinase Raf, 222 Serine/threonine kinases, 213 Serpentine receptors, 209 Sheets, cytoskeletal, 81-88 Snoods, 74

INDEX

Specializations in cytoskeletal function during early development, 59-112 in chordates, 77-100 actin mRNA, 78 alpha-actinin, 85 amphibians, 88-100 {see also "Amphibians...") ascidians, 77-80 blastocyst formation, 80-83 [Ca'li, 87 embryonic compaction, 80-81 fodrin, 85 in hamsters, 81-86 histone mRNA, 78 kinesin, 87 mammals, 80-88 {see also "Mammals...") Maturation Promoting Factor (MPF), 92 in mice, 81-88 myosin, 85 Pentament, 81 PKX agonists, 87-88 poly(A)^RNA, 78 sheets, 81-88 sperm penetration, 80, 85 striated filament bundles, 86 Styela, 11-iQ vimentin, 91 vinculin, 85,90 Xenopus laevis, 88-100 eggs and embryos, developmental problems of, 62-64 blastomeres, 63 cytoskeletal specializations, conservation of, 63-64 sperm penetration, 62-64 introduction, 59-62 in cell mobility, 60, 62 in cell shape, 60 growth factor receptor kinase, 61

287

Index

GTP-binding proteins, 61 intracellular transport, 60-61 kinesin, 61 limitations of cell culture method, 62 protein and mRNA localization, 60, 61 tyrosyl kinase, 61 in nonchordates, 64-77 actin, 67, 73-75 aphidicolin, 67 bicoid mRNA, 66, 72 ([Ca^^li), transient increase in, 76-77 cellularization process, 69-71 cyclin B mRNA, 72 cytochalasin B, 66 Drosophila, (A-ll echinoderms, 73-77 insects, 64-72 karyokinesis, 64-65, 68 mutants, Drosophila, study of, 71 myosin, 68-70, 72 no(i mutations, 71 nullo, 69 nurse cells, 64 PKC, 77 sea urchin, 73-77 serendipity alpha, 69 shibire ^voitm, 71 snoods, 74 spectrin, 66, 68, 73-74 star fish, 73-76 villin,75,77 summary, 100 Spectrin, 66, 68, 73-74, 144-153, 222, 227 i8-, 211-212 Sperm penetration, developmental problems of egg and embryo and, 62-64

Styela, cytoskeleton in early development of, 77-80 T cell receptor, 209 Talin, 171, 175-178, 184-187, 193 Tau, 136 Taxol, 86, 126, 244, 251, 253, 256, 260 Tensin, 178-181, 187, 218 Tenuin, 182 TGN, 242-243, 254 Thrombin, 190-191 Thrombospondin, 172 trans-Golgi network, 242-243, 254 Transcription factors, 208, 210 Transferrin, 261-262 TritonX-100,84,87 Tropomyosin, 211 Tubulin, 243-244 Tumor cells, 23-24 Tween-20, 84, 87 Tyrosine kinase, 171, 213 Src gene family, 213-219 Tyrosine phosphorylation, 189 Tyrosyl kinase, 61 Tyrphostin, 213 Uvomorulin, 150, 161 Vanadate, 131 VASP, 182 Video microscopy, 114, 124-125 {see also "Microtubulebased...") Villin, 75 Vimentin, 35, 84, 91 mRNA, 44-45 Vinblastine, 155, 157, 244, 256 Vincristine, 244 VincuUn, 85, 90, 171, 177-181, 186187, 190 expression, regulation of, 184

288

Vitronectin, 168, 172-174, 189-191 von Willebrand's factor, 172 Warburg effect, 24 Xenopus laevis, 88-100 (see also "Specializations...")

INDEX

XMAP, 99 Zygote, 62-63 Zyxin, 179, 182

The Cytoskeleton A Multi-Volume Treatise Edited by J.E. Hesketh, Rowett Research Institute, Aberdeen and Ian Pryme, Department of Biochemistry and Moiecular Biology, University of Bergen This four volume treatise will cover aspects of the cytoskeleton from basic biochemistry and cell biology to those of relevance to medical science. The first volume of the treatise deals with the structural aspects of the cytoskeleton: the characteristics of the filaments and their components, the organization of the genes, motor proteins, and interactions with membranes. The second volume deals with the functions of the cytoskeleton indifferent cellular processes such as cell compartmentation and organanelle transport, secretion and cell attachment. In volume three the functional theme is continued but in this case the emphasis Is on the cytoskeleton in different tissues such as bone, liver, and intestine. Lastly, in the fourth volume a selection of pathological situations which are related to defects in the cytoskeleton are discussed. Contributions from leading scientists working in internationally renowned research centres have been commissioned. These articles will not only describe the background necessary to introduce the topical area and present sufficient details of recent work so as to give an up to date account of the various concepts and hypotheses involved, but the authors will also where appropriate give their own research perspective on the subject. The chapters will thus represent the cutting edge of research, emphasizing directions in which the subjects are developing and where future progress will prove fruitful. The Editors' envisage of the books will be of interest to postgraduate student/post-doctoral workers both actively involved in cytoskeleton research and other areas of cell biology. By including physiological and clinical aspects of the subject, the hope is that the volumes will also be of use in the teaching of medical students. Volume 1, Structure and Assembly 1995,300 pp. ISBN 1-55938-687-8

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CONTENTS: Preface, Sudarshan K. Malhotra. Early Appearance of Neuropeptides during Development: Neuromodulatory and Neurotrophic Roles, Susan Kentroti and Antonia Vernadakis. Role of Membrane Calpain (CA21"Activated Neutral Proteinase) in Central Nervous System, Naren L Banik and Edward L Hogan. Synaptic Ribbons of the Mammalian Pineal Gland: Enigmatic Organelles of Poorly Understood Function, Kunwar P. Bhatnagar. Structural Characteristics of the Transcription Factor Tfiiia and Its Interaction With Nucleic Acids, MarkL Brown and David P. Bazett-Jones. The Nuclear Estrogen Receptor of the Mammalian Uterus: Role in Transcriptional and Post-Transcriptional Control of Gene Expression, Raghava V. Thampan. Peroxisomes in Biology and Medicine, Inderjit Singh. Biological Basis of Hypertrophic Scarring, Paul G. Scott, Aziz Ghahry, Margaret H. Chambers, and Edward E. Tredget. Multiple Origins of Intravacuolar Protein Accumulation of Plant Cells, Eliot M. Herman. Controls of Germination in Noncereal Monocotyledons, Darleen A. DeMason. Also Available: Volumes 1-2 (1991-1993)

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PART A — CONTENTS: Preface, Bhagirath Singh. Expression and Function of IgM and IgD in Normal and Immunoglobulin-Transgenic Mice, Robert Brink, Christopher C. Goodnow, Jeffrey Crosbie, Elizabeth Adams, and Anthony Basten. Signal Transduction by the B-Cell Antigen Receptor (mig), Anthony L DeFranco, Michael R. Gold, William R. Hempel, Debbie Law, Dawne M. Page, Linda Matsuuchl, and Vivien Ghana. Antigen-Specific B-Cell Signaling and Desensltization, Alan H. La zarus and Terry L. Delovitch. CD23 and CD72: C-Type Lectins and B-Lymphocyte Regulation, John Gordon. Generation and Characterization of Functionally Distinct Subpopulations of B Cells, Richard R. Hardy and Kyoko Hayakawa. A New View of B-Cell Activation, Philip D. Hodgkln and Marilyn R. Kehry. Interactions Between Signals Generated Via Surface Ig and IL-4 Receptors in the Regulation of B-Cell Activation and Unresponsiveness, Gerry G.B. Klaus, Margaret M. Harnett and Robert D. Brines. Subject Index. Volume 1 Part B, 219 pp. ISBN 1-55938-614-2

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PART B — CONTENTS: Preface, Bhagirath Singh. Molecular Basis for Unresponsiveness and Tolerance Induction in Immature Stage B Lymphocytes, John G. Monroe, Amy J. Yellen-Shaw and Vicki L. Seyfert. Helper T Cell Signaling of B Cell Growth and Differentiation, Randolph J. Noelle and E. Charles Snow. Synergy Between Cell Contact and Cytokines in B Cell Activation, Trevor Owens, Diane Heath, andJohanne Poudrler. Expression of CD45 Isoforms (Leukocyte Common Antigen) and Adhesion Molecules During Normal and Abnormal Human B Lymphocyte Development, Una M. Pllarski and Gitte S. Jensen. B Cell Tolerance: Life or Death, David W. Scott, Mellnda Borrello, Lleh-Bang Llou, Yao Xlao-rui, and Garvin L. Warner. Natural History of Signaling Events in B Cells, Nicholas R. St. C. Sinclair. Subject Index.

Advances In Cell and Molecular Biology of Membranes and Organelles (Previosly published as Advances In Cell and Molecular Biology of Membranes) Edited by Alan M. Tartakoff, Institute of Pathology, Case Western Reserve University Volume 4, Protein Export and Membrane Biogenesis 1995,276 pp. $97.50 ISBN 1-55938-924-9

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Edited by Ross E. Dalbey, Department of Chemistry, The Ohio State University CONTENTS: Introduction to the Series, Alan M. Tartakoff. Preface, Ross E. Dalbey. Membrane Protein Assembly, Paul Whitley and Gunnar von Heijne. Membrane Insertion of Small Proteins: Evolutionary and Functional Aspects, Dorothee Kiefer and Andreas Kuhn. Protein Translocation Genetics, Koreaki Ito. Biochemical Analyses of Components Comprising the Protein Translocation Machinery of Escherichia coli, Shin-ichi Matsuyama and Shoji Mizushima. Pigment Protein Complex Assembly Ih Rhodobacter sphaeroides and Rhodobacter capsulatus, AmyR. Vargas and Samuel Kaplan. Identification and Reconstitution of Anion Exchange Mechanisms in Bacteria, AtuI Varadhachary and Peter C. Maloney Helix Packing in the C-Terminal Half of Lactose Permease, H. Ronald Kaback, Kirsten Jung, Heinrich Jung, Jianhua WU^ Gilbert C. Prive, and Kevin Zen. Export and Assembly of Outer Membrane Proteins in E. coli, Jan Tommassen and Hans de Cock. StructureFunction Relationships in the Membrane Channel Porin, Georg E. Schulz. Role of Phospholipids in Escherichia coli Cell Function, William Dowhan. Mechanism of Transmembrane Signaling in Osmoregulation, Alfaan A. Rampersaud. Index. Also Available: Volumes 1-3 (1993-1994)

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Advances in Cell Aging and Gerontology Edited by Paola S. Timiras, Department of Molecular and Cell Biology, University of California at Berkeley ar\6 E. Edward Bittar, Department of Physiology, University of Wisconsin, Madison Volume 1, In preparation, Fall 1996 ISBN 1-55938-631-2

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CONTENTS: The Cell Aging Process, Paola S. Timiras. Genes, Hormones and Aging, Peter J. Hormsby. Erythrocyte Aging, G. Bartosz. The Influence of Human Growth Hormone in Elderly Men, K. R. Shetty and David Rudman. The Cardiovascular System in Aging, James R. Docherty. Neuronal Calcium Regulation in Aging Brain, Mary L Michaelis and J. Huschenbett. Neuronal Aging and Alzheimer's Disease, G.J.C.G.M. Bosman and W.J. Grip. The Neurobiology of LateLife Psychosis, Frank W. Brown. Medicine in Elderly People, Peter H. Millard. Geriatric Psychiatry, D.G. Wilkinson.

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