Advances in Molecular and Cell Biology Volume 26: Cell Polarity

ADVANCES IN MOLECULAR AND CELL BIOLOGY CELL Volume 26

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ADVANCES IN MOLECULAR AND CELL BIOLOGY CELL POLARITY Series Editor:

E. EDWARD BITTAR Department of Physiology University of Wisconsin Madison, Wisconsin

Guest Editor:

JAMES R. BARTLES Department of Cell and Molecular Biology hbrthwestern University School of Medicine Chicago, lllinois

~

VOLUME 26

1998

@JAI PRESS INC. Stamford, Connecticut

London, England

Copyright 0 19981Al P R K S INC. 7 00 Prospect Street Stamford, Connecticut 06901 )A1 PRESS LJD. 38 Javistock Street Covent Garden London WC2f 7PB England All rights reserved. No part of this publication may be reproduced, stored on a retrieval system, or transmitted in any way, or by any means, electronic, mechanical, photocopying, recording, filming or otherwise without prior permission in writing from the publisher. ISBN: 0-7623-0381-6 Manufactured in the United States of America

CONTENTS

vii

LIST OF CONTRIBUTORS PREFACE James R. Bartles

ix

CELL POLARITY IN THE BUDDING YEAST SACCHAROMYCES CEREVlSlAE Christine Costigan and Michael Snyder

1

CELL POLARITY AND MOUSE EARLY DEVELOPMENT Tom P. Flemin Elizabeth Butler, lane Collins, Bfav Sheth, and Arthur E. Wild

67

SIGNALS AND MECHANISMS OF SORTING IN EPITHELIAL POLARITY Cara 1. Cottardi and Michael]. Caplan

95

THE GENERATION OF POLARITY IN NEURONAL CELLS Sharon K. Powell and Rodolfo 1. Rivas

133

POLARITY AND DEVELOPMENT OF THE CELL SURFACE IN SKELETAL MUSCLE Annelise 0.lorgensen

157

POLARITY AND POLARIZATION OF FIBROBLASTS IN CULTURE Albert K . Harris

201

INDEX

253

V

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LIST OF CONTRIBUTORS

Elizabeth Butler

Department of Obstetrics and Gynecology University of Southampton Southampton, England

Michael 1. Caplan

Department of Cellular and Molecular Physiology Yale University School of Medicine New Haven, Connecticut

lane Collins

Department of Pathology University of Southampton Southampton, England

Christine Costigan

Department of Biology Yale University New Haven, Connecticut

Tom P. Fleming

School of Biological Sciences University of Southampton Southampton, England

Cara 1. Cottardi

Department of Cell Biology Memorial Sloan Kettering Cancer Center New York. New York

Albert K. Harris

Department of Biology University of North Carolina Chapel Hill, North Carolina

Annelise 0.Jorgensen

Department of Anatomy and Cell Biology University of Toronto Toronto, Ontario, Canada

vii

...

LIST OF CONTRIBUTORS

Vlll

Sharon K. Powell

Department of Zoology University of Maryland College Park, Maryland

Rodolfo /. Rivas

Department of Zoology University of Maryland College Park, Maryland

Bhav Sheth

School of Biological Sciences University of Southampton Southampton, England

Michael Snyder

Department of Biology Yale University New Haven, Connecticut

Arthur E. Wild

School of Biological Sciences University of Southampton Southampton, England

PREFACE Few cells conform to the stereotype of the spherical blob hastily scribbled on chalkboards and, regrettably, sometimes even displayed prominently in textbooks. Instead, real cells display a remarkable degree of structural and functional asymmetry. In modern cell biological parlance, this asymmetry has come to be lumped under the general heading of “cell polarity.” Cell polarity is by no means restricted to the cells of tissues and organs, but can also be displayed by cells that lead a more solitary existence. The amazing extent to which cell morphology is correlated with function has long been a source of inspiration for biologists. Today the fascination continues unabated in the field of cell polarity, where it is fueled by an ever-deepening appreciation for the ways that fundamental cellular processes, such as membrane trafficking and cytoskeleton assembly, contribute to the establishment and maintenance of cell polarity. In the ensuing chapters, a collection of experts will summarize and interpret the findings obtained from basic research on cell polarity in a diverse array of experimental systems. James R. Bartles Guest Editor

IX

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CELL POLARITY IN THE BUDDING YEAST SACCHAROMYCES CEREVlSlAE

Christine Costigan and Michael Snyder

I . Overview . . . . . . . . . . . . . . . . . . . . . . . . ..................... 2 II. Budding During Vegetative Growth. . . . . . . . . . . . . . . . . . . . . . . . . ..................... 4 A. Cytology of the Budding Process . . . B. Components Important for Bud Form . . . . . . . . .24 C. Pathway of Assembly of Components D. Bud Site Selection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Coordination of Bud Initiation with Cell Cycle and Growth Control . . . . . 33 F. Other Signalling Mechanisms Underlie the Maintenance of Polarity. . . . . 36 G. Segregation of Organelles into the Newly Formed Bud. H . Speculations about Bud Formation and Growth. . . . . . I. Summary of the Budding Process. . . . . . . . . . . . . . . . . . . . . . . . Ill. Pseudohyphal Growth . . . . . . . . . . . . . . A. Polarized Growth and Divisions in Pse B. Known Inhibitors and Enhancers of Pseudohyphal Growth. . . . . . . . . . . . 42 C. Pseudohyphal Morphogenesis May Result from Differential Regulation of Components Conserved . .44 with Bud and Mating Projection Formation . . . . . . . . . . . . . . . .

Advances in Molecular and Cell Biology Volume 26, pages 1-66. Copyright 0 1998 by JAI Press Inc. All right of reproduction in any form reserved. ISBN: 0-7623-0381-6

1

CHRISTINE COSTIGAN and MICHAEL SNYDER

2

IV. Mating Projection Formatio A. Cytology of Mating Proje B. Cell Signalling in Response to Phero

. . . . . . . . . . . . . . . . . . 46 C. Pheromone Concentrat Initiation and Shape of D. Pheromone Concentration and Gradient Direct the Orientation of Polarized Growth . . . . . . . . . . . . . . . . . . . . . . . 48 E. Projection Orientation Is Controlled by Pheromone Receptors and Other Components at the Cortex F. Components Important for Projection Formation G. Summary and Model of How Projection Formation and Orientation Occurs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 V. Conclusion. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .53

1.

OVERVIEW

Polarized cell growth and polarized cell division are two basic processes important for both unicellular and multicellular organisms. Polarized cell growth is crucial for producing precise cellular structures and shapes that help mediate the specialized functions of distinct cell types (Drubin, 1991; Madden et al., 1992; Nelson, 1992). For example, polarized cell growth in pollen cells (Bedinger et al., 1994) and nerve cells (Eisen, 1994) mediates contact with specific targets; in epithelial cells polarized growth is essential for generating discrete cellular domains with distinct functions (Hubbard and Stieger, 1989). Polarized cell divisions occur at critical times in development and are responsible for directing appropriate cell-cell contacts, mediating growth in a specific direction, and establishing cytoplasmic differences between two daughter cells (Hyman and Stearns, 1992; Strome, 1993). Thus, both polarized growth and polarized divisions are essential for the development of tissues and entire organisms. How sites of polarized growth and division are selected and how subsequent growth events are directed toward these sites are processes that are only now beginning to be elucidated. Succhromyces cerevisiae undergoes both polarized cell growth and polarized cell division and is a useful organism for studying these processes (Drubin, 1991; Madden et al., 1992). Yeast cells undergo polarized cell growth at several times during the life cycle (Figure 1): during vegetative and pseudohyphal growth, and before mating. In the presence of ample nutrients S. cerevisiae cells propagate vegetatively by budding. At the end of G1 polarized growth initiates by formation of a bud on one edge of the cell. Cell growth is directed primarily into the bud which grows and enlarges until cytokinesis. The position of the bud ultimately determines the plane of cell division. The location of the bud site is either proximal or distal to the previous bud site, and is determined by the mating type locus and the pedigree of the cell (Freifelder, 1960; Hicks et al., 1977; Snyder, 1989).

Cell Polarity in Yeast

3 Vegetative Growth

/

Pseudohyphal Growth

O - d 5 a cell

Figure 7.

a cell

Three phases of the yeast life cycle during which polarized growth is

important. When yeast cells are exposed to less favorable growth conditions, such as limited nutrients, they can undergo pseudohyphal growth (Scherr and Weaver, 1953; Gimeno et al., 1992,1993). The cells still propagate by budding but they assume a dramatically elongated shape, fail to detach after cytokinesis, and form long chains which spread across a surface and invade solid medium. As in vegetative growth, both polarized growth and polarized cell divisions are important for pseudohyphal growth. The ability to both bud, that is, undergo asymmetric growth, and to form sequential buds in a specific direction are thought to enhance the ability of unicellular organisms to maximize access to nutrients in a spreading microcolony of either vegetative or pseudohyphal cells (Gimeno et al., 1992, 1993; Madden et al., 1992). During mating, after exposure to pheromone, yeast cells also exhibit polarized growth (Sprague and Thorner, 1992). The cells arrest in G1 and form a projection on one edge of the cell. Projections from each mating partner grow toward one another until they contact. The cell walls then break down at the contact site, and nuclear fusion (karyogamy) follows. The cell surface location of projection formation is apparently determined in response to local mating pheromone gradients generated by the mating partners (Jackson and Hartwell, 1990a, b; Segall, 1993), and is irrespective of previous growth sites (Madden and Snyder, 1992). Thus, during mating the sites of polarized growth are not predetermined, but rather are “plastic” as compared to the “hard wired’ or predetermined sites used during budding. Cell

CHRISTINE COSTICAN and MICHAEL SNYDER

4

morphogenesis during mating is reminiscent of cell differentiation in multicellular organisms, in which cells arrest cell division in response to an extracellular signal and differentiate into a specialized cell type. During the past six years there has been a vast expansion in our knowledge of both the basic processes and molecular components which control and carry out polarized growth in yeast at different times during the life cycle. This article reviews the different steps and components involved in polarized growth in vegetative, pseudohyphal, and mating yeast cells.

II.

BUDDING DURING VEGETATIVE GROWTH

Polarized growth during budding is composed of several steps (Madden et al., 1992). These include the decision to initiate growth, the selection of a site, the polarization of cytoskeletal and secretory components toward the growth site and subsequent growth events, and finally the segregation of cellular organelles such as the nucleus and mitochondria into the newly formed bud. Many components involved in yeast polarized growth participate in several of these steps. Prior to reviewing the steps involved in budding, the cytology of budding and components involved in this process are first discussed.

A.

Cytology of the Budding Process

Bud emergence begins at the GUS transition, at which time a small protrusion grows from one pole of the ellipsoidal cell. As the bud grows and enlarges, growth is thought to occur primarily at the bud tip. Labeling experiments indicate that new cell wall material is preferentially deposited at the bud tip (Tkacz and Lampen, 1972), and a variety of components important for growth of the bud concentrate at the tip (see below). As the bud nears its maximum size, there appears to be a transient phase of isotropic (i.e. uniform) growth in which cell wall deposition and growth components are localized throughout the bud. Once the bud reaches its final size, cytokinesis ensues. The secretory apparatus is presumably important for synthesizing and directing new plasma membrane and cell wall components to the growing bud tip (Schekman and Novick, 1981). Histochemical staining for alkaline phosphatase, a secreted protein, reveals preferential staining of the bud indicating that secretion is polarized within a vegetatively growing yeast cell (Field and Schekman, 1980). Prior to bud emergence, 3 W O nm vesicles accumulate at one edge of the cell (Matile et al., 1969; Byers, 1981), and during bud formation and growth the nascent bud is densely filled with membranous organelles (Matile et al., 1969; Byers, 1981; Preuss et al., 1991,1992). As the bud enlarges relatively fewer vesicles are present, and these vesicles appear to be in the process of fusing with the plasma membrane of the bud (Preuss et al., 1992).

Cell Polarity in Yeast

5

The secretory apparatus is nonrandomly organized in vegetatively growing yeast cells. Immunogold labeling experiments using antibodies to different proteins that reside within the endoplasmic reticulum and Golgi revealed that a significant portion (though not all) of each of these organelles concentrates at the incipient bud site and is segregated into the bud as it emerges (Preuss et al., 1991, 1992). The positioning of a portion of the secretory machinery within the bud is expected to enhance the efficiency with which components can be rapidly synthesized and incorporated into the site of new growth. In addition to the many vesicles in the bud, a series of small (1 50-250 nm in depth) membranous invaginations are observed which are surrounded by an electron dense material containing actin (Mulholland et al., 1994; see below). These invaginations have been hypothesized to be sites where localized cell surface growth occurs (Mulholland et al., 1994). It has been speculated that for cells with high turgor pressure such as yeast, deposition of plasma membrane and cell wall components into reinforced invaginations and their subsequent incorporation into the cell surface during growth might be an important mechanism for maintaining cell integrity. Direct deposition of components into the cell wall might cause cell lysis as the existing cell wall is locally degraded to allow incorporation of new components (Mulholland et al., 1994). One complication with this proposal is that the distribution of actin within the bud is much broader and organized into foci in comparison with the more diffuse localization of many other growth components (Spa2p, Myo2p, Cmdlp; see below). An alternative possibility is that the invaginations represent endocytic intermediates (Mulholland et al., 1994). Consistent with either of these proposed functions (in cell wall biosynthesis or endocytosis), strains carrying certain actin mutations are hypersensitive to osmotic stress and also exhibit defects in endocytosis (Novick and Botstein, 1985; Kubler and Riezman, 1993; see below). An important feature of budding is the constriction at the neck; without it, the bud would not be a bud. As the bud grows substantially both in length and diameter the neck enlarges relatively slightly. Chitin, a cell wall polysaccharide, is deposited as a thick ring around the neck outside the plasma membrane (Hayashibe and Katohda, 1973). Electron dense 10 nm filaments, regularly spaced 28 nm apart, also encircle the neck and lie just inside the plasma membrane (Byers and Goetsch, 1976). Presumably the combination of the chitin ring plus the 10 nm filaments provides structural support for maintaining the constriction at the neck. Ultimately, after cytokinesis, the chitin ring is asymmetrically partitioned; it will remain as a large annular bud scar [ 1.2 pm and 2 pm in diameter in haploids and diploids, respectively (Streiblova, 1970)] on the surface of the mother cell. The surface of the daughter cell is left with a region that is primarily devoid of chitin and enriched for mannan proteins; it appears as a brightly staining region when stained with fluorescein-conjugated Concanavalin A (FITC-ConA) (Lew and Reed, 1993). During budding, the constriction at the neck ultimately becomes the site of cytokinesis. This site is chosen early in the cell cycle and components important for

6

CHRISTINE COSTIGAN and MICHAEL SNYDER

neck formation and cytokinesis begin assembling before substantial progression through S phase (Hayashibe and Katohda, 1973; Ford and Pringle, 1991; Kim et al., 1991; see below). The establishment of the site of cytokinesis early in the cell cycle distinguishes budding yeast from many other organisms in which the cytokinesis site is determined relatively late, at anaphase. In these organisms the position of the mitotic spindle apparatus determines the site of cytokinesis (Rappaport, 1986).

B. Components Important for Bud Formation Analysis of components important for budding has revealed many aspects of the bud formation process. Polarized secretion and growth requires actin and a wide variety of actin-interacting proteins. In addition, a number of proteins including Bemlp and Spa2p localize at the growing surface of the cell and participate in polarized growth, perhaps as components or regulators of the cytoskeleton (Snyder, 1989, 1991; Pringle et al., 1995). Small GTP-binding proteins regulate bud formation and growth by modulating cytoskeletal function and secretion. Finally, components at the bud neck which constitute the 10 nm ring of filaments and the chitin ring are important both for bud morphology and ultimately for cytokinesis. Cytoskeletal and Secretory Components The Actin Cytoskeleton. Actin plays an important role in budding and polarity in yeast. Fluorescence microscopy with anti-actin antibodies or with rhodamine conjugated-phalloidin, which binds F actin, reveals that actin occurs in two forms in yeast. Actin patches (or spots) are preferentially located near the cell surface of the bud, and actin cables run longitudinally along the mothedbud axis near the cell surface (Figure 2; Adams and Pringle, 1984; Kilmartin and Adams, 1984). Actin patches form a ring at the incipient bud site in unbudded cells, they accumulate in the bud, particularly at the tip, in budded cells, and finally they localize at the neck between the mother cell and bud during cytokinesis (Adams and Pringle, 1984; Kilmartin and Adams, 1984). Biochemically, actin patches and cables are not identical. Some actin-binding proteins are present only in patches, and others only in cables (see below); presumably this reflects differences in actin function and/or assembly at these two sites. Recent electron microscopic analysis indicates that actin patches correspond to electron dense material surrounding the membrane invaginations described above (Mulholland et al., 1994). 7 nm actin filaments emanating from the cytoplasm frequently intersect the invaginations (Mulholland et al., 1994). Mulholland et al. speculate that the actin filaments might coil around the membrane invaginations and be contiguous with the cytoplasmic actin filaments (Figure 3).

7

Cell Polarity in Yeast

0 Veslcle

Actln patch MyoZp. Cmdlp. Bemlp, SpePp. Smylp, Cdc42p. Rholp, Secrlp -Actln cables -Mlcrotubules D

2‘

Vegetative

Mating

Figure 2. Localizations of several cytoskeletal and growth components in vegetative cells and mating cells. (Localizations of MyoZp, Smyl p, Rho1 p, and Sec4p in mating cells have not been determined.)

Yeast contains two actin genes, ACT1 andACT2; each of these genes is essential (Shortle et al., 1982; Schwob andMartin, 1992).ACTl encodes aprotein most homologous to the abundant actin isoform found in all eucaryotes (Gallwitz and Sures, 1980; Ng and Abelson, 1980). Actlp is the major actin isoform visualized by fluorescence microscopy because cells expressing only a mutant Act lp, that does not bind phalloidin in vitro, contain wild-type actin structures and do not stain with rhodamine conjugated-phalloidin (Drubin et al., 1993). ACT2 encodes a highly divergent actin-related protein, which is only 47% identical to Actlp (Schwob and Martin, 1992). The role of the Act2 protein in yeast is not known. Spores deleted for theACT2 gene, uct2-A cells, germinate and arrest as large budded cells, suggesting that Act2p is required during nuclear division (Schwob and Martin, 1992). Mutational analysis ofACTI indicates that actin plays a role in polarized growth and secretion in yeast. Thus far 21 different mutations of the ACT1 gene that cause growth inhibition at high temperatures (i.e., temperature sensitive or 7s” alleles) have been generated, the genetic lesion characterized, and the mutant phenotype analyzed (Shortle et al., 1984; Novick and Botstein, 1985; Dunn and Shortle, 1990; Johannes and Gallwitz, 1991; Wertman et al., 1992; Drubin et al., 1993). A large fraction of these mutants (76%) are cold-sensitive andor sensitive to high osmolarity (Novick and Botstein, 1985; Wertman et al., 1992). Two uctl(ts) mutants which have been studied extensively, actl - I and actl -2, exhibit polarized growth defects at the restrictive temperature: the asymmetric distribution of actin is disrupted, cells accumulate in the unbudded phase of the cell cycle, chitin is delocalized, and cell lysis is apparent (Novick and Botstein, 1985). Moreover, cell sizes are very heterogeneous, and large mother cells containing small buds are often present; these characteristics are expected of cells with polarized growth defects in which the mother cell grows at the expense of the bud (described also forABPl overexpression strains (Drubin et al., 1988; see Table 1). Other classes of uctl(ts) mutants exhibit morpho-

8

CHRISTINE COSTIGAN and MICHAEL SNYDER Cell Wall

1

Actin Patch

Actin Cable

Figure 3. A possible model for adin structure at the cortex. lnvaginations of the cell surface are encased by actin filaments. Myo2p might direct secretion by transporting membrane vesicles along cytoplasmic actin filaments that are contiguous with those encircling membrane invaginations. logical defects such as angular shapes, elongated buds, enlarged necks, and aberrant cell sizes (Wertman et al., 1992; Drubin et al., 1993). Actin participates in polarized growth and morphogenesis, at least in part, through its role in secretion (Novick and Botstein, 1985; Novick et al., 1989; Johnston et al., 1991). The a c t l ( t s ) mutants, act1 -1 and a c t l - 2 , accumulate abnormally high levels of the mature, glycosylated form of invertase and secretory vesicles at the restrictive temperature (Novick and Botstein, 1985). One postulate for how actin participates in polarized secretion is that the longitudinally arrayed actin cables provide a surface for the movement of secretory vesicles to the cell surface growth site. This hypothesis was tested by analysis of yeast strains deleted for either tropomysin ( t p m l - A mutants) or actin capping protein (cap1 - A cap2-A cells) which exhibit few, if any, actin cables during exponential growth (Liu and Bretscher, 1989b; Amatruda et al., 1992). Secretion of invertase in these mutants is not inhibited (Amatruda et al., 1992; Liu and Bretscher, 1992), and the mutants still form buds and exhibit growth localized at the bud tip, as visualized FITC-ConA labeling. Therefore, either residual actin cables are still present which function in polarized secretion, or actin cables are not essential for this process (Liu and Bretscher, 1989a; Amatruda et al., 1990, 1992; Liu and Bretscher, 1992). However, further analysis of t p m l - A mutant cells revealed that polarized secretion is not completely normal in the absence of actin cables. Exponentially growing t p m l - A cells accumulate vesicles which are likely to be late stage secretory vesicles, and secretion to the growing mating projection is delayed and partially delocalized in tpml - A cells at the restrictive temperature (Liu and Bretscher, 1992). Thus, although polarized secretion can occur in the absence of detectable actin cables, they probably contribute at least partially to this process.

Cell Polarity i n Yeast

9

In addition to its role in secretion, actin might be important for organization of the mitochondria within the cell. Mitochondria are usually associated with actin cables, and the distribution of mitochondria is often altered in act1 mutant cells (Drubin et al., 1993). Thus, actin may interact with and organize several types of membranous organelles within the cell. Myosins. Several myosins have been identified in yeast (Watts et al., 1987; Johnston et al., 1991;Haarer et al., 1994). The myosin type V homolog, Myo2p, is an unconventional myosin that is a candidate motor for transporting secretory vesicles to the cell surface along actin filaments (Figure 3; Johnston et al., 1991; Cheney et al., 1993). MY02 is essential for growth and extensive mutant analysis has been performed on cells carrying a myo2(ts) allele, myo2-66 (Prendergast et al., 1990; Johnston et al., 1991). At the restrictive temperature these cells are largely unbudded and enlarged because they grow isotropically, accumulate membrane vesicles in the cytoplasm, and are frequently multinucleate because the nuclear division cycle continues in the absence of budding (Prendergast et al., 1990; Johnston et al., 1991). Johnston et al. postulate that my02 cells are defective in polarizing secretion to a single site on the cell surface; secretion therefore occurs in these mutants by a random diffusion mechanism (Johnston et al., 1991). Myo2p localizes to growth sites: as a cap at the incipient bud site, to the tip of the bud in small budded cells, and to the neck at cytokinesis (Figure 2; Lillie and Brown, 1994). A direct association of Myo2p with vesicles has not been demonstrated. Nevertheless, consistent with the possibility that Myo2p interacts with actin filaments is the observation that the combination of myo2-66 and a deletion of the tropomyosin gene (tpml-Amutants, which, as noted above, have few actin cables; see Table 1) results in cell inviability at normally permissive temperatures for growth (Liu and Bretscher, 1992). The underlying assumption here is that even at permissive temperatures for growth, the myo2-66 mutant exhibits reduced protein activity. Myo2p Function is Probabfy Regufated by Calmodulin (Cmdlp). The function of Myo2p is likely to be regulated by yeast calmodulin, Cmdlp. Cmdlp concentrates at sites of localized cell surface growth in a similar pattern to that of Myo2p (Figure 2; Brockerhoff and Davis, 1992; Sun et al., 1992), and certain calmodulin mutants exhibit polarized growth defects including a failure to form buds (Davis, 1992; Ohya and Botstein, 1994), similar to the defect of my02 mutants. CMDI and MY02 interact genetically: cmdl mutants exhibit allele-specific poor growth or inviability phenotypes in combination with the ts allele, my02-66 (Brockerhoff et al., 1994). Cmdlp and Myo2p also interact biochemically: Myo2p coimmunoprecipitates with Cmdlp from yeast cell extracts and Cmdlp binds Myo2p in a gel overlay assay (Brockerhoff et al., 1994). Although Cmdlp is a Ca++-binding protein and Ca++appears to have some role in budding (see below), Cmdlp regulation of Myo2p is likely to be Ca++-independent(Davis et al., 1986; Brockerhoff et

10

CHRISTINE COSTIGAN and MICHAEL SNYDER

al., 1994). Identification of what aspect of Myo2p function is modulated by Cmdlp binding awaits a more precise description of how Myo2p participates in polarized growth. Actin-binding Proteins. In addition to myosins, a number of structural proteins which bind actin have been identified in yeast. As in multicellular organisms, the yeast actin-binding proteins appear to regulate the assembly and organization ofactin arrays. Mutant analyses, immunolocalization data, and effects of the yeast actin-binding proteins on actin in vitro suggest that, as might beexpected, these proteins participate in polarized growth by modulating actin function (see Table 1; reviewed in Welch et al., 1994). Actin-binding proteins which have been isolated and characterized i n yeast include cofilin, capping protein, fimbrin (Sac6p), profilin, tropomyosin, and Abplp (Drubin et al., 1988, 1990; Magdolen et al., 1988; Liu and Bretscher, 1989a,b; Haarer et al., 1990; Amatruda et al., 1990; Adams et al., 1991; Amatruda and Cooper, 1992; Amatruda et al., 1992; Liu and Bretscher, 1992; Moon et al., 1993). After biochemical purification from yeast, these proteins have all been demonstrated to share the functions of their homologs in other organisms (Liu and Bretscher, 1989a,b; Haarer et al., 1990; Adams et al., 1991; Amatruda and Cooper, 1992; Moon et al., 1993). For instance, yeast profilin inhibits actin polymerization and actin ATPase activity (Haarer et al., 1990) and yeast fimbrin bundles actin filaments (Adams et al., 1991). In vivo, Abplp, cofilin and capping protein localize only to actin patches (Drubin et al., 1988; Amatruda and Cooper, 1992; Moon et al., 1993), tropomyosin localizes to actin cables (Liu and Bretscher, 1989a), and fimbrin (Sac6p) localizes to both actin patches and cables (Drubin et al., 1988). These various localization patterns presumably reflect the different roles of these proteins in regulating actin function or assembly at these sites. In spite of the divergent functions that the actin-binding proteins carry out in vitro, and in many cases their different localization patterns, the actin-binding protein mutants often share certain characteristic “actinoid” defects. These include depolarization of actin spots, loss or disorganization of actin cables, changes in cell morphology and size, and sensitivity to stress conditions such as high osmolarity or high temperature (see Table 1 and references therein). Interestingly, most of the actin-binding structural proteins that have been identified are not essential for viability. To date the only exception to this rule is cofilin (Moon et al., 1993). Other yeast actin-binding proteins including Caplp and Cap2p, Sac6p, Pfylp, and Tpmlp are dispensable for growth except under stress conditions (Liu and Bretscher, 1989a; Amatruda et al., 1990; Haarer et al., 1990; Adams et al., 1991; Amatruda et al., 1992; Liu and Bretscher, 1992). Abplp, which is an actin-binding protein that is presently unique to yeast, is apparently completely dispensable for growth, even under temperature stress (Drubin et al., 1988, 1990). It therefore seems likely that many of the actin-binding proteins, and

Table 7. Activity In Vitro

Proteins That Bind Actin and/or Ceneticallv Interact with Actin Localization in Essential Vegetative Cells

Gene

Protein

ABPl

Binds F-actin Abpl p, contains a SH3 domain, cofilin homology domain

no

Anclp, actin Does not bind noncomplementing actin protein; component of TAF (11) complex

no

ANC2

Anc2p

n.t.

n.t.

ANC3

Anc3p

n.t.

ANC4

Anc4p

n.t.

ANC7

Vegetative Phenotype of Mutant

Synthetic lethal with*

anclA is ts, exhibits actin depolarization, swollen or elongated cells; ancl-1 is osm’

anc4, ski76

n.t.

anc2 is ts

anc3

(Vinh et al., 1993; Welch et al., 1993)

n.t.

n.t.

anc3 is cs

anc2

(Vinh et al., 1993; Welch et al., 1993)

n.t.

n.t.

anc4 is ts

ancl, tpm7A

(Vinh et al., 1993; Welch et al., 1993)

no

Actin patches, except actin patches at cytokinesis

cap strains are ts on high osmolarity, cells are rounded, exhibit chitin and actin depolarization, actin cables are diminished or absent, cell sizes are heterogeneous

sac64

(Amatruda et al., 1990; Amatruda and Cooper, 1992; Amatruda et al., 1992; Adams et al., 1993; Karpova et al., 1993)

Nucleus

slaZA

p subunits of Increases the capping protein, critical respectively concentration for actin polymerization lowers the viscosity of an

(Vinh et al., 1993; Welch et

al., 1993; Welch and Drubin, 1994)

_L

CAP2

(Drubin et al., 1988; Drubin et al., 1990; Adams et al., 1993; Holtzman et al., 1993; Moon et al., 1993; Mulholland et al., 1994)

sac6A, slalA,

-1

C A P l , a and

Reference

abpl-A, no detectable defects; ABPl overexpression strain is ts, exhibits actin depolarization, enlarged cells, bipolar budding in haploids

Actin patches

slcl, dC2

continued

Table 7. Activity In Vitro

Gene

Protein

COF7

cofilin

Effects on actin are consistent with actin filament severing activity

PN7

profilin

Localization in Essential Vegetative Cells

Synthetic lethal with*

Reference

Actin patches

ts mutants accumulate actin patches in mother cell

(Moon et al., 1993; Mulholland et al., 1994; Lappalainen and Drubin, 1997)

Slows the rate no of actin polymerization and inhibits actin ATPase activity

n.t

pfy7A is ts, osm’, exhibits chitin and actin depolarization, cells are often multinucleate, sizes are heterogeneous

(Haarer et al., 1990; Haarer et al., 1993)

RAH7, Rahp, suppressors RAH2, of act7 (osm’) RAH3

n.t.

n.t.

n.t.

rah3 cells are ts, slightly osm’, exhibit actin and chitin depolarization

(Chowdhury et al., 1992)

SAC7

n.t.

no

Endoplasmic reticulum and Golgi

sac7A is cs, exhibits adin depolarization, loss of actin cables, inositol auxotrophy; certain sac7 alleles suppress both actl(ts) and sec74A (Sec74p is a phosphatidylinositol/phosphatidylcholine transfer protein which is important for Golgi function)

(Cleves et al., 1989; Novick et al., 1989; Whitters et al,, 1993)

Saclp, suppressor of actl(ts)

yes

(Continued) Vegetative Phenotype of Mutant

-L

w

(Novick et al., 1989)

SAC2, SAC3

Sac2p and Sac3p, suppressors of act 1(t.5)

n.t.

n.t.

n.t.

sac2 and sac3 are cs, exhibit actin depolarization; sac2 mutants accumulate secretory vesicles

SAC6

fimbrin

Bundles actin filaments

no

Actin patches and cables

sac6A is ts, exhibits actin depolarization, susceptibility to lysis, defects in endocytosis, cells are rounded, sizes are heterogeneous

SAC7

Sac7p, suppressor of act 1(ts)

n.t.

no

n.t.

sac7A mutants are cs, lack actin cables, cells are rounded

SlAl

Slal p, synthetic lethal with abplA, contains three SH3 domains

n.t.

no

n.t.

s/alA is ts, exhibits actin depolarization with actin chunks (enlarged actin patches), cells are rounded, heterogeneous in size

abp 1A, anclA

(Holtzman et al., 1993; Welch and Drubin, 1994)

SM2

Slazp, talin homology at C-terminus

n.t.

no

n.t.

s/a2A is ts, exhibits actin depolarization, cells are rounded, often enlarged and rnultinucleate

abplA

(Holtzman et al., 1993)

SLC1

Slcl p, synthetic lethal with cap2

n.t.

n.t

n.t.

s/cl is weakly ts,actin is cap2A depolarized, cells are susceptible to lysis, form long chains with attached. elongated cells I

abp 1A, anc3, cap4 SIC2

(Drubin et al., 1988; Adams and Bostein, 1989; Adams et al., 1991; A d a m et al., 1993; Karpova et al., 1993; Kubler and Riezman, 1993; Vinh et al., 1993)

(Dunn and Shortle, 1990)

(Karpova et al., 1993)

"

continued

Table 1.

(Continued)

localization in Essential Vegetative Cells

Vegetative Phenotype of Mutant slc2 is ts, weakly OSm’, actin cables are absent, actin patches are depolarized, cells are susceptible to lysis, form long chains with attached, elongated cells

Gene

Protein

Activity In Vitro

SLCZ

Slc2p

n.t.

n.t.

n.t.

TPMl

tropomyosin

1Tpmlp molecule binds 3 actin monomers; binding requires divalent cations

no

Colocalizes with tprnlA ts, actin cables are very actin cables diminished or absent, chitin is depolarized, cells are rounded, heterogeneous in size, secretory vesicles accumulate

Synthetic lethal with* cap24 sacs6A

Reference (Karpova et al , 1993)

J

P

Key: cs: cold sensitive, ts: high temperature sensitive, osm’: high osmolarity sensitive, n.t.: not tested.

* Genetic backgrounds in which the mutant is inviable

anc4, myo2-66

(Liu and Bretscher, 1989a; Liu and Bretscher, 1989b; Liu and Bretscher, 1992; Vinh et al., 1993)

Cell Polarity in Yeast

15

the actin-interacting proteins (described below), carry out at least partially overlapping functions in yeast cell growth. Consistent with this theory, various double mutant combinations of the actin-binding protein mutants and/or actininteracting protein mutants exhibit severely reduced growth or inviability under permissive conditions for growth of the single mutants (Table 1). Another possible reason for the mild phenotypes of the actin-binding protein mutants is that, in some cases, additional homologs of a single protein might exist that have not yet been identified. For example, a second tropomyosin gene has been cloned from yeast (Drees et al., 1993). The examination of interactions between the genes encoding actin-binding proteins, including double mutant analysis and genetic suppression studies, has only recently been initiated. It is expected that more information about actin-binding protein function will be gained from such experiments, thereby clarifying the specific roles of each of these protein in polarized growth. Furthermore, the analysis of interactions of these proteins with the large number of defined actin mutants (Shortle et al., 1984; Novick and Botstein, 1985; Johannes and Gallwitz, 1991; Wertman et al., 1992; Drubin et al., 1993) should contribute significantly to what is known about the mechanisms by which these proteins regulate actin function in wivo. Genes which Genetically Interact with Actin and Actin-binding Proteins. Genetic screens and the isolation of extragenic suppressors have been used to identify a number of genes which genetically interact with actin. Mutants in the ANC genes exhibit allele-specific noncomplementation with actl mutants in a d / + and+ diploids (Vinh et al., 1993; Welch et al., 1993). sucmutants and ruh mutants were isolated as extragenic suppressors of the high temperature sensitivity and osmosensitivity, respectively, of actl mutants (Adams and Botstein, 1989; Novick et al., 1989; Dunn and Shortle, 1990; Chowdhury et al., 1992). In addition to the genes which interact with actin, several genes were identified through their interactions with genes encoding actin-binding proteins. Mutations in the SLA genes and SLC genes are not lethal in wild-type backgrounds but cause lethality (“synthetic lethality”) in combination with abpl-A (Holtzman et al., 1993) and cap2-A (Karpova et al., 1993), respectively. Conditional alleles of most of these genetically interacting loci have been characterized phenotypically; many cause actin depolarization, aberrant cell morphologies, and other actinoid phenotypes under restrictive conditions and synthetic lethality with other actin-interacting mutants under normally permissive conditions (see Table 1). Compared to the actin-binding proteins much less is known about the molecular structure, intracellular localization, and functions of most of the Anc, Sac, Rah, Sla, and SICproteins. Moreover, it is still possible that some of these proteins are identical either to one another, to one of the actin-binding proteins, or to other proteins important in polarized growth processes. Some exceptions are Saclp and the Sla proteins. S a c l p has been extensively characterized and is likely to interact with actin indirectly. It localizes to the Golgi and has been speculated to alter

16

CHRISTINE COSTIGAN and MICHAEL SNYDER

phosphoinositide levels and thereby the activities of various actin-binding proteins (Cleves et al., 1989; Novick et al., 1989; Whitters et al., 1993). Slalp and S la2p share sequence similarities with other proteins implicated in cytoskeletal function, and s l a l - A and s l a 2 - A mutants exhibit defects and synthetic lethal interactions which are suggestive of a role in polarized growth (Holtzman et al., 1993). Hence Slalp and Sla2p are more likely to directly affect actin cytoskeletal function. In summary, the phenotypes of the actin-binding protein mutants and the actininteracting protein mutants implicate these proteins in polarized cell growth. How these proteins are regulated and the specific polarized growth processes in which they participate remain to be elucidated. Bern I p and Spa2p Are Two Additional Cortical Proteins which Participate in Budding. Bemlp was identified by genetic interactions with the bud emergence

genes (see below) and has been postulated to link the membrane cytoskeleton with growth site determinants and thereby to effect cytoskeletal changes required for morphogenesis (Chenevert et al., 1992). Like Myo2p and Cmdlp, Bemlp localizes to growth sites in unbudded and small budded cells (Figure 2; Brockerhoff and Davis, 1992; Sun et al., 1992; Lillie and Brown, 1994; Pringleet al., 1995). The predicted Beml protein sequence contains SH3 domains (Chenevert et al., 1992), like many other proteins believed to be associated with the membrane cytoskeleton. The participation of Bemlp in polarized growth is shown by the phenotype of beml-A cells. Exponentially growing beml - A cells often have aberrant morphologies, are enlarged, have delocalized actin and chitin distributions, and are frequently multinucleate (Bender and Pringle, 1991;Chenevert et al., 1992). BEMl is essential for growth at high temperatures, and at the restrictive temperature arrested beml cells are mostly unbudded (Bender and Pringle, 1991; Chenevert et al., 1992). It is possible that Bemlp affects the cytoskeleton only indirectly, and that it serves as an anchor at cortical growth sites with which regulatory proteins associate, thus ensuring crucial regulatory interactions occur at the proper site. Like Bemlp, Myo2p, and Cmdlp, Spa2p also localizes to growth sites in unbudded and small budded cells (Figure 2; Snyder, 1989; Snyder et al., 1991; Brockerhoff and Davis, 1992; Sun et al., 1992; Lillie and Brown, 1994; K. Corrado and J. Pringle, personal communication). In addition Spa2p is also present in the buds of medium-sized and most large budded cells. It is diffuse as buds approach their maximum size, probably coincident with the transition to isotropic bud growth, and then it is present at the neck during cytokinesis, although its prevalence at this location varies with the particular strain background (Snyder, 1989; Snyder et al., 1991). Spa2p is predicted to have a coiled-coil domain, and it has been suggested that Spa2p might be a cytoskeletal protein (Gehrung and Snyder, 1990). Deletion of SPA2 causes only mild defects during vegetative growth: cells are marginally rounder than wild-type cells and they have mild cytokinesis and bud site selection defects (Snyder, 1989; Snyder et al., 1991; Flescher et al., 1993).

Ceff Pofarity in Yeast

17

spa2-A beml-A mutants are inviable indicating that Spa2p and Bemlp may carry out overlapping functions in polarized growth (Costigan et al., 1992). In contrast, deletion of SPA2 in a strain containing a mutant form of the polarized protein, Cmdlp, has no effect on growth of the cmdl mutant (Brockerhoff et al., 1994). This suggests that either the Spa2p and Cmdlp functions are entirely independent, or that Spa2p function requires Cmdlp and thus is already abolished in the cmdl mutant. The Putative Neck filament Proteins. Another important class of proteins that participate in budding are the closely related Cdc3, Cdc10, Cdcl 1, and Cdcl2 proteins. These proteins each accumulate as a ring at the incipient bud site well before bud emergence, remain at the neck during bud formation and growth, and persist through cytokinesis (Haarer and Pringle, 1987; Ford and Pringle, 1991; Kim et al., 1991; H. Kim, S. Ketcham, B. Haarer, and J. Pringle, personal communication). Cdc3p, CdclOp, Cdcl lp, and Cdcl2p are thought to comprise the ring of 10 nm neck filaments observed by electron microscopy at the mother-daughter constriction (Byers, 1981). The CDC3 and CDC12 genes are essential for yeast cell growth; CDCIO and CDCII are not, but deletion of either of these genes causes severe growth defects (Flescher et al., 1993; J. Pringle, personal communication). Ts mutations in any of the putative neck filament genes cause a loss of the filaments at the restrictive temperature and failure to undergo cytokinesis, resulting in long chains of connected cells (Hartwell, 1971; Byers, 1981). Homologs of these proteins have been identified in a variety of organisms including mammals (Kato, 1991), C. albicans (DiDomenico et al., 1994), and Drosophila (Neufeld and Rubin, 1994). In Drosophila these proteins also participate in cytokinesis, indicating a general role for these proteins i n this process (Neufeld and Rubin, 1994). The Cdc3, CdclO, Cdcll, and Cdcl2 proteins are predicted to bind GTP (Flescher et al., 1993). It has been speculated that these proteins may form filaments through the hydrolysis of GTP, analogous to the proposed role for nucleotide hydrolysis in other filament-forming proteins (e.g., actin and tubulin) (Flescher et al., 1993). The GTP-binding protein, FtsZ, is important for septation in E. coli (de Boer et al., 1992; RayChaudhuri and Park, 1992) suggesting a universal role for GTPbinding proteins in cytokinesis. Microtubules May Play an Ancillary Role in Bud Growth. Microtubules are not essential for bud formation in yeast. tub2 mutants, which are defective in the single yeast P-tubulin gene, and cells treated with the microtubule-depolymerizing drug, nocodazole, still form buds normally (Huffaker et al., 1988; Jacobs et al., 1988). However, it is possible that microtubules play an ancillary role in bud formation and/or growth. Cytological evidence indicates that microtubules extend into the bud (Figure 2; Byers and Goetsch, 1975), and myo2(ts) mutant defects can be suppressed by multiple copies of SMYI, which encodes a kinesin homolog (Lillie andBrown, 1992). Smylp, like Myo2p, localizes to cell growth sites (Figure 2; Lillie and Brown, 1994) and a SMYl deletion causes no detectable defects in wild-type

18

CHRISTINE COSTIGAN and MICHAEL SNYDER

cells, yet it causes inviability in a myo2(ts) strain (Lillie and Brown, 1992). These various data suggest that actin-based motility and microtubule-based motility may have partially overlapping roles in polarized growth, with actin playing the predominant role. Among other possibilities, Smy l p and Myo2p might both transport secretory vesicles to growth sites. Redundant actin-based and microtubule-based motility systems have been described in squid axoplasm, where a single organelle can have both actin-based and microtubule-based motors on the surface (Kuznetsov et al., 1992). Thus, both actin and tubulin may have redundant functions in vesicle transport in a variety of systems. Secretory Components. As might be expected from the polarized localization of the secretory apparatus, several proteins involved in the late steps of secretion have recently been shown to exhibit a polarized distribution (Schekman and Novick, 1981; Salminen and Novick, 1989; Novick and Brennwald, 1993; D. TerBush and P. Novick, personal communication). Late steps in secretion are mediated by the Sec4p GTP-binding protein, which is associated with late secretory vesicles and the plasma membrane (Salminen and Novick, 1987; Goud et al., 1988). Activated, GTP-bound Sec4p has been hypothesized to function in exocytosis by facilitating the targeting of secretory vesicles to the vesicle fusion site on the plasma membrane (Walworth et al., 1989; Brennwald and Novick, 1993). It is speculated that Sec4p regulates the interaction between integral membrane protein tags on the vesicles, “v-SNARE” proteins, and recognition molecules on the plasma membrane, the “t-SNARES” (Novick and Brennwald, 1993). Sec4p-GTP is thought to be required for V-SNARE activity (Brennwald and Novick, 1993; Novick and Brennwald, 1993), and upon vesicle fusion at the plasma membrane, GTP is hydrolyzed on Sec4p, thus allowing Sec4p release and recycling for subsequent cycles of vesicle targeting (Walworth et al., 1989). Sec6p, Sec8p, and SeclSp are also involved in the late steps in secretion and form part of a large complex that is peripherally associated with the plasma membrane (Salminen and Novick, 1987; Bowser and Novick, 1991; Bowser et al., 1992). Immunofluorescence experiments show that Sec4p (Figure 2; Novick and Brennwald, 1993), Sec6p (TerBush and Novick, 1995), Sec8p (D. TerBush and Novick, 1995), and SeclSp (Salminen and Novick, 1989) each localize to the incipient bud site in unbudded cells and the tip of the bud in budded cells. Presumably the late secretory components are targeted to a site on the plasma membrane specified by the cortical cytoskeleton. Rho CTPase Cycles that Regulate Budding

In mammalian cells, the Rho (Ras-homologous) GTP-binding proteins appear to regulate actin cytoskeletal dynamics and cell morphology (reviewed in Chrzanowska-Wodnicka and Burridge, 1992). A number of GTP-binding proteins which are related to members of the Rho family and function in bud formation a n d o r growth have been identified in yeast. In many cases regulators of these pro-

Cell Polarity in Yeast

19

teins have also been identified and characterized. It is probable that these GTPases function in targeting components to specific subcellular locations, analagous to the functions proposed for a wide variety of other GTPases, for example, Sec4p (Walworth et al., 1989; Novick and Brennwald, 1993), EF-Tu (Kaziro, 1978), and Ras (Leevers et al., 1994). The Cdc42p GTPase is Essential for Polarized Growth and Budding. Cdc42p is a Rho homologous protein that was identified in a genetic screen for cdc (cell division cycle) mutants defective in bud emergence (Adams et al., 1990;Johnson and Pringle, 1990). Ts cdc42 mutants arrest at the restrictive temperature as unbudded cells (Adams et al., 1990). Although they have stopped dividing the mutant cells are biosynthetically active and continue to grow isotropically; they become enlarged, often multinucleate, exhibit delocalized chitin and actin patches over the entire cell surface, and have few, if any, actin cables (Adams et al., 1990).Dominant activating alleles of CDC42 can cause cells to form multiple buds (Ziman et al., 1991), consistent with its essential role in polarized growth and bud emergence. Functional homologs of Cdc42p have been isolated from a wide variety of other organisms, including humans (Shinjo et al., 1990) and C. elegans (Chen et al., 1993), suggesting that the function of Cdc42p might be widely conserved in eucaryotes. Cdc42p is another member of a growing class of proteins which localize at growth sites (see Figure 2). It is present as a patch at the incipient bud site in unbudded cells, at the tip of the bud in budded cells, and sometimes is detected at the neck at cytokinesis (Ziman et al., 1993). Immunoelectron microscopic analysis revealed that Cdc42p localizes to the membrane invaginations in the bud which have also been associated with actin (Ziman et al., 1993;Mulholland et al., 1994). Consistent with its localization pattern at the cell periphery, Cdc42p is associated with membranes (Ziman et al., 1993). Regulators of Cdc42p. Cdc42p activity is apparently regulated by the other bud formation proteins, Cdc24p and Cdc43p. Ts cdc24 and cdc43 mutants exhibit identical polarized growth defects as cdc42 mutants (Hartwell et al., 1973; Johnston et al., 1977; Sloat and Pringle, 1978; Sloat et al., 1981; Adams and Pringle, 1984; Adams et al., 1990). Several lines of evidence show that Cdc43p is the geranylgeranyl transferase (GGT) for Cdc42p. In common with other Ras superfamily proteins, Cdc42p contains a consensus sequence for isoprenylation at its carboxy terminus (-C-A-A-X; A = aliphatic) (Hall, 1990; Johnson and Pringle, 1990; Clarke, 1992). The particular motif found at the carboxy terminus of Cdc42p is -C-A-A-L (Johnson and Pringle, 1990), making Cdc42p a candidate for prenylation by type I geranylgeranyltransferase (GGT I) (Clarke, 1992). Mutagenesis of the conserved cysteine to serine abolishes function of Cdc42p (Ziman et al., 1991) and the tight association of Cdc42p with membranes (Ziman et al., 1993), suggesting that the association of Cdc42p with membranes is essential for Cdc42p function. Yeast express GGT I activity which is dependent on CDC43 (Finegold et

20

CHRISTINE COSTICAN and MICHAEL SNYDER

al., 1991). Based on the predicted protein sequence and in vitro functional analysis, Cdc43p is the p subunit, which, together with the Ram2p a subunit constitutes the yeast GGT I (Ohya et al., 1991; Mayer et al., 1992). Combination of ts alleles of CDC42 and CDC43 results in synthetic lethality at normally permissive temperatures for either mutant alone, suggesting that their gene products interact (Adams et al., 1990). Furthermore, simultaneous overexpression of CDC42 and RHO1 (another yeast Rho protein with a carboxy terminal C-A-A-L motif; see below) rescues the lethality of a cdc43-A mutant (Ohya et al., 1993). Simultaneous overexpression of Cdc42p and Rholp that have been mutated at the carboxy terminus to carry a consensus sequence for farnesylation, rather than geranylgeranylation, restores growth of the cdc43-A mutant to wild-type levels (Ohya et al., 1993). These experiments suggest that Cdc42p and Rholp are the essential substrates of Cdc43p/Ram2p, and that the requirement for Cdc43p/Ram2p is bypassed when Cdc42p and Rho l p are prenylated by farnesyltransferase. As with other Rho proteins, the guanine nucleotide bound state of Cdc42p appears to be essential for its function (Ziman et al., 1991). Cdc24p is likely to be the GDP dissociation stimulator for Cdc42p. Cdc24p shares a region of similarity with the Dbl oncogene product which promotes GDP release from the human Cdc42p homolog, Cdc42Hs (Hart et al., 1991; Ron et al., 1991). In vitro, yeast Cdc24p specifically stimulates guanine nucleotide exchange on Cdc42p, and not on two other yeast GTPases implicated in budding, RsrlplBudlp or Rholp (Zheng et al., 1994). In agreement with the model that the essential function of Cdc24p is to activate Cdc42p, overexpression of CDC42 allows growth of a cdc24(ts) mutant at the restrictive temperature (Bender and Pringle, 1989). Bem3p contains a Rho GTPase activating domain and is a likely GTPase activating protein (GAP) for Cdc42p (Zheng et al., 1994). The GAP domain of Bem3p can bind to and stimulate the GTPase activity of Cdc42p in vitro, whereas the GAP domain of Bem2p, another protein with a Rho GAP domain (see below) cannot (Zheng et al., 1994). It is likely that there are other GAPSfor Cdc42p that have yet to be identified because unlike CDC24, CDC42, and CDC43, BEM3 is not an essential gene (Coleman et al., 1986; Johnson and Pringle, 1990; Miyamoto et al., 1991; Ohya et al., 1991; Zheng et al., 1994). Deletion of BEM3 results in no detectable growth or cytoskeletal defects (Zheng et al., 1994). Genes which interact with the CDC24, CDC42, and CDC43 gene consort have been identified. When present in multiple copies two genes, MSBI and MSB2, suppress the cdc24(ts) growth defect; multicopy MSBl plasmids can also suppress cdc42(ts) defects (Bender and Pringle, 1989). MSBl and MSB2 are not essential for yeast cell growth, and msbl-A, msb2-A and msbl-A msb2-A strains exhibit no detectable growth defects under a variety of conditions (Bender and Pringle, 1991; Bender and Pringle, 1992; A. Bender, unpublished data, cited in Bender and Pringle, 1992). Despite the lack of obvious defects, Msblp probably participates in cell growth, because deletion of MSBl is lethal in combination with a mutation in the BEMl gene (Bender and Pringle, 1991). The predicted protein sequence of

Cell Polarity in Yeast

21

Msblp exhibits no distinguishing features or homologies with other known proteins (Bender and Pringle, 1991), while that of Msb2p contains two putative transmembrane domains (Bender and Pringle, 1992). Model for the Cdc42p CTPase Cycle. One model for Cdc42p function in polarization of growth is as follows (see Figure 4). The Cdc42p GTPase might target cytoskeletal Components to specific sites on the cortex or helps assemble precomplexes (see below). Binding of a Cdc42p-associated complex at the cortex would be coupled to GTP hydrolysis on Cdc42p. Cortical markers recognized by Cdc42p could include the putative Cdc42p GAP, Bem3p, and might be established, in part, by the Rsrlp(Bud1p) GTPase (see below). Alternatively, Cdc42p-GTP itself might not be important for recognition of the cortical marker, but might facilitate an interaction between a targeting molecule and the cortical marker, analagous to the hypothesized role of Sec4p in exocytosis (see above; Novick and Brennwald, 1993). The components brought to the cortex by Cdc42p might include actininteracting proteins, actin, or even Rholp (see below). Polarized growth occurs in the absence of GTP hydrolysis on Cdc42p: constitutively activated cdc42 mutants can polarize and form buds (Ziman et al., 1991). However it is likely that GTP hydrolysis on Cdc42p enhances polarization because constitutively active cdc42 mutants do exhibit substantial overall growth and enlargement, suggesting an excess of undirected, isotropic growth (Ziman et al., 1991). It is possible that Cdc42p also promotes general surface growth, and that in this respect its function partially overlaps that proposed for Rholp (see below). GTP hydrolysis might facilitate docking of growth components at the cortex andor recycling of Cdc42p which would be important for subsequent rounds of Cdc42p targeting activity. Rholp is Also lmplicated in Bud Growth. In addition to Cdc42p, there are a number of other yeast Rho proteins, including Rholp, Rho2p, Rho3p, and Rho4p, that are involved in budding. Rholp is unique among the Rho proteins in that, like Cdc42p, it is essential for growth (Madaule et al., 1987; Yamochi et al., 1994). Also like Cdc42p, Rholp accumulates at growth sites: the incipient bud site, the tip of the bud in budded cells, and the neck at cytokinesis (Figure 2; Yamochi et al., 1994). Rholp has also been reported to associate with the Golgi and post-Golgi secretory vesicles (McCaffrey et al., 1991), but these findings have been questioned (Yamochi et al., 1994). As suggested by the subcellular localization pattern of Rholp, the protein functions in budding (Yamochi et al., 1994). Furthermore, at the restrictive temperature, rhol(ts)cells arrest primarily as small budded cells and undergo cell lysis; the nuclear cycle continues and a large fraction of the cells have replicated their DNA (Yamochi et al., 1994). The rhol(ts) arrest phenotype is similar to that of cells depleted of Pkclp (Levin et al., 1990; Levin and Bartlett-Heubusch, 1992) and, in fact, Rho l p physically binds Pkc l p and is required for its activity (Nonaka et al., 1996). Thus, one target of Rholp is Pkclp. Another is glucan synthase (Qadota et al., 1996). It is likely that other targets exist as well.

BUD SITE SELECTION GTPASE CYCLE

-

CMOKlNESlS

I

I RHO1 AND CDC42 GTPASE CYCLES

6. CDC42

POLARIZED GROWTH

Figure 4. Model of GTPase cycles that are important in budding (see text for details). The Rsrl p(Bud1 p) CTPase directs bud site selection. Rsrl p(Bud1p) may be present throughout the plasma membrane, become locally activated in the vicinity of the cytokinesis tag, and participate in targeting the growth effector to the cortex near the cytokinesis tag. The growth effector may be a Cdc42p effector and/or regulator such as Beml p or Cdc24p (Park et al., 1997). Binding of the growth effector at the cortex could be coupled to GTP hydrolysis on Rsrl p(Bud1p). The Cdc42p and Rholp GTPases function in polarized growth of the bud. Several different models of Cdc42p and Rhol p function can be imagined (see text). In the model shown here, cytoskeletal and/or growth components are assembled into complexes in the cytoplasm. Cytoskeletal components might include actin, actin-binding proteins, Beml p, and Spa2p; growth components could include proteins important for secretion. Complex assembly might be mediated by active, Rhol p-CTP. Alternatively Cdc4Zp-GTP might assemble Rhol p and other growth components, and Rhol p-CTP might be active at the cortex in recruiting actin cytoskeletal components and regulatingtheir assembly. Targeting of the complexes to the cortical growth effector is mediated by Cdc42p. In addition to these roles, Rhol p also serves as an activator of Pkcl p and glucan synthase (see text). The Sec4p CTPase functions in the late steps of secretion (not shown, see text for details) and i s polarized to the growth site directly or indirectly by the Rsrl p(Bud1p), Cdc42p, and Rhol p CTPases. 22

Cell Polarity in Yeast

23

Rho l p activity might be regulated in vivo by Bem2p GAP activity.Bem2p is important for bud formation (Bender and Pringle, 1991). Deletion mutants of BEM2 are inhibited for growth at h g h temperatures and at permissive temperatures hem2 mutants are often enlarged and multinucleate (Bender and Pringle, 1991). In vitro Bem2p activates GTP hydrolysis on Rholp (Y. Zheng, R. Cerione, and A. Bender, unpublished data, cited in Zheng et al., 1994) and consistent with Bem2p serving as a GAP for Rholp, hyperactivated rhol mutants resemble bem2-A mutants: cells are enlarged and exhibit bud formation defects (J. S. Johnson, A. M. Myers, M. McCaffrey, P. Boquet, and P. Madaule, manuscript in preparation, cited in McCaffrey et al., 1991). The carboxy terminus of Rho l p contains a consensus sequence for geranylgeranylation (Madaule et al., 1987; Clarke, 1992) and mutation of this sequence prevents proper localization of Rholp (Yamochi et al., 1994). Cdc43p is the presumptive GGT I for Rholp, and for the related protein, Rho2p (Qadota et al., 1992; Ohya et al., 1993). RHO2 is not required for cell growth (Madaule et al., 1987), and its function is unknown. It has been suggested that Rholp and Rho2p might directly or indirectly control Cdc43p/Ram2p activity through feedback regulation because multiple copies of RHOl and RHO2 are able to enhance Cdc43p/Ram2p GGT I activity on other substrates in vivo (Qadota et al., 1992). Rho3p and Rho4p Are Also Implicated in Budding. The Rho3 and Rho4 proteins are also implicated in bud growth and their functions are partially overlapping (Matsui and Toh-e, 1992a,b) rho3-A rho4-A double mutants are viable but fail to grow at high temperatures at which either single mutant can grow. Furthermore, the rho3-A mutant forms abnormally small colonies at semipermissive temperatures and overexpression of RHO4 restores rho3-A growth to approximately wild-type levels. As is true for other Rho proteins, Rho3p and Rho4p are likely to be prenylated. Their carboxy termini contain the motif, -C-A-A-M, characteristic of proteins which are farnesylated (Clarke, 1992; Matsui and Toh-e, 1992a). Like rhol(ts)cells (Yamochi et al., 1994), rho3-A rho4-Adouble mutants accumulate as small budded cells and lyse at restrictive temperatures for growth (Matsui and Toh-e, 1992b). This defect is partially suppressible by growth of cells in high osmolarity media (Matsui and Toh-e, 1992b). However, even under osmotic stabilization rho3-A rho4-A cells exhibit polarity defects. They become rounded lose chitin and actin polarity, and frequently become multinucleate (Matsui and Toh-e, 1993b). Overexpression of either of two cell polarity genes, CDC42 and BEMI, restores cell polarity and growth to rho3-A rho4-Acells (Matsui and Toh-e, 1992b). Thus Rho3p and Rho4p may function in polarized growth by interaction with growth components at the cell cortex. In spite of some phenotypic similarities between the rhol and rho3 rho4 mutants, neither overexpression of RHOl or RHO2 can rescue the growth defects of rho3-Aor of rho4-A cells (Matsui and Toh-e, 1992b, Y. Matsui, personal communication, cited in Matsui and Toh-e, 1992b). Thus, at least by this criteria, the functions of Rholp and Rho2p are nonoverlapping with Rho3p and Rho4p.

24

CHRISTINE COSTICAN and MICHAEL SNYDER

Summary of the CTPase Cycles that Are Important in Budding. In summary, at least four GTPase cycles are important for budding. As described above, the Cdc42p GTPase functions in polarization of components (Adams et al., 1990; Johnson and Pringle, 1990),the Rho1 p GTPase is implicated in both polarization of components and in bud growth (Madaule et al., 1987; Yamochi et al., 1994), and the Sec4p GTPase functions in bud growth (Novick et al., 1980; Salminen and Novick, 1987).A fourth cycle, the Rsrlp(Bud1p) GTPase cycle, functions in bud site selection (see below; Bender and Pringle, 1989; Chant and Herskowitz, 1991). The activities of these GTPases are controlled at a number of levels. They are all spatially restricted and accumulate in discrete subcellular domains: the sites of cell growth in the cases of Cdc42p (Ziman et al., 1993) and Rholp (Yamochi et al., 1994),late secretory vesicles and sites of cell growth in the case of Sec4p (Goud et al., 1988; Novick and Brennwald, 1993), and perhaps the entire cell surface in the case of Rsrlp(Bud1p) (see below). Localization, which is mediated in part by prenylation, has been demonstrated to be essential for GTPase function in all cases in which it has been examined. The transitions between guanine nucleotide bound states are obviously also important for GTPase function, and a number of potential regulators of GTP/GDP cycling have been identified for the various GTPases. One possible model for how these GTPases interact with each other and function in budding is as follows (Figure 4). The Rsrlp(Bud1p) GTPase establishes acortical marker to which Cdc42p targets cytoskeletal and/or growth components. Rholp promotes surface growth by activation of specific enzymes, Pkclp and glucan synthase, and perhaps by regulating actin assembly. Secretory components, including Sec4p, are polarized by the asymmetric distribution of the actin cytoskeleton and growth is consequently directed to the defined cortical site.

C. Pathway of Assembly of Components at the Incipient Bud Site Arrival of Components at the Bud Site Precedes Bud Emergence

Understanding the steps involved in polarized growth will require a description of the temporal and spatial order in which components assemble at the bud site. In synchronized cell populations, actin, Spa2p, and the putative neck filament proteins Cdc3p, CdclOp, Cdcl lp, and Cdcl2p all assemble at the presumptive bud site at approximately the same time, well in advance of budemergence (Snyder et al., 1991;Ford and Pringle, 1991;Kim et al., 1991; H. Kim, S. Ketcham, B. Haarer, and J. Pringle, personal communication). Cmdlp, Smy Ip, and Myo2p are reported to localize shortly after actin accumulation (Brockerhoff and Davis, 1992; Lillie and Brown, 1994). Although the data clearly indicate that significant assembly of growth components occurs prior to bud emergence, the relative order of assembly of components should be considered tentative because (1)scoring of actin patches as polarized is somewhat difficult in the early stages of budding and (2) different antibody sensitivities might influence the ability to detect early assembly of small amounts of growth components.

Cell Polarity in Yeast

25

The dependence of localization of Spa2p, the neck filament proteins, Cmdlp, Smy lp, and Myo2p upon actin, and vice versa, have been investigated using mutant strains. From these studies it is clear that localization of actin does not depend upon Spa2p or the neck filament proteins, and the localization of Spa2p and the neck filament proteins, Cdc3p and CdclOp, does not require actin (Adams and Pringle, 1984; Snyder et al., 1991; H. Kim, S. Ketcham, B. Haarer, and J. Pringle, personal communication). However, it remains possible that establishment of Spa2p and neck filament protein polarity depends upon actin, but once they arrive at the growth site their maintenance at that region is achieved by some other means. In the case of Cmdlp, Myo2p and actin, most actl cells exhibit delocalization of Cmdlp and Myo2p (Brockerhoff and Davis, 1992; Lillie and Brown, 1994); in most cmdl and my02 cells, actin is substantially depolarized (Johnston et al., 1991; Brockerhoff and Davis, 1992). Thus Cmdlp and Myo2p are interdependent with actin for proper localization. Interestingly, proper localization of Cmdlp is not required for actin polarization. Some separation-of-function mutations of Cmdlp abolish Cmdlp localization but do not affect actin polarity (Ohya and Botstein, 1994). Finally, Smy l p is delocalized in actl mutants but in smyl cells actin polarity is normal (Lillie and Brown, 1992; Lillie and Brown, 1994), indicating that actin controls Smy l p localization. These various dependency relationships will have to be investigated more thoroughly using synchronized ceI 1 populations in order to distinguish cytoskeletal requirements at specific times in the cell cycle. Furthermore, actin dependencies will have to be examined more systematically. In these studies several different actl mutants were used, and were examined under a variety of different conditions (e.g., differing lengths of time at the restrictive temperature for growth). It is likely that these variations yielded different forms and severities of actin cytoskeletal defects, and this may have contributed to the observed variations in actin dependencies. In contrast to their variable dependencies on ACTl, all polarized proteins which have been tested, Spa2p, Cdc3p and CdclOp, Cmdlp, and actin, are dependent on Cdc24p activity for proper localization (Adams and Pringle, 1984; Snyder et al., 1991; Sun et al., 1992; H. Kim, S. Ketcham, B. Haarer, and J. Pringle, personal communication). This suggests that Cdc42pKdc24p has a central role in directing polarization whereas the role of actin is to help mediate Cdc42p function. In summary, it is expected that this general strategy of double mutant analysis will be auseful one for understanding the pathway by which components assemble at the bud site. As additional proteins involved in bud emergence are identified, it should be possible to determine the steps by which bud site assembly occurs. Arrival of Components at the Bud Site Precedes Orientation of the SPB and Microt ubules

Initial electron microscopic studies of yeast containing nascent buds revealed that the microtubule organizing center of yeast, the spindle pole body (SPB) is ori-

26

CHRISTINE COSTIGAN and MICHAEL SNYDER

ented toward the newly forming bud and microtubules emanate toward the nascent bud (Byers and Goetsch, 1975; Byers, 1981). This observation led to the longstanding hypothesis that the SPB andor its associated microtubules were important for specifying the incipient bud site and organizing components at that site (Byers, 1981). In cells that undergo axial budding (see below), at the end of cytokinesis the SPB resides on the opposite side of the nucleus from the next site of bud formation. Thus, sometime during the unbudded phase of the cell cycle the SPB reorients 180” toward the incipient bud site (either by rotation of the nucleus, or by movement of the SPB through the nuclear envelope). This SPB reorientation is at least partially dependent upon microtubules; cells treated with the microtubule depolymerizing drug, nocodazole, frequently fail to reorient their SPBs (Jacobs et al., 1988). The chronology of growth component assembly at the incipient bud site relative to SPB reorientation and microtubule extension toward that site was determined using synchronized cells and immunofluorescence with anti-tubulin antibodies and with anti-Spa2p antibodies (as a marker for early assembly of components involved in budding). The SPB and microtubule polarization was found to occur after assembly of components at the incipient bud site (Figure 5; Snyder et al., 1991). Thus, in contrast to what was speculated (Byers, 1981), polarization of growth components is directed by component(s) at the cell cortex and not the other way around (Snyder et al., 1991; Madden et al., 1992; Flescher et al., 1993). Experiments in a variety of other eucaryotic cell types, most notably in Chaetopterus oocytes and C. elegans embryos (Lutz et al., 1988; Hyman, 1989), reveal that the regulation of SPBkentrosome positioning by cortical components is likely to be a widespread mechanism in eucaryotic nuclear division (reviewed in Strome, 1993). D. Bud Site Selection Budding Patterns

As noted above, the position of the bud is not random (see Figure 6). Haploid MATu and MATa cells undergo axial budding in which mother cells form buds adjacent to the previous bud site and daughter cells bud adjacent to the birth scar (i.e., proximal sites) (Freifelder, 1960; Hicks et al., 1977; Snyder, 1989). Diploid MATdMATa cells exhibit a bipolar pattern: mother cells usually bud at proximal sites, but daughter cells preferentially form buds opposite their birth scar (i.e., distal sites) (Freifelder, 1960; Hicks et al., 1977; Snyder, 1989). Diploid mother cells which have already budded many times, “old mothers,” show reduced fidelity of budding at proximal sites (Freifelder, 1960; Snyder, 1989; M. Snyder, unpublished) and bud at the secondary, distal sites. Seemingly contradictory reports, in which diploid mothers are described as budding with equal frequency from either pole (Chant and Herskowitz, 1991), are based on observations of populations in which new mothers and old mothers are not distinguished.

Cell Polarity in Yeast

27

ii( Microtubule

IiI,

ER

Actin-cortex

CORTCAL ROTATION

Figure 5. lmmunofluorescence experiments with anti-tubulin and anti-Spa2p antibodies reveal that the assembly of growth components at the incipient bud site precedes reorientation of the SPB. In the predominant pathway of assembly (shown on the left), (1)the Spa2p patch forms on the edge of the cell early in the unbudded phase, while the SPB still resides on the distal side of the nucleus, (2) next, a long microtubule bundle emanating from the distal SPB intersects the Spa2p patch, and (3) then the SPB orients so that it is proximal to the Spa2p patch and the microtubule bundle emanating from the SPB intersectsthe Spa2p patch. Two infrequently detected deviations from this pathway are shown on the right. First, early in the unbudded phase a small fraction of cells have two adjacent Spa2p patches. One patch is thought to represent residual material left from the previous site of cytokinesis and the second patch is believed to be material that accumulates at the incipient bud site. Thus, new material is sometimes deposited before the previous material from the site of cytokinesis has disappeared. Second, a few early GI cells have the SPB proximal to the incipient bud site without an intersecting microtubule bundle. Thus, sometimes the SPB orients prior to microtubule intersection. (Figure modified from Snyder et al., 1991.)

The difference between haploid and diploid budding patterns presumably reflects their different functions. Haploid budding patterns may have evolved to facilitate diploidization (Nasmyth, 1982), because diploid strains are expected to be more resistant to environmental stresses than haploid strains. S. cerevisiae strains

CHRISTINE COSTICAN and MICHAEL SNYDER

28

found in the wild are normally homothallic, that is, in the haploid state they readily switch their mating type. In homothallic haploid yeast the mother cell switches mating type after budding; the new daughter cell does not switch. Thus starting from a single haploid cell, axial budding and mating type switching produce a cluster of four cells, in which two pairs of cells of opposite mating type are positioned adjacent to one another. In Figure 6, M and D2 would be one mating type; D1 and D 1.1 the other mating type. This configuration is expected to promote efficient mating. Natural selection against selfing suggests that, if this model is correct, other mechanisms must exist that counterbalance the bias toward mother-daughter matings conferred by axial budding patterns when nonisogenic mating partners are available. Diploid budding patterns are hypothesized to facilitate foraging (Madden et al., 1992; Gimeno et al., 1992). The bipolar budding pattern allows the growing yeast colony to spread efficiently across a solid surface, and hence increase access to nutrients. Such spreading is particularly evident in diploid yeast growing as pseudohyphae (see below; Gimeno et al., 1992). Budding patterns are altered in response to environmental conditions. Haploid cells have been reported to bud at random or distal sites under glucose-limiting conditions (Thompson and Wheals, 1980), and preferentially bud distal to the previous bud site when exiting stationary phase (Madden and Snyder, 1992). We speculate that these alterations in the normal budding patterns are strategies for escaping environmental stress. For example, choice of random or distal sites in nutrient-starved haploid yeast cells might increase the chances of locating more favorable nutrient conditions. Similarly, upon osmotic stress diploid cells bud increasingly at distal

-

Haploid Proximal Sites

W D l .1

-

Diploid ProximaVDistal Sites

Figure 6. Yeast exhibit defined budding patterns. Haploid MATa and MATa cells undergo axial budding in which mother cells form buds adjacentto the previous bud site and daughter cells bud adjacent to the birth scar (i.e., proximal sites). Diploid MATaIMAJa cells exhibita bipolar pattern: Mother cells usually bud at proximal sites, but daughter cells preferentially form buds opposite their birth scar (i.e., distal sites). (Figure modified from Snyder, 1989.)

Cell Polarity in Yeast

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sites, and this has been proposed to be a mechanism that allows the growing colony to “move” away (Brewster and Gustin, 1994). A Cytokinesis Tag Directs New Bud Formation to a Site That is Proximal to the Previous Bud Site A cytokinesis tag has been postulated to determine the axial budding pattern (Snyder et al., 1991; Madden et al., 1992; Flescher et al., 1993). In this model, critical components from the previous site of cytokinesis nucleate assembly of components involved in bud formation at an adjacent site on the cortex (Snyderet al., 1991; Chant and Herskowitz, 1991; Madden et al., 1992). Since many components involved in the formation of a new bud are also present at the site of cytokinesis, these components can simply assemble on the cortex adjacent to the cytokinesis site. When the tag is lost or modified, for example by entry into stationary phase, budding occurs at the secondary, distal sites (Snyder et al., 1991; Madden et al., 1992; Flescher et a]., 1993). A number of lines of evidence are consistent with the hypothesis that a cytokinesis tag directs budding at proximal sites. First, the budding patterns themselves demonstrate that new buds form next to previous sites (Freifelder, 1960). Second, components at the site of cytokinesis persist into the next cell cycle and new components assemble next to them (Figure 5; Snyder et al., 1991; Kim et al., 1991). Third, as noted above, the SPB reorientation experiments demonstrate that polarity starts at the cortex, not at the SPB (Snyderet al., 1991). Finally, mutations in genes encoding the putative 10 nm neck filament proteins, Cdc3p, CdclOp, Cdcllp and Cdc12p, often cause buds to form at distal sites instead of proximal sites (Flescher et al., 1993). These latter data indicate that the putative neck filament proteins are important for bud site selection (Flescher et al., 1993). In diploid cells the frequency of cytokinesis tag-directed bud site selection is apparently lower. Diploid mother cells usually bud at proximal sites, but diploid daughter cells bud at distal sites. Diploid daughter cells may bud at distal sites because they have a longer G1 than mother cells; this prolonged period is expected to result in a greater chance of losing (or modifying) the tag or growth components at the neck (Snyder et al., 1991;Madden et al., 1992; Flescher et al., 1993). Consistent with this possibility, old mothers exhibit a reduced fidelity of axial budding compared to new mothers (Freifelder, 1960;Snyder, 1989; M. Snyder, unpublished) and GI is longer in old mothers compared to new mothers (Egilmez and Jazwinski, 1989; Flescher et al., 1993). Although a similar difference in G1 duration exists between haploid mothers and daughters, haploid daughters usually bud at proximal sites. It is possible that the difference in fidelity of proximal budding in haploids and diploids is due to differences in the cytokinesis tag. In haploid cells, modifications (or loss of modifications) in structures at the neck might make the tag a high affinity site for assembly of new components; these modifications may not be present in diploid cells. Such modifications could be either the presence ofproteins at the neck

30

CHRISTINE COSTIGAN and MICHAEL SNYDER

region in haploid cells that are not present in diploid cells (Chant and Herskowitz, 1991) or modifications of existing proteins. Selection of Sites which are Distal to the Previous Bud Site Several observations suggest that in the absence of the cytokinesis tag there are secondary sites at the distal pole. Diploid daughters preferentially bud at a site that is distal to the previous bud site (Freifelder, 1960; Hicks et al., 1977; Snyder, 1989). In addition, in certain axial budding mutants, bud3 and bud4, the ratios of distal budding to proximal budding are >1.2 in both haploid mothers and daughters (Chant and Herskowitz, 1991). Finally, proteins usually found polarized to cortical growth sites are absent or delocalized in haploid strains which have entered stationary phase (Snyder et al., 1991;M. Snyder, unpublished results), and these cells preferentially choose distal sites after dilution into fresh medium (Madden and Snyder, 1992). This latter observation also indicates that the proximal sites are transient. Two mechanisms that might direct distal budding have been proposed. Distal budding might be directed by proximity to the SPB (Snyder et al., 1991; Drubin, 1991; Madden et al., 1992; Madden and Snyder, 1992). After cytokinesis the SPB lies on the opposite side of the nucleus from the previous bud site and it is possible that microtubules or other SPB-associated components could organize assembly of growth components at the cortex near the SPB (Byers, 1981; Drubin, 1991; Snyder et al., 1991; Madden et al., 1992; Madden and Snyder, 1992). Treatment with nocodazole does not prevent preferential distal budding of daughter diploid cells, suggesting that microtubules are not required for choosing this site (Jacobs et al., 1988; Drubin, 1991). However, these experiments are not definitive because the sites may be marked by the SPB and/or its associated microtubules prior to nocodazole treatment. Alternatively, distal budding might be driven by the presence of growth components remaining from previous cell surface growth events (Chant and Herskowitz, 1991). Such components might begin assembling a new bud or could be recognized by the bud formation machinery as targets for assembling a new bud at the distal site. Although this mechanism may be important for choice of distal bud sites in daughters, it is not likely to explain preferential distal budding of mother cells, for example in cells reentering the cell cycle from stationary phase (Madden and Snyder, 1992). SPB-directed bud site selection or some other mechanism may direct distal budding in these cells. Finally, it is possible that other mechanisms might also contribute to bud site selection. For example, the poles could be more susceptible to turgor pressure than the other surfaces of the cell because of subtle differences in cell wall architecture. Proteins which Might Be important for the Cytokinesis Tag Several components have been identified which may directly or indirectly form the cytokinesis tag. Cdc3p, Cdc lop, and Cdc 1 1p are present at the neck at cytokine-

Cell Polarity in Yeast

31

sis and remain at the cortex into G1 of the next cell cycle (Haarer and Pringle, 1987; Ford and Pringle, 1991; Kim et al., 1991; Kim, Haarer, and Pringle, personal communication). Their localizations and involvement in axial budding make the neck filament proteins good candidates for components of, or proteins which interact with, the cytokinesis tag (Flescher et al., 1993). Other proteins important in axial budding include Bud3p and Bud4p (Chant and Herskowitz, 1991). In haploid bud3 and bud4 mutants, both mother cells and daughter cells frequently form buds at distal sites (Chant andHerskowitz, 1991). Bud3p and Bud4p have recently been localized to the neck at cytokinesis (J. Chant), suggesting that they are also part of the cytokinesis tag. A CTPase Cycle is lrnportant for Bud Site Selection

Mutants which exhibit random budding patterns are predicted to be defective in targeting bud formation proteins to one of the poles of the cell and/or in assembling bud formation components at the bud site (Flescher et a]., 1993). These defects might lead to the formation of aberrant complexes which localize randomly to initiate budding (Snyder et al., 1991; Madden et al., 1992). Selection of bud sites is likely to be a guanine nucleotide-regulated event mediated by Rsrlp(Budlp), BudSp, and Bud2p. Mutants of RSRl ( B U D ] ) ,BUD2, or BUD5 exhibit random budding, and bud scars are scattered over the entire cell surface, not just at the poles (Bender and Pringle, 1989; Chant et al., 1991; Chant and Herskowitz, 1991; Park et al., 1993). Moreover distal/proximal pole budding in bud3 and bud4 mutants requires the functions of Rsrlp(Bud1p) and Bud2p (Chant and Herskowitz, 1991). The predicted protein sequences of Rsrlp(Budlp), BudSp, and Bud2p are that of a Ras-type GTPase of the rap1 family (Bender and Pringle, 1989; Hall, 1990), a GDP dissociation stimulator (Chant et al., 1991), and a GAP (Park et al., 1993), respectively. These proteins are predicted to directly interact in vivo. Mutant analysis of Rsrlp(Bud1p) indicates that its ability to bind and hydrolyze GTP is essential for its function (Ruggieri et al., 1992). BudSp is likely to act as a GDP dissociation stimulator for Rsrlp(Bud1p): genetic tests demonstrate that Rsrlp(Bud1p) function requires BudSp, and this requirement is diminished for a mutationally activated form of Rsrlp(Bud1p) (Bender, 1993). Bud2p is a GAP for Rsrlp(Bud1p) in vitro (Park et al., 1993), and genetic interactions between BUD2 and RSRl (BUD1) suggest that Bud2p is a negative regulator of Rsrlp(Bud1p) in vivo (Bender, 1993). Thus BudSp and Bud2p modulate the guanine nucleotide bound state of Rsrlp(Bud1p) and are required forRsrlp(Bud1p) function i n budsite selection. In mammalian neutrophils, the Rsrlp(Bud1p) homolog, Rapl, is associated with components of the superoxide-generating system which is locally activated at the plasma membrane (Quinn et a]., 1989). Thus other members of the Rsrl(Bud1p) family are also implicated i n processes that are polarized within the cell.

32

CHRISTINE COSTICAN and MICHAEL SNYDER

Rsrlp(Bud1p) may function in targeting growth components/regulators to specific cortical sites, for example to the site marked by the cytokinesis tag in the case of proximal budding (see Figure 4). Possible components targeted by Rsrlp(Bud1p) include regulators of Cdc42p or other polarized growth components (Park et al., 1997). Rsrlp(Bud1p) localizes in a punctate pattern over the entire cell surface (Michelitch and Chant, 1996). The localizations of BudSp and Bud2p have not been reported, although Bud2p is present in cell membrane fractions (Park et al., 1993). A possible model for how Rsrlp(Bud1p) functions in targeting is as follows (see Figure 4 legend for other possibilities). Rsrlp(Bud1p) might be locally activated at specific cortical sites either because Bud2p and/or BudSp is localized at the cytokinesis tag or locally activated at that site. Activated Rsrlp(Bud1p)-GTP may recruit a growth effector from the cytoplasm, for example, a Cdc42p regulator or even Cdc42p. Alternatively, Rsrl p(Bud lp) might regulate the targeting of the effector molecule. Docking of the growth effector at the cortex is coupled to Bud2p-stimulated GTP hydrolysis on Rsrlp(Bud1p). GTP hydrolysis on Rsrlp(Bud lp) might be required for attachment of the growth effector at the cortex and/or for recycling of Rsrlp(Bud1p) and another round of effector targeting. Bud Emergence and Cytoskeletal Genes which Function in Bud Site Selection

Several genes which are important for bud emergence may also play roles in bud site selection, as would be expected for such closely related processes. Two different cdc24 mutants, cdc24-3 and cdc244, and a bem2 mutant exhibit random budding patterns in both haploids and homozygous diploids (Sloat et al., 1981; Yi-Jun Sheu, K. Madden, and M. Snyder, unpublished data). Overexpression of CDC42 causes random budding in both haploids and diploids (Johnson and Pringle, 1990). The budding pattern defects of these various mutants may reflect a direct role for the wild-type gene in bud site selection, for example as part of a complex that is targeted to bud sites by Rsrlp(Bud1p). Consistent with the hypothesis that bud emergence components interact with bud site selection components is the observation that these components interact genetically. The ts growth defect of a cdc24 strain are partially suppressed by multiple copies of RSRl(BUD1) (Bender and Pringle, 1989), andmutations inBUD5slightly enhance the growth defects of beml mutants (Chant et al., 1991). Aberrations in actin cytoskeletal proteins and other potential cortical proteins can also influence budding patterns. Actin hemizygotes (MATdMATa A C T l h c t l -A) and homozygous diploids of certain act1 alleles (MATdMATa actl/actl) contain bud scars randomly distributed on their surfaces (Drubin et al., 1993). Similarly, the PFYl and CAP(SRV2) genes are important for bud site selection at all sites (Cap(Srv2p): adenylyl cyclase-associated protein is described below; Haarer et al., 1990; Vojtek et al., 1991). spa2 mutants show reduced fidelity of axial budding, par-

Cell Polarity in Yeast

33

ticularly in diploid mother cells (Snyder, 1989). Diploid rvs167 cells bud randomly whereas haploid rvs167 cells exhibit normal axial budding (Bauer et al., 1993; described further below). Finally, overexpression of Abp 1 protein causes haploid cells to lose the fidelity of axial budding and bud scars are seen clustered at both poles of the cells (Drubin et al., 1988). Thus, a variety of bud formation components are implicated in bud site selection. As noted above, one possible explanation for how each of these mutations/overexpression constructs affect bud site selection is that they disrupt targeting or assembly of growth components at the bud site. Some of these are present at the neck at cytokinesis, such as Spa2p and actin (Adams and Pringle, 1984; Kilmartin and Adams, 1984; Snyder, 1989; Snyder et al., 1991), and could potentially also affect the cytokinesis tag.

E.

Coordination of Bud Initiation with Cell Cycle and Growth Control

Bud emergence depends on the execution of “Start,” and budding is therefore coordinated with growth control (Pringle and Hartwell, 198 1). Start is the step in G 1 at which the cell commits to a new cell cycle (Pringle and Hartwell, 198 1 ; Bartlett and Nurse, 1990). For execution of Start the cell must have attained a minimum critical size and there must be sufficient nutrients available (Johnston et al., 1977; Pringle and Hartwell, 198 1 ; Bartlett and Nurse, 1990). Cdc28p, in conjunction with the GI cyclins, Cln Ip, Cln2p, and Cln3p, mediates progression through Start (Pringle and Hartwell, 1981; Richardson et al., 1989; Bartlett andNurse, 1990). As described in detail below, Cdc28p-Clnp participates in bud morphogenesis by contributing to polarization in G1 and carrying out an essential function in formation of the bud neck. Cdc28p-Clnp, Functions in Bud Morphogenesis

Cdc28p-Clnp is implicated in polarity establishment in GI. At the restrictive temperature, cdc28(ts) mutants fail to form buds and the establishment of actin polarization in G1 is activated by Cdc28p and Cln proteins (Pringle and Hartwell, 198 1; Lew and Reed, 1993). In addition, CLNl and CLN2 interact genetically with the bud site selection genes. Introduction of a BUD2 mutation into clnl cln2 cells results in an accumulation of large, round, unbudded multinucleate cells and temperature-dependent inviability (Benton et al., 1993; Cvrckova and Nasmyth, 1993). The bud2 clizl cln2 inviability is alleviated by mutating RSRl(BUDI), suggesting that inviability might result from active, Rsrlp(Bud 1p)-GTP titrating out components required for budding (Benton et al., 1993). In agreement with this, the bud2 clnl cln2 mutant is partially rescued by introduction of a multicopy plasmid carrying CDC42 or by a centromeric plasmid carrying BEMl (Benton et al., 1993). These observed genetic interactions suggest that Cdc28p-Clnlp and Cdc28-Cln2p directly or indirectly participate in the establishment of polarized growth; in the ab-

34

CHRISTINE COSTICAN and MICHAEL SNYDER

sence of this regulation polarized growth is less efficient, thus titration of Cdc42p and Bemlp activity by hyperactivated Rsrlp(Bud1p) abolishes the cell’s competence for bud emergence. A CDC28KLN-independent pathway for polarity establishment also exists (Snyder et al., 1991; Madden et al., 1992; Lew and Reed, 1993). Independent pathways are suggested by the G1 arrest phenotype of both cdc28 mutants and clnl cln2 cln3 mutants, in which polarized growth is delayed, but does occur, resulting in cells with an elongated shmoo-like morphology (Pringle and Hartwell, 1981; Richardson et al., 1989; Lew and Reed, 1993; shmoos are described below). The normal sites of cell growth are utilized in cdc28 cells: haploid cells form projections proximal to the previous bud site (Madden and Snyder, 1992). Furthermore, growth components are polarized: Spa2p and actin are present at the projection tips (Snyder et al., 1991; Madden and Snyder, 1992; Lew and Reed, 1993). The delay in polarization of cdc28 mutants at the restrictive temperature cannot be shortened by overexpression of cyclins, thus it is unlikely that the polarized growth reflects residual function of the mutant Cdc28p at the restrictive temperature (Lew and Reed, 1993). Cdc28p-Clnp also has an essential role in neck assembly in G1, as suggested by the absence of constrictions in G1-arrested cdc28 and clnl cln2 cln3 mutants which still polarize (Pringle and Hartwell, 1981; Snyder et al., 1991; Madden and Snyder, 1992; Lew and Reed, 1993). Furthermore, a number of mutants isolated as synthetic lethal loci with clnl cln2 exhibit cytokinesis defects in the clnl cln2 background (Benton et al., 1993; Cvrckova and Nasmyth, 1993). Perhaps Cdc28p-Clnp directly or indirectly regulates Cdc3p, CdclOp, Cdcl Ip, Cdcl2p and/or other proteins at the neck, and thus mediates proper assembly of the neck cytoskeleton for the nascent bud. The Slt2p MAPK Pathway May Direct Cdc28p-Clnp-independent Polarized Growth

A candidate CDC28-independent pathway underlying polarized growth is the SLT2(MPKI) MAPK (mitogen-activated protein kinase) pathway. The SLT2(MPKI) MAPK pathway is composed of an array of protein kinases, which by genetic tests appear to function in the following order: Pkclp (protein kinase C) -++Slklp(Bck1p) -+ Mkkp -+ Slt2p(Mpklp) (Levin et al., 1990; Torres et al., 1991; Costigan et al., 1992; Lee and Levin, 1992; Levin and BartlettHeubusch, 1992; Paravicini et al., 1992; Irie et al., 1993; Lee et al., 1993; Levin andErrede, 1993;Mazzoni et al., 1993;Costigan et al., 1994).The growthdefects of cdc28(rs) strains that arrest at Start are enhanced by mutation of SLT2 (Mazzoni et al., 1993). The role of the SLT2 pathway in polarized growth and budding is indicated by several lines of evidence: pkcl(ts) mutants, like rhol(rs) mutants, arrest at the restrictive temperature with small buds (Levin et al., 1990; Levin and BartlettHeubusch, 1992; Paravicini et al., 1992; Yamochi et al., 1994); electron micro-

Cell Polarity 117 Yeast

35

scopic analysis indicates that these cells have defective cell walls (Paravicini et al., 1992; Levin and Errede, 1993). A variety of polarity defects have been observed in slkl-A and slt2-A mutants. For example slkl-A and slt2-A mutants are inviable in combination with a deletion of the SPA2 polarity gene, have delocalized actin distributions, accumulate secretory vesicles, and many cells are unusually small or have aberrant shapes (Costigan et al., 1992; Mazzoni et al., 1993; Costigan et al., 1994). One mechanism by which the Slt2p pathway regulates polarized cell growth is through phosphorylation of SBFfMaddenet al., 1997). SBFis a transcription factor complex that is important for transcription of G1 cyclins (which in turn promote polarized growth) and cell wall synthesis genes. Thus, a likely manner in which the Slt2p pathway functions is to activate SBF, which in turn leads to increased expression of G1 cylins and cell wall synthesis components that activate polarized cell growth (Madden et al., 1997). The SLT2 MAPKpathway is also important for nutrient sensing and growth control (Costigan et al., 1992; Mazzoni et al., 1993; Costigan et al., 1994; Costigan and Snyder, 1994); after incubation in starvation medium, slkl-A and d t 2 - A mutants fail to enter stationary phase as assessed by a variety of criteria. Thus, the SLT2 MAPK pathway appears to be a common regulatory pathway for both morphogenesis and growth control. General Mechanisms Linking Growth Control and Morphogenesis

In addition to components of the SLT2 pathway, several other genes have been identified that affect both morphogenesis and growth control. The spu2-A mutant was the first example of such a mutant to be identified. In addition to polarity defects, the spu2-A mutant exhibits a mild defect in cell cycle arrest (see above; Snyder, 1989). Rvs167p is another candidate cytoskeletal component which also is important for growth control (Bauer et al., 1993). The predicted Rvs167p protein sequence contains an SH3 domain, and cells lacking Rvs167p function exhibit morphogenic and cytoskeletal defects upon osmotic stress or starvation. rvs167 cells are also defective in nutrient sensing and in cell cycle arrest. Cap(Srv2p), adenylate cyclase-associated protein, provides one of the best characterized example of a protein with dual functions in growth control and morphogenesis. cup(srv2) mutants exhibit nutrient sensing defects and Cap(Srv2p) interacts with the RaskAMP-dependent protein kinase pathway: it can modulate Ras activation of adenylate cyclase and is detected in a complex with adenylate cyclase (Field et al., 1990; Fedor-Chaiken et al., 1990; Wang et al., 1992). cup(srv2) mutants also have cell polarity defects, including aberrant morphologies, actin depolarization, and loss of budding pattern fidelity (Field et al., 1990; Vojtek et al., 1991). Finally, CAP(SRV2) genetically interacts with PFYl (Vojtek et al., 1991). There are two possible mechanisms by which these components might affect both morphogenesis and nutrient sensing. First, nutrient signalling and the cellular response to starvation might be particularly sensitive to perturbations of a protein’s

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function in morphogenesis. The cell surface is both the site of important events in polarized growth and the interface at which the cell receives and responds to signals from the environment. A variety of nutrient transporters and permeases are found at the cell surface, as well as components of the RaskAMP-dependent protein kinase pathway, a key regulatory pathway in yeast nutrient sensing (Broach, 1991). The proper organization of these components in the membrane of the emergent bud is likely to be critical for nutrient sensing. Mutations that alter the membrane cytoskeleton in the growth region might affect the organization and/or proper functioning of transporters or signalling components. Alternatively, components implicated in both morphogenesis and nutrient sensing might function in processes that actively coordinate growth signals with morphological events. It is expected that changes in the growth state of the cell will be transmitted into morphological changes by modification of cytoskeletal and membrane components at the cell perimeter. By analogy, in many mammalian cell types signalling for proliferation or differentiation, for example by src family protein kineses, is accompanied by modification of components at the cell perimeter (Cooper, 1990). Calcium Regulation of Budding

Another candidate regulatory mechanism for coordination of polarized growth with the cell cycle and growth control is through Ca++-regulatedchanges in protein activities. Ca++ uptake increases for a short period at the start of budding (Saavedra-Molina et al., 1983), and intracellular Ca++levels are likely to regulate events in G I . Exponentially growing cells depleted of Ca++by simultaneous addition of a Ca++ionophore and a Ca++chelator transiently arrest in G I (Iida et al., 1990a,b). This Ca++depletion apparently does not have a general effect on growth as the rate of protein synthesis is the same in Ca++-depletedand untreated cells, consistent with Ca++playing a regulatory role in G1 (Iida et al., 1990a,b). One budding component that might be regulated by Ca++fluxes is Cdc24p. The predicted protein sequence of Cdc24p contains two putative Ca++-bindingdomains (Miyamoto et al., 1987) and strains with aparticular allele of CDC24 do not grow in medium containing high CaCI, concentrations (Ohya et al., 1986a,b). Ca++may also regulate Pkc I p activity, and perhaps the Slt2p MAPK pathway. The growth defects of conditional pkcl alleles are suppressible by exogenous Ca++, suggesting that Ca++stimulates Pkclp (Levin and Bartlett-Heubusch, 1992).

F.

Other Signalling Mechanisms Underlie the Maintenance of Polarity

Growth lnhibitors and Environmental Factors Affect the Maintenance of Cell Polarity

Cell polarity and morphogenesis are affected by the growth state of the cell. Treatment of cells with general growth inhibitors such as the protein synthesis in-

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hibitor, cycloheximide, and the mitochondria1 respiration inhibitor, sodium azide, cause negative effects on integrity of the actin cytoskeleton (Novick et al., 1989). Environmental stresses such as heat shock and osmotic shock cause a transient growth arrest (Johnston and Singer, 1980; Chowdhury et al., 1992); and these stresses also cause transient actin depolarization (Chowdhury et al., 1992; Palmer et al., 1992). For the case of osmotic shock, the transient growth arrest is approximately temporally correlated with the transient actin depolarization (Chowdhury et al., 1992). Hoglp MAPK Pathway The HOG1 MAPK pathway, which is required for growth on high osmolarity media, is involved in maintenance of cell polarity (Brewsteret al., 1993; Brewster and Gustin, 1994). PBS2(HOG4) encodes a MAPK kinase homolog which under conditions of high osmolarity is believed to activate, Hoglp, a MAPK (Brewster et al., 1993). Polarity defects are observed in hog mutants upon transfer from permissive to restrictive osmolarity (Brewster and Gustin, 1994). As is observed in wild-type cells (Chowdhury et al., 1992), actin polarity is lost: actin cables disappear and in budded cells actin patches become delocalized throughout the mother cell and bud. However, unlike wild-type cells, hog mutants do not recover actin polarity in the bud and reinitiate bud growth. Rather, the mechanism (which is likely to be based on information at the cortex) for reestablishment of polarity in osmotically shocked wild-type cells is lost or absent in hog mutants. Upon osmotic shock hog mutants reinitiate growth at a new site on the mother cell. Interestingly, the fidelity of budding patterns is lost in selection of this new site and budding occurs at random sites in haploid cells. Possible structural protein substrates for the Hoglp pathway could include Pfylp, Caplp, Cap2p, Rah3p, Rvs167p, and even actin, all of which have been shown to be important for growth on high osmolarity media (Novick and Botstein, 1985; Haarer et al., 1990; Amatrudaet al., 1992; Chowdhury etal., 1992; Bauer et al., 1993). Interestingly, mutations in RSRI/BUDl, BUD2, BUD3, BUD4, and BUD5 do not cause defects in reinitiation of polarized growth on high osmolarity medium (Brewster and Gustin, 1994). Thus although the Bud proteins are important for establishing growth sites (Bender and Pringle, 1989; Chant et al., 1991;Chant and Herskowitz, 1991), they are not essential for maintaining them under osmotic stress.

C . Segregation of Organelles into the Newly Formed Bud After bud formation, organelles which reside in the mother cell must be properly partitioned into the bud. Mitochondria and the vacuole probably begin to appear in the bud after the secretory vesicles but prior to, or approximately coincident with, the time when the bud is about one half the size of the mother cell (Weisman et al., 1987; Stevens, 1981). The nucleus is segregated into the bud only after the bud is

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near or at its maximum size (Pringle and Hartwell, 1981). Several different cytoskeletal elements are important in organelle segregation.

Segregation of the Nucleus The replicated nucleus is segregated into the bud via cytoplasmic microtubules, and several steps are involved in this process (Figure 7; reviewed in Page and Snyder, 1993).As described above, prior to bud formation the SPB orients toward the

Figure 7. The replicated nucleus is segregated into the bud via cytoplasmic microtubules. In C1 the SPB orients toward the incipient bud site and cytoplasmic microtubules emanating from the SPB intersect this region. The SPB may initiate duplication prior to (shown on the right) or after (shown on the left) orientation. Sometime in late S or C2 a short spindle i s set up, and the nucleus migrates from the mother cell to the neck of the bud. Upon nuclear division, the newly formed SPB is segregated into the bud (Vallen et al., 1992). Cytoplasmic microtubules are thoughtto be important for SPB orientation toward the incipient bud site, migration of the nucleus to the neck, and positioning of the dividing nucleus within the budding cell. (Figure reproduced, with permission, from Page and Snyder, 1993.1

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incipient bud site and cytoplasmic microtubules emanating from the SPB intersect this region (Snyder et al., 1991). After the GUS transition, the SPB duplicates 1 (Peterson and Ris, 1976). Sometime later in late S or G2 a short spindle is set up and the nucleus migrates from the mother cell to the neck of the bud (Peterson and Ris, 1976; Kilmartin and Adams, 1984). When the bud has neared or reached its maximum size, the spindle elongates to the full length of the cell and the nucleus divides between the mother cell and the bud (Peterson and Ris, 1976; Kilmartin and Adams, 1984). The yeast nucleus remains intact during mitosis. Cytoplasmic microtubules are thought to be important for SPB orientation toward the incipient bud site, migration of the nucleus to the neck, and positioning of the dividing nucleus within the dumbbell-shaped cell (reviewed in Page and Snyder, 1993). Cells treated with nocodazole and rub2(ts) mutants incubated at the restrictive temperature fail to orient their SPB toward the bud and do not migrate their nucleus toward the neck (Huffaker et al., 1988; Jacobs et al., 1988). In addition a tub2(ts)mutant which specifically lacks cytoplasmic microtubules at the restrictive temperature, fails to position the spindle apparatus (and nucleus) between the mother cell and the bud, and often undergoes nuclear division in either the mother cell or the bud (Palmer et al., 1992; Sullivan and Huffaker, 1992). Microtubule capture sites at the cell cortex are thought to interact with the ends of cytoplasmic microtubules, thereby stabilizing them and maintaining the orientation of the SPB and spindle apparatus. Many late G1 cells contain a long microtubule bundle emanating from the SPB toward the vicinity of the incipient bud site, intersecting the cortex in that region (Snyder et al., 1991). Because microtubules of other organisms are unstable both in vivo and in vitro (e.g., Caplow, 1992), these long microtubule bundles are proposed to be stabilized by microtubule capture sites at the cortex (Snyder et al., 1991). Capture sites in the bud and mother cell would be important for interacting with the cytoplasmic microtubules emanating from the SPBs and thus for orienting and maintaining the spindle position between the mother cell and bud. Actin is likely to directly or indirectly contribute to establishing the putative microtubule capture site because spindle orientation is lost in actl(ts) mutants and the cells become multinucleate (Palmer et al., 1992; Sullivan and Huffaker, 1992; Drubin et al., 1993). Segregation of the Mitochondria and the Vacuole

In addition to secretory organelles, the newly emergent bud contains mitochondria and a vacuole (Stevens, 1981; Weisman et al., 1987). In contrast to their essential role in nuclear segregation, microtubules are not required for vacuolar or mitochondrial segregation (Huffaker et al., 1988; Jacobs eta]., 1988; Weisman and Wickner, 1988). Several mutants that affect mitochondrial segregation have been identified. One of these carries a mutation in the M D M l gene, which encodes a yeast intermediate filament protein (McConnell et al., 1990; McConnell and Yaffe, 1992; McConnell

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and Yaffe, 1993). Mdml mutants fail to segregate mitochondria into the bud. Nuclear segregation also fails in these mutants, although cytoplasmic microtubules extend into the bud and the nuclei do divide (McConnell et al., 1990). As noted above, mitochondria1 organization may also be controlled in part by the actin cytoskeleton (Drubin et al., 1993). Once buds have reached half the mother cell diameter they nearly always contain a vacuole (Weisman et al., 1987). As the bud grows, the bud vacuole enlarges and accumulates vacuolar components from the mother cell vacuole (Weisman et al., 1987). Tracks containing mother cell vacuole contents are detected between the bud vacuole and the mother cell vacuole, suggesting that the bud vacuole is generated by vesicle transport from the mother cell vacuole and/or by a continuous network of vacuolar structure extending from the mother cell to the bud (Weisman and Wickner, 1988; Raymond et al., 1990). Although the detailed mechanisms governing vacuolar segregation have yet to be elucidated, many genes important for proper vacuolar inheritance have been identified (Herman and Em, 1990; Raymond et al., 1990; Weisman et al., 1990; Raymondet al., 1992; Weisman and Wickner, 1992). In cases where it has been examined, these mutants do not exhibit defects in nuclear or mitochondria segregation, indicating that the vacuolar segregation mechanism is at least partially independent from the mechanisms of segregation of these other organelles.

H. Speculations about Bud Formation and Growth Two fundamental questions concerning budding remain to be addressed. First, how do cells form one and only one bud in each division cycle? Second, how does a cell form buds of uniform size? Formation of a single bud many involve the assembly of “precomplexes.” We speculate that a singleprecomplex is formed which is then is then targeted to the cortical site (the cytokinesis tag), presumably by Budlp. Preferential assembly of growth components into a precomplex which is targeted to the growth site rather than directly onto the cortical tag itself (which could be very large) would ensure that only one bud is formed. Mutants that disrupt the precomplex might form multiple buds. In this respect, we note that a mutant containing a particular spa2 allele, spa2-7, occasionally forms multiple buds upon entry and sometimes exit from stationary phase (Snyder, 1989). It is possible that the precomplexes are not formed properly in this mutant, and under conditions where cell polarity is reduced (e.g., loss of the cytokinesis tag), multiple buds form from several defective precomplexes. In vegetative cells, the bud size is relatively uniform, and interestingly, in cells such as cdc4 mutants which form multiple elongated buds (Hartwelt et al., I973), the buds are generally also of uniform size (albeit larger than vegetative cells, perhaps through the prolonged activation of the Clns (Lew andReed, 1993). Hence we propose that aregulatory mechanism must exist for monitoring bud size. Onepossible mechanism to explain uniform bud size is that an intracellular gradient is moni-

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tored. Signals from the tip of the bud (perhaps from ion or nutrient channels) may be sensed in the mother portion of the budded cell. When the signal receivied is weak, either the cell reduces growth (as occurs in nuclear division cdc and tub mutants) or the Cdc28p cell cycle machinery signals the onset of cytokinesis, assuming that nuclear division steps are completed. Greater understanding of each of these issues (single buds and uniform bud size) will require additional studies.

I.

Summary of the Budding Process

In summary, a variety of steps in the budding process have been elucidated and many structural and regulatory components involved in this process have been identified. Bud site selection involves a cytokinesis tag for bud formation at proximal sites and secondary mechanisms for distal sites. Polarization of actin and the secretory apparatus is important for bud formation and growth, and a number of regulators of the actin cytoskeleton have been identified. At least four GTPase cycles participate in various aspects of the budding process. The Rsrl p(Bud1p) GTPase functions in bud site selection (Bender and Pringle, 1989; Chant and Herskowitz, 1991), the Cdc42p GTPase functions in polarization of growth (Adams et al., 1990; Johnson and Pringle, 1990), the Rholp GTPase is also implicated in both polarization of cytoskeletal components and in bud growth (Madaule et al., 1987; Yamochi et al., 1994), and the Sec4p GTPase functions in late steps of secretion that are required for growth (Novick et al., 1980; Salminen and Novick, 1987). A number of regulators of these GTP-binding protein have been identified. Finally various candidates for regulators of bud formation and growth have been identified, including the Cdc28-cyclin complex, Pkc l p and the Slt2p MAPK pathway, Ca2+ fluxes, and the Hoglp MAPK pathway. There are many aspects of the budding process which remain to be clarified. For most known proteins, their specific functions are not well understood, and there are presumably many components yet to be identified. The mechanism(s) by which different proteins target to the growth site and then assemble to mediate bud formation and growth is not known. Finally, the controls that regulate execution of the different steps in budding, including the onset of bud formation and termination of bud growth, have not been defined.

111.

PSEUDOHYPHAL GROWTH

Under appropriate inducing conditions, for example nitrogen limitation, S. cerevisiae undergoes a transition to a specialized type of budding in which it forms pseudohyphae (Scherr and Weaver, 1953; Brown and Hough, 1965; Gimeno et al., 1992; Gimeno et al., 1993). Pseudohyphal growth requires both polarized cell divisions (Gimeno et al., 1992) and polarized cell growth. It is distinguished from “sated” vegetative growth (i.e., growth in favorable environmental conditions) by the for-

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mation of elongated cells which often remain attached after cytokinesis and thus form long chains (pseudohyphae) and by the ability to invade solid media, suggesting that the cells secrete degradative enzymes (Gimeno et al., 1992; Gimeno et al., 1993). By analogy with vegetative budding, commitment to pseudohyphal growth probably occurs at Start, although this has not been shown. Pseudohyphal growth has a number of advantages for yeast. First, it increases the surface to volume ratio of the cell, thus expanding the surface area for nutrient uptake from a nutrient-limited environment. Second, it provides a means of efficiently spreading, increasing the chance of contacting a better nutrient source. A.

Polarized Growth and Divisions in Pseudohyphal Cells

Pseudohyphal cells divide by budding and growth is at least partially polarized to one pole of the cell as the bud grows. The details of polarized growth in pseudohyphal cells have not been described, and the roles of only a few well-characterized cell polarity genes, for example SPAZ and SLAZ, have not been reported (Yang et al., 1997; Roemer et al., 1998). However it is known that polarized divisions, specifically bipolar budding patterns, are critical for pseudohyphal growth. Diploid cells that express the dominant negative mutant form of RSRl(BUD1) bud randomly and are unable to grow as pseudohyphae (Bender and Pringle, 1989; Chant and Herskowitz, 1991; Gimeno et al., 1992; Ruggieri et al., 1992). A unique aspect of pseudohyphal budding is the frequent exhibition of apical dominance, in which daughter cells divide to form pseudohyphal cells whereas mother cells stop dividing or divide to produce cells typical of (sated) vegetative growth (Gimeno et al., 1992; Gimeno et al., 1993). The effect of apical dominance is to suppress branching of the pseudohypha. Its origin may be genetic, or alternatively, it may be determined in part by nutrient availability; cells furthest from the apex of the hypha are in a nutrient-depleted environment and thus do not bud. It has been suggested that pseudohyphal growth is a mechanism to efficiently deliver vegetative cells to new colonization sites, because cells which are distant from the apex in the chain often bud to form cells typical of (sated) vegetative growth (Gimeno et al., 1992; Gimeno et a]., 1993). These cells can bud to form more vegetative cells, and make up the majority of the biomass of a pseudohypha (Gimeno et al., 1992; Gimeno et al., 1993).

B.

Known Inhibitors and Enhancers of Pseudohyphal Growth

The molecular mechanism by which changes in nutrient conditions bring about the dramatic changes in cell morphology and growth which constitute the pseudohyphal transition is being actively researched. Thus far, the genes which have been identified as affecting pseudohyphal growth can be separated into three groups: (1) those which are likely to affect nutrient sensing (MSNUPHD2, SHR3, and RAS2 (Gimeno et al., 1992; Gimeno and Fink, 1994), (2) those which are likely to participate in the

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control of gene expression (Liu et al., 1993; Gimeno and Fink, 1994), and (3) those which affect protein phosphorylation states (Blacketer et al., 1993). Information about the targets of transcriptional and posttranslational control by the latter groups of regulators is currently lacking. Whether these genes primarily affect morphogenesis or growth control events that are critical for pseudohyphal growth cannot be determined, as relatively little is known about pseudohyphal transitions. A Kinase Cascade and Transcriptional Regulation in Pseudohyphal Growth

Liu et al. reported the surprising finding that a subset of components of the MAPK pathway which functions in pheromone-induced transcriptional activation (described below; Sprague and Thorner, 1992; Kurjan, 1993) also are required for the pseudohyphal transition (Liu et al., 1993). Activation of the mating response requires a linear array of protein kineses: Ste2Op + Stel l p + Ste7p -+ Fus3p, Ksslp. Fus3p and Ksslp are MAP kinase homologs which activate a transcription factor, Stel2p. Cells deleted for Ste20p, Stel lp, Ste7p, and Stel2p function fail to undergo pseudohyphal transitions, and dominant activated alleles of STEll and STEl2 cause enhanced pseudohyphal growth in wild-type cells. Genetic analyses indicates that the order of function in pseudohyphal growth is conserved with mating: Ste20p + Stel l p + Ste7p -+ Stel2p. Interestingly, neither the pheromone receptors and heterotrimeric G protein (which function upstream of Ste2Op during mating), nor the Fus3p and Ksslp MAPKs are required during pseudohyphal growth. The alternative upstream signalling and downstream targets of this pathway during pseudohyphal growth are not known. In mating, both morphogenesis and cell cycle control are regulated by the mating MAPK pathway through the activity of the Stel2p transcription factor. This pathway, perhaps by activation of a different MAP kinase(s), may play a similar central regulatory role in pseudohyphal growth. Another possible transcriptional regulator of pseudohyphal growth, Phd lp, was recovered in a scrcen for genes which when present in multiple copies enhance pseudohyphal growth (Gimeno and Fink, 1994). Phdlp is a nuclear protein which contains a proline-rich putative activation domain and a region of significant similarity with the DNA-binding domains of the yeast transcription factors, Mbplp and Swi4p (Primig et al., 1992; Koch et al., 1993). Overexpression of PHDI causes pseudohyphal growth of diploids even on noninducing, rich media; in haploid cells it causes cell elongation. Other Proteins that Function in Pseudohyphal Growth

Four loci ELMI, ELM2, ELM3, and CDC.55 have been described that are potential negative regulators of pseudohyphal growth (Blacketer et al., 1993). ELM1 and CDC5.5 encode a putative protein kinase (Blacketer et al., 1993) and protein phophatase subunit (Healy et al., 1991), respectively; ELM2 and ELM3 are less well characterized. Homozygous diploid elm1 -A mutants exhibit a constitutive pseudo-

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hyphal phenotype: on rich media cells are elongated, remain attached after cytokinesis, form colonies from which filaments of cells radiate out, and invade the agar (Blacketer et al., 1993). Homozygous diploid elm2 and elm3 mutants exhibit the same constitutive pseudohyphal phenotype on rich media and also enhance the efficiency of pseudohyphal transition even when present in the heterozygous state (Blacketer et al., 1993). Genetic interactions between CDC5.5 and ELM1 suggest that they carry out overlapping functions in sated, vegetative growth (Blacketer et al., 1993). In addition, heterozygous cdc55/+ cells undergo amore rapid pseudohyphal transition than wild-type cells (Blacketer et al., 1993) and cdc.55 mutants at the restrictive temperature for growth are elongate and remain attached after cytokinesis (Healy et al., 1991). Thus Elmlp- and Cdc55p-regulated protein phosphorylation states are likely to be important for pseudohyphal growth.

C.

Pseudohyphal Morphogenesis May Result from Differential Regulation of Components Conserved with Bud and Mating Projection Formation

It is anticipated that the processes which underlie polarized growth in vegetative cells will be conserved in pseudohyphal cells. The similar random budding phenotype of rsrl/budl mutants in both pseudohyphal and vegetative growth supports this theory (Bender and Pringle, 1989; Chant andHerskowitz, 1991; Gimeno et al., 1992; Ruggieri et al., 1992). The observed differences in shape between vegetative and pseudohyphal cells probably reflect differences in assembly of cytoskeletal and cell wall structure. This might be accomplished by several means. Regulation of actin and/or the neck filaments might be a primary determinant of morphology. Certain act1 mutants exhibit distinct cell morphologies including elongated cells (Drubin et al., 1993) (like pseudohyphal cells). In addition, cdc3(ts), csclO(ts), cdcl l(ts), and cdcl2(ts) mutants incubated at restrictive temperatures form long chains of connected cells, each with an elongated morphology (Hartwell, 1971; Flescher et al., 1993), that is highly reminiscent of pseudohyphal cells. Perhaps limiting nitrogen is sensed by the cell and translated into changes in actin dynamics, actin-binding protein activities, and/or regulation of the neck filament proteins, and this leads to the different morphologies apparent in vegetative and pseudohyphal cells. It is also possible that nitrogen limitation affects the activities of enzymes which act on the cell wall (Bartnicki-Garcia and McMurrough, 1971), and this leads to the changes in cell shape and alterations in cell physiology which characterize pseudohyphal growth.

IV. MATING PROJECTIONFORMATION Projection formation during mating (reviewed in Sprague and Thorner, 1992) has many similarities to budding (Drubin, 1991; Madden et al., 1992). Like budding, projection formation involves several steps including a commitment to growth, se-

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lection of a growth site, polarization of the cytoskeleton and secretory apparatus to growth sites, and polarized movement of organelles. Many components involved in budding carry out similar functions in projection formation. However, the following description of polarized growth during mating is necessarily incomplete. Extensive genetic screens to identify essential components for projection formation have not been performed and, furthermore, many proteins and regulatory pathways important in budding have not been examined for their function in morphogenesis during mating. A.

Cytology of Mating Projection Formation

In response to mating pheromone from a cell of the opposite mating type, a haploid cell arrests in GI and initiates projection formation on one edge of the cell. Growth is concentrated in the apical portion of the emerging projection (see Figure 2). Polarized growth components such as Spa2p and Bemlp localize to the projection tip (Snyder, 1989; Gehrung and Snyder, 1990; J. Chenevert, personal communication cited in Sprague and Thorner, 1992) and new cell wall mannan is deposited diffusely in the apical portion of the projection (Tkacz and MacKay, 1979). Polarized secretion directs growth. Membrane vesicles accumulate in the emergent projection and are also present, though in relatively less abundance, as the projection enlarges (Baba et al., 1989). Growth of the projection results in an elongated cell, called a “shmoo,” with a cylindrical or pear-shaped morphology (Moore, 1983; Segall, 1993). Efficient mating requires that at least one partner form shmoos well (Gehrung and Snyder, 1990; Chenevert et al., 1992).A critical difference between projection formation and budding is the absence of a tight constriction at the base of the growth site. Although there may be some constriction, the width of the base is rarely less than the width of the projection. Chitin and the neck filament proteins, Cdc3p, CdclOp, Cdcl lp, and Cdc12p, which are important constituents of the constriction in budding cells, exhibit a relatively diffuse localization to the apical portion of the mating projection (SchekmanandBrawley, 1979;FordandPringle, 1991; Kimetal., 1991;H. Kim,S. Ketcham, B. Haarer, and J. Pringle, personal communication). It is possible that these components contribute a structural role in defining the shape of the projection. Alternatively, although these proteins are polarized they may be inactive in projection morphogenesis due to lack of Cdc28Klnp activity (see below). The projection can grow until it reaches several times the length of the original cell. At the projection tip the cell wall is relatively thin and contains a diffuse outer layer (Lipke et al., 1976; Baba et al., 1989). Organelles such as the nucleus and mitochondriaare polarized toward the projection tip (Haseket al., 1987; Babaet al., 1989; Gehrung and Snyder, 1990), whereas vacuoles become fragmented and are localized in the posterior of the shmoo relative to the projection tip (Baba et al., 1989). Ultimately two mating cells fuse at the tips of their projections in a process that involves localized cell wall degradation and plasma membrane fusion. Cytoplas-

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mic fusion is followed by karyogamy. The resulting diploid nucleus undergoes DNA replication and mitosis, and is segregated into the zygotic bud. Both parental vacuoles contribute components to the single vacuole in the zygotic bud (Weisman and Wickner, 1988); in contrast parental mitochondria often do not fuse in the zygote or zygotic bud, and diploid progeny often inherit mitochondria from only one parent (Lukins et al., 1973). B.

Cell Signalling in Response to Pheromone

A great deal is known concerning the signal transduction process required for mating (reviewed in (Sprague and Thorner, 1992; Kurjan, 1993). Mating pheromones are recognized by cell type-specific receptors present on the cell surface (Hagen et al., 1986;Jackson et al., 1991). The STE2 gene product is expressed in MATa cells and binds a-factor and the STE3 gene product is expressed in MATa cells and binds a-factor. Binding of pheromone activates a G protein-coupled MAPK pathway. As noted above, the component kineses of this pathway and their relative order of action are: Ste20p + Stel l p -+ Ste7p -+ Ksslp, Fus3p (Rhodes et al., 1990; Cairns et al., 1992; Gartner et al., 1992; Leberer et al., 1992; Stevenson et al., 1992). Ksslp and Fus3p are MAPKs (Boulton et al., 199 1 ; Gartner et al., 1992). Fus3p and Kss 1p carry out overlapping functions in activating Stel 2p, a transcription factor, that mediates transcriptional activation of pheromone-induced genes (Errede and Ammerer, 1989; Elion et al., 1991; Song et al., 1991). Transcription of the pheromone and pheromone receptor genes are each induced by the pheromone response, thereby amplifying the signal (Strazdis and MacKay, 1983; Nakayama et al., 1985; Hagen et al., 1986; Sprague and Thorner, 1992). Fus3p has an additional essential function in bringing about cell cycle arrest by inactivating the G1 cyclins (Elion et al., 1991). Although the mechanism for this is not completely clear, Fus3p acts in part in conjunction with the pheromone-induced Far1 protein (Chang and Herskowitz, 1990; Peter et al., 1993; Valdivieso et al., 1993). A Fus3p-independent mechanism for inactivation of a subset of the G1 cyclins is also likely to exist (Elion et al., 1991). C.

Pheromone Concentration Controls the Initiation and Shape of Projection Formation

The yeast pheromone response depends upon the pheromone concentration (see Figure 8; Moore, 1983). When exposed to low concentrations of pheromone, cells undergo G1 arrest and pheromone-induced transcriptional activation. The cells initially grow isotropically; after prolonged exposure to pheromone the cells gradually polarize growth and their shapes change from “round” to “ovoid’ (Moore, 1983). These ovoid cells resemble the “cylindrical” shmoos observed by Segall in cells responding to a spatial gradient of low pheromone concentrations (Segall,

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High concentrations of pheromone

Polarized growth at the proximal site

Low concentration pheromone gradient

Transient depolarization followed by gradual polarization of growth toward the high end of the pheromone gradient

Figure 8. The pheromone concentration and gradient determine the timing of the onset of polarized growth, selection of the growth site, and morphology of the mating projection. In high uniform concentrations of pheromone, cells initiate projection formation rapidly upon cell cycle arrest at sites determined by the bud site selection mechanism, that is, they form projections at sites adjacent to the previous bud site. The projection formed is thin and pointed. In contrast, in gradients of low concentrations of pheromone, cells arrest and undergo transient depolarization and isotropic growth. Ultimately, growth is polarized toward the high end of the pheromone gradient, and a broad, rounded projection is formed.

1993). As long as cells treated with pheromone are arrested in GI, they will ultimately undergo polarized cell growth (L. Vallier, M. Snyder, and J. Segall, personal communication). At high pheromone concentrations (10-1 00 times higher than those described above) cells also arrest in G1 and activate the mating response (Moore, 1983; Madden and Snyder, 1992; Segall, 1993). However, these cells polarize rapidly and the projections formed are much thinner than those of cells which polarize after treatment with low concentrations of pheromone (Moore, 1983; Segall, 1993). Thus, both the onset of polarized growth and the morphology of the projection are dependent on pheromone concentration.

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D.

Pheromone Concentration and Gradient Direct the Orientation of Polarized Growth

A number of lines of evidence show that the yeast cell can respond to the local pheromone gradient, and can initiate projection formation anywhere on the surface in response to that gradient. First, in competition assays cells preferentially mate with strains producing higher levels of pheromone (Jackson and Hartwell, 1990a,b). Second, in cells from mating mixtures sites of projection formation are random with respect to previous bud sites, presumably because the cell responds to the local pheromone gradient generated by a cell of the opposite mating type (Madden and Snyder, 1992). Finally, cells in pheromone gradients form and orient their projections toward the highest concentration of pheromone (Segall, 1993). Thus, projection site selection is “plastic” compared to the “hard-wired’’ mechanism directing bud site selection. Projection growth toward a mating partner is likely to be optimal in spatial gradients of relatively low pheromone concentration because these conditions induce a delay in the initiation of polarized growth, transient depolarization of actin, and isotropic growth of the cell (Moore, 1983; Madden and Snyder, 1992). Depolarization may be accompanied by some modification of the cytokinesis tag, thus allowing the cell to respond to external cues i.e. the pheromone gradient of a mating partner (Madden and Snyder, 1992). In addition, the isotropic growth probably serves to increase the surface area over which the cell can respond to and transmit pheromones. It is likely that very high levels of pheromone saturate the receptors so that cells can no longer detect gradients. Cells form and orient projections randomly when exposed to high concentration pheromone gradients (Segall, 1993). Under these circumstances cells probably grow at sites specified by the bud site selection mechanism. Consistent with this hypothesis in the presence of uniform high pheromone concentrations the bud site selection mechanism determines the site of projection formation, and projections usually form at sites proximal to previous bud sites (Madden and Snyder, 1992). The saturation effect can explain why mutants which are supersensitive to pheromone are defective in two types of orientation assays. Supersensitive strains d o not orient projections along pheromone gradients at concentrations in which wild-type cells are able to orient, but they do orient correctly if the gradient is lowered in pheromone concentration (Segall, 1993). Supersensitive strains are also defective in discrimination between pheromone-producing and non-producing partners in competition assays (Jackson and Hartwell, 1990a). Presumably the pheromone produced by the normal mating partner saturates the signalling system in the supersensitive cell and induces immediate polarization at sites that are random with respect to the location of the pheromone-producing partner.

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Projection Orientation is Controlled by Pheromone Receptors and Other Components at the Cortex

Mating partner discrimination probably reflects the cell’s ability to form and orient its projection in the direction of the mating partner producing the most pheromone. In competition assays using cells that constitutively express the mating program, discrimination is dependent upon the pheromone receptor and, to a lesser extent, onACT1, MY02, and CHCl (Jackson et al., 1991). In contrast, microtubules are not needed for discrimination and tub2 mutants behave like wild-type cells in these assays (Jackson et al., 1991). Moreover, cells treated with nocodazole still orient projections toward the high end of a pheromone gradient (Segall, 1993). Thus the pheromone receptor, actin, Myo2p and clathrin heavy chain are all implicated in the process of directing projection growth in response to a spatial gradient of pheromone whereas microtubules are not important for this process. A model describing how some of these components might function is described below.

F.

Components Important for Projection Formation

Projection formation requires many of the same components that are important in budding, including actin, actin-interacting proteins, cortical proteins such as Spa2p and Bemlp, and regulatory proteins which bind GTP or Ca++.Certain mutations in cell polarity genes have a relatively greater impact on morphogenesis in mating than they do in budding, for example spa2-A and beml-s (see below, (Snyder, 1989; Gehrung and Snyder, 1990; Chenevert et al., 1992). This might reflect either a specific function of the protein in forming projections or, more likely, a greater sensitivity of the projection formation process to mutations that affect polarized growth. Finally, the pheromone receptors function in projection growth, in addition to their roles in signal transduction and projection orientation, described above. Cytoskeletal Components The Actin Cytoskeleton. In mating cells, actin has a characteristic asymmetric distribution. Actin cables are oriented toward the projection tip, and actin patches accumulate at the tip of the projection, preferentially toward the cell surface (Figure 2; Hasek et al., 1987; Gehrung and Snyder, 1990; Read et al., 1992). Actin is required for organizing the mating projection (Read et al., 1992). At high temperatures, pheromone-treated actl(ts) cells arrest in G 1 but exhibit moderate to severe defects in cell polarization and projection formation. A significant fraction of a c t l cells remain round or oval in shape, and fail to polarize either actin patches or Spa2p. Actin cables appear to be dispensable for projection formation since a c t l -4 mutants lack actin cables at high temperatures, but are able to form projections, albeit inefficiently (Dunn and Shortle, 1990; Read et al., 1992). Analogous to the role

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described for actin in budding, actin probably participates in polarized secretion toward the projection tip. Given the importance of actin to projection formation, it is expected that actininteracting proteins also participate in projection formation. Indeed, this has proven to be true for some of the few actin-binding proteins which have been examined for roles in mating. tpml-A mutants arrest properly in response to pheromone, but even at high pheromone concentrations only a subset of tpml-A mutants are able to polarize actin and undergo some form of polarized growth (Liu and Bretscher, 1992). In addition, bilateral tpml - A matings often fail, probably because of a defect in cell fusion (Liu and Bretscher, 1992). Another actin-binding protein mutant, capZ-A, exhibits only mild defects (Amatruda et al., 1992). Mutation in CAP2 causes a slight bilateral mating defect, and projection formation and polarization of actin are indistinguishable from wild-type. Myo2p has not been directly examined for its role in projection formation and mating. However, it is possible that Myo2p might carry out actin-based membrane trafficking events in projection formation, similar to its hypothesized role in polarized growth in budding (Prendergast et al., 1990; Johnston et al., 1991). The presumptive Myo2p regulator, Cmdlp (Brockerhoff et al., 1994; Ohya and Botstein, 1994), is present during mating and is polarized to mating projection tips (Figure 2; Brockerhoff and Davis, 1992). Microtubules Are Polarized in Making Projections and Are Important for SPB Orientation. Polarization of actin is accompanied by polarization of the SPB and

microtubules in a distribution that is reminiscent of that seen in budding cells (Drubin, 1991; Madden et al., 1992). The SPB lies at the point in the nuclear envelope nearest to the growth site and microtubules emanating from the SPB orient toward, and often extend to, the projection tip (Figure 2; Byers and Goetsch, 1975; Gehrung and Snyder, 1990; Snyder et al., 1991; Read et al., 1992). Some microtubules are also observed to orient away from the projection tip, toward the cell body (Gehrung and Snyder, 1990; Read et al., 1992). There is some evidence that the microtubules which intersect the region of the projection tip are preferentially stabilized (Read et al., 1992), indicating that there may be microtubule capture sites in that region. As in budding, microtubules are not important for polarized growth in mating. Cells treated with microtubule depolymerizing drugs and tub2 mutants arrest in response to pheromone and form normal mating projections (Haseket al., 1987;Read et al., 1992). However, polarization of the SPB within the projection tip requires microtubules and can be inhibited by actin cytoskeletal defects (Hasek et al., 1987; Read et al., 1992). This is analogous to the primary importance of microtubules, and contributing role of actin, to nuclear segregation in budding (Huffaker et al., 1988; Jacobs et al., 1988; Palmeret al., 1992; Drubin et al., 1993). SPB orientation toward the projection tip is presumably important for karyogamy after cell fusion. After cells mate the SPBs of the two nuclei face one another and microtubules from the two SPBs form an interdigitating array (Byers and Goetsch, 1975). The nuclei are

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thought to be drawn together by sliding microtubules past one another (see Page and Snyder, 1993); fusion occurs at the SPBs (Byers and Goetsch, 1975). Karyogamy requires microtubules and is mediated by a kinesin-related microtubule motor protein (Delgado and Conde, 1984; Meluh and Rose, 1990). Cortical Proteins which Contribute to Projection Morphogenesis. Spa2p and Bemlp are found at the tip of the mating projection and have important roles in projection formation (Figure 2; Snyder, 1989; Gehrung and Snyder, 1990; Chenevertet al., 1992; Chenevert et al., 1994; J. Chenevert, personal communication cited in Sprague and Thorner, 1992). Strains lacking Spa2p, spa2-A, or expressing certain hypomorphic forms of Bemlp, beml -s,arrest as unbudded cells and enlarge when treated with mating pheromone. However, they are defective in properly polarizing actin and forming mating projections. Pheromone-treated spa2-A mutants are predominantly round or oval, though sometimes they form broad, rounded projections; in these cells the degree of actin depolarization roughly correlates with the failure to form a projection (Gehrung and Snyder, 1990; Chenevert et al., 1994). Pheromone-treated beml -s mutants appear to be uniformly round with depolarized actin (Chenevert et al., 1992; Chenevert et al., 1994). Both spa2 and beml mutants exhibit bilateral mating defects and for spa2 cells the mating defect on solid medium is increased wt-.:n cells are more dilute (Gehrung and Snyder, 1990; Chenevert et al., 1992; Chenevert et al., 1994). These phenotypes are consistent with the expectation that efficient mating requires polarization of at least one mating partner (Gehrung and Snyder, 1990). The pheromone receptor, which like Spa2p and Bemlp concentrates at the projection growth site (Jackson et al., 1991), is also important in projection formation. Konopka et al. report that cells expressing a truncation of the carboxy terminal cytoplasmic tail of the Ste2p a factor receptor, Ste2p-T326, arrest in response to pheromone and enlarge but fail to form projections, remaining round or oval in shape (Konopka et al., 1988). When these mutants are examined after being treated with pheromone for extended time periods, they ultimately do form projections; hence they are delayed in projection formation compared to wild-type cells, but do ultimately undergo polarized growth (L. Vallier, J. Segall, and M. Snyder, unpublished data). Genes that Regulate Budding Are Also Implicated in Projection Morphogene-

The two Rho proteins, Rholp and Cdc42p, are essential for budding (Madaule et al., 1987; Adams et al., 1990; Johnson and Pringle, 1990; Yamochi et al., 1994) and preliminary evidence suggests that they are also important in projection formation. cdc24(ts)mutants fail to form projections at semi-permissive or restrictive temperatures for vegetative growth, and fail to mate at the restrictive temperature (Reid and Hartwell, 1977; Field and Schekman, 1980; Chenevert et al., 1994). In budding Cdc24p apparently serves as a guanine (GDP) dissociation stimulator forCdc42p(Hartetal., 1991;Ronetal., 1991;Zhengetal., 1994);Cdc42pcan then

5;s.

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bind GTP and presumably activate growth events at the cortex. During projection formation Cdc42p is specifically localized at the tip of the mating projection (Figure 2; Ziman et al., 1993), thus it is plausible that it functions in projection growth and is an essential substrate for Cdc24p activity in mating. Projection formation is accompanied by Ca++transport into the cell and a large increase in intracellular Ca++ (Ohsumi and Anraku, 1985; Iida et a]., 1990b). Cdc24p contains Ca++binding domains (Miyamoto et al., 1987) and thus its activity may be modulated by the changing Ca++concentration. Mutants in RHO1 have not been examined with respect to projection formation, however mutants of the putative Rholp GAP, Bem2p (Bender and Pringle, 1991; Y. Zheng, R. Cerione, and A. Bender, unpublished data, cited in Zheng et al., 1994), cause projection formation defects (Yi-Jun Sheu, K. Madden, and M. Snyder, unpublished data). Thus two essential Rho proteins for budding, Cdc42p and Rholp, are also implicated in polarized growth in mating. Components of the Slt2p(Mpklp) MAPK pathway that functions downstream of Pkclp, are required for projection formation (Torres eta]., 1991; Costigan et al., 1992; Lee and Levin, 1992; Levin and Bartlett-Heubusch, 1992; Irie et a]., 1993; Lee et al., 1993; Levin and Errede, 1993; Mazzoni et al., 1993). s l k l - A and slt2-A mutants polarize, but form aberrant truncated projections (Costigan et al., 1992; Mazzoni et al., 1993). The Slt2p(Mpklp) pathway might regulate either cytoskeletal or cell wall components and thus affect projection formation. A genetic screen has identified two other mutants, pea2 and tnyl, which exhibit defects in projection formation (Chenevert et al., 1994). These mutants also exhibit mild unilateral mating defects. Pea2p localizes to growth sites similar to Spa2p, and both proteins interact physically (Valtz et al., 1996; Sheu, Y.J., and Snyder, M., unpublished). It is expected that further characterization of all the projection formation mutants, as well as other mutants implicated in polarized growth during bud formation, wiIl provide insight into the process of polarized growth during mating. C.

Summary and Model of How Projection Formation and Orientation Occurs

As is described above, many aspects of projection formation closely resemble budding. Actin, Spa2p, Bemlp, Cmdlp, microtubules, and Cdc42p have similar polarized distributions and presumably similar functions in both processes. Furthermore, regulatory factors such as Cdc24p, Bem2p, and the Slt2p MAPK pathway all function in both projection formation and in budding. Organelles are polarized in both buds and shmoos, and the microtubule cytoskeleton has a primary role in nuclear positioning in both processes. Hence a likely mechanism for projection formation that incorporates what is known about polarized growth during budding is as follows (modified from Moore, 1983; Madden and Snyder, 1992; Segall, 1993). When a cell encounters a spatial gradient of relatively low pheromone concentration, signalling from the pheromone receptor is modest, and the cell arrests in G 1

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and undergoes transient depolarization (Moore, 1983; Madden and Snyder, 1992). Consequently cortical growth sites determined by the bud site selection mechanism are modified (Madden and Snyder, 1992) and/or are overridden by signals from the mating response. The levels of mating pheromones are induced, and hence the concentration of the pheromone gradient between two responding cells is likely to increase with time, thus enhancing their growth toward each other (Strazdis and MacKay, 1983; Madden and Snyder, 1992). The G1-arrested cell continues to grow and secrete pheromone receptors. Receptor activation on the cell surface closest to the pheromone source will be slightly greater than activation at other surfaces of the cell, because there the frequency of receptor occupancy will be slightly higher. Consequently slightly more growth components and newly synthesized pheromone receptors will be targeted toward this surface, which will, in turn, amplify signalling and thus further stimulate growth at this surface relative to other surfaces of the cell. The accumulation of pheromone receptors and/or the intracellular signalling generated by receptors presumably specify the cortical site of polarized growth; How this occurs is not known. The receptor itself might interact with growth components at the surface, or activate adjacent effectors at the surface that participate in recruitment andlor assembly of new growth components (See Roemer et al., 1996). Alternatively, ligand-induced receptor endocytosis might indirectly polarize secretion to specific cortical sites, simply by causing increased assembly of components important for both processes, for example actin and Myo2p (Novick and Botstein, 1985; Johnston et al., 1991; Kubler and Riezman, 1993; Govindan and Novick, 1993). Nevertheless, once the cortical site is established by the mating pheromone receptor or its signal, downstream events involved in the assembly of the projection are likely to be similar to the events in bud site assembly. Cdc42p-GTP probably participates in targeting growth and/or cytoskeletal components to the cortical site and may also participate in overall surface growth. Rholp is likely to be activated either in the cytoplasm or at the cortex, and participate in surface growth by assembling growth and/or cytoskeletal components. Rholp and/or Rho 1p assembled with complexes may be targeted by Cdc42p to growth sites. The resulting polarization and assembly of cytoskeletal and growth components such as actin, actin regulators, secretory components, and cortical growth factors at the cortical site drives growth of the projection.

V.

CONCLUSION

Bud formation and projection formation are distinct mechanisms for polarized cell growth, and in the past ten years a wealth of information about how these processes occur has been generated. Many of the components involved in polarized cell growth in yeast, including both structural components such as actin and regulatory components such as MAP kinase pathways and Rho proteins, are remarkably con-

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served in structure and in function with other eucaryotic cell types. The ease with which components can be identified and analyzed in S. cerevisiae will continue to make this organism a useful model for identifying new components, determining how they function, and elucidating the molecular pathway by which polarized cell growth occurs and is regulated in eucaryotes.

ACKNOWLEDGMENTS We thank our numerous colleagues for communication of unpublished results. S. Erdmann, K. Madden, and L. Vallier provided critical comments on the manuscript. Research from our laboratory was supported by NIH grant GM36494. C.C. was supported in part by a John Woodruff Simpson Fellowship award.

REFERENCES Adams,A. & Pringle, J. (1984). Relationshipofactin and tubulin distribution to bud growth in wild-type and morphogenetic-mutant Succharomyces cerevisiae. J. Cell Biol. 98,934-945. Adams, A. E., Cooper, J . A. & Drubin, D. G. (1993). Unexpected combinations of null mutations in genes encoding the actin cytoskeleton are lethal in yeast. Mol. Biol. Cell 4 , 4 5 9 4 6 8 . Adams, A. E. M. & Botstein, D. (1989). Dominant suppressors of yeast actin mutations that are reciprocally suppressed. Genetics 121, 675-683. Adams, A. E. M., Botstein, D. & Drubin, D. G. (1991). Requirement of yeast fimbrin for actin organization and morphogenesis in vivo. Nature 354,404408. Adams, A. E. M., Johnson, D. I., Longnecker, R. M., Sloat, B. F. & Pringle, J. R. (1990). CDC42 and CDC43, two additional genes involved in budding and the establishment of cell polarity in the yeast Succhuromycescerevtsiue.J. Cell Biol. 1 1 1, 131-142. Amatruda, J. F., Cannon, J. F., Tatchell, K., Hug, C. & Cooper, J. A. (1990). Disruption of the actin cytoskeleton in yeast capping protein mutants. Nature 344, 352-354. Amatruda, J. F. & Cooper, J. A. (1 992). Purification, characterization, and immunofluorescence localization of Saccharomyces cerevisiae capping protein. J. Cell Biol. 117, 1067-1076. Amatruda, J. F., Gattermeir, D. J., Karpova, T. S. & Cooper, J . A. (1992). Effects of null mutations and overexpression of capping protein on morphogenesis, actin distribution, and polarized secretion in yeast. J. Cell Biol. 119, 1151-1 162. Baba, M., Baba, N., Ohsumi, Y., Kanaya, K. & Osumi, M. (1989). Three-dimensional analysis of morphogenesis induced by mating pheromone a factor in Saccharomyces cerevisiae. J. Cell Sci. 94,207-216. Bartlett, R & Nurse, P. (1990). Yeast as a model system for understanding the control of DNA replication in Eukaryotes. BioEssays 12,457-463. Bartnicki-Garcia, S. & McMurrough, I. (1971). Biochemistry of morphogenesis in yeasts. In. The Yeasts (Rose, A. H. & Harrison, J. S., Eds.), Vol. 2, pp. 441492. Academic Press, New York. Bauer, F., Urdaci, M., Aigle, M. & Crouzet, M. (1993). Alteration of a yeast SH3 protein leads to conditional viability with defects in cytoskeletal and budding patterns. Mol. Cell. Biol. 13, 507&5084. Bedinger, P. A,, Hardeman, K. J. & Loukides, C. A. (1994). Travelling in style: the cell biology of pollen. Trends Cell Biol. 4, 132-138. Bender, A. (1993). Genetic evidence for the roles of the bud-site-selection genes BUDS and BUD2 in control of the Rsrlp (Budlp) GTPase in yeast Proc. Nail. Acad. Sci. USA 90,9926-9929.

Cell Polarity in Yeast

55

Bender, A. & Pringle, J. R. (1989). Multicopy suppression of the cdc24 budding defect in yeast by CDC42 and three newly identified genes including the ras-related gene RSRI. Proc. Natl. Acad. Sci. USA 86,9976-9980. Bender,A. & Pringle, J. R. (1991). Useofascreen forsyntheticlethal andmulticopysuppresseemutants to identify two new genes involved in morphogenesis in Saccharomyces cerevisiae. Mol Cell. Biol. 11, 1295-1305. Bender, A. & Pringle, J. R. (1992). A SerRhr-rich multicopy suppressor of a cdc24 bud emergence defect. Yeast 8 , 3 15-323. Benton, B. K., Tinkelenberg, A. H., Jean, D., Plump, S. D. & Cross, F. R. (1993). Genetic analysis of CldCdc28 regulation of cell morphogenesis in budding yeast. EMBO J. 12,5267-5275. Blacketer, M. J., Koehler, C. M., Coats, S. G., Myers, A. M. & Madaule, P. (1993). Regulation of dimorphism in Saccharomyces cerevisiae: involvement of the novel protein kinase homolog Elmlp and protein phosphatase2A. Mol. Cell. Biol. 13, 5567-5581. Boulton,T. G.,Nye, S. H., Robbins,D. J., Ip,N. Y., Radziejewska,E.,Morgenbesser,S. D.,DePinho,R. A,, Panayotatos, N., Cobb, M. H. & Yancopoulos, G. D. (1991). ERKs: a family of protein serinehhreonine kineses that are activated and tyrosine phosphorylated in response to insulin and NGF. Cell 65,663-675. Bowser, R., Muller, H., Govindan, B. & Novick, P. (1992). Sec8p and Secl5p are components of a plasma membrane-associated 19.5s particle that may function downstream of Sec4p to control exocytosis. J. Cell Biol. 118, 1041-1056. Bowser, R. & Novick, P. (1991). Secl5 protein, an essential component of the exocytotic apparatus, is associated with the plasma membrane and with a soluble 19.5 S particle. J. Cell Biol. 112, 1 1 17-1 131. Brennwald, P. & Novick, P. (1993). Interactions of three domains distinguishing the Ras-related GTP-binding proteins Yptl and Sec4. Nature 362, 56Cb563. Brewster, J. L. & Gustin, M. C. (1994). Positioning of cell growth and division after osmotic stress requires a MAP kinase pathway. Yeast 10,4255439. Brewster, J. L., Valoir, T. D., Dwyer, N. D., Winter, E. & Gustin, M. C. (1993). An osmosensingsignal transduction pathway in yeast. Science 259, 1760-1763. Broach, J. R. (1991). RAS genes in Saccharomyces cerevisiae: signal transduction in search of a pathway. Trends Genet. 7,28-33. Brockerhoff, S. E. & Davis, T. N. (1992). Calmodulin concentrates at regions of cell growth in Succharomyces cerevisiae. J. Cell Biol. 118,619-629. Brockerhoff, S. E., Stevens, R. C. & Davis, T. N. (1994). The unconventional myosin, MyoZp, is a calmodulin target at sites of cell growth in Succharomyces cerevisiae. J. Cell Biol. 124, 3 15-323. Brown, C. M. & Hough, J . S. (1965). Elongation of yeast cells in continuous culture. Nature 206, 676-678. Byers, B. (1981). Cytology of the yeast life cycle. In: The Molecular Biology of the Yeast Sacciiaromyces: Life Cycle and Inheritance (Strathern, J.N., Jones, E. & Broach, J., Eds.), pp. 59-96, Cold Spring Harbor Laboratory. New York. Byers, B. & Goetsch, L. (1975). Behavior of the spindle plaques in the cell cycle and conjugation of Saccharomyces cerevisiae. J. Bacteriol. 124, 5 1 1-523. Byers, B. & Goetsch, L. (1976). A highly ordered ring of membrane-associated filaments in budding yeast. J. Cell Biol. 69, 717-721. Cairns, B. R., Ramer, S. W. & Kornberg, R. D. (1992). Order of action of components in the yeast pheromone response pathway revealed with a dominant allele of the STEl 1 kinase and the multiple phosphorylation of the STE7 kinase. Genes & Dev. 6, 1305-1318. Caplow, M. (1992). Microtubule dynamics. Curr. Opin. Cell Biol. 4, 58-65. Cliang, F. & Herskowitz, I. (1990). Identification of a gene necessary for cell cycle arrest by a negative growth factor of yeast: FAR1 is an inhibitor of a GI cyclin, CLNZ. Cell 63,999-101 1.

56

CHRISTINE COSTICAN and MICHAEL SNYDER

Chant, J., Corrado, K., Pringle, J. R. & Herskowitz, 1. (1991). The yeast BUDS gene, which encodes a putative GDP-GTP exchange factor, is necessary for bud-site selection and interacts with bud-formation gene BEMI. Cell 65, 1213-1224. Chant,J. &Herskowitz,l. (1991).Geneticcontrol ofbud-siteselection in yeastby asetofgeneproducts that comprise a morphogenetic pathway. Cell 65, 1203-1212. Chant, J., Mischkle, M., Mitchell, E., Herskowitz, I., & Pringle, J. R. (1995). Role of Bud3p in producing the axial budding pattern of yeast. J. Cell Biol. 129, 767-778. Chen, W., Lim, H. H. & Lim, L. (1993). The CDC42 homolog from C. elegans. J. Biol. Chem. 268, 13280-1 3285. Chenevert, J., Corrado, K., Bender, A,, Pringle, J . & Herskowitz, I. (1992). A yeast gene (BEMI) necessary for cell polarization whose product contains two SH3 domains. Nature 356, 77-79. Chenevert, J., Valtz, N. & Herskowitz, I. (1994). Identification of genes required for normal pheromone-induced cell polarization in Succharomyces cerevisiae. Genetics 136, 1287-1297. Cheney, R. E., Riley, M. A. & Mooseker, M. S. (1993). Phylogenetic analysis of the myosin superfamily. Cell Motil. Cytoskel. 24,215-223. Chowdhury, S., Smith, K. W. & Gustin, M. C. (1992). Osmotic stress and the yeast cytoskeleton: phenotype-specific suppression of an actin mutation. J. Cell Biol. 118, 561-571. Chrzanowska-Wodnickq M. & Burridge, K. (1992). Rho, rac, and the actin cytoskeleton. Bioessays 14, 777-778. Clarke, S. (1992). Protein isoprenylation and methylation at carboxyterminal cysteine residues. Ann. Rev. Biochem. 61,355-386. Cleves, A. E., Novick, P. J. & Bankaitis, V. A. (1989). Mutations in the SAC1 gene suppress defects in yeast Golgi and yeast actin function. J. Cell Biol. 109,2939-2950. Coleman, K. G., Steensma, H. Y., Kaback, D. B. & Pringle, J. R. (1986). Molecular cloning of chromosome IDNA from Succharomyces cerevuiae: isolation and characterization of the CDC24 gene and adjacent regions of the chromosome. Mol. Cell. Biol. 6,451W525. Cooper, J. A. (1990). The src family of protein-tyrosine kineses. in Peptides and protein phosphorylation, CRC Press, Boca Raton, Fla., Ed. B. E. Kemp 85-1 13. Costigan, C., Gehrung, S. & Snyder, M. (1992). A synthetic lethal screen identifies SLKI, a novel protein kinase homolog implicated in yeast cell morphogenesis and cell growth. Mol. Cell. Biol. 12, 1162-1178. Costigan, C., Kolodrubetz, D. & Snyder, M. (1994). NHP6A and NHP6B, which encode HMGl-like proteins, function downstream in the yeast SLT2 MAPK pathway. Mol. Cell. Biol. 14, 239 1-2403. Costigan, C. & Snyder, M. (1994). SLKI, a homolog of MAP kinase activators, mediates nutrient sensing independently of the yeast CAMP-dependent protein kinase pathway. Mol. Gen. Genet. 243,286296. Cvrckova, F. & Nasmyth, K. (1993). Yeast G 1 cyclins CLNI and CLN2 and a GAP-like protein have a role in bud formation. EMBO J. 12, 5277-5286. Davis, T. N. (1992). A temperature-sensitive calmodulin mutant loses viability during mitosis. J. Cell Biol. 118,607-617. Davis. T. N., Urdea, M. S., Masiarz, E. R. & Thorner, J. (1986). Isolation ofthe yeast calmodulin gene: calmodulin is an essential protein. Cell 47, 423431. de Boer, P., Crossley, R. & Rothfield, L. (1992). The essential bacterial cell-division protein FtsZ is a GTPase. Nature 359,254-256 Delgado, M. A. & Conde, J . (1984). Benomyl prevents nuclear fusion in Succhuromyces cerevisue. Mol. Gen. Genet. 193, 188-189. DiDomenico, B. J., Brown,N. H., Lupisella, J., Greene, J. R., Yanko, M. & Koltin, Y. (1994). Homologs of the yeast neck filament associated genes: isolation and sequencee analysis of Cundidu ulbrcuns CDC3 and CDCIO. Mol. Gen. Genet. 242,689498.

Cell Polarity in Yeast

57

Drees, B., Bretscher, A,, Brown, C & Barrell, B. (1993). S. cerevisiue contains asecond tropomyosin, encoded by TPM2, which is functionally distinct from the TPMl gene. Mol. Biol. Cell 4, 12a. Drubin, D. (1991). Development o f cell polarity in budding yeast. Cell 65, 1093-1096. Drubin, D. G., Jones, H. D. & Wertman, K. F. (1993). Actin structure and function: roles in mitochondria1 organization and morphogenesis in budding yeast and identification o f the phalloidin-binding site. Mol. Biol. Cell. 4, 1277-1294. Drubin, D. G., Miller, K. G. & Botstein, D. (1988). Yeast actin-binding proteins: evidence for arole in morphogenesis. J. Cell Biol. 107, 2551-2561. Drubin, D. G., Mulholland,J.,Zhu,S. & Botstein, D. (1990). Homologyofayeastactin-bindingprotein to signal transduction proteins and myosin-I. Nature 343,288-290. Dunn, T. M. & Shortle, D. (1990). Null alleles o f S A C 7 suppress temperature-sensitive actin mutations in Succhuromyces cerevisiue. Mol. Cell. Biol. 10,2308-2314. Egilmez, N. & Jazwinski, S. M. (1989). Evidence for the involvement of a cytoplasmic factor in the aging of the yeast Succhuromyces cerevisiue. J. Bacteriol. 171,3742. Eisen, J. S. (1994). Development of motoneuronal phenotype. Ann. Rev. Neur. 17, 1-30. Elion, E., Brill, J. & Fink, G. (1991). FUS3 represses CLNl and CLN2 and in concert with KSSl promotes signal transduction. Proc. Natl. Acad. Sci. USA 88,9392-9396. Errede, B. & Ammerer, G. (1989). STE12, a protein involved in cell-type specific transcription and signal transduction in yeast is part o f protein-DNA complexes. Genes & Dev. 3, 1349-1361. Fedor-Chaiken, M., Deschenes, R. J. & BroachJ R. (1990).SRV2, agenerequiredforRASactivationof adenylate cyclase in yeast. Cell 61,329-340. Field, C. & Schekman, R. (1980). Localized secretion of acid phosphatase reflects the pattern of cell surface growth in Succhuromyces cerevisiue. J. Cell Biol. 86, 123-128. Field, J., Vojtek, A,, Ballester, R., Bolger, G., Colicelli, J., Ferguson, K., Gerst, J., Kataoka, T., Michaeli,T., Powers, S., Riggs, M., Rodgers, L., Wieland, I., Wheland, B. & Wigler, M. (1990). Cloning and characterization of CAP, the S. cerevisiue gene encoding the 70kd adenylyl cyclase-associated protein. Cell 61, 3 19-327. Finegold, A. A,, Johnson, D. I., Farnsworth, C. C., Gelb, M. H., Judd, S. R., Glomset, J. A. & Tamanoi, F. (1 991). Protein gernanylgeranyltransferase of Succhuromyces cerevisiue is specific for Cys-Xaa-Xaa-Leu motif proteins and requires the CDC43 gene product but not the DPRl gene product. Proc. Natl. Acad. Sci. USA 88,44484452. Flescher, E. G., Madden, K . & Snyder, M. (1993) Components required for cytokinesis are important for bud site selection in yeast. J . Cell Biol. 122, 373-386. Ford, S. & Pringle, J. (1991). Cellular morphogenesis in the Succhuromyces cerevisiue cell cycle: Localization of the CDCl I gene product and the timing o f events at the budding site. Devel. Genet. 12,281-292. Freifelder, D. (1960). Bud position in Succhuromyces cerevisiue. J. Bacteriol. 124, 5 1 1-523. Gallwitz,D. & Sures, I. (1980). Structureofasplitgene: completenucleotidesequenceoftheactingene in Succharomyces cerevisrue. Proc. Natl. Acad. Sci. USA 77,2546-2550. Gartner, A,, Nasmyth, K. & Ammerer, G. (1992). Signal transduction in Succhuromyces cerevisrue requires tyrosine and threonine phophorylationofFUS3 and KSSI. Genes & Dev. 6, 128Ck1292. Gehrung, S. & Snyder, M (1990). The SPA2 gene of Succhuromyces cerevisiue is important for pheromone-induced morphogenesis and efficient mating. J. Cell Biol. I 1 I , 145 1-1464. Gimeno, C., I,jungdahl, P. O., Styles, C. A. & Fink, G. R. (1993). Characterization ofSuccharomyces cerevisrue pseudohyphal growth. In: Dimorphic Fungi in Biology and Medicine (Vanden, H., Bossche, F., Odds, C. & Kerridge, D., Eds.) pp. 83-103. Plenum Press, New York. Gimeno, C. J . & Fink, G. R. (1994). Induction of pseudohyphal growth by overexpression ofPHDI, a Succhuromyces cerevisiue gene related to transcriptional regulators of fungal development. Mol Cell Biol. 14.2100-21 12. Gimeno,C. J.,Ljuiigdahl,P.O.,Styles,C.A . &Fink,G. R.(1992). Unipolarcelldivisionsin theyeastS. cerevisrue lead to filamentous growth: regulation by starvation and RAS. Cell 68, 1077-1090.

58

CHRISTINE COSTIGAN and MICHAEL SNYDER

Goud, B., Salminen, A,, Walworth, N. C. & Novick, P. J. (1988). A GTP-binding protein required for secretion rapidly associates with secretory vesicles and the plasma membrane in yeast. Cell 53, 753-768. Govindan, B. & Novick, P. (1993). The role ofMYO2 in vesicular transport in S. cerevisiae. Mol. Biol. Cell 4, 910a. Haarer, B. & Pringle, J. R. (1987). Immunofluorescence localization of the Saccharomyces cerevisiae CDCl2 gene product to the vicinity of the 10 nm filaments in the mother-bud neck. Mol. Cell. Biol. 7,3678-3687. Haarer, B. K., Lillie, S. H., Adams, A. E., Magdolen, V., Bandlow, W. & Brown, S . S. (1990). Purification of profilin from Saccharomyces cerevrsiae and analysis of profilin-deficient cells. J. Cell Biol. 110, 105-1 14. Haarer, B. K , Petzold, A,, Lillie, S. H. & Brown, S. S. (1994). IdentificationofMYOI,a second class V myosin gene in yeast. J. Cell Sci. 107, 1055-1064. Haarer, B. K., Petzold, A. S . & Brown, S. S. (1993). Mutational analysis of yeast profilin. Mol. Cell. Biol. 13, 7864-7873. Hagen, D. C., McCaffrey, G. & Sprague, G. F. (1986). Evidence the yeast STE3 gene encodes a receptor for the peptide pheromone a factor: Gene sequence and implications for the structure of the presumed receptor. Proc. Natl. Acad. Sci. USA 83, 1418-1422. Hall, A. (1990). The cellular functions of small GTP-binding proteins. Science 249,635-640. Hart, M. J., Eva, A., Evans, T., Aaronson, S. A. & Cerione, R. A. (1991). Catalysis ofguanine nucleotide exchange on the CDCI2Hs protein by the dbl oncogene product. Nature 354,3 11-3 14. Hartwell, L. H. (1971). Genetic control ofthe cell division cycle in yeast. IV. Genes controlling bud emergence and cytokinesis. Exp. Cell. Res. 69,265-276. Hartwell, L. H., Mortimer, R. K., Culotti, J. & Culotti, M. (1973). Genetic control of the cell division cycle in yeast. V. Genetic analysis of cdc mutants. Genetics 74,267-286. Hasek, J., Rupes, I., Svobodova, J. & Streiblova, E. (1987). Tubulin and actin topology during zygote formation of Saccharomyces cerevisiae. J. Gen. Microbiol. 133,3355-3363. Hayashibe, M. & Katohda, S. (1973). Initiation ofbudding and chitin-ring. J. Gen. Appl. Microbiol. 19, 23-39. Healy, A. M., Zolnierowicz, S., Stapleton, A. E., Goebl, M., DePaoli-Roach, A. A. & Pringle, J. R. (1991). CDCSS, a Saccharomyces cerevisiae gene involved in cellular morphogenesis: Identification, characterization, and homology to the B subunit of mammalian type 2A protein phosphatase. Mol. Cell. Biol. 1I , 5767-5780. Herman, P. K. & Emr, S. D. (1990). Characterization of VfS34,a gene required for vacuolar protein sorting and vacuole segregation in Saccharomuyces cerevisiae. Mol. Cell. Biol. 10, 6742-6754. Hicks, J. B., Strathern, J. N. & Herskowitz, 1. (1977). Interconversionofyeast mating types. 111. Action of the homothallism (HO) gene in cells homozygous for the mating type locus. Genetics 85, 395405. Holtzman, D. A,, Yang, S. & Drubin, D. G. (1993). Synthetic-lethal interactions identify two novel genes, SLAl and SLA2, that control membrane cytoskeleton assembly in Saccharomyces cerevisiae. J. Cell Biol. 122, 635-644. Hubbard, A. L. & Stieger, B. (1989). Biogenesis of endogenous plasmamembrane proteins in epithelial cells. Ann. Rev. Physiol. 5 1, 755-770. Huffaker, T. C., Thomas, J. H. & Botstein, D. (1988). Diverse effects of P-tubulin mutations on microtubule formation and function. J. Cell Biol. 106, 1997-2010. Hyman, A. (1989). Centrosome movement in the early divisions of Caenorhabditis elegans: A cortical site determining centrosome position. J. Cell Biol. 109, 1185-1 193. Hyman, A. A. & Steams, T. (1992). Spindle positioning and cell polarity. Curr. Biol. 2 , 4 6 9 4 7 1. lida, H., Sakaguchi, S . , Yagawa, Y. & Anraku, Y. (1990a).Cell cyclecontrol byCa++inSucchuromyces cerevisiae. J. Biol. Chem. 265,21216-21222.

Cell Polarity i n Yeast

59

lida, H., Yagawa, Y. & Anraku, Y. (1990b). Essential role for induced Ca++influx followed by [Ca++] rise in maintainingviability ofyeast cells late in the mating pheromone response pathway. J. Biol. Chem. 265, 13391-13399. hie, K., Takase, M., Lee, K., Levin, D., Araki, H., Matsumoto, K. & Oshima, Y. (1993). MKKl and MKK2, which encode Saccharomyces cerevisiae mitogen-activated protein kinase-kinase homologs, function in the pathway mediated by protein kinase C. Mol. Cell. Biol. 13, 3076-3083. Jackson, C. L. & Hartwell, L. H. (l990a). Courtship in S. cerevisiae: both cell types choose mating partners by responding to the strongest pheromone signal. Cell 63, 1039-1051. Jackson, C. L. & Hartwell, L. H. (1990b). Courtship in Saccharomyces cerevisiae: an early cell-cell interaction during mating. Mol. Cell. Biol. 10,2202-2213. Jackson, C. L., Konopka, J. B. & Hartwell, L. H. (1991). S. cerevisiae a-pheromone receptors activate a novel signal transduction pathway for mating partner discrimination. Cell 67, 389-402. Jacobs,C. W., Adams, A. E. M., Szaniszl0,P. J. & Pringle, J. R. (1988). Functionsofmicrotubules in the Sacchuromyces cerevisiae cell cycle. J. Cell Biol. 107, 1409-1426. Johannes, F.-J. & Gallwitz, D. (1991). Site-directed mutagenesis ofthe yeast actin gene: a test for actin function in vivo. EMBO J. 10, 3951-3958. Johnson, D. 1. & Pringle, J. R. (1990). Molecular characterization of CDC42 a Saccharomyces cerevisiae gene involved in the development of cell polarity. J. Cell Biol. 11 I , 143-152. Johnston, G. C., Prendergast, J. A. & Singer, R. A. (1991). The Saccharomyces cerevisiae MY02 gene encodes an essential myosin for vectorial transport of vesicles. J. Cell Biol. 113, 539-55 1. Johnston, G. C., Pringle, J. R. & Hartwell, L. H. (1977). Coordination ofgrowth with cell division in the yeast Succhuromyces cerevisiae. Exp. Cell Res. 105,79-98. Johnston, G. C. & Singer, R. A. (1980). Ribosomal precursor RNA metabolism and cell division in the yeast Saccharomyces cerevisiue. Mol. Gen. Genet. 178,357-360. Karpova, T. S., Lepetit, M. M. & Cooper, J. A. (1993). Mutations that enhance the cup2 null mutant phenotype in Saccharomyces cerevisiue affect the actin cytoskeleton, morphogenesis, and pattern of growth. Genetics 135,693-709. Kato, K. (1991). Sequence analysis oftwenty mouse brain cDNA clones selected by specificexpression patterns. J. Neurosci. 2, 7 0 6 7 I I . Kaziro, Y. (1978). The role of guanosine 5'-triphosphate in polypeptide chain elongation. Biochim. Biophys. Acta 505,95-127. Kilmartin, J. V. &Adams,AE. M. (1984). Structuralrearrangementsoftubulinandactinduringthecell cycle of the yeast Saccharomyces. J. Cell Biol. 98,922-933. Kim, H. B., Haarer, B. K. & Pringle, J. R. (1991). Cellular Morphogenesis in the Saccharomyces cerevisiae cell cycle: localization of the CDC3 gene product and the timing of events at the budding site. J. Cell Biol. 112, 535-544. Koch, C., Moll, T., Neuberg, M., Ahorn, H. & Nasmyth, K. (1993). A role for the transcription factors Mbpl and Swi4 in progression from G I to S Science 261, 1551-1557. Konopka, J. B., Jenness, D. D. & Hartwell, L. H. (1988). TheC-terminus o f t h e S . cerevisiae a-pheromone receptor mediates an adaptive response to pheromone. Cell 54, 609-620. Kubler, E. & Riezman, H. (1993). Actin and fimbrin are required for the internalization step of endocytosis in yeast. EMBO J. 12,2855-2862. Kurjan, J. (1993). The pheromone response pathway in Saccharomyces cerevisiae. Ann. Rev. Genet. 27, 147-179. Kuznetsov, S. A., Langford, G. M. & Weiss, D. G. (1992). Actin-dependent organelle movement in squid axoplasm. Nature 356, 722-725. Lappalainen, P. & Drubin, D. G (1997) Cofilin promotes rapid actin filament turnover in vivo. Nature 389,211-214.

60

CHRISTINE COSTICAN and MICHAEL SNYDER

Leberer, E., Dignard, D., Harcus, D., Thomas, D. Y. & Whiteway, M. (1992). The protein kinase homologue STE20p is required to link the yeast pheromone response G-protein py subunits to downstream signalling components. EMBO J. 1 I , 48 154824. Lee, K., h e , K., Gotoh, Y., Watanabe, Y., Araki, H.,Nishida, E., Matsumoto, K. & Levin, D. (1993). A yeast mitogen-activated protein kinase homolog (Mpklp) mediates signalling by protein kinase C Mol. Cell. Biol. 13, 3067-3075. Lee, K. & Levin, D. (1992). Dominant mutations in a gene encoding a putative protein kinase (ECKI) bypass the requirement for a Succhuromyces cerevisiue protein kinase C homolog. Mol. Cell. Biol. 12, 172-182. Leevers, S. J., Paterson, H. F. & Marshall, C. J. (1994). Requirement for Ras in Raf activation is overcome by targeting Raf to the plasma membrane. Nature 369,411414. Levin,D. &Errede,B.(1993).AmultitudeofMAPkinaseactivationpathways.J.NIHRes. 5,49-52. Levin, D., Fields, F. O., Kunisawa, R., Bishop, J. M. & Thomer, J. (1990). A candidate protein kinaseC gene, PKCl, is required for the S. cerevisiue cell cycle. Cell 62, 213-224. Levin, D. E. & Bartlett-Heubusch, E. (1992). Mutants in the S. cerevisiue PKCI gene display a cell cycle-specific osmotic stability defect. J. Cell Biol. 116, 1221-1229. Lew, D. J, & Reed, S. I . (1993). Morphogenesis in theyeast cell cycle: regulation by Cdc28 and cycling. J. Cell Biol. 120, 1305-1320. Lillie, S. H. & Brown, S. S. (1992). Suppression o f a myosin defect by a kinesin-related gene. Nature 356,358-361. Lillie, S. H. & Brown, S. S. (1994). Immunofluorescence localization of the unconventional myosin, MyoZp, and the putative kinesin-related protein, Smy Ip, to the same regions of polarized growth in Succhuromyces cerevisiue. J. Cell Biol. 125,825-842. Lipke, P. N., Taylor, A. & Ballou, C. E. (1976). Morphogenic effects of a-factor on Succhuromyces cerevisiue a cells. J. Bacteriol. 127, 610-618. Liu, H. & Bretscher, A. (1989a). Disruption of the single tropomyosin gene in yeast results in the disappearance of actin cables from the cytoskeleton. Cell 57,233-242. Liu, H. & Bretscher, A. (1989b). Purification of tropomyosin from Succhuromyces cerevisiue and identification of related proteins in Schizosucchuromyces and Physurum. Proc. Nat. Acad. Sci. U.S.A. 86,90-93. Liu, H. & Bretscher, A. (1992). Characterization of TPMl disrupted yeast cells indicates an involvement of tropomyosin in directed vesicular transport. J. Cell Biol. 118,285-299. Liu, H., Styles,C. A. & Fink, G. R. (1 993). Elementsofthe yeast pheromone response pathway required for filamentous growth of diploids. Science 262, 1741-1744. Lukins, H. B., Tate, J. R., Saunders, G. W. & Linnane, A. W. (1973). Mitochondria1recombination: the segregation of parental and recombinant mitochondria1 genotypes during vegetative division of yeast. Mol. Gen. Genet. 120, 17-25. Lutz, D. A,, Hamaguchi, Y. & Inoue, S. (1988). Micromanipulation studies of the asymmetric positioning of the maturation spindle in Chueropferus sp. oocytes. I. Anchorage of the spindle to the cortex and migration of a displaced spindle. Cell Motil. Cytoskel. 1 I , 83-96. Madaule, P., Axel, R. & Myers, A. M. (1987). Characterization oftwo members of the rho gene family from the yeast Succhuromyces cerevisiue. Proc. Natl. Acad. Sci. USA 84,779-783. Madden, K., Costigan, C. & Snyder, M. (1992). Cell polarity and morphogenesis in Succhuromyces cerevisiue. Trends Cell Biol. 2,22-29. Madden, K . & Snyder, M. (1992). Specification of sites of polarized growth in Succhuromyces cerevisiue and the influence of external factors on site selection. Mol. Biol. Cell 3, 1025-1 035. Magdolen, V., Oechsner, U., Muller, G. & Bandlow, W. (1988). The intron-containing gene for yeast profilin (PFYf) encodes avital function. Mol. Cell. Biol. 8, 5108-51 15. Matile, P., Moor, H. & Robinow, C. F. (1969). Yeast cytology. In: The Yeasts(Rose, A. H., & Harrison, J . S., eds.). pp. 219-302, Academic Press, New York.

Cell Polarity in Yeast

61

Matsui, Y. & Toh-e, A. (1992a). Isolation and characterization of two novel ras superfamily genes in Saccharomyces cerevurae. Gene 114,4349. Matsui, Y. & Toh-e, A. (1992b). Yeast RHO3 and RHO4 ras super-family genes are necessary for bud growth, and their defect is suppressed by a high dose of bud formation genes CDC42 and BEMI. Mol. Cell. Biol. 12, 569e5699. Mayer, M. L., Caplin, B. E. & Marshall, M. S. (1992). CDC43 and RAM2 encode the polypeptide subunits of a yeast type I protein geranylgeranyltransferase.J. Biol. Chem. 267,20589-20593. Mazzoni, C., Zarzov, P., Rambourg, A. & Mann,C. (1993). SLTZ(MPK1) MAP kinase homolog is involved in polarized cell growth in Saccharomyces cerevisiae. J. Cell Biol. 123, 182 1-1 823.

McCaffrey, M., Johnson, J. S., Goud, B., Myers, A. M., Rossier, J., Poppoff, M. R., Madaule, P. & Boquet, P. (1991). The small GTP-binding protein Rholp is localizedon the Golgi apparatus and post-Golgi vesicles in Saccharomyces cerevisiae. J. Cell Biol. 115,309-3 19. McConnell, S. J., Stewart, L. C., Talin, A. & Yaffe, M. P. (1990). Temperature-sensitive yeastmutants defective in mitochondrial inheritance. J. Cell Biol. 11 1,967-976. McConnell, S. J. & Yaffe, M. P. (1992). Nuclear and mitochondrial inheritance in yeast depends on novel cytoplasmic structures defined by the MDMl protein. J. Cell Biol. 118,385-395. McConnell, S. J. & Yaffe, M. P. (1993). Intermediate filament formation by a yeast protein essential for organelle inheritance. Science 260,687-689. Meluh, P. B. &Rose, M. D. (1990). KAR3, a kinesin-related gene required for yeast nuclear fusion. Cell 60, 1029-1 04 1.

Michelitch, M. & Chant, J. (1996). A mechanism of Budlp GTPase action suggested by mutational analysis and immunolocalization. Curr. Biol. 6, 4 4 W 5 4 . Miyamoto, S., Ohya, Y., Ohsumi, Y. & Anraku, Y. (1987). Nucleotide sequence ofthe CLS4 (CDC24) gene of Saccharomyces cerevisiae. Gene 54, 125-1 32. Miyamoto, S., Ohya, Y., Sano, Y., Sakaguchi, S., Iida, H. & Anraku, Y. (1991). A DBL-homologous region ofthe yeast CLSWCDC24gene product is important for calcium-modulated bud assembly. Biochem. Biophys. Res. Comm. 181,604610. Moon, A. L ,Janmey, P. A,, Louie, K. A. & Drubin, D. G. (1993). Cofilin is an essential component of the yeast cortical cytoskeleton. J. Cell Biol. 120,421-435. Moore, S. A. (1983). Comparison of dose-response curves for a-factor-induced cell division arrest, agglutination, and projection formation of yeast. J . Biol. Chem. 258, 13848-13856. Mulholland, J., Preuss, D., Moon, A,, Wong, A,, Drubin, D. & Botstein, D. (1994). Ultrastructureofthe yeastactincytoske1etonanditsassociationwiththeplasmamembrane.J.Cell Biol. 125,381-391. Nakayama, N., Miyajima, A. & Arai, K. (1985). Nucleotide sequences of STE2 and STE3, cell type-specific sterile genes from Saccharomyces cerevisiae. EMBO J . 4,2643-2648. Nasmyth, K. A. (1982). Molecular genetics of yeast mating type. Ann. Rev. Genet. 16,439-500. Nelson, W. J. (1992). Regulation of cell surface polarity from bacteria to mammals. Science 258, 948-95 5.

Neufeld, T. P. & Rubin, G. M. (1994). The Drosophila peanut gene is required for cytokinesis and encodes a protein similar to yeast putative bud neck filament proteins. Cell 77,371-379. Ng, R. & Abelson, J. (1980). Isolation of the gene for actin in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 77,3912-3916. Nonaka, H., Tanaka, K., Hirano, H., Fujiwara, T., Kohno, H., Umikawa, M., Mino, A., & Takai, Y. (1995). A downstream target of RHO1 small GTP binding protein is PKC 1, a homolog of protein kinase C, which leads to activation of the MPKl kinase cascade in Saccharomyces cerevisiae. EMBO J. 14,5931-5938. Novick, P. & Botstein, D. (1985). Phenotypicanalysisoftemperature-sensitiveyeastactinmutants.Cel1 40,405-416.

Novick, P. & Brennwald, P. (1993). Friends and family: the role ofthe Rab GTPases in vesicular traffic. Cell 75, 597-601.

62

CHRISTINE COSTIGAN and MICHAEL SNYDER

Novick, P., Field, C. & Schekman, R. (1980). ldentification of 23 complementation groups required for post-translational events in the yeast secretory pathway. Cell 21, 205-215. Novick, P., Osmond, B. C. & Botstein, D. (1989). Suppressors ofyeast actin mutations. Genetics 121, 659474. Ohsumi, Y. & Anraku, Y. (1985). Specific induction of Ca++ transport activity in MATa cells of Succhuromycescerevisiaeby amating pheromone, a-factor. J. Biol. Chem. 260,10482-10486. Ohya, Y. & Botstein, D. (1994). Diverse essential functions revealed by complementing yeast calmodulin mutants. Science 263,963-966. Ohya, Y., Goebl, M., Goodman, L. E., Petersen-Bjorn, S., Friesen, J . D., Tamanoi, F. & Anraku, Y, (1991). Yeast CALI is astructural and functional homologue to the DPRl (RAM) gene involved in rus processing. J. Biol. Chem. 266, 1235G12360. Ohya, Y., Miyamoto, S., Ohsumi, Y. & Anraku, Y. (1986a). Calcium-sensitive cls4 mutant of Succhuromyces cerevisiue with a defect in bud formation. J. Bacteriol. 165,28-33. Ohya, Y., Ohsumi, Y. & Anraku, Y. (1986b). Isolation and characterization of calcium-sensitive mutants ofSuccharomyces cerevisiue. J. Gen. Microbiol. 132,979-988. Ohya, Y., Qadota, H., Anraku, Y., Pringle, J. R. & Botstein, D. (1993). Suppression of yeast geranylgeranyl transferase 1 defect by alternative prenylation of two target GTPases, Rholp and Cdc42p. Mol. Biol. Cell 4, 1017-1025. Page, B. & Snyder, M. (1993). Chromosome segregation in yeast. Ann. Rev. Microbiol.47,201-23 1. Palmer, R. E., Sullivan,D. S., Huffaker, T. & Koshland, D. (1 992). Role ofastral microtubules and actin in spindle orientation and migration in the budding yeast, Succhuromycescereviszue.J . Cell Biol. 119, 583-593. Paravicini,G., Cooper, M., Friedli, L., Smith, D. J., Carpentier,J.-L., Klig, L. S. &Payton, M. A. (1992). The osmotic integrity ofthe yeast cell requires a functional PKCl gene product. Mol. Cell. Biol. 12,48964905. Park, H. O., Bi, E., Pringle, J. R., & Herskowitz, 1. (1997). Two active states of the Ras-related Budl/Rsrl protein bind to different effectors to determine yeast cell polarity. Proc. Natl. Acad. Sci. USA 94,44634468. Park, H. O., Chant, J. & Herskowitz, I. (1993). BUD2 encodes a GTPase-activating protein for Budl/Rsrl necessary for proper bud-site selection in yeast. Nature 365,269-274. Peter, M., Gartner, A,, Horecka, J., Ammerer, G. & Herskowitz, I. (1993). FAR1 links the signal transduction pathway to the cell cycle machinery in yeast. Cell 73, 747-760. Peterson, J . B. & Ris, H. (1976). Electron microscopic study ofthe spindle and chromosome movement in the yeast Succhmomyces cerevisioe. J. Cell Sci. 22,219-242. Prendergast, J. P., Murray, L. E., Rowley, A. R., Carruthers, D. R., Singer, R. A. &Johnston, G. C. (1990). Size selection identifies new genes that regulate Succharomyces cerevrsiae cell proliferation. Genetics 124, 81-90. Preuss, D., Mulholland, J., Franzusoff, A,, Segev, N. & Botstein, D. (1992). Characterization of the SucchuromycesGolgi complex through the cell cycle by immunoelectron microscopy. Mol. Biol. Cell 3,789-803. Preuss, D., Mulholland, J., Kaiser, C. A,, Orlean, P., Albright, C., Rose, M. D., Robbins, P. W. & Botstein, D. (1991). Structure of the yeast endoplasmic reticulum: localization of ER proteins using immunofluorescence and immunolocalization microscopy. Yeast 7, 891-91 1. Primig, M., Sockanathan, S., Auer, H. & Nasmyth, K. (1992). Anatomy of a transcription factor important for the Start of the cell cycle in Succhuromyces cerevrsiue. Nature 358, 593-597. Pringle, J . R., Bi, E., Harkins, H. A,, Zahner, J . E., DeVirgilio, C., Chant, J., Corrado. K. & Fares. H. (1995) Establishment of cell polarity in yeast Cold Spring Harbor Sym Quant Biology 60, 729-744 Pringle, J. R. & Hartwell, L H. (1981). The Succhuromyces cerevisiae cell cycle. In: The Molecular Biology ofthe Yeast Succhuromyces: Life Cycle and Inheritance (Strathern, J. N., Jones, E. W. Broach, J. R., eds.). pp. 97-142. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.

Cell Polarity in Yeast

63

Qadota H., Ishii, I., Fujiyama, A,, Ohya, Y. & Anraku, Y . (1992). R H O gene products, putative small GTP-binding proteins, are important for activation of the CALI/CDC43 gene product, a protein geranylgeranyl-transferase in Succhuromyces cerevisiue. Yeast 8,735-74 1. Qadota, H., Python, C. P., Inoue, S. B., Arisawa, M., Anraku, Y., Zheng, Y., Wananabe,T., Levin, D. E., & Ohya, Y. (1996). IdentificationofyeastRholpGTPaseas aregulatorysubunitof 1,3-B-Glucan Synthase. Science 272,279-28 1. Quinn, M. T., Parkos, C. A., Walker, L., Orkin, S. H., Dinauer, M. C. & Jesaitis, A. J. (1989). Association of a Ras-related protein with cytochrome b of human neutrophils. Nature 342, 198-200. Rappaport, R. (1986). Establishment of the mechanism of cytokinesis in animal cells. Int. Rev. Cytol. 105,245-28 1. RayChaudhuri, D. & Park, J. T. (1992). Escherichia cold cell-division geneJisZ encodes a novel GTP-binding protein. Nature 259,251-254. Raymond, C. K., Howald-Stevenson, I., Vater, C. A. &Stevens, T. (1992). Morphologicalclassification of the yeast vacuolar protein sorting mutants: evidence for a prevacuolar compartment in Class E vps mutants. Mol. Biol. Cell 3, 1389-1402. Raymond, C. K., O’Hara, P. J., Eichinger, G., Rothman, J. &Stevens, T. H. (1990). Molecular analysis of the yeast VPS3 gene and the role of its product in vacuolar protein sorting and vacuolar segregation during the cell cycle. J. Cell Biol. 11 1, 877-892. Read, E. B., Okamura, H. H. & Drubin, D. G. (1992). Actin- and tubulin-dependent functions during Sacchuromyces cerevisiae mating projection formation. Mol. Biol. Cell 3,429-444. Reid, B. J. & Hartwell, L. H. (1977). Regulationofmatinginthe cell cycleofSaccharomycescerevisiue. J. Cell Biol. 75,355-365. Rhodes, N., Connell, L. & Errede, B.(1990). STEl 1 is a protein kinase required for cell-type-specific transcription and signal transduction in yeast. Genes Dev. 4, 1862-1874. Richardson, H. E., Wittenberg, C., Cross, F. & Reed, S. I. (1989). An essential G I function for cyclin-like proteins in yeast. Cell 59, 1127-1 133. Ridley, A. J. & Hall, A. (1992). The small GTP-binding protein Rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell 70,389-399. Roemer, T., Vallier, L., Sheu, Y.-J., & Snyder, M. (1998). The Spa2p-related protein Sphlp is important for polarized growth in yeast. J. Cell Sci. I 1 I , 479494. Roemer, T., Vallier, L. G., & Snyder, M. (1996). Selection of polarized growth sites in yeast. Trends Cell Biol. 6,434-441. Ron, D.,Zannini, M., Lewis, M., Wickner, R. B., Hunt, L. T., Graziani, G., Tronick, S. R., Aaronson, S. A. & Eva, A . (1991). A region ofproto-Dbl essential for its transforming activity shows sequence similarity to a yeast cell cycle gene, CDC24, and the human breakpoint cluster gene, BCR. New Biol. 3,372-379. Ruggieri, R., Bender, A,, Matsui, Y., Powers, S., Takai, Y., Pringle, J. R.J. Matsumoto, K. (1992). RSRI, a ras-like gene homologous to Krev-1 (smg2lA/raplA): role in the development of cell polarity and interactions with the Ras pathway in Sacchuromyces cerevuzae. Mol. Cell. Biol. 12,758-766. Saavedra-Molina, A,, Villalobos, R. & Borbolla, M. (1983). Calcium uptake during the cell cycle of Succhuromyces cerevisiae FEBS Lett. 160, 195-197. Sanders, S. L. & Herskowitz, I. (1996). The Bud4 protein of yeast, required for axial budding, is localized to the mother-bud neck in a cell cycle-dependent manner. J. Cell Biol. 134,413-427. Salrninen, A. & Novick, P. (1987). A ras-like protein is required for a post-Golgi event in yeast secretion. Cell 49, 527-538. Salminen, A. & Novick, P. (1989) The Secl5 protein responds to the function of the GTP binding protein, Sec4, to control vesicular traffic in yeast. J. Cell Biol. 109, 1023-1036. Schekrnan, R. & Brawley, V. (1979). Localized deposition ofchitin on the yeast cell surface in response to mating pheromone. Proc. Nat. Acad. Sci. U S A . 76,645-649.

64

CHRISTINE COSTICAN and MICHAEL SNYDER

Schekman, R. & Novick, P. (1981). The secretory process and yeast cell-surface assembly. In: The Molecular Biology of the Yeast Succhuromyces: Life Cycle and Inheritance (Strathern, J . N., Jones, E., & Broach, J., Eds.), pp. 361-398. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Scherr, G. H. & Weaver, R. H. (1953). The dimorphism phenomenon in yeasts. Bacteriol. Rev. 17, 5 1-92. Schwob, E. & Martin, R. P. (1992). New yeast actin-like gene required late in the cell cycle. Nature 355, 179-1 82. Segall, J. E. (1993). Polarization of yeast cells in spatial gradients ofa-factor. Proc. Natl. Acad. Sci. 90, 8332-8336. Shinjo,K.,Koland, J. G.,Hart,M. J.,Narasimhan,V., Johnson,D. I . , Evans,T. &Cerione,R. A. (1990). Molecular cloning of the gene for the human placental GTP-binding protein, Gp (G25K): identification of this GTP-binding protein as the human homolog of the yeast cell division cycle protein, CDC42. Proc. Natl. Acad. Sci. USA 87,9853-9857. Shortle, D., Haber, J. E. & Botstein, D. (1982). Lethal disruption ofthe yeast actin gene by integrative DNA transformation. 217,371-373. Shortle, D., Novick, P. & Botstein, D. (1984). Construction and genetic characterization of temperature-sensitive mutant alleles of the yeast actin gene. Proc. Natl. Acad. Sci. USA 81, 48894893. Sloat, B. & Pringle, J. (1978). A mutant of yeast defective in cellular morphogenesis. Science 200, 1171-1173. Sloat, B. F., Adams, A. & Pringle, J. R. (1981). Roles of the CDC24 gene product in cellular morphogenesis during the Succhuromyces cerewisiae cell cycle. J. Cell Biol. 89,395405. Snyder, M. (1989). The SPA2 protein of yeast localizes to sites of cell growth. J. Cell Biol. 108, 14 19-1429. Snyder, M., Gehrung, S. & Page, B. D. (1991). Studies concerning the temporal and genetic control of cell polarity in Sacchuromyces cerevisiue. J. Cell Biol. 114,515-532. Song, O., Dolan, J. W., Yuan, Y. 0. &Fields, S. (1991). Pheromone-dependent phosphorylation of the yeast STEl2 protein correlates with transcriptional activation. Genes & Dev. 5, 741-750. Sprague, G. F. & Thorner, J. (1992). Pheromone response and signal transduction during the mating process of Succharomyces cerevisiue. In: The Molecular Biology of the Yeast Succharomyces (Broach, J. R., Pringle, J. R. &Jones, E. W., Eds.), pp. 657-744. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Stevens, B. (1981). Mitochondria1 structure. In: The Molecular Biology of the Yeast Succharomyces cerevisiae: Life Cycle and Inheritance (Strathern, J . N., Jones, E. W. & Broach, J . R., Eds.), pp. 471-504. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Stevenson, B. J., Rhodes, N., Errede, B. & Sprague, G . F. (1992). Constitutive mutants of the protein kinase STEl 1 activate the yeast pheromone response pathway in the absence of G protein. Genes & Dev. 6 , 1293-1304. Strazdis, J. R. & MacKay, V. L. (1983). Induction ofyeast mating pheromone a-factor by a cells. Nature 305,543-545. Streiblova, E. (1970). Study of scar formation in the life cycle of heterothallic Saccharomyces cerevisiae. Canad. J. Microbiol. 16, 827-831. Strome, S. (1993). Determination of cleavage planes. Cell 72,3-6. Sullivan, D. S. & Huffaker, T. C. (1992). Astral microtubules are not required for anaphase B in Saccharomyces cerevisiae. J. Cell Biol. 119, 379-388. Sun, G. H., Ohya, Y. & Aiiraku, Y. (1992). Yeast calmodulin localizes to sites of cell growth. Protoplasma 166, 110-113. Thompson, P. W. & Wheals, A. E. (1980). Asymmetrical division of Succharomyces cerevisiae ~n glucose-limited chemostat culture. J. Gen. Microbiol. 121,401409.

Cell Polarity in Yeast

65

TerBush. D. R.. & Novick, P. (1995). Sec6, Sec8, and Secl5 are components of amultisubunitcomplex which localizes to small bud tips in Succharomycescerevisiae. J. Cell Biol. 130,299-312. Tkacz, J. S. & Lampen, J. 0.(1972). Wall replication inSucchuromycesspecies: use offluorescein-conjugated concanavalin A to reveal the site of mannan insertion. J. Gen. Microbiol. 72,243-247. Tkacz, J. S . & MacKay, V. L. (1979). Sexual conjugation in yeast: Cell surface changes in response to the action of mating hormones. J. Cell Biol. 80,326333. Torres, L., Martin, H.,Garcia-Saez, M. l.,Arroyo, J., Molina, M., Sanchez, M. &Nombela,C. (1991). A protein kinase gene complements the lytic phenotype ofSacchuromyces cerevisiae lyf2 mutants. Mol. Microbiol. 5,2845-2854. Valdivieso, M. H., Sugimoto, K., Jahng, K. Y., Fernandes, P. M. & Wittenberg, C. (1993). FAR1 is required for posttranscriptional regulation of CLN2 gene expression in response to mating pheromone. Mol. Cell. Biol. 13, 1013-1022. Vallen, E. A., Scherson, T. Y., Roberts, T., Zee, K . V. & Rose, M. D. (1992). Asymmetric mitotic segregation of the yeast spindle pole body. Cell 69, 505-515. Valtz, N. & Herskowitz, I . (1996). Pea2 protein o f yeast is localized to sites of polarized growth and is required for efficient mating and bipolar budding. J . Cell Biol. 135, 725-739. Vinh, D. B. N., Welch, M. D., Corsi, A. K., Wertman, K. F. & Drubin, D. G. (1993). Genetic evidence for functional interactions between actin noncomplementing (Anc) gene products and actin cytoskeletal proteins in Saccharomyces cerevisiue. Genetics 135,275-286. Vojtek, A,, Haarer, B., Field, J., Gerst, J., Pollard, T. D., Brown, S. & Wigler, M. (1991). Evidence for a functional link between profilin and CAP in the yeast S. cerevisiae. Cell 66,497-505. Walworth, N. C., Goud, B., Kabcenell, A. K. & Novick, P. J. (1989). Mutational analysis of SEC4 suggests a cyclical mechanism for the regulation ofvesicular traffic. EMBO J. 8, 1685-1693. Wang, J., Suzuki, N. & Kataoka, T. (1992). The 70-kilodalton adenylyl cyclase-associatedproteinIS not essential for interaction of Sacchuromyces cerevisiue adenylyl cyclase with RAS proteins. Mol. Cell. Biol. 12,49374945. Watts, F. Z., Shiels, G . & Orr, E. (1987). The yeast MY01 gene encoding amyosin-like protein required for cell division. EMBO J. 6, 3499-3505. Weisman, L. S., Bacallao, R. & Wickner, W. (1987). Multiple methods ofvisualizing the yeast vacuole permit evaluation of its morphology and inheritance during the cell cycle. J. Cell Biol. 105, 1539-1 547. Weisman, L. S., Emr, S. D. & Wickner, W. T. (1990). Mutants ofSaccharomycescerevisiaethat block intervacuole vesicular traffic and vacuole division and segregation. Proc. Natl. Acad. Sci. USA 87, 1076-1080. Weisman, L. S. & Wickner, W. (1988). lntervacuole exchange in the yeast zygote: a new pathway in organelle communication Science 241, 589-591 Weisman, L S. & Wickner, W. (1992). Molecular characterizationof VACI, agenerequired forvacuole inheritance and vacuole protein sorting. J . Biol. Chem. 267, 618-623. Welch, M D. & Drubin, D. (1994). A nuclear protein with sequence similarity to proteins implicated in human acute leukemias is important for cellular morphogenesis and actin cytoskeletal function in Saccharomyces cerevisiae. Mol. Biol. Cell 5, 61 7 4 2 2 . Welch, M. D., Holtzman, D. A. & Drubin, D. G. (1994). The yeast actin cytoskeleton. Curr. Opin. Cell Biol. 6, 110-1 19. Welch, M. D , Vinh, B. B. N., Okamura, H. H. & Drubin, D. G. (1993). Screens for extragenicmutations that fail to complement act1 alleles identify genes that are important for actin function in Saccharomyces cerevisiue. Genetics 135,265-274. Wertman, K F., Drubin, D. G. & Botstein, D. (1992). Systematic mutational analysis ofthe yeast ACT1 gene. Genetics 132,337-350. Whitters, E. A., Cleves, A. E., McGee, T. P., Skinner, H. B. & Bankaitis, V. A. (1993). SAClp is an integral membrane protein that influences the cellular requirement for phospholipid transfer protein function and inositol in yeast. J Cell Biol. 122, 79-94.

66

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Yamochi, W., Tanaka, K., Nonaka, H., Maeda, A., Musha, T. & Takai, Y. (1994). Growth site localization of Rho1 small GTP-binding protein and its involvement in bud formation in Succhuromyces cerevzsiue. J. Cell Biol. 125, iO77-1093. Yang, S., Ayscough, K. R. & Drubin, D. G. (1997). A role for the actin cytoskeleton ofSucchuromyces cerevisiue in bipolar bud-site selection. J. Cell Biol. 136, 111-123. Zheng, Y., Cerione, R. & Bender, A. (1994). Control of the yeast bud-site assembly GTPase Cdc42. J. Biol. Chem. 269,2369-2372. Ziman, M., O’Brien, J. M., Ouellette, L. A,, Church, W. R. & Johnson, D. I . (1991). Mutational analysis of CDC42Sc. a Succhuromyces cerevisiue gene that encodes a putative GTP-binding protein involved in the control of cell polarity. Mol. Cell. Biol. 11, 3537-3544. Ziman, M., Preuss, D., Mulholland, J., O’Brien, J. M., Botstein, D. &Johnson, D. I. (1993). Subcellular localization of Cdc42p, a Succhuromyces cerevisiue GTP-binding protein involved in the control ofcell polarity. Mol. Biol. Cell 4, 1307-1316.

CELL POLARITY AND MOUSE EARLY DEVELOPMENT

Tom P. Fleming, Elizabeth Butler, Jane Collins, Bhav Sheth, and Arthur E. Wild

I. Introduction

........................ A. Cell Polarity and Compaction B. Regulation of Cell Polarity . . . . . . . . . . . . . . . . . . . . . . . . ........ Stage in Epithelial Differentiation. . . . . . .

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Advances in Molecular and Cell Biology Volume 26, pages 67-94. Copyright 0 1998 by JAI Press Inc. All right of reproductionin any form reserved. ISBN: 0-7623-0381-6

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I.

INTRODUCTION

Cell polarity occurs for the first time during mouse embryogenesis at the 8-cell stage, about 2.5 days after fertilization. This event is critical for the formation of the blastocyst and its subsequent development. First, cell polarity initiates the program of differentiation of the trophectoderm epithelium which forms the wall of the blastocyst. This tissue generates the blastocoel, regulates vectorial exchange of metabolites with the embryo interior, constitutes the embryonic surface that engages in uterine attachment, and, after implantation, gives rise to the chorio-allantoic placenta. Second, cell polarity underlies the concurrent program of cell diversification in the embryo in which differentiative cell divisions lead to the formation and segregation of the earliest embryonic tissues, the outer trophectoderm and the enclosed inner cell mass (ICM) from which the entire fetus is derived. Cell polarity is therefore of fundamental importance as a developmental mechanism in mammals. In this chapter, we first review the cell biological characteristics of cell polarity during mouse cleavage and the consequences for blastocyst differentiation and tissue diversification. Second, we consider the influence of polarity at a molecular level with respect to the differentiation of multimolecular adhesive junctions in trophectoderm and the origin of differential gene expression in the embryo. Different aspects of early mouse development have been reviewed elsewhere recently (Kimber, 1990; Wiley et al., 1990; Cruz, 1992; Fleming, 1992; Fleming et al., 1992, 1993a, 1994; Gueth-Hallonet and Maro, 1992; Watson, 1992; Collins and Fleming, 1995).

II.

CYTOLOGICAL ASPECTS OF CELL POLARITY AND TISSUE SEGREGATION A.

Cell Polarity and Compaction

Fertilization of the mouse egg is followed by three reduction cleavage divisions to produce an embryo composed of 8 spherical, loosely-associated and nonpolarized blastomeres (Figure 1A). By this stage, the embryo has activated its own genome (2-cell stage; Flach et al., 1982), degraded nearly all maternal transcripts (Paynton et al., 1988), and has transcribed and translated most of the proteins required to initiate cell polarity and differentiation (Levy et al., 1986). The switch in blastomere phenotype from nonpolar to polar at the 8-cell stage is comprehensive in nature and coincides with the onset of cell-cell adhesion, these combined events being referred to as “compaction” since blastomere outlines become indistinct as adhesive contacts form (Figures 1B-D). Blastomere polarity at compaction is detectable both within the deeper cytoskeletal and cytoplasmic zones and within the cell cortex and membrane. Thus, microfilaments and microtubules polymerize predominantly in the apical (outer-facing) cytoplasm (Johnson and Maro, 1984; Houliston et al., 1987) al-

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Figure 7. (A-C) Scanningelectron micrographs, of &cell embryos following removal of the zona pellucida (from Fleming et al., 1986a).(A) before compaction has occurred, each blastomere is non-adhesive and uniformly microvillous.(B) after compaction, blastomeres adhere closely together and display a pole of microvilli.(C) compact embryo following exposure to calcium-free medium, causingloss of adhesion and showing apical pole of microvilli. (D)Transmission electron micrograph of one blastomere from a compact 8-cell embryo (from Fleming and Pickering, 1985) showing apical pole of microvilli (M), and clustering of endocytic organelles (E) in the apical cytoplasm. Arrowheads indicate position of adhesive, non-microvillous, basolateral membrane. Bar = 10pm.

though a sub-population of stable acetylated microtubules polarizes basally (Houliston and Maro, 1989). Actin-associated proteins polarize mostly in the apico-lateral region (Sober, 1983; Lehtonen et al., 1988; Slager et al., 1992), nuclei relocate to the basal cytoplasm (Reeve and Kelly, 1983), endosomes and clathrincoated vesicles redistribute from a nonpolar distribution and become localized mostly in the apical cytoplasm (Reeve, 1981;Fleming and Pickering, 1985;Mar0 et dl., 1985; Figure lD), while mitochondria become cortically localized (Batten et al., 1987). Polarization of cytoplasmic components is dependent upon cytoskeletal organization and is consistently modified or inhibited by reagents affecting microfilament or microtubule integrity (Johnson and Maro, 1985; Fleming et al., 1986a,b). Polarization of the cytocortex (membrane and underlying cytoskeleton) is intimately associated with the initiation of cell-cell adhesion mediated by the calcium-dependent cadherin, uvomorulin (E-cadherin), which becomes localized at cell-cell contact sites at compaction (Hyafil et al., 1980; Peyrieras et al., 1983; Vestweber et al., 1987). This redistribution may be promoted by increased stabili-

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zation (cytoskeletal anchorage?) of uvomorulin at contact sites and loss of stability at contact-free cell surfaces (Clayton et al., 1993). Uvomorulin appears to be the only mediator of cell adhesion at compaction although N-CAM (calciumindependent neural cell adhesion molecule) is also present in embryos at this time (Kimber et al., 1994). The newly-formed adhesive basolateral cell surfaces form functional gap junctions (Ducibella and Anderson, 1975; Magnuson et al., 1977; Lo and Gilula, 1979; McLachlin et al., 1983; Goodall and Johnson, 1984; Pratt, 1985) and, at their apicolateral border, focal tight junctions emerge (Ducibella and Anderson, 1975; Magnuson et al., 1977; Pratt, 1985) containing the marker protein ZO-1 (Heming et al.. 1989). Most significantly, a distinct apical cytocortex is formed at compaction, comprising a pole of microvilli (Ducibella et al., 1977;Handyside, 1980; Reeve and Ziomek, 1981; Figure 1). Unlike cytoplasmic polarity, the essential features of the apical microvillous pole can form and be maintained in the presence of cytoskeleton-disrupting agents (Johnson and Maro, 1984, 1985; Fleming et al., 1986a,b). Taken together, 8-cell blastomeres at compaction reorganize into a polarized proto-epithelial phenotype that marks the beginning of trophectoderm differentiation. We next consider the mechanisms by which the spatial patterning of cell polarity at compaction and the timing of its expression in the fourth cell cycle may be controlled. B.

Regulation of Cell Polarity

What is the role of cell adhesion in the establishment of blastomere polarity? In undisturbed 8-cell embryos, the apicobasal axis of polarity develops with respect to cell-cell contact sites (i.e., apical microvilli form opposite the contact points) and is not predetermined before the fourth cell cycle (Ziomek and Johnson, 1980; Johnson and Ziomek, 1981a; Figure 2). Culturing embryos in the absence of calcium or in the presence of antibodies against the ectodomain of uvomorulin, or in cytochalasin, inhibits or reverses the adhesive component of compaction (Pratt et al., 1982; Shirayoshi et al., 1983; Johnson et al., 1986; Figures lC, 2C) as do peptides containing the cadherin HAV recognition sequence (Blaschuk et al., 1990). Under conditions of uvomorulin neutralization, cell polarity can still occur for most components but does so over a more protracted time period and usually displays a random axis with respect to sites of cell-cell contact (Fleming et al., 1986a,b, 1989; Johnson et al., 1986; Figure 2C). The capacity of blastomeres to polarize is therefore “programmed” independently from their capacity to initiate adhesion, although the latter process would normally act to catalyze and synchronize cell polarity and to define in a permissive way the orientation of its axis (Johnson et aI., 1986). What is the relationship between the different features of cell polarity that are established at compaction following receipt of the “permissive” inductive signal mediated by uvomorulin adhesion? A number of experimental approaches indicate that the apical cytocortical domain of the cells (essentially where the apical pole of

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non-polar

polar

d

Figure2 (A-C) Disaggregated8-cell blastomeres labelled with FITC-concanavalinA to reveal distribution of microvilli (from Fleming et al., 1986a). (A) before compaction, staining is uniform.(B) after compaction, staining (microvillous pole) i s localised opposite the point of intercellular adhesion.(C) blastomeres exposed to cytochalasin fail to adhere together (see b) but can still generate microvillous polarity, along a random axis not related to the point of cell-cell contact (top blastomere). (D) Cell adhesion leads to conversion of 8-cell blastomeres from non-polar to polar phenotype; see text for consideration of mechanisms. Bar = IOpm.

microvilli forms) acts as a stable "memory" of the axis of polarity throughout the period of trophectoderm differentiation and is responsible for organizing polarization within the deeper cytoplasm (reviewed in Fleming, 1992; Figure 2D). Thus, in experimental conditions where microvillous polarity develops in the absence of cytoplasmic polarity (e.g., cytochalasin treatment, disrupting microfilaments, see earlier), returning embryos to normal medium permits cytoplasmic polarity to occur and, significantly, along the axis already defined by the pole of microvilli (Johnson and Maro, 1985). Similarly, cytoplasmic polarity, but not cortical polarity, is dissipated as 8-cell blastomeres enter mitosis, but is reestablished in the next interphase again along the axis defined by the stable microvillous pole (Fleming and Pickering, 1985; Maro et al., 1985; Johnson et al., 1988). Also, in heterokaryons formed by fusion of polarized 8- or 16-cell blastomeres with nonpolar 4-cell blastomeres, the apical cytocortex of the polarized cell induced polarization of cytoplasmic components derived from the 4-cell (Wiley and Obasaju, 1988). The question of how a contact-mediated signal might lead to a polarized cytocortical organization remains elusive. One possibility is that signal transduction

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mechanisms involving protein kinases may be involved in the propagation of a basal contact signal towards the apical domain in the plane of the membrane (reviewed in Fleming, 1992; Figure 2D; also see below). Alternatively, or in addition, transcellular ion currents, carried largely by Na+ ions and generated by the restricted membrane distribution of appropriate ion transporters, may be responsible for initiating a polarized state (Nuccitelli and Wiley, 1985; Wiley and Obasaju, 1988, 1989; reviewed in Wiley et al., 1990; Figure 2D). The identification of a Na+/glucose cotransporter at the apical microvillous domain of blastomeres from compaction onwards may be significant in initiating a Na+-based transcellular ion current (Wiley et al., 1991). For most events of compaction to proceed, neither proximate transcription nor translation are required (Kidder and McLachlin, 1985; Levy et al., 1986). suggesting that the timing of compaction at the 8-cell stage occurs through posttranslational modification of existing proteins. This is borne out in biogenetic studies on the uvomorulin adhesion system. Uvomorulin expression occurs both in unfertilized eggs and throughout cleavage, ruling out its biosynthesis as a mechanism initiating adhesion at compaction (Vestweber et al., 1987; Clayton et al., 1993). Prior to fertilization, however, uvomorulin is not transported to the cell surface, this is achieved from the zygote stage onwards (Clayton et al., 1993). Uvomorulin interacts with catenin proteins at its cytoplasmic tail which mediate the interaction with actin filaments, a requirement for adhesive function in tissue culture cells (reviewed in Takeichi, 1991; Geiger and Ayalon, 1992; Grunwald, 1993). Both a- and p-catenin are detectable during early development by immunoblotting and immunocytochemistry before compaction, indicating that their expression does not regulate uvomorulin adhesion at compaction (Ohsugi et al., 1996; J. Lewthwaite and T. Fleming, manuscript in preparation). It has been proposed that modifications which initiate compaction are prevented from occurring until the 8-cell stage by the synthesis of arestraining factor since, in the absence of protein synthesis, compaction takes place prematurely (Levy et al., 1986). Evidence suggests that phosphorylation events may be an important posttranslational mechanism to initiate compaction. The use of phorbol ester to activate protein kinase C (PKC) caused premature compaction of 4-cell embryos, coincident with redistribution of uvomorulin to regions of cell contact (Winkel et al., 1990; see also Bloom, 1989). Winkel et al. (1990) proposed that PKC activation may be an integral part of the cell-cell signaling mechanism that leads to the lifting of the putative compaction restraining factor. However, premature compaction induced by phorbol ester comprises uvomorulin-mediated cell adhesion but not polarization of blastomeres. A role for phosphorylation in the adhesive component of compaction is also supported by the fact that a number of cellular proteins become phosphorylated at the time of compaction or in response to agents that manipulate compaction (Bloom and McConnell, 1990; Bloom, 1991). Significantly, uvomorulin itself becomes phosphorylated for the first time at the beginning of the 8-cell stage (Sefton et al., 1992). However, recent studies indicate that the signaling path-

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way regulating compaction is more complex than first anticipated. First, the kinase inhibitor staurosporine, like the PKC activator phorbol ester, has been shown in embryos to induce premature adhesion mediated by uvomorulin but not premature polarization (O’Sullivan et al., 1993). Staurosporine is a potent inhibitor of PKC but is not entirely specific for this kinase group (Herbert et al., 1990). Second, treatment of embryos with 6-dimethyl-aminopurine (6-DMAP), a serine-threonine kinase inhibitor, also causes adhesion (but not polarization) to occur prematurely, again mediated by uvomorulin (Aghion et al., 1994). Collectively, these results suggest that both phosphorylation and dephosphorylation reactions contribute to the regulation of adhesion at compaction although the mechanism initiating cell polarity at compaction (involving transcellular ion current?) is yet to be identified. C.

Consequences of Cell Polarity

Cell polarity at compaction marks the initiation of trophectoderm differentiation and provides blastomeres with the essential spatial organization to give rise to divergent cell lineages. Division of polarized 8-cell blastomeres results in the formation of two distinct phenotypes in the 16-cell morula, a population of larger outer polar cells surrounding a group of smaller nonpolar cells (Handyside, 1980; Johnson and Ziomek, 1981b; Reeve and Ziomek, 1981; Figure 3). Various studies have demonstrated that the outer cell population tends to give rise to trophectoderm while the internal cells tend to form the ICM of the blastocyst (Tarkowski and Wroblewski. 1967: Hillman et al., 1972; Handyside and Johnson, 1978; Ziomek and Johnson, 1981; Balakier and Pedersen, 1982; Pedersen et al., 1986; Fleming, 1987a). These two cell types are distinct from the moment of their formation following differential inheritance of polarized cellular domains within parental 8-cell blastomeres (Johnson and Ziomek, 1981b). Thus, most, but not all, 8-cell blastomeres divide along an axis perpendicular to the axis of polarity, generating a polar cell which incorporates the apical pole of microvilli and a nonpolarcell incorporating the basal region (defined as a differenriarive division). A minority of blastomeres divide conservatively, parallel to the axis of polarity, such that the apical pole is bisected and inherited by both daughter cells (Johnson and Ziomek, 1981b). Thus, although cytoplasmic polarity is dissipated during mitosis, cytocortical polarity is maintained (Johnson et al., 1988) and provides the basis for establishment of separate trophectoderm and ICM cell lineages during late cleavage (Figure 3). The behavior of newly-formed polar and nonpolar 16-cell blastomeres ensure that their relative position within the embryo is maintained. Polar cells in culture tend to adhere to and envelop the nonpolar cells by virtue of the localization of uvomorulin, which is found on all cell surfaces except the apical membrane of polar cells (Ziomek and Johnson, 1981; Kimber et al., 1982; Surani and Handyside, 1983; Vestweber et al., 1987). During the fifth cell cycle, the outer polar cells continue and indeed extend their program of epithelial polarization (see later) while the internal cells remain nonpolar and gradually acquire the characteristics of ICM

T.F? FLEMING, E. BUTLER, J. COLLINS, B. SHETH, and A.E WILD

74 8-cell

16-Cell

32-cell early

co rnpaction

morula

blastocyst

Diagrammatic representation of the role of cell polarity in tissue formation and segregation during mouse cleavage (8- to 32-cell stage). Whole embryos shown above; the division plane options of individual polar cells from corresponding stage shown below. Trophectoderm lineage unshaded, ICM lineage shaded. Polar cells at the end of either 8- or 16-cell stages can divide along conservative or differentiative division planes to generate the blastocyst tissues. See text for details. Figure 3.

(Handyside and Johnson, 1978). At the end of the fifth cell cycle, polar cells can again divide either by differentiative or conservative divisions to yield either polar and nonpolar or two polar daughter cells, respectively. As in the previous cycle, nonpolar 32-cell blastomeres are located in the embryo interior, are surrounded by outer polar 32-cell blastomeres, and represent a second and final allocation of cells to the ICM lineage. Why should the mechanism of cell polarity and differentiative division be utilized twice to generate the ICM? Do the two rounds of differentiative cleavage contribute in distinct ways to early tissue segregation? Although it has been shown that the first allocation (8- to 16-cell transition) involves rnostjlastomeres dividing differentiatively, the numbers can vary considerably between embryos, from four to all eight (Johnson and Ziomek, 1981b; Balakier and Pedersen, 1982; Pedersen et al., 1986;Fleming, 1987a).This indicates that there is little control in situ on the orientation of cytokinesis in polarized 8-cell blastomeres. However, Pickering et al. (1988) have postulated from very convincing data that more advanced blastomeres (ie., those entering the fourth cell cycle earlier in typically asynchronous embryos) also engage in adhesive contacts at compaction earlier, thereby positioning deeper within the embryo and acquiring a smaller polarized apical cytocortical domain. This last attribute would in turn lead to an increased likelihood of such blastomeres dividing differentiatively rather than conservatively and would provide a cell bio-

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logical explanation of the long-standing observation that more advanced blastomeres tend to allocate disproportionately more progeny to the ICM than do later-dividing blastomeres (Surani and Barton, 1984;Garbutt et al., 1987; Pickering et al., 1988; Sutherland et al., 1990). Despite this effect of temporal order, the variable number of inner and outer 16cell blastomeres in embryos suggests that the primary role of the first round of differentiative division is to establish two populations of phenotypically distinct blastomeres in different locations within the embryo. Experimental evidence from isolated polar 16-cell blastomeres either cultured alone or in combination with other cells, has indicated that, in contrast to the first round, cell contact patterns could have a significant effect on the orientation of cleavage in the second round of differentiative division. Thus, polar 16-cell blastomeres were less likely to divide differentiatively when combined with other blastomeres, particularly nonpolar blastomeres, than when cultured alone (Johnson and Ziomek, 1983). The simplest interpretation of this phenomenon is that adhesive interactions with other blastomeres influence cell shape which in turn influence the orientation of the spindle. Moreover, support for a role for cell shape in the regulation of division plane orientation in situ has been forthcoming. It has been shown that the number of polar 16cell blastomeres dividing differentiatively in intact embryos is inversely related to the number of inner cells present within the morula (Fleming, 1987a). Thus, if the first allocation to the ICM is relatively small (fewer than normal 8-cell blastomeres dividing differentiatively) then the embryo can compensate by a relatively large allocation in the second round, and vice versa (Fleming, 1987a; Figure 4). This endogenous regulation mechanism can be best understood if the shape of polar 16-cell blastomeres is considered. A smaller than normal first allocation will lead to a larger population of polar cells enveloping this smaller core and displaying a more columnar disposition. Conversely, a large first allocation will lead to fewer polar cells of more flattened appearance enveloping a larger core (Figure 4). These shape changes, without further elaboration, could modify polar cell division orientations to ensure consistency in the cell population sizes forming the trophectoderm and ICM tissues of the blastocyst during the 32-cell stage. The use of cell polarity to establish tissue diversity by differentiative division can therefore be viewed as a two-phase event, the first concerned with establishing qualitative differences between cells (8- to 16-cell transition), the second with establishing quantitative differences between them (16- to 32-cell transition; Fleming, 1987a). D. Cell Polarity in 16-cell Blastomeres: An Intermediate Stage in Epithelial Differentiation

Maturation of the comprehensive features of cell polarity first observed at compaction continue during 16- and 32-cell cycles in outer blastomeres. At the 16-cell stage, the basolateral membrane of polar cells acquires a more complex molecular

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16-cell morula

polar cell division plane

Dff

more

Con

less

Diff

less

Con

more

32-cell early blastocyst

Figure 4.

Diagrammatic representation of the relationship between polar cell division plane and blastocyst tissue sizes. 16-cell morulae vary substantially in the number of outer polar (unshaded) and inner non-polar (shaded) cells present, indicating inter-embryonic variation in the proportion of &cell blastomeres dividing differentiatively. The polar cells within morulae containing relatively few inner cells (top) tend to divide differentiatively (Diff) more frequently and conservatively (Con) less frequently than do the polar cells within morulae containing more inner cells (bottom). This distinction can be accounted for by polar cell shape in srtu and can regulate quantitatively trophectoderm and ICM tissue sizes in the blastocyst. See text for details.

organization. The apicolateral tight junction extends from a series of focal contacts to a discontinuous zonular configuration and becomes more complex in molecular composition (discussed later; Fleming et al., 1989, 1993b; Javed et al., 1993; Sheth et al., 1997). The adherens junction becomes distinct ultrastructurally (Reima, 1990) and may include increased assembly of myosin and actin at the cytoplasmic face (Slager et al., 1992). Calcium-independent adhesion systems have been identified to function in 16-cell morulae, particularly those based on highly branched lactosaminoglycans; their neutralization results in loss of adhesion between blastomeres (Bird and Kimber, 1984; Rastan et al., 1985; Bayna et al., 1988; Fenderson et al., 1990; Kimber, 1990). The extracellular matrix, particularly laminin, is expressed at this time yet its contribution to cell polarity and trophectoderm differentiation remains to be defined (Leivo et al., 1980; Cooper and MacQueen, 1983; Leivo and Wartiovaara, 1989; Thorsteinsdottir, 1992; Hierck et al., 1993; reviewed in Damsky et al., 1993). The laminin receptor component ahintegrin, is expressed on membranes throughout cleavage but does not begin to localize to basal surfaces until laminin A-chain is present, a feature indicative of the formation of the first basement membrane (Hierck et al., 1993).

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Polarity in the cytoplasm is extended such that preferential apical endocytic activity and apical endosome clustering is further enhanced relative to basolateral regions (Fleming and Pickering, 1985). The stability of this polarity is also increased, with the previously microtubular control of endosome distribution being supplemented by microfilament interactions (Fleming et al., 1986b). A secondary lysosome compartment, polarized to the basal cytoplasm, becomes detectable for the first time in this cell cycle (Fleming and Pickering, 1985; Figure 5). Polarized transcytosis via the endosomal compartment and membrane recycling pathways exist at this stage and may be involved in the stabilization of polarized membrane domains (Fleming and Goodall. 1986; Fleming, 1987b). In addition, Golgi bodies (Fleming and Pickering, 1985; Maro et al., 1985), mitochondria and lipid droplets (Wiley and Eglitis, 1981; Batten et a]., 1987) all polarize in the basal cytoplasm, and there is an increase in the assembly of cytokeratin filaments in cytocortical and perinuclear locations (Chisholm and Houliston, 1987; Emerson, 1988).

Figure 5. Maturation of endocytic system in polar cells during trophectoderm differentiation. (A) Secondary lysosomes (L) form during the 14-ceIl stage and polarise in the basal cytoplasm of each polar cell; endosomes (E) remain polarised in the apical cytoplasm. Bar = 5pm (B)Endocytic polarity is also present in blastocyst trophectoderm cells. (C)Trophectoderm cells demonstrate preferential endocytic activity from the apical membrane. Endocytic pathways across the epithelium, as well as recycling and lysosomal pathways (arrowed), all involve obligatory sorting at endosomes, suggesting these embryonic cells are capable of maintaining the distinct composition of apical and basolateral membrane domains. After Fleming and Goodall (1986) and Fleming (1987b).

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Cell Polarity at the 32-cell Stage: Completion of Epithelial Biogenesis

Following division to the 32-cell stage, the polarization process culminates in the formation of the trophectoderm as a discrete epithelium as the blastocoel cavity is generated by vectorial fluid transport. However, cavity formation in the very early blastocyst is believed to involve the intercellular accumulation of water derived from the oxidation of cytoplasmic lipid before vectorial transport, mediated by the activity of Na+, K+-ATPase located on basolateral membranes, takes over (Wiley and Eglitis, 1981, Wiley, 1984). Na+ K+-ATPase activity has been detected along trophectoderm basolateral membranes from the 32-cell stage by ultrastructural cytochemistry (Vorbrodt et al., 1977) and by immunofluorescence microscopy (Watson and Kidder, 1988). In the latter study, thecatalytic nonglycosylated a subunit of Na+,K+-ATPase was first detectable as cytoplasmic foci in late morulae and redistributed to basolateral membranes just at the time cavitation began. However, the a subunit is transcribed from early cleavage (Watson et al., 1990b; Gardiner et al., 1990; MacPhee et al., 1994) and nonfluorescent techniques indicate that low levels of a subunit protein are present well in advance of cavitation (Gardiner et al., 1990; Van Winkle and Campione, 1991). Regulation of Na+, K+ATPase activity and basolateral localization may therefore be achieved by late expression of the glycosylated p subunit in the morula (Gardiner et al., 1990; Watson et al., 1990b; reviewed in Watson, 1992). The importance of basolateral localization of Na+, K+-ATPase in mediating cavitation has been demonstrated by the inhibitory effects of particular ionic conditions and ouabain, a specific inhibitor of the enzyme (DiZio and Tasco, 1977; Wiley, 1984; Manejwala et al., 1989). Culture media ion substitution experiments implicated Na+ (and not C l - ) ions as the major contributors to the osmotic gradient that drives water across the trophectoderm (Manejwala et al., 1989). Use of specific inhibitors suggests that transport of Na' into trophectoderm cells is carrier-mediated and may involve several apical routes of entry including Na+, K+ exchangers and Na+ channels (Manejwala et al., 1989). Na+-coupled amino acid and glucose transporters, though present, are not thought to play a significant role in blastocoel formation (DiZio and Tasco, 1977; Manejwala et al., 1989; Wiley et al., 1991). Physiological regulation of Na+,K+-ATPaseactivity at cavitation is mediated by CAMP since experimental elevation of this intracellular signaling pathway stimulates both Na+ uptake by trophectoderm and the rate of blastocoel accumulation (Manejwala et al., 1986; Manejwala and Schultz, 1989). Other important steps in the maturation of the polarized trophectoderm phenotype occur during the 32-cell stage and contribute to the functional capacity of the epithelium to engage in vectorial transport processes. Most significantly, tightjunctions become fully zonular in organization in freeze fracture replicas and functional as a permeability seal (Ducibella et al. 1975; Magnuson et al., 1977; Pratt, 1985; see later). The completion of tight junction formation in trophectoderm coincides with

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the polarized distribution of a number of membrane proteins involved in vectorial transport processes. In addition to Na+, K+-ATPase(see above), the Na+-independent GLUT2 glucose cotransporter is localized on trophectoderm basolateral membranes (Aghayan et al., 1992). This transporter may function in regulating glucose delivery to the blastocoel and ICM, demonstrating a further vectorial transport role for the polarized trophectoderm in controling the metabolic requirements of the ICM (Hewitson and Leese, 1993; Brison et al., 1993). This role is further served by an increase in the rate of endocytic activity in the trophectoderm compared with earlier stages, with endocytosis occurring preferentially at the apical surface (Fleming and Pickering, 1985; Fleming and Goodall, 1986; Pemble and Kaye, 1986; Figures 5B,C). Coordinated with endocytic polarization, the trophectoderm also engages in polarized secretion of polypeptides at apical (uterine) and basal (blastocoel) surfaces (Dardik and Schultz, 1991), an activity that is enhanced by transforming growth factor a (Dardik et al., 1993). The membrane distribution of growth factor receptors is also polarized in trophectoderm cells with the EGF receptor localized preferentially in the apical membrane (Wiley et al., 1992; Adamson, 1993) and the insulin receptor in the basolateral membrane (Heyner et al., 1989; Smith et al., 1993). These and other receptors interact with their ligands causing enhancement of trophectoderm metabolic, vectorial, and endocytic activity, including transcytosis of growth factors and their stimulation of ICM proliferation andmetabolism (Heyner et al., 1989; Harvey and Kaye, 1990, 1992; Kaye et al., 1992; Brice et al., 1993; Dunglison and Kaye, 1993; Smith et al., 1993; Shi et al., 1994). Desmosome formation, together with the assembly of major desmosomal cadherin glycoproteins and plaque proteins, occurs for the first time at punctate sites between apposed trophectoderm basolateral membranes at the 32-cell stage (Fleming et al., 1991; Collins et al., 1995; discussed later). These junctions associate with cytokeratin filaments synthesized within the cytoplasm and form i n particular when blastocoel accumulation is underway, indicating a role in stabilizing the new epithelium from stresses imposed by the expansion of the blastocoel (Fleming et al., 1991). Cavitation also coincides with increased laminin and type I11 collagen expression and deposition into basement membrane (Leivo et al., 1980; Sherman et al., 1980; Hierck et al., 1993). Recently, we have shown that a high molecular weight (330-380 kDa) membrane glycoProtein with certain characteristics of gp330 of the Heymann nephritis antigen complex (Orlando et al., 1992) and recognized by the monoclonal antibody 283D3 (Meads and Wild, 1993) is expressed along trophectoderm apical membranes from the 32-cell stage and relocates into the apical endosomal compartment from the time that blastocoel fluid accumulates; inhibition of cavitation by ouabain treatment disturbs 283D3 antigen redistribution (E. Butler, A. Wild, and T. Fleming, manuscript i n preparation). Although the precise function of this glycoprotein has yet to be defined, its activity provides a further example of cavitation coinciding with spatial reorganizaiion of blastomeres.

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MOLECULAR ASPECTS OF CELL POLARITY AND TISSUE SEGREGATION

Cell biological studies of cell polarity and its contribution to the differentiation and diversification of tissues in the preimplantation embryo have demonstrated the following: (a) cell polarity, although programmed independently of cell interactions, is dependent upon asymmetric adhesive cell contacts to define the orientation of its axis; (b) cell polarity is a stable state and represents the proto-epithelial phenotype that is extended and elaborated during late cleavage, culminating in the formation of the polarized trophectoderm epithelium; and (c) cell polarity is utilized by the embryo as a mechanism to generate phenotypically distinct cell subpopulations by differentiative division. These cell populations express many identical proteins but two-dimensional gel electrophoresis or immunolocalization studies have demonstrated that some polypeptides are specific to either trophectoderm or ICM, or their polar and nonpolar progenitors in 16-cell morulae (Van Blerkom et al., 1976; Handyside and Johnson, 1978; Johnson, 1979; Slager et al., 1991). The glycogen content of the two tissues is also distinct (Edirisinghe et al., 1984). In order to identify how these basic cellular mechanisms might regulate at the molecular level the emergence and differentiation of the two tissues in the blastocyst, in recent years we have focused our studies on the expression and distribution of cell adhesions systems during cleavage. Many elegant studies have demonstrated that the pattern of cell adhesion expression, particularly that of cadherins, has a pivotal role in the formation and segregation of tissues during development (reviewed in Takeichi, 1991). As discussed earlier, cell adhesion mediated by uvomorulinlcatenin is initiated at compaction and continues to operate throughout preimplantation development, causing adhesion between all blastomeres of the blastocyst (Vestweber et al., 1987). Uvomorulinkatenin adhesion is therefore not expressed tissue specifically in the early embryo. A.

Tight Junction

Although analysis of the uvomorulin adhesion system has not contributed to our understanding of tissue divergence, biosynthetic studies on the tight junction adhesion system have been more fruitful in this respect. Recently, it has become apparent that the apicolateral tight junction is a multimolecular complex. It is composed of at least one transmembrane protein, occludin, and several cytoplasmic “plaque” proteins including ZO- 1, localized very close to the membrane domain, and cingulin, located more internally and possibly also interacting with actin filaments (reviewed in Anderson et al., 1993; Citi, 1993;Furuse et al.. 1993). As discussed above, tightjunction formation begins at compaction but is not complete until about the 32-cell stage, approximately 24 hours later, when blastocoel fluid accumulation occurs (reviewed in Fleming et al., 1993a, 1994; Collins and Fleming, 1995). Thus, the intramembraneous component (although not yet identified in molecular terms) assembles at the

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apicolateral region of cellkell contact between %cell blastomeres at compaction (Ducibella and Anderson, 1975; Pratt, 1985), ZO-1 protein assembles at this site about 1-2 hours later (Fleming et al., 1989), and cingulin assembles some 10 hours later again and usually during the 16-cell stage (Fleming et al. 1993b; Figure 6). Fi-

Figure 6.

Tight junction formation and tissue segregation during cleavage. (A) ZO-1 protein (arrows) assembles for the first time at discontinuous sites along the apicolateral margin between &cell blastomeres after compaction has occurred; here shown in two isolated cells. (B) Cingulin protein (arrows) assembles apicolaterally for the first time usually during the 16-cell stage; here shown in cluster of four 16-cell blastomeres (from Fleming et al., 1993b). (C) At the blastocyst stage, ZO-1 (and cingulin, not shown) is distributed as a continuous belt around the apicolateral contact site between trophectoderm cells, here shown en face. (D,E) Blastocyst viewed by confocal brightfield and fluorescent imaging showing ZO-1 staining essentially restricted to the trophectoderm layer (arrowheads) and absent from ICM (I). Bar = 20pm (A,B same mag).

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nally, a new splice variant of ZO- 1 is transcribed for the first time at approximately the 32-cell stage and may have important implications for the functional activity of the tight junction. This isoform, ZO-la+, assembles at the tight junction at the early 32cell stage after intracellular association with occludin; the two proteins assembly at the membrane as a complex (Sheth et al., 1997). This event appears to complete tight junction formation such that blastocoel fluid accumulation occurs almost immediately afterward. The sequential nature of the molecular assembly of the junction appears to be coordinated by sequential expression of the proteins involved. Thus, synthesis of ZO-1 (now known to the ZO1 a-isoform) is detectable before synthesis of cingulin from the embryonic genome (Fleming et al., 1989; Javed et al., 1993), and then finally Z O - l a + (Sheth et al., 1997). Significantly, tight junction assembly is specific to the polar cell lineage generating the trophectoderm and does not occur in the ICM. Thus, both ZO- 1 isoforms and cingulin are detectable immunocytochemically in trophectoderm and are essentially absent from the ICM (Figures 6C-E, but see below). However, since the expression of tightjunction constituents spans the period during which differentiative divisions give rise to these blastocyst tissues, there is the opportunity to investigate, in biogenetic terms, the basis of tight junction tissue specificity. We have utilized synchronized clusters of isolatTd blastomeres to determine whether ZO- 1 is inherited by one or both daughter cells following differentiative division (Fleming and Hay, 1991). We have shown that at 8- to 16-cell and at 16- to 32-cell cycles, ZO-I a-isoform is inherited by both polar and nonpolar daughter cells following division, ruling out a mechanism based upon differential inheritance to explain trophectoderm-specificity of tight junction formation (Figure 7). In such polar: nonpolar cell clusters, putative apicolateral tight junction contacts containing ZO-1 are transient and remain intact only as long as the nonpolar cell retains a contact-free membrane face. Once this is lost, as occurs in intact embryos as nonpolar cells become internalized and surrounded by polar cells, then the tight junction link between polar: nonpolar sister cells is rapidly degraded with ZO-1 fragmenting into a series of randomly distributed membrane-associated foci before disappearing altogether (Fleming and Hay, 1991; Figure 7). Conversely, po1ar:polar daughter cells of conservative divisions (from 8- to 16-cell cycles onwards) establish stable apicolateral tight junctions containing ZO- 1 .This can be explained by their retention of a contact-free membrane face, the nonadhesive apical membrane domain, which ensures that they remain in an outer position in the intact embryo. The capacity to assemble a stable multimolecular tightjunction only in the outer epithelial lineage therefore appears to be regulated by cell position, (interpreted by cell contact asymmetry) rather than by a mechanism involving differential inheritance. However, it should be noted that cell positionperse is in fact regulated by differential inheritance of the apical polar domain (see earlier). A positional model to explain tight junction tissue specificity is supported by experiments in which ICMs are immunosurgically isolated from early blastocysts and cultured in vitro. Here, the outer ICM cells, now in an outer position and experiencing a contact-free sur-

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figure 7. ZO-I localization following differentiative division of polar &cell biastomeres (after Fleming and Hay, 1991). (A,B) 2/16 couplet 5 hours after differentiative division stained with TRITC-concanavalin A (A) to identify polar cell (left, with microvillous pole) and with anti ZO-1 antibody (B) showing membrane assembly at the contact site (arrow). Here, the non-polar cell still retains a contact-free membrane face. (C,D) 2/16 couplet 8 hours after differentiative division; TRITC-concanavalin A staining (C) identifies the surface of the polar cell which has now entirely enveloped the non-polar cell seen in (D)which shows ZO-1 at the contact site between cells (arrows). Here, ZO-1 appears fragmented and soon disappears since the nonpolar cell no longer possesses a contact-free membrane face. (E,F) Early 4/16 cell cluster comprising three outer polar cells and one central nonpolar cell (derived from two &cell sister blastomeres, one dividing differentiatively, the other conservatively). The nonpolar cell isstill not fully enveloped and retains a contact-free membrane face and displays ZO-1 at contact sites with polar cells (arrows). ( G I ) Later 4/16 cell cluster comprising two outer polar cells and two inner nonpolar cells (derived from two &cell blastomeres, both having divided differentiatively). Here, both nonpolar cells are now fully enclosed and are negative for ZO-1 which is found only at contact sites between polar cells (arrows) shown en face (H) and in midsection (I) of cluster. Bar = 1 Oprn.

face, rapidly reassemble an apicolateral tight junction belt containing ZO-1 (Fleming and Hay, 1991). We next investigated the effect that cell position might have on the synthesis of tight junction-associated proteins. We have shown by immunoprecipitation that

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cingulin synthesis in metabolically-labeled blastocysts is significantly greater, up to a 15-fold difference, in the trophectoderm than the ICM (Javed et al., 1993). Thus, it appears that loss of cell contact asymmetry in the ICM lineage not only leads to loss of assembly competence but also to down-regulation of expression of tightjunction components. Recently, we have analyzed whether the reduction in expression of tight junction proteins in the ICM was regulated by transcriptional or translational mechanisms. Utilizing the RT-PCR technique, transcripts for both isoforms of ZO-1 were detected in both trophectoderm and ICM of early blastocysts, suggesting that the reduction in ICM expression is controlled by reduction in mRNA translation (Sheth et al., 1997). Moreover, the reformation of a ZO-1containing tightjunction network in outer cells of isolated ICMs in culture is insensitive to a-amanitin treatment, confirming that new transcription is not required for this up-regulation event (Fleming and Hay, 1991). Why might transcripts for tight junction proteins be retained within the ICM? Presumably such transcripts (as well as protein, see above) will be inherited by differentiative divisions during the morula stage, but we have not yet established whether ICM nuclei in situ engage in transcription of mRNA for tight junction constituents. We suspect that these transcripts may serve a role in the developmental program undertaken by the ICM. In the late blastocyst, a new epithelium, the primary endoderm (progenitor of extra-embryonic parietal and visceral endoderm tissues), is delaminated at the blastocoelic face of the ICM and typically contains tight junctions (Nadijcka and Hillman, 1975; reviewed in Gardner and Beddington, 1988). We have proposed that the same pool of transcripts may be utilized for the expression of tight junctions in both trophectoderm and primary endoderm epithelia (Fleming and Hay, 1991; Fleming et al., 1993a). Thus, during blastocyst expansion, this pool would be maintained in a state of low translation by the presence of cellular processes, derived from nearby trophectoderm cells, that cover the blastocoelic face of the ICM and prohibit the formation of contact-free membrane surfaces at this site (Fleming et al., 1984; shown in Figures 3 and 4). These processes withdraw in the late blastocyst, concommitant with the differentiation of primary endoderm and up-regulation of tight junction expression and assembly. Taken together, our analysis of the tight junction adhesion system during blastocyst formation has demonstrated that cell polarity plays a significant role in its maturation and tissue specificity. The sequential pattern of tight junction protein expression and membrane assembly is dependent upon the continued presence of a contact-free cell surface provided by the nonadhesive apical membrane domain. The loss of such a domain, as occurs in cells entering the ICM lineage, leads to reduction or cessation in synthesis of tight junction proteins and their disassembly at membrane contact sites. This down-regulation is reversible if a contact-free surface is reestablished. However, these dramatic changes in biosynthetic events do not appear to be coordinatined with, or dependent upon, changes in transcriptional activity of tight junction constituents.

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85

Desmosomes

The formation of disc-shaped desmosome adhesive junctions represents a late maturation step in the cell polarization program underlying trophectoderm differentiation and occurs during the 32-cell stage Just as tight junction formation is completed and blastocoel fluid accumulation has initiated (see above). The principal components of desmosome junctions are two adhesive transmembrane glycoproteins belonging to the cadherin superfamily (desmoglein and desmocollins) and three associated cytoplasmic proteins (plakoglobin, desmoplakin I and 11) which together form a plaque into which cytokeratin filaments insert (reviewed in Schwarz et al., 1991; Buxton and Magee, 1992; Garrod, 1993). Like tight junctions, desmosomes are trophectoderm-specific in the blastocyst (Ducibella et al., 1975; Magnuson et a]., 1977; Fleming et al., 1991) but appear to have a different mechanism for regulating their initial construction. Metabolic labeling and immunoprecipitation analysis of carefully-staged preimplantation embryos has revealed that the desmosomal plaque constituents are first synthesized during cleavage (8- and 16-cell stages) and in advance of the desmosomal cadherins (32-cell stage). However, all major constituents first assemble along apposed trophectoderm membrane contact sites at approximately the same time (32-cell stage) when desmosomes first form (Fleming et al., 199 1). These biogenetic characteristics indicate that the initiation of desmosoma1 cadherin expression may regulate the timing of desmosome formation which may utilize preexisting nonassembled plaque components. Desmosome formation therefore occurs rapidly, within a single cell cycle, and is mediated by the delay in availability of the membrane-spanning components whereas the tight junction is formedprogressively with cytoplasmic plaque constituents assembling in a sequential manner during cleavage. These differing strategies may reflect the difference in shape, size, and morphology of these junction types (discussed in Fleming et al., 1993a) but ensure that they become functional at about the same time, as the trophectoderm vectorial transport activity gets underway. The biogenetic control of the timing of desmosome formation i n the early embryo has been studied further by analyzing the timing of transcription of desmocollin, the desmosomal cadherin that has been implicated a significant role in desmosome adhesion. The two alternatively-spliced variants (a and b; Collins et al., 1991) of the mouse desmocollin gene 2 (DSC2) are transcribed coordinately from the embryonic genome beginning at the late 16-cell stage or the early 32-cell stage (Collins et al., 1995; Figure 8A). This transcriptional event correlates well with the onset of desmocollin translation (Fleming et al., 199 1 ; see above). Taken together, our data strongly suggests that desmosome formation in trophectoderm is controled by desmocollin transcriptional activation.

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T.f? FLEMING, E. BUTLER, J. COLLINS, B. SHETH, and A.E WILD

Reverse transcriptase PCR amplification of DSC2 (desmocollin)transcripts in embryos and blastomeres. (A) Lanes labeled 1 are DNA amplified from single 16-cell morulae showing either negative or weak positive signal indicative of the initiation of DSC2 transcription from the embryonic genome (confirmed by sequencing of product). 3 , three 16-cell morulae, B, single early blastocyst, where more product has been amplified; C, control samples minus template but with complete reaction. Arrowheads adjacent to marker lanes indicate 564bp. (B) DSC2 transcript detection in single trophectoderm (T) or ICM ( I ) cells derived from early blastocysts. Trophectoderm cells consistently demonstrate the presence of DSC2 mRNA whereas only a minority of ICM cells do. B, single intact blastocyst; C, controls minus template as above. Arrowheads adjacent to marker lanes indicate 600 bp. After Collins et al. (1 994).

Figure 8.

The mechanism regulating desmosome tissue specificity has also been investigated. The presence of DSC2 mRNA is detectable by a sensitive modification of the RT-PCR technique within single blastomeres of known phenotype isolated from early blastocysts. This analysis has demonstrated that all trophectoderm cells contain DSC2 mRNA but in most ICM cells (approximately 75%) the transcript is not detectable despite the reliable identification of uvomorulin mRNA in all cells irrespective of their phenotype (Collins et al., 1995; Figure 8B). Desmosome tissue specificity in the early embryo can best be explained therefore by differential transcription of the desmocollin gene in the early blastocyst. Moreover, the detection of DSC2 mRNA in a minority of ICM cells need not represent “leakiness” in regulation of transcriptional activity. This proportion of ICM cells (25%) is exactly the average proportion known to be allocated from the polar lineage following the second round of differentiative divisions at the 16- to 32-cell stage (Fleming, 1987a). Since DSC2 transcription may just precede this division cycle, the ICM DSC2 mRNA pool is presumably generated by inheritance from outer polar cells rather than by inherent transcription. Significantly, isolation and culture of ICMs from early blastocysts leads to a substantial increase in the level of DSC2 mRNA detected and to the expression and assembly of desmocollin protein at putative desmosomes present between outer cells (Collins et al., 1995).Thus, the positional signal of the presence or absence of a contact-free membrane face identified to regulate tight junction protein expression also appears to operate to regulate desmocollin transcription and translation.

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CONCLUSIONS

Cell polarity established at compaction in the mouse embryo can be viewed as the foundation of several interrelated processes governing blastocyst formation. Blastomere polarization is likely to be of widespread importance in the early development of eutherian mammals since it occurs in several species other than the mouse (e.g., Koyama et al., 1994). In the mouse, it is a stable state in that polarized cells have not been shown to lose their polarity even in isolation. In the intact embryo, this stability is conducive with the polar lineage maintaining cell position and orientation, and in acquiring further structural and molecular features of polarity with time, such as occurs in intercellular junction formation. The molecular nature of this stability is unknown, but given the relative importance of the cytocortex over cytoplasmic domains of the cell in establishing and maintaining polarity, the apical membrane and cortex may be regarded as the most likely center where stability is controlled. The microvilli of the apical pole, unlike others on blastomere membranes, are not disrupted by prolonged cytochalasin treatment (Pratt et al., 1982; Fleming et al., 1986a), which suggest the actin bundles that structure them are biogenetically stable, turning over very slowly. The composition and molecular organization of actin-associated proteins in the apical cortex may therefore be important in conferring stability to cell polarity (see Johnson et al., 1988). In addition to providing a stable "framework" upon which the polarized cellular organization of the trophectoderm can be manifest, cell polarity also regulates tissue divergence by differentiative division. Our studies on cell adhesion maturation have identified that the presence or absence of a contact-free membrane surface on blastomeres controls up- or down-regulation of gene and protein expression which in turn underlies tissue divergence. It appears, therefore, that the pattern of blastomere biogenesis is controlled by whether or not it inherits part or all of the apicaI cortical domain which will ensure the continuance of a contact-free cell surface. Our next task will be to unravel the molecular signaling pathway leading from cellular interaction pattern to gene expression pattern.

ACKNOWLEDGMENTS We are grateful to the Wellcome Trust, the Medical Research Council, the Science and EngineeringResearch Council, and the Wessex Medical Trust for funding of research in our laboratory and for provision of studentships. We thank many collaborators for gifts of precious antibodies, Sue Pickeringfor scanningEM micrographs,and Mark Hay for his skill in generating the computerized diagrams.

REFERENCES Adamson, E.D. (1993).Activities of growth factors in preimplantation embryos. J. Cell, Biochem. 53, 280-287.

88

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Aghayan, M., Rao, L.V., Smith, R.M., Jarett, L., Charron, M.J., Thorens, B. & Heyner, S. (1992). Developmental expression and cellular localization of glucose transporter molecules during mouse preimplantation development. Development 115,305-312. Aghion, J., Gueth-Hallonet, C., Anthony, C., Gros, D. & Maro, B. (1994). Cell adhesion and gap junction formation in the early mouse embryo are inducedprematurelyby 6-DMAP in the absence of E-cadherin phosphorylation. J. Cell Sci. 107, 1369-1379. Anderson, J.M., Balda, M.S. & Fanning, A S . (1993). The structure and regulation of tight junctions. Curr. Opin. Cell Biol. 5, 772-778. Balakier, H. & Pedersen, R.A. (1982). Allocation ofcells to inner cell mass and trophectoderm lineages in preimplantation mouse embryos. Dev. Biol. 90,352-362. Batten, B.E., Albertini, D.F. & Ducibella, T. (1987). Patterns of organelle distribution in mouse embryos during preimplantation development. Amer. J. Anat. 178,204213. Bayna, E.M., Sharper, J.H. & Shur, B. (1988). Temporally specific involvement of cell surface 8-1,4 galactosyltransferase during mouse embryo morula compaction, Cell 53, 145-157. Bird, J.M. & Kimber, S.J. (1984). Oligosaccharides containing fucose linked a(1-3) and a(I-4) to N-acetylglucosamine cause decompaction of mouse morulae. Dev. Biol. 104,449460. Blaschuk, O.W., Sullivan, R., David, S. & Pouliot Y. (1990). Identification of a cadherin cell adhesion recognition sequence. Dev. Biol. 139,227-229. Bloom, T.L. (1989). The effects of phorbol ester on mouse blastomeres: a role for protein kinase C in compaction? Development 106, 159-171. Bloom, T.L. (1991). Experimental manipulation of compaction of the mouse embryo alters patterns of protein phosphorylation. Molec. Reprod. Devel. 28, 23g244. Bloom, T.L. & McConnell, J. (1990). Changes in protein phosphorylation associated with compaction of the mouse preimplantation embryo. Molec. Reprod. Devel. 26, 199-210. Brice, E.C., Wu, J.X., Muraro, R., Adamson, E.D. & Wiley, L.M. (1993). Modulation of mouse preimplantation development by epidermal growth factor receptor antibodies, antisense RNA, and deoxyoligonucleotides. Devel. Genet. 14, 174-1 84. Brison, D.R., Hewitson, L.C. & Leese, H.J. (1993). Glucose, pyravate, and lactate concentrations in the blastocoel cavity of rat and mouse embryos. Molec. Reprod. Devel. 35,227-232. Buxton, R.S. & Magee, A.I. (1992). Structure and interactionsofdesmosomal and other cadherins. Sem. Cell Biol. 3, 157-167. Chisholm, J.C. & Houliston, E. (1987). Cytokeratin filament assembly in the preimplantation mouse embryo. Development 101,565-582. Citi, S. (1993). The molecular organization of tightjunctions. J. Cell Biol. 121,485489. Clayton, L., Stinchcombe, S.V. & Johnson, M.H. (1993). Cell surface localisation and stability of uvomorulin during early mouse development. Zygote 1,333-344. Collins, J.E. & Fleming, T.P. (1995). Epithelial differentiation in the mouse preimplantation embryo: Making adhesive cell contacts for the first time. Trends. Biochem. Sci. 20, 307-312. Collins, J.E., Legan, P.K., Kenny, T.P., Macgarvie, J., Holton, J.L. & Garrod, D.R. (1991). Cloning and sequencing of desmosomal glycoproteins 2 and 3 (desmocollins): cadherin-like desmosomal adhesion molecules with heterogeneous cytoplasmic domains. J. Cell Biol. 113,381-391. Collins, J,E., Lorimer, J.E., Garrod, D.R., Pidsley, S., Buxton, R. & Fleming, T.P. (1995). Regulation of desmocollin transcription in mouse preimplantation embryos. Development l 2 t , 743-753. Cooper, A.R. & MacQueen, H.A. (1983). Subunits of laminin are differentially synthesised in mouse eggs and early embryos. Dev. Biol. 96,467471. Cruz, Y.P. (1992). Role of ultrastructural studies in the analysis of cell lineage in the mammalian pre-implantation embryo. Microsc. Res. Tech. 22, 103-125. Damsky, C., Sutherland, A. & Fisher, S. (1993). Extracellular matrix 5: adhesive interactions in early mammalian embryogenesis, implantation, and placentation. FASEB J. 7, 1320-1329.

Cell Polarity and Mouse Early Development

89

Dardik, A. & Schultz, R.M. (1991). Protein secretion by the mouse blastocyst: differences in the polypeptide composition secreted into the blastocoel and medium. Biol. Reprod. 45,328-333. Dardik, A,, Doherty, A.S. & Schultz, R.M. (1993). Protein secretion by the mouse blastocyst: stimulatory effect on secretion into the blastocoel by transforming growth factor-a. Molec. Reprod. Devel. 34,396401. DiZio, S.M. & Tasca, R.J. (1977). Sodium-dependentaminoacid transport in preimplantationembryos. 111. Na+, K+-ATPase linked mechanism in blastocysts. Dev. Biol. 59, 198-205. Ducibella, T. & Anderson, E. (1975). Cell shape and membrane changes in the 8-cell mouse embryo: prerequisites for morphogenesis of the blastocyst. Dev. Biol. 47.45-58. Ducibella, T., Albertini, D.F., Anderson, E. & Biggers, J.D. (1975). The preimplantation mammalian embryo: characterization of intercellular junctions and their appearance during development. Dev. Biol. 45,231-250. Ducibella, T., Ukena, T., Karnovsky, M. & Anderson, E. (1977). Changes in cell surface and cortical cytoplasmic organization during early embryogenesis in the preimplantation mouse embryo. J. Cell Biol. 74, 153-167. Dunglison, G.F. & Kaye, P.L. (1993). Insulin regulates protein metabolism in mouse blastocysts. Molec. Reprod. Devel. 36, 4 2 4 8 . Edirisinghe, W.R., Wales, R.G. &Pike, I.L. (1984). Studies ofthe distributionofglycogen between the inner cell mass and trophoblast cells of mouse embryos. J. Reprod. Fert. 71, 533-538. Emerson, J.A. (1988). Disruption of the cytokeratin filament network in the preimplantation mouse embryo. Development 104,219-234. Fenderson, B.A., Eddy, E.M. & Hakomori, S. (1990). Glycoconjugate expression during embryogenesis and its biological significance. BioEssays 12, 173-179. Flach,G., Johnson,M.H.,Braude,P.R. ,Taylor,R. &Bolton,V.N. (1982).Thetransitionfrommaternal to embryonic control in the 2-cell mouse embryo. EMBO J. 1,681-686. Fleming, T.P. (1987a). A quantitative analysis ofcell allocation to trophectoderm and inner cell mass in the mouse blastocyst. Dev. Biol. 119, 520-531. Fleming, T.P. (1987b). Endocytosis and epithetial biogenesis in the mouse early embryo. BioEssays 4, 105-109. Fleming, T.P. (1992). Trophectoderm biogenesis in the preimplantation mouse embryo. In: Epithelial Organization and Development (T.P. Fleming, Ed.), pp. 1 1 1-136. Chapman and Hall, London. Fleming, T.P. & Goodall, H. (1986). Endocytic traffic in trophectoderm and polarised blastomeres of the mouse preimplantation embryo. Anat. Rec. 216,490-503. Fleming, T.P. & Hay, M.J. (1991). Tissue-specific control of expression of the tight junction polypeptide ZO-1 in the mouse early embryo. Development 113,295-304. Fleming, T.P. & Pickering, S.J. (1985). Maturationand polarizationofthe endocytoticsystem in outside blastomeres duringmouse preimplantation development. J. Embryol.exp. Morph. 89,175-208. Fleming, T.P., Warren, P.D., Chisholm, J.C. & Johnson, M.H. (1984). Trophectodermal processes regulate the expression of totipotency within the inner cell mass of the mouse expending blastocyst. J. Embryol. exp. Morph. 84, 63-90. Fleming, T.P., Pickering, S.J., Qasim, F. & Maro, B. (1986a). The generation ofcell surface polarity in mouse 8-cell blastomeres: the role of cortical microfilaments analysed using cytochalasin D. J. Embryol. exp. Morph. 95, 169-191. Fleming, T.P ,Cannon, P M. & Pickering, S.J. (1986b). The cytoskeleton, endocytosis and cell polarity in the mouse preimplantation embryo. Dev. Biol. I 13,406419. Fleming, T.P., McConnell, J., Johnson, M.H. &Stevenson, B.R. (1989). Development oftightjunctions de novo in the mouse early embryo: control of assembly of the tight junction-specific protein, ZO-1. J . Cell Biol. 108, 1407-1418. Fleming, T.P., Garrod, D.R. & Elsmore, A.J (1991). Desmosome biogenesis in the mouse preimplantation embryo. Development 1 12,527-539.

90

T.P FLEMING, E. BUTLER, J. COLLINS, B. SHETH, and A.E WILD

Fleming, T.P., Javed, Q. & Hay, M. (1992). Epithelial differentiation and intercellular junction formation in the mouse early embryo. Development supplement 105-1 12. Fleming, T.P., Javed, Q., Collins, J. & Hay M. (l993a). Biogenesis of structural intercellularjunctions during cleavage in the mouse embryo. J. Cell Science supplement 17, 119-125. Fleming, T.P., Hay, M.J., Javed, Q. & Citi, S. (1993b). Localization oftightjunction protein cingulin is temporally and spatially regulated during early mouse development. Development 117, 1135-1 144. Fleming, T.P., Butler E., Lei, X., Collins, J., Javed, Q., Sheth, B., Stoddart, N., Wild, A. & Hay, M. (1994). Molecular maturation of cell adhesion systems during mouse early development. Histochemistry 101, 1-7. Furuse, M., Hirase, T., Itoh, M., Nagahchi, A., Yonemura, S., Tsukita, S. & Tsukita, S. (1993). Occludin: a novel integral membrane protein localizing at tight junctions. J. Cell Biol. 123, 1777-1788. Garbutt, L.C., Johnson, M.H. &George, M.A. (1987). When and how does cell division order influence cell allocation to the inner cell mass of the mouse blastocyst? Development 100,325-332. Gardiner, C.S., Williams, J.S. & Menino, A.R. (1990). Sodium/potassium adenosine triphosphatase aand p-subunit and a-subunit mRNA levels during mouse embryo development in vifro. Biol. Reprod. 43,788-794. Gardner, R.L. & Beddington, R.S.P. (1988). Multi-lineage “stem” cells in the mammalian embryo. J. Cell Sci. supplement 10, 11-27. Garrod, D.R. (1993). Desmosomes and hemidesmosames. Curr. Opin. Cell Biol. 5 , 3 W O . Geiger, B. & Ayalon, 0. (1992). Cadherins. Ann. Rev. Cell Biol. 8,307-332. Goodall, H. & Johnson, M.H. (1984). The nature of intercellular coupling within the preimplantation mouse embryo. J. Embryol. exp. Morph. 79, 53-76. Grunwald, G.B. (1993). The structural and functional analysis of cadherin calcium-dependent cell adhesion molecules, Curr. Opin. Cell Biol. 5 , 797-805. Gueth-Hallonet, C. & Maro, B. (1992). Cell polarity and cell diversification during early mouse embryogenesis. Trends Genet. 8,274279. Handyside, A.H. (1980). Distribution of antibody- and lectin-binding sites on dissociated blastomeres of mouse morulae: evidence for polarization at compaction. J . Embryol. exp. Morph. 60,99-116. Handyside, A.H. & Johnson, M.H. (1978). Temporal and spatial patterns of the synthesis of tissue-specific polypeptides in the preimplantation mouse embryo. J. Embryol. exp. Morph. 44, 191-199. Harvey, M.C. & Kaye, P.L. (1990). Insulin increases the number ofthe inner cell mass and stimulates morphological development of mouse blastocysts in vifro. Development 110,963-967. Harvey, M.C. & Kaye, P.L. (1992). IGF-2 stimulates growth and metabolism ofearly mouse embryos. Mechanisms Devel. 38, 169-174. Herbert, J.M., Augereau, J.M., Gleye, J. & Maffrand, J.P. (1990). Chelerythrine is apotentand specific inhibitor of protein kinase C. Biochem. Biophys. Res. Commun. 172,993-999. Hewitson, L.C. & Leese, H.J. (1993). Energy metabolism of the trophectoderm and inner cell mass of the mouse blastocyst. J. Exp. Zool. 267,337-343. Heyner, S., Rao, L.V., Jarett, L. & Smith, R.M. (1989). Preimplantation mouse embryos internalize maternal insulin via receptor-mediated endocytosis: pattern of uptake and functional correlations. Dev. Biol. 134,48-58. Hierak, B.P., Thorsteinsdottir, S., Niessen, C.M., Freund, E., Iperen, L.V., Feyen, A,, Hogervorst, F., Poelmann, R.E., Mummery, C.L. & Sonnenberg, A. (1 993). Variants ofthe a6pIlaminin receptor in early murine development: distribution, molecular cloning and chromosomal localization of the mouse integrin a6subunit. Cell Adhesion and Regulation 1,33-53. Hillman, N., Sherman, M.1. & Graham, C. (1972). The effect of spatial arrangement on cell determination during mouse development. J. Embryol. exp. Morph. 28, 263-278.

Cell Polarity and Mouse Early Development

91

Houliston, E. & Maro, B. (I 989). Posttranslational modification of distinct microtubule subpopulations during cell polarization and differentiation in the mouse preimplantation embryo. J. Cell Biol. 108,543-551. Houliston, E., Pickering, S.J. & Maro, B. (1987). Redistribution of microtubules and pericentriolar material during the developmentof polarity inmouse blastomeres. J. Cell Biol. 104,1299-1 308. Hyafil, F., Morello, D., Babinet, C. & Jacob, F. (1980). A cell surface glycoprotein involved in the compaction of embryonal carcinoma cells and cleavage stage embryos. Cell 21,927-934. Javed, Q., Fleming, T.P., Hay, M. & Citi, S. (1993). Tight junction protein cingulin is expressed by maternal and embryonic genomes during early mouse development. Development I 17,1145-1 I5 I. Johnson, M.H. (1979). Molecular differentiation of inside cells and inner cell masses isolated from the preimplantation mouse embryo. J. Embryol. exp. Morph. 53,335-344. Johnson, M.H. & Maro, B. (1 984). The distribution of cytoplasmic actin in mouse 8-cell blastomeres. J. Embryol. exp. Morph. 82,97-I 17. Johnson, M.H. & Maro, B. (1985). A dissection ofthe mechanisms generating and stabilisingpolarity in mouse 8- and 16-cell blastomeres: the role ofcytoskeletal elements. J. Embryol. exp. Morph. 90, 3 11-334. Johnson, M.H. & Ziomek, C.A. (1981a). Induction of polarity in mouse 8-cell blastomeres: specificity, geometry and stability. J. Cell. Biol. 91, 303-308. Johnson, M.H. & Ziomek, C.A. (198 1b). The foundation of two distinct cell lineages within the mouse morula. Cell 24, 71-80. Johnson, M.H. & Zionek, C.A. (1983). Cell interactions influence the fate of mouse blastomeres undergoing the transition from the 16- to the 32-cell stage. Dev. Biol. 95,211-218. Johnson, M.H., Maro, B. & Takeichi, M. (1986). The role of cell adhesion in the synchronization and orientation of polarization in 8-cell mouse blastomeres. J. Embryol. exp. Morph. 93,239-255. Johnson, M.H., Pickering, S.J., Dhiman, A,, Radcliffe, G.S. & Maro, B. (1988). Cytocortical organisation during natural and prolonged mitosis in mouse 8-cell blastomeres Development 102, 143-158. Kaye, P.L.,Bell, K.L., Beebe, L.F.S., Dunglison, G.F., Gardner,H.G. & Harvey, M.B. (1992). Insulin and IGFs in preimplantation development. Reprod. Fertil. Devel. 65,367-375. Kidder, G.M. & McLachlin, J.R. (1985). Timing of transcription and protein synthesis underlying inorphogenesis in preimplantation mouse embryos. Dev. Biol. 1 12,265-275. Kimber, S.J. (1990). Glycoconjugates and cell surface interactions in pre- and peri-implantation mammalian embryonic development. Int. Rev. Cytol. 120, 53-167. Kimber, S.J., Surani, M.A.H. & Barton, S.C. (1982). Interactions ofblastomeres suggest changes in cell surface adhesiveness during the formation of inner cell mass and trophectoderm in the preimplantation mouse embryo. J. Embryol. exp. Morph. 70, 133-152. Kimber, S.J., Bentley, J., Ciemerych, M., Moller, C.J. & Bock, E. (1994). Expression of N-CAM in fertilized pre- and peri-implantation and parthenogenetically activated mouse embryos. Eur. J. Cell Biol. 63, 102-1 13. Lehtonen, E., Ordonez, G. & Reima, I. (1988). Cytoskeleton in preimplantation mouse development. Cell Differentiation 24, 165-1 78. Leivo, 1. & Wartiovaara, J. (1989). Basement membrane matrices in mouse embryogenesis, teratocarcinoma differentiation and in neuromuscular maturation. Int. J. Dev. Biol. 33, 81-89. Leivo, I., Vaheri, A., Timpl, R. & Wartiovaara, J. (1980). Appearance and distribution ofcollagens and laminin in the early mouse embryo. Dev. Biol. 76, 1OC-114. &Maro,B. (1986). Control ofthetimingofcompaction: amajor Levy, J.B.,Johnson,M.H.,Goodall,H. developmental transition in mouse early development. J. Embryol. exp. Morph. 95,213-237. LO,C.W. & Gilula,N.B. (1979). Gap junctional communication in the preimplantationmouse embryo. Cell 18, 399409. MacPhee, D.J., Barr, K.J., De Sousa, P.A., Todd, S.D.L. &Kidder, G.M. (1994). RegulationofNa+, K+-ATPase alpha subunit gene expression during mouse preimplantation development. Dev. Biol. 162,259-266.

92

T.P FLEMING, E. BUTLER, J. COLLINS, 6.SHETH, and A.E WILD

Magnuson, T., Demsey, A. & Stackpole, C.W. (1977). Characterization of intercellularjunctions in the preimplantation mouse embryo by freeze-fracture and thin-section electron microscopy. Dev. Biol. 61, 252-261. Manejwala, F.M. & Schultz, R.M. (1989). Blastocoel expansion in the preimplantation mouse embryo: stimulation of sodium uptake by CAMPand possible involvement of CAMP-dependent protein kinase. Dev. Biol. 136, 560-563. Manejwala, F.M., Cragoe, E.J. & Schultz, R.M. (1989). Blastocoel expansion in the preimplantation mouse embryo: role of extracellular sodium and chloride and possible apical routes oftheir entry. Dev. Biol. 133,210-220. Manejwala, F.M., Kali, E. & Schultz, R.M. (1986). Developmentofactivatable adenylate cyclase in the preimplantation mouse embryo and a role for cyclic AMP in blastocoel formation. Cell 46, 95-1 03, Maro, B., Johnson, M.H., Pickering, S.J. & Louvard, D. (1985). Changes in the distribution of mernbraneous organelles during mouse early development. J. Embryol. exp. Morph. 90, 287-309. McLachlin, J.R., Caveney, S. & Kidder, G.M. (1983). Control ofgapjunction formation in early mouse embryos. Dev. Biol. 98, 155-164. Meads, T.J. & Wild, A.E. (1993). Apical expression of an antigen common to rabbit yolk sac endoderm and kidney proximal tubule epithelium. J. Reprod. Immunol. 23,247-264. Nadijcka, M. & Hillman, N. (1974). Ultrastructural studies of the mouse blastocyst substages. J. Embryol. exp. Morph. 32,675495. Nuccitelli, R. & Wiley, L. (1985). Polarity of isolated blastomeres from mouse morulae: detection of transcellular ion currents. Dev Biol. 109,452463. Ohsugi, M., Hwang, S.Y., Butz, S., Knowles, B.B., Solter, D. & Kemler, R. (1996). Expression and cell-membrane localization of catenins during mouse preimplantation development. Devel. Dynam. 206,391-402. Orlando, R.A., Kerjaschki, D., Kurihara, H., Biemesderfer, D. & Farquhar, M.G. (1992). gp330 associates with a 44-kDa protein in the rat kidney to form the Heymann nephritis antigenic complex. Proc. Natl. Acad. Sci. USA 89,6698-6702. O’Sullivan, D.M., Johnson, M.H. & McConnell, J.M.L. (1993). Staurosporine advances interblastomeric flattening of the mouse embryo. Zygote 1, 103-1 12. Paynton, B.V., Rempel, R. & Bachvarova, R. (1988). Changes in state of adenylation and time course of degradation of maternal mRNAs during oocyte maturation and early embryonic development in the mouse. Dev. Biol. 129,304-314. Pedersen, R.A., Wu, K. & Balakier, H. (1986). Origin of the inner cell mass in mouse embryos: cell lineage analysis by microinjection. Dev. Biol. 117, 581-595. Pemble, L.B. & Kaye, P.L. (1986). Whole protein uptake and metabolism by mouse blastocysts. J. Reprod. Fert. 78, 149-157. Peyrieras, N., Hyafil, F., Louvard, D., Ploegh, H. L. & Jacob, F. (1983). Uvomorulin: a non-integral membrane protein of early mouse embryo. Proc. Natl. Acad. Sci USA 80,6274-6277. Pickering, S.J., Maro, B., Johnson, M.H. & Skepper, J.N. (1988). The influence of cell contact on the division of mouse 8-cell blastomeres. Development 103,353-363. Pratt, H.P.M. (1985). Membrane organization in the preimplantation mouse embryo. J. Embryol. exp. Morph. 90, 101-121. Pratt, H.P.M., Ziomek, C.A., Reeve, W.J.D. & Johnson, M.H. (1982). Compaction of the mouse embryo: an analysis of its components. J. Embryol. exp. Morph. 70, 1 13-132. Rastan, S., Thorpe, S.J., Scudder, P., Brown, S., Gooi, H.C. & Feizi, T. (1985). Cell interactions in preimplantation embryos: evidence for involvement of saccharides of the poly-N-acetyllactosamineseries. J. Embryol. exp. Morph. 87, 115-128. Reeve, W.J.D. (1981). Cytoplasmic polarity develops at compaction in rat and mouse embryos. J. Embryol exp. Morph. 62,351-367.

Cell Polarity and Mouse Early Development

93

Reeve, W.J.D. & Kelly, F. (1983). Nuclear position in cells ofthe mouse early embryo. J. Embryol exp. Morph. 75, 117-139. Reeve, W.J.D. & Ziomek, C.A. (I98 I). Distribution ofmicrovilli on dissociated blastomeres from mouse embryos: evidence for surface polarization at compaction. J. Embryol. Exp. Morph. 62,339-350. Reima, I. (1990). Maintenance of compaction and adherent-type junctions in mouse morula-stage embryos. Cell. Differen. 29, 143-153. Schwarz, M., Owaribe, K., Kartenkeck, J, & Franke, W.W. (1991). Desmosomes and hemidesmosomes: constitutive molecular components. Ann. Rev. Cell Biol. 6,461491. Sefton, M., Johnson, M.H. & Clayton, L. (1992). Synthesis and phosphorylation of uvomorulin during mouse early development, Development I 15,313-318. Sherman, M.I., Gay, R., Gay, S. & Miller, E.J. (1980). Association ofcollagen with preimplantationand periimplantation mouse embryos. Dev. Biol. 74,470-478. Sheth, B., Fesenko, I., Collins, J.E., Moran, B., Wild, A.E., Anderson, J.M. & Fleming, T.P. (1997). Tight junction assembly during mouse blastocyst formation is regulated by late expression of ZO-la+ isoform. Development 124,2027-2037. Shi, C.Z., Collins, H.W., Buettger, C.W., Garside, W.T., Matschinsky, F.M. & Heyner, S. (1994). Insulin family growth factors have specific effects on protein synthesis in preimplantation mouse embryos. Molec. Reprod. Devel. 37,398-406. Shirayoshi, Y., Okada, T.S. & Takeichi, M. (1983). The calcium-dependent cell-cell adhesion system regulates inner cell mass formation and cell surface polarization in early mouse development. Cell 35,631438. Slager, H.G., Lawson, K.A., Van Den Eijnden-Van Raaij, A.J.M., DeLaat, S.W. & Mummery, C. ( 1991). Differential localization of TGF-P2 in mouse preimplantation and early post-implantation development. Dev. Biol. 145,205-218. Slager, H.G., Good, M.J., Schaart, G., Groenewoud, J.S. & Mummery, C.L. (1992). Organization of non-muscle myosin during early murine embryonic differentiation. Differentiation 50,47-56. Smith, R.M., Garside, W.T., Aghayan, M., Shi, C.Z., Shah, N., Jarett, L. & Heyner, S. (1993). Mouse preimplantation embryos exhibit receptor-mediated binding and transcytosis of maternal insulin-like growth factor I . Biol. Reprod. 49, 1-12. Sobel, J.S. (1983). Localization of myosin in the preimplantation mouse embryo. Dev. Biol. 95, 227-23 1. Surani, M.A.H. &Barton, S.C. (1984). Spatial distributionofblastomeres is dependent on cell division order and interactions in mouse morulae. Dev. Biol. 102,335-343. Surani, M.A.H. & Handyside, A.H. (1983). Reassortment of cells according to position in mouse morulae. J. Embryol. exp. Morph. 225,505-51 1. Sutherland,A.E., Speed, T.P. & Calarco, P.G. (1990). Inner cell allocation in the mouse morula: the role of orientated division during fourth cleavage. Dev. Biol. 137, 13-25. Takeichi, M. (1991). Cadherin cell adhesion receptors as a morphogenetic regulator. Science 251, 1451-1455. Tarkowski, A.K. & Wroblewska, J. (1967). Development of blastomeres of mouse eggs isolated at the 4- and 8-cell stage. J. Embryol. exp. Morph. 18, 155-180. Thorsteinsdottir, S. (1992). Basement membrane and fibronectin matrix are distinct entities in the developing mouse blastocyst. Anat. Rec. 232, 141-149. Van Blerkom, J., Barton, S.C. &Johnson, M.H. (1976). Molecular differentiation in the preimplantation mouse embryo. Nature 259,3 19-32 I . Van Winkle, L.J. & Campione, A.L. (1991). Ouabain-sensitive Rb+ uptake in mouse eggs and preimplantation conceptuses. Dev. Biol. 146, 158-166. Vestweber, D.,Gossler, A,, Boller, K. & Kemler, R. (1987). Expressionanddistributionofcell adhesion molecule uvomorulin in mouse preimplantation embryos. Dev. Biol. 124,45 1 4 5 6 . Vorbrodt, A,, Konwinski, M., Solter, D. & Koprowski, H. (1977). Ultrastructural cytochemistry of membrane-bound phosphatases in preimplantation mouse embryos. Dev. Biol. 55, 1 17-1 34.

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Watson, A.J. (1992). The cell biologyofblastocyst development. Molec. Reprod. Devel. 33,492-504. Watson, A.J. & Kidder, G.M. (1988). Immunofluorescence assessment ofthe timing of appearance and cellular distribution of NdK-ATPase during mouse embryogenesis. Dev. Biol. 126, 80-90. Watson, A.J., Damsky, C.H. & Kidder, G.M. (1990a). Differentiation of an epithelium: factors affecting the polarized distribution of Na+, K+-ATPase in mouse trophectoderm. Dev. Biol. 141, 104-1 14. Watson. A.J., Pape, C.. Emanuel, J.R., Levenson, R. & Kidder, G.M. (1990b). Expression of Na,K-ATPase a and p subunit genes during preimplantation development of the mouse. Devel. Genet. 1 1 , 4 4 8 . Wiley, L.M. (1984). Cavitation in the mouse preimplantation embryo: NdK-ATPase and the origin of nascent blastocoele fluid. Dev. Biol. 105,33&342. Wiley, L.M. & Eglitis, M.A. (1981). Cell surface and cytoskeletal elements: cavitation in the mouse preimplantation embryo. Dev. Biol. 86,493-501. Wiley, L.M. & Obasaju, M F. (1988). Induction of cytoplasmic polarity in heterokaryons of mouse 4-cell-stage blastomeres fused with 8-cell- and 16-cell-stage blastomeres. Dev. Biol. 130, 276-284. Wiley, L.M. & Obasaju, M.F. (1989). Effects of phlorizin and ouabain on the polarity of mouse 4-ce11/16-celI stage blastomere Ireterokaryons. Dev. Biol. 133,375-384. Wiley, L.M., Kidder, G.M. & Watson, A.J. (1990) Cell polarity and development of the first epithelium. BioEssays 12,67-73. Wiley, L.M., Lever, J.E., Pape, C. & Kidder, G.M. (1991). Antibodies to a renal Na+/glucose cotransport system localize to the apical plasma membrane domain of polar mouse embryo blastomeres. Dev. Biol. 143, 149-161. Wiley, L.M., Wu, J.X., Harari, I. &.Adamson, E.D. (1992). Epidermal growth factor receptor mRNA and protein increase after the four-cell preimplantation stage in murine development. Dev. Biol. 149,247-260. Winkel, G.K., Ferguson, J.E., Takeichi, M. & Nuccitelli, R. (1990). Activation of protein kinase C triggers premature compaction in the 4-cell stage mouse embryo. Dev. Biol. 138, 1-15. Ziomek, C.A. & Johnson, M.H. (1980). Cell surface interactions induce polarization of mouse 8-cell blastomeres at compaction. Cell 21, 935-942. Ziomek, C.A. & Johnson, M.H. (1981). Properties of polar and apolar cells from the 16-cell mouse morula Roux’s Arch. Dev. Biol, 190,287-296.

SIGNALS A N D MECHANISMS OF SORTING IN EPITHELIAL POLARITY

Cara J. Cottardi and Michael J. Caplan

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 A. Epithelial Membrane Polarity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Sorting Pathways in Polarized Cells . . . . . . . . . . . . . . . . . . . . 99 C. The Sorting Signal Paradigm. . . . . . II. Sorting Signals in Epithelial Membrane Polarity A. Considerations Relevant to the Study of Sorting Signals . . . . . . . . . . . . . 102 B. "Default" Sorting Pathways and the interpretation of Sorting Si C. Multiplicity of Signals and Epithelial Polarity, . . . . . . . . . . . . . D. The Discovery of Distinct Basolateral Targeting Determinants. E. Apical Sorting: GPI-linkages and Glycosphingolipids. . . . . . . . F. Tissue and Cell-Type Specificity of Membrane Polarity. . . . . . . . . . . . . . 112 Ill. Polarized Sorting and Targeting Machinery: Elements of the lntracellular Protei A. CTPases and Epithelial Polarity 6 . Rabs . . . . . . . . . . . . . . . . . . . . C. The SNARE Paradigm and Epithelial Polarity D. Insights from the Membrane Traffic-Perturbing Reagent, BFA . . . . . . . . . 11 9

Advances in Molecular and Cell Biology Volume 26, pages 95-131. Copyright 0 1998 by JAI Press Inc. All right of reproduction in any form reserved. ISBN: 0-7623-0381-6

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E. BFA, Vesicle Bud Formation, and Polarized Trafficking Events . . . . . . . . 119 F. Heterotrimeric G Proteins and Sorting . . . . . . . . . . . . . . . . . . . . . . . . . . 121 G. Insights from Genetic Models. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121

1.

INTRODUCTION

Polarized epithelial cells have long captured the attention of cell biologists and cell physiologists. This is largely because the architecture of these cells so tellingly bespeaks their function. At the electron microscopic level, one of the most apparent and fundamental features of this cell type is its polarized organization of intracellular organelles and its structually and compositionally distinct lumenal (apical) and serosal (basolateral) plasma membrane domains (Figures 1A, B). Through the eyes of the physiologist, the polarized epithelial phenotype is an absolute necessity for organ system function. In the most general sense, these cells organize to form a continuous, single layer of cells, or epithelium, which serves as a semi-permeable barrier between apposing and biologically distinct compartments. Within the tubules of the nephron, these cells orchestrate complex ion-transporting processes that ultimately control the overall fluid balance of the organism. At the surface of the gastrointestinal tract, specialized versions of this cell type control the digestion, absorption and immuno-protection of the organism. Thus while polarized epithelial cells can carry out myriad functions, they share one defining feature: a structural polarity which serves their underlying functional polarity. A.

Epithelial Membrane Polarity

The differential distribution of membrane proteins between the plasmalemmal surfaces of polarized epithelial cells enables these cells to both respond to and effect changes upon their environment in a directed fashion. The gastric parietal cell of the stomach, for example, contains a population of H,K-ATPase-rich vesicles. Upon stimulation, these vesicles fuse selectively with the lumenal membrane, resulting in the massive apical secretion of HCl which initiates digestion. Without two important elements of the polarized phenotype, that is, junctional integrity and the precision of this membrane insertion, proton pumps might be delivered to a compartment which would be adversely affected by the secretion of acid. Another illustration of the utility of the polarized phenotype is provided by the principal cells of the kidney, which carry out net sodium absorption through a mechanism which is entirely dependent upon the polarized distribution of two membrane proteins. Sodium absorption is stimulated by the hormone aldosterone, which increases the amount or activity of Na,K-ATPase at the basolateral surface, while increasing the number or activity of apical sodium channels and thus the sodium conductance of the lumenal membrane (Doucet and Barlet-Bas, 1989). Because the Na,K-ATPase generates low intracellular { Na+}, sodium is

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Figure 7. (A) Light micrograph showing the salient features of a polarized epithelium. individual polarized epithelial cells bounded by a junctional complex (jc) come together to form a simple columnar sheet. This sheet, or epithelium, sits on a basement membrane (bm) and serves as a semi-permeable barrier between the lumen (Lcontinuous with the outside world of an organism) and the interstitium (In- interior) of an organism’s tissues. (Photo courtesy of Dr. Marian Neutra, Children’s Hospital, Boston, MA). (6) Electronmicrograph showing the unique morphological features of a polarized epithelial cell. The cell’s apical (Ap) membrane surface is equipped with numerous microvillar (mv) bundles. The basolateral (BI) domain of this particular cell is characterized by extensive interdigitations with the adjacent cell’s lateral membrane. These morphologically distinct apical and basolateral membrane domains are separated by a unique ultrastructure known as the tight junction (tj). This structure is just visible as an area of close, uniform membrane apposition located at the apices between adjacent epithelial cells. (Photo courtesyof Dr. Marian Neutra, Children’s Hospital, Boston, MA).

able to pass from the lumen of the kidney tubule through apical sodium channels and into the cytoplasm down its electrochemical gradient. The Na+ is then pumped across the basolateral membrane and into the interstitum by the sodium pump and is ultimately prevented from leaking back into the lumen by impermeable tight junctions. Therefore, it is the differential assignment of Na’ channels to the apical surface and Na,K-ATPase molecules to the basolateral domain that ensures the vectoriality of this transport process. How the polarized cell assigns these two proteins (and apical and basolateral membrane proteins in general) to their respective surface domains has been the subject of much investigation and is the general focus of this review. It is perhaps important to point out that the fundamental questions of plasma membrane protein aniosotropy are not unique to surface membrane proteins or even to the study of epithelial polarity. The Golgi apparatus, for example, is a polarized

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organelle whose cis- and trans-most cisternae are structurally and biochemically distinct. This organization is thought to enable the ordered addition and trimming of glycoprotein sugar residues as they traverse the stacked cisternae. As is clearly represented in the breadth of topics covered in this book, numerous cell types adopt a polarized state for some functional purpose. The propagation of a nervous impulse from dendrite to axon requires compositionally different membrane proteins in each of these domains, while the localization of determinants to specific parts of an egg’s cytoplasm gives rise to cells with different growth potentials and the necessary assymetries required for embryo development . What we hope will become clear in this chapter and related chapters in this book is that we are beginning to appreciate the universality of polarity. The mechanisms involved in establishing and maintaining the polarized state appear to be so fundamental that some of the schemes through which a cell is able to localize a particular protein to a given cellular domain are turning out to be conserved between epithelia and neurons, and even between epithelia and yeast. While the need for protein asymmetries in development, or membrane polarity in epithelial transport is clear, the means through which it is achieved are only beginning to be elucidated. Before we embark upon our review of the field, we first introduce the conceptual framework onto which the results in this field are organized and interpreted. First, a protein destined to accumulate with a polarized distribution needs to be recognized as different from other proteins. We presume that what is recognized is some structural aspect of the protein itself. We refer to that part of the protein that is recognized for polarized localization as a sorting signal or localization determinant. These two terms are often used interchangeably, but in fact there is a subtle difference between the two. “Sorting signal” is often taken to imply a signal that is recognized and acted upon before the protein is delivered to its ultimate residence. Sorting signals are thought to be those signals that enable a cohort of similar proteins with similar destinations to be sorted and sifted away from all of the other molecules traversing the biosynthetic pathway at the same time. A “localization determinant” is perhaps a more general term that carries fewer mechanistic implications. It is defined here as the determinant that specifies a protein’s polarized distribution, but it does not make a distinction between recognition that takes place before the protein has reached its final destination or after (e.g., through a selective retention mechanism). The proteins which serve to recognize a particular signal and act upon it are generally referred to as sorting machinery. Often, a distinction is made in the literature between “sorting” and “targeting machinery.” In these cases, the sorting machinery is exclusively those elements which recognize the sorting signal. Any downstream effectors of this sorter that orchestrate the vectorial directing of a vesicle to its final destination are referred to as targetting machinery. A simple schematic of these elements is presented in Figure 2. As is discussed in the second half of this review, we know much more about general targeting machinery than the sorters themselves.

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domain-specific vesicle

t C. Sorter for membrane protein

Golgi

Figure 2. Conceptual framework for sorting in polarized cells. This illustration offers one of many possible ways to think about how a secretory or membrane protein could be sorted into a vesicle. It is presumed that the "sorters" will recognize a sorting signal ("1 embedded within the protein structure. It seems likely that this recognition event would need to take place in the lumen of the Golgi for a secretory protein, but this might not be necessary for a membrane protein, which could interact with a sorter from either a lumenal- or cytoplasmic-facing signal domain. Ultimately, the sorted protein(s) could be contained within a "domain-specific vesicle," which would then be targetted (with the help of protein targetting machinery X, Y, and Z)to the appropriate apical or basolateral surface domain.

B.

Sorting Pathways in Polarized Cells

It is thought that proteins destined for either the apical or basolateral domain of a polarized cell occupy the same Golgi cisternae during their biosynthesis ( M a t h and Simons, 1984; Misek et al., 1984; Rindler et al., 1984; Fuller et al., 1985; Pfeffer et al., 1985). Immunoelectron microscopic studies performed on nonpolarized

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endocrine cells which manifest two biochemically and kinetically different secretory pathways suggested that the process of sorting components away from one another takes place at the TGN (Orci et al., 1987; Tooze et al., 1987). However, recent studies have demonstrated that sorting may not take place exclusively at the TGN. Sorting mechanisms have been suggested to take effect as early along the biosynthetic pathway as the ER (Balch et al., 1994) as well as at the recycling endosome (Matter et al., 1993; Matter and Mellman, 1994). In hepatocytes, sorting appears to occur after all newly synthesized membrane proteins are delivered to the basolateral plasmamembrane (Bartles et al., 1987). Similar delivery routes have been detected in polarized intestinal epithelial cell lines (Matter et al., 1990). Finally, in at least one subclone of the canine renal MDCK cell line, sorting may take place both at the Golgi as well as at the level of the plasma membrane. While most proteins in this cell line are sorted in the TGN, the Na,K-ATPase can be preferentially localized to the basolateral membrane through domain-specific stabilization mechanisms after random insertion into both plasmamembrane domains (Hammerton et al., 1991; Siemers et al., 1993). Apically and basolaterally sorted proteins have been shown to be packaged into distinct classes of Golgi-derived vesicles (Wandinger-Ness et al., 1990) which are ultimately targeted to their appropriate domains. Recently it has been shown that membrane and secretory proteins are segregated into distinct vesicular carriers upon transport from the Golgi to the basolateral surface of hepatocytes (Saucan and Palade, 1994) The extent to whch distinct basolateral (or apical) proteins are cosorted and incorporated within the same vesicle either due to common localization signals or the ability to co-aggregate has not yet been determined. After proteins are sorted, the targeting of a vesicle to a particular surface domain can occur directly (vectorially) from the TGN to the apical domain (Matlin and Simons, 1984; Rindler et al., 1984; Fuller et al., 1985), basolateral domain (Caplan et al., 1986) or indirectly as has been shown for the poly-immunoglobulin receptor (pIgR) (Mostov and Deitcher, 1986). In the latter case, the protein is first targeted to the basolateral surface where the receptor can bind its ligand and is then transported to the apical surface via a process known as transcytosis (reviewed in Mostov and Simister, 1985). As noted above, in hepatocytes all apical proteins studied to date make use of this indirect pathway for apical delivery (Bartles et al., 1987), while cell lines derived from intestine and kidney can employ both routes for surface delivery (Matter et al., 1990; Casanovaet al., 1991;Low et al., 1991) While the details of the routes have been determined for a number of sorting pathways, the molecular signals and recognition components which control each of them are not well understood. The search for these molecular signals and recognition components has been the focus of much study over the last 15 years. During this period, the subjects of protein sorting and epithelial polarity have been extensively reviewed. Several of these reviews are listed here for those seeking more background on specific aspects of this field: for general reviews on protein sorting pathways (Burgess and Kelly,

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1987); general concepts of sorting and targeting (Caplan and Matlin, 1989); a discussion of the mechanisms required for the establishment and maintenance of epithelial polarity (Rodriguez-Boulan and Nelson, 1989); polarized transport of surface porteins and lipids in epithelial cells (Simons and Wandinger-Ness, 1990); comparative epithelial and neuronal polarity (Rodriguez-Boulan and Powell, 1992); the generality of the polarized phenotype (Nelson, 1992); cytoskeleton as a component of the protein sorting machinery (Mays et al., 1994); summary of the few known sorting signals in polarized epithelial cells (Mostov et al., 1992); common signals involved in sorting from the TGN and endosomes (Matter and Mellman, 1994). Perhaps now more than ever before, it is becoming a rather daunting task to provide a synthesis of the observations relevant to the study of epithelial polarity. This is in part due to the fact that important insights into the mechanisms of sorting are being contributed by fields that are not exclusively focussed on epithelial biology. As we discussed in this review, some important contributions are emerging from studies of endocytosis, secretion in yeast and neurons, and the sorting of yeast lysosoma1 enzymes (see Chapter I of this volume), in addition to more “classical” approaches to epithelial polarity. In this review, we explore the current paradigm that the generation and maintainance of distinct membraneous compartments requires “sorting signals,” the recognition domains embedded within the amino acid sequence or polypeptide structure of the protein, and “sorting machinery,” the proteins which interpret and act upon these signals. In the first half, we review and categorize the signals that have begun to be elucidated, as well as discuss the approaches and difficulties associated with finding and interpreting sorting signals. While the polarity field itself has not yet succeeded in characterizing the definitive sorting machinery, numerous components of the membrane budding and fusion apparatus are rapidly being elucidated. We have chosen to review some of the important findings in the field of membrane transport, and in particular examine the potential roles that GTP-binding proteins of the rab, ARF and heterotrimeric classes may play. We also discuss a class of proteins referred to as adaptins as well as the implications that the SNARE hypothesis may have for epithelial polarity. Although these components have not been shown to be directly involved in sorting per se, it is becoming increasingly clear that in a general sense, the composition of the membrane vesicle budding and fusion machinery may be part of the overall apparatus which “acts upon” the sorted species and contributes to domain specific surface targeting.

C. The Sorting Signal Paradigm AS stated above, the paradigm for conceptualizing the mechanisms responsible for biosynthetic sorting requires that each protein contains signal information embedded within its polypeptide sequence/structure (sorting signal) which is interpreted and acted upon by components referred to as sorting machinery. This scheme

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takes its cue from the process through which ribosomes translating secretory and membrane proteins are targeted to the endoplasmic reticulum to initiate cotranslational protein translocation (Blobel, 1980). Prior to the elucidation of this process, it was suggested that protein targeting might require cellular sorting machinery to recognize certain signals which would be shared by proteins with common destinations (Blobel, 1980). Shortly after this suggestion, it became clear that targeting to the RER, mitochondria and chloroplasts required short, contiguous, N-terminal signal peptides (reviewed in Burgess and Kelly, 1987). In the case of the former, the signal was recognized by a receptor, SRP (Lingappa et al., 1978; von Heijne, 1984; Kurzchalia et al., 1986; Walter and Lingappa, 1986). Subsequently, a number of short, contiguous amino acid domains have been shown to play a role in later stages of post-synthetic targeting. These include: (1) the KDEL and adenovirus E l 9 signals which ensure the retention or recapture of resident ER proteins (Munro and Pelham, 1987; Nilsson et al., 1989); ( 2 ) a transmembrane domain signal responsible for Golgi retention (Swift & Machamer, 1991; Machamer, 1993); (3) the cluster of positively charged lysine residues (SV40-NLS) sufficient for nuclear targeting (Richardson et al., 1986); (4) the critical tyrosine/ “tight-turn’’ structural motif which can mediate localization to clathrin coated-pits (Goldstein et al., 1985; Pearse and Robinson, 1990; Collawn et al., 1991); and (5) the discovery that lysosomal hydrolases were targeted to lysosomes through the recognition of a phosphorylated sugar residue (mannose-6-phosphate; reviewed by Kornfeld and Mellman, 1989). In several of these cases receptors for these signals have been well-characterized: the signal recognition particle (SRP) for secretory and membrane proteins (Walter and Lingappa, 1986), the mannose-6-phosphate receptor (M6PR) for the targeting of lysosomal hydrolases to the lysosome (Sly and Fischer, 1982; VonFiguraandHasilik, 1986), the KDELreceptor (Tanget al., 1993) and the adaptins which couple coated pit localization sequences to clathrin cages (Pearse and Robinson, 1990; Robinson, 1994).

II.

SORTING SIGNALS IN EPITHELIAL MEMBRANE POLARITY A.

Considerations Relevant to the Study of Sorting Signals

The search for definitive signals which mediate the delivery of proteins to a particular epithelial surface domain has proven to be quite difficult. This is due in part to general limitations imposed by certain molecular biological approaches, as well as to some inherent difficulties specific to the investigation of epithelial polarity. Our goal in this section is to outline reasonable criteria for the identification of a sorting signal. The observation that the influenza and vesicular stomatitis viruses bud from opposite surface domains of polarized MDCK cells (Madin Darby Canine Kidney)

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(Rodriguez-Boulan and Sabatini, 1978) spawned an extensive search in which chimeric and deletion analyses were applied to the problem of identifying the underlying apical and basolateral sorting signals (reviewed in Caplan and Matlin, 1989). These efforts to characterize sorting signals have generally involved the generation of chimeric or truncated contructs prepared from portions of apical and basolateral membrane proteins. Through analysis of the subcellular distributions of the resulting proteins, sorting information can, at least in theory, be assigned to particular portions of the parent molecules. While a large number of chimeric and truncated viral glycoproteins have been generated and analyzed, it has been difficult to interpret many of the resultant observations. With the benefit of hindsight, we now know that these difficulties can be attributed to a number issues that we discuss in more detail below (including the tertiary stuctures of the experimental constructs, the confounding possibilities introduced by uncharacterized default pathways, and the potential for multiple and hierarchical signals to be embodied within the structures of the studied proteins). Until recently (Thomas and Roth, 1994), the analysis of viral spike glycoproteins did not produce a definitive sorting signal. Much of the uncertainty associated with this work is likely attributable to the fact that these studies engineered chimeras from portions of structurally dissimilar molecules. The tertiary structures of the resultant chimeras may thus differ substantially from those of either parent molecule, which may in turn exert unpredictable effects upon sorting behavior. Clearly, if sorting signals are formed from domains arising from noncontiguous regions of a polypeptide, for example, in much the same manner that heterotrimeric G proteins are thought to “see” their effectors (Berlot and Bourne, 1992), or in the way that the human growth hormone receptor (hGHbp) is thought to interact with its ligand (Cunningham and Wells, 1989), it is easy to imagine how the structural integrity of the putative sorting signal could become compromised in a chimeric construct. While producing a rough map of the signal-bearing domain of a protein can be relatively straight forward, determining the exact residues which constitute the signal is turning out to require a collaboration between many different types of mutagenesis approaches. Often, contradicting results can arise from alanine scanning, truncation and point mutation/deletion mutagenesis, since a mutated protein can manifest impaired sorting behavior even though the altered residues are not part of the actual sorting signal (Aroeti et al., 1993). It is becoming clear that a judicious and thorough comparison of many different types of mutagenesis approaches may be necessary to determine definitively the key residues necessary for sorting. B.

”Default” Sorting Pathways and the Interpretation of Sorting Signals

Perhaps another difficulty in looking for apical or basolateral sorting signals is that the default pathway for “signal-less” membrane proteins is still not known. A protein that is sorted “by default” is, by definition, unable to interact with and be

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acted upon by any sorting machinery whatsoever. In theory, at least, such “unsorted’’ proteins may be distributed with polarity, depending on the nature and characteristics of the membrane vesicular traffic arising from the Golgi complex in a particular cell type. Obviously, if the localization of a protein construct under study is identical to that produced by the cell’s default pathway, elucidation of a signal will be difficult, since elimination of the signal will not alter the protein’s distribution. Thus, one can appreciate the difficulty in assigning localization information to a particular domain in the context of an undefined default pathway. This caveat accounts for at least some of the reasons which explain why a definitive basolateral sorting signal in the C-terminal domain of VSV-G protein took so long to discern. In the following example we summarize the HA-VSVG spike glycoprotein chimera literature as a means to illustrate the difficulties in interpretating these types of studies. When acDNA encoding the influenza HA was expressed in MDCK cells, the encoded protein localized to the apical membrane (Roth et al., 1983), while a cDNA encoding the VSVG polypeptide produced a protein that is localized to the basolateral domain (Gottlieb et al., 1986b; Stephens and Compans, 1986). When truncation mutants were expressed in which soluble ectodomain versions of these proteins were synthesized, the VSVG ectodomain was secreted from both apical and basolateral domains (Stephens and Compans, 1986; Gonzalez et al., 1987) while the HA ectodomain was predominantly secreted from the apical domain (Gonzalez, et al., 1987; Roth et al., 1987b). Based on evidence that the default pathway for secreted proteins leads to nonpolarized secretion from both surface domains (Kondor-Koch et al., 1985; Gottlieb et al., 1986a; Caplan et al., 1987), it was reasoned that the ectodomain of HA encodes an apical sorting signal while the VSVG ectodomain lacks signal information. This was further confirmed by the observation that a hybrid HA-VSVG protein comprising the HA ectodomain fused to the VSVG transmembrane and cytoplasmic tail region was targeted to the apical membrane (McQueen et al., 1986; Roth et al., 1987a). But if the VSVG ectodomain is randomly secreted and the VSVG tail domain fused to HA is apical, which domain of VSVG encodes basolateral sorting information? The complementary hybrid comprised of the ectodomain of VSVG (presumably signal-less) tethered to the HA transmembrane and tail region (perhaps also signal-less) was targeted either to the basolateral membrane or to both surface domains (McQueen et al., 1986; Puddington et a]., 1987; Roth et a]., 1987a; Compton et al., 1989). The interpretation of the behavior of this chimera was clearly complicated; it was suggested that this protein could be pursuing its distribution by default. (As discussed above, the default pathway for membrane proteins is still not defined in polarized cells). An alternative interpretation was that the VSVG ectodomain indeed contains basolateral sorting information, but that perhaps this domain needs to be tethered to the plasmamembrane with a transmembrane anchor in order to interact with its presumptive sorting machinery. This interpretation, however, was proved incorrect by the observation that the anchoring of this ectodomain to the membrane through a

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lipid-linkage resulted in apical targeting (Brown et al., 1989). Interestingly, when the ectodomain of the normally apical placental alkaline phosphatase (PLAP) was attached to the VSVG transmembrane and cytosolic tail domains (which were though to lack a dominant signal), the resulting chimeric protein was targeted basolaterally. It is difficult to reconcile the HA-VSVG and PLAP-VSVG chimeras without invoking hierarchical and competing signals. Recently, a basolateral targeting signal has been precisely localized to the cytoplasmic domain of the VSVG protein (Thomas and Roth, 1994). In light of the vicissitudes which attended the interpretation of each round of chimeric constructs discussed above, it was certainly unexpected that definitive sorting information would be localized to the cytoplasmic tail of VSVG. The nature and function of this signal will be discussed in depth below. The preceding discussion was presented simply to reinforce the caveat that the default pathway, protein structural considerations and the possible interactions between “dominant” and “recessive” sorting signals can considerably cloud the interpretation of chimera experiments.

C. Multiplicity of Signals and Epithelial Polarity Recent studies of the polymeric immunoglobulin receptor (pIgR), the low density lipoprotein receptor (LDLR) and polytopic hetero-oligomeric proteins (H,KATPase and Na,K-ATPase) suggest that individual proteins can interact in multiple and complex fashions with the machinery responsible for surface targeting. It is becoming increasingly clear that there can be an array of signals encoded within an individual protein, and the sorting problem is becoming evermore complicated by the apparent redundancy, multiplicity and hierarchical nature of these signals (Matter et al., 1992; Mostov et al., 1992). For example, Brewer and Roth’s (1991) demonstration that they could completely overwhelm the apical signal present in the HA ectodomain and redirect it to the basolateral surface by changing a single amino acid in this protein’s cytoplasmic tail strongly suggests that multiple signals present in a single protein can interact in a heirarchical fashion. The newly created cytoplasmic signal is dominant over the presumed apical sorting signal present in the ectodomain of HA. As discussed below, the LDL receptor has been shown to encode redundant, basolateral sorting information, since either of two cytoplasmic determinants could independently mediate basolateral delivery (Matter et a]., 1992). Moreover, the protein may also contain acryptic apical sorting signal in its ectodomain, since a cytoplasmic tail-minus construct of this protein (CT12) is sorted with great efficiency to the apical membrane in MDCK cells (Matter et al., 1992). An ectodomain apical localization signal has also been found within the pIgR, whose initial surface delivery is to the basolateral plasmalemma. Why do these proteins need multiple signals? What does the LDLR gain by expressing two basolateral localization signals? Recent studies (discussed in greater detail in the following section) have more finely decoded these two signals and are revealing functional differences. For ex-

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ample, the “membrane proximal determinant” encodes coated-pit internalization information, while the “membrane distal determinant” appears to ensure efficient sorting from a basolateral endosome back to the basolateral surface (Matter et al., 1993). Analysis of the sorting behavior of multisubunit ion pumps provides further insight into the possible utility of multiple signals (reviewed in Gottardi et al., 1993) The gastric H,K-ATPase and the Na,K-ATPase are close cousins in the large family of P-type ion transporting ATPases. Both are composed of 100 kDa a-subunits and heavily glycosylated 55 kDa P-subunits. They share similar reaction mechanisms and catalytic properties and, not surprisingly, are highly homologous at the amino acid sequence level. The a-subunits are -65% identical, whereas the P-polypeptides manifest roughly 40% identity. While the Na,K-ATPase is a basolateral protein in most polarized epithelial cell types (with the exception of neural epithelia such as choroid plexus and retinal pigment epithelium), the H,K-ATPase occupies the apical membrane and a pre-apical storage compartment in gastric parietal cells. Hormonal stimulation of gastric acid secretion induces fusion of the membrane vesicles which comprise the intracellular reservoir with the plasma membrane, resulting in delivery of the H,K-ATPase to the apical cell surface. During the interdigestive period, the H,K-ATPase is re-endocytosed and returned to its storage compartment. Chimera studies reveal that each subunit of the H,K-ATPase possesses a sorting signal which participates in regulating this complex traffic (Gottardi and Caplan, 1993). The a-subunit is endowed with a dominant apical targeting signal, which can drive the apical sorting of chimeric pumps expressed in both MDCK and LLC-PKl renal epithelial cells. The P-subunit of the H,K-ATPase possesses a tyrosine-based endocytosis signal (Roush et al., manuscript submitted). This signal causes the protein to be sorted basolaterally when it is expressed in MDCK cells and apically when it is expressed in LLC-PK1 cells. The Na,K-ATPase P-subunit does not possess a similar sequence domain. It seems likely that the two H,K-ATPase signals participate in distinct stages of pump sorting in the gastric parietal cells. The apical signal in the a-subunit probably mediates the sorting of the entire complex to the apical membrane or the pre-apical storage compartment, whereas the P-subunit signal is responsible for ensuring the re-internalization of the pump following the cessation of secretagogue stimulation (Courtois-Coutry et al., 1997). It remains to be determined why the P-subunit’s tyrosine-based signal is differentially interpreted by MDCK and LLC-PK1 cells. Investigation of this phenomenon may shed light on the nature and function of the epithelial sorting machinery. This apparent trend towards a multiplicity of signals is not entirely surprising, since many proteins are required to perform highly sophisticated feats of membrane targeting during the course of their transits throughout the endomembranous networks of the cell. For example, the pIgR receptor expressed in its native hepatocytes or by transfection in MDCK cells travels first to the basolateral membrane to pick up ligand and is then transported to the apical surface domain. It appears that an apical sorting signal in this protein’s ectodomain might be required for basolateral to

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apical transcytosis, while a basolateral signal in the cytoplasmic domain ensures the initial basolateral delivery. Unlike proteins that are constitutively expressed at one surface domain, a number of distinct and individually acting signals are necessary to orchestrate the more complicated surface targeting events displayed by pIgR receptor, and other molecules like it. Obviously, the hierarchical (both temporal and spatial) regulation of each signal will be of utmost importance in ensuring that a protein follows a physiologically relevent trafficking pathway. Recent evidence, for example, demonstrates that the pIg receptor undergoes phosphorylation on a cytosolic serine residue around the time that it is delivered to the basolateral surface (Larkin et al., 1986). This phosphorylation event appears to inactivate the protein’s basolateral signal and thus permit its transcytosis to the apical membrane (Casanova et al., 1990).

D.

The Discovery of Distinct Basolateral Targeting Determinants

Perhaps not surprisingly, the greatest advances in the elucidation of sorting signals have been made with single membrane-spanning monomeric or homooligomeric proteins (e.g., pIgR, LDL-R, TfR). With these molecules the requirements for surface expression are easily met and the effects of mutagenesis on tertiary structure can be assessed through well-characterized functional assays, such as receptor-ligand or antibody binding. Through deletion analysis and heterologous expression in MDCK cells, it was determined that the pIgR (Casanova et al., 1991) and the LDLR (Hunziker et al., 1991) each contained basolateral targeting determinants which mapped to short, contiguous regions of their cytoplasmic domains (Table 1). Both signals could be grafted onto heterologous proteins and cause them to be targeted to the basolateral surface, supporting the idea that each determinant was truly an autonomous basolateral sorting signal. Exhaustive mutagenesis studies have more finely mapped each of these determinants. The LDLR possesses two distinct basolateral targeting determinants, one that is “coated-pit related” (proximal determinant) and another which is tyrosine-dependent but not capable of mediating localization into coated-pits (distal determinant) (Matter et al., 1992; 1993; 1994). Interestingly, the polymeric immunoglobulin receptor (pIgR) signal may constitute yet another class of basolateral targetting determinant, since it shares little in the way of sequence homology with either determinant of the LDLR and shows weak tyrosine dependence (Aroeti et al., 1993). The general characteristics of these three determinants and the degree to which they are related are only beginning to be eluciated (Matter et al., 1994; Thomas and Roth, 1994). An attempt to categorize these basolateral sorting determinants has been made by Matter et al. (1994) and is summarized in Table 1. Before discussing the nature of the “coated-pit related” basolateral targeting determinant, it is necessary to be familiar with the signals that are known to mediate the accumulation of plasma membrane receptors into clathrin-coated pits (Goldstein et al., 1985). It is now generally accepted that tyrosine- and dileucine-

Table 7. Coated Pit Unrealted localization Signals

Classification of Basolateral Sorting Determinants

Strong Jyrosine Dependence LDL receptor- distal determinant

--_

-- Q D C Y S Y PS R Q M V S L E D D V A -

LDL-R (distal)

Transferrin receptor (TfR) Weak Tyrosine Dependence plg receptpr (plgR)

-

-- R H R R N V D R V S I G S Y R T -- including downstream acidic residues

plgR

Coated-Pit Related Localization Signals Jyrosine Dependent A

0

D

LDL receptor- proximal determinant

LDL-R (proximal)

Hemagglutinin (HA)-Y543

HA-Y54 3 VSVG

VSV G Lysosomal membrane glycoproteins (Igp-Nlamp-1) Lysosomal acid phosphatase (LAP) Asialoglycoprotein receptor (ASGP-R) Nerve growth factor receptor (NGF-R)

Igp-120 LAP

-- N F D N P V Y Q K T T ? D E V H --

-- N G S L Q Y R I C I --- C I K L K H T K KB Q I Y T D I EM --- K R S H A CY Q T I --

-- Q A Q P P G Y R H V A D G E D H A --

ASGP-R (H1 subunit)

MT K EY Q DLQ H L D N E E S D H H .

Jyrosine hdependent

IgG Fc receptor (di-leucine dependent)

FcRILB2

-- NI ! I li S L 1 K H -- (including downstream acidic residues)

Notes: This table was adapted from that of Matter et al., 1994 and Matter and Mellman, 1994. The classification of signals (e.g., coated-pit related, unrelated, etc.) is shown on the left column, while the actual signal for most of the proteins is shown on the right. Critical residues for basolateralsorting are shown in bold face t.,pe, while other important residuesare underlined.The dashes (-1 present over the residues E D D of the LDL-R (distal)signal and the E D E of the LDL-R (proximal)signal denote that these amino acids are acidic and important for sorting signal function.

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containing sequence motifs present in the cytoplasmic tails of a number of coated-pit clustering proteins serve as the critical recognition elements for the adaptor components of clathrin coats (Pearse and Robinson, 1990; Trowbridge, 1991). More recently, numerous studies have demonstrated a strong relationship between the signals which mediate localization into coated pits and a subset of those involved in basolateral targeting (Brewer and Roth, 1991; Hunziker et al., 1991; LeBivic et al., 1991). For example, Brewer and Roth (1991) found that the apically targeted HA molecule could be completely rerouted to the basolateral membrane by replacing a strategically localized cysteine residue (cys 543) with a tyrosine in the cytoplasmic domain. This tyrosine was also sufficient to localize this protein into coated-pits and direct the protein’s incorporation into endosomes. This observation that an endocytosis signal might also double as a basolateral targeting signal led to the suggestion that the recognition determinants for endocytosis and for TGN-to-basolateral targeting might be similar or identical to one another. Thorough mutagenesis studies on the coated-pit localization and basolateral sorting determinants of HA-Y543 (Thomas and Roth, 1994; Lin et al., 1997), VSVG protein (Thomas et al., 1993), and the LDLR (Matter et al., 1994), however, have led to a revision of this initial interpretation. It turns out that the “endocytosis signal” of both the HA-YS43 and the LDLR (proximal signal) can be resolved into two overlapping but distinct signal components. In other words, there is information recognized for endocytosis that is distinct from that recognized for basolateral sorting, even though the sequences are in part superimposed and share marked similarity. Table 2 shows the systematic mutagenesis that ultimately unraveled this relationship. Brewer and Roth (1991) found that HA-Y543 is capable of both basolateral sorting and endocytosis. The second generation mutant HA-Y543,RS46, however, behaved as aprotein that was capable of endocytosis, but whose basolateral localization was inhibited (Lin et al., 1997). Similar results were found with the LDLR proximal determinant. Matter and colleagues (1994) showed that the truncation mutant CT27 was basolaterally targetted and rapidly endocytosed, while the removal of terminal acidic residues in CT22 proTable 2. Evidence that Coated-Pit-Related Determinants Encode Two Distinct Signals. Protein

Signal Region

Cell Surface Localization

Endocytosis Competent! no

HA HA-Y543

-N -N

CSL Q C R IC I C SL Q Y R 1 C I

apical basolateral

Yes

HA-Y543, R546

-N

G SLQY R I R I

inhibited

Yes

basolateral apical

Yes ves

LDL-R (Proximal Determinant) -N F D N P V Y Q K T T E D E V H CT-27 -N F D N P V Y Q K T T CT-22 Notes:

This table summarizes data from Thomas and Roth, (1994)and Matter et al., (1993)demonstratingthat the coated-pit related basolateral sorting determinant could be resolved into two distinct signals, one which mediates basolateral localization, the other which mediates endocytosis.

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duced a protein that was not capable of basolateral targetting, but could nonetheless be endocytosed. Thus, the initial correlation between endocytosis signals and basolateral targeting has now resolved into two distinct but overlapping signals that can share common residues for their respective activities. The implications of this result are very exciting for the field of epithelial polarity. First, they suggest that the signals for basolateral sortingkargetting may be structurally similar to signals for clathrin-coated pit localization and endocytosis. The involvement of similar signals suggests that the sortinghecognition molecules themselves may be related. At least for endocytosis signals, there is evidence in favor of clathrin “adaptins” (of the AP2 plasma membrane class) playing a role in recognizing these sequences (Pearse, 1988; Glickman et al., 1989; Beltzer and Spiess, 1991; Sorkin and Carpenter, 1993; Sosa et al., 1993). In light of the recent characterization of adaptin related molecules (COPS, discussed in section 111, below), it has been suggested that a family of structurally and functionally similar sorting adaptors may serve as the sorting machinery which interacts with these basolateral sorting signals (Matter et al., 1994). The findings support the more general contention that sorting at the level of the TGN may be mechanistically similar to that at the level of the endosome (Matter et al., 1993, 1994). Taken altogether, there now appear to be two general classes of basolateral targeting determinants. One of these is biochemically related to the signals that mediate sorting into coated pits. This type of signal can be colinear with an endocytosis determinant and may share the critical tyrosine residue required for the activity of both, but it is nonetheless distinct and dissociable from an endocytosis signal. The second class of basolateral targeting determinants appears to be unrelated to clathrin-coated pit localization signals, although it may also strongly depend on a tyrosine for activity. This second type of determinant appears to be unique to the LDLR, pIgR (Casanovaet al., 1991) and the TW (Dargement et al., 1993), although these signals share no primary sequence homology with one another. It is possible however, that this second determinant present in these three proteins may be mutually similar in three-dimensional structure but not in primary sequence. In this context it is important to note that adaptor proteins are thought to recognize tyrosine residues in the context of a tight turn, which can be achieved by many different primary sequences (Glickman et al., 1989; Collawn et al., 1990, 1991; Bansal and Gierasch, 1991). More detailed analyses are revealing that while the dependency on tyrosine is crucial, other residues which are acidic and C-terminal to the tyrosine are also important. Matter et al. (1994) demonstrated that the clusters of two or more acidic amino acids downstream from a tyrosine, phenylalanine or di-leucine are important for signal function (see Table 1). While the authors of this study have argued that it is premature to propose a common motif characteristic of all basolateral targeting determinants, they have found that this critical aromatic amino acid followed by acidic residues can be discerned in the cytoplasmic domains of many known proteins which are targeted to the basolateral membrane of MDCK cells, including Ecadherin, transferrin receptor, cation-independent and dependent mannose-6-

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phosphate receptors, LAP, pIgR and FcRIIB2. (see discussion of Matter et al., 1994). As these authors have suggested, it will be exciting to define mutations that will prevent the recognition of these sequences so that the identification and characterization of the molecules which serve to interact with and interpret these signals can be facilitated. E.

Apical Sorting: GPI-linkages and Glycosphingolipids

An ever growing list of proteins are anchored to membranes through a covalent attachment to glycosylphosphatidylinositol or GPI. Proteins of this class are initially synthesized on bound polysomes as transmembrane polypeptides and, while still resident within the ER, are cleaved from their transmembrane portions and transferred covalently to lumenally facing glycosyl-phosphatidylinositol molecules (Cross, 1990). GPI-anchored proteins are widely distributed with respect to both cell type and function. Members of this class of proteins include protozoal surface coat proteins (e.g, the variant surface glycoproteins of trypanosomes), differentiation antigens (e.g., Thy- I), adhesion molecules (e.g., the GPI-linked isoform of N-CAM), hydrolases (e.g., alkaline phosphatase and Snucleotidase), and receptors (folate receptor). The functional advantages that this membrane linkage confers upon a particular protein is presently unclear, and has been the focus of a great deal of attention (reviewed in Brown, 1992). In general, the GPI-linkage has been suggested to be important for enabling proteins to “c1uster”at a surface density much higher than is possible for single-pass transmembrane proteins (Hooper, 1992). Studies have also shown that these clusters of GPI-anchored proteins may be important for certain cell surface signal transducing events (reviewed in Anderson, 1993). GPI-linked proteins captured the attention of epithelial biologists because of their polarized distribution in MDCK cells (Lisanti et al., 1988) and other cultured epithelial cell lines (Lisanti et al., 1990). The nearly exclusive correlation of membrane anchoring via GPI with apical localization raised the question as to whether or not the GPI membrane anchor was itself a signal for apical targeting. Chimeric analyses showed clearly that the GPI-linkage is sufficient for apical targeting in MDCK cells (Brown et al., 1989; Lisanti et al., 1989a,b). Of course, in the absence of a known default pathway for membrane proteins, it remains formally possible that the GPI-anchor prevents a protein from gaining entry into the basolateral sorting pathway. Moreover, the fact that the cytoplasmic tail-minus versions of the LDL and pIg receptors are directly targetted to the apical membrane is consistent with the possibility that apical sorting occurs by default (discussed in Matter and Mellman, 1994). Nonetheless, the GPI-linkage is the field’s best accepted apical localization signal characterized to date. Interestingly, glycosphingolipids (GSLs) share the apical preference of GPI-linked proteins and are generally found exclusively in the outer leaflets of the apical membranes of MDCK cells. The means through which GPI-anchored proteins and glycosphingolipids (GSLs) are sorted and subsequently targetted to the apical membrane are poorly un-

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derstood. It has been shown that GSLs manifest biophysical properties which enable them to self-associate or form clusters in the plane of the membrane (Thompson and Tillack, 1985). These properties have been invoked to support the proposal that GSL clustering occurs at the level of the TGN, and that newly synthesized GPI-linked proteins might co-cluster with these lipids (Simons and van Meer, 1988). It has been further suggested that apically-destined transmembrane proteins could similarly be sorted through an ability to co-cluster with GSLs and GPI-linked proteins (Simons and Wandinger-Ness, 1990). According to this model, apical sorting could take place through selective inclusion within these GSL microdomains, while certain basolateral membrane protein components would be sorted by selective exclusion. However, it should be pointed out that there is still no experimental evidence showing that the GSL clusters are important for apical sorting. One cell line in particular suggests that the role of GSLs in sorting of GPI-anchored proteins may be more complex. A rat thyroid epithelial cell line (FRT) distributes its GSLs and GPI-anchored proteins to the basolateral surface while the polarized distribution of a number of transmembrane proteins is identical to that of MDCKcells (Zurzolo et al., 1993). This suggests that at least some of the apical proteins analyzed (e.g., HA) do not partition with basolaterally directed GSLs. The FRT cell line will serve as an excellent tool for furthering our understanding about the role of glycolipid clustering in the sorting of proteins and lipids in polarized epithelial cells. F.

Tissue and Cell-Type Specificity of Membrane Polarity

Most of the early studies in epithelial polarity used the kidney-derived MDCK cell line as their workbench. However, the last six years has seen the introduction of a number of new cell culture models into the field: CaCo2 (Pinto et al., 1983; Matter et al., 1990; Costa de Beauregard et al., 1995); HT-29 and T-84 (human intestinal epithelial), (Madara et al., 1987; Polak-Charcon et al., 1989; Mikogami et al., 1994);LLC-PK1 (pig kidney proximal tubule) (Hull et al., 1976; Gstrauthaler et al., 1985; Gottardi and Caplan, 1993; Gottardi et al., 1995); MDBK (Madin-Darby bovine kidney) (Furuse et al., 1994), FRT (Fischer rat thyroid) (Zurzolo et al., 1993), as well as primary cultures of choroid plexus and retinal pigmented epithelium (Marrs et al., 1993). As we have discussed in the first half of this review, we arejust beginning to elucidate the nature of certain “apical” and “basolateral” sorting signals. However, the “nonstandard” sorting of GPI-link proteins in FRT cells mentioned above, and the fact that a number of proteins display tissue and cell-type specific membrane localizations (see Table 3), calls into question the ways in which we think about polarized sorting signals and the mechanisms of sorting. As shown in Table 3, there are notable differences in the localization of certain membrane proteins expressed in different tissue cell-types. The Na,K-ATPase, nearly ubiquitously expressed at the basolateral domain of most polarized cell types, is localized to the lumenal (apical) domain of both retinal pigmented epithelial and chorid plexus cells (Wright, 1972; Steinberg and Miller, 1979; Spector and

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Table 3. Tissue Specificity of Sorting Protein

TissuelCell Type

Apical

intestine

LDL-R

liver proximal kidney

+

intestine, liver, kidney choroid plexus Na,K-ATPase

retinal pigmented epithelium

+ +

Basolateral

GPI-lin ked proteins

+

C

d-e

+

vacuolar H+ATPase References:

kidney: alpha intercalateed cells beta intercallated cells

f

I

+ +

Folicular Eopithelium of Drosophila

+

g-’

Neuron

Gut epithelium of Drosophila

b

+

Fischer Rat Thyroid (FRT) epithelial cells

Dendrite Ref.

+ +

neuron most polarized cell culture models, e.g., MDCK I & 11, LLC-PKI, Caco-2, native hepatocytes

Axon

+

k I I

+

m-n

+ +

m-n

(a) Pathak et al., 1990, (b)Almen and Stirling, 1984; (c) Quinton et al., 1973; (d)Steinbergand Miller, 1979; (el Rizzolo, 1990; (0 Pietrini et al., 1992; (g) Lisanti et al., 1988; (h) Lisanti et al., 1989a,b, (i) Lisanti et al., 1990; (1) Zurzolo et al., 1993; (k)Dotti et al., 1991; (I)Shiel and Caplan, 1995a,b; (m) Schwartz et al., 1985; (n) Brown et al., 1988.

Johanson, 1989; Gundersen et al., 1991). When the cDNA encoding the LDL receptor was placed under the control of a metallothionein promoter and employed in the generation of a transgenic mouse, the receptor was expressed at the basolateral domains of liver and intestinal epithelial cells, but unexpectedly localized to the apical domains of proximal kidney tubule cells (Pathak et al., 1990). The polarized budding of certain viruses and the localization of their respective spike glycoproteins was shown to vary considerably between kidney derived MDCK and thyroidderived FRT cells (Zurzolo et al., 1992a). In some instances, ashift in the type of targeting pathway used by a protein can depend on the differentiated state of the cell culture (Zurzolo et al., 1992b). Furthermore, the polarized localization of a particular GPI-linked protein was found to be developmentally regulated in Drosophila embryos (Shiel and Caplan, 1995a). Finally, a remarkable flexibility and “plastic-

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ity” of protein sorting has been suggested to be present in kidney intercalated cells, which appear to direct the vacuolar proton pump to either surface domain, depending on particular environmental cues (Schwartz et al., 1985; Brown et al., 1988a) At the present time, we have little understanding of the signals or sorting mechanisms that mediate the differential sorting of the same protein in distinct cellular types. Are different signals recognized by the different epithelial cells or is the same signal interpreted differently? Is the sorting machinery itself different between polarized cells, or is the sorting machinery basically conserved between different cell-types while its regulation, adaptation, or wiring to the targeting machinery is different? Evidence discussed in the second half of this review on the rab family of proteins suggests that elements of the targeting machinery are in fact highly conserved between different cell types, and it is the cell-type specific adaptation of this machinery which accounts for differences. Nonetheless, it is becoming clear that the sorting of a particular protein can be a highly idiosyncratic feature of each polarized cell model. The observation that different epithelial cell lines may handle the same protein (or the same signal) differently has to reflect more than a mere capriciousness of epithelial cells in culture. Each of the cultured cell models employed in polarity studies derive from and reflect some of the differentiated features of a tissue or organ system. Accordingly, the sorting behavior observed in a particular cell type needs to be evaluated in the context of this cell’s functional history. For example, is this cell derived from a tissue specialized for apical secretion or apical endocytosis? Studies of the sorting of ion-transporting ATPase molecules expressed in distal tubule-derived MDCK and proximal tubule derived-LLC-PK1 kidney cells suggest that the distinct cell surface distributions which an ATPase subunit achieve in these two lines are consistent with established physiologic differences between the distal and proximal tubule epithelial cells (Roush et al., manuscript submitted). These observations have led to the suggestion that sorting mediates delivery to functionally defined rather than topographically defined domains (Gottardi and Caplan, 1993a).

111. POLARIZED SORTING AND TARGETTING MACHINERY: ELEMENTS OF THE INTRACELLULAR PROTEIN TRANSPORT MACHINERY? It is becoming quite clear that the findings in the field of intracellular protein transport (reviewed by Rothman, 1994 and by Mellman, 1994) will prove to be extremely valuable to the discipline of epithelial polarity. In this field, the convergence of studies on synaptic vesicle (regulated) secretion in neurons, constitutive secretion in yeast, and intra-Golgi transport have led to the rapid identifcation and characterization of the basic components necessary for vesicle formation, docking and fusion. Clearly, the general components of the bud-

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115

ding and docking machinery lie at the heart of any transport process, whether we are considering the transport of a membrane protein from ER to Golgi, or a secretory protein from the TGN to a particular cell surface. In the following sections we touch upon some of the key discoveries in the field of intracellular transport and focus on the relevant molecules that may contribute to polarized sorting and delivery processes. A.

GTPases and Epithelial Polarity

One of the recent paradigms in intracellularprotein transport is based on the concept that vesicle shuttling between different organellar compartments is regulated through the coordinated efforts of different GTP-binding proteins. There are two broad classes of GTP-binding proteins which have been shown to regulate membrane trafficking events; the small G proteins (rabs and ARF) reviewed by (Donaldson and Klausner, 1994; Pfeffer, 1992,1994) and the trimeric G proteins (reviewed by Bomsel and Mostov, 1992). B.

Rabs

The role of a GTP-binding protein in regulating vesicular transport was first realized with the analysis of one of the temperature sensitive SEC (secretory) mutants in yeast (Salminen and Novick, 1987). Sec4 mutants display a rather striking accumulation of secretory vesicles when cultured at the restrictive temperature. The cloning, sequencing, and characterization of the SEC4 gene revealed that it encoded a ras-like or ‘small’ GTP-binding protein which was present on the surfaces of the vesicles and could bind and hydrolyze GTP (Salminen and Novick, 1987; Goud et al., 1988; Kabcenell et al., 1990). Since the phenotype of cells bearing mutant sec4 is the accumulation of transport vesicles, it was apparent that SEC4 is necessary for the targeting and/or fusion of secretory vesicles with the plasma membrane. Similar results were found with another yeast protein YPTl(48% identical to SEC4), which in its mutant form inhibited vesicular transport between the ER and Golgi complex (Gallwitz et al., 1983; Segev et al., 1988). The suggestion that two small GTPases were important in the regulation of two different vesicular transport events in yeast led to the hypothesis that each step in vesicular traffic was regulated by a specific GTPase (Bourne et al., 1990). These ras-like GTPases are known to zdopt either of two distinct conformations, depending upon whether or not they are complexed with GTP or GDP. Consequently, these GTPases have been postulated to serve as key regulators or “molecular switches” for membrane fission and fusion events. The apparent generality of ras-like GTPase in yeast, as revealed by sec4 and yptl, inspired asearch for these proteins’ mammalian counterparts. To date, 30 YPTl/SECCrelated proteins have been identified and are often referred to as rab proteins (“Ras-like” proteins from rat brain) reviewed in (Balch, 1990; Hall, 1990; Goud and McCaffrey, 1991; Zerial and Stenmark, 1994).

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A number of rabs have been localized to specific organelles within the cell and through the combined efforts of in vitro and in vivo approaches have been shown to regulate membrane traffic between these organelles (reviewed in Zerial and Stenmark, 1994). How this class of molecules contributes to the overall fidelity of membrane trafficking events is still unclear (Rothman, 1994). The idea that specific rab proteins regulate distinct steps along the transport pathway (e.g., rabl always regulates ER to Golgi traffic, whether in a kidney cell or neuron) led to the hypothesis that cells which contain unique, cell-type specific transport processes might be regulated by disinct rabs. Indeed, the best example of this is the family of rab3 isofoms which have been found to be localized within cells which are well-adapted for regulated secretory events. Rab3a has been suggested to be important in the regulation of Caz+dependent secretion in neuronal (Fischer von Mollard et al., 1991), neuroendocrine (Darchen et al., 1990) and endocrine cell types (Mizoguchi et al., 1989). Interestingly, an isoform of rab3a, rab3d, has been localized to the glucose transporter-containing vesicles of adipocytes, which are known to undergo regulated exocytosis after insulin stimulation (Baldini et al., 1992). Thus, despite cell-specific differences, or vesicle-content differences, these regulated pathways rely on similar rabs (rab3). Thus, distinct regulated exocytic events in a variety of cell types make use of similar molecular machinery (Lutcke et al., 1993). In this context, it has been speculated that polarized epithelial cells, with their distinct apical and basolateral targeting pathways, may employ epithelia-specific rab molecules. Recent data suggest that this may be true. There are four rabs which have been implicated in polarized epithelial-specific functions: rab 17, rab 3b, rabl3, and rab8. Of the four, only rabl7 is truly specific to polarized epitheiial cells. In the developing kidney, rabl7 mRNA is detected only after mesenchyme is induced to differentiate into polarized epithelial structures (Lutcke et al., 1993). Interestingly, rabl7 induction was shown to occur just prior to the appearance of apical markers and has therefore been suggested to be involved in the generation of apicalhasolateral polarity in these cells. Rab 17 localizes to the basolateral membrane and to electron dense tubules near the apical membrane. Since rab proteins have been shown to regulate transport between the subcellular compartments with which they associate, it has been suggested that rabl7 regulates epithelial transcytosis. As we stated previously, two isoforms of rab3 (3A and 3D) have been implicated in the regulated exocytosis events shared by neuronal, endocrine and adipocyte cell types. Interestingly, another isoform of rab3,3b, has been shown to be specific for polarized epithelial cells and is exclusively localized to the apical pole of cells, near the tight junctions (Weber et al., 1994). Rabl3, like rab3b, also accumulates at the apical poles of polarized cells and co-localizes with the tight junction associated protein, ZO-1 (Zahraoui et al., 1994). It has been suggested that these two rabs could regulate events necessary for the establishment of polarity. For example, since the localization of both rabs are completely dependent on the presence the of cell-cell contacts, it is possible that these mole-

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cules control the recruitment of membrane protein-containing vesicles required for establishing the tight junction “fence,” a structure thought to maintain the distinct protein and lipid compositions of apical and basolateral membranes (Dragsten et al., 1981). It has also been proposed that these rabs control general vesicle targetting to the apical membrane (Zahraoui et al., 1994). This hypothesis was based on two independent observations. It has been shown that an apical membrane protein (aminopeptidase) inserts preferentially into the apical membrane at regions of cell-cell contact in MDCK cells (Louvard, 1980). Furthermore, under conditions in which MDCK cells are denied intercellular contacts, apical proteins appear to be sorted and retained within a large subapical vacuolar compartment (vacuolar apical compartment, or VAC) which, after initiation of cell-cell contact, is inserted preferentially at regions of cell-cell contact (Vega-Salas et al., 1988). Taken together, the localization of r a b l 3 and rab3b at this region of cell contact places these monomeric GTPases in a position to regulate the delivery of apical proteins to the cell surface (Zahraoui et al., 1994). Moreover, the localization of a regulated, exocytic compartmentspecific rab (rab3) to a subdomain of the apical membrane of polarized cells is intriguing and suggests possible functional relationships between these subcellular compartments. The last rab worth exploring in the context of epithelial polarity is rab8. While rab8 is not solely expressed in polarized cells, it is the only rab that has been functionally implicated in vectorial targeting. A peptide derived from the C-terminal region of rab8 can inhibit basolateral but not apical transport of membrane proteins in a permeabilized-MDCK cell assay (Huber et al., 1993a). Interestingly, rab8 can also regulate membrane transport to the dendritic plasma membranes of neurons in culture; antisense rab8 oligonucleotides decrease the level of viral glycoprotein transported to this domain (Huber et al., 1993b). This observation is consistent with the model which suggests that the mechanisms which produce axoddendrite polarity in neurons may be similar to those involved in apicallbasolateral polarity in epithelia (Simons et al., 1992). Taken together, the identification of a polarized epithelia-specific rab (rab 17), and the localization of other rabs to specific polarized epithelial domains (rab 13 and rab3B, apical; rab8, basolateral) suggests that rabs may regulate specific pathways in polarized epithelial cells. For the epithelial cell biologist, the obvious question is, “What brings about the pathway-specific localizations of rab proteins in polarized epithelial cells?” It has been demonstrated that the carboxy-terminal regions of rab proteins are responsible for their unique cellular localizations (Chavrier et al., 1991). It has been suggested that organelle-specific receptors exist which recognize the C-terminal domains of these molecules. At least in terms of polarized cells, it would be tempting to speculate that identification of such receptors for rabl3,3b and rab8 will bring us one step closer to an understanding of the overall machinery that orchestrates domain-specific vesicle formation and targeting.

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Recent evidence, however, suggests that rabs may not provide the primary level of specificity in membrane targeting events (Brennwald and Novick, 1993; reviewed by Rothman and Warren, 1994). As we discuss below, a new class of proteins, the SNARES, may provide the necessary specificity for vesicle-membrane targeting events throughout the cell.

C. The SNARE Paradigm and Epithelial Polarity The SNARE hypothesis for vesicle targeting arose from research in three related fields: synaptic vesicle release in neurons, transport between cisternae of the Golgi, and secretion in yeast. Briefly, a number of synaptic proteins were discovered to be important for the regulated fusion of synaptic vesicles with their targets on the pre-synaptic plasma membranes (reviewed by Pevsner and Scheller, 1994). Homologues of these proteins were found in yeast and shown to be required for constitutive vesicle transport (Aalto et al., 1993). At the same time, key elements of the general machinery for intracellular membrane fusion were being elucidated. In all three .cases, membrane fusion requires an NEM-sensitive factor (NSF), adaptors that link NSF to membrane proteins (SNAPS: Soluble NSF Attachment Proteins) and the membrane receptors for the NSF-SNAP complexes (SNARES: SNAP Receptors) (reviewed in Rothman and Warren, 1994). Distinct SNARE proteins are present in the membranes of the vesicle and the target. The SNARE hypothesis stipulates that each transport vesicle is endowed with its own vesicle- (v-) SNARE (or VAMP-like molecule) that can specifically interact with its cognate target- ( t - )SNARE (or syntaxin/SNAP25-like protein). This ‘pairing’ could ensure vesicleharget membrane specificity, while a general fusion apparatus consisting of NSF and SNAPs could be used throught the cell (Sollner et al., 1993). In the context of epithelial polarity, this hypothesis suggests that vectorial targeting of apical and basolateral proteins will require distinct V-SNARES. Interesting recent data suggest that the situation in at least one epithelial cell type may be somewhat more complicated. When the surface delivery of membrane proteins is examined in MDCK cells permeabilized at their apical or basolateral surfaces with streptolysin 0, it appears that basolateral transport involves all of the machinery discussed above. Toxins which cleave SNAREs inhibit basolateral delivery, as do antibodies directed against SNAPs. In contrast, apical protein insertion is unaffected by these reagents. Isolation of apically-bound vesicles from MDCK cells reveals the presence of high concentrations of an adducin homologue in their surface membranes. Adducins are calcium-dependent phospholipid binding proteins thought to be involved in a number of membrane fusion events (Ilkonen et al., 1995). It would appear, therefore, that completely distinct classes of vesicular targeting and fusion machinery may operate in the two membrane delivery pathways present in polarized epithelial cells.

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Insights from the Membrane Traffic-Perturbing Reagent, BFA

In the absence of a readily available genetic system with which to identify the genes and gene products necessary for such higher eukarotic functions as transcytosis or polarized targeting, epithelial cell biologists have been resigned to the prospect of “poking” at the epithelial cell with various reagents and watching how it responds. Reagents which prevent the polymerization of actin (Gottlieb et al., 1993; Jackmon et al., 1994) and tubulin (Achler et al., 1989; Parczyk et al., 1989), toxins which modify a particular class of G proteins (Stow et al., 1991; Pimplikar and Simons, 1993b), or toxins that inactivate the VAMP, syntaxin and SNAP-25 molecules described above, second messanger stimulators, analogues of the messangers themselves (Apodaca et al., 1994; Cardone et al., 1994; Hansen and Casanova, 1994) and the remarkable fungal metabolite brefeldin A (BFA) are all being incorporated into the repetoire of tools which we hope will enable us to gleen more information from a particular transport pathway. Those interested in polarized and nonpolarized cell functions alike have made use of such cell-perturbing reagents. Since the focus of this review is epithelial polarity, we have chosen to summarize some of the studies which are providing insights about the mechanisms of polarized sorting and targeting.

E.

BFA, Vesicle Bud Formation, and Polarized Trafficking Events

Brefeldin A is a fungal metabolite that endeared itself to cell biologists because of its dramatic effect on the protein secretory pathway (reviewed in Klausner et al., 1992). Protein secretion is inhibited by BFA: membrane trafficking out of the ER is blocked and the Golgi appears to breakdown and become redistributed into the ER (Lippincott-Schwartz et a]., 1989). Before Golgi redistribution, BFA causes this organelle to form tubular extensions which are devoid of any cytoplasmic (nonclathrin) “coat” material (Lippincott-Schwartz et al., 1990). It has been shown that these morphological changes are not restricted to the Golgi but rather are observed in a number of organelles of the endomembranous network such as endosomes, lysosomes and the TGN (Hunziker et al., 1991; Lippincott-Schwartz et al., 1991; Wood et al., 1991), suggesting that the BFA “effector” might play a role in membrane transport events all over the cell. Perhaps surprisingly, while membrane transport phenomena are remarkably altered in the presence of BFA, several processes are clearly unaffected, including receptor mediated endocytosis and endocytic recycling (Lippincott-Schwartz et al., 1991). From the standpoint of sorting and polarized delivery, BFA’s most interesting property is its ability to differentially affect polarized cell surface targeting events. For example, Low and colleagues ( I 991,1992) determined a concentration of BFA where ER-Golgi trafficking was not inhibited, so that delivery from the TGN to the surface could be assayed for BFA sensitivity. Interestingly, BFA inhibited the apical delivery of both endogenous, MDCK secretory proteins (199 1) and the membrane

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protein DPPIV (1992) while also enhancing their mis-delivery to the basolateral surface. Basolateral targeting of the endogenous MDCK protein, uvomorulin, was not affected under these conditions. Taken together, it would seem that a target molecule for BFA action exists that is exclusively involved in directing apical vesicles or which is simply more sensitive to the effects of BFA than similar molecules participating in the basolateral pathway. Either way, these results provide a hint that there are indeed molecular differences between these two targeting pathways. It is important to add that in addition to inhibiting the exocytic apical pathway in MDCK cells, basolateral to apical transcytosis is also inhibited by this drug (Hunziker et al., 1991; Low et al., 1992). These findings have led to the suggestion that sorting mechanisms for apically destined proteins, whether along the exocytic or the transcytotic pathway may be functionally and biochemically similar (Hunziker et al., 1991). The fact that the loss of the structural integrity of the Golgi induced by BFA correlates with a striking absence of its characteristic “coat” (observed at the EM level) led to the idea that coat proteins might be rendered non-functional due to BFA action. Through a number of studies (reviewed by Donaldson et al., 1992; Helms and Rothman, 1992; Klausner et al., 1992; Rothman & Orci, 1992) molecules which make up this “coat” were identified and characterized (e.g., PCOP and A m ) . An “order of events” necessary for vesicle budding emerged from these studies and is outlined below. ARF is a GTP-binding protein loosely related to ras and distinct from the family of rabs. In its GTP-bound state, it is capable of associating with the membrane by virtue of its myristoyl group, while its GDP-bound form is soluble and not membrane bound. ARF binding to membranes appears to be the signal for coatomer binding, that is, the binding of PCOP in addition to other as yet uncharacterized coat proteins. Coatomer binding is believed to be absolutely necessary for vesicle budding. Therefore proper coatomer assembly would be required for any event downstream of budding, such as targeting. Recently, it has been determined that BFA inhibits coatomer assembly and vesicle formation through ARF, by essentially allowing it to remain in its GDP-bound or inactive form. There exists a class of proteins which are able to catalyze the exchange of GDP for GTP called guanine nucleotide exchange factors (GNE). BFA has been proposed to antagonize the action of a GNE on ARF, thus preventing coatomer assembly and membrane budding (Donaldson et al., 1992; Helms and Rothman, 1992). With the recent identification of an ever-growing family of new ARF-related genes (Kahn et al., 1991) and the speculation that different COPs may exist in the control of membrane budding events from different organelles (Matter and Mellman, 1994), there is growing excitement that ARFs and COPs will turn-out to be essential components for regulating a particular level of specificity inherent to membrane targetting events. In the context of BFAs affect on apical sorting and targeting in polarized MDCK cells (Low et al., 1991, 1992), it is likely that distinct ARFkoatomer complexes regulate the budding of apical and basolaterallydestined vesicles from the TGN. Moreover, the fact that significant missorting into

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the basolateral pathway was observed in the presence of BFA (Low et al., 1992) suggests that coatomer assembly may be inextricably linked to proper secretory and membrane protein sorting.

F.

Heterotrimeric G Proteins and Sorting

It has been known for some time that members of the heterotrimeric family of G proteins are associated not only with the plasma membrane but also with intracellular membranes (reviewed by Bomsel and Mostov, 1992). A number of toxins (cholera, pertussis and mastoparan) known to activate or inhibit various classes of G proteins have been applied to studies of polarized sorting and targeting. Stow et al. (1991) found that overexpression of Gai-3 in polarized LLC-PK1 cells significantly reduced the level of constitutive basolateral secretion of an extracellular matrix component, heparan sulfate proteoglycan. Pertussis toxin, which ADP-ribosylates and inactivates the a-subunits of the G a i/o class of heterotrimeric G proteins, relieved this inhibition. Similarly, Pimplikar and Simons (1993) suggested that Gi and Gs may differentially regulate the trafficking of apical and basolateral vesicles in SLO-permeabilized MDCK cells, while Leyte et al. (1992) found that Gi/o and Gs associated with the TGN could oppositely regulate constitutive secretory vesicle formation. It should be noted that in no case did the G protein related inhibition or stimulation appear to affect the actual sorting or missorting of apical or basolaterally destined proteins (in contrast to the BFA results discussed above (Low et al., 1992), but rather may only affect the rate or “efficiency” of sortingkargetting (Pimplikar and Simons, 1993a). A possible link between heterotrimeric G proteins and coatomer formatiodvesicle budding was provided by Ktistakis et al., (1992). This group found that activation of a G a protein with mastoparan promoted PCOP binding and prevented BFAinduced effects. Pretreatment of cells with pertussis toxin, which is known to specifically affect G a i subclass of heterotrimerics, prevented mastoparan’s antagonizing effects on BFA. Stated more simply, these results showed that activation of a pertussis-toxin-sensitive Ga promotes the binding of PCOP to Golgi membranes and thus antagonizes the action of BFA. The authors of this study suggest further that different subclasses or isoforms of G a could be responsible for some of the differences in BFA-sensitivities observed between cell types and organellar membranes. These key observations have led to the idea that heterotrimeric G proteins, by virtue of their membrane topology would be ideal candidates for coordinating the transfer of sorting information to the cytoplasmic surface of the TGN necessary for vesicle budding (Bomsel and Mostov, 1992; Ktistakis et al., 1992). C.

Insights from Genetic Models

The outer surface of a fruit fly embryo is composed of a monolayer of polarized epithelial cells. The apical membranes of these epithelial cells face the outer shell,

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or chorion, while their basolateral surfaces face the embryonic interior and yolk space. Invaginations of this surface epithelium give rise rise to all of the embryo’s internal tissue structures (for review see Shiel and Caplan, 1995b). Recent investigations have examined the mechanisms through which proteins are sorted in these epithelial cells. Human placental alkaline phosphatase (PLAP) is a GPI-linked protein which has been shown to be sorted to the apical plasma membrane when it is expressed by transfection in MDCK cells. A chimeric construct of PLAP, in which the GPIlinkage domain is replaced by the transmembrane and cytoplasmic domains of the VSV G protein (PLAPG), is sorted to the basolateral surfaces of MDCK cells (Brown et al., 1989). These two proteins have been expressed under the control of heat shock promoters in transgenic flies and their distributions have been examined in embryonic epithelia throughout embryogenesis (Shiel and Caplan, 1995a). As would be expected, the PLAPG protein is restricted to basolateral surfaces throughout ontogeny in the surface epithelial cells as well as in the internal epithelia which derive from invaginations of the surface cells. Surprisingly, PLAP was also restricted to a basolateral distribution in the surface epithelial cells in both early and late stage embryos. Biochemical experiments demonstrated that this mis-sorting of the PLAP protein can not be attributed to problems with the addition of the GPI-linkage, since at all embryonic stages PLAP is correctly glipiated. Internal epithelial cells sorted PLAP exclusively to their apical surfaces. Since in many cases internal epithelia form from surface epithelia without undergoing any mitosis (e.g., salivary gland), essentially the same epithelial cell is capable of differentially sorting PLAP depending on that cell’s physical position within the embryo. Examination of epithelia undergoing invagination (e.g., ventral furrow, tracheal placode) demonstrate that the transition in PLAP sorting occurs in the early stages of the invagination process. While the mechanism responsible for this switch remains unclear, the power of Drosophilu genetics will hopefully allow the cellular components responsible for this transition to be readily identified. It is likely that the isolation of the proteins responsible for this phenomenon will shed light on the Drosophilu as well as on the mammalian epithelial sorting machinery. A Drosophilu mutation whose phenotype includes peturbations of the polarized organization of the surface epithelial cells has recently been identified and characterized at the molecular level (Knust et al., 1993; Wodarz et al., 1993). The crumbs gene encodes a transmembrane protein which is normally expressed in the apical membranes of surface and internal epithelial cells. Mutation of the crumbs gene results in a loss of crumbs polarity and markedly alters embryonic morphology. Genetic studies have demonstrated that the crumbs gene product is necessary not only for its own apical sorting, but for the apical delivery of other proteins as well. Furthermore, the product of the stardust gene appears to interact with the crumbs protein and also appears to participate in apical sorting. Understanding these proteins’ biochemical functions and their intermolecular associations will undoubtedly provide enormous insight into the cellular components responsible for generating and

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maintaining the polarized phenotype. Hopefully, the development of genetic approaches such as these, in concert with the continuing refinement of in vitro and model systems, will allow us to develop a clear and fundamental understanding of how epithelial cells produce their remarkable asymmetry.

REFERENCES Aalto, M. K., Ronne, H. & Keranen, S. (1993). Yeast syntaxins Ssolp and Sso2p belong to a family of related membrane proteins that function in vesicular transport. EMBO J. 12,40954104. Achler, C., Filmer, D., Merte, C. & Drenckhahn, D. (1989). Role ofmicrotubules in polarized delivery of apical membrane proteins to the brush border ofthe intestinal epithelium. J. Cell Biol. 109(1), 179-1 89. Almers, W. & Stirling, C. (1984). Distribution of transport proteins over animal cell membranes. J. Membr. Biol. 77, 169-186. Anderson, R. G. W. (1993). Podocytosis of small molecules and ions by caveolae. Trends Biochem. Sci. 3,69-72. Apodaca, G., Enrich, C. & Mostov, K. E. (1994). The calmodulin antagonist, W-13, alters transcytosis, recyclingand morphology ofthe endocyticpathway in Madine-Darby canine kidney cells. J. Biol. Chem. 269(29), 19005-19013. Aroeti, B., Kosen, P. A,, Kuntz, I . D., Cohen, F. E. & Mostov, K. E. (1993). Mutational and secondary structural analysis of the basolateral sorting signal of the polymeric immunoglobulin receptor. J. Cell Biol. 123, 1149-1 160. Balch, W. E. (1990). Trends Biochem. Sci. 15,473477. Balch, W. E., McCaffery, J. M., Plutner, H. & Farquhar, M. G. (1994). Vesicular stomatitis virus glycoprotein is sorted and concentrated during export from the endoplasmic reticulum. Cell 76, 841-852. Baldini, G., Hohl, T., Lin, H. Y. & Lodish, H. F. (1992). Cloning of a rab3 isotype predominately expressed in adipocytes. Proc. Natl. Acad. Sci. USA 89, 5049-5052. Bansal, A. & Gierasch, L. M. (1991). The NPXY internalization signal of the LDL receptor adopts a reverse turn conformation. Cell 67, 1195-1201 Bartles, J., Ferracci, H., Steiger, B. & Hubbard, A. (1987). Biogenesis of the rat hepatocyte plasma membrane in vivo: Comparison of the pathways taken by apical and basolateral proteins using subcellular fractionation. 1. Cell Biol. 105, 1241-1251. Beltzer, J. & Spiess, M. (1991) In vifrobinding oftheasiatoglycoproteinreceptor to the betaadaptin of plasma membrane coated vesicles. EMBO 10, 3735-3742. Berlot, C. &Bourne, H. (1992). Identificationofeffector-activatingresiduesofGsa. Cell 68,9 I 1-922. Blobel, G. (1980). lntracellular protein topogenesis. Proc. Natl. Acad. Sci. USA 77, 149&1500. Bomsel, M. & Mostov, K. E. (1992). Role of heterotrimeric G proteins in membrane traffic. Mol. Biol. Cell 3, 1317-1328. Bourne, H. R., Sanders, D. A. & McCormick, F. (1990). The GTPase superfamily: aconserved switch for diverse cell functions. Nature 348, 125-13 1. Brennwald, P. & Novick, P. J. (1993). Interactions of three domains distinguishing the ras-related GTP-binding proteins Yptl and Sec4. Nature 362, 56C-563. Brewer, C. B. & Roth, M. G. (1991). A single amino acid change in the cytoplasmic domain alters the polarized delivery of influenza virus hemagglutinin. Journal of Cell Biology 1 14(3), 413421. Brown, D. & Rose, J. K. (1992). Sorting of GPI-anchored proteins to glycolipid-enriched membrane subdomains during transport to the cell surface. Cell 68,533-544. Brown, D., Hirsch, S. & Gluck, S. (1988). An H+-ATPase in opposite plasma membrane domains in kidney epithelial cell subpopulations.Nature 33 1, 622-624.

124

CARA J. COTTARDI and MICHAEL J. CAPLAN

Brown, D. A. (1992). Interactions between GPI-anchored proteins and membrane lipids. Trends Cell Biol. 2,338-343. Brown, D. A., Crise, B. & Rose, J. K. (1989). Mechanism of membrane anchoring affects polarized expression oftwo proteins in MDCK cells. Science 245, 1499-1501. Burgess, T. L. & Kelly, R. B. (1987). Constitutive and regulated secretion of proteins. Annu. Rev. Cell Biol. 3,243-293. Caplan, M. J., Anderson, H. C., Palade, G. E. & Jamieson, J. D. (1986). Intracellular sorting and polarized cell surface delivery of NqK-ATPase, an endogenous component of MDCK cell basolateral plasma membranes. Cell 46, 623-63 1. Caplan, M. J., Stow, J., Newman, A,, Madri, J., Anderson, H., Farquhar, M., Palade, G. & Jamieson, J. D. (1987). Dependence on pH of polarized sorting of secreted proteins. Nature 329,632-635. Caplan, M. J. & M a t h , K. S. (1989). Sortingofmembrane and secretoryproteins in polarizedepithelial cells. In Functional Epithelial Cells in Culture, (K. S. Matlin & J. Valentich, Eds.), pp. 71-127. New York: Liss, Alan R. Cardone, M. H., Smith, B. L., Song, W., Mochly-Rosen, D. & Mostov, K. E. (1994). Phorbol myristate acetate-mediated stimulation of transcytosis and apical recycling in MDCK cells. J. Cell Biol. 124(5), 717-727. Casanova, J., Breitfeld, P., Ross, S. & Mostov, K. (1990). Phosphorylation of the polymeric immunoglobulin receptor required for its efficient transcytosis. Science 248, 742-746. Casanova, J. E., Apodaca, G. & Mostov, K. E. (1991). An autonomous signal for basolateral sorting in the cytoplasmic domain of the polymeric immunoglobulin receptor. Cell 66, 65-75. Chavrier, P., Gorvel, J. P., Stelzar, E., Simons, K., Gruenberg, J. & Zerial, M. (1991). Hypervariable C-terminal domain of rab proteins acts as a targeting signal. Nature 353,769-772. Collawn, J. F., Stangel, M., Kuhn, L. A,, Esekogwu, V., Jing, S., Trowbridge, I. S. & Tainer, J. A. (1990). Transferrin receptor internalization sequence YXRF implicates a tight turn as the structural recognition motif for endocytosis. Cell 63, 1061-1072. Collawn, J. F., Kuhn, L. A,, Liu, L.-F. S., Tainer, J. A. & Trowbridge, I. S. (1991). TransplantedLDL and mannose-6-phosphate receptor internalization signals promote high-efficiency endocytosis of the transferrin receptor. EMBO J. 10,3247-3254. Compton,T.,Ivanov, I. E., Gottlieb,T., Rindler,M., Adesnik, M. & Sabatini,D. (1989). Asortingsignal for the basolateral delivery of the vesicular stomatitis virus (VSV) G protein lies in its luminal domain: Analysisofthe targetingofVSV G-influenzahemagglutininchimeras.Proc. Natl. Acad. Sci. USA 86,4112-41 16. Costa de Beauregard, M.-A,, Pringault, E., Robine, S. & Louvard, D. (1995). Suppression of villin expression by antisense RNA impairs brush border assembly in polarized epithelial intestinal cells. EMBO J. 14(3), 409421. Courtois-Coutry, N., Roush, D., Rajendran, V., McCarthy, J. B., Geibel, J., Kashgarian, M. & Caplan, M. J. (1997). A tyrosine-based signal targets H,K-ATPase to a regulated compartment and is required for the cessation of gastric acid secretion. Cell 90, 501-510. Cross, G. A. M. (1990). Glycolipid anchoring of plasma membrane proteins. Annu. Rev. Cell Biol. 6, 1-39. Cunningham, B. & Wells, J. (1989). High-resolution epitope mapping of hGH-receptor interactions by alanine-scanning mutagenesis. Science 244, 1081-1085. Darchen, F., Zahraoui, A,, Hammel, F., Monteils, M.-P., Tavitian, A. & Scherman, D. (1990). Association of the GTP-binding protein rab3A with bovine adrenal chromaffin granules. Proc. Natl. Acad. Sci. USA 87(5692-5696). Dargement, C., Le Bivic, A., Rothenberger, S., lacopetta, 8.& Kuehn, L. C. (1993). The internalization signal and the phosphorylation site of transferrin receptor are distinct from the main basolateral sorting information. EMBO J. 12, 1713-1721. Donaldson, J. G. & Klausner, R. D. (1994). ARF: a key regulatory switch in membrane traffic and organelle structure. Cum. Op. Cell Biol. 6, 527-532.

Epithelial Polarity

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Donaldson, J. G., Finazzi, D. & Klausner, R. D. (1992). Brefeldin A inhibits Golgi membrane-catalysed exchange of guanine nucleotide onto ARF protein. Nature 360,35&353. Dotti, C. G., Parton, R. G. & Simons, K. (1991). Polarized sorting ofglypiated proteins in hippocampal neurons. Nature 349, 158-1 61. Doucet, A. & Barlet-Bas, C. (1989). Involvement of Na,K-ATPase in antinatriuretic action of mineralocorticoids in mammalian kidney. Curr. Top. Membr. Trans. 34, 185-208. Dragsten, P. R., Blumenthal, R. & Handler, J. S. (1981). Membrane assymetry in epithelia: Is the tight junction a barrier to diffusion in the plasma membrane? Nature 294,718-722. Fischer von Mollard, G., Sudhof, T. & Jahn, R. (1991). A small GTP-binding protein dissociates from synaptic vesicles during exocytosis. Nature 349, 79-81, Fuller, S., Bravo, R. & Simons, K. (1985). An enzymatic assay reveals that proteins destined for the apical or basolateral domains of an epithelial cell line share the same late Golgi compartments. EMBO J. 4,297-307. Furuse, M., Itoh, M., Hirase, T., Nagafuchi, A., Yonemura, S., Tsukita, S. & Tsukita, S. (1994). Direct association of occludin with ZO-1 and its possible involvement in the localization of occludin at tightjunctions. J. Cell Biol. 127(6), 1617-1626. Gallwitz, D., Donrath, C. & Sander, C. (1983). A yeast gene encoding a protein homologous to the human c-hashas proto-oncogene product. Nature 306,704-709. Glickman, J., Conibear, E. & Pearse, M. (1989). Specificity of binding of clathrin adaptors to signals on the mannose-6-phosphate/insulin-likegrowth factor 11 receptor. EMBO J. 8, 1041-1047. Goldstein,J. L., Brown, M. S.,Anderson, R. G. W., Russell, D. W. & Schneider, W. J. (1985).Receptor-mediated endocytosis: concepts emerging h m the LDL receptor system. h.Rev. Cell Biol. I , 1-39. Gonzalez, A,, Rizzolo, L., Rindler, M., Adesnik, M., Sabatini, D. D. & Gotlieb,T. (1987). Nonpolarized secretion of truncated forms of the influenza hemagglutinin and the vesicular stomatitis virus G protein from MDCK cells. Proc. Natl. Acad. Sci. USA 84, 3738-3742. Gottardi, C. J. & Caplan, M. J. (1993). An ion transporting ATPase encodes multiple localization signals. J. Cell Biol. 121(2), 283-293. Gottardi, C. J., Dunbar, L. A. & Caplan, M. J. (1995). Biotinylation and assessment of membrane polarity: caveats and methodological conerns. Am. J. Physiol. 268 (Renal Fluid Electrolyte Physiol. 37), F285-F295. Gottardi, C. J., Pietrini, G., Roush, D. L. & Caplan, M. J. (1993). Sorting of ion transport proteins in polarized cells. J. Cell Science Suppl. 17, 13-20. Gottlieb, T., Beaudry, G., Rizzolo, L., Colman, A,, Rindler, M., Adesnik, M. & Sabatini, D. (1986a). Secretion of endogenous and exogenous proteins from polarized MDCK monolayers. Proc. Natl. Acad. Sci. USA 83,2100-2104. Gottlieb, T., Gonzalez, A., Rizzolo, L., Rindler, M. J., Adesnik, M. & Sabatini, D. D. (1986b). Sorting and endocytosis of viral glycoproteins in transfected polarized epithelial cells. J. Cell Biol. 102, 1242-1255. Gottlieb, T., Ivanov, 1. E., Adesnik, M. & Sabatini, D. D. (1993). Actin microfilaments play a critical role in endocytosis at the apical but not the basolateral surface ofpolarized epithelial cells. J. Cell Biol. 1993(3), 695-710. Goud, B. & McCaffrey, M. (1991). Small GTP-binding proteins and their role in transport. Curr. Op. Cell Biol. 3,626-633. Goud, B., Salimen, A,, Walworth, N. C. & Novick, P. J. (1988). A GTP-binding protein required for secretion rapidly associates with secretory vesicles and the plasma membrane in yeast. Cell 53, 753-7 68. Gstrauthaler, G., Pfaller, W. & Kotanko, P. (1985). Biochemical characterization ofrenal epithelial cell cultures (LLC-PK1 and MDCK). Am. J. Physiol. 248, F536F544. Gundersen, D., Orlowski, J. & Rodriguez-Boulan, E. (1991). Apical polarity ofNa,K-ATPase in retinal pigment epithelium is linked to a reversal of the ankyrin-fodrin submembrane cytoskeleton. J. Cell Biol. 112(5), 863-872.

126

C A M J. GOTTARDI and MICHAEL J. CAPLAN

Hall, A. (1990). Science 249,635-640. Hammerton, R. W., Krzeminski, K. A,, Mays, R. W., Ryan, T. A,, Wollner, D. A. & Nelson, W. J. (1991). Mechanism for regulatingcell surfacedistributionofNa,K-ATPaseinpolarizedepithelial cells. Science 254, 847-850. Hansen, S. H. &Casanova, J. E. (1994). Gs alphastimulates transcytosis and apical secretion in MDCK cells through CAMPand protein kinase A. J. Cell Biol. 126(3), 677487. Helms, J. B. & Rothman, J. E. (1992). Inhibition by Brefeldin A of a Golgi membrane enzyme that catalyzes exchange of guanine nucleotide bound onto ARF protein. Nature 360,352-354. Hooper, N. M. (1992). More than just a membrane anchor. Curr. Biol. 2( 1 I), 617-619. Huber, L. A,, Pimplikar, S., Parton, R. G., Virta, H., Zerial, M. & Simons, K. (1993a). Rab 8, a small GTPase involved in vesicular traffic between the TGN and the basolateral plasma membrane. J. Cell Biol. 123(1), 3 5 4 5 . Huber, L. A,, de Hoop, M. J., Dupree, P., Zerial, M., Simons, K. & Dotti, C. (1993b). Protein transport to the dentritic plasma membrane of cultured neurons is regulated by rab8p. J. Cell Biol. 123(1), 47-55. Hull, R. N., Cherry, W. R. & Weaver, G. W. (1976). The origin and characteristics of a pig cell strain, LLC-PK 1. In Vitro 12,67&677. Hunziker, W., Harter, C., Matter, K. & Mellman, I. (1991). Basolateral sorting in MDCK cells requires a distinct cytoplasmic domain determinant. Cell 66,907-920. Ilkonen, E., Tagaya, M., Ullrich, O., Montecucco, C. & Simons, K. (1995). Different requirements for NSF, SNAP, andrab proteins in apical and basolateral transport inMDCK cells. Cell 81,571-580. Jackmon, M. R., Shurety, W., Ellis, J. A. & Luzio, J. P. (1994). Inhibition of apical but not basolateral endocytosisofricinandfolateinCaco-2 cells by cytochalasinD. J. Cell Sci. 107(9),2547-2556. Kabcenell, A., Goud, B., Northup, J. K. & Novick, P. J. (1990). Binding and hydrolysis of guanine nucleotides by sec4p, ayeast protein involved in the regulation of vesiculartraffic. J. Biol. Chem. 265,9366-9372. Kahn, R. A,, Kern, F. G., Clark, J., Gelman, E. P. & Ralka, C. (1991). Human ADP-ribosylationfactors: A functionally conserved family of GTP-binding proteins. J. Biol. Chem. 266(4), 26062614, Klausner, R. D., Donaldson, J. G. & Lippincott-Schwartz, J. (1992). Brefeldin A: insights into the control of membrane traffic and organelle structure. J. Cell Biol. 116(5), 1071-1080. Knust, E., Tepass, U. & Wodarz, A. (1993). crumbs and stardust, two genes of Drosophila required for the development of epithelial cell polarity. Development Suppl., 261-268. Kondor-Koch, C., Bravo, R., Fuller, S., Cutler, D. & Garoff, H. (1985). Exocytotic pathways exist to both the apical and the basolateral cell surface of the polarized epithelial cell MDCK. Cell 43, 297-306. Kornfeld, S. & Mellman, I . (1989). The biogenesis of lysosomes. Annu. Rev. Cell Biol. 5,483-525. Ktistakis, N. T., Linder, M. E. & Roth, M. G. (1992). Action of brefeldin A blocked by activation of a pertussis-toxin-sensitive G protein. Nature 356, 344-346. Kurzchalia, T. V., Wiedmann, M., Girshovich, A. S., Bochkareva, E. S., Bielka, H. & Rapoport, T. A. (1986). The signal sequence of nascent preprolactin interacts with the 54K polypeptide of the signal recognition particle. Nature 320, 634-636. Larkin, J., Sztul, E. S. & Palade, G. E. (1986). Phosphorylationofthe rat hepaticpolymericIgA receptor. Proc. Natl. Acad. Sci. USA 83,47954763. LeBivic, A., Sambuy, Y , Patzak, A,, Patil, N., Chao, M. & Rodriguez-Boulan, E. (1991). An internal deletion in the cytoplasmic tail reverses the apical localization of human NGF receptor in transfected MDCK cells. J. Cell Biol. 115, 607-618. Leyte, A., Barr, F. A,, Kehlenbach, R. H. & Huttner, W. B. (1992). Multiple trimeric G-proteins on the trans-Golgi network exert stimulatory and inhibitory effects on secretory vesicle formation, EMBO J. I1(13), 47954804. Lin, S., Naim, H. Y. & Roth, M. G. (1997). Tyrosine-dependent basolateral sortingsignals are distinct from tyrosine-dependent internalization signals. J . Biol. Chem. 272, 26300-26305.

Epithelial Polarity

127

Lingappa, V. R., Katz, F. N., Lodish, H. F. & Blobel, G. (1978). A signal sequence for the insertion of a transmembrane glycoprotein. J. Biol. Chem. 253, 8667-8670. Lippincott-Schwartz, J., Yuan, L., Bonifacino, J. & Klausner, R. D. (1989). Rapid redistribution of Golgi proteins into the ER in cells treated with brefeldin A: evidence for membrane cycling from Golgi to ER. Cell 56, 801-813. Lippincott-Schwartz, J., Donaldson, J. G., Schweizer, A,, Berger, E. G., Hauri, H. P., Yuan, L. C. & Klausner, R. D. (1990). Microtubule-dependentretrogradetransport of proteins into the ER in the presence of brefeldin A suggests an ER recycling pathway. Cell 60, 821-836. Lippincott-Schwartz, J., Yuan, L., Tipper, C., Amherdt, M., Orci, L. & Klausner, R. D. (1991). Brefeldin A’s effects on endosomes, lysosomes, and the TGN suggest a general mechanism for regulating organelle structure and membrane traffic. Cell 67,601-616. Lisanti, M., Sargiacomo, M., Graeve, L. & Rodriguez-Boulan, E. (1988). Polarized apical distribution of glycosyl-phoshatidylinositol-anchoredproteinsin a renal epithelial cell line. Proc. Natl. Acad. Sci. USA 85,9557-9561. Lisanti, M., Caras, I., Davitz, M. & Rodriguez-Boulan, E. (1989a). A glycophospholipid membrane anchor acts as an apical targeting signal in polarized epithelial cells. J. Cell Biol. 109, 2145-2 156. Lisanti, M., Le Bivic, A,, Sargiacomo, M. & Rodriguez-Boulan, E. (1989b). Steady-state distribution and biogenesis of endogenous MDCK-glycoproteins: Evidence for intracellular sorting and polarized cell surface delivery. J. Cell Biol. 109,2117-2127. Lisanti, M., Le Bivic, A,, Saltiel, A. R. & Rodriguez-Boulan, E. (1990). Preferred apical distribution of glycosyl-phosphatidylinositol (GPI) anchored proteins: a highly conserved feature of the polarized epithelial cell phenotype. J. Membr. Biol. 113, 155-167. Louvard, D. (1980). Apical membrane aminopeptidase appears at site of cell-cell contact in cultured kidney epithelial cells. Proc. Natl. Acad. Sci. USA 77(7), 41324136. Low, S. H., Tang, B. L., Wong, S. H. & Hong, W. (1992). Selective inhibition ofprotein targeting to the apical domain of MDCK cells by Brefeldin A. J. Cell Biol. 118(1), 51-62. Low, S H., Wong, S. H., Tang, B. L., Tan, P., Subramaniam, V. N. & Hong, W. (1991). Inhibition by BrefeldinA ofprotein secretion from the apical cell surfaceofMadin-Darby CanineKidney cells. J. Biol. Chem. 266(27), 17729-17732. Lutcke, A., Jansson, S., Parton,R. G., Chavrier, J., Valencia, A., Huber, L. A., Lehtonen, E. &Zerial, M. (1993). Rabl7, a novel small GTPase, is specific for epithelial cells and is induced during cell polarization. J. Cell Biol. 121(3), 553-564. Machamer, C. E. (1993). Targeting and retentioon ofGolgi membrane proteins. Curr Op Cell Biol5(4), 606412. Madara, J. L., Stafford, J., Dharmsathaphorn, K. & Carlson, S. (1987). Structural analysis o f a human intestinal epithelial cell line. Gastroenterology 92, 1 133-1 145. Marrs, J. A., Napolitano, E. W., Murphy-Erdosh, C., Mays, R. W., Reichardt, L. F. & Nelson, W. J. (1993). Distinguishing roles of the membrane-cytoskeleton and cadherin mediated cell-cell adhesion in generating different Na,K-ATPase distributions in polarized epithelia. 1. Cell Biol. 123, 149-164. M a t h , K. S. & Simons, K. (1984). Sortingofan apical plasmamembraneglycoproteinoccurs before it reaches the cell surface in cultured epithelial cells. J. Cell Biol. 99,2131-2139. Matter, K., Brauchbar, M., Bucher, K. & Hauri, H. P. (1990). Sortingofendogenous plasmamembrane proteins occurs from two sites in cultured human intestinal epithelial cells (Caco-2). Cell 60, 429437. Matter, K., Hunziker, W. & Mellman, 1. (1992). Basolateral sorting of LDL receptor in MDCK cells: The cytoplasmic domain contains two tyrosine-dependent targeting determinants. Cell 71, 741-753. Matter, K. & Mellman, I. (1994). Mechanisms ofcell polarity: sorting and transport in epithelial cells. Curr. Op. Cell Biol. 6, 545-554.

128

CARA J. COTTARDI and MICHAEL J. CAPLAN

Matter, K., Whitney, J. A., Yamamoto, E. M. & Mellman, 1. (1993). Common signals control low density lipoprotein receptor sorting in endosomes and the Golgi complex of MDCK cells. Cell 74, 1053-1064. Matter, K., Yamamoto, E. M. & Mellman, I. (1994). Structural requirements and sequence motifs for polarized sorting and endocytosis of LDL and Fc receptors in MDCK cells. J. Cell Biol. 126, 991-1 004. Mays,R. W.,Beck,K.A. &Nelson, W. J. (1994).Organizationandfunctionofthecytoskeletoninpolarized epithelial cells: a component of the protein sorting machinery. Curr. Op. Cell Biol. 6, 1624. McQueen, N., Nayak, D., Stephens, E. B. & Compans, R. W. (1986). Polarizedexpressionofa chimeric protein in which the transmembrane and cytoplasmic doamins of influenza hemagglutinin have beenreplaced by those of the vesicular stomatitis G protein. Proc. Natl. Acad. Sci. USA 83, 93 18-9322. Mellman, I. (1994). Membranes and sorting. C u r . Op. Cell Biol. 6,497498. Mikogami, T., Heyman, M., Spik, G. & Desjeux, J. F. (1994). Apical-to-basal transepithelial transport of human lactoferrin in the intestinal cell line HT-29c1.19A. Am. J. Physiol. 267, G308-G315. Misek, D. E., Bard, E. & Rodriguez-Boulan, E. J. (1984). Biogenesis of epithelial cell polarity: Intracellular sorting and vectorial exocytosis of an apical plasmamembrane glycoprotein.Cell 39, 537-546. Mizoguchi, A., Kim, S., Ueda, T. & Takai, Y. (1989). Tissue distribution of smg p25A, a ras p21-like GTP-binding protein, studied by use of a specific monoclonal antibody. Biochem. Biophys. Res. Commun. 162, 1438-1445. Mostov, K. E. & Deitcher, D. L. (1986). Polymeric immunoglobulin receptor expressed in MDCK cells transcytoses IgA. Cell 46,613-621. Mostov, K. & Simister, N. E. (1985). Transcytosis. Cell 43,389-390. Mostov, K., Apodaca, G. & Aroeti, B. (1992). Plasma membrane protein sorting in polarized epithelial cells. J. Cell Biol. 116, 577-583. Munro, S. & Pelham, H. (1987). A C-terminal signal prevents secretion of luminal ER proteins. Cell 48, 899-907. Nelson, W. J. (1992). Regulation of cell surface polarity from bacteria to mammals. Science 258, 948-955, Nilsson, T., Jackson, M. & Peterson, P. A. (1989). Short cytoplasmic sequences serve as retention signals for transmembrane proteins in the endoplasmic reticulum. Cell 58, 707-718. Orci, L., Ravazzola, M., Amherdt, M., Perrelet, A., Powell, S. K., Quinn, D. L. & Moore, H.-P. H. (1987). The trans-most cisternae of the Golgi complex: A compartment for sorting of secretory and plasma membrane proteins. Cell 51, 1039-1 05 1. Parczyk, K., Haase, W. & Kondor-Koch, C. (1989). Microtubules are involved in the secretion of proteins at the apical cell surface of the polarized epithelial cell, Madin-Darby canine kidney. J. Biol. Chem. 264(28), 16837-16846. Pathak, R. K., Yokode, M., Hammer, R. E., Hofmann, S. L., Brown, M. S.,Goldstein, J. L. &Anderson, R. G. W. (1990). Tissue-specific sorting of the human LDL receptor in polarized epithelia of transgenic mice. Journal of Cell Biology 11 1,347-359. Pearse, B. & Robinson, M. S. (1990). Clathrin, adaptors, andsorting. Ann Rev. Cell Biol. 6,15 1-17 1. Pearse, B. M. F. (1988). Receptors compete for adaptors found in plasmamembranecoated pits. EMBO J. 7,333 1-3336. Pevsner, J. & Scheller, R. H. (1994). Mechanisms of vesicle docking and fusion: insights from the nervous system. Curr. Op. Cell Biol. 6, 555-560. Pfeffer, S. R. (1992). GTP-bindingproteins inintracellulartransport.Trendsincell Biology2,4146. Pfeffer, S. R. (1994). Rab GTPases: master regulators of membrane traficking. Curr. Op. Cell Biol. 6 , 522-526. Pfeffer, S., Fuller, S. & Simons, K. (1985). Intracellular sorting and basolateral appearance of the G protein of vesicular stomatitis virus in MDCK cells. J. Cell Biol. 101, 4 7 M 7 6 .

Epithelial Polarity

129

Pietrini,G., Matteoli, M., Banker,G. &Caplan,M. (1992). IsoformsoftheNa,K-ATPase are presentin both axons and dendrites of hippocampal neurons in culture. Proc. Natl. Acad. Sci. USA 89, 84 14-84 18. Pimplikar, S. & Simons, K. (1993a). Regulation of apical transport in epithelial cells by a Gs class of heterotrimeric G protein. Nature 362,456-458. Pimplikar, S. W. & Simons, K. (1993b). Role of heterotrimeric G proteins in polarized membrane transport. J. Cell Science Suppl. 17, 27-32. Pinto, C., Robine-Leon, S., Appay, M.-D., Kedinger, M., Triadou, N., Dassaulx, E., Lacroix, B., Simon-Assmann, D., Haffen, K., Fogh, J. & Zweibaum, A. (1983). Biol. Cell. 47,323-330. Polak-Charcon, S., Hekmati, U. & Ben-Shaul, Y. (1989). The effect ofmodifying the culture medium on cell polarity in a human colon cell line. Cell Differentiation and Development 26(2), 119-129. Puddington, L., Woodgett, C. & Rose, J. K. (1987). Replacement of the cytoplasmic domain alters sorting of a viral glycoprotein in polarized cells. Proc. Natl. Acad. Sci. USA 84,275&2760. Quinton, P. M., Wright, E. M. & Tormey, J. M. (1973). Localization of sodium pumps in the choroid plexus epithelium. J. Cell Biol. 58, 724-730. Richardson, W. D., Roberts, B. L. & Smith, A. E. (1986).Nuclear localization signals in polyomavirus Large-T. Cell 44, 77-85. Rindler, M. J., Ivanov, 1. E., Plesken, H., Rodriguez-Boulan, E. J. & Sabatini, D. D. (1984). Viral glycoproteins destined for apical or basolateral plasma membrane domains traverse the same Golgi apparatus during their intracellular transport in doubly infected Madin-Darby Canine Kidney cells. J. Cell Biol. 98, 1304-1319. Rizzolo, L. J. (1990). The distribution of Na,K-ATPase in the retinal pigmented epithelium from chicken embryo is polarized in vivo but not in primary cell culture. Exp. Eye Res. 51, 435-446. Robinson, M. S. (1994). The roleofclathrin, adaptors and dynamin inendocytosis. Curr.Op. Cell Biol. 6,538-544. Rodriguez-Boulan, E. & Nelson, W. J. (1989). Morphogenesis of the polarized epithelial cell phenotype. Science 245,71&725. Rodriguez-Boulan, E. & Powell, S. K . (1992). Polarity of epithelial and neuronal cells. Ann. Rev. Cell Biol. 8, 395-427. Rodriguez-Boulan, E. & Sabatini, D. (1 978). Asymmetric budding of viruses in epithelial monolayers: A model system for study of epithelial polarity. Proc. Natl. Acad. Sci. USA 75, 5071-5075. Roth, M., Compans, R., Giusti, L., Davis, A., Nayak, D., Gething, M. J. & Sambrook, J. S. (1983). Influenza virus hemagglutinin expression is polarized in cells infected with recombinant SV40 viruses carrying cloned hemagglutinin DNA. Cell 33,435443. Roth, M. G., Gunderson, D., Patil,N. & Rodriguez-Boulan, E. (l987a). The large extracellular domain is sufficient for the correct sorting of secreted or chimeric influenza virus hemagglutinins in polarized monkey kidney cells. J. Cell Biol. 104,769-782. Roth, M. G., Gunderson, D., Patil, N. & Rodriguez-Boulan, E. J. (1987b).The large external domain is sufficient for the correct sorting of secreted or chimeric influenza virus hemagglutinins in polarized monkey kidney cells. J . Cell Biol. 104, 769-782. Rothman, J. E. (1994). Mechanisms of intracellular protein transport. Nature 3 7 2 , 5 5 4 3 . Rothman, J. E. &Orci, L. (1992). Molecular dissectionofthesecretory pathway. Nature355,409-415. Rothman, J. E. & Warren, G. (1994). Implications ofthe SNARE hypothesis for intracellularmembrane topology and dynamics. Curr. Biol. 4(3), 22G233. Salminen, A. & Novick, P. (1987). A ras-like protein is required for a postGolgi event in yeast secretion. Cell 47, 527-538. Saucan, L. & Palade, G. E. (1994). Membrane and secretory proteins are transported from the Golgi complex to the sinusoidal plasmalemmaofhepatocytes by distinct vesicular carriers. J. Cell Biol. 125(4), 733-741.

130

CARA J. COTTARDI and MICHAEL J. CAPLAN

Schwartz, G., Barasch, J . & Al-Awqati, Q. (1985). Plasticity of functional epithelial polarity. Nature 318,368-371. Segev, N., Mulholland, J. & Botstein, D. (1988). The yeast GTP-binding YPTl protein and a mammalian counterpart are associated with the secretion machinery. Cell 52,915-924. Shiel, M. J. & Caplan, M. J. (1995a). Developmental regulation of membrane protein sorting in Drosophila embryos. Am. J. Physiol. 269(Cell Physiol. 38), C207<216. Shiel, M. J. & Caplan, M. J. (1995b). The generation of epithelial polarity in mammalian and Drosophila embryos. Sem. in Dev. Biol. 6: 39-46. Siemers, K., Wilson, R., Mays, R. W., Ryan, T. A,, Wollner, D. A. &Nelson, W. .I. (1993). Delivery of the Na,K-ATPase in polarized epithelial cells. Science 260, 554-556. Simons, K. & van Meer, G. (1988). Lipid sorting in epithelia. Biochemistry 27,6197-6202. Simons, K. & Wandinger-Ness, A. (1990). Polarized sorting in epithelia. Cell 62,207-210. Simons, K., Dupree, P., Fiedler, K., Huber, L. A., Kobayashi, T., Kurchalia, T., Olkkonen, V., Pimplikar, S., Parton, R. & Dotti, C. (1992). Biogenesis of cell-surface polarity in epithelial cells and neurons. Cold Spring Harbor Symposia on Quantitative Biology 57,611419. Sly, W. S. & Fischer, H. D. (1982). The phosphomannosyl recognition system for intracellular transport of lysosomal enzymes. J. Cell. Biochem. 18(1), 67-85. Sollner, T., Whiteheart, S. W., Brunner, M., Erdjument-Bromage, H., Geromanos, S., Tempsf P. & Rothman, J. E. (1993). SNAP receptors implicated in vesicle targeting and fusion. Nature 362,3 1%323. Sorkin, A. & Carpenter, G. (1993). Interaction of activated EGF receptor with coated pit adaptins. Science 261 (6 1 2 415). Sosa, M. A,, Schmit, B., von Figura, K. & Hille-Rehfeld, A. (1993). In vitro binding of plasma-coated vesicle adaptors to the cytoplasmic domain of lysosomal acid phosphatase. J. Biol. Chem. 268(17), 12537-12543. Spector, R. & Johanson, C. E. (1989). The mammalian choroid plexus. Sci. Am. 131,68-74. Steinberg, R. M. & Miller, S. S. (1979). Transport and membrane properties of the retinal pigment epithelium. In The Retinal Pigment Epithelium K. M. Z. a. M. F. Marmor, Eds., pp. 205-225. Harvard University Press, Cambridge, MA. Stephens, E. B. & Compans, R. W. (1986). Nonpolarized expression of a secreted murine leukemia virus glycoprotein in polarized epithelial cells. Cell 47, 1053-1059. Stow, J., de Almeida, J. B., Narula, N., Holtzman, E., Ercolani, L. & Ausiello, D. (1991). A heterotrimeric G protein, Gai-3, on golgi membranes regulates the secretion o f a heparan sulfate proteoglycan in LLC-PKI epithelial cells. J. Cell Biol. 114, 1113-1 124. Swift, A. M. & Machamer, C. E. (1991). A Golgi retention signal in a membrane-spanning domain of coronavirus El protein. J. Cell Biol. 1lS(l), 19-30. Tang, B. L., Wong, S. H., Qi, X. L., Low, S. H. & Hong, W. (1993). Molecularcloning,characterization, subcellular localization and dynamics of p23, the mammalian KDEL receptor. J. Cell Biol. I20(2), 325-328. Thomas, D. C., Brewer, C. B. & Roth, M. G. (1993). Vesicular stomatitis virus glycoprotein contains a dominant cytoplasmic basolateral sorting signal critically dependent on tyrosine. J. Biol. Chem. 268,33 13-3320. Thomas, D. C. & Roth, M. G. (1994). The basolateral targetingsignal inthe cytoplasmicdomainofVSV G protein resembles a variety of intracellular targeting motifs related by primary sequence but having diverse targeting activities. J. Biol. Chem. 269, 15732-15739. Thompson, T. E. & Tillack, T. W. (1985). Organization of glycosphingolipids in bilayers and plasmamembranes of mammalian cells. Annu. Rev. Biophys. Chem. 14,361-386. Tooze, J., Tooze, S. A. & Fuller, S. (1987). Sorting of progeny coronavirus from condensed secretory proteins at the exit from the trans Golgi network of AtT 20 cells. J. Cell Biol. 105, 1215-1226. Trowbridge, I. S. (1991). Endocytosis and signals for internalization. Curr. Op. Cell Biol. 3(4), 634-641.

Epithelial Polarity

131

Vega-Salas, D. E., Salas, P. J. I. & Rodriguez-Boulan, E. (1988). Exocytosis of vacuolar apical compartment (VAC): A c e l k e l l contact controlled mechanism for the establishmentofthe apical plasma membrane domain in epithelial cells. J . Cell Biol. 107, 1717-1728. von Figura, K. & Hasilik, A. (1986). Lysosomal enzymes and their receptors. Ann Rev Biochem 55, 167-1 93. vonHeijne, G. (1984). AnalysisofthedistributionofchargedresiduesintheN-terminal regionofsignal sequences: Implications for protein export in prokaryotic and eukaryotic cells. EMBO J. 3, 23 15-23 18. Walter, P. & Lingappa, V. R. (1986). Mechanisms of protein translocation across the endoplasmic reticulum. Annu. Rev. Cell Biol. 2,499-516. Wandinger-Ness, A,, Bebbet, M. K., Antony, C. & Simons, K. (1990). Distinct transport vesicles mediate the delivery of plasmamembrane proteins to the apical and basolateral domains of MDCK cells. J. Cell Biol. 11 1,987-1000. Weber, E., Berta, G., Tousson,A., St. John, P., Green, M. W., Gopalokrishnan, U., Jilling, T., Sorscher, E. J., Elton, T. S., Abrahamson, D. R. & Kirk, K. (1994). Expression and polarized targeting of a rab3 isoform in epithelial cells. J. Cell Biol. 125(3), 583-594. Wodarz, A,, Grawe, F. & Knust, E. (1993). CRUMBS is involved in the control of apical protein targeting during Drosophrlu epithelial development, Mechanisms of Development 44(2-3), 175-187. Wood, S. A,, Park, J. E. &Brown, W. J. (1991). Brefeldin A causes amicrotubule-mediatedfusionofthe trans-Golgi network and early endosomes. Cell 67,591400. Wright,E. M. (1972). Mechanismsofiontransportacrossthe choroidplexus. J. Physiol. 226,545-571. Zahraoui, A,, Joberty, G., Arpin, M., Fontaine, J. J., Hellio, R., Tavitian, A. & Louvard, D. (1994). A small rab GTPase is distributed in cytoplasmic vesicles in non-polarized cells but colocalizes with the tightjunction marker 20-1 in polarized epithelial cells. J. Cell Biol. 124, 101-1 15. Zerial, M. & Stenmark, H. (1994). Rab GTPases invesiculartransport. Curr. Op. Cell Biol. 6,613420. Zurzolo, C., Polistina, C., Saini, M., Gentile, R., Aloj, L., Migliaccio, G., Bonatti, S. & Nitsch, L. (1992a). Opposite polarity of virus budding and of viral envelope glycoprotein distribution in epithelial cells derived from different tissues. J. Cell Biol. 117(3), 551-564. Zurzolo, C., LeBivic, A,, Quaroni, A,, Nitsch, L. & Rodriguez-Boulan, E. (1992b). Modulation of transcytotic and direct targeting pathways in a polarized thyroid cell line. EMBO J. 11(6), 2337-2344. Zurzolo, C., Lisanti, M. P., Caras, I. W., Nitsch, L. & Rodriguez-Boulan, E. (1993). Glycosylphosphatidylinositol-anchoredproteins are preferentially targeted to the basolateral surface in Fischer rat thyroid epithelial cells. J. Cell Biol. 121(5), 1031-1039.

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S NEURONAL CELLS

Sharon K. Powell and Rodolfo J. Rivas

.............. A. General Axonal and Dendritic Differences. . . . . . . . . . . . . . . ,134 B. Axonal and Dendritic Cytoskeletons: . . . . . . . . 135 Different Directions Using Different MAPS. C. Differential Protein Localization at the Axonal and Dendritic Domains. . 136 D. Epithelial Cell and Neuronal Polarity: Are They Really the Same Ill. Mechanisms for the Generation of Neuronal Polarity: W A. Intracellular Targeting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. RNA Localization: A Dendrite-Specific Mechanism?. C. Selective Retention . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Intrinsic Factors in Neuronal Polarity: What the Cells Kn V. Conclusions/Future Directions . . . . . . . . . . . . . . . . . . .

Advances in Molecular and Cell Biology Volume 26, pages 133-156. Copyright 0 1998 by JAI Press Inc. All right of reproduction in any form reserved. ISBN: 0-7623-0381-6

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1.

INTRODUCTION

Ramon y Cajal first elucidated the Principle of Dynamic Polarization, which states that the cytoarchitecture of neuronal cells is specialized for communication, with information flowing from the receiving sites of the neuron (usually the dendrites and cell body) to the axon, where the action potential is initiated and propagated undirectionally to pre-synaptic release sites in the axon terminal (Kandel et al., 1991). Although this principle may be too simplistic to explain many aspects of complex synaptic interactions (for discussion see Craig and Banker, 1994), it is particularly useful for directing thinking about the molecular basis of neuronal polarity. Reflecting the specialized functional polarity of the neuron are distinct differences between axons and dendrites in morphology, cytoskeletal composition, and distribution of organelles and membrane-associated proteins. Morphological and biochemical differences between axons and dendrites are well documented and have been extensively summarized in recent reviews (Black and Baas, 1989; Sargent, 1989; Craig et al., 1992; Rodriguez-Boulan and Powell, 1992; Kelly and Grote, 1993; Craig and Banker, 1994). Molecular mechanisms that lead to the formation of axons and dendrites and to the differential targeting and segregation of proteins and lipids to the two compartments are less well understood. In this review, rather than providing an extensive summary of the literature, we concentrate on a number of recent developments that have provided clues to the molecular mechanisms underlying the generation of polarity in neuronal cells.

II.

NEURONAL POLARITY: WHAT WE KNOW A.

General Axonal and Dendritic Differences

Neurons are considered to be divided broadly into two domains, the axonal domain and the somato-dendritic domain. These two domains differ in a number of features. Usually only one axon extends from the cell body; this long process is often unbranched or infrequently branched until it reaches its synaptic target, where it divides into fine branches with swellings at the pre-synaptic terminals that are the specialized sites for neurotransmitter release (Kandel et al., 1991). Dendrites, in contrast to the axon, are usually post-synaptic; these shorter processes are highly branched and often terminate with a tapered ending (Sargent, 1989). Dendrites, both functionally and structurally, are in many ways an extension of the cell body, sharing with it Golgi elements and smooth and rough endoplasmic reticulum (Peters et al., 1976). Axons, in contrast, do not contain rough endoplasmic reticulum or Golgi elements. Examination of mature neurons reveals that both axons and dendrites may be further divided into sub-domains. In myelinated axons, for example, proteins such as components of the Na+channel are selectively localized at the nodes of Ranvier (Ellisman and Levinson, 1982; Haimovich et al., 1984). Other axonal proteins, such as the synapsins, are particularly concentrated at synaptic densities (for review, see

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Kelly, 1993). Within the somatodendritic compartment, certain neurotransmitter receptors, such as the metabotropic glutamate receptor mGluR6, are concentrated at the post-synaptic site (Nomura et al., 1994). Sub-domain localization in the somatodendritic compartment also extends to mRNA localization. The majority of mRNAs are confined to the cell body, but some mRNAs (including the mRNA for the dendrite-specific microtubule associated protein, MAP-2) extend into the dendritic tree (Steward and Banker, 1992). B.

Axonal and Dendritic Cytoskeletons: Different Directions Using Different MAPs

The distinct differences in morphology and distribution of membranous organelles in axons and dendrites are reflected by underlying differences in the organization and composition of the cytoskeleton. In the axon, microtubules are arrayed in overlapping bundles, with the plus-ends of the tubulin polymers oriented distal to the cell body (Burton and Paige, 198 1; Heidemann et al., 1981). In contrast, in dendrites microtubules are arrayed in a mixed orientation, with both plus- and minusendmicrotubules orienteddistal to the cell body (Burton, 1988; Baas et al., 1988). Axons and dendrites also contain distinct sets of abundant neuron-specific microtubule-associated proteins, or MAPs, which promote microtubule assembly and stability (Ginzburg, 1991; Goedartet al., 1991; Matus, 1994). Axons are characteristically enriched in the microtubule-associated tau protein (Binder et al., 1985; Peng et al., 1986), while dendrites contain the related protein, MAP-2 (Goedertet al., 1991). MAP-2 and tau may contribute generally to the support of neuronal processes by making the microtubules they contain longer, more stable, and stiffer than those in non-neuronal cells (Matus, 1994). When Sf9 cells, which normally have a rounded morphology, were infected with a baculovirus containing tau cDNA, the cells elaborated long, slender, relatively unbranched processes resembling the axons of neurons in culture (Knops et al., 1991). Indeed, inhibition of tau expression by the addition of tau anti-sense oligonudeotides to cerebellar neurons in culture prevents axon formation (Caceres and Kosik, 1990). Although the extension of initial exploratory neurites occurred normally after tau anti-sense treatment, the cells failed to elaborate stable axons, indicating that the conversion of initial neurites (with unstable microtubules) to the mature axon (with stable microtubules) was inhibited (Caceres and Kosik, 1990). In contrast to results using tau anti-sense oligonucleotides, MAP-2 anti-sense treatment does block initial establishment of neurites (Caceres et al., 1992). The differences in microtubule orientation and MAP composition between axons and dendrites suggests that these differences may be used by membranous organelles and secretory vesicles for specific microtubule-based targeting to the axonal or dendritic compartments. The microtubule-based motor protein, kinesin, has been shown to associate with vesicles and has the proper specificity to participate in the transport of vesicles to distal sites of the axon (Vale et al., 1985; Vale, 1987; Schroer and Sheetz, 1991). Kinesin-coated objects move undirectionally toward the plus-ends of

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microtubules. This direction corresponds to anterograde axonal transport in neurons, since axonal microtubules have their plus-ends oriented distally.A number of experiments further support the idea that kinesin transports organelles in an anterograde direction in vivo (Schroer, 1991; Schroer and Sheetz, 1991). Although a single kinesin motor had been thought to serve all cell types, a second conventional kinesin heavy chain (KHC) is specifically expressed in neurons (Niclas et al., 1994). In hippocampal neurons in culture, the ubiquitously expressed KHC (uKHC) is distributed uniformly throughout the neuron, whereas the neuronal KHC is selectively concentrated in the cell body (Niclas et al., 1994). Inhibition of uKHC expression using anti-sense oligonucleotides caused the failure of the axon specific proteins, GAP-43 and synapsin I, to move out of the cell body (Ferreira et al., 1992). Despite the inhibition of the transport of specific proteins, the neurons could still extend processes and acquire an asymmetric morphology. As with microtubules, filamentous actin is present in all regions of the neuron, but is particularly concentrated in certain neuronal subdomains, such as in the peripheral region of extending axonal growth cones (Forscher and Smith, 1988; Mitchison and Kirschner, 1988; Bridgman and Dailey, 1989; Rivas et al., 1992) and in the cortical region of the synapse (Hirokawa et al., 1989). As with microtubuleassociated proteins, it is possible that certain actin binding proteins may be specifically localized to axons or dendrites. Recently, squid axoplasmic organelles have been shown to move on invisible tracks that are sensitive to cytochalasin, suggesting that actin filaments may be an important component of vesicular transport in the axon (Kuznetsov et al., 1992). Using an in vitro assay that permits observation of axoplasmic organelles moving on unipolar actin bundles, Bearer et al. (1993) demonstrated that axoplasmic organelles move on actin filaments in the barbed-end direction, suggesting the presence of a myosin motor on the organelles. Myosin may thus be involved in transport of axcplasmic organelles through the axoplasm, their movement through the cortical actin in the synapse, or some other aspect of axonal function (Bearer et al., 1993). Axons and dendrites both contain neurofilaments, which are cytoskeletal elements of the intermediate filament type (Liem, 1993). Dendrites, in contrast to the axon, often contain relatively few neurofilaments (Matus, 1994).The heavily phosphorylated forms of neurofilament proteins are preferentially localized to the axon (Lee et al., 1987; Lein and Higgins, 1989; Pennypacker et al., 1991).

C.

Differential Protein Localization at the Axonal and Dendritic Domains

Both the initial formation of appropriate neural circuits as well as the directional nature of synaptic transmission rely on the specific localization of proteins to the axonal or dendritic domains. The localization of three classes of proteins are particularly important to consider with respect to the development of neuronal polarity: (i) receptors and ion channels involved in synaptic transmission; (ii) cell adhesion molecules involved in neuronal cell morphogenesis and in axon out-

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growth, fasciculation, pathway selection, and target recognition (Jessell, 1991); (iii) intracellular effector molecules that transmit signals (either synaptic or adhesive) from the extracellular environment into the cell. In this section, the emphasis is placed on where these molecules are localized; possible mechanisms directing their polarized expression are discussed in a later section. The major excitatory transmitter in the central nervous system is glutamate; the major inhibitory transmitters are GABA and glycine (Kandel et al., 1991). The receptors for all three neurotransmitters are multisubunit transmembrane proteins, and as might be expected, all three receptor types are post-synaptic in most neurons and are usually localized to the somatodendritic domain (Triller et al. 1985; Seitandou et al., 1988; Somogyi et al., 1989; Killisch et a]., 1991; Baude et al., 1993; Craig et al., 1993; Nomura et al., 1994). Ion channels also show domain specific localization. The type I and I11 a subunits of the Na+channel are localized to the somatodendritic domain; the type I1 a subunit is axonal and may function in the low threshold needed for action potential generation in the initial segment of the axon (Westenbroek et al., 1989; 1992). Two isoforms of rat K+channels exhibit domain specific localization and function: Kv1.4 is an axonal pre-synaptic K+channel subunit involved in control of neurotransmitter release, and Kv4.2 is a dendritic K+channel subunit potentially important in regulating post-synaptic excitablity (Sheng et al., 1992). The localization of adhesion molecules is much less easy to predict on the basis of the physiological polarity of the neuron. Four major classes of cell surface adhesion receptors have been identified: the cadherins, the immunoglobulin (Ig) superfamily, the selectins, and the integrins (Hynes and Lander, 1992). Of these, the members of the Ig superfamily have been best characterized in terms of axonal or dendritic localization. Members of the Ig superfamily that are membrane spanning proteins have been reported to be either somatodendritic, such as NCAM,*,,(Persohn and Schachner, 1990) or axonai, such as L l (Persohn and Schachner, 1990). One member of the Ig superfamily that is linked to the membrane by a glycosyl phosphatidyinositol (GPI) anchor, TAG-1 (Furley et al., 1990; Yamamoto et al., 1986,1990)has been consistently reported to be localized to the axonal membranes of various neuronal cell types. Less is known about the localization of integrins, the majority of which bind to various extracellular matrix (ECM) molecules and mediate cell-matrix interactions (Hynes and Lander, 1992). ECM molecules directly influence the number and character of neuritic processes extended by different types of neurons i n culture (Bruckenstein and Higgins, 1988a,b;Chamak and Prochiantz, 1989; Lein and Higgins, 1989; Craig and Banker, 1994) and therefore can also directly influence the development of neuronal polarity. Intracellular effector molecules are the least well characterized in terms of axonal and dendritic localization. GAP-43 is a lipid-linked cytoplasmic protein specifically localized to axonal growth cones. GAP-43 has the properties of a membrane-bound regulatory protein that may regulate the growth cone actin cytoskeleton (Skene, 1989; Kelly and Grote, 1993). Another cytoplasmic protein, the

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RII subunit of CAMPkinase, binds to MAP-2 and is localized to the somatodendritic domain (DeCamilli et al., 1986). D.

Epithelial Cell and Neuronal Polarity: Are They Really the Same?

It has recently been proposed that neuronal cells, which arise from neuroepithelial precursors during development, use sorting mechanisms analogous to those present in polarized epithelial cells (Dotti and Simons, 1990; Dotti et al., 1991). According to this idea, axonal protein targeting is analogous to protein targeting to the apical surface of epithelial cells; targeting to the somatodendriticdomain is analogous to targeting to the basolateral surface (see Table IA). Comparison of the localization of several proteins in both neurons and epithelial cells supports this idea: proteins such as influenzahemagglutinin (HA) and Thy-1 are localized to the apical surface of epithelial cells (Rodriguez-Boulan and Sabatini, 1978; Powell et al., 1991b) and have been reported to be localized to the axonal domain of cultured hippocampal neurons (Dotti and Simons, 1990, Dotti et al., 1991). The vesicular stomatitis virus (VSV) G protein is targeted to the basolateral surface (Rodriguez-Boulan and Sabatini, 1978) and has been reported to be localized to the somatodendritic domain of hippocampal neurons (Dotti and Simons, 1990). Localization studies of the transferrin receptor (Fuller and Simons, 1986; Cameron et al., 1991) and the 180 kDa isoform of N-CAM (Persohn and Schachner, 1990; Powell et al., 1991a) are also consistent with the epithelialneuronal analogy. Since many components of the intracellular protein trafficking machinery are common to both neuronal and epithelial cells, as well as to other Table 7.

ComDarison

Protein

of Protein Localization in Epithelial and Neuronal Cells Localization in Neurons

Localization in Epithelia

Dendrites’ Axon’

Apical’

A. VSV-G

HA

Basolateral’

Thy-I

Axon’ (H)

Apical4

TAG-I Transferrin receptor

k o n 5 (GI

Apical6

Dendrites’

Basolateral’

N-CAM 180

Dendrites’

Basolateral’”

GABA, receptor

Dendrites”

Basolateral’*

Thy-1

N o t Polar’ (G)

Apical4

NA’K’-ATPase

N o t polar”(H)

Bas~lateral”,’~

6.

Note:

Source of neuronal localizationdata is from primary cultures of either hippocampalnerons (H)or cerebellar granule neurons (C).All epithelial data is from MDCK cell studies. References: 1. Dotti and Simons, 1990. 2. Rodrigues-Boulanand Sabatini, 1978. 3. Dotti and Simons, 1991. 4. Powell et at., 1991b. 5 . Powell et at., 1997. 6. Powell et at. Submitted. 7.Cameron et at., 1991. 8. Fuller and Simons, 1986. 9. Persohn and Schachner, 1990. 10. Powell et at., 1991a. 11. Killissch et at., 1991. 12. PerezVelasquez and Angelides, 1993. 13.Pietrini et at., 1992. 14. Caplan et at., 1986.

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139

non-polarized cells (Bennett and Scheller, 1993), it is probable that some mechanisms are shared among the different cell types However, recent data indicates that the epitheliaheuronal analogy does not hold true in every case (Table 1B). For example, different isoforms of the Na’, K+-ATPase are expressed in the membranes of both axons and dendrites of mature cultured hippocampal neurons (Pietrini et al., 1992). These same isofonns were found to be localized exclusively to the basolateral surface when expressed endogenously or by stable transfection in renal epithelial cells (Caplan et al., 1986;Pietrini et al., 1992). Thy-1, which is expressed apically in epithelial cells, has been reported to be localized to both axons and dendrites in different neuronal cell types (Morris et al., 1985; Xue et al., 1990). In addition to the conflicting results from localization studies, there are several structural differences between epithelial and neuronal cells which suggest that the two cell types do not have completely analogous membrane trafficking mechanisms. The apical and axonal surfaces differ in the orientation of microtubules; in the axon, microtubules are arrayed with their plus-ends distal to the cell body (Mitchison and Kirschner, 1988), whereas in epithelial cells, microtubules are arrayed with their minus-ends toward the apical surface (Bacallao et al., 1989). Therefore, microtubule orientation differs in the direction of vesicular traffic from the Golgi towards the apical surface or distal tip of the axon; different microtubuleassociated motors would therefore be required for apical-specific or axonalspecific transport of vesicles. The neuronal morphology is spatially more complex than that of epithelial cells, with the axon extending up to 1 meter from the cell body. A physical barrier to diffusion between the axon and dendrites (analogous to the epithelial tightjunction) has been proposed (Kobayashi et al., 1992) but remains completely uncharacterized; however, the need for such a barrier in neurons has been questioned (Futerman et al., 1993). These results suggest that additional mechanisms, other than a simple apical=and basolateral=somatodendritic analogy, may be necessary to explain the polarized targeting of proteins in neurons. Recent work demonstrates that even amongst different epithelial cell types there is heterogeneity in targeting signals and transport pathways (the “flexible epithelial phenotype”) (Rodriguez-Boulan and Powell, 1992; Zurzolo et al, 1993). It would be surprising if the great heterogeneity in morphology between different types of neurons was not also reflected in the use of different targeting mechanisms. Perhaps a variety of cell type specific variations are imposed on a basic constitutive secretory pathway (common to all cells); these variations would confer specialized properties, such as polarized secretion and regulated release, to particular cells.

111.

MECHANISMS FOR THE GENERATION OF NEURONAL POLARITY: WHAT WE SUSPECT

In epithelial cells, both apical and basolateral transport are signal-mediated processes and sorting generally occurs intracellularly (Rodriguez-Boulan and Pow-

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140

ell, 1992). In neuronal cells, the mechanisms resulting in the polarized distribution of proteins to axons and dendrites are not well understood; in this section, we propose three mechanisms to explain polarized expression in neurons. The relative contributions of each of these mechanisms are, at this point in time, completely unknown. A.

lntracellular Targeting

Possible Targeting Signals The GPI anchor and lipid sorting in neurons: As described above, some similarities in targeting pathways have been described for epithelial and neuronal cells. Proteins linked to the membrane by a glycosyl phosphatidyinositol lipid moiety (GPI-linked) proteins are preferentially targeted to the apical surface of epithelial cells (Rodriguez-Boulan and Powell, 1992). Based upon studies of a GPI-linked member of the Ig superfamily,Thy- 1, in cultured hippocampal neurons, it was proposed that the GPI-anchor acts as an axonal targeting signal in neurons equivalent to its role as an apical targeting signal in epithelial cells (Dotti and Simons, 1991). Other studies have demonstrated that another GPI-anchored adhesion molecule, TAG-1, is also localized to the axon (Furley et al., 1990), supporting the general idea that GPI-linked proteins are localized to the axon. However, several reports call the role of the GPI anchor in axonal targeting into question. An in vivo study of the GPI-anchored adhesion molecule, F3F11, in the cerebellar cortex demonstrated that this protein is restricted to the parallel fiber axons of granule cells; however, it is present in the axon, cell body, and dendrites of another cerebellar interneuron, the Golgi cell (Faivre-Sarrailh et al., 1992; FaivreSarrailh and Rougon, 1993). In contrast to its localization in cultured hippocampal neurons, Thy-1 has been reported to be uniformly distributed to both axons and dendrites in cerebellar Purkinje and olfactory neurons in vivo (Morris et al., 1985; Xue et al., 1990) and in cerebellar granule neurons in vitru (Powell et al., 1997). The localization of GPI-anchored proteins may also vary among different GPIlinked proteins in the same cell as well as among different neuronal cell types. Recent results with low density cultures of mouse cerebellar granule neurons demonstrate that at least one GPI-anchored protein is targeted specifically to the axonal domain; two other GPI-anchored proteins are non-polarized in their distribution, suggesting that signals other than the GPI-anchor are required for axonal expression (Powell et al., 1997). Therefore, it appears that in neuronal cells GPI-anchored proteins can be polarized or unpolarized, and can be found in all domains of the membrane (summarized in Table 2). Does the GPI anchor have any role in axonal targeting? There are two possible explanations for the diverse results reported. The first is that the axonal localization of GPI-anchored proteins such as TAG- 1 is due to a signal other than the GPI-anchor (cf. Powell et al., 1991b). Since a number of axonal cell adhesion mole-

141

Polarity in Neuronal Cells

Table 2.

Polarity of CPI-anchored Proteins in Neuronal Cells

Protein

Localization

Cell Type

In Vitrolln Vivo

Reference

Thy-I

axonal

hippocampal

in vitro

Dotti & Simons, 1990

nonpolar

cerebellar

in vitro

Powell et al., 1997

axonal

cerebellar

in vitro

Powell et al., 1997

cerebellar

in vivo

Stottman and Rivas, 1998

TAG-I

spinal cord

in vivo

Furley et al., 1990

PrPC

nonpolar

cerebellar

in vitro

Powell et al., 1997

F2/F11

axonal

cerebellar (granule cells)

in vivo

Faivre-Sarrailh et al., 1992

nonpolar

cerebellar (interneurons)

in vivo

Faivre-Sarrailh et al., 1992

cules are members of the Ig superfamily, a conserved Ig domain may contain proteinaceous axonal targeting information. The second possibility is that the non-axonal localization of certain GPI-anchored proteins in neurons is the result of competition between the GPI-anchor (acting as an axonal targeting signal) and other signals, as yet undefined, present in the molecule. These two possibilities can be tested by expression of genetically modified forms of some of these proteins in neuronal cells. One other piece of the puzzle is still missing in neuronal cells. In epithelial cells, clustering of GPI-anchored proteins with glycolipids, which are also preferentially expressed on the apical surface, is proposed to be involved in apical sorting (Simons and Van Meer, 1988; Van Meer, 1989,1993).Recent studies havedemonstrated that complexes containing GPI-anchored proteins and glycolipids form during transport to the apical surface (Brown and Rose, 1992; Fielder et al., 1993; Lisanti et al, 1993; Zurzolo et al., 1994). In neuronal cells, a polarized distribution of the glycolipids themselves has not yet been demonstrated. An inhibitor of sphingolipid synthesis selectively inhibits axon outgrowth in cultures of hippocampal neurons, suggesting that this class of glycolipids is preferentially inserted into the axon (Hare1 and Futerman, 1993). It will be of interest to determine if GPI-anchored proteins that are polarized to the axon also form aggregates with glycolipids during transport.

Proteinaceous signals for targeting in neurons: In several instances, ion channels which are highly related in primary structure have been found to be differentially targeted to pre- or post-synaptic membrane specializations. Comparison of the sequences of such proteins has implicated certain regions in directing their differential distribution. For example, the Na' channel subunits RI (dendritic) and R I1 (axonal) exhibit 85% sequence similarity but are specifically localized to either of the two domains (Westenbroek et al., 1989). Another example is provided by the fast transient or A type subgroup of voltage-sensitive potassium channels, which contain a central core region with six membrane spanning domains. Targeting sig-

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SHARON K. POWELL and RODOLFO J. RIVAS

nals responsible for the differential axonal and dendritic distributions of two forms of this channel may be present in the poorly conserved, cytoplasmic-facing N and C terminal regions of these proteins (Sheng et al., 1992; Sheng et al., 1994). These regions could act directly as targeting signals, or could serve to modulate the assembly of these proteins (along with components of the sub-membrane cytoskeleton) into domain specific complexes, as has been shown for isoforms of the GABA, receptor (Perez-Velasquez and Angelides, 1993). As described above, a number of cell adhesion molecules of the Ig superfamily are localized to the axon (Ranscht, 1988; Gennarini et al., 1989; Brummendorf and Rathjen, 1993), suggesting the existence of a possible axonal targeting signal in a conserved Ig domain. However, a new member of the Ig superfamily, telencephalin, has been described which is found exclusively on the cell body and dendrites of neurons of the telencephalon (Yoshihara et al., 1994). Comparison of the sequence of this protein with other Ig superfamily members expressed on the axon may help to identify which domains are important for axonal targeting. lntracellular Targeting: The Molecular Players

Although there have been relatively few biochemical studies of protein targeting mechanisms in neurons, several recent studies of other non-neuronal cell types (both polarized and non-polarized) have implicated a variety of proteins in the regulation of membrane trafficking. As described here, many of these proteins are also expressed in neuronal cells, but their role in polarized traffic in the neuron remains to be elucidated.

Rabs: Small GTP-binding proteins of the rab family were first identified as regulators of the secretory process in yeast, and have since been found to exist in multiple forms in most higher cells (reviewed in Simons and Zerial, 1993). Rab proteins follow a cycle of GTP-binding/membrane attachment followed by GTP hydrolysishelease; rabs specific for a variety of intracellular transport steps have been identified. Many (of the more than 20 described) are expressed in all cell types and are implicated in processes common to all cells, such as ER to Golgi transport, intercisternal Golgi transport, and endocytosis. In polarized epithelial cells, functional studies in a permeabilized cell system indicate that rab 8 regulates basolateral transport (although it should be noted that rab8 is expressed in non-polarized cells as well) (Huber et al, 1993a). Rab 17 is the only epithelial-specific rab identified to date and is thought to function in transcytotic delivery between the apical and basolateral surfaces (Lutcke et al., 1993); no rab specific for the direct apical pathway has yet been identified. Several rabs have been identified in neuronal cells, but again, most of these occur in secretory pathways common to other cell types. Huber et al. (1993b) showed that rab 8 is polarized to the dendrites of cultured hippocampal neurons; in the only functional study to date, these workers also showed that anti-sense oligonucleotides directed against

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rab8 affect the dendritic transport of both rab 8 and another dendritic protein. Thus, at least one component of the membrane traffic machinery is localized specifically to dendrites. Rab3a is the only rab to date found to be specific for neural and endocrine tissues, and has been proposed to be a negative regulator of regulated secretion in neuroendocrine cells (Johannes et a], 1994). Rab 3a also associates with synaptic vesicles in neurons, and is thought to be associated with themembrane fusion complex (see below).

Synaptic vesicle proteins: The synaptic vesicle has for some time been viewed as a highly specialized machine for membrane fusion and recycling, and a variety of proteins specific for synaptic vesicles have been isolated by investigators interested in synaptic transmission. However, recent work suggests that the molecular machinery present on synaptic vesicles may regulate a variety of membrane fusion events common to all cell types. Much of this evidence comes from work using in vitro assays for protein transport, which have provided a powerful tool for the identification of the molecular machinery involved in intracellular transport steps. Rothman and co-workers have painstakingly identified several factors required for intercisternal transport in the Golgi (reviewed in Rothman and Orci, 1992). Quite unexpectedly, the membrane receptors for some of these soluble factors were identified as proteins previously identified as abundant constituents of synaptic vesicles (Sollner et al., 1993a; Sudhof et a]., 1993; Whiteheart et al., 1993) (see Table 3). One of the first components of the intracellular transport machinery identified using Rothman’s in vitro assay was a soluble N-ethlymaleimide Sensitive Factor (NSF). More recent work has identified the receptors for NSF which mediate its association with vesicles the a, p, and y forms of SNAP (Soluble NSF-Attachment Protein). The receptors for SNAPS (SNARES) occur in both vesicle-bound (v-SNARE; i.e., synaptobrevin) and target membrane bound (&SNARE; i.e., SNAP-25, syntaxin) forms. The membrane fusion cycle is initiated by the binding of NSF to the SNAPS, which in turn interact with the SNAREs to form an ATPdependent, 20s docking/fusion complex (Sollner et al., 1993b). Rab 3a (Johannes eta]., 1994) and synaptotagmin (Bennett et al., 1992) also associate, possibly transiently, with this complex; both of these proteins are proposed to act as negative regulators of spontaneous release (see Figure 1) (DeBelIo et al., 1993; Elferink et al., 1993; DiAntonio and Schwaartz, 1994),allowing the coupling of excitation and secretion necessary for regulating synaptic transmission. Synaptotagmin binds directly to the SNARE complex, blocking the a-SNAP binding site to block fusion; the nature of the signal displacing synaptotagmin so fusion can occur is not yet known. In parallel with the in vitro transport assays in non-neuronal cells which have identified the role of the SNARES in intracellular transport, several studies identifying have identified synaptobrevin (v-SNARE), syntaxin and SNAP-25 (t-SNARES) as the cellular targets of certain neurotoxins. The pathological effect of these neurotox-

SHARON K. POWELL and R O D O L F O 1. RlVAS

144

Transport Vesicle

Transport Vesicle

synaptobrevin

synaptobrevin synaptotagmin

ZOS

Fusion mmpkx

~

a SNAP

synaptotwmin

Target

msmbrans

syntaxin synaptobrevin

Figure 1.

Table 3.

Synaptic Vesicle Proteins Involved in lntracellular Transport

Protein

Vesicle or Target Membrane

Homologs

Transport Step

synaptotagmin Synaptobrevin

vesicle vesicle

-

SNAP-25 syntaxin A,B

target target

-

Golgi-PM. endocytic recycling Colgi-PM. ER-Colgi ER-Golgi PM. fusion ER-Golgi Golgi-vacuole

Note:

cellubrevin SEC 22 (yeast) BET1 (yeast) BOSI (yeast) SED5 (yeast) PEP12 (yeast)

References as in text. PM. = plasma membrane.

ins is to inhibit neurotransmitterrelease, confirming that these proteins are critical for exocytosis in neuronal cells (Schiavo et al., 1992; Blasi et al., 1993a,b;Bennett et al., 1993; McMahon et al., 1993). Genetic studies in yeast have also identified homologs of synaptic vesicle proteins that function in a variety of intracellular transport steps (Table 3) (Bennett and Scheller, 1993).

CaveolinNIP21: Caveolin is a 21kD protein isolated as an abundant component of the specialized membrane invaginationscalled caveolae (Anderson, 1993). Caveolae are thought to play a role in the uptake of small molecules (Rothberg et al., 1992) and also in intracellular signalling (Glenney and Soppet, 1992; Sargiacomo et al., 1993). There are two lines of evidence suggesting a role for this protein in membrane trafficking. First, caveolin was independently isolated as a component of transport vesicles in polarized epithelial cells (Glenney, 1992; Kurzchalia et al., 1992). Second, it appears that in caveolae, GPI-anchored proteins are found in aggregates that contain caveolin (Lisanti et al., 1993; Sargiacomo et al., 1993). Although caveolae have not been described in neuronal cells, caveolin is expressed in neurons (Simons et al., 1992; Powell and Rivas, unpublished). A direct role for this protein in trafficking remains to be elucidated.

Polarity in Neuronal Cells

B.

145

RNA Localization: A Dendrite-Specific Mechanism?

Another important mechanism for the spatial regulation of protein expression in eukaryotic cells is selective RNA localization, which has been most thoroughly described in Drosophilu and Xenopus eggs and early embryos (reviewed in Kislauski and Singer, 1992; Stephenson and Pokrywka, 1992; Steward and Banker, 1992). In neuronal cells, most mRNAs are found in the cell body; mRNAs are typically excluded from the axonal compartment. A few mRNAs have been reported to be localized to dendrites, most notably the mRNA for the dendritic microtubule-associated protein, MAP-2 (Bruckenstein et al., 1990). Most translatable mRNAs are associated with the cytoskeleton (Singer, 1992); segregation of mRNA into the cell body or dendrites is time-dependent and requires microtubules (Bassell et al. 1994). Polyribosomes are preferentially localized at post-synaptic sites on dendritic spines (as well as at synapses on the dendritic shaft and in the proximal portion of the axon), possibly providing a mechanism for regulating protein synthesis at a single synapse in response to activity (Steward and Banker, 1992). It has been suggested that polyribosomes are excluded from the mature axon because of the tight bundling of microtubules (Baas et al., 1987). Axonal localization of mRNA has also been reported for an axonal protein (Litman et al., 1993); however, since axons appear to lack polyribosomes, it is not clear if these mRNAs can be utilized for domain-specific synthesis. How are mRNAs localized to the dendrites? First, the mRNA must have a signal which allows its specific recognition and transport into the dendrites. Second, the RNA must be selectively retained at the appropriate location. Cis-acting sequences in the 3'-untranslated regions appear to be necessary for selective localization (Papandrikopoulu et al., 1989). In various systems, both microtubules and actin filaments have been shown to be required for RNA localization (Singer, 1992). In Drosophilu, several gene products have been shown to be required for localization of specific transcripts (Stephenson and Pokrywka, 1992). C.

Selective Retention

The specific localization of a number of proteins in neurons is mediated by either direct or indirect interaction with the cytoskeleton. In several cases, retention is mediated by interactions with cell surface proteins which in turn interact with other proteins or environmental cues. Thus, the use of selective retention as a localization mechanism in neurons can quite clearly be affected by factors external to the cell. An k yrinfSpectn'n

In many cell types, macromolecular complexes including the molecules ankyrin and spectrin are responsible for the indirect association of surface proteins with the

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cytoskeleton (Lazarides and Nelson, 1985;Bennett and Gilligan, 1993). Both ankyrin and spectrin are derived from multiple genes and transcripts, and exhibit isoform-specific subcellular localization in neurons (Kordeli et al., 1990; Chan et al., 1993; Lambert and Bennett, 1993a; Malchiodi-Albedi et al., 1993), making them candidate molecules for directing the polarized localization of specific proteins that associate with them. Spectrin is a flexible rod shaped molecule which occurs as a heterotetramer of a and p subunits; the p subunit contains the actin binding domain (Bennett and Gilligan, 1993). Although there is evidence for direct interaction of spectrin with some integral membrane proteins, the association of cell surface proteins with the sub-membrane cytoskeleton is generally mediated by ankyrin (Gumbiner, 1993). Ankyrin is a peripheral membrane protein with an N-terminal domain containing 24 tandem repeats that mediates binding to the membrane and a C-terminal domain that associates with spectrin (Bennett and Gilligan, 1993). Ankyrin has been shown to directly associate with the erythrocyte anion exchanger, voltage-dependent sodium channels, and several cell adhesion molecules of the nervous system, such as L1 and neurofascin (Davis et al., 1993 ). Ankyrin occurs in several isoforms which are selectively localized to cell bodies and dendrites, premyelinated axons, or to the Nodes of Ranvier in myelinated axons (ankyrin,,,,,) (Kordeli et al., 1990). Transgenic mice defective in ankyrin expression exhibit neurological dysfunction, demonstrating the importance of these proteins in proper neuronal function (Lambert and Bennett, 1993b). Both ankyrin and spectrin have been implicated in conveying signals from the environment to the interior of the cell (Bennett and Gilligan, 1993; Gumbiner, 1993). Notch, a Drosophila protein containing ankyrin repeats in its cytoplasmic domain, is involved in the determination of neuronal cell fate, suggesting a role for this domain in the interpretation of localized cues during development (Fortini and Artavanis-Tsakonas, 1993; Diederich et al, 1994). External Factors Directing Selective Retention

Although in many instances it is clear that neurons can exhibit a well polarized morphology in the absence of cellkell interactions, specific interactions with other cells and with the extracellular matrix play an important role in the directing the localization of proteins at pre- and post-synaptic specializations. The molecular basis of this process is best understood at the neuromuscular synaptic junction. Acetylcholine receptors (AChR) are found in clusters of extremely high densities at postsynaptic sites of skeletal muscle. A 220 kDa protein purified from the basal lamina of the neuromuscular junction, agrin, mediates aggregation of the AChR. A receptor for agrin has recently been identified as a a-dystroglycan (Campanelli et al., 1994; Gee et al., 1994; Sealock and Froehner, 1994;), a protein identified via its association with dystrophin. Dystrophin is a spectrin-related protein expressed in skeletal muscle and is defective in Duchenne’s ivIuscular Dystrophy patients (Mat-

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sumara and Campbell, 1994). Thus, the agrin-dystroglycan interaction may serve to direct the assembly of a synapse-specific submembrane cytoskeleton.

IV.

INTRINSIC FACTORS IN NEURONAL POLARITY: WHAT THE CELLS KNOW

The study of neuronal polarity obviously necessitates the establishment of cultures with well differentiated axons and dendrites; most cell lines of neuronal origin do not exhibit well defined axons and dendrites and are thus not optimal models of neuronal polarity (Craig and Banker, 1994). Dissociated primary neuronal cells in culture, which do exhibit well defined axons and dendrites, have therefore been used extensively for studies of neuronal polarity, in particular the low density embryonic hippocampal neuronal culture system developed by Gary Banker and his colleagues (reviewed in Craig and Banker, 1994). The typical pattern of the development of axons and dendrites in this system is illustrated in Figure 2. In the first few hours after plating, the cells extend a lamellipodium around the cell body. After approximately 12 hours in culture, several short “minor processes” are extended from the cell body. Within 1-2 days, one of these processes begins to elongate more than the others; this process eventually differentiates into an axon (Dotti and Banker, 1987). With continued time in culture, the axon and dendrites continue to elongate, and the cells survive in a highly differentiated state, forming synapses, for two or more weeks. At the time that the axon is first identifiable morphologically, molecular markers for axons and dendrites begin to segregate (Dotti et al., 1987; Days in culture Hippocampal pyramidal

- ~ -

0.25

Figure 2.

0.5

1.5

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Godin and Banker, 1989). A similar pattern of development is also exhibited by some other neuronal cell types in low density dissociated culture, such as sympathetic neurons (Bruckenstein and Higgins, 1988 a,b). Although the hippocampal system is the “gold standard” in the field, it is important to emphasize that there is not a single “typical” neuronal morphology. Cell type specific differences in neuronal morphology in vivo are reflected when neurons are cultured at low density in vitro. We have been studying the development of polarity in low density cultures of mouse cerebellar granule cells (Powell et al., 1997). Granule cell development in low density culture can be described in five stages (Figure 2) corresponding precisely to the defined stages of morphogenesis and migration these cells undergo in vivo (Ramon y Cajal, 1894; Jacobson, 1991; Gao and Hatten, 1993). In contrast to hippocampal pyramidal neurons, granule cells first extend a single process, becoming unipolar. After approximately 2 days in culture, the cells extend a second process, becoming bipolar. After 3-4 days in culture, the cells develop a more complex morphology, with a branch point forming in one process quite similar to the characteristic T-shaped axons that these cells form in viva. After 4-5 days in culture, shorter dendritic processes form around the cell body. Accompanying the morphological development of polarity is the segregation of molecular markers for axons and dendrites (Tau-1 and MAP-2, respectively) (Powell et al., 1997). How are these cell type-specific differences in neuronal morphology maintained in low density culture, in the absence of the highly specific cell-cell interactions and other environmental influences the neurons are exposed to in their region of origin? Although the cells in the two systems described above survive and differentiate when removed from the animal, the neurons almost certainly received temporally regulated environmental cues for differentiation prior to placement in culture (for a review, see McConnell, 1991). It should also be noted that the conditions required for the growth and differentiation of neurons at low density often require that a glial feeder or conditioned medium be present, again suggesting that biochemical cues from the environment are necessary development of polarity. For some cell types, conditioned medium is not sufficient: specific heterotypic interactions between cell types are required for complete morphogenesis in culture. This has been demonstrated for cerebellar Purkinje cells, which require contact with granule cells for complete dendritic differentiation (Baptista et al, 1994). Perhaps the most interesting question is how secretory processes are regulated to assist in the formation of characteristic polarized neuronal morphologies. HOW do hippocampal neurons initiate neurite outgrowth at a number of sites in the plasma membrane, first becoming multipolar? In contrast, why do cerebellar granule neurons initiate neurite outgrowth at only one site? Is there perhaps a small GTP-binding protein that is involved in site selection for new membrane insertion and that is differentially regulated between the two cell types? To what extent are these processes influenced by extrinsic factors in different cell types? The answers to these exciting questions will require the adaptation of a variety of mo-

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lecular biological and biochemical techniques standard in other cell types to neuronal cells.

V.

CONCLUSIONS/FUTURE DIRECTIONS

We still know very little about the mechanisms by which proteins are selectively delivered to the axonal and dendritic domains. Neurons contain many proteins known t o play a role in intracellular targeting in other cell types, but for the most part the function of these proteins in neurons has not been directly tested. This is largely due to technical difficulties in obtaining sufficient amounts of purified populations of primary neurons for biochemical assays. At least for the time being, studies of neuronal polarity are dependent on molecular biological approaches, such as the use of anti-sense oligonucleotides to inhibit the expression of known membrane trafficking components and the use of viral vectors to express exogenous proteins.

ACKNOWLEDGMENTS We gratefully acknowledge the support of Dr. Mary E. Hatten and Dr. Enrique Rodriguez-Boulan. We thank Drs. Phyllis Faust and Charles Yeaman for reading the manuscript. Supported by SpinaI Cord Research Foundation grant #1152 (SKP).

REFERENCES Anderson, R. G. W. (1993). Plasmalemmal caveolae and GPI-anchored membrane proteins. Current Biol. 5, 647-652. Baas, P. W., Deitch, J. S., Black, M. M. & Banker, G. A. (1988). Polarity orientation ofmicrotubules in hippocampal neurons: Uniformity in the axon and nonuniformity in the dendrite. Proc. Natl. Acad. Sci. USA 85,8335-8339. Baas, P.W., Sinclair, G. I. & Heidemann, S. R. (1987). Role of microtubules in the cytoplasmic compartmentation of neurons. Brain. Res. 420,73-8 1. Bacallao, R., Antony, C., Dotti, C. G., Karsenti, E., Stelzer, E. H. K. & Simons, K. (1989). The subcellular organization of Madine Darby Canine Kidney cells during the formation of a polarized epithelium. J. Cell Biol. 109,2817-2832. Baptists, C. A,, Hatten, M. E., Blazeski, R. & Mason, C. A. (1994). Cell-cell interactions influence survival and differentiation of purified Purkinje cells in vitro. Neuron 12,243-260. Bassell, G. J., Singer, R. H. & Kosik, K. S. (1994). Association ofPoly(A) mRNA with microtubules in cultured neurons. Neuron 12, 571-582. Baude, A., Nusser, Z., Roberts, J. D. B., Mulvihill, E., McIlhinney, R. A. J. & Somogyi, P. (1993). The metabotropic glutamate receptor (mGluRla) is concentrated at perisynaptic membrane of neuronal subpopulations as detected by imrnunogold reaction. Neuron 11,771-787. Bearer, E. L., DiGiorgis, J. A., Bodner, R. A,, Kao, A. W., & Reese, T. S. (1993). Evidence for myosin motors on organelles in squid axoplasm. Proc. Natl. Acad. Sci. 90, 11252-1256.

150

SHARON K. POWELL and RODOLFO J. RIVAS

Bennett, M. K., Calakos, N., Kreiner, T. & Scheller, R. H. (1992). Synaptic vesicle proteins interact to form a multimeric complex. J. Cell Biol. 116, 761-775. Bennett, M. K. & Scheller, R. H. (1 993). The molecular machinery for secretion is conserved from yeast to neurons. Proc. Natl. Acad. Sci. USA 90,2559-2563. Bennett, M. K., Garcia-Arraras, J. E., Elferink, L. A., Peterson, K., Fleming, A. M., Hazuka, C. D. & Scheller, R. H., (1993). The syntaxin family of vesicular transport receptors. Cell 74, 863-873. Bennett, V. & Gilligan, D. M. (1993). The spectrin-based membrane cytoskeleton and micron-scale organization of the plsma membrane. Ann. Rev. Cell Biol. 9 , 2 7 4 6 . Binder, L. I. Frankfurter, A. & Rebhun, L. 1. (1985). The distribution oftau in the mammalian nervous system. J. Cell Biol. 101, 1371-1378. Black, M. M. & Baas, P. W. (1989). The basis of polarity in neurons. TINS 12,211-214. Blasi,l.,Chapman,E. R., Link, E., Binz,T., Yamasaki,S., DeCamilli, P., Sudhof,T. C.,Niemann,H. & Jahn, R. (1993a). Botulinum neurotoxin A selectively cleaves the synaptic protein SNAP-25. Nature 365, 16@163. Blasi, J., Chapman, E. R., Yamasaki, S., Binz, T., Niernann, H. & Jahn, R. (1993b). Botulinum neurotoxin CI blocks neurotransmitter release by means of cleaving HPC-l/syntaxin. EMBO J. 12,4821-4828. Bridgrnan, P. C. & Dailey, M. E. (1989). The organization of myosin and actin in rapid frozen nerve growth cones. J. Cell Biol. 108,95-109. Brown, D. A. & Rose, J. K. (1992). Sorting of GPI-anchored proteins to glycolipid-enriched membrane subdomains during transport to the apical cell surface. Cell 68, 533-544. Bruckenstein, D. A. & Higgins, D. (1988a). Morphological differentiationofembryonic rat sympathetic neurons in tissue culture. I. Conditions in which neurons form axons but not dendrites. Dev. Biol. 128,324-336. Bruckenstein, D. A. & Higgins, D. (1988b). Morphological differentiation of embryonic rat sympathetic neurons in tissue culture. II. Serum promotes dendritic growth. Dev. Biol. 128, 337-348. Bruckenstein, D. A., Lein, P. J., Higgins, D. & Fremeau, Jr., R. T. (1990). Distinctspatial localizationof specific mRNAs in cultured sympathetic neurons. Neuron 5,809-81 9. Brurnmendorf,T. & Rathjen, F. G. (1993). Axonal glycoproteinswith immunoglobulin-andfibronectin Type 111-related domains in vertebrates: structural features, binding activities, and signal transduction. J. Neurochem. 61, 1207-1219. Burton, P. R. (1988). Dendrites of mitral cell neurons contain microtubules of opposite polarity. Brain Res. 473, 1078-1 11 5. Burton, P. R. & Paige, J. L. (1981). Polarity of axoplasmic microtubules in the olfactory nerve of the frog. Proc. Natl. Acad. Sci. USA 78,3269-3273. Caceres, A. & Kosik, K. S. (1990). Inhibition ofneuronal polarity by tau anti-sense oligonucleotides in primary cerebellar neurons. Nature 343,461-463. Caceres, A,, Mautino, J. & Kosik, K. S. (1992). Suppression of MAP2 in cultured cerebellar macroneurons inhibits minor neurite formation. Neuron 9,607-618. Cameron, P. L., Sudhof, T. C., Jahn, R. & De Camilli, P. (1991). Colocalization of synaptophysin with transferrin receptors: implications for synaptic vesicle biogenesis. J. Cell Biol. 115, 15 1-164. Campanelli, J. T., Roberds, S. L., Campbell, K. P. & Scheller, R. H. (1994). A role for dystrophin-associated glycoproteins and utrophin in agrin-induced AChR clustering. Cell 77, 663474. Caplan, M. J., Anderson, H. C., Palade, G. E. & Jamieson, J. D. (1986). Intracellular sorting and polarized surface delivery of Na+K+-ATPase, an endogenous component of Madine-Darby Canine Kidney cell membranes. Cell 36,623431. Charnak, B. & Prochiantz, A. (1989). Influence of extracellular matrix proteins on the expression of neuronal polarity. Development 106,483491.

Polarity in Neuronal Cells

151

Chan, W., Kordeli, E. &Bennett, V. (1993). 440-kD ankyrinB: Structure ofthe major developmentally regulated domainandselective localization in unmyelinated axons. J. Cell Biol. 123,1463-1473. Craig, A. M. & Banker, G. (1994). Neuronal polarity. Ann. Rev. Neurosci. 17,267-310. Craig, A. M., Jareb, M. & Banker, G. (1992). Neuronal polarity. Curr. Opin. Neurobiol. 2,602-606. Craig, A. M., Blackstone, C. D., Huganir, R. L. & Banker, G. (1993). The distribution of glutamate receptors in cultured rat hippocampal neurons: Post-synaptic clustering of AMPA-selective subunits. Neuron 10, 1055-1068. 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. DeBello, W. M., Betz, H. & Augustine, G. J. (1993). Synaptotagmin and neurotransmitterrelease. Cell 74,947-950. DeCamilli, P., Moretti, M., Donini, D., Walter, U. & Lohmann, S. M. (1986). Heterogeneous distribution of CAMP receptor protein RII in the nervous system: evidence for its intracellular accumulation on microtubules, microtubule-organizing centers, and in the area of the Golgi complex. J. Cell Biol. 103, 189-203. DiAntonio, A. & Schwaartz, T. L. (1994). The effect on synaptic physiology of synaptotagrnin mutations in Drosophilu. Neuron 12,909-920. Diederich, R. J., Matsuno, K., Hing, H. & Artavanis-Tsakonas, S. (1994). Cytosolic interaction between deltex and Notch ankyrin repeats implicates deltex in theNotch signalling pathway. Development 120,473481. Dotti, C.G. & Banker, G. A. (1987). Experimentally induced alteration in the polarity of developing neurons. Nature 330,254-256. Dotti, C. G., Banker, G. A. & Binder, L. 1. (1987). The expression and distribution of the microtubule-associated proteins tau and microtubule-associated protein 2 in hippocampal neurons in situ and in cell culture. Neurosci. 23, 121-130. Dotti, C. G. & Simons, K. (1990). Polarized sorting of viral glycoproteins to the axon and dendrites of hippocampal neurons in culture. Cell 62,63-72. Dotti, C. G., Parton, R. G. & Simons, K. (1991). Polarizedsortingofglypiated proteins in hippocampal neurons. Nature 349, 158-161. Elferink, L. A,, Peterson, M. R. & Scheller, R. H. (1993). A role for synaptotagmin (p65) in regulated exocytosis. Cell 72, 153-159. Ellisman, M. H. & Levinson, S. R. (1982). lmmunocytochernical localization of sodium channel distributions in the excitable membranesofEZectrophoruselectrrcus.Proc. Nati. Acad. Sci. USA 79,6707-671 I . Faivre-Sarrailh, C., Gennarini, G., Goridis, C. & Rougon, G. (1992). F3/Fll cell surface molecule expression in the developing mouse cerebellum is polarized at synaptic sites and within granule cells. J. Neurosci. 12,257-267. Faivre-Sarrailh, C. & Rougon, G. (1993). Are the glypiated adhesion molecules preferentially targeted to the axonal compartment? Molec. Neurobiol. 7 , 4 9 6 0 . Ferreira, A.,Niclas, J., Vale, R. D., Banker, G. & Kosik, K. S. (1992). Suppressionofkinesinexpression using anti-sense oligonucleotides.J. Cell Biol. 117, 595406. Fielder, K., Kobayashi, T., Kurzchalia, T. V. & Simons, K. (1993). Glycosphingolipid-enriched, detergent-insoluble complexes in protein sorting in epithelial cells. Biochemistry 32, 6365-6373. Forscher, P. & Smith S. (1988). Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J. Cell Biol. 107, 1505-1516. Fortini, M. E & Artavanis-Tsakonas, S. (1 993). Notch: neurogenesis is only part ofthe picture. Cell 75, 1245-1247. Fuller, S. D & Simons, K. (1986). Transferrin receptor polarity and recycling accuracy in “tight” and ‘‘leaky’’ strains of Madine-Darby Canine Kidney cells. J. Cell Biol. 103, 1767-1779.

152

SHARON K. POWELL and RODOLFO J. RIVAS

Furley, A J., Morton, Manalo, D., Karagogeos, D., Dodd, J. & Jessell, T. M. (1990). The axonal glycoprotein TAG-I is an immunoglobulin superfamily member with neurite outgrowth-promoting activity. Cell 61 157-170. Futerman, A. H., Khanin, R. & Segal, L. A. (1993). Lipid diffusion in neurons. Nature 362, 119. Gao, W. Q. & Hatten, M. E. (1993). Neuronal differentiation rescued by implantation ofweaver granule cell precursors into wild-type cerebellar cortex. Science 260,367-369. Gee, S. H., Montanaro, F., Lindenbaum, M. H. & Carbonetto, S. (1994). Dystroglycan-a, a dystrophin-associated glycoprotein, is a functional agrin receptor. Cell 77,675-686. Gennarini, G., Cibelli, G, Rougon, G., Mattei, M.-G. & Goridis, C. (1989). The mouse neuronal cell surface protein F3: a phosphatidylinositol-anchoredmember of the immunoglobulin superfamily related to contactin. J. Cell Biol. 109, 775-788. Glenney, J. R. (1992). The sequence of human caveolin reveals identity with VIP 21, a component of transport vesicles. FEBS Lett. 3 1 4 , 4 5 4 8 . Glenney, J. R. & Soppet, D. (1992). Sequence and expression of caveolin, a protein component of caveolae plasma membrane domains phosphorylated on tyrosine in Rous sarcoma virus-transformed fibroblasts. PNAS 89, 10517-10521. Ginzburg, I. (1991). Neuronal polarity: Targeting of microtubule components into axons and dendrites. TIBS 16,257-261. Goedert, M., Crowther, R. A. & Gamer, C. C. (1991). Molecular characterization of microtubule-associated proteins tau and MAP-2. TINS 14, 193-199. Goslin, K. & Banker, G. (1989). Experimental observations on the development of polarity by hippocampal neurons in culture. J. Cell Biol. 108, 1507-1516. Gumbiner, B. M. (1993). Proteins associated with the cytoplasmic surface of adhesion molecules. Neuron 1I, 55 1-564. Haimovich, B., Bonilla, E., Casadei, J. & Barchi, R. (1984). Immunocytochemical localization of the mammalian voltage sensitive sodium channel using polyclonal antibodies against the purified protein. J. Neurosci. 4,2259-2268. Harel, R. & Futerman, A. H. (1993). Inhibition of sphingolipid synthesis affects axonal outgrowth in cultured hippocampal neurons. J. Biol. Chem. 268, 14476-14481. Heidemann, S. R., Landers, J. M. & Hamborg, M. A. (1981). Polarity orientationofaxonal microtubules J. Cell Biol. 91,661465. Hirokawa, N., Sobue, K., Kanda, K., Harada, A. & Yorifuji, H. (1989). The cytoskeletal cytoarchitecture of the pre-synaptic terminal and molecular structure of synapsin I. J. Cell Biol. 108, 11 1-126. Huber, L. A,, Pimplikar, S., Parton, R. G., Virta, H., Zerial, M. & Simons, K. (1993a). Rab8, a small GTPase involved in vesicular traffic between the TGN and the basolateral plasma membrane. J. Cell Biol. 123,3545. Huber, L. A., de Hoop, J., Dupree, P.,Zerial, M. & Simons, K. (l993b). Protein transport to the dendritic plasma membrane is regulated by rab8p. J. Cell Biol. 123,47-55. Hynes, R. 0.& Lander, A. D. (1992). Contact and adhesive specificities in the associations, migrations, and targeting of cells and axons. Cell 68,303-322. Jacobson, M. (1991). Developmental Neurobiology. Plenum Press, New York, NY. Jessell, T. M. (1991). Adhesion molecules and the hierarchy of neural development. Neuron I , 3-13. Johannes, L., Lledo, P.-M., Roa, M., Vincent, J.-D., Henry, J.-P., & Darchen, F. (1994). The GTPase Rab3a negatively controlscalciumdependent exocytosisin neuroendocrinecells. EMBO J. 13,2029-2037. Kandel, E. R., Schwa&, J. H. & Jessell, T. M. (1991). Principles of Neural Science. Appleton and Lange, Nonvalk, CT. Kelly, R. B. (1993). Storage and release of neurotransmitters. Cell 72Neuron 10 (Suppl.), 43-53. Kelly, R. B. & Grote, E. (1993). Protein targeting in the neuron. Ann. Rev. Neurosci. 16,95-127. Killisch, I., Dotti, C. G., Laurie, D. J., Luddens, H. & Seeburg, P. H. (1991). Expression patterns of GABA-A receptor subtypes in developing hippocampal neurons. Neuron 7,927-936.

Polarity in Neuronal Cells

153

Kislauski, E. H. & Singer, R. H. (1992). Determinants of mRNA localization. Curr. Op. Cell Biol. 4, 975-978. Knops, J., Kosik, K. S., Lee, G., Pardee, J. D., Cohen-Gould, L. & McConlogue, L. (1991). Overexpression of tau in a non neuronal cell induces long cellular processes. J. Cell Biol. 114, 725-733. Kobayashi, T., Storrie, B., Simons, K. & Dotti, C. G. (1992). A functional barrier to movement of lipids in polarized neurons. Nature 359,647450. Kordeli, E., Davis J., Trapp, B. & Bennett, V. (1990). An isoform of ankyrin is localized at nodes of Ranvier in myelinated axons of central and peripheral nerves. J. Cell Biol. 110, 1341-1352. Kurzchalia, T. V., Dupree, P., Parton, R. G., Kellner, R., Virta, H., Lehnert, M. & Simons, K. (1992). VIP21, a 21-kD membrane protein is an integral component of trans-Golgi network-derived transport vesicles. J. Cell Biol. 118, 1003-1014. Kuznetsov, S.A., Langford, G. M., & Weiss, D.G. (1992). Actin-dependent organelle movement in squid axoplasm. Nature 356,722-725. Lambert, S. &Bennett, V. (1993a). Post-mitoticexpressionofankyrin-Randbeta-R-spectrin in discrete neuronal populations of the rat brain. J. Neurosci. 13,3725-3735. Lambert, S. &Bennett, V. (1993b). Fromanemiatocerebellardysfunction: areviewofthe ankyringene family. Eur. J. Biochem. 21 1, 1-6. Lazarides, E. & Nelson, W. J. (1985). Expression and assembly of the erythroid membrane-skeletal proteins ankyrin (goblin) and spectrin in the morphogenesis of chick neurons. J. Cell. Biochem. 27,423441. Lee, V. M., Carden, M. J., Schlaepfer, W. W. & Trojanowski, J. Q. (1987). Monoclonal antibodies distinguish sevral differentially phosphorylated states of the two largest rat neurofilament subunits (NF-H andNF-M) and demonstrate their existence in the normal nervous system of adult rats. J. Neurosci. 7, 3474-3488. Liem, R. K. H. (1 993). Molecular biology of neuronal intermediate filaments. Curr. Opin. Cell Biol. 5, 12-16. Lein, P. J. & Higgins, D. (1989). Laminin and basement membrane extract have different effects on axonal and dendritic outgrowth from embryonic sympathetic neurons in vitro. Dev. Biol. 136, 330-345. Lisanti, M. P., Tang, Z. L. & Sargiacomo, M. (1993). Caveolin forms a hetero-oligomeric protein complex that interacts with an apical GPI-linked proteins: implications for the biogenesis of caveolae. J. Cell Biol. 123,595-604. Litman, P., Barg, J., Rindzoonski, L. & Ginzburg, I. (1993). Subcellular localization of tau mRNA in differentiating neuronal cell culture: Implications for neuronal polarity. Neuron 10,627-638. Lutcke, A., Jansson, S. J., Parton, R. G., Chavrier, P., Valencia, A,, Huber, L. A. Lehtonen, E. & Zerial, M. (1993). Rab 17, anovel small GTP-ase, is specific for epithelial cells and is induced during cell polarization. J. Cell Biol. 121, 553-564. Malchiodi-Albedi,F.,Ceccarini,M., Winkelman, J. C., Morrow, J. S. & Petrucci,T. (1993). The270kD splice variant of erythrocyte P-spectrin (PlZ2) segregates in vivo and in vitro to specific domains of cerebellar neurons. J. Cell Sci. 106, 67-78. Matsumara, K. & Campbell, K. P. (1994). Dystrophin-glycoprotein complex: its role in the molecular pathogenesis of muscular dystrophies. Muscle Nerve 17,2-15. Matus, A. (1994). Stiff microtubules and neuronal morphology. TINS 17, 19-22. McConnell, S. K. (I99 I). The genetationofneuronal diversity in the central nervous system. Ann. Rev. Neurosci. 14,269-300. McMahan, U. J. & Wallace, B.G. (1989). Molecules in the basal lamina that direct the formation of synaptic specializations at neuromuscularjunctions. Dev. Neurosci. 11,227-247. McMahon, H. T., Ushkaryov, Y. A., Edelmann, L., Link, E.;Binz,T.,Niemann, H., Jahn, R. & Sudhof, T. C. (1993). Cellubrevin is a ubiquitous tetanus-toxin substrate homologous to a putative synaptic vesicle fusion protein. Nature 364,346349.

154

SHARON K. POWELL and RODOLFO 1. RIVAS

Mitchison,T. & Kirschner, M. (1988). Cytoskeletal dynamics and nervegrowth. Neuron I , 761-772. Morris, R. J., Beech, J. N., Barber, P. C. & Raisman, G. (1985). Early stage of Purkinje cell maturation demonstratedby Thy-l immunohistochemistryon postnatal rat cerebellum.J. Neurocytol. 14,427452. Niclas, J., Navone, F., Hom-Booher, N. & Vale, R. D. (1994). Cloning and localization of a conventional kinesin motor expressed exclusively in neurons. Neuron 12, 1059-1072. Nomura, A., Shigemoto, R., Nakamura, Y., Okamoto, N., Mizuno, N. & Nakanishi, S. (1994). Developmentally regulated post synaptic localization of a metabotropic glutamate receptor in rat rod bipolar cells. Cell 77, 361-369. Papandrikopoulu, A., Doll, T., Tucker, R. P., Garner, C. G. & Matus, A. (1989). Embryonic MAP-2 lacks the cross-linking sidearm sequences and dendritic targeting signal of adult MAP-2. Nature 340,65&652. Peng, I., Binder, L. I. &Black, M. M. (1986). Biochemical and immunological analysis ofcytoskeletal domains of neurons. J. Cell Biol. 102,252-262. Pennypacker, K., Fischer, 1. & Levitt, P. (1991). Early in vitro genesis and differentiation ofaxons and dendrites by hippocampal neurons analyzed quantitatively with neurofilament-H and microtubule-associated protein 2 antibodies. Experimental Neurology 1 1 I , 25-35. Perez-Velasquez, J. L., & Angelides, K. J. (1993). Assembty of GABA, receptor subunits determines sorting and localization in polarized cells. Nature 361,457460. Persohn, E. & Schachner, M. (1990). Immunohistological localization of the neural adhesion molecules LI and N-CAM in the developins hippocampus ofthe mouse. J. Neurocyte 19, 807-819 Peters, A,, Palay, S. L. & deF. Webster, H. (1976). The Fine Structure of the Nervous System: The Neurons and Supporting Cells. W.B. Saunders, New York. Pietrini, G., Matteoli, M., Banker, G. & Caplan, M. J. (1992). Isoforms of the Na+,K+-ATPase are present in both axons and dendrites ofhippocampal neurons in culture. Proc. Natl. Acad. Sci. USA 89,84148418, Powell, S.K., Cunningham, B. A,, Edelman, G. M. & Rodriguez-Boulan, E. (1991a). Transmembrane and GPI anchored forms of NCAM are targeted to opposite domains of polarized epithelial cells. Nature 353, 76-77. Powell, S. K., Lisanti, M. P. & Rodriguez-Boulan, E. (1991b). Thy-I expresses two signals for apical localization in epithelial cells. Am. J. Physiol. 260, C71 S-C720. Powell, S. K., Rivas, R. J., Rodriguez-Boulan, E. & Hutten, M. (1997). Development of polarity in cerebellar granule neurons in vitro. J. Neurobiol. 32,223-236. Ramon y Cajal, S. (1894).New IdeasontheStructureoftheNervous System inMan andVertebrates(tr. Swanson, N. & Swanson, L. W. 1990). MIT Press, Cambridge, MA. Ranscht, B. (1988). Sequence of contactin, a 130-kD glycoprotein concentrated in areas of interneuronal contact, defines a new member of the immunoglobulin supergene family in the nervous system. J. Cell Biol. 107, 1561-1573. Rivas, R. J., Burmeister, D. W. & Goldberg, D. J. (1992). Rapid effects of laminin on the growth cone. Neuron 8, 107-1 15. Rodriguez-Boulan,E. & Sabatini, D. (1978). Asymmetric budding ofviruses in epithelial monolayers: a model system for study ofepithelial polarity. Proc. Natl. Acad. Sci. 75, 5071-5075. Rodriguez-Boulan, E. & Powell, S. K. (1992). Polarity of epithelial and neuronal cells. Ann. Rev. Cell Biol. 8, 395427. Rothberg, K. G., Heuser, J . E., Donzell, W. C., Ying, Y. S., Glenney, J. R. & Anderson, R. G. (1992). Caveolin, a protein component of caveolae membrane coats. Cell 68,673-682. Rothman, J. E. &Orci, L. (1992). Moleculardissectionofthesecretorypathway.Nature355,409~15. Sargent, P. (1989). What distinguishes axons from dendrites?Neurons know more than we do. TINS 12, 203-205. Sargiacomo, M., Sudol, M., Tang, 2. & Lisanti, M. P., (1993). Signal tranducing molecules and glycosyl-phosphatidyl-inositollinked proteins form a caveolin-rich insoluble complex in MDCK cells. J . Cell Biol. 122, 789-807.

Polarity in Neuronal Cells

155

Schiavo, G., Benfati, F., Poulain, B., Rossetto, O., Polverino de Laureto, P., DasGupta, B. R. & Montecucco, C. (1992). Tetanus and botulinum B neurotoxins block transmitter release by proteolytic cleavage of synaptobrevin. Nature 359,832-835. Op.CellBiol.3,133-137. Schroer,T. A.(1991).Associationofmotorproteinswithmembranes.Curr. Schroer, T. A. & Sheetz, M. P. (1991). Functions of microtubule-based motors. Ann. Rev. Phys. 53, 629-652. Sealock, R. & Froehner, S.C. (1994). Dystrophin-associated proteins and synapse formation: is a-dystroglycan the agrin receptor? Cell 77,617-619. Seitandou, T., Triller, A. & Korn, H. (1988). Distribution of glycine receptors on the membrane of a central neuron: an immunoelectron microscopy study. J. Neurosci. 8,43 194333. Sheng. M., Tsaur, M.-L., Jan, Y. N. & Jan, L. Y. (1992). Subcellular segregation of two A-type K+ channel proteins in rat central neurons. Neuron 9,271-284. Sheng. M., Tsaur, M.-L., Jan, Y. N. & Jan, L. Y. (1994). Contrasting subcellular localization of the Kv1.2 K+ channel subunit in different neurons of the rat brain. J. Neurosci. 14, 2408-24 17. Simons, K. & Van Meer, G. (1988). Lipid sorting in epithelial cells. Biochem. 27; 197-202. Simons, K. & Zerial, M. (1993). Rab proteins and the road maps for intracellular transport. Neuron 11: 789-799. Simons, K., Dupree, P., Fiedler, K., Huber, L. A,, Kobayashi, T., Kurzchalia, T., Olkkonen, V., Pimplikar, S., Parton, R. & Dotti, C. (1992). Biogenesis of cell surface polarity in epithelial cells and neurons. Cold Spring Harbor Symp. Quant. Biol. 57,611-619. Singer, R. H. (1992). The cytoskeleton and mRNA localization. Curr. Op. Cell Biol. 4, 15-19. Skene, J. H. (1989). Axonal growth-associated proteins. Ann. Rev. Neurosci. 12., 127-156. Sollner, T., Whiteheart, S. W., Brunner, M., Erdjument-Bromage, H., Geromanos, S., Tempst, P. & Rothman, J. E. (1993a). SNAP receptors implicated in vesicle targeting and fusion. Nature 362, 318-324. Sollner, T., Bennett, M. K., Whiteheart, S. W., Scheller, R. H. & Rothman, J. E. (1993b). A protein assembly-dissassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 75, 409418. Somogyi, P., Takagi, H., Richards, J. G. & Mohler, H. (1989). Subcellular localization of benzodiazepine/GABA-A receptors in the cerebellum of rat, cat, and monkey using monoclonal antibodies. J. Neurosci. 9,2197-2209. Stephenson, E. C. & Pokrywka, N. J. (1992). Localization of bicoid message during Drosophila oogenesis. Curr. Top. Dev. Biol. 26,23-24. Steward. 0. & Banker, G. A. (1 992). Getting the message from the gene to the synapse: sorting and intracellular transport of RNA in neurons. TINS 15, 180-186. Stottman, R. S. & Rivas, R. J. (1998). Relationship of Purkinje cell dendritic development to the distribution of TAG-I and synaptophysin in the developing rat cerebellar cortex: A confocal microscopy study. J. Comp. Neurol. (in press). Sudhof, T. C., De Camilli, P., Niemann, H. & Jahn, R. (1993). Membrane fusion machinery: insights from synaptic proteins. Cell 75, 1 4 . Triller, A., Cluzeaud, F., Pfeiffer, F., Betz, H. & Korn, H. (1985). Distribution of glycine receptors at central synapses: an immunoelectron microscopy study. J. Cell Biol. 101,683-688. 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. Vale, R. D. (1987). lntracellular transport using microtubule-based motors. Ann. Rev. Cell Bio. 3, 347-378. Van Meer, G. (1989). Lipid traffic in animal cells. Ann. Rev. Cell Biol. 5,247-275. Van Meer, G. (1993). Transport and sorting of membrane lipids. Cum. Op. Cell Biol. 5,661473. Westenbroek, R. E., Merrick, D. K. & Catterall, W. A. (1989). Differential subcellular localization of the RI and RII Na+ channel subtypes in central neurons. Neuron 3,695-704.

SHARON K. POWELL and RODOLFO J. RIVAS

156

Westenbroek, R. E., Noebels, J. L. & Catterall, W. A. (1992). Elevated expression of type 11 Na+ channels in hypomyelinated axons of shiverer mouse brain. J. Neurosci. 12,2259-2267. Whiteheart,S.W., Griff, I. C., Brunner, M., Clary, D. O., Mayer,T., Buthrow, S. A. &Rothman, J. E. (1993). SNAP family ofNSF attachment proteins includes a brain-specific isofonn. Nature 362,353-355. Xue, G. P., Calvert, R. A. & Morris, R. J. (1990). Expression of the neuronal cell surface glycoprotein Thy- 1 is under post-transcriptional control and is spatially regulated in the developing olfactory system. Development 198,851-864. Yamamoto, M., Boyer, A. M., Crandall, J. E., Edwards, M. & Tanaka, H. (1986). Distribution of stage specific neurite-associated proteins in the developing murine nervous system recognized by a monoclonal antibody. J. Neurosci. 6,3576-3594. Yamamoto, M., Hassinger, L. & Crandall, J. E. (1990). Ultrastructural localization of stage-specific neurite-associated proteins in the developing rat cerebral and cerebellar cortices. J. Neurocytol. 19,619-627.

Yoshihara, Y., Oka, S.,Nemoto, Y., Watanabe, Y.,Nagata, S., Kagamiyama, H. & Mori, K. (1994). An I-CAM related neuronal glycoprotein, telencephalin, with brain-segment specific expression. Neuron 12,541-553. Zurzolo, C., Lisanti, M. P., Caras, 1. W., Nitsch, L. & Rodriguez-Boulan, (1993). Glycosyl-phosphatidyl inositol-anchored proteins are preferentially targeted to the basolateral surface in Fischer rat thyroid cells. J. Cell Biol. 121, 1031-1039. Zurzolo, C., van’t Hof, W., van Meer, G. & Rodriguez-Boulan, E. (1994). VIP2l/caveolin, glycosphingolipid clusters and the sorting of glycosyl phosphatidyl-inositol anchored proteins in epithelial ells. EMBO J. 13,42-53.

POLARITY AND DEVELOPMENT OF THE CELL SURFACE IN SKELETAL MUSCLE

Annelise 0.Jorgensen

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 II. Organization of Skeletal Muscle Fibers in Muscle Tissue . . . . . . . . . . . . . . . 159 Ill. Polarity of Structurally Distinct but Continuous Domains of the Cell Surface of Adult Skeletal Myofibers . . . . . . . . . . . . . . .161 A. Ultrastructure. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 B. Protein Composition of Transverse (T)-Tubules and Surface Sarcolemma 165 IV. Development of a Polarized Cell Surface Membrane in Skeletal Muscle . . . 178 A. Morphological Aspects. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 B. Molecular Approaches. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181

1.

INTRODUCTION

Skeletal muscle fibers are highly polarized. They are specialized to contract in response to depolarizationof the plasma membrane. For this purpose, skeletalmuscle fibers contain a large number of contractile apparati called myofibrils that contract in response to a rapid increase in the concentration of cytosolic CaZ+.In resting Advances in Molecular and Cell Biology Volume 26, pages 157-199. Copyright Q 1998 by JAI Press Inc. All right of reproductionin any form reserved. ISBN: 0-7623-0381-6

157

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skeletal muscle, Ca2+is stored in the sarcoplasmic reticulum (SR), a specialized form of endoplasmic reticulum, and is released in response to depolarization of the plasma membrane. The polarity of the SR is well established with respect to its ultrastructure (Franzini-Armstrong, 1980, 1994),protein composition (Fleischer and Inui, 1989; Lytton and MacLennan, 1991) and the distribution of its proteins in two distinct domains (Fleischer and Inui, 1989; Lytton and MacLennan, 1991; Franzini-Armstrong and Jorgensen, 1994). Similarly, ultrastructural studies have demonstrated that the plasma membrane in skeletal muscle fibers is polarized and composed of two structurally distinct but continuous domains, namely the surface sarcolemma (SL) and its tubular invaginations called transverse (T)-tubules (Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994). Early physiological studies imply that T-tubules are specialized for signal transduction events essential for excitation-contraction (E-C) coupling (Peachey and FranziniArmstrong, 1983; Rios et al., 1992; Franzini-Armstrong, 1994). In contrast, the specific association of the surface SL with a basal lamina on the extracellular side (Sanes et al., 1986; Sanes, 1994) and the presence of various filamentous structures on the cytoplasmic side (Pierobon-Bormioli, 1981) clearly pointed to a specific role for the surface SL in linking myofibrils to the internal side of the surface SL and in attaching the muscle fiber itself to the surrounding extracelluar matrix. The polarity of the surface SL and the T-tubules with respect to structure and function in turn predicted that these two domains of the plasma membrane would each have unique proteins, as has been demonstrated, for distinct domains of the plasma membrane in other polarized cells including epithelial cells and neurons (Rodriguez-Boulan and Powell, 1992). However knowledge of the protein composition of the distinct domains of the plasma membrane in skeletal muscle has been slow to accumulate. Advances in monoclonal antibody technology and molecular biological techniques have been key to obtaining specific probes with which one can begin to identify and characterize cell surface proteins of skeletal muscle and to determine their precise subcellular distribution in relation to the structurally distinct domains of the cell surface. Since T-tubular membranes assemble de novo during skeletal muscle development both in situ (Kelly, 1971) and in culture (Ishikawa, 1968), it has been feasible to begin to probe the spatial-temporal accumulation of T-tubular and surface SL specific proteins during the biogenesis of T-tubules and triads. The present difficulty in obtaining adequate amounts of highly purified surface SL and Ttubular membranes from skeletal muscle cells developing in situ and in culture combined with the fact that these developing systems are not highly synchronized have so far impeded detailed studies of the dynamic aspects of the assembly of Ttubular or surface SL specific proteins into their target membrane domains. The use of mutants that either lack or have defective genes coding for some of the subunits of hetero-oligomeric protein complexes specific for either T-tubules (Powell, 1990; Flucher, 1992;Franzini-Armstrong and Jorgensen, 1994) or the surface SL (Matsumura and Campbell, 1994; Campbell, 1995) have provided some clues regarding the sequence of some of the events leading to the accumulation of T-tubular and sur-

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face SL proteins into their target membranes. It is likely that identification and characterization of additional T-tubular and SL proteins combined with molecular biological approaches will soon enable us to study the roles of the various proteins in the assembly and function of the polarized cell surface membrane in skeletal muscle. Many excellent reviews have been published on various aspects of the cell surface in adult and developing skeletal muscle. Most of these are devoted to specific aspects of either the surface SL (e.g., the dystrophin glycoprotein complex, see Ervasti and Campbell, 1993a; Tinsley et al., 1994, Campbell, 1995), the membrane cytoskeleton (Small et al., 1992), the assembly of costameres (Fulton, 1993), or of the T-tubules (e.g., the structure and development of the excitation-contraction coupling units, see Franzini-Armstrong and Jorgensen, 1994),the structure and function of the 1,4-dihydropyridine receptor (Campbell et al., 1988, Catterall, 1991). Although features of each of these two membrane systems are often compared, only rarely are they presented in the same review. The purpose of this chapter is to provide an overview of the polarized features of the surface SL and T-tubular membranes with respect to their ultrastructure, protein composition and development. Due to the large volume of literature, regrettably only representative reviews and references have been quoted.

II.

O R G A N I Z A T I O N OF SKELETAL MUSCLE FIBERS IN MUSCLE TISSUE

Skeletal muscle fibers (cells) are very large cylindrically shaped multinucleated cells. Their nuclei are located at the cell periphery. With few exceptions the cells are arranged in bundles parallel to one another and to the longitudinal axis of the muscle fiber. They are embedded in connective tissue containing blood vessels of various sizes and axons of the motor neurons that innervate them. The axon of one motor neuron branches and innervates many skeletal muscle fibers (Cormack, 1993). Depolarization of the plasma membrane by the innervating motor neuron results in the release of the neurotransmitter acetylcholine from the nerve terminal. Acetylcholine binds to acetylcholine-receptors localized on the tip of the folds of the specialized domain of the skeletal muscle membrane called the neuromuscular junction (Engel, 1994). This event, in turn, elicits the depolarization of the surface SL and its transversely oriented invagina;ions, the T-tubules (Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994). This produces a message that is transmitted from T-tubules to the closely apposed specialized domain of the internal Ca2+-storagecompartment called the sarcoplasmic reticulum (Rios et al., 1992). The release of Ca2+from the specialized domain of the sarcoplasmic reticulum (SR) called the junctional SR (j-SR) into the cytosol triggers the contraction of myofibrils (Peachey and Franzini-Armstrong, 1983; Rios et al., 1992; FranziniArmstrong, 1994) (see Figure 1 for nomenclature).

8 Figure 1. (A) Diagrammatic model of the structural relationship between three distinct but continuous cell surface domains in adult skeletal muscle’ namely transverse (T)-tubules, surface sarcolemma (SL) and caveolae. The close apposition of T-tubules to junctional sarcoplasmic reticulum (j-SR) (triad) and the position of A-bands and I-bands in relation to that of T-tubules are also illustrated. (B)Model of a junctional complex between T-tubules and junctional SR called the triad showing the relative distribution of the ryanodine receptor, the 1,4-dihydropyridine receptor (DHPR) and TS28 in this complex. (C)Model showing the distribution of costameres at the Z-line and M-line domains of the surface sarcolemma in adult skeletal muscle.

161

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111. POLARITY OF STRUCTURALLY DISTINCT BUT CONTINUOUS DOMAINS OF THE CELL SURFACE OF ADULT SKELETAL MYOFIBERS A.

Ultrastructure

The cell surface of skeletal myofibers is composed of three major domains, namely the surface SL and its two structurally distinct types of invaginations

C

Basement membrane

-

h

__-I.---

Surface sarcolemma (SL)

_/--

/.-_.---

I

Caveohe connecting T-tUbUle to SL

1 Costameres:

T-tubule

-;:I. *-

\

Z-line

M-line

} A-band } I-band

'Peripheral myofibril Figure 7 continued

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(Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994) (Figures 1A and 1C). One of these constitutes the extensive tubular invaginations called transverse (T)-tubules (Franzini-Armstrong and Porter, 1964; Huxley, 1964; Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994), while the other one is composed of a large number of small flask-shaped invaginations called caveolae (Dulhunty and Franzini-Armstrong, 1975; Franzini-Armstrong, 1994). Although the depolarization event is transmitted both along the surface SL and into T-tubules, the T-tubules alone transmit the depolarization elicited signal to the closely apposed junctional SR that in turn releases Ca2+essential for contraction (Fleischer and Inui, 1989; Rios et al., 1992). Additional specialized domains of the surface SL include the postsynaptic membrane of the neuromuscularjunction (Engel, 1994) and the myotendenous junctions (sites of end-on insertion of myofibrils into the sarcolemma) (Horwitz et al., 1994). Their structural and molecular features are outside the scope of this chapter. The ultrastructural features of the surface SL (Horwitz et al., 1994), T-tubules (Peachey and Franzini-Armstrong, 1983; Franzini-Amstrong, 1994), and the junctional complexes between T-tubules and the junctional SR (triads) (Franzini-Armstrong and Jorgensen, 1994; Franzini-Armstrong, 1994) have been extensively reviewed. A diagram depicting some unique structural features of these membrane systems and their association with the sarcoplasmic reticulum is presented in Figure 1.

SurfaceSarcolemma (SL) Morphological studies have shown that the extracellular side of the surface SL, in contrast to that of T-tubules, is associated with a basement membrane composed of extracellular matrix proteins including laminin, fibronectin, collagen IV and heparan sulphate (Sanes et al., 1986; Sanes, 1994) (Figure 1A and 1C). On the cytoplasmic side of the surface SL, thin fibrous structures bridge the Z-lines and the Mlines of peripheral myofibrils to the surface SL (Pierobon-Bormioli, 198I ) (Figure 4). It has been proposed that those fibrous structures correspond to costameres (Craig and Pardo, 1983; Pardo et al., 1983a,b) (Figure IC). During contraction, which results in shortening of the I-band but not the A-band, this attachment of myofibrils to the SL at the Z-lines results in ‘festooning’ of the surface SL between neighboring Z-lines as the Z-lines are pulled closer together. Thus it appears that the surface SL plays important roles in maintaining the proper structural organization between myofibrils and the surface SL, as well as in the appropriate organization of muscle fibers in the connective tissue via the basement membrane. In contrast to Ttubules, the surface SL in adult skeletal muscle is only rarely observed to be closely apposed to junctional sarcoplasmic reticulum (Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994; Horwitzet a]., 1994; see section on T-tubules below, p. 163) implying that the surface SL does not directly transmit signals to the SR resulting in the release of Ca2+essential to induce contraction. Freeze fracture stud-

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ies have shown that the density and distribution of integral membrane particles of the surface SL are clearly distinct from those of T-tubules and caveolae (Dulhunty and Franzini-Armstrong, 1975; Severs, 1988; Franzini-Armstrong, 1975). Transverse m-Tubules

T-tubules are continuous with the surface SL (Franzini-Armstrong and Porter, 1964; Huxley, 1964) to which they are often observed to be connected via single or series of interconnected caveolae as detailed below (Zampighi et al., 1975; Franzini-Armstrong, 1986)(Figure 1A). T-tubules form extensive tubular invaginations at regular intervals along the myofibers and extend transversely to the center of the myofiber (Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994). Their surface area in fast skeletal fibers is approximately 10 fold higher than that of the surface SL (Eisenberg, 1983). They surround individual myofibrils at the level of the interface between A- and I-bands in mammalian skeletal muscle (Figure 1A) but at the level of the center of the I-band in frog skeletal muscles (Peachey and Franzini-Armstrong, 1983; Franzini-Armstrong, 1994). The significance of this difference in the distribution of T-tubules is presently unknown. T-tubules are closely apposed and physically connected to a specialized domain of the SR called the terminal cisternae, thus forming a junctional complex called the triad (Figure 1B) (Peachey and FranziniArmstrong, 1983; Franzini-Armstrong, 1994). It follows that T-tubules themselves are composed of two distinct domains, namely the junctional and non- junctional T-tubular membranes (Franzini-Armstrong, 1975). The terminal cisternae of the sarcoplasmic reticulum are specialized to release Ca2+stored in their lumen via the feet structures (Franzini-Armstrong and Jorgensen, 1994) corresponding to the Ca2+-release channel/ryanodine receptor, a marker protein of junctional SR, (Fleischer and Inui, 1989; Franzini-Armstrong and Jorgensen, 1994). Feet structures protrude from the junctional face of the terminal cisternae bridging the narrow gap between the two membrane systems (Franzini-Armstrong and Jorgensen, 1994). Freeze fracture replicas of T-tubules from toadfish skeletal muscle show that integral membrane particles are distinctly distributed in the junctional and nonjunctional domains of T-tubules (Block et al., 1988).In the junctional domain, tetrads of particles are aligned in double rows. In contrast, integral membrane particles are not detected in the non-junctional region of T-tubules. Interestingly the position of every other foot structure in the junctional SR closely apposed to T-tubules corresponds precisely to that of a tetrad of integral membrane particles in the junctional T-tubule membrane. This observation strongly supports the hypothesis of E-C coupling proposing that the charge movement associated with the signal transduction event in the triad is accomplished by a direct or indirect physical interaction between a tetrad and a foot, respectively localized to junctional T-tubules and junctional SR (Rios et al., 1992; Schneider, 1994). It has been suggested that tetrads in T-tubules (Block et al., 1988) represent the 1,bdihydropyridine receptor/Ca*+-cha-

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nnel, a marker protein of T-tubules in adult skeletal muscle (Jorgensen et al., 1989; Flucher et al., 1990; Franzini-Armstrong and Jorgensen, 1994) essential for transmission of the signal elicited in T-tubules (Numa, et al., 1990; Rios et al., 1992) to the Ca2+-releasechannel/ryanodine receptor (feet) in the junctional SR (FranziniArmstrong and Jorgensen, 1994). Caveolae

Caveolae are small (80 x 90nm) flask-shaped invaginations of the surface SL (Dulhunty and Franzini-Armstrong, 1975; Zampighi et al., 1975). Some are single, others are arranged in linear and branched rows in the subsarcolemma with a single common opening to the sarcolemmal surface (Figures 1A and 1C). The density of caveolae openings to the surface SL varies between species (e.g., Dulhunty and Franzini-Armstrong, 1975;Bonilla et al., 1981).The average density of caveolae in frog sartorius (Dulhunty and Franzini-Armstrong, 1975) and human quadriceps muscles (Bonilla et al., 1981) are respectively, 37/pm2 and 18/pm2.Their total surfacearea is substantial, namely -60% of that of the surface SL in frog sartorius muscle (Dulhunty and Franzini-Armstrong, 1975). The finding that caveolae membrane (Dulhunty and Franzini-Armstrong, 1975; Bonilla et al., 198l), like nonjunctional T-tubules, contain very few intramembranous particles as compared to the surface SL (Franzini-Armstrong, 1975) suggests that caveolae and surface SL are designed to carry out distinct functions. Morphological infusion studies with electron dense markers suggest that all caveolae are continuous with the cell surface implying that skeletal muscle caveolae are static structures (Severs, 1988). This is in sharp contrast to caveolae in endothelial cells where morphological studies suggest that they are dynamic structures engaged in transcytosis (Severs, 1988). Various roles have been proposed for caveolae in skeletal muscle (Dulhunty and Franzini-Armstrong, 1975; Zampighi et al., 1975; Severs, 1988). A role for caveolae as connectors between T-tubules and surface SL is supported by ultrastructural studies showing that T-tubules frequently terminate in caveolae that are continuous with other caveolae and with the surface SL [Zampighi et al., 1975; FranziniArmstrong, 1986 (Fig. 13)] (Figures. 1A and 1C). This function is consistent with the apparent static nature of skeletal muscle caveolae. Furthermore, it has been difficult to demonstrate direct continuity between T-tubules and the surface SL of vertebrate skeletal muscle. Nonetheless physiological and morphological studies suggested that only a subpopulation of caveolae terminate in T-tubules thus raising the possibility that distinct populations of caveolae with distinct functions may be present in skeletal muscle (Zampighi et al., 1975).Alternatively,the interconnected caveolae providing continuity between T-tubules and surface SL could branch at the cell periphery and give rise to more than one caveola opening per T-tubule (Figure 1). This possibility is supported by images from thick sections of frog skeletal muscle showing that tortuous T-tubules close to the surface SL are often associated with a large number of caveolae (Franzini-Armstrong, 1986).

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Recent studies on the protein composition of caveolae in other cell types including endothelial-rich tissues and smooth muscle tissue suggest that caveolae are distinct multifunctional membrane domains engaged in various kinds of ion transport and signal transduction events (Anderson, 1993; Lisanti et. al., 1994). It remains to be determined whether there are one or more types of caveolae in skeletal muscle and, if so, whether they are engaged in one or more of the functions previously shown to be associated with caveolae in endothelial and smooth muscle cells. The low frequency of intramembranous particles in the nonjunctional T-tubules (Franzini-Armstrong, 1975) and caveolae (Dulhunty and Franzini-Armstrong, 1975) may reflect some common molecular characteristics of these two domains of the plasma membrane in skeletal muscle.

B.

Protein Composition of Transverse (T)-Tubules and Surface Sarcolemma (SL)

The surface SL and the T-tubules each constitute at most 5 to 10 percent, while the sarcoplasmic reticulum constitute approximately 90% of the total membrane fraction of skeletal muscle tissue. Combined with the fact that all the T-tubular (Lau et al., 1977; Rosemblatt et. al., 1981; Sabbadini and Dahms, 1989) and surface SL proteins (Seiler and Fleischer, 1988) appear to be minor components of their respective isolated membrane fractions have made it difficult to identify and characterize proteins unique to either T-tubules or the SL before the advent of monoclonal antibody technology. Nonetheless, to elucidate the molecular basis for the major function(s) of Ttubules in E-C coupling, several laboratories have developed procedures for the isolation and characterization of purified T-tubular membranes. (Lau et al., 1977; Rosemblatt et al., 1981; Sabbadini and Dahms, 1989). Generally two distinct approaches have been used. In one procedure T-tubular membrane vesicles are isolated from purified triads dissociated by a French press (Lau et al., 1977). Alternatively T-tubules can be obtained from calcium phosphate-loaded sarcoplasmic reticulum by sucrose density centrifugation (Rosemblatt et al., 1981). Both preparations are characterized by a high concentration of binding activity for 1,4dihydropyridine. However the content of other enzymatic activities considered to be markers of the plasma membrane (e.g. Na", K+-ATPaseand Mg2+-ATPase)differs considerably. The reasons for these differences are presently unknown. Examination of the protein composition of isolated T-tubular membrane fractions by SDS-polyacrylamide gel electrophoresis have generally shown that they contain about 15-25 proteins, most of which appear to constitute a small proportion of the total protein. Much effort has been made to characterize the major proteins in the T-tubular membrane fractions. However due to the fact that T-tubular membranes constitute a small proportion of the total membrane protein in skeletal muscle, conclusions regarding the uniqueness of the proteins in isolated T-tubules have been difficult to confirm by immunocytochemical labeling, because of the difficulty of

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obtaining sufficient amounts of purified protein for the production and characterization of specific antibodies. Similar difficulties were encountered in original attempts to identify membrane proteins specific to the surface SL. Approaches

The advent of hybridoma monoclonal antibody technology provided an alternative and powerful approach for the preparation of monoclonal antibodies specific to proteins present in either isolated T-tubular or surface SL membranes. In turn, immunocytochemical labeling of tissue sections with these probes can show whether or not a particular protein present in for example the T-tubular membrane fraction is indeed a component of T-tubules and if so whether or not it is specific for this membrane (e.g., Jorgensen et al., 1989; Flucher et al., 1990; Jorgensen et al., 1990; Ohlendieck et al., 1991). Furthermore, a monoclonal antibody can in principle be used to immunoaffinity purify and characterize the protein it identifies. In cases where the protein is a subunit of a hetero-oligomeric protein, the monoclonal antibody may also be used in the identification of the other subunits of this complex (e.g., Ervasti et al., 1990). Finally, monoclonal antibodies specific for either T-tubules or surface SL may be useful for the purification and characterization of the membrane domain in which its antigen resides (Ohlendieck et al., 1991). In the present review only proteins demonstrated by immunocytochemical labeling techniques to be preferentially distributed to either T-tubules or the surface SL of skeletal muscle fibers will be included. Protein Markers Of T-Tubules The ?,4-DihydropyridineReceptor (DHPR). The best characterized T-tubular specific protein is the 1,4dihydropyridinereceptor (DHPR) (Campbell et al., 1988; Catterall, 1988, 1991) believed to correspond to the tetrad detected in the junctional T-tubular membrane of adult skeletal muscle by in situ freeze-fracture studies (Block et al., 1988). The DHPR is essential for E X coupling (Numa et al., 1990; Rios et al., 1992). It is generally accepted that it functions as a “voltage sensor” gating the signal elicited by the depolarization of T-tubules and transmitted directly or indirectly to the foot structure (RR/Ca2+-releasechannel) in the closely apposed junctional SR which, in turn, releases Ca2+required for contraction (Rios et al., 1992). In contrast the L-type Ca2+-channelactivity of the DHPR does not appear to be an essential link in the E-C coupling pathway in skeletal muscle (Rios et al., 1992). The DHPR has been purified from skeletal muscle triads and T-tubular membrane fractions (Campbell et al., 1988; Catterall, 1988 and 1991). It is a hetero-oligomeric protein complex composed of equimolar amounts of four subunits called a , (170 kDa), a2/6(175 kDa), p (52 kDa) and y (32 kDa). Determination of the subcellulardistribution of subunits of the DHPR by immunofluorescence labeling of a,(Jorgensen et al., 1989), a2(Flucher et al., 1990), and

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p (Jorgensen et al., 1989), and by immuno-electronmicroscopical labeling of the a,-(Jorgensen et al., 1989; Flucher et al., 1990) and a,-subunits (Flucher et al., 1990) confirmed that the DHPR is indeed densely distributed in T-tubules and apparently absent from the surface SL. Unexpectedly the a,-subunit was also detected over some but not all subsarcolemmal vesicles (Jorgensen et al., 1989). It was proposed that these vesicles may correspond to a subpopulation of the caveolae prominently distributed in the subsarcolemma in skeletal muscle. Although one study reported that approximately 50% of the colloidal gold particles in the triad region might be in the non-junctional region of the T-tubules (Jorgensen et al., 1989) a subsequent immunoelectronmicroscopical study of 1-2 day old skeletal muscle (Jorgensen et al., 1995) strongly suggested that amajority of the DHPRs are confined to the junctional domain of T-tubules. In agreement with the immunolocalization studies summarized above, none of the subunits of the DHPR was detected by Western blotting of highly purified sarcolemmal vesicles obtained from skeletal muscle by selective fractionation by wheat germ agglutination (Ohlendieck et al., 1991). These highly purified surface SL vesicles contained high amounts of all the components of the dystrophin glycoprotein complex shown by imm.unolocalization to be specifically localized to the surface SL in skeletal muscle but not detected in interior regions where T-tubules are localized (Ohlendieck et al., 1991). The a,-subunit is essential for the function of the DHPR (Rios et al., 1992; Numa et al., 1990). Its cDNA sequence predicts a polypeptide of -212 kDa with four homologous domains each containing six transmembrane segments (Miller, 1994; Tanabe et al., 1988). Nonetheless two isoforms of the a,-subunit with M,s of 212 kDa and 170 kDa have been identified (De Jongh et al., 1989). In purified Ttubules the 170 kDa a,-subunit constitutes at least 90% of the total a,-subunit. The fact that only one mRNA of the a,-subunit has been identified, combined with immunochemical analysis of the two subunits suggests that the 170 kDa a,-subunit is formed by proteolytic cleavage of -300 amino acids from the COOH-terminal end of the 212 kD a,-subunit. It has been proposed that the 212 kDa and 170 kDa a,-subunits may respectively represent the Caz+-channeland voltage sensor functions associated with the DHPR. However transfection of dysgenic muscle cells that lack the a,-subunit, with a cDNA construct coding for the 170 kDa a,-subunit showed that this 170 kDa a,-subunit can perform both functions (Numa et al., 1990). These studies also predict that monoclonal antibodies that identify epitopes on the 17OkDa a,-subunit will alsoidentify the 212 kDa a,-subunit. Although distinct isoforms of the a,DHPR subunit are also present in other cell types including smooth and cardiac muscle, the similarity of the amino acid sequences of hydrophilic portions of these isoforms is low (Miller, 1994). The a,/6 -subunit is a heterodimer with an apparent molecular mass of 175 kDa (Campbell et al., 1988; Catterall, 1991; Miller, 1994). Both the a*- and the &-subunitsare encoded by a single gene that predicts a M,of 125 kDa (Catterall, 1988; Jay et al., 1991; De Jongh et al., 1990). az-and 6-subunits, respectively, form

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the N-terminal and C-terminal portions of the encoded peptide. Upon reduction the a,/b-subunit dissociates into the a,-peptide with an M, of 150 kDa and the &subunits composed of three peptides of 25,22, and 17 kDa (Catterall, 1988; Jay et. al., 1991). Both the a,-subunit and the &subunit are highly glycosylated. The three &peptides appear to have identical core peptides that are heterogeneously glycosylated. Although the presence of hydrophobic sequences in both the a,- and &peptides suggest that both peptides are transmembrane polypeptides (De Jongh et al., 1990), biochemical studies imply that the 6-peptide is the membrane anchor for the extracellularly located a,-subunit (Jay et al., 1991). Further analysis by, for example, immunofluorescence labeling of cultured muscle cells with site specific antibodies to various domains of each of the a,- and &peptides may provide more conclusive results regarding the topology of the various regions of this protein. The ability of the a2/6-subunitto enhance the expression of the a,-subunit when coexpressed in L-cells implies a functional interaction between the a,- and the a2/6-subunits.Proteins very similar to the a,/&-subunit in skeletal muscle are present in a large number of tissues (Miller, 1994). The cDNA of the @-subunitpredicts a 50 kDa protein without a transmembrane segment (Ruth et al., 1989). Recent molecular biological studies show that the P-subunit binds to a conserved motif on the cytoplasmic loop between repeats I and I1 of the a,-subunit (Pragnell et al., 1994). The finding that mutations in this motif decrease the ability of the P-subunit to modulate the chemical activity of the a,-subunit (e.g., stimulation of peak currents) implies an essential role for the P-subunit in the function of the DHPR. It was previously shown that co-expression of the P- and a,-subunits in stably transfected L-cells dramatically increased the DHP binding sites as well as the peak Ca2+current implying that P-subunits enhanced the targeting of a,-subunits to the plasma membrane (Snutch and Reiner, 1992). However studies by Neeley et al., (1993) showed that although the peak currents were increased the tail currents remained the same in the presence and absence of co-expression of the P-subunit (Neely et al., 1993). These results imply that the P-subunit modulates the activity of the a,-subunit by inducing a conformational change as opposed to increasing the number of the a,-subunits targeted to the SL. The y subunit. Its cDNA sequence predicts a 25 kDa (32 kDa) hydrophobic polypeptide that contains four transmembrane domains and two N-linked glycosylation sites (Jay et al., 1990). These findings are consistent with biochemical studies showing that this subunit is a glycosylated hydrophobic polypeptide. Its function is presently unknown. Interestingly, it has only been detected in skeletal muscle (Miller, 1994). The potential roles of they subunit in for example the assembly, targeting and function of DHPR remains to be explored. TS 28. Defined by mAb IXEII,, TS28 is a minor protein component of Ttubular membranes from rabbit skeletal muscle. It has an apparent molecular mass of 28 kDa (Jorgensen et al., 1990).

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Immunofluorescence and immunoelectronmicroscopical studies with mAb IXEII, (Jorgensen et al., 1990) showed that the TS28 like the subunits of the DHPR is mainly localized to T-tubules while only a small amount is localized to some subsarcolemmal vesicular structures possibly corresponding to the subgroups of caveolae connecting the T-tubules with the sarcolemma in skeletal muscle. In contrast TS28 was not detected over the surface SL. However subsequent immunoelectron microscopical studies of skeletal muscle in 1-2 day old rabbits where focal triads are interconnected by extensive stretches of nonjunctional T-tubules clearly showed that TS28 in contrast to the alsubunit of the DHPR is densely distributed in the nonjunctional domain of T-tubules (Jorgensen et al., 1995). Since junctional and nonjunctional domains of T-tubules are in close vicinity to one another it is difficult by this approach to exclude the possibility that TS28 is present in the junctional domain of T-tubules. In conclusion, the immunocytochemical studies showed that TS28 and the a,-a*-, and P-subunits of the DHPR are localized to Ttubules but not detected in the surface SL. They also show that TS28 is localized to the nonjunctional domain of T-tubules while the a,-DHPR is preferentially localized to the junctional domain of T-tubules. TS28 is a minor 28 kDa integral membrane protein present in isolated Ttubular membrane vesicles (Ohlendieck et al., 1991; Jorgensen et al., 1990) and not detected by immunoblotting of highly purified sarcolemmal membrane vesicles ( Ohlendieck et al., 1991). This protein does not copurify with the DHPR nor does an antibody to TS28 bind to any of the subunits of the DHPR. Although the function of TS28 is presently unknown, it has been proposed that it corresponds to a 28 kDa protein in isolated T-tubular membrane vesicles that is a substrate of an endogenous protein kinase C (Salvatori et al., 1993). It has also been proposed that TS28 corresponds to a 28 kDa GTP-binding protein enriched in isolated T-tubular membranes (Doucet and Tuana, 1991). It remains to be determined whether TS28 indeed corresponds to either of these two proteins. The protein has been immunoaffinity purified (Mason and Jorgensen, manuscript in preparation). Biochemical characterization of the purified TS28 as well as cloning and sequencing of the cDNA of TS28 should provide clues regarding these possibilities. Protein Kinase C. It has recently been reported on the basis of immunocytochemical studies that protein kinase C is densely distributed in regions of the muscle fiber corresponding to the location of T-tubules (Salvatori et al., 1993). This finding combined with the observation that the activation of a protein kinase C endogenous to isolated T-tubular membranes resulted in the phosphorylation of several T-tubular proteins including a 28 kDa protein lead the authors to propose that this 28 kDa protein may represent the 28 kDa protein called TS28 (Jorgensen et al., 1990)shown to be localized to the nonjunctional domain of T-tubules in sifu (Jorgensen et al., 1995) and not detected in the surface SL (Jorgensen et al., 1990). Further studies will be required to determine if this is indeed the case.

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MgL+-ATPase. On the basis of biochemical and immunocytochemical studies it has been reported that a Mg2+-ATPaseof 82 kDa is a specific component of skeletal T-tubules, where it appears to be specifically associated with the junctional Ttubules (Sabbadini and Dahms, 1989). Surprisingly recent comparison of the amino acid sequence deduced from the cDNA sequence of the Mg2+-ATPaseto known cDNA sequences showed the sequence of the presumptive Mg2+-ATPaseto be identical to that of T-cadherin (Cunningham et al., 1993; Ranscht and DoursZimmermann, 1991). The likelihood that T-cadherin is a component of T-tubules would seem slim in view of the fact that cadherins are supposed to function as cellcell adhesion molecules. Further studies will be required to resolve this apparent discrepancy. Protein Markers o f the Surface Sarcolernrna Three Distinct ffetero-oligornericTransmembrane Cytoskeletal Complexes are Unique to the SurfaceSarcolernrna (SU. Ultrastructural studies showing that myofibrils underlying the sarcolemma are uniquely attached to the surface SL (Pierobon-Bormioli, 1981) predicted that surface SL proteins involved in this attachment would be specific for the surface SL and not detected in T-tubules. So far a majority of the proteins demonstrated by immunolocalization to be uniquely associated with the surface SL and not detected in T-tubules can indeed be considered to be components of one of three well characterized hetero-oligomeric transmembrane protein complexes engaged in distinct plasma membrane cytoskeleton interactions (Luna and Hitt, 1992). Two of these are transmembrane complexes that bridge membrane cytoskeletal components to the extracellular basement membrane via transmembrane proteins. They include: (1) the integrin-actin filament based complex characteristic of the cell-matrix adhesion site in cultured cells called focal contact (Figure 2A) (Luna and Hitt, 1992; Burridge et al., 1988; Simon et al., 1991); and (2) the dystrophin-glycoprotein complex characteristic of the surface SL of striated muscle (Figure 2B) (Ervasti and Campbell, 1993; Tinsley et al., 1994). The third complex is the spectrin-actin filament based cortical cytoskeleton present in most eucaryotic cells (Bennett, 1990a,b; Bennett and Lambert, 1991; Luna and Hitt, 1992). In red cells and epithelial cells the spectrin-based cortical cytoskeleton anchor specifically interacts with certain integral membrane proteins of the plasma membrane thereby inhibiting the lateral diffusion of these proteins in the plane of the lipid bilayer (Figure 2C). Specific aspects of the structure, function, interaction, and subcellular distribution of components of each of these transmembrane protein complexes in skeletal myofibers and other cells and tissues have been detailed in previous reviews (e.g., Burridge et al., 1988; Bennett, 1990a,b; Bennett and Lambert, 1991; Simon et al., 1991; Luna and Hitt, 1992; Ervasti and Campbell, 1993a; Tinsley et al., 1994). Here only features of each of these complexes that illustrate (1) their unique distribution i n the surface SL, (2) their contribution to the

A

F-nttio

./

\

B

F-A&

Fkai.

Figure 2. (A)Diagrammatic model showing the major protein Components of the integrin-actin filament based complex of focal contacts in cultured cells and their proposed interactions with each other and the extracellular matrix components of the substratum. (Adapted from Luna and Hitt, 1992.) (B)Diagrammatic representation of the Components of the dystrophin-glycoprotein complex, their interactions with each other, with components of the extracellular matrix and with cortical F-actin. (Adapted with permission from Matsumura and Campbell, 1994.) (C)Diagrammatic representation of some of the components of the spectrin-ankyrin based cytoskeletal complex and its interaction with transmembrane proteins of the red cell membrane. (Adapted from Luna and Hitt, 1992.) 171

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172

C

Glycophorin C

/

Anion exchs

Figure 2 continued

polarity of the skeletal muscle cell surface, (3) their potential usefulness as probes to study the processes involved in the biogenesis and maintenance of the polarized cell surface in skeletal muscle, and (4) the potential interaction between the distinct hetero-oligomeric protein complexes are discussed. A diagrammatic representation of the topology and interaction of some of the protein components of each of these hetero-oligomeric transmembrane complexes are presented in Figure 2. lmmunofluorescence Studies Localize Components of All Three Transmembrane Cytoskeletal Complexes To Costameres. The term costamere was coined by Craig and collaborators (Craig and Pardo, 1983; Pardo et. al., 1983a) who were the first to show that vinculin and y-actin (both components of the integrin-based hetero-oligomeric transmembrane complex) co-distribute with each other and with spectrin (Craig and Pardo, 1983) in transversely oriented “hoops” located at the cytoplasmic subdomain of the SL overlying the Z-line domains of peripheral myofibrils. They furthermore proposed that costameres represent the subsarcolemmal cytoskeletal network previously observed by ultrastructural studies to link the surface SL (Figure 1C) to the Z-line domains of the underlying (peripheral) myofibrils (Pierobon-Bormioli, 1981). More recent immunofluorescence studies using confocal microscopy have suggested that costamere associated proteins, (e.g., dystrophin, vinculin, and P-spectrin) are arranged in a two-dimensional grid-like network composed of alternating “heavy” and “light” sub-sarcolemmal “hoops” respectively associated with domains of the surface SL overlying the Z- and M-lines of peripheral myofibrils (Porter et al., 1992) (Figure 1C). The grid-like arrangement is accomplished by rnterconnecting the transversely oriented “hoops” by longitudinally oriented narrow strands.

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The observation that the distribution of costamere proteins, as imaged by immunofluorescence confocal microscopy, correspond quite closely to the distribution of the filaments bridging myofibrils to domains of the surface SL overlying the Z- and M-lines of peripheral myofibrils (Pierobon-Bormioli, 198l), supports the idea that the components of the three transmembrane cytoskeletal complexes and the filamentous structures bridging myofibrils to the SL are integral components of costameres. lmmunoelectron Microscopy: Are Transmembrane-Cytoskeletal Complexes Preferentially Localized to Costameres ? Although immunoelectron microscopi-

cal studies by Shear and Bloch (1985) showed that vinculin (a component of the integrin-based complex) is densely distributed in electron dense structures associated with domains of the sarcolemma overlying the Z-lines in skeletal muscle, other immunoelectromicroscopical (IEM) studies (Geiger et al., 1980; Jorgensen and Shens unpublished observations) did not detect vinculin over costameres in cardiac muscle as predicted by immunofluorescence studies (Pardo et al., 1983b), even though the same IEM studies showed vinculin to be densely distributed in the submembranous domains of the adherens junctions of the intercalated disc in the same fiber. Furthermore immunoelectronmicroscopical studies with antibodies to dystrophin (Watkins et al., 1988; Cullen et al., 1990; Byers et al., 1991) and to the dystrophin associated protein called 43 DAG (Cullen et al., 1994) did not show a preferential distribution of these proteins in domains overlying the Z-line and the M-line of peripheral myofibril as predicted on the basis of results from immunofluorescence localization of these and other proteins. Rather it appeared as if these proteins were fairly uniformily distributed over the surface sarcolemma. Further studies will be required to resolve the apparent discrepancy between the distribution of dystrophins and the 43 DAG as observed at the light and electron microscopical levels of resolution. The apparent absence of vinculin at costameres as suggested by some immunoelectron microscopical studies might be the result of steric hindrance. Thus it will be important to use antibodies to different epitopes of vinculin. The Integrin-Actin Filament-Based Complex. The proposed topology and interactions of components of the integrin-actin filament based complex in focal contacts have been reviewed in detail elsewhere and are presented diagrammatically in Figure 2A (Burridge et al., 1988; Simon et al., 1991; Luna and Hitt, 1992). Only a brief overview is presented here as an introduction to the presence of these components in skeletal muscle. Integrins are members of a large family of heterodimeric transmembrane proteins each composed of an a-and a P-subunit (Hynes, 1992). There are numerous isoforms of both subunits specific for particular cell types and subcellular distributions. In focal contacts integrin mediates interaction between extracellular matrix proteins (e.g. fibronectin and laminin) and components of a dense cytoplasmic plaque into which bundles of actin filaments insert. In vitro stud-

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ies have demonstrated that integrin interacts with the cytoplasmic plaque proteins talin and a-actinin. Talin in turn has distinct domains for interaction with vinculin and actin filaments while a-actinin interacts with actin filaments. It is generally believed that the interaction between the cytosolic tail of the 0-subunit of integrin and the plaque proteins talin and a-actinin does not occur until integrin molecules cluster in response to binding to their extracellular ligand (e.g., fibronectin or laminin). This in turn leads to the assembly of additional attachment proteins including vinculin thus linking integrin to cortical actin filaments. The relatively dense distribution of integrin (Volk et al., 1990; Knudsen and Horwitz, 1994); talin (Belkin et al., 1986), vinculin (Porter et al., 1992; Craig and Pardo, 1983) and y-actin (Craig and Pardo, 1983) over costameres in skeletal muscle as determined by immunolocalization studies is consistent with the idea that they are components of an integrin-actinfilamentbased transmembranecomplex in this domain of the sarcolemma, analogous to the well characterized cell-matrix adhesion site in cultured cells called focal contacts (Figure 2A). A role for an integrin-actin filament based transmembranecomplex in the attachment of myofibrils to the sarcolemma is strongly supported by studies on Dmsophila embryos containing either normal and/or mutant versions of the P,-subunit of costamere specific integrin showing that fibronectin induced clustering of integrin is essential for sarcomere alignment and possibily for the subsequent lateral attachment of myofibrils to costameres in developing muscle (Volk eta]., 1990)(see section below on the Integrin-Actin Filament BasedComplex, p. 21 1). Since it is well established that focal contacts in cultured cells represent sites of signal transduction (Hynes, 1992; Romer et al., 1992), it is reasonable to anticipate that the presumptive integrin-actin filament based complex in costameres in adult skeletal muscle in addition to playing a potential role in the attachment of myofibrils to the SL may also play important roles in signal transduction events. In this regard it is noteworthy that a study on cultured cardiac muscle cells showed that costameres are sites of transmembrane force transduction during muscle contraction (Danowski et al., 1992). The Dystrophin-Clycoprotein Complex. The dystrophin-glycoprotein complex (DGC) is a large hetero-oligomeric complex that spans the surface SL and provides a direct physical bridge between laminin in the basement membrane and y-actin filaments of the subsarcolemmal cortical cytoskeleton in skeletal muscle (Ervasti and Campbell, 1993; Tinsley et al., 1994)(Figure 2B). Identificationand characterizationof h s protein complex in skeletal muscle began with the identification of dystrophin (Hoffman et al., 1987; Anderson and Kunkel, 1992;Worton, 1994,reviews), the protein encoded by the Duchenne Muscular Dystrophy gene and the demonstration that this protein is localized to the surface SL but not detected over T-tubules in normal adult skeletal muscle in situ (Matsumura and Campbell, 1994; review). Subsequent biochemical studies demonstrated that dystrophin is tightly associated with a protein complex composed of at least six sarcolemmal membrane proteins (Ervasti and Campbell, 1993;Tinsley et al., 1994; reviews). In contrast

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to the integrin-actinfilament based complex described above and the spectrin-basedcytoskeleton described in the next section, the dystrophin-glycoprotein complex has been purified as an intact complex from the sarcolemma of skeletal muscle (Ervasti et al., 1990;YoshidaandOzawa, 1990,ErvastiandCampbell, 1991;Ohlendiecketal., 1991). This complex contains a 156 kDa glycoprotein (1 56 DAG) called a-dystroglycan, a 59 kDa triplet protein (59 DAP) called syntrophin (a,, b, and P2), a SO kDa glycoprotein (50 DAG) called adhalin, a 43 kDa glycoprotein (43 DAG) called P-dystroglycan,a 35 kDa glycoprotein (35 DAG), and a 25 kDa protein (25 DAP). The proposed topology and interactions of the components of the dystrophin-glycoproteincomplex in relation to each other and the sarcolemma is illustratedin Figure 2B. The 50 DAG (adhalin),the 43 DAG (P-dystroglycan), the 35 DAG and the 25 DAP are transmembrane proteins. The 59 DAP triplet is like dystrophin, an extrinsicprotein on the cytoplasmic side of the sarcolemma; while the a-dystroglycan (156 DAG) is an extracellular glycoprotein shown to bind to laminin (Ibraghlmov-Beskrovnayaet al., 1992),a component specific to the basal lamina. The studies documenting the molecular and functional characteristics of this complex has been reviewed (Ervasti and Campbell, 1993a; Tinsley et al., 1994; Campbell, 1995). Furthermore, dystrophin has been purified from highly purified surface SL of skeletal muscle and shown to represent about 5% of the total membrane protein (Ervasti et al., 1991). Immunolocalization studies confirmed that the dystrophin associated glycoproteindprotein (156 DAG, 59 DAP, SO DAG, 43 DAG and 35 DAG) like dystrophin arc densely distributed at the sarcolemma but are not detected in interior regions of the muscle fibers where T-tubules are located (Ervasti et al., 1990; Jorgensen et al., 1990; Ervasti and Campbell, 1991; Ohlendick et al., 1991). Confocal imaging of immunofluorescently-labeled cryosections from skeletal muscle suggested that dystrophin is preferentially localized to domains of the surface SL corresponding to the distribution of costameres (Porteret al., 1992;Masuda et al., 1992; Straub et al., 1992). It remains to be explored whether the dystrophin associated proteins also localize to costamere domains of the surface SL. Immunoelectron microscopical studies with antibodies to dystrophin (Watkins et al., 1988; Cullen et al., 1990; Byers eta]., 1991) and to the dystrophin associated protein, 43 DAG (Cullen et al., 1994) confirmed that they are confined to the sarcolemma. However they did not show a preferential distribution of these proteins in domains overlying the Z-line and the M-line of peripheral myofibril as predicted on the basis of results from the immunofluorescence localization studies of dystrophin. Rather it appeared as if these proteins were fairly uniformily distributed over the surface SL. The explanation of this apparent discrepancy between the distribution of dystrophin and the 43 DAG as observed at the light and electron microscopical levels of resolution is presently unknown but might in part be caused by the relatively low intensity of labeling observed by immunoelectron microscopy. Dystrophin is a cytoskeletal protein that contains four structurally distinct domains (Ervasti and Campbell, 1993a; Tinsley et al., 1994): (1) The NH,-terminal domain contains regions homologous to some actin-binding proteins including

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a-actinin and P-spectrin, (2) a series of 24 repeats, (3) a cysteine rich domain, and (4) the COOH-terminal domain that lacks homology to previously reported peptide sequences. It has been shown that the NH,-terminal domain of dystrophin can bind to actin (Ervasti and Campbell, 1993b) (Figure 2B), while the cysteine rich domain and part of the COOH-terminal domain are essential for dystrophin interaction with the glycoprotein complex (Ervasti and Campbell, 1993a). The finding that dystrophin molecules lacking the carboxyl-terminal end still localize to the SL raises the possibility that dystrophin associates with the SL by binding to y-actin of the cortical membrane cytoskeleton (Ervasti and Campbell, 1993a). a- and P-dystroglycans (156 DAG and 43 DAG, respectively) are products of a single gene that encodes a 97 kDa precursor protein (Ibraghimov-Beskrovnaya et al., 1992). In agreement with the previous biochemical studies the NHy terminal portion of this precursor is processed into a 156 kDa glycoprotein that lacks transmembrane domains, while the COOH-terminal domain is processed into a 43 kDa glycoprotein containing a single transmembrane domain. a-Dystroglycan binds to laminin. Northern and Western blotting analyses have demonstrated that the a-and P-dystroglycans are expressed in a wide variety of muscle and non-muscle tissues. The cDNA sequence of adhalin (50 DAG) predicts a protein with 387 amino acids, a signal sequence, one transmembrane domain and two potential N-linked glycosylation sites (Roberds, et al., 1993). In contrast to a- and P-dystroglycan, adhalin is only expressed in skeletal and cardiac muscles and in some but not all smooth muscles (Roberds, et al., 1993). Sequencing and expression of the cDNA show that the syntrophin (59 DAP) triplet is heterogeneous (Yang et al., 1994). Sequencing of the cDNA of the 35 DAG and the 25 DAP remains to be determined. With respect to future studies aimed at probing the targeting of SL specific proteins to their special domains on the surface SL, it is of particular interest that immunochemical and immunocytochemical studies suggest that the 35 DAG (Yamamoto et al., 1994) like adhalin (Roberds et al., 1993; Yamamoto et al., 1994) is mainly confined to striated (cardiac and skeletal) muscles. The Spectrin-Ankyrin-based Membrane Cytoskeletal Complex. The spectrinankynn-based membrane cytoskeletalcomplex was first identifiedand characterizedas a complex of proteins associated with the cytoplasmic side of the red cell membrane (Figure 2C) (Bennett, 1990a,b; Bennett and Lambert,1991; reviews). Spectrin is the major protein of this complex which also includes ankynn (Bennett, 1992),the band 4.1 protein, short y-actin filaments and several other actin binding proteins. It is generally accepted that the spectrin-ankyrin-basedcytoskeleton stabilizes the red cell membrane since its dissociationfrom the membrane leads to vesiculation of the strippedredcell membrane. Spectrin is a heterodimeric protein (a,P) that self-assembles head-to-head. The formed tetramers interact indirectly tail-to-tailvia the band 4.1 protein, y-actin filaments and other proteins to form the two-dimensional network that constitutes the spectrinankynn-based membrane cytoskeleton of the red cell membrane. The spectrin-

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ankyrin-based cytoskeleton is attached to two integral membrane proteins namely band 3 protein (the anion exchanger) and the band 4.1 protein. These attachments in turn inhibit lateral diffusion of these integral membrane proteins in the lipid bilayer. The presence of spectrin-like and ankyrin-like proteins in most cell types including skeletal muscle have been known for some time (Small et al., 1992; Bennett, 1990a,b; Bennett and Lambert, 1991). Immunofluorescence localization studies of skeletal muscle have generally demonstrated that spectrin-like (aand p) (Craig and Pardo, 1983; Repasky et al., 1982; Nelson and Lazarides, 1983) and ankyrin-like components (Nelson and Lazarides, 1984) are localized to costamere domains of the sarcolemma and not detected in internal regions of the fiber where T-tubules are located. One study reported that ankyrin was detected in domains corresponding to the distribution of T-tubules in adult chicken skeletal muscle (Flucher et al., 1990). This was surprising since spectrin has not been detected in this domain. However recent molecular biological studies have shown that certain repeats of amino acid sequences of ankyrin are also detected in proteins otherwise unrelated to ankyrin (Liou and Baltimore, 1993). Thus, it will be important to determine whether the molecular characteristics of this ankyrin-like protein localized to the T-tubular domain of skeletal muscle indeed represents ankyrin or whether it corresponds to a protein that is functionally unrelated to ankyrin. Molecular characterization of the spectrin-like (Small et al., 1992; Vybiral et al., 1992; Winkelmann et al., 1990) and ankyrin-like proteins (Birkenmeier et al., 1993) in skeletal muscle by immunoblotting and/or Northern blotting imply that there are several distinct isoforms. The significance of these distinct isoforms remains to be determined. Although the function of spectrin in various cell types including skeletal muscle may vary, it is likely that it plays a role in anchoring certain integral membrane proteins to subdomains of the plasma membrane. However, so far integral membrane proteins anchored to the spectrin-ankyrin-based cytoskeleton of the surface SL have not been identified. Interestingly, the Na+/K+ATPase is detected in both the sarcolemma and T-tubules in skeletal muscle (Ohlendieck et al., 1991) and apparently not anchored by the spectrin-based cytoskeleton confined to the sarcolemma in this cell. This is in contrast to some epithelial cells where anchoring of the Na+/K+-ATPaseby the spectrin-ankyrin-based cytoskeleton is proposed to lead to the accumulation of this protein in the basolateral domain (Nelson et al., 1990), even though it is incorporated with equal efficiency into the apical and basolateral domains of epithelial cells (e.g., MDCK) (Hammerton et al., 1991). The identification of cDNAs unique to 8-spectrin (Winkelmann et al., 1990) and to 3 distinct ankyrins (Birkenmeier et al., 1993) in skeletal muscle suggest that a better understanding of the structure, function, and subcellular distribution of these two proteins as well as their potential interaction with each other and with integral membrane proteins of the skeletal sarcolemma is forthcoming. The lntegrin-700A Filament Based Complex: Is It a Component of Costameres?

Hemidesmosomes are small circular domains (diameter: 0.4 pm) of the plasma

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membrane in some stratified squamous epithelia (Schwarz et al., 1990). Like focal contacts they attach the cell to the basement membrane. They contain an integrin (c(,P,)-IOOA filament transmembrane complex (Stepp et al., 1990; Kurpakus et al., I99 1). Immunofluorescence labeling studies have shown that desmin, and other subunits of 100 A filaments (Small et al., 1992; Lazarides, 1978; Craig and Pardo, 1983) besides their localization to the interfibrillar space at the level of Z-disk are also localized to costamere domains in adult cardiac muscle. This finding points to the possibility that lOOA filaments may be anchored to costamere domains of the sarcolemma via integrins. It remains to be determined whether this is indeed the case. This possibility is of considerable interest in view of a potential role for vinculin and desmin in the assembly of costamerzs proposed to link the surface SL to the Z- and Mline domains of the myofibrils underlying the sarcolemma (see p. 21 1 and Figure 5). Do the Three Transmembrane Cytoskeletal Complexes in Costameres Interact with Each Other! Costameres were originally reported as vinculin-containing “hoops” localized to the cytoplasmic side of the subsarcolemma at the level of Z-lines and proposed to bridge the Z-lines of the subsarcolemmal myofibrils tg the sarcolemma (Pardo et al., 1983a). One study showed that dystrophin co-localized with vinculin and P-spectrin (muscle specific) not only at the Z-line domain of the sarcolemma but also at the level of M-lines as well as in occasional longitudinally oriented strands thus forming a 2-D network in the subsarcolemma (Porter et al., 1992). Since vinculin co-distributes with dystrophin at all three sites, it was proposed that the distribution of dystrophin corresponds to the distribution of costameres in skeletal muscle. However, another study found that dystrophin and vinculin co-localized in some but not all of the dystrophin-containing domains (Masuda et al., 1992). The significance of the co-distribution of the components of the three distinct transmembrane cytoskeletal complexes is presently unknown but might imply interactions between the three systems. However, it was clearly shown that the accumulation of P-spectrin and vinculin in constameres occurs independently of the presence of dystrophin (Porter et al., 1992). Thus the absence of dystrophin in, for example mdx mice did not alter the distribution of either spectrin or vinculin. It remains to be determined whether dystrophin-associated proteins codistribute with dystrophin in the costameres.

IV.

DEVELOPMENT OF A POLARIZED CELL SURFACE MEMBRANE IN SKELETAL MUSCLE

The present knowledge of the cellular processes responsible for the biogenesis and maintenance of a polarized cell surface membrane in skeletal muscle is sparse compared to our knowledge of those involved in the establishment and maintenance of the apical and basolateral domains of the plasma membrane in epithelial cells (e.g. Rodriguez-Boulan and Powell, 1992; Matter and Mellman, 1994; reviews).

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Nonetheless, recent studies using immunocytochemical and molecular biological approaches to probe the temporal appearance and subcellular distribution of surface SL and T-tubules specific proteins in relation to those of ultrastructural and physiological hallmarks of muscle development have provided some insights regarding the sequence of some of the events leading to a polarized cell surface in developing skeletal muscle. Furthermore, the availability of mutants to some of the T-tubular and surface SL specific proteins have recently provided interesting clues regarding the potential role of particular proteins in the biogenesis and maintenance of a polarized cell surface in skeletal muscle. So far the temporal appearance and subcellular distribution of proteins unique to either surface SL or to T-tubular membranes have only occasionally been compared to each other in developing muscle (Yuan et al., 1990; Flucheret al., 1991b;Flucher, 1992).Rather the spatial-temporal distribution of T-tubular proteins have mainly been compared to those ofjunctional SR proteins and to that of ultrastructural hallmarks such as the appearance of Ttubules and triads (Flucher, 1992; Franzini-Armstrong and Jorgensen, 1994), while the spatial-temporal appearance of sarcolemmal costamere proteins have mainly been correlated with those of cross-striated myofibrils and spontaneous contractile activity in developing muscle (Morris and Fulton, 1994; Volk et al., 1990). In an attempt to correlate current knowledge of the temporal appearance and subcellular distribution of unique T-tubular proteins to those of unique surface SL proteins, the data available has, when feasible, been related either to each other or to at least one of the following morphological and physiological hallmarks of muscle development outlined below. This data is summarized in Figure 3. The chosen hallmarks are presented below in the sequence in which they appear during muscle development. They include the onset of (1) fusion of myoblasts into myotubes, ( 2 ) T-tubule formation, ( 3 ) triad formation, (4) myofilament attachment to ends of myotubes, (5) appearance of cross-striated myofibrils (alignment of neighboring myofibrils), (6) spontaneous contractile activity, and (7) costamere organization. A.

Morphological Aspects

Temporal Appearance of Caveolae, T-Tubules and Triads

The formation of functional myotubes requires the division, differentiation and subsequent fusion of mononucleated presumptive myoblasts into multinucleated myotubes (Hauschka, 1994). T-tubules are not detected until after the onset of fusion (Kelly, 1971; Ishikawa, 1968). Furthermore, their first appearance in early myotubes is preceded by the appearance of peripheral junctional complexes between the SL and junctional SR called peripheral E-C coupling units (FranziniArmstrong and Jorgensen, 1994) and by a gradual increase in the number of caveolae observed to be arranged either in clusters or as short beaded tubular structures at the cell periphery (Kelly, 1971). The finding that both caveolae and T-tubules were infused by electron dense markers supported the proposal that caveolae are at first

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STRUCTURAL ANC PHYSIOLOGICAL HALLMARKS

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MIDDLE STAGE

LATER STAGES

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DYSTFtORIIN GLYCOPROTEIN COMPLD:

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Figure 3. Tentative correlation of the temporal appearance and subcellular distribution of T-tubular and costamere proteins with the temporal appearance of some structural and physiological hallmarks of skeletal muscle development. References: 1. Jorgensen and Arnold, unpublished results. 2. Yuan, 1991; Flucher et al., 1991.3. Yuan etal., 1990,1991. 4. Yaun et al., 1990. 5. Takemitsu et al., 1991; Zhao et al., 1993; Clerk et al., 1993. 6. Hagiwara et al., 1989; Prelle et al., 1991;Park-Matsumotoet al., 1991;Heiman-Pattersonet al., 1990; Miranda etal., 1998; Klamutetal., 1989. 7. Volketal., 1990.8. Morris and Fulton, 1994 (reviews the data on the assembly of cytoskeletal costamere proteins). 180

181

Cell Surface in Skeletal Muscle

initiation sites for T-tubule assembly and subsequently remain as connectors between the sarcolemma and the T-tubules. At first, forming T-tubules are oriented parallel to the long axis of the fibers and in close vicinity to the SL (Kelly, 1971). Shortly after the first appearance of short T-tubular invaginations focal internal junctional complexes between T-tubules and junctional SR were observed (early triads). As development proceeded, T-tubules were observed to branch, and gradually become transversely oriented while the number and size of focal junctional complexes increased dramatically (Kelly, 1971) (Figure 3). Models of J-Tubule Biogenesis

Originally two models for the assembly of T-tubules were proposed on the basis of ultrastructural studies of muscle developing in culture. The “add-on” model proposed that the formation of T-tubules occurs by sequential fusion of newly formed membrane vesicles with caveolae (Schiaffino et al., 1977). The “swept-in” model proposed that T-tubules form by continuous extension of short sarcolemmal membrane invaginations (Ishikawa, 1968). In the latter model it is proposed that the increase in area of the cell surface membrane occurs by random fusion of newly formed membrane vesicles with the sarcolemma. Temporal Appearance of Cross-striated Myofibrils and Spontaneous Contractility

Following onset of fusion of muscle cells developing in culture early myofibrils begin to form and attach end-on to small domains of the SL at the end of forming myotubes. Since sarcomeres of neighboring early myofibrils are not yet in register, these developing myofibers do not appear cross-striated when imaged by phasecontrast microscopy. As development proceeds lateral alignment of sarcomeres in neighboring myofibrils is now reflected as cross-striated appearance of myofibers. Appearance of cross-striations precedes the onset of spontaneous contractions (Cossette and Vincent, 1991). The onset of both events precedes the earliest detection of cytoskeletal costamere proteins at the the I-band domains of the SL in developing skeletal muscle (Figure 3) (reviewed in Moms and Fulton, 1994).

B.

Molecular Approaches

The availability of antibodies and cDNAs to protein markers of T-tubules, junctional SR and surface SL has enabled the use of immunofluorescence and immunoelectron microscopical labeling as well as in situ hybridization to begin to delineate the sequence of events leading to the accumulation of these proteins and in some cases their respective mRNAs at their respective target membrane domains during the de novo formation of T-tubules, triads and costameres and thus to the development of a polarized skeletal muscle surface membrane. The availability of

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spontaneous and designed mutant animals lacking particular proteins unique to either T-tubules or to the surface SL has also been useful in beginning to delineate some of the key events leading to the accumulation of T-tubular and surface SL proteins into their target domains. Temporal Appearance of T-Tubular Proteins TS28. Immunofluorescence labeling has demonstrated that TS28 so far is the only T-tubular specific protein uniformly distributed in nonjunctional domains of Ttubules from the onset of their de novo biogenesis in skeletal muscle developing in situ (Yuan, et al., 1990) (Figure 4A) and in culture (Wilson et al., 1992). Since TS28 is not detected in the surface SL by immunofluorescence labeling at anytime before or during T-tubule formation, it was proposed that TS28 is an inherent component of a subset of Golgi derived transfer vesicles that according to the add-on model of Ttubule biogenesis (Schiaffino et al., 1977) initiate this event by first fusing with preformed caveolae at the cell periphery followed by sequential fusion of TS28 containing transfer vesicles to this domain (Yuan, et al., 1990) (Figure 4A). However since this study determined the steady-state distribution of TS28 in developing skeletal muscle, it remains to be determined whether the specific accumulation of TS28 in forming T-tubules is accomplished by direct targeted delivery of TS28 to T-tubules as they form or whether TS28 is delivered uniformly to both T-tubules and the SL but preferentially retained at T-tubules, as demonstrated for the Na+/K+ATPase in some epithelial cells (e.g., MDCK) (Hammerton et al., 1991; Nelson et al., 1990). The first possibility would correspond to the add-on model proposed by Schiaftino et al., (Schiaffino et al., 1977). The second alternative would correspond to the original pull-in model proposed by Ishikawa (Ishikawa, 1968). A third distinct model for T-tubule formation has been proposed by Flucher et al (Flucher et al. 1991b) on the basis of similar studies with a polyclonal antibody (prepared to purified T-tubular membranes) that specifically labels T-tubules in adult skeletal muscle. This probe labels an internal network in both myoblasts and myotubes before the appearance of T-tubules continuous with the surface SL. This finding suggests that first a closed internal T-tubular network is formed that subsequently fuses with the surface SL to form an open T-tubule system. It remains to be determined whether such a closed internal T-tubular network is detected before the onset of T-tubular formation in developing skeletal muscle in situ. The potential function of TS28 in the assembly of T-tubules is presently unknown. The cloning, sequencing and availability of TS28-cDNA should greatly facilitate studies designed to determine the structure and function of this protein, including its potential role in the biogenesis of T-tubules. ?,4-Dihydropyridine Receptor (DHPR). In contrast to TS28 (a T-tubular protein), the a,-subunit of the 1,4-dihydropyridine receptor (a,-DHPR) accumulates in discrete foci at the cell periphery before the onset of T-tubule formation (Yuan et al.,

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A

Figure 4. (A)Model for the assembly of SL50 and TS28 into sarcolemma and transverse tubules. First, SL50 begins to be synthesized on membrane-bound polysomes and then i s transported to the subsarcolemmal region ofthe myotubes (Step 1).Next, SL50-containing vesicles fuse with the sarcolemma, resulting in, first, a nonuniform distribution of SL50 (step 2). Then the SL50 quickly diffuses laterally into the lipid bilayer of the sarcolemma, where it remains uniformly distributed during all subsequent stages of myofiber development (step 3). As this continues to occur (step 41, TS28 synthesized on membrane-bound polysomes is transported to the subsarcolemmal region in transfer vesicles (step 5). To initiate the formation of a particular transverse tubule, a TS28-containing vesicle (step 6) fuses with the sarcolemma, forming a TS28-containing caveola (step 7). However, TS28 in the lipid bilayer of the caveola is somehow prevented from lateral diffusion into the lateral portion of the sarcolemma. Subsequently, additional TS28-containing transfer vesicles (step 8) fuse first with the TS28-containing caveola (step 7), forming short tubular invaginations in the subsarcolemmal region (step 9). Repeated fusions of TS28-containing transfer vesicles with the short TS28-containing tubular invaginationsresult in the formation of further extended transverse tubules (step 10). These are, at first, oriented parallel to the long axis in the subsarcolemmal region of the myotubes (not shown). Eventually they become organized into a transversely oriented chickenwire-like network surrounding the myofibrils present throughout the cytosol. (Diagram and legend reprinted with permission from Jorgensen et al., 1990). (B) One plausible scheme for the biogenesisof T-tubules and triads in developing skeletal muscle. First, a,-DHPR (Al) and ryanodine receptors (BI) are assumed to be synthesized on membrane-bound polyribosomes, incorporated into unique transfer vesicles, and distributed (A2 and 82, respectively).Next, TS28-containingvesicles (C1) fuse firstwith the sarcolemma to form a caveolae (C2). Sequential fusions of TS28-containingvesiclesto the caveolae result in the formation of a tubular invagination into the myotube (C3-C5).As this occurs, ryanodine receptor-containing vesicles are incorporated into forming SR (B3), followed by complex formation between ryanodine receptors in SR and the a,-DHPR-containing transfer vesicles (AB4) in the outer zone of the cytosol. In turn, the a,-DHPR-containing vesicle of this complex fuses with the TS28-containing forming T-tq-bules (ABCG),thus, incorporatingthe a,-DHPR-containingvesiclesinto a discrete region of the forming T-tubules (ABC7) and forming a junctional complex between the T-tubule and the SR. (Diagram and Legend reprinted with permissionfrom Yuan et al., 1991.)

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B

S K E L E r A L MUSCLE

Figure 4 continued

1991) (Figure 4B). Since the a,-DHPR co-distributes with the ryanodine receptor (RR; a marker of the junctional SR) (Yuan et al., 1991) from the onset of its first appearance in cultured myoblasts (Jorgensen and Arnold, manuscript in preparation), it is likely that at least some of these foci correspond to peripheraljunctional complexes transiently present at the SL before the onset of formation of T-tubules.However, this has not yet been shown directly by immunoelectronmicroscopicallabeling with antibodies that identifies extracellular epitopes of for example the a , - and a,-subunits of the DHPR. Nonetheless, freeze fracture studies have shown that tetrads believed to represent DHPRs are clustered on the SL and possibly represent peripheral couplings (Takekura et al., 1994; Franzini-Armstrong et al., 1991). Furthermore, studies by Marks et a1 (Marks et al., 1991) showed that the appearances of the mRNAs of the DHPR and the ryanodine receptor in a non-fusing differentiating muscle cell line are coincident with each other and with the appearance of feet structures at peripheral junctional complexes. Since “primitive”junctional complexes that lack feet (ryanodine receptors) are present prior to the induction of the synthesis of ryanodine receptor and the DHPR, newly synthesized ryanodine receptor and DHPR may be incorporated directly into these structures, thus forming a peripheral junctional complex. Alternatively, newly synthesized ryanodine receptor and DHPR may be assembled during the de novo assembly of an upgraded form of junctional complex. Once T-tubules begin to form, some of the DHPR and ryanodine receptor containing foci co-distribute with discrete domains of T-tubules labeled for TS28 indicating that these foci may represent internal couplings between T-tubules and junctional SR (Yuan et al., 1991). The finding that some of these foci are present further into the cytoplasm where T-tubules are not detected suggested that these foci represent ‘triad precursors’ proposed to be composed of two closely apposed vesicles, one of which contains the DHPR while the other contains the RR (Figure 4B). It was further hypothesized that the DHPR containing vesicles would become

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incorporated into discrete domains of T-tubules and as a result of the association of each DHPR containing foci with a ryanodine receptor containing membrane structure, form an internal junctional complex where TS28 and the DHPR are respectively localized to the nonjunctional and junctional domain of forming T-tubules and triads (Figure 4B). If correct, this hypothesis implies that TS28 and the DHPR are sorted and transported to T-tubules in distinct vesicles. Alternatively, the ryanodine receptor and the DHPR could first become incorporated into SR and the surface SL/T-tubules, respectively, at a density too low to be detected by immunofluorescence labeling. The appearance of DHPR and ryanodine receptor containing foci would then exclusively represent peripheral and internal complexes formed as spontaneous clustering of the ryanodine receptor in the SR closely apposed to either surface SL or T-tubules, would induce clustering of DHPR thus forming a junctional complex (Franzini-Armstrong and Jorgensen, 1994). Regardless of the pathway leading to the accumulation of the a,DHPR first in peripheral couplings and after the onset of T-tubule formation into triads, a mechanism for targeting the ryanodine receptor containing SR or the triad precursor to Ttubules and not to the surface SL is required. It will be interesting to explore the possibility that TS28 and potentially associated proteins may play a role in this targeting process. It was originally proposed that the DHPR is an essential link in the formation of junctional complexes (Yuan et al., 1991).However similar studies of dysgenic (Flucher et al., 1991a) and crooked neck skeletal muscle (Airey et al., 1993a,b)that respectively lacks the a,-DHPR and the p-isoform of the ryanodine receptor while expressingclose to the normal amount of other SR proteins (Airey et al., 1993a,b;Knudson et al., 1989) suggested that neither the a,-DHPR nor the ryanodine receptor is essential for the formation of junctional complexes and have provided some insights into the role of the a,-DHPR and the ryanodine receptor in the assembly of triads. Ultrastructural studies indicate that dysgenic muscle in situ and in culture contain a normal complement of peripheral couplings that lack tetrads while only a few internal couplings are detected (Pincon-Raymond et al., 1985; Franzini-Armstrong et al., 1991). Furthermore foci immunolabeled for the ryanodine receptor are also detected in some dysgenic myotubes (Flucher et al., 1991a). Since some of these foci are oriented transversely in relation to the longitudinal axis of the myotubes, it was proposed that they represent internal couplings (triads) lacking the DHPR. Similarly a study on crooked neck muscle reported a normal peripheral distribution of a,-DHPR in the absence of p ryanodine receptorrequired for EC coupling (Airey et al., 1993a,b). These studies support the conclusion that the accumulation of either DHPRs or ryanodine receptors in discrete cytoplasmic foci possibly representing peripheral and/or internal junctional complexes does not require either the ryanodine receptor or the DHPR, respectively. However since there are fewer internal complexes at least in dysgenic fibers, it has been proposed that the presence of the a,-DHPR may be important for the stability of this complex. Interestingly the studies in dysgenic muscle also showed that the a,-DHPR is essential for targeting

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the a,-subunit of the IP-DHPR to the RR containing foci (Flucher et al., 1991a). The possibility that a cardiac like a,-DHPR subunit may substitute for the skeletal a,-DHPR at least in peripheral couplings is supported by the finding that mRNA of the a,-DHPR is prominently expressed in normal and dysgenic muscle cultures at early stages of development (Chaudhari and Beam, 1993). The finding that the ryanodine receptor and the DHPR can each accumulate in discrete clusters in the cytoplasm in the respective absence of the DHPR and the ryanodine receptor, is consistent with the idea that both proteins are “passive passengers” in the organelle(s) transporting them to the site of incorporation into T-tubules and SR, respectively, and that other components of these transport vesicles target the assembly of these vesicles into triad structures regardless of the presence of the ryanodine receptor and the DHPR. The observation that the ryanodine receptor and the DHPR containing foci codistribute from the onset of their appearance in mononucleated myoblasts and myotubes suggest that their assembly into triad structures occur simultaneously. Whether or not it occurs by simultaneous fusion of distinct or associated DHPR and ryanodine receptor containing vesicles with junctional SR and T-tubular membranes respectively remains to be determined. However the ability of the two molecules to cluster independently would seem to exclude the model proposing that clustering of the ryanodine receptor in the SR induce clustering of DHPR uniformly distributed in SL and T-tubules (Franzini-Armstrong and Jorgensen, 1994). Irrespective of which model is correct, the almost exclusive presence of internal junctional complexes (triads) after the onset of formation of T-tubules, strongly points to the need for a mechanism that targets junctional SR and/or triad precursors to forming T-tubules and not to the SL where junctional complexes form before T-tubules begin to form. In view of a recent proposal for mechanisms responsible for targeted vesicle fusion by Rothman and colleagues (Sollner et al., 1993a,b; Rothman and Warren, 1994), it is tempting to speculate that T-tubule specific proteins such as TS28 may function as receptors for this purpose. Alternatively the finding that T-tubule formation is preceded by the specific accumulation in the sarcolemma of adhalin (previously called SL50, Yuan et al., 1990), a muscle specific integral membrane protein of the dystrophin glycoprotein complex, raises the possibility that components of one or more of the membrane cytoskeleton systems specifically localized to the sarcolemma may directly or indirectly play a role in the preferential formation ofjunctional complexes (triad) at T-tubules by, for example, inhibiting the apposition of junctional SR to the sarcolemma. Assembly of Sarcolemmal Protein Markers (Costamere Proteins) into Costameres

So far the temporal appearance and subcellular distribution of only a small number of costamere proteins have been examined during muscle development. Representative results of these studies are presented in Figure 3. Collectively they show that accumulation of some costamere proteins (vimentin, desmin, vinculin) in

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costamere domains at the surface SL (the domains overlying the Z-lines of peripheral myofibrils) does not begin in a particular myotube until sarcomere alignment (cross striated appearance of myofibers observed by phase contrast microscopy) and spontaneous contractile activity have occurred in this fiber (Cossette and Vincent, 1991;Morris and Fulton, 1994). However, other studies using muscle cultures from Drosophilu embryos with normal and mutant versions of the costamere specific isoform of P-integrin imply that fibronectin induced clustering of costamere specific integrin is essential for the alignment of sarcomeres in neighboring myofibrils (Volk et al., 1990). Although a solution to this apparent riddle is not yet at hand, two sets of experiments presented below provide clues regarding the sequence of some of the key events and cellular processes involved in the targeting and assembly of costamere proteins into costameres at the surface SL (Figure 5).

The Integrin-Actin Filament Based Complex. Several studies have demonstrated that interaction between integrin and their extracellular ligands (e.g., fibronectin) play key roles in various aspects of myogenesis including the end-on and lateral attachment of myofibrils to the SL (Knudsen andHorwitz, 1994). It has been reported that integrin clusters at costameres and at myotendenous junctions where Z-lines and ends of myofibrils are respectively attached laterally (Figure 1C) and end-on to the SL in adult skeletal muscle (Knudsen and Horwitz, 1994). During

~ 1 liilegrin ) randomly distributed in the SL

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Fibronectin induced integrin clustering

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(3)Assembly of early integrin-actin J filament -a-actinin complexes randomly distributed in the SL ?

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D)* Interaction between periphernl components of t

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E)*

of'oo 'lamentsubunits (eg. vimentin, desmin) and other components of the integrin-actin filament complex (eg.vinculin) can begin

It i s presently unknown whether step D precedes or follows step E

Figure 5. A plausible diagrammatic model for the sequence of some of the events leading to the assembly of early costameres (see text for detailed description of model).

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muscle development in Drosophilu clustering of integrin occurs at “myotendenous junctions” (end-on attachment sites for myofibrils) before they occur at domains of the surface SL above the Z-line domains of peripheral myofibrils (site of forming costameres?) (Volk et al., 1990). (a) Clustering of costamere-specific integrins: A series of studies on Drosophilu embryo muscle cells with normal and mutant (null) versions of costamerespecific P-integrin showed that fibronectin induced clustering of costamere specific integrin is essential for the alignment at Z-lines in neighboring myofibrils and possibly for the lateral attachment of myofibrils to costameres during the development of skeletal muscle (Volk et al., 1990). This essential role of fibronectin was supported by the finding that sarcomere alignment and clustering of costamere specific integrin (diffusely distributed in sarcolemma) at the sarcolemma did not occur in normal myotubes unless fibronectin was added to the culture medium. Interestingly, a laminin substrate was sufficient for fusion to occur and for integrin to cluster at the sites of the SL where end-on attachment of myofibrils occur. The essential requirement for integrin in the fibronectin dependent sarcomere alignment and possibly the subsequent lateral attachment of myofibrils to the SL was demonstrated in muscle cells from a null mutant of Drosophilu that lacks the costamere-specific p-integrin and fails to align sarcomeres. Other preliminary results described in the same study (Volk et al., 1990) indicated that an a-actinin null mutant of Drosophilu also fails to form and align sarcomeres but do form fibronectin induced integrin clusters. However, the integrin clusters are randomly distributed at the SL, thus implying that Z-disc components of peripheral myofibrils may play arole in the alignment of integrin into costamere domains of the surface SL (Z-disc positions). A similar study with a C. eleguns (nematode) null mutant for vinculin showed that vinculin is essential for muscle contraction. However since the absence of vinculin also prevented the end-on attachment of myofibrils to the SL, the potential role of vinculin in sarcomere alignment and the attachment of myofibrils to costameres cannot be discerned from this study (Barstead and Waterston, 1991). Combined, the results from the Drosophilu mutant experiments summarized above (Volk et al., 1990) would indicate that random clustering of integrin occurs independently of sarcomere alignment and that alignment of integrin clusters over Zline domains of the sarcolemma (costameres) may be dictated by the position of Zlines in aligned myofibrils (Figure 5). Possibly the alignment of integrin clusters is accomplished by physical interaction between components associated with Z-lines of peripheral myofibrils and components of randomly distributed integrin clusters in the SL, thereby anchoring the randomly clustered integrins to the costamere domains of the surface SL. However, the sarcomere alignment must occur first. The studies summarized above point to the possibility that fibronectin induced clustering of costamere specific p-integnn is essential for the alignment of sarcomeres. The mechanism(s) involved in this process remains to be identified. However, since focal contacts are sites for transmembrane signal transduction (Hynes, 1992; Romer et al.,

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1992), it is plausible that fibronectin induced clustering of integrin may lead to a signal tranduction event that produces a yet unknown signal or a series of signals leading to the alignment of sarcomeres including Z-lines of neighboring myofibrils. Clues regarding some of the cellular processes enabling Z-line domains of the aligned myofibrils to dictate the alignment of integrin clusters and the assembly of associated cytoskeletal proteins into the integrin-actin filament based transmembrane complexes at costamere domains are provided by the results of experiments summarized below. The current knowledge regarding the temporal appearance of cytoskeletal costamere proteins in developing skeletal muscle in culture and in situ was reviewed by Morris and Fulton (1994). Briefly, detailed examination of the temporal appearance and subcellular distribution of vinculin (a component of the integrin-actin filament based transmembrane complex) and other cytoskeletal costamere proteins at Z-lines by double immunofluorescencelabeling showed that sarcomere alignment and spontaneous contractions precede the first detection of several cytoskeletal costamere proteins at costamere domains of the surface SL of chick skeletal muscle cells developing in culture (Morris and Fulton, 1994; Cossette and Vincent, 1991). This implies that these cytoskeletal costamere proteins including vinculin are not required for sarcomere alignment (Morris and Fulton, 1994). Furthermore, they show very clearly that a highly ordered process is responsible for the assembly of vinculin and other cytoskeletal costamere proteins into the costamere domains of the surface SL (Figure 3). Thus vimentin (a subunit of lOOA filaments) was the first protein detected at the domain of the surface SL overlying I-bands and at the periphery of Z-discs ( Cripe et al., 1993; Morris and Fulton, 1994). Next, desmin appeared at these domains (Fulton, 1993). Subsequently vimentin disappeared as vinculin clustered at costamere domains, while desmin remained at costamere domains of the SL and at the periphery of Z-discs ( Cripe et al., 1993;Morris and Fulton, 1994). Although the temporal appearance of integrin was not examined in the above studies, it is anticipated on the basis of the studies by Volk et al. (1990) that costamere specific clusters of integrin appear in the sarcolemma before vinculin appears since sarcomere alignment and spontaneous contractility precedes this event. However, it would be important to determine if this is indeed the case.

(b) Targetingof cytoskeletalproteins to sites of costamere assembly: How are cytoskeletal costamere proteins targeted to their site of assembly? Since vinculin, vimentin and desmin are “cytosolic” proteins it is generally believed that they are released into the cytosol from free polysomes and then diffuse to their site of assembly. Alternatively recent studies by Fulton and colleagues support the idea that these costamere proteins are assembled co-translationally into their target domains of the surface SL (Cripe et al., 1993; Morris and Fulton, 1994), since it was shown that mRNAs of each of these costamere proteins first appeared shortly after their

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corresponding proteins and codistributed with their corresponding proteins in the I-band domain (Morris and Fulton, 1994, see Figure 3). Relevant to understanding how for example vimentin mRNA is targeted to the site of vimentin assembly, it has been demonstrated that mRNAs coding for several distinct cytoskeletal proteins accumulate at the site of assembly of the respective proteins, suggesting that co-translateral assembly occurs in several types of cells (Morris and Fulton, 1994; Singer, 1992; review). The mechanism responsible for targeting cytoskeletal costamere proteins to their site of assembly is presently unknown. However it has been proposed that the nascent chain of these proteins carry a target sequence (Singer, 1992). Alternatively recent studies (Kisilauskis et al., 1993, 1994) have shown that mRNAs encoding distinct actin isoforms accumulate at the assembly site of the particular isoform they encode. These studies also showed that mRNAs of distinct actin isoforms are targeted to their particular site of assembly by the 3'-untranslated region of the corresponding mRNA. This is a very attractive and testable concept that could explain in part how cytoskeletal costamere proteins accumulate at the site of their assembly. Undoubtedly, other mechanisms also play a role in this complicated process. The molecular nature of the targets for the assembly of each of the cytoskeletal costamere proteins/mRNAs and in particular the target of the first component to accumulate at the costamere domains remains to be determined. It is plausible that a physical link between clusters of integrin and Z-line components is established as soon as sarcomere alignment has occurred. On the basis of demonstrated interactions in vitro between integrin and associated cytoskeletal proteins (Figure 2A), it is tempting to speculate that muscle a-actinin (an integral component of Z-lines) is instrumental in anchoring integrin clusters possibly via a non-muscle a-actinin-y -actin linker into costamere domains of the surface SL (Figure 5). This proposed series of events also supports the idea that the position of the Z-line and I-bands of contracting myofibrils dictate the position of costamere assembly which in turn may dictate targeting of other costamere components as suggested by Morris and Fulton (1994).

(c)A plausible model of costamereassembly: Assuming that targeting and assembly of components of the integrin-actin filament based complex into costameres occur in a similar manner in developing skeletal muscle of most species including Drosophila and chicken the following events are tentatively proposed as steps in its assembly on the basis of the studies summarized above and on current concepts regarding the composition and interactions of proteins in focal contacts of cultured cells (Burridge et al., 1988; Simon et al., 1991; Luna and Hitt, 1992; Romer et al., 1992) (Figure 5). First, costamere specific integrin is incorporated into the surface SL of newly formed myotubes (Figure 5; Step A l ) while nonaligned myofibrils accumulate in the cytoplasm (Figure 5, A2). Interaction of the costamere specific isoform of integrin with extracellular fibronectin leads to the formation of randomly distributed clusters of integrin in the sarcolemma (Figure 5; B). Since T-tubules lack fi-

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bronectin these clusters are presumably confined to the surface SL. The event of integrin clustering is aprerequisite for the yet unknown signal or series of signals, that induce the alignment of sarcomeres (Figure 5 ; Cl) in newly formed myofibrils already attached end-on to the sarcolemma. Presumably the event of integrin clustering also leads to the onset of the assembly of the integrin-actin filament based complex (Figure 5 ; C2). In this scheme it is proposed that only nonmuscle a-actinin and y-actin filament association with clustered integrin would be required. The alignment of sarcomeres is in turn essential for the anchoring of integrin-actin filament based complexes into the costamere domains of the surface SL. Most likely the position of the Z-line domains of the aligned myofibrils dictates the site of accumulation of the integrin-actin filament based complex (Figure 5 ; D). This possibility is supported by the finding that the synthesis and accumulation of vinculin and possibly other components of the integrin-actin filament based complex are targeted to costamere domains of the surface SL (Figure 5 ; E). It is proposed that once the cytoskeletal components of this complex essential for the formation of a physical link between the Z-disc and integrin clusters assemble at this site, their interaction with each other and components of the Z-disc (e.g., muscle a-actinin) will lead to the alignment of clustered integrins at the position of the costamere domains of the surface SL (i.e., domains of SL overlying Z-lines). It is presently unknown whether the accumulation of vinculin and lOOA filament subunits at costamere positions precedes or follows the proposed formation of a physical link between myofibrils and the surface SL. This model is clearly highly speculative and is only intended as a plausible framework for further studies designed to determine when and how costamere proteins assemble to form the proposed physical link between myofibrils and the surface SL.

Dystrophin Clycoprotein Complex. Some studies have examined the temporal appearance and subcellular distribution of components of the dystrophin-glycoprotein complex in developing muscle. However since the different components of this complex have so far not been compared to each other or to other markers of the surface SL in the same experimental setting, it is at present difficult to correlate the temporal appearance and distribution of subunits of the dystrophin-glycoprotein complex to those of other costamere proteins presented above. Thus an extensive review of this area awaits additional correlative studies. Comparison of the temporal appearance and subscellular distribution of adhalin (50 DAG; called SL 50 in the first study) to those of TS28 (a marker of T-tubules) during skeletal muscle development in situ (Yuan et al., 1990) showed that adhalin is first detected in the SL of early myotubes before the onset of T-tubule formation (Figure 4A) which presumably precedes the onset of sarcomere alignment and spontaneous contractile activity in developing muscle cells (Figure 3). Following the appearance of T-tubules and at all subsequent stages of development adhalin appeared to be confined to the SL and not detected in interior regions of developing muscle where T-tubules are located. It is presently unknown whether adhalin is

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sorted to the surface SL or becomes anchored there by interaction with for example cytoskeletal and/or extracellular components of the dystrophin-glycoprotein complex. However, since the cDNA of adhalin has been cloned and sequenced (Roberds et al., 1993), it should be feasible to begin to identify domains of this glycoprotein essential for its targeting and accumulation in the surface SL. Immunolocalization studies have shown that the dystrophin-like molecule called utrophin is detected in the SL of mouse and human muscle early on during muscledevelopment (Takemitsuetal., 1991;Clerket al., 1993;Zhaoetal., 1993). It remains in the SL until the onset of nerve innervation after which utrophin is confined to the neuromuscular junction in normal innervated skeletal muscle. Immunolocalization of dystrophin in skeletal muscle developing in situ (Hagiwara et al., 1989; Prelle et al., 1991) and in culture (Miranda et al., 1988; Klamut et al., 1989; Heiman-Patterson et al., 1990; Park-Matsumoto et al., 1991) have shown that dystrophin is at first diffusely distributed in early myotubes and only gradually becomes associated with the SL first in a patchy distribution but later appears to be homogeneously distributed in the surface SL. Similar studies of aneural and innervated human skeletal muscle cells developing in culture suggested that contractile activity play an essential role in the homogeneous distribution of dystrophin in the surface SL (Park-Matsumoto et al., 1991). Conclusions regarding the assembly of the utrophin/dystrophin-glycoprotein complex and its anticipated accumulation into costameres in developing muscle awaits further correlative studies.

ACKNOWLEDGMENTS Results reported from my laboratory were supported by grants in-aid from the Medical Research Council of Canada.

REFERENCES Airey, J.A., Baring, M.D., Beck,C.F.,Chelliah, Y., Deerinck,T.J., Ellisman, M.H., Houenou, L.J., McKemy, D.D., Sutko, J.L. & Talvenheimo, J. (1993a). Failure to make normal a ryanodine receptor is an early event associated withthe crookedneckdwarf(cn)mutation inchicken.Dev. Dynamics 197,169-188. J.L.,McKemy,D.D. Airey, J.A., Deerinck,T.J., Ellisman,M.H.,Houenou,L.J.,Ivanenko,A.,Kenyon, & Sutko, J.L. (1993b). Crooked neck dwarf(cn) mutant chicken skeletal muscle cells in low density primary cultures fail to express normal a ryanodine receptor and exhibit a partial mutant phenotype. Dev. Dynamics 197, 189-202. Anderson, M.S. & Kunkel, L.M. (1992). The molecular and biochemical basis of Duchenne Muscular Dystrophy. Trends in Biochem. Sci. 17,289-292. (Review) Anderson, R.G.W. (1993). Caveolae: Where incoming and outgoing messengers meet. Proc. Natl. Acad. Sci. USA 90, 10909-10913. (Review) Barstead, R.J. & Waterston, R.H. (1991). Vinculin is essential for muscle function in the nematode. J. Cell Biol. 114(4), 715-724. Belkin, A.M., Zhidkova, N.I. & Koteliansky, V.E. (1986). Localization of talin in skeletal and cardiac muscle. FEBS Letters 200(1), 32-36.

Cell Surface in Skeletal Muscle

193

Bennett, V. (1990a). Spectrin-based membrane skeleton: a multipotential adaptor between plasma membrane and cytoplasm. Physiol. Rev. 70(4), 1029-1065. (Review) Bennett, V. (1 990b). Spectrin: astructural mediator between diverse plasma membrane proteins and the cytoplasm. Curr. Opin. Cell Biol. 2(1), 51-56. (Review) Bennett, V. (1992). Ankyrins: Adaptors between diverse plasmamembrane proteins and the cytoplasm. J. Biol. Chem. 267(3), 8703-8706. Bennett, V. & Lambert, S. (1991). The spectrin skeleton: From red cells to brain. J. Clin. Invest. 87, 1483-1489. (Review) Birkenmeier, C.S., White, R.A., Peters, L.L., Hall, E.J., Lux, S.E. & Barker, J.E. (1993). Complex patterns of sequence variation and multiple 5' and 3' ends are found among transcripts of the erythroid ankyrin gene. J. Biol. Chem. 268(13), 9533-9540. Block, B.A., Imagawa, T., Leung, A,, Campbell, K.P., Franzini-Armstrong, C. (1988). Structural evidence for direct interaction between the molecular components of the transverse tubuleslsarcoplasmicreticulum junction in skeletal muscle. J. Cell Biol. 107,2587-2600. Bonilla, E., Fischbeck, K. & Schotland, D.L. (1981). Freeze-fracture studies of muscle caveolae in human Muscular Dystrophy. Am. J. Pathol. 104, 167-173. Burridge, K., Fath, K., Kelly, T., Nuckolls, G. & Turner, C. (1988). Focal adhesions: transmembrane junctions between the extracellular matrix and the cytoskeleton. Annu. Rev. Cell Biol. 4, 487-525. (Review) Byers, T.J., Kinkel, L.M. & Watkins, S.C. (1991). The subcellular distribution of dystrophin in mouse skeletal, cardiac, and smooth muscle. J. Cell Biol. 115(2), 41 1 4 2 1 . Campbell, K.P. (1995). Three muscular dystrophies: loss of cytoskeleton-extracellularmatrix linkage. Cell 80(5), 675-679. (Review) Campbell, K.P., Leung, A.T. & Sharp, A.H. (1988). The biochemistry and molecular biology of the dihydropyridine-sensitivecalcium channel. TINS 11(10), 425430. (Review) Catterall, W.A. (1988). Structure and function of voltage-sensitive ion channels. Science 242(4875), 5@-61. (Review) Catterall, W.A. (1991). Excitation-contraction coupling in vertebrate skeletal muscle: a tale of two calcium channels. Cell 64, 871-874. (Review) Chaudhari, N. & Beam, K.G. (1993). mRNA for cardiac calcium channel is expressed during development of skeletal muscle. Dev. Biol. 155, 507-515. Clerk, A., Morris, G.E, Dubowitz, V., Davies, K.E. & Sewry, C.A. (1993). Dystrophin-related protein, utrophin, in normal and dystrophic human fetal skeletal muscle. Histochem. J. 25, 554-561. Cormack, D.H. (1993). Muscle tissue. In: Essential Histology, pp. 215-232. Lippincott, Philadelphia. Cossette, L.J. & Vincent, M. (1991). Expression of a developmentally regulated cross-linking intermediate filament-associated protein (IFAPa-400) during the replacement of vimentin for desmin in muscle cell differentiation. J. Cell Sci. 98(2), 251-260. Craig, S.W. & Pardo, J.V. (1983). Gamma actin, spectrin, and intermediate filament proteins colocalize with vinculin at costameres, myofibril-to-sarcolemma attachment sites. Cell Motility 3, 449-462. Cripe, L., Morris, E. & Fulton, A.B. (1993). Vimentin mRNA location changes during muscle development. Proc. Natl. Acad. Sci. USA 90,2724-2728. Cullen, M.J., Walsh, J., Nicholson, L.V.B. & Harris, J.B. (1990). Ultrastructural localization of dystrophin in human muscle by using gold immunolabelling. Proc. R. SOC.Lond. B 240, 197-210. Cullen, M.J., Walsh, J. & Nicholson, L.V.B. (1994). lmmunogold localization of the 43-kDa dystroglycan at the plasma membrane in control and dystrophic human muscle. Acta Neuropathol. 87,349-354. Cunningham, H.B., Yazaki, P.J., Domingo, R.C., Oades, K.V., Bohlen, H., Sabbadini, R.A. & Dahms, A S . (1993). The skeletal muscle transverse tubular Mg-ATPase: identity with Mg-ATPases of smooth muscle and brain. Arch. Biochem. Biophys. 303(1), 3 2 4 3 .

194

ANNELISE 0. JORCENSEN

Danowski, B.A., Imanaka-Yoshida, K., Sanger, J.M. & Sanger, J.W. (1992). Costameres are sites of force transmission to the substratum in adult rat cardiomyocytes. J. Cell Biol. 118(6), 1411-1420. De Jongh, K.S., Merrick, D.K. & Catterall, W.A. (1989). Subunits of purified calcium channels: a 212-kDa form of alpha 1 and partial amino acid sequence of a phosphorylation site of an independent beta subunit. Proc. Natl. Acad. Sci. USA 86( l), 8585-8589. De Jongh, K.S., Warner, C. & Catterall, W.A. (1990). Subunits of purified calcium channels. Alpha 2 and delta are encoded by the same gene. J. Biol. Chem. 265(25), 14738-14741. Doucet, J-P. & Tuana., B.S. (1991). Identification of low molecular weight GTP-binding proteins and their sites of interaction in subcellular fractions from skeletal muscle. J. of Biol. Chem. 266(26), 17613-1 7620. Dulhunty, A.F. &Franzini-Armstrong,C. (1975). The relativecontributionsofthe folds andcaveolaeto the surface membrane of frog skeletal muscle fibres at different sarcomere lengths. J. Physiol. 250,513-539. Eisenberg, B.R. (1983). Quantitative ultrastructure of mammalian skeletal muscle. In: Handbook of Physiology, Section 10, pp. 73-1 12. Peachey, L.D., Adrian, R.H. &Geiger, S.R. eds., American Physiological Society, Bethesda, Maryland. (Review) Engel, A.G. (1994). The neuromuscular junction. In: Myology: Basic and Clinical, 2nd Edition, Volume 1. pp. 261-302. Engel, A.G. & Franzini-Armstrong, C. eds., McGraw-Hill, Inc. New York. (Review) Ervasti, J.M. &Campbell, K.P. (1993a). Dystrophinand the membrane skeleton. Curr. Opin. Cell Biol. 5, 82-87. (Review) Ervasti, J.M. & Campbell, K.P. (1993b). a role for the dystrophin-glycoprotein complex as a transmembrane linker between laminin and actin. J. Cell Biol. 122(4), 809-823. Ervasti, J.M., Ohlendieck, K., Kahl, S.D., Gaver, M.G. & Campbell, K.P. (1990). Deficiency of a glycoproteincomponentofthedystrophincomp1exindystrophicmuscle.Nature 345,315-319. Ervasti, J.M., Kahl, S.D. & Campbell, K.P. (1991). Purification of dystrophin from skeletal muscle. J. Biol. Chem. 26,9161-9165. Fleischer, S. & Inui, M. (1989). Biochemistry and biophysics ofexcitation-contractioncoupling. Annu. Rev. Biophys., Biophys. Chem. 18,333-364. (Review) Flucher, B.E. (1992). Structural analysis of muscle development: transverse tubules, sarcoplasmic reticulum, and the triad. Dev. Biol. 154,245-260. (Review) Flucher, B.E., Morton, M.E., Friehner, S.C. &Daniels, M.P. (1990). Localization ofthe alpha, and alpha, subunits of the dihydropyridine receptor and ankyrin in skeletal muscle. Neuron 5,339-351. Flucher, B.E., Phillips, J.L. & Powell, J. (1991a). Dihydropyridine receptor alpha subunits in normal and dysgenic muscle in vitro: expression of alpha, is required for proper targetingand distribution of alpha,. J . Cell Biol. 115, 1345-1356. Flucher, B.E.,Terasaki, M., Chin, H.M., Beeler, T.J. &Daniels, M.P. (1991 b). Biogenesis oftransverse tubules in skeletal muscle m vitro. Dev. Biol. 145, 77-95. Franzini-Armstrong, C. (1975). Membrane particles and transmission at the triad. Fed. Proc. 34(5), 1382-1 389. (Review) Franzini-Armstrong, C. (1980). Structure of sarcoplasmic reticulum. Fed. Proc. 39(7), 2403-2409. (Review) Franzini-Armstrong, C. (1986). The sarcoplasmic reticulum and the transverse tubules. In: Myology: Basic and Clinical, (Engel, A.G. & Banker, B.O., Eds.). McGraw-Hill, New York. (Review) Franzini-Armtrong, C. (1994). The sarcoplasmic reticulum and the transverse tubules. In: Myology: Basic and Clinical, (Engel, A.G. & Franzini-Armstrong, C. Eds.), 2nd Ed., Vol. I, pp. 176-199. McGraw-Hill, New York. (Review) Franzini-Armstrong, C. & Jorgensen, A.O. (1994). Structure and development of E-C coupling units in skeletal muscle. Annu. Rev. Physiol. 56, 509-534 (Review)

Cell Surface in Skeletal Muscle

195

Franzini-Armstrong, C. & Porter, K.R. ( I 964). Sarcolemmal invaginations constituting the T-system in fish muscle fibers. J. Cell Biol. 22,675-696. Franzini-Armstrong,C., Pincon-Raymond, M. & Rieger, F. (1991). Muscle fibers from dysgenic mouse in vivo lack a surface component of peripheral couplings. Dev. Biol. 146, 364-376. Fulton, A.B. (1993). Spatial organization ofthe synthesis of cytoskeletal proteins. J. of Cell. Biochem. 52, 148-152. (Review) Geiger, B., Tokuyasu, K.T., Dutton, A.H. & Singer, S.J. (1980). Vinculin, an intracellular protein localized at specialized sites where microfilament bundles terminate at cell membranes. Proc. Natl. Acad. Sci. USA 77(7),4127-4131. Hagiwara, Y., Yoshida,M.,Nonaka,I. &Ozawa,E. (1989). Developmental expressionofdystrophinon the plasmamembrane of rat muscle cells. Protoplasma 151, 11-18. Hammerton,R.W.,Krzeminski,K.A.,Mays,R.W.,Ryan,T.A., Wollner,D.A. &Nelson, W.J. (1991). Mechanism for regulating cell surface distribution of Na+, K+-ATPase in polarized epithelial cells. Science 254, 847-850. Hauschka, S.D. (1994). The embryonicoriginofmuscle. In: Myology: Basic and Clinical (Engel, A.G. & Franzini-Armstrong, C., Eds.) 2nd Edn., pp. 3-73. McGraw-Hill, Inc. New York. (Review) J.E. &Tamoush, A.J. (1990). Heiman-Patterson,T.D, Krupa,T.,Davis, K.F.,Fishbeck,K.F.,Sylvester, Immunohistochemistry and Western analysis of dystrophin in normal and DMD muscle culture. J. Neurol. Sci. 98,252-253. Hoffman, E.P., Brown, R.H. Jr. & Kunkel, L.M. (1987). Dystrophin: the protein product of the Duchenne muscular dystrophy locus. Cell 5 1(6), 919-928. Horwitz, A.F., Schotland, D.L. & Franzini-Armstrong, C. (1994). The plasmamembrane ofthe muscle fiber: composition and structure. In: Myology: Basic and Clinical (Engel, A.G. & Franzini-Armstrong, C., Eds.), 2nd Edn., Vol. 1, pp. 200-222. McGraw-Hill, New York. (Review) Huxley, H.E. (1964). Evidence for continuity between the central elements of the triads and extracellular space in frog sartorius muscle. Nature 4937, 1067-1071. Hynes, R.O. (1992). Integrins: versatility, modulation, and signalling in cell adhesion. Cell 69(1), 11-25. (Review) Ibraghimov-Beskrovnaya, O., Ervasti, J.M., Leveille, C.L., Slaughter, C.A., Sernett, S.W. & Cambpell, K.P. (1992). Primary structure of dystrophin-associated glycoproteins linking dystrophin to the extracellular matrix. Nature 355,696702. Ishikawa, H. (1968). Formation of elaborate networks of T-system tubules in cultured skeletal muscle with special reference to the T-system formation. J. Cell Biol. 38, 5 1 4 6 . Jay, S.D., Ellis, S.B., McCue, A.F., Williams, M.E., Vedvick, T.S., Harpold, M.M. & Campbell, K.P. (1990). Primary structure of the a-subunit of the DHP-sensitive calcium channel from skeletal muscle. Science, 248, 4 9 W 9 2 . Jay, S.D., Sharp,A.H.,Kahl, S.D.,Vedvick,T.S.,Harpold, M.M. &Campbell, K.P. (1991). Structural characterization of the dihydropyridine-sensitive calcium channel alpha 2-subunit and the associated delta peptides. J. Biol. Chem. 266(5), 3287-3293. Jorgensen, A.O., Shen, AC.-Y., Arnold, W., Leung, A.T. & Campbell, K.T. (1989). Subcellular distribution of the 1,4-dihydropyridine receptor in rabbit skeletal muscle in situ: an immunofluorescence and immunogold labeling study. J. Cell Biol. 109, 135-147. Jorgensen, A.O., Arnold, W., Shen, AC.-Y, Yuan, S., Gaver, M. & Campbell, K.P. (1990). Identification of novel proteins unique to either transverse tubules (TS28) or the sarcolemma (SL50) in rabbit skeletal muscle. J. Cell Biol. 110, 1173-1 185. Jorgensen, A.O., Shen, AC.-Y. & Campbell, K.P. (1995). Distinct distribution of TS28 and the a,-subunit of the LCdihydropyridine receptor (a,-DHPR) in transverse (T) tubules of developing rabbit skeletal muscle. Biophys. J. 68(2), A348 Kelly, A.M. (197 1). Sarcoplasmic reticulum and T-tubules in differentiating rat skeletal muscle. J. Cell Biol. 4 9 , 3 5 4 4 .

196

ANNELISE 0. JORCENSEN

Kislauskis, E.H., Li, Z., Singer, R.H. & Taneja, K.L. (1993). Isoform-specific3'-untranslatedsequences sort a-cardiac and P-cytoplasmic actin messenger RNAs to different cytoplasmic compartments. J. Cell Biol. 123(1), 165-172. Kislauskis, E.H., Zhu, X. & Singer, R.H. (1994). Sequences responsible for intracellular localization of p-actin messenger RNA also affect cell phenotype. J. Cell Biol. 127(2), 441451. Klamut, H.J., Zubrzycka-Gaarn, E.E., Bulman, D.E., Malhotra, S.B., Bodrug, S.E., Worton, R.G. & Ray, P.N. (1989). Myogenic regulation of dystrophin gene expression. British Med. Bull 45(3), 681-702. (Review) Knudsen,K.A. & Horwitz, A.F. (1994). The plasmamembraneofthemuscle fiber: adhesionmolecules. In: Myology: Basic and Clincial (Engel, A.G. & Franzini-Armstrong, C., Eds.), 2nd Edn., pp. 223-241. McGraw-Hill, New York. (Review) Knudson, C.M., Chaudari, N., Sharp, A.H., Powell, J.A., Beam, K.G. & Campbell, K.P. (1989). Specific absence of the alpha, subunit of the dihydropyridine receptor in mice with muscular dysgenesis. J. Biol. Chem. 264, 1345-1348. Kurpakus, M.A., Quaranta, V. &Jones, J.C.R. (1991). Surface relocation of alpha6 Beta4 integrins and assembly ofhemidesmosomes in an invitro model ofwound healing. J. Cell Biol. 115,1737-1750. Lau, Y.H., Caswell, A.H. & Brunschwig, J.P. (1977). Isolation oftransverse tubules by fractionation of triad junctions of skeletal muscle. J. Biol. Chem. 252, 5565-5574. Lazarides, E. (1978). The distribution of desmin (100 A) filaments in primary cultures of embryonic chick cardiac cells. Exp. Cell Res. 112, 265-273. Liou, H-C. & Baltimore, D. (1993). Regulation ofthe NF-xB/rel transcription factor and IxB inhibitor system. Curr. Opin. Cell Biol. 5,477487. (Review) Lisanti, M.P., Scherer, P.E., Tang, Z. & Sargiacomo, M. (1994). Caveolae, caveolin and caveolin-rich membrane domains: a signalling hypothesis. Trends Cell Biol. 4,23 1-235. (Review) Luna, E.J. & Hitt, A.L. (1992). Cytoskeleton-plasma membrane interactions. Science 258, 955-963. (Review) Lytton, J. & MacLennan, D.H. (1991). Sarcoplasmic Reticulum, In: The Heart and Cardiovascular System (Fozzard, H.A., Haber, E., Jennings, R.B., Katz, A.M. & Morgan, H.E., Eds.), Vol. 2, pp. 1203-1222. Raven Press, New York. Marks, A.R., Taubman, M.B., Saito, A., Dai, Y. & Fleischer, S. (1991). The ryanodine receptor/junctional channel complex is regulated by growth factors in a myogenic cell line. J. Cell Biol. 114,303-312. Masuda, T., Fujimaki, N., Ozawa, E. & Ishikawa, H. (1992). Confocal laser microscopy of dystrophin localization in guinea pig skeletal muscle fibers. J. Cell Biol. 119(3), 543-548. Matsumura, K. & Campbell, K.P. (1994). Dystrophin-glycoprotein complex: Its role in the molecular pathogenesis of muscular dystrophies. Muscle & Nerve 17,2-15. (Review) Matter, K. & Mellman, I. (1994). Mechanisms of cell polarity: sorting and transport in epithelial cells. Curr. Opin. Cell Biol. 6, 545-554. (Review) Miller, R.J. (1994). Voltage-sensitive Ca2+channels. J. Biol. Chem. 267(3), 1403-1406. (Review) Miranda, A.F., Bonilla, E., Martucci, G., Moraes, C.T., Hays, A.P. & DiMauro, S. (1988). Immunocytochemical study of dystrophin in muscle culture from patients with Duchenne muscular dystrophy and unaffected control patients. Am. J. Pathol. 132,410-416. Morris, E.J. & Fulton, A.B. (1994). Rearrangement ofmRNAs for costamere proteins duringcostamere development in cultured skeletal muscle from chicken. J. Cell Sci. 107, 377-386. (Review) Neely, A,, Wei X., Olcese, R., Birnbaumer, L. & Stefani, E. (1993). Potentiation by the betasubunit of the ratio of the ionic current to the charge movement in the cardiac calcium channel. Science 262(5133), 575-578. Nelson, W.J. & Lazarides, E. (1983). Expression of the beta subunit of spectrin in nonerythroid cells. Proc. Natl. Acad. Sci. USA 80(2), 363-367. Nelson, W.J. & Lazarides, E. (1984). Goblin (ankyin) in striated muscle: Identification of the potential membrane receptor for erythroidspectrin in muscle cells. Proc. Natl. Acad. Sci. USA 81,3292-3296.

Cell Surface in Skeletal Muscle

197

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. Numa, S., Tanabe,T., Takeshima, H., Mikami, A.,Niidome,T. &Nishimura,S., Adams, B.A. & Beam, K.G. (1990). Molecular insights into excitation-contraction coupling. Cold Spring Harb. Sympos. Quant. Biol. 55, 1-7. (Review) Ohlendieck, K., Ervasti, J.M., Snook, J.B. &Campbell, K.P. (1991). Dystrophin-glycoproteincomplex is highly enriched in isolated skeletal muscle sarcolemma. J. Cell Biol. 112( I), 135-148. Pardo, J.V., Siciliano, J.D. & Craig, S.W. (1983a). A vinculin-containing cortical lattice in skeletal muscle: transverse lattice elements (“costameres”) mark sites of attachment between myofibrils and sarcolemma. Proc. Natl. Acad. Sci. USA 80, 1008-1012. Pardo, J.V., Siciliano, J.D. & Craig, S.W. (1983b). Vinculin is a component ofan extensive network of myofibril-sarcolemma attachment regions in cardiac muscle fibers. J. Cell Biol. 97, 1081-1 088. Park-Matsumoto, Y.C., Kameda,N., Kobayashi, T. & Tsukagoshi, H. (1991). Developmental study of the expression of dystrophin in cultured human muscle aneurally and innervated with fetal rat spinal cord. Brain Res. 565,28&289. Peachey, L.D. & Franzini-Armstrong, C. (1983). Structure and function of membrane systems of skeletal muscle cells. In: Handbook of Physiology Skeletal Muscle, pp. 23-71, (Peachey, L.D., Adrian, R.H. & Geiger, S.R., Eds.), Section 10. American Physiological Society, Bethesda, Maryland. (Review) Pierobon-Bormioli, S. (1981). Transverse sarcomere filamentous sytems: “Z- and M-cables.” J. Mus. Res. Cell Motil. 2, 401413. Pincon-Raymond, M., Rieger, F., Fosset, M. & Lazdunski, M. (1985). Abnormal transverse tubule system and abnormal amount of receptors for Ca2+ channel inhibitors of the dihydropyridine family in skeletal muscle from mice with embryonic muscular dysgenesis. Dev. Biol. 112, 458466. Porter, G.A., Dmytrenko, G.M., Winkelmann, J.C. & Bloch, R.J. (1992). Dystrophin colocalizes with P-spectrin in distinct subsarcolemmal domains in mammalian skeletal muscle. J. Cell Biol. 117(5), 997-1005. Powell, J.A. (1990). Muscular dysgenesis: a model system for studying skeletal muscle development. FASEB J. 4,2798-2808. (Review) Pragnell, M., De Waard, M., Mori, Y., Tanabe, T., Snutch, T.P. & Campbell, K.P. (1994). Calcium channel beta-subunit binds to a conserved motif in the 1-11 cytoplasmic linker of the alpha I-subunit. Nature 368(6466) 67-70. Prelle, A., Chianese, L., Moggio, M., Gallanti, A,, Sciacco, M., Checcarelli,N., Comi, G., Scarpini, E., Bonilla, E. & Scarlato, G. (1991). Appearance and localization of dystrophin in normal human fetal muscle. Int. J. Devl. Neuroscience 9(6), 607412. Ranscht, B. & Dours-Zimmermann, M.T. (1991). T-Cadherin, a novel cadherin cell adhesion molecule in the nervous system lacks the conserved cytoplasmic region. Neuron 7,391-402. Repasky, E.A., Granger, B.L. & Lazarides, E. (1982). Widespread occurrence of avian spectrin in nonerythroid cells. Cell 29(3), 821-833. Rios, E., Pizarro, G. & Stefani, E. (1992). Charge movement and the nature of signal transduction in skeletal muscle excitation-contraction coupling. Ann. Rev. Physiol. 54, 109-133. (Review) Roberds, S.L. & Anderson, R.D., Ibraghimov-Beskrovnaya, 0. & Campbell, K.P. (1993). Primary structure and muscle-specific expression of the 50-kDa dystrophin-associated glycoprotein (adhalin). J. Biol. Chem. 268(32), 2373S23742. Rodriguez-Boulan, E. & S.K. Powell. (1992). Polarity ofEpithelial andNeuronal Cells. Ann. Rev. Cell Biol. 8,395-428. (Review)

198

ANNELISE 0.JORGENSEN

Romer, L.H., Burridge, K. & Turner, C.E. (1992). Signaling between the extracellular matrix and the cytoskeleton: tyrosine phosphorylation and focal adhesion assembly. Cold Spring Harb. Symp. Quant. Biol. 57, 193-202. Rosemblatt, M., Hidalgo, C., Vergara, C. & Ikemoto, N. (1981). Immunological and biochemical properties of transverse tubule membranes isolated from rabbit skeletal muscle. J. Biol. Chem. 256( 15), 8140-8 148. Rothman, J.E. & Warren, G. (1994). Implications ofthe SNARE hypothesis for intracellularmembrane topology and dynamics. Curr. Biol. 4(3), 22&233. (Review) Ruth, P., Rohrkasten, A., Biel, M., Bosse, E., Regulla, S., Meyer, H.E., Flockerzi, V. & Hofmann, F. (1989). Primary structure ofthe beta subunit of the DHP-sensitive calcium channel from skeletal muscle. Science 245(4922), 11 15-1 118. Sabbadini, R.A. & Dahms, A.S. (1989). Biochemical properties of isolated transverse tubular membranes. J. Bioenerg. Biomemb. 21(2), 163-213. (Review) Salvatori, S., Furlan, S., Millikin, B., Sabbadini, R., Betto, R., Margreth, A. & Salviati, G. (1993). Localization of protein kinase C in skeletal muscle T-tubule membranes. Biochem. & Biophys. Res. Comm., pp. 1073-1080. Sanes, J.R. (1994). The extracellular matrix. In: Myology: Basic and Clinical, 2nd Edition, Volume I, pp. 242-260. Engel, A.G. & Franzini-Armstrong,C.,Eds., McGraw-Hill,. New York. (Review) Sanes, J.R., Schachner, M. & Covault, J. (1986). Expression of several adhesive macromolecules (N-CAM, L1, J1, NILE, uvomorulin, laminin, fibronectin, and a heparan sulfate proteoglycan) in embryonic, adult, and denervated adult skeletal muscle. J. Cell Biol. 102,42&43 1. Schneider, M.F. (1994). Control of calcium release in functioning skeletal muscle fibers. Annu. Rev. Physiol. 56,463484. (Review) Schiaffno, S., Cantini, M. & Sartore, S. (1977). T-system formation in cultured rat skeletal tissue. Tiss. Cell 9,437446. Schwarz, M.A., Owaribe, K., Kartenbeck, J. & Franke, W.E. (1990). Desmosomes and hemidesmosomes: constitutive molecular components. Ann. Rev. Cell Biol. 6, 461491. (Review) Seiler, S. & Fleischer, S. (1988). Isolation and characterization of sarcolemmal vesicles from rabbit fast skeletal muscle. Methods Enzymol. 157,2636. (Review) Severs, N.J. (1988). Caveolae: static inpocketings of the plasma membrane, dynamic vesicles or plain artifact? J. Cell Sci. 90,341-348. (Review) Shear, C.R. & Bloch, R.J. (1985). Vinculin in subsarcolemmal densities in chicken skeletal muscle: localization and relationship to intracellular and extracellular structures. J. Cell Biol. 101, 24&2 5 6. Simon, K.O.,Otey, C.A., Pavalko, F.M. & Burridge, K. (1991) Protein interactions linking actin to the plasma membrane in focal adhesions. Curr. Topics Memb. 38, 57-64. (Review) Singer, R.H. (1992). The cytoskeleton and mRNA localization. Curr. Opin. Cell Biol. 4, 15-19. (Review) Small, J.V., Furst D.O. & Thornell, L.E. (1992). The cytoskeletal lattice of muscle cells. Eur. J. Biochem. 208(3), 559-572. Snutch, T.P. & Reiner, P.B. (1992). Ca2+ channels: diversity of form and knction. Curr. Opin. Neurobiol. 2(3), 247-253. (Review) Sollner, T., Whiteheart, S.W., Brunner, M., Ejument-Bromage, H., Geromanos, S., Tempts, P. & Rothman, J.E. (1993a). SNAP receptors implicated in vesicle targetting and fusion. Nature 362, 3 18-324. Sollner, T., Bennett, M.K., Whiteheart, S.W., Scheller, R.H. & Rothman, J.E. (1993b). A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation and fusion. Cell 75,409-41 8. Stepp, M.A., Spurr-Michaud, S., Tisdale, A., Elwell, J. & Gipson, I.K. (1990). a6P4 integrin heterodimer is a.component of hemidesmosomes. Proc. Natl. Acad. Sci. USA 87, 8970-8974.

Cell Surface in Skeletal Muscle

199

Straub, V., Bittner, RE., Leger, J.J. & Voit, T. (1992). Direct visualization ofthe dystrophin network on skeletal muscle fiber membrane. J. Cell Biol. 119(5), 1183-1 191. Bennett, L., Tanabe, T., Beam, K.G. & Franzini-Armstrong, C. (1994). Restoration of Takekura, H., junctional tetrads in dysgenic myotubes by dihydropyridine receptor cDNA. Biophys. J. 67, 793-803. Takemitsu, M., Ishiura, S., Koga, R., Kamakura, K.: Arahata, K., Nonaka, I. & Sugita, H. (1991). Dystrophin-related protein in the fetal and denervated skeletal muscles of normal and mdr mice. Biochem. Biophys. Res. Comm. 180(3), 1179-1186. Tanabe, T., Beam, K.G, Powell, J.A. & Numa, S. (1988). Restoration of excitatiowcontraction coupling and slow calcium current in dysgenic muscle by dihydropyridine receptor complementary DNA. Nature 336(6195), 134-139. Tinsley, J.M., Blake, D.J., Zuellig, R.A. & Davies, K.E. (1994). Increasing complexity of the dystrophin-associated protein complex. Proc. Natl. Acad. Sci. USA 9 I, 8307-83 13. (Review) Volk, T., Fessler, L.I. & Fessler, J.H. (1990). A role for integrin in the formation of sarcomeric cytoarchitecture. Cell, 63, 525-536. Vybiral, T., Winkelmann, J.C., Roberts, R., Joe, E-H., Casey, D.L., Williams, J.K. & Epstein, H.F. (1992). Human cardiac and skeletal muscle spectrins: differential expression and localization. Cell Motil. Cytoskel. 21,293-304. Watkins. S.C., Hoffman, E.P., Slayter, H.S. & Kunkel, L.M. (1988). Immunoelectron microscopic localization of dystrophin in myofibres. Nature 333, 863-866. Wilson, P.,Arnold, W. & Jorgensen, A.O. (1992). Early biogenesis of transverse tubules (T-T) in developing rabbit skeletal muscle in vitro. Mol. Biol. Cell (Suppl.) 3, 256a. Winkelmann, J.C., Costa, F.F., Linzie, B.L. & Forget, B.G. (1990). !3 Spectrin in human skeletal muscle. J. Biol. Chem. 265(33), 20449-20454. Worton, R.G. (1994). Dystrophin: the long and short of it. J. Clin. Invest. 93,4. Yamamoto, H., Mizuno, Y., Hayashi, K., Nonaka, I., Yoshida, M. & Ozawa, E. (1994). Expression of dystrophin-associated protein 35 DAG (A4) and 50 DAG (A2) is confined to striated muscles. J. Biochem. 115( I), 162-167. Yang, B., Ibraghimov-Beskrovnaya, O., Moomaw, C.R., Slaughter, C.A. & Campbell, K.P. (1994). Heterogeneity of the 59 k-Da dystrophin-associated protein revealed by cDNA cloning and expression. J. Biol. Chem. 269(8), 6040-6044. Yoshida, M. & Ozawa, E. (1990). Glycoprotein complex anchoring dystrophin to sarcolemma. J. Biochem. 108,748-752. Yuan, S., Arnold, W. & Jorgensen, A.O. (1990). Biogenesis of transverse tubules: immunocytochemical localization of a transverse tubular protein (TS28) and a sarcolemmal protein (SL50) in rabbit skeletal muscle developing in situ. J. Cell Biol. 110, 1187-1 198. Yuan, S., Arnold, W. & Jorgensen, A.O. (1991). Biogenesis of transverse tubules and triads: immunolocalization of the 1,4-dihydropyridine receptor, TS28, and the ryanodine receptor in rabbit skeletal muscle developing in situ. J. Cell Biol. 112,289-301. Zampighi, G., Vergara, J. & Ramon, F. (1975). On the connection between the transverse tubules and the plasma membrane in frog semitendinosus skeletal muscle. J. Cell Biol. 64, 734-740. Zhao, J., Yoshioka, K., Miike, T. & Miyatake, M. (1993). Developmental studies ofdystrophin-positive fibers in mdx, and DRP localization. J. Neurol. Sci. 114, 104-108.

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POLARITY AND POLARIZATION OF FIBROBLASTS IN CULTURE

Albert K. Harris

I. Introduction . . . . .................................. II. Description of Fib ity (Special Characteristics of Each Type of Margin) ..................... A. "Type A" Margins. . . . . . . . . . . . . . . . . ......................... B. "Type 6" Margins C. "Type C" Margins . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Contact Guidance. . . . . . . . . . . . . . . .................. IV. Contact Inhibition and Contact Paralysis. . . . . . . . . . . . . . . . . . . . . V. Calvanotaxis: Effects of Electric Fields on Fibroblast Polarity VI. Microtubules in the VII. Questions About the Autonomy of Fibroblast Polarity . . VIII. Conclusions and Prospects . . . .........................

Advances in Molecular and Cell Biology Volume 26, pages 201-252. Copyright 0 1998 by JAI Press Inc. All right of reproduction in any form reserved. ISBN: 0-7623-0381-6

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1.

INTRODUCTION

Fibroblasts are the most familiar of tissue culture cells. They are the ubiquitous, nondescript, generally stellate cells that quickly come to dominate primary cultures derived from explants of almost any organ or tissue. Many of the classical studies of tissue cell locomotion, such as those of Michael Abercrombie and Joan Heaysman (1953, 1954), concentrated on fibroblasts from primary cultures, with chick embryo heart fibroblasts being the most frequently used. In addition, many “established’’ cell lines of fibroblasts have been developed and are very widely used in research; examples include the various 3T3 lines, among many others. To attempt any very precise definition of exactly what is, and what is not, a fibroblast would be premature. Molecular criteria will eventually be found, such as possession of intermediate filaments of the vimentin class (as opposed to those of the keratin type characteristic of epithelial cells), although vimentin has turned up in other cell types, including endothelia. The secretion of collagen and other fibers was once assumed to be a special property of fibroblasts, until it was shown that this is also done by epithelial cells, among others. The approximately synonymous term “mechanocyte” failed to catch on as a substitute for fibroblast, but would have been a more accurate reflection of what seems to be the real function and special role of these cells, namely the exertion of mechanical forces. Using flexible rubber substrata and gels of fibrin and reprecipitated collagen, it has been shown that fibroblasts exert pulling forces hundreds of times stronger than needed for their own propulsion, with these forces being capable of large scale reorganization of collagen fibers over distances as great as several centimeters (Stopak and Harris, 1982). The main function of the phenomenon we see as fibroblast ‘‘locomotion’’ may really be mechanical reorganization of extracellular matrix materials. Fibroblasts try to pull surrounding materials rearward past their cell bodies, for purposes believed to include wound closure. Because of the rigidity of glass, polystyrene and other artificial culture substrata, however. all we usually see is that the fibroblasts stretch themselves out flat and crawl around jerkily. The polarity of fibroblasts is primarily a matter of directional protrusion combined with the exertion of strong pulling forces in the direction opposite to this protrusion. This is easiest to visualize when a fibroblast happens to have only a single front end, advancing in front, with a single trailing margin at the opposite end. Unfortunately, such nice and simple “typical fibroblasts” tend to be more the exception than the rule. Even when a given fibroblast does happen to be monopodial in this way, this is usually only transient. In contrast to polymorphonuclear leucocytes, for example, where monopodial behavior is the norm, fibroblasts are neither consistently nor persistently monopodial, with distinct and long-lasting front and rear ends. Instead, most fibroblasts are irregularly multipolar. Two, three (or more) parts of a given cell’s margins typically crawl off in different directions at the same time, stretching the cell body between them. The usual stellate shape of fibroblasts results from their being stretched between several competing “front” ends, (Figures 1,2).

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figure 7.

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3T3 fibroblast, seen by ordinary phase contrast.

The same 3T3 fibroblast seen by interference reflection microscopy, in which the areas of contact and adhesion between the substratum and the cell’s lower surface show up as grey and black areas. figure 2.

It should also be noted that even quite small fragments of fibroblasts, lacking nuclei, tend to adopt very much these same types of polarity (Gelfand et al. 1985). The very useful term “leading lamella” was introduced by Abercrombie et al. (1970a, 1970b) to refer to these “front” ends, and is equally applicable whether there happens to be only one of them (in a monopodial cell), or when a cell happens to have two or more leading lamellae at the same time. For example, the fibroblast shown in figures one and two happens to have four leading lamellae. It is not even

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unusual for a fibroblast to have one single broad leading lamella extending all the way, 360°, around its periphery. When trypsinized fibroblasts are plated out onto a new substratum, most of them will pass through such a stage. Freshly-plated cells spread initially along all parts of their periphery, thus passing through a characteristic “fried egg” morphology a few hours after first being plated out (see Witkowski and Broughton, 197I). Subsequently, this single leading lamella breaks up into two or three regions that continue to advance outward, separated by parts of the margin that retract inward. Another case where leading lamellae occupy the entire cell margin is when some kinds of fibroblasts are exposed to antimicrotubule drugs (Vasiliev et al. 1970), as is discussed more fully in a later section. Net locomotion, when it occurs, results from imbalances between leading lamellae, for example when those along one side of a given cell are stronger than those along the opposite side. Such a dominant leading lamella, or (often) groups of adjacent leading lamellae, therefore become the temporary front end of the cell, in the sense of pulling everything else along behind them. At the opposite, trailing side of the cell, one frequently observes parts of the cell margin progressively being pulled loose from their adhesions to the substratum, often breaking these adhesions a few at a time in a jerky fashion, sometimes with long strands of plasma membrane (called “retraction fibers”) being pulled out behind the cell as its margin passively retreats (Harris, 1990). As I am trying to emphasize, the essential feature of fibroblast polarity is the subdivision of each cell’s lateral margins into several relatively distinct kinds of behavior (three main kinds, I think) . Some parts of the margin advance actively outward. Other parts of the margin are dragged passively behind as a sort of tail. And, the third class of margins are those that sag inward in long smooth catenary-like curves along the sides. With the goal of streamlining discussion, I propose simply to call these types of cell margins “A,” “B,” and “C.” The “type A” margins are the ones that Advance outward; the type B margins are the ones trailing Behind; while the type C margins are the ones that sag inward like Catenary curves, (Figure 3). This nomenclature is meant only to be provisional, and to focus attention on such questions as which particular molecules need to be concentrated near a given part of a cell’s margin in order to cause that region to protrude forward as a “type A margin.” When the molecular basis of these three different behaviors becomes understood, then we can name them accordingly. For example, the type A margins may then be called “actin reassembly margins,” or perhaps “type I myosin margins.” Meanwhile, using simply A, B, and C is intended to minimize circumlocution. The kinds of questions we need to ask include the following. What special molecules (type I myosins, perhaps?), or molecular activities (such as assembly of gactin into fibers) areresponsible for causing a given one part of a cell’s margin to become “type A’? Likewise, why does treatment with microtubule poisons tend to cause type A activity to spread out, and why does this occur in certain cell lines much more than in those from primary cultures? In the phenomenon called “contact guidance,” why do ridges and grooves cause cells to concentrate type A activity to

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Figure3.

Diagram of a typical fibroblast, crawling on a flat glass or plastic substratum ir tissue culture, with the three characteristic types of cell margins that are described in the text as type A, type 6 , and type C. Type A margins advance forward, with protrusive activity, ruffling, actin assembly and type I myosin being concentrated there. Type E margins are those that retreat, but do retain adhesions to the substratum, while type C margins are mostly non-adherent and sag passively inward in smooth concave curves Fibroblast polarization is essentially a matter of separating these three sets of activities and properties into different parts of each cell’s margins.

the parts touching grooves and ridges? Conversely, in the twin phenomena of contact paralysis and contact inhibition, why is it that cell-cell contact so often causes type A margins to switch over to type B or even to type C behavior? With the molecular aspects of propulsion now becoming so much clearer, and with the accumulation of descriptions of new cytoskeletal proteins and their distributions, the time now seems to be ripe for explaining whole-cell behaviors in terms of molecules and forces. I hope that this review can help focus attention on solvable problems in this area. Some widely-used textbooks describe the locomotion of fibroblasts and other tissue cells as consisting of an alternating cycle of forward protrusion of filopodia, adhesion of these filopodia to the substratum, and contraction of these filopodia, so as to pull the cell body forward. Unfortunately, this “reach-grab-pull” concept is a misleading oversimplification, essentially an “Aesop’s fable,” especially as applied to fibroblasts. Notice that it would lead you to expect all of the following (none of which is true): (1) that cells lacking filopodia could not spread or crawl; (2) that cytoplasmic protrusions should be concentrated along the cell’s leading margins, and therefore that the presence of such protrusions along a certain part of a cell’s margin would be evidence that that must be the front end or leading edge; (3) that rearward force-exertion would be concentrated in the area right at the cell margin, between the tips and the bases of these filopodia; and (4) that force exertion would be pulsatory, varying with time in synchrony with the supposed cycle of protrusion, attachment and contraction. Not one of these predictions is true. (1) It is not unusual for actively crawling fibroblasts to lack filopodia. (2) Where concentrations of cytoplasmic extensions are observed, this most often corresponds to the rear end of the fibroblast (rather than the front end), with these extensions being retraction fibers in the process of being pulled out uassivelv from retreatinn (“tvue B”) cell margins.

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( 3 ) Most of the pulling forces exerted by fibroblasts are transmitted to the substratum 5, 10 or more millimicrons behind the advancing (type A) cell margins, being exerted tangentially through the plasma membrane. &furthermore, (4) even though the magnitude of these pulling forces can vary with time, such variations are not in any regular cycle correlated with filopodial extension and withdrawal. My own experience and belief is that filopodia have little or nothing to do with fibroblast locomotion, despite the undoubted importance of such protrusions in nerve growth cones. However useful for satisfying students’ curiosity, the “reach-grab-pull” idea makes too many incorrect predictions to be anything but an obstacle to understanding.

11.

DESCRIPTION OF FIBROBLAST POLARITY (SPECIAL CHARACTERISTICS OF EACH TYPE OF MARGIN) A.

”Type A“ Margins

These are the parts of the cell margins that advance forward across substrata. Their forward progress is usually oscillatory, with periods of advance alternating with periods of retreat, in the category of “two steps forward and one step back,” with the durations of each type of behavior being on the order of a few minutes. Even during these small retreats, these type margins retain the peculiarity of being the only parts of the cell margin where stiff protrusions extend out in front of the most distal adhesions to the substratum. Thin forward protrusions, called lamellipodia, are the most characteristic feature of these margins. Lamellipodia are very thin, as little as 0.1 micrometers in thickness (Abercrombie et al., 1970a, 1971), but can be 5 , 10, 15 or more micrometers in width and extend out 5 or more micrometers in front of the cell, usually bending slowly backwards in a type of movement called “ruffling” (Abercrombie et al., 1970b), which is familiar to anyone who has ever seen a time lapse film of any type of crawling tissue cells. Lamellipodia are filled with a meshwork of actin fibers, and mechanically are surprisingly stiff, in the sense of resisting bending by glass microneedle. Lamellipodia may seem to be flopping around limply, but really they are stiff and their bending is an active process. Lamellipodia are packed tightly with meshworks of actin fibers, as seen in transmission electron microscopy sections (Heath and Dunn, 1978; Small, 1981) as well as when cells are stained with fluorescently labeled phalloidin, apeptide mushroom toxin which binds strongly to filamentous actin (as well as promoting assembly of actin into fibers). There is also evidence that the outermost parts of type A margins are sites of actin assembly (Wang, 1985; Svitkinaet al., 1986; Symons and Mitchison, 1991), as well as that actin flows centripetally from there (Fisher et al., 1988). Other cytoplasmic components are excluded from the lamellipodia, including not just mitochondria and vesicles but even microtubules and conventional type I1 my-

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osins. In contrast, type I myosins have recently been found concentrated specifically along the outer margins of fibroblast lamellipodia (Wagner et al., 1992). Two other classes of cell surface protrusion, in addition to lamellipodia, are often concentrated along the type A margins of fibroblasts. Filopodia are spike-like protrusions, which are likewise packed with fibrous actin, and likewise extend out ahead, fold back, and move centripetally across cell’s upper surface, often being intermingled with lamellipodia, like fingers extending from the hand. The third class of cell surface protrusion consists of hemispherical herniations in the plasma membrane, called blebs, ranging from diameters of as little as one micrometer up to 5 or even 10 micrometers. Blebs can bubble forth fromjust about any part of a fibroblast’s surface, but are often concentrated right along the advancing “type A” margins, intermingled with lamellipodia. If a cell is blebbing only slightly, at a few locations, these will almost always be concentrated right at the tips of the type A margins, although these parts of the cell surface were either under especially strong outward pressure, or were particularly weak in resisting widely distributed outward forces. Each bleb forms rapidly, in a few seconds, and then gradually shrinks and disappears over a period of a minute or more. Fibroblasts can be easily stimulated to switch from forming lamellipodia to forming blebs; one way to do this is to make the culture medium more hypertonic and another is to decrease the stretching tension in the cell body. A wave of spontaneous blebbing occurs each time a fibroblast undergoes partial detachment and retraction of one or more of its leading lamellae; not just the lamellipodia at the tip of the retracted lamella, but also those along the edges of this cell’s other lamellae, will be replaced with blebs until re-spreading has restored a state of tensile stress. Likewise, when trypsinized fibroblasts are plated out, they first bleb and only later ruffle. The typical sequence is for each new bleb to be a little smaller in diameter than those previously formed, until lamellipodia replace the (now very tiny) blebs. The mechanistic significance of this and other easily-observed transitions has yet to be discovered. Besides their special powers of outward protrusion, type A margins also have special adhesive properties, in particular the propensity to initiate new adhesions (Harris, 1973b; Heath and Dunn, 1978). The optical technique called interference reflection microscopy (IRM) makes visible the closeness of approach between the cell’s lower plasma membrane and glass, plastic, or even silicone rubber substrata. The closest contacts appear black by this technique, while slightly wider gaps appear gray (Abercrombie and Dunn, 1975; Izzard and Lochner, 1976, 1980). Both these types of cell-substratum contact are concentrated behind the type A margins of fibroblasts, the black “focal adhesion plaques,” appearing as many small spots elongated perpendicular to the nearby cell margin and around a micrometer in width and several micrometers in length, scattered within a much wider gray contact area. I once directly tested what had seemed to me the naive assumption that the black contacts were very strong adhesions while the gray contacts were weak adhesions. Using thin glass microneedles and a micromanipulator, I peeled fibroblasts

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away from glass, styrene and rubber substrata. As it turned out, however, the results indicated that the usual assumptions about relative adhesive strengths, however naive, seem to be absolutely correct! The gray areas can be easily peeled away from the substratum, while the black contacts cannot be detached without ripping away that part of the plasma membrane (except with silicone rubber substrata, in which case you can often pop loose adhesion plaques without ripping the membrane). In addition, I found that new black adhesions cannot be created simply by pushing down on the top of the cell until previously nonadherent areas of plasma membrane on the bottom are pushed close enough to the substratum to appear black by interference reflection. Even after being pushed close to glass substrata for as long as several minutes, such areas immediately moved back to their former separation distance once the pressure is removed. This supports the idea that there is something specially adhesive about the plasma membrane components at the focal contacts, and it has indeed been shown that fibronectin receptors (integrins) become concentrated there (Burridge and Fath, 1989). On the other hand, it remains to be clarified what is special about the membrane making up the broad gray contacts; one possibility is that these areas are likewise integrin-rich, but with a lesser degree of concen tration. Time lapse films of the interference reflection images of fibroblasts have to be made using long time exposures, at moderate light levels, since the intense light ordinarily used for IRM will radically change cell behavior. But with time exposures, one can easily observe a close correlation between type A margin behavior and the presence of the broad gray contact areas. These areas are very reminiscent of radar images shown on television newscasts of bands of thunderstorms. Wherever new contacts develop, this is a sure sign that a part of the cell margin that had previously been inactive (type B or C) is about to become a type A margin and begin protruding outward. Conversely, quiescence and shrinkage of these gray contact areas is the first indication that a type A cell margin is about to change into a type B region. The molecular basis of these changes deserves study. As regards force exertion, this aspect of fibroblast polarity has been studied by culturing the cells on flexible substrata of plasma clots, collagen gels and very thin sheets of silicone rubber (Figure 4). These forces are called “traction”, and should be not be confused with contraction, in that the parts of the cells exerting the traction forces do not shorten in length (do not contract), but rather become more and more elongated and stretched as a cumulative result of their exertion of these traction forces. The active displacement of visible particles and lectins (Abercrombie et al., 1 9 7 0 ~ 1972; ; Harris and Dunn, 1972) attached to the outer surface of the plasma membrane (sometimes called “capping”) seems to be caused by the same mechanism of force exertion, since the forces are exerted in the same directions and locations. Along type A margins, the traction forces are consistently directed inward, in the direction exactly opposite the direction of outward spreading. This is mostly perpendicular to the leading margin, but often with a degree of lateral convergence.

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Figure 4. Low magnification view of chick heart fibroblasts cultured on a silicone

rubber substratum. Note the wrinkles in the rubber layer, which are produced by the cells’ traction forces, by means of which the cells spread and crawl about. This is shown by the patterns of wrinkles formed in silicone rubber substrata, where the compression wrinkles formed behind the type A margins are mostly oriented perpendicular to the advancing margin, but often with some wrinkling perpendicular to this as well. Particles attached to both the upper and lower surfaces of fibroblasts are actively transported directly rearward away from type A margins, although often with some tendency of their paths to converge somewhat. On the top, the particles are transported centripetally, usually accumulating near the nucleus, but sometimes also accumulating near the type C and type B margins. As regards cytoplasmic proteins, the concentration of actin as a meshwork along type A margins has already been mentioned. Based on time lapse videos made with very high contrast differential interference contrast optics by Heath and Holifield (I 991), a copy of which they kindly provided to me, it appears that this actin network flows steadily rearward away from fibroblast type A margins at several micrometers per minute (almost exactly as observed in neuronal bag cells by Forscher and Smith, 1988). This would necessitate a high rate of assembly of actin monomers into fibers, with this assembly apparently being concentrated right along the type A margins, apparently at the outer side of the lamellipodia. This concentration of actin assembly may be the most fundamental peculiarity of the type A margin, and be responsible both for the forward protrusion and mechanical stiffening of the lamellipodia, as well as for the traction forces exerted directly rearward away from the margin. Behind the type A margins, the so-called “stress fibers” are formed consisting of actin combined with type I1 myosin, tropomyosin, alpha actinin and other muscle-related proteins. These stress fibers terminate at the focal (black) adhesion plaques mentioned above, where they attach at the inner surface of the plasma membrane. The linking proteins talin and vinculin are also concentrated just inside the plasma membrane at these

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plaques. Talin is also concentrated along the lamellipodium itself, in other words right at the type A margin (Burridge and Connell, 1983). At least a few membrane proteins develop special geometric arrangements relative to type A margins, becoming progressively depleted in concentration there, so as to form dramatic concentration gradients detectable by staining with fluorescent antibodies (Holifield et al., 1990; Ishihara et al., 1990). The formation of these gradients by a given membrane protein is evidence that it is one of the “tine proteins” serving to transmit traction forces tangentially through the plasma membrane, from the rearward-flowing actin networks just inside the plasma membrane to adherent objects and substrata on the outside. B.

“Type B“ Margins

These are the retreating margins that form the rear ends of crawling fibroblasts. The process of their retraction has been analysed by Chen (198 1). Their two most characteristic features are the formation of retraction fibers, rather than lamellipodia or filopodia, and the locations of the black-appearing adhesion plaque cellsubstratum adhesions right at the extreme margins of the cell. This is in contrast to these adhesions’ locations along type A margins, where they are many micrometers behind the cell margin. Another difference is that the broad gray adhesions are usually absent behind type B margins. In other words, whereas behind type A margins, the black adhesions are well behind the edge of the cell, and are surrounded by gray adhesions, in the case of type B margins, the edge of the cell retreats right up to the black adhesions, and the gray-appearing ones are absent. In my experience watching time lapse films of fibroblasts’ IRM images, the disappearance of these broad gray contacts behind an actively spreading type A margin is a sure sign that it is about to become converted into a type B margin. Conversely, it sometimes happens that the latter become reactivated into type A margins, with one of the first signs being a reappearance of gray contacts. Especially in scanning electron micrographs, retraction fibers are often mistaken for filopodia. This can be very misleading indeed, given that retraction fibers are peculiar to trailing margins while filopodia are characteristic of advancing margins. To confuse the one for the other is to mistake the front end for the rear. Retraction fibers are attached to the substratum at their distal ends, which are often somewhat expanded where they adhere, somewhat resembling suction cups. Retraction fibers can be 5, 10,20 or more micrometers in length, in contrast to filopodia which are only a few micrometers long. Another difference is that filopodia are stiff, whereas retraction fibers are flexible, bend easily, and recoil if cut. It is also not unusual for blebs to form along type B margins of fibroblasts. But lamellipodia only form there rarely, and are a sign that the margin is about to become reactivated again as a type A margin. Type B margins are the rarest and most transient of our 3 categories, at least in fibroblasts. Note, for example, that the cell shown in Figures 1 and 2 has only type A

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and type C margins. Examples of type B margins can be seen in the cell at the extreme left of Figure 3, and also in Figure 18 (in a later section, on mirror image symmetry) at the extreme lower right end of the right-hand cell and the extreme lower left end of the left-hand cell. When the molecular basis of fibroblast polarity becomes better understood, it may be that “Type B” margins will be regarded as no more than an inactive form of a type A margin, or perhaps as essentially a transition stage in the conversion of type A margins to type C margins. On the other hand, there are some other cell types, polyrnorphonuclear leucocytes being a good example, for which it is normal to possess a well demarcated and more or less permanent “type B” retreating cell margin. This is accompanied by a higher degree of directionality and polarity than is characteristic of fibroblasts. Whether well-formed “type B” margins should be considered merely a side effect of greater polarization, or perhaps alternatively as part of the causation of that polarization, are questions that will have to be left to future studies.

C. “Type C” Margins These margins are concave, usually smoothly curved, have few if any adhesions to the substratum, and are quiescent, although sometimes with intermittent formation of small lamellipodia. The cytoplasm seems to be stretched tight along the axis parallel

Figure 5. Low magnification view of several chick heart fibroblasts cultured on a haptotactic substratum consisting of elongate rectangles (10 micrometers wide and 100 micrometers long) of vacuum-evaporated palladium metal overlying a nonwettable (and relatively nonadhesive) polystyrene culture dish. The cells elongate along these rectangles, and only extend short distances off of them. Had these adhesive islands been made smaller than the space occupied by a spread cell, cells would spread off them. In fact, when initially plated onto such rectangles, those cells that start out trying to spread prependicular to the long axes of the rectanglesoften do cross the gaps from one island to the next.

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Figure 6.

Low magnification view of a single 3T3 fibroblast cultured on a haptotactic substratum consisting of a series of small squares of vacuum-evaporated palladium metal overlying a nonwettable (and relatively non-adhesive) polystyrene culture dish. Because the individual 10 micrometer squares are much smaller than the area that such a cell would normally occupy on a homogeneously adhesive substratum, the cell eventually occupies 8 or 9 of them, becoming stretched between them into spindly shapes very unusual for these cells.

to the cell margin, which is often much thcker than either of the other two types of margin, often appearing phase dense and refractile in phase contrast microscopy (Harris, 1973b). Bundles of actin fibers are concentrated in these parts of the marginal cytoplasm, and microtubules are often found there, with both the actin fibers and the microtubules having their long axes oriented parallel to the margin (Figures 5, 6). Zand and Albrecht-Buehler (1989,1992) have made special studies of these marginal bundles of actin, reporting that they have properties somewhat different from the usual actin stress fibers in more central areas. In particular, they report that the bundles at the margins are more resistant than ordinary stress fibers to disruption, for example by the drug cytochalasin. They also studied the effects of using micromanipulation to disturb these margins (which they called “webbed edges”).

111.

CONTACT GUIDANCE

Fibroblasts, as well as other tissue cells, respond to fibrous or grooved substrata by aligning themselves along the axes of the fibers or grooves (Figure 7). This orientation response has been observed since the very earliest tissue culture studies (Harrison 1914), and was given the name contact guidance by Paul Weiss (1934, 1961), who studied the phenomenon intensively for many years and made it the basis of important theories about embryonic tissue development and the guidance of nerve

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fibers. When fibroblasts are cultured in or on gels made of clotted plasma (fibrin) or reprecipitated collagen, the cells’ individual morphologies become much more elongate than when flat glass and plastic surfaces are used as culture substrata (Figure 8). These differences in cell morphology are presumed to be due to contact guidance, and make cultured cells look much more like the mesenchymal cells one

Figure 7. Single frame from a time lapse film showing fibroblasts crawling about on a thin layer of clotted fibrin (within which small particles of carbon black had been placed in order to be able to trace the directions and relative strengths of the cells’ propulsive traction forces).

figure 8. Three chick heart fibroblasts cultured inside a gel made of reprecipitated collagen from a rat tail tendon. Notice how the cells adopt extremely elongated shapes when cultured in or on collagen substrata, in contrast to the flattened stellate and polygonal shapes these same cells would have adopted on a flat glass or plastic su bstratum.

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sees in histological sections. This implies that contact guidance must be a primary determinant of cell shape in vivo, where collagen (and, in wounds, fibrin) would form the substrata in which they spread and crawl . In collaboration with Garber, Weiss showed that the degree of directional polarization of fibroblasts plated out in plasma clots depends strongly on the conditions of clotting. When there are many small fibers, the cells become more stellate than when the plasma is clotted in such a way as to produce larger fibers. In the latter case, the fibroblasts become more bipolar (Weiss and Garber, 1952; Garber, 1952). It should be pointed out, however, that these authors’ interpretation of these results in terms of capillarity has long since been overtaken by demonstrations that fibroblast locomotion is an active process, not a form of passive wetting. One can produce contact guidance experimentally by plating fibroblasts out onto such things as diffraction gratings (see Figure 9), fish scales (at least some of which have aligned collagen fibres on one of their surfaces; see Weiss 1961), grooved surfaces (Rovensky and Slavnaja, 1974)), plastic surfaces scratched directionally with abrasives, thin glass fibers, and lengths of spiders’ web (Harrison, 1914) among many other materials. The cells’ dimensional range of sensitivity seems to span a surprisingly broad scale. At the large end of the scale, alignment will occur on cylindrical fibers as much as 200 micrometers in diameter (Dunn and Heath, 1976); while at the small end of the scale, they can also align in response to grooves almost too narrow and shallow to see with amicroscope (O’Hara and Buck, 1979). In either case, the net result can fairly be described as that of favoring development of what we are calling type A cell margins on those sides of each cell that are directed along the fibers or grooves. This results in polarization of the individual cell morphologies into bipolar shapes, each with its long axis oriented parallel to the fibers or grooves. Likewise, the cells’ direction of locomotion also becomes

Figure 9. Low magnification view of the alignment of chick heart fibroblasts along the grooves and ridges of a glass diffraction grating.

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strongly constrained to this axis, along which the individual cells shuttle back and forth. Weiss’s own mechanistic explanation for this phenomenon was based on erroneous notions about cell spreading being a passive wetting of the substratum by a sort of directional capillarity, rather than as an active process of locomotion. Nevertheless, many of Weiss’ proposals about control functions for contact guidance during embryonic development remain tenable. These proposals have been supported by observations of positive feedback cycles in which fibroblast traction pulls collagen fibers into alignment and this progressive alignment of fibers stimulates further alignment of fibroblasts by contact guidance (Stopak and Harris, 1982). In addition, Campbell and Marcum (1980) have documented an actual case of contact guidance of migrating cells inside living Hydra. On the other hand, it is interesting to note that in the development of arteries and arterioles, fibroblasts as well as smooth muscle cells line up circumferentially (rather than longitudinally) around the cylindrical shape of these vessels. Although this would seem to indicate some sort of “reverse contact guidance,” I have plated fragments of small arteries onto glass cylinders and found their cells to align parallel to these cylinders’ axes. The morphogenetic function of contact guidance thus confronts us with some interesting problems in the control of cell polarization. One attempt to account for contact guidance in terms of modern knowledge of cell locomotion was that of Dunn and Heath (1976). Their idea was that protrusion or contraction might inhibited in those parts of cells which happen to become bent by the curvature of the substratum. For example, bending might interfere with either the formation or the contraction of actomyosin fibers. On a cylindrical surface, the least bent sides of the cell would be those facing along the axis of the cylinder (curvature being zero in that direction). If the activities characteristic of type A margins were to become concentrated along these (least bent) sides of the cell, this would explain the bipolar elongation that one observes. This hypothesis was supported by elegant experiments in which fibroblasts were cultured on special substrata made to have sharp ridge lines (shaped like the roof of a house). When those cells that spanned this ridge line were closely examined, their cytoplasmic actin fibers were found not to cross from one side to the another. This indicates either that actin fibers cannot extend around bends, or that there is some kind of preferential elimination of those that are bent. Either should have the effect of biasing locomotory activities in the direction of least substratum curvature. Dunn and Brown (1986) have also developed geometrical criteria for describing and comparing degrees of cell alignment in response to contact guidance. A rather different explanatory hypothesis was proposed by O’Hara and Buck ( 1979). Their idea concerned cell-substratum adhesions rather than the cytoskeleton, and related more specifically to the situation on fibrous and finely grooved substrata, as opposed to ones that are smooth but curved. They reasoned that since the adhesion plaques formed by most fibroblasts are elongated in the direction of spreading, often being several microns long and only one micron or less in width,

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then the formation of such adhesions would be hindered on substrata having grooves or fibers with lateral spacing of a few microns or less. Although the cells would be able to form adhesion plaques behind the parts of their margins facing along the axes of the grooves or fibers, formation of such adhesions would be difficult or impossible in directions oblique to this axis. As these authors showed, and I can attest from parallel studies of my own, fibroblasts will align strongly in response to grooves that are exceedingly narrow, shallow, and closely spaced. Carborundum and optical rouge are sold in a wide range of different grain sizes for the purpose of lens and mirror grinding by amateur astronomers: by directionally wiping these different abrasives across glass and polystyrene surfaces, one can easily make substrata having many parallel grooves, with the widths and depths of these grooves varying according to which abrasive is used. Fibroblasts orient quite strongly in response to extremely fine grooves with spacings and depths on the sub-micron scale. Buck (1980) also showed that when fibroblasts are cultured on rubber surfaces, and the rubber subjected to repeated cycles of stretching and relaxation, then the fibroblasts will line up perpendicular to the direction of this stretching. I would argue that this is simply due to differential breakage of cell-substratum adhesions. Those adhesions on the sides of the cells directed parallel to the stretching will be subject to much larger stress than those on the sides of the cells perpendicular to this direction. More recent studies of cellular responses to fine grooves have used ultrafine photoengraving techniques and photoresists, originally developed for the manufacture of printing plates, but now used to make computer chips. These methods allow construction of grooved surfaces with precisely consistent geometries and dimensions on the submicrometer scale. This technique has used extensively by Clark and his collaborators (1987, 1990, 1991, 1992, also see Britland et al., 1992, and Curtis Clark, 1990, and Dow et al., 1987). They found that some cells can orient in response to parallel arrays of grooves that are only about a tenth of a micrometer deep, and spaced only a quarter of a micrometer apart. As Dunn has discussed in arecent review (Dunn, 199l), this ability of fibroblasts to orient in response to extremely shallow grooves and ridges is difficult to explain by the actin bending theory that he and Heath had previously suggested. It is hard to see why such extremely fine order striations would be able to produce enough bending of cytoplasmic actin fibers to bias cell locomotion directionally. An important question here is the extent to which the cell’s lower surface bridges over the grooves, as opposed to extending down into them. Dunn and Brown (1986) found some evidence that cells do extend down into shallow grooves, as did Brunette (1986; see also Brunette et al., 1983; Oakley and Brunette, 1993). In these studies, fibroblasts were sometimes able to form focal adhesions to the bottoms and sides of such grooves. In addition, Clark et al. (1991) reported that some fibroblasts orient more in response to deep grooves than to shallow ones of the same width, which would be hard to explain if the cells were merely bridging over the tops. Adam Cur-

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tis has been developing a new type of hypothetical mechanism, based on the idea that sharp bends in substratum shape could locally favor either actin condensation or the formation of new substratum adhesions. This would fit in well with the recent discoveries by Depasquale and Izzard (1987,1991) on these marginal condensation events (see also Izzard and Izzard, 1987; Izzard, 1988). If high contrast DIC observations such as theirs were to be carried out on finely grooved substrata such as those of Clark et al., perhaps the underlying mechanisms would reveal themselves. Recent work by Oakley and Brunette (1993) employed confocal microscopy to study the sequences of rearrangements and realignments of cytoskeletal proteins in human gum fibroblasts plated out onto the surfaces of finely grooved titanium substrata. They report that microtubules are the first cytoskeletal components to align, that these become positioned along the bottoms of the grooves, and that this occurs as early as 20 minutes after the cells have been plated out. Not until 40 minutes to an hour after plating could aligned actin fibers be detected; and not until 3 hours after plating were aligned adhesion plaques apparent. They also note the interesting fact that the early position of the microtubule organizing center was essentially random relative to the nucleus and the direction of cell alignment, as if its role were merely passive. These exciting results seem to point to microtubules themselves as important in detecting substratum alignment as well as controlling morphological responses. Drawing upon the literature of plant cell wall morphogenesis, they suggest that the microtubules’ behavior may be a matter of minimizing shear stress. I believe this would be closely analogous to the hypothesis proposed by Dunn and Heath (discussed above), except that it would be the microtubules rather than the actin fibers that are being disturbed along the direction of substratum curvature. It is intriguing that the current hypotheses all look at contact guidance from the point of view of the inhibition of cell extension in the directions perpendicular to the fibers or grooves. Although such inhibition definitely occurs, perhaps more attention should be given to the converse, that is the question whether the process of cell extension may somehow favored along the axes of fibers and grooves. This is often what it looks like, at least when when cells are plated out on aligned collagen or on scratched plastic surfaces. It would seem to be a simple matter to plate cells out along a boundary between scratched and smooth areas, and simply observe the relative rates of spreading, ( I ) along the grooves, (2) across the grooves, and (3) onto the smooth areas. Certainly 1 would be faster than 2, but might it not also be faster than 3? Brunette (1988) has already demonstrated what happens when fibroblasts cross from an area of grooved substratum to an adjacent area in which the grooves were oriented in the perpendicular axis: their leading lamellae reorient even before their cell bodies do. It may be worth noting that fibroblasts respond very differently to fine-order surface roughness (pits and bumps) than they do to striations (grooves and ridges). Roughness actually inhibits fibroblast locomotion at least as strongly as striations seem to favor it (Rich and Harris, 1981). Nor do the dimensions of this surface roughness have to be very large; roughness having sub-micrometer dimensions is

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more inhibitory that larger-scale hills and valleys. If you drag the rounded end of a glass rod lightly across a polystyrene surface, you can produce bands a few micrometers in width whose roughness is barely visible by the highest resolution phase contrast microscopy, but which will totally block the locomotion of nearly all fibroblasts. We named this effect “rugophobia”, and it may contain clues as to the mechanism of contact guidance. The alternative hypotheses discussed above would seem to predict responses to fine-order surface roughness. Adhesion plaques wouldn’t be able to form, while actin fibers would be bent. In the case of Curtis’ forthcoming model, one could ask whether and why an elongate ridge should favor the condensation of actin or adhesion molecules, while spikes and pits having similarly microscopic dimensions seem to have the exact opposite effect. Presumably, the photoengraving techniques of Clark et al. could be used to produce substrata with two oblique or perpendicular sets of grooves. Would fibroblasts be as unable to spread on this as they are on a roughened surface? Or might it simultaneously favor spreading in both perpendicular directions? Incidentally, macrophages are very sensitive to contact guidance along fine grooves, but respond to roughened surfaces in exactly the opposite way as fibroblasts-macrophages actually accumulate on the roughened areas and refuse to crawl off of them (Rich and Harris, 1981)! As discussed below, macrophages also respond peculiarly to voltage gradients, where they move toward the positive pole rather than the negative one (Orida, 1980). These are a few of many puzzling cases in which the directional responses of macrophages are exactly the opposite to those of fibroblasts. It is hard to imagine what the causes might be, much less what single difference in cellular properties could explain all three differences in cell behavior. Nevertheless, for any possible molecular mechanism, either for contact guidance or these other two polarization phenomena, it may well be that the oppositeness of the macrophages’ responses can provide the proverbial “exception that proves the rule.” As regards the lower limit of a fibroblast’s sensitivity to the shape of its substratum, some of the most interesting studies were those of Murray Rosenberg (1962, 1963). These used the “Langmuir trough” method as described by Blodgett (1935) to coat glass surfaces with multiple monolayers of a long-chain fatty acid. The fatty acid is first allowed to form amonolayer on the surface of a small dish of water; then the slide or cover slip is dipped very slowly down into the water, lifted out again, dipped in again. If everything is done correctly, a single monolayer of the fatty acid, on the order of only 2 nanometers thick (one five-hundreth of amicrometer), will be deposited onto the glass surface each time it traverses the water surface. This is not quite as easy as it sounds, as the glass surface has to be extremely clean, the water has to be very pure, and the rate at which the glass is raised and lowered should be very steady and very slow (in the range of millimeters per minute: you need some kind of electric motor to do it right). My former colleague Mark Leader and I were allowed by the UNC Chemistry Department to use their facilities to replicate Rosenberg’s classic experiments. The

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most interesting of his experiments involved depositingjust a few layers of behenic acid, then scraping narrow lines (about 10 micrometers wide) through these layers, and then depositing more layers of behenic acid on top of the previous layers and the scratches. What made his results so interesting was his observation that when fibroblasts were then cultured on these surfaces, the fibroblasts accumulated at the sites of the scratches and lined up along them. The fibroblasts did this even when the scratches had been made after the deposition of only a very few layers of the fatty acid, and even though this scratching had been followed by the deposition of dozens of further layers. These results seemed to indicate either that the cells were responding to grooves only tiny fractions of a micrometer in thickness, or that the cells were capable of some kind of long range sensing of the slight differences in substratum thickness. Among researchers on cell motility, Rosenberg’s remarkable discoveries were sometimes compared with the fairy tale about “the princess and the pea”, in which true royalty is identified by the abiliiy to sense a single pea even through a dozen mattresses stacked on top of one another. How fibroblasts might be able to sense shallow grooves through many overlying monolayers was long a topic of inconclusive debates. Our own attempts (A. Harris and M. Leader, unpublished) to replicate these results pointed toward an explanation different than any considered previously. The answer hinges on a subtle aspect of the Langmuir trough method that seems not have occurred to anyone, namely that each successive layer of fatty acid is deposited with a reversed polarity, that between one dip and the next it is the hydrophobic face that forms the surface, and thus scratching down to the underlying glass creates a discontinuity, a hydrophilic line surrounded by a hydrophobic background. The first, third, fifth and successive odd-numbered layers are oriented with their hydrophilic, carboxylic acid surfaces facing toward the glass, and their hydrophobic faces outward. The second, fourth and other even numbered layers have the opposite orientation. Hydrophilic surfaces face toward other hydrophilic surfaces, and hydrophobic surfaces likewise face toward other hydrophobic surfaces. As pointed out by both Langmuir and Blodgett, because glass itself is hydrophilic, this means that the very first layer has its hydrophilic layer facing down toward the glass surface. This first layer is deposited when the glass surface is lifted out of the water after the first dip. The significance of all this for Rosenberg’s experiments may not yet be altogether obvious. The crucial point is that when you have deposited one or more layers of fatty acid onto glass, and then scratch this surface while it is out of the water, then the outermost layer will therefore always be oriented with its hydrophobic surface outward (Figure 10). This means that when you scratch down to the glass through however many fatty acid layers you have deposited, you create a hydrophilic line surrounded by a hydrophobic surface. This, by itself, accounts for the ability of fibroblasts to detect, accumulate and align along scratches. But what about the other experiments, in which Rosenberg made the scratches but then dipped the scratched monolayers in and out many more times? The point is that

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figure 10. Diagrammatic cross section of the sequential deposition of fatty acid monolayers onto a glass sheet by the Langmuir trough method. Because glass surfaces are hydrophilic, the first layer of fatty acid is deposited acid-end-downward when the glass is pulled out of the water the first time (c). When it is dipped the second time (d), a second layer of fatty acid is then deposited with its hydrophobic surface facing the hydrophobic surface of the first layer. Then when the glass is pulled back out the second time (d), a third layeroffattyacid isdeposited, like the first layer with its hydrophobicend facingout away from the glass. This means that scratches made between dips create a hydrophilic area (of uncoated glass) surrounded by the hydrophilic face of a fatty acid sheet. Nor can subsequent dips be expected to correct this situation. These considerations suggest a simple explanation for Rosenberg’s classical experiments showing that fibroblasts accumulate in the scratched areas.

monolayer deposition is not at all like applying a coat of paint. For one thing, it is much more sensitive to local surface properties. The reversal of polarity at the scratch site means that further monolayers cannot be expected to deposit over the hydrophilic strip you have created. The principles of monolayer formation predict that this scratched area will just be skipped over, in the same way that no monolayer is deposited when the glass first enters the water. In order to cover the scratch, the additional monolayers would either have had to reverse their polarities (flip upside-down) at the scratch site, or they would have to bridge passively across the gap. Such bridging would require the hydrophobic ends of the behenic acid molecules to stick to the hydrophilic glass. Even in the former case, you would still be left with a hydrophobic surface surrounding a hydrophilic strip where the scratch had been. I conclude from all this that the accumulation of cells at the scratch site is not due to any long range effects, nor to the depth of the groove, but merely to the scratched areas not really having been coated with any of the subsequent layers of fatty acid, and remaining as uncoated hydrophilic glass (Figure 11). Although our observa-

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Figure 71. Low magnification view of the accumulation and alignment of chick heart fibroblasts along a pair of prependicular scratches in a monomolecular layer of behenic acid deposited onto the surface of a glass cover slip. The fibroblasts are preferentially adhesive to the underlying glass, uncovered by the scratches, in preference to the intact layers of behenic acid.

tions matched Rosenberg’s (in the sense that fibroblasts did accumulate and align in the scratches), we believe this was because subsequent layers skipped the scratched areas. By watching the meniscus through a hand lens during deposition, we could see differences in its shape at the scratch sites. Likewise, observations of the coated surfaces with an interference reflection microscope indicated that the scratched regions remained uncoated even after many subsequent dips. I therefore conclude that these classical results were essentially artifactual.

IV.

CONTACT INHIBITION AND CONTACT PARALYSIS

When a leading lamella of one fibroblast crawls into contact with another cell, there is usually an inhibition of its further locomotion in that direction. The result is sometimes for the fibroblast’s locomotion to be stopped altogether, at least for a while; but more usually the result is a change in direction, away from the site ofcontact with the other cell (Figure 12). Especially when the collision is “head on,” the fibroblast’s leading margin stops its forward advance, becoming converted to an inactive state (converting what had been a type A margin into a type B margin). In these cases, the cell as a whole becomes temporarily immobile, but then usually reverses direction. All such events are examples of the phenomenon of contact inhibition, originally discovered by Abercrombie and Heaysman (1953, 1954). In addition to these changes in directions and speeds of locomotion, one usually can also observe inhibition of whatever ruffling movements or protrusion of blebs

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Figure 12. Part of a single frame from a 16-mm time lapse film, showing a high magnification view of the occurrence of contact inhibition (and also contact paralysis) beween the advancing margins of two 3T3 fibroblasts.

had been occumng along the parts of the two cells’ margins where the collision occurred (Abercrombie and Ambrose, 1958). This is called “contact paralysis”; or at least it should be; people too often lumpeverything together, using the term contact inhibition interchangeably for both the inhibition of ruffling and for the inhibition of locomotion itself. This term has even been applied to the inhibition of cell growth in division that often occurs in crowded cultures. Such usage implies that these different phenomena share common mechanisms, and reactions against this carelessness have led many to the opposite extreme, the assumption that the mechanisms are not related. We shall be careful with these distinctions here, despite my own expectation that close causal linkages will eventually be found, not only between contact inhibition of locomotion and contact paralysis of ruffling and other cell surface movements, but even between these phenomena and the inhibition of cell cycling in crowded cell populations. Abercrombie and Heaysman’s (1953, 1954) original discovery of contact inhibition was based on a long series of meticulous statistical analyses of cell positions and speeds of fibroblasts’ movement in time-lapse films. In particular, they studied how cell speeds varied as functions of local cell population densities. In one set of experiments, fibroblasts crawling radially outward from tissue explants were shown to be deflected laterally where two such explants were placed close to one another. In otherexperiments, overlaps between fibroblasts were shown to be much less frequent than they should have been if cell positioning were entirely random, and it was found that average speeds of fibroblast locomotion were inversely proportional to the number of other cells with which a fibroblast is in contact. These and related studies have been reviewed in detail by Abercrombie (1967, 1970), by Heaysman (1978), and by Harris (1 974).

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It is important to emphasize that Abercrombie and Heaysman never claimed that complete stoppages of locomotion result from contacts, only that locomotion is reduced and its direction changed. For example, fibroblasts having contacts with as many as six immediate neighsors (most of which would therefore be completely surrounded by other cells) nevertheless continued to crawl at about half the speed of equivalent fibroblasts that had no cell-cell contacts. Unfortunately, their work has too often been misunderstood and over-simplified by others, who assumed that fibroblast locomotion is entirely prevented by cell-cell contact. The essential point is that locomotion is slowed and redirected: in other words, its polarity is changed. Subsequent studies showed that at least some cancerous cells have a much reduced sensitivity to contact inhibition (Abercrombie et al., 1957), suggesting that the invasiveness of cancerous cells may be partly attributable to an abnormally increased ability to continue locomotory activities (those characteristic of the type A margins) directly adjacent to cell-cell contacts. The general idea is that the function of contact inhibition would be to slow or redirect locomotion by normal cells in proportion to their contacts with other cells, thus allowing them to crawl into wounds or other gaps, but not otherwise to move excessively. Conversely, more recent studies by Paddock and Dunn (1986) suggest that the locomotory activities of some cancerous cells may actually be stimulated where they make contact with fibroblasts . An analogous sort of “reverse contact inhibition” also occurs with some macrophages, in that they accumulate preferentially under fibroblasts (Harris, 1974). Those lines or lunds of cells that are susceptible to contact inhibition are usually spoken of as being “contact inhibited,” and those with less susceptibility, such as various kinds of white blood cells and cancerous cells are said to be “non-contact inhibited.” The greatest obstacle to further progress has been not knowing exactly which intracellular events are being inhibited, or otherwise altered, so as to slow and redirect locomotion (much less why normal fibroblasts should be so much more sensitive to this inhibition than are either leucocytes or cancer cells). The formation and backfolding of lamellipodia (i.e., “ruffling”) are not the only types of cell surface movements that have been shown to be inhibited in areas of cell-cell contact. Bleb protrusion has also been found to undergo a similar paralysis, as has phagocytosis (Vasiliev et al., 1975), as well as the retrograde transport of such labels as Concanavalin A (Vasiliev et aI., 1976). &, as was first pointed out by Trinkaus et al., (1971), contact paralysis is usually very tightly localized to the actual region of cell-cell contact. Paralysis does not spread to the adjacent areas, which often continue to ruffle or bleb indefinitely; this localization can be taken as evidence that the mechanism of inhibition is mechanical in nature, rather than being based on some kind of diffusible chemical. At the time of the original demonstration of contact paralysis by Abercrombie and Ambrose (1958), it was strongly suspected that the cell surface movements being paralyzed represented some kind of peristalsis, such that these movements of the cell surfaces would be a necessary part of the mechanism for exerting propulsive forces. It thus seemed to make sense that inhibiting these surface move-

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ments would directly cause inhibition of locomotion: like stopping the wheel of a car from turning. Certainly most people who are shown time lapse films of contact paralysis get a subjective impression that the phenomenon represents some kind of “turning off’ of the cellular motors responsible for locomotion. As naive as the peristalsis idea has proven to be, this subjective impression may not be so wrong after all. What is actually being inhibited is more likely to be polymerization of actin monomers into networks, or rearward transport of tine proteins, or something along those lines. A frequent result of contact inhibition is that fibroblasts tend to become monopolar along boundaries between high and low population densities. In other words, when a fibroblast has many neighbors on one side, and few or no neighbors on the opposite side, the contact inhibition of its leading lamellae along the crowded side tends to polarize locomotion in the direction of the fewest neighbors. This occurs along the boundaries of explants and where cells have been scraped away from monolayers that had previously been confluent. The effect is the locomotion of fibroblasts and other cells into areas of lower population density. Inside the body, this would help to fill in wounds, which is presumed to be one of the main reasons for the evolution of this property. Anyone wanting to study fibroblasts that are nicely polarized and monopolar would do well to use cells with strong susceptibilities to contact inhibition, and then concentrate their attention along boundaries between areas of high and low population densities. Contact inhibition can also contribute to the alignment of bipolar fibroblasts into tracts within which all the cells are nearly parallel to one another. Fibroblasts explanted from certain tissues (the kidneys), including those of certain established cell lines (BHK), are especially prone to this type of alignment, which has been most extensively described in the studies of Elsdale (1968). The experiments of Erickson (1978a) showed how this alignment could be explained in terms of contact inhibition. As she demonstrated, when one fibroblast advances obliquely into contact with the side of another, then one side of its leading lamalla contacts the other cell first, and is therefore inhibited, while the other side of this same leading lamella continues its outward advance. This inhibition of one side of the leading lamella, but not the other, tends to make the advancing cell turn slightly away from the contact, thus bringing itself more nearly parallel with the side of the other cell. When the contacted side of the leading lamella ceases forward spreading, but the noncontacted side continues to crawl forward and pulls somewhat sideways, away from the contact site, then the result tends to rotate the axis of the crawling cell toward the direction parallel to the axis of the cell that it has contacted, as diagrammed in Figure 13. This makes mechanical sense, especially if we imagine that each side of a leading lamella pulls slightly laterally, somewhat like the side horses of a Russian troika, so that weakening one side will rotate the direction of maximum force exertion, away from the weakened, inhibited side. Indeed, subsequent observations with silicone rubber substrata do indicate that individual leading lamellae of fibroblasts frequently exertjust this sort of convergent force; in particular, some of their

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Figure 13. How contact inhibition can cause progressive alignment between fibroblasts, based on the observations of Carol Erickson. When one cell collides with another at too large an angle (top left), it reverses direction (top right). But when the collision occurs at a shallow angle (bottom left), the result is inhibition of its leading lamella on one side more than the other, so that the colliding cell turns slightly into a parallel direction (bottom right).

compression wrinkles are oriented perpendicular to the nearest advancing cell margin. As already mentioned, the mechanistic relation between contact paralysis and contact inhibition remains very uncertain. We very much need to find out exactly what molecular events are blocked near the contacts, what mechanism blocks them, and how cell-cell contact is able to switch what we are calling type A margins into inactive or retreating (type B or even type C ) types of behavior. As already mentioned, a likely explanation is that actin assembly somehow becomes locally blocked; but experimental proof of this is not yet available. Michael Abercrombie himself favored the idea that cell-cell contact induces some kind of increased cytoplasmic contractility, so that the cells are induced to pull apart. This would explain the frequent retraction of cells soon after contact (called “contact retraction”), which is usually away from the site of the cellkell contact, but is sometimes directly toward the contact site. Using flexible substrata and other criteria, I was never able to find direct evidence for such any consistent strengthening of fibroblasts’ longitudinal contractility following contact with another cell, and proposed the alternative hypothesis that contact retraction may instead be caused by induction of weakening of cell-substratum adhesions near sites of cell-cell contact (Harris, 1973b). The idea was that this weakening of cell-substratum adhesions would permit the fibroblast’s pre-existing (and ever-present) contractility to cause retraction. However, this would not explain why cells sometimes contract toward the site of contact inhibition, and when Abercrombie and Dunn (1975) used interference reflection microscopy to look for evidence of the inhibition of cell-substratum adhesions that I had postulated, they failed to find it. They used mercury arc illumination, however, the extreme brightness and near-ultraviolet components of

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which interfere with the normal formation and breakage of cell adhesions. (This is itself a phenomenon worthy of more study: for example, leucocytes illuminated by an arc become immobilized and trapped in the light beam, because they cannot detach their old adhesions!) My own experience is that when interference reflection is carried out with sufficiently dim illumination from ordinary tungsten incandescent lamps, you can often see marked inhibition of cell-substratum contact areas just distal to sites of contact inhibition. Obviously, this is a topic deserving further study. A more important and controversial question concerns overlapping between cells, and the extent to which increased overlapping among cancerously transformed fibroblasts is really a consequence of decreased sensitivity to contact inhibition. It had been long assumed that overlapping between adjacent cells could only be the result of active locomotion of one cell up onto the surface of its neighbor, which would necessarily constitute a failure of contact inhibition. However, it was eventually realized that only a small fraction of the lower surfaces of most fibroblasts is actually adhering to a glass or plastic substratum (Harris, 1973a), with what we are calling the “type C” margins offering relatively unobstructed avenues to the type A margins of neighboring cells to crawl underneath (Boyde et al., 1969). Detailed analyses of time lapse films of polyoma virus transformed fibroblasts revealed that most apparent overlaps are really cases of “under-lapping”, in which the advancing cell has continued to use the glass or plastic as its substratum, and has merely crawled into the unoccupied area underneath its neighbor’s type C margins (Guelstein et al., 1973;Di Pasguale and Bell, 1974;Bell, 1977; Erickson, 1978b). In those cases where an advancing type A margin of one cell encountered a type A margin of one of its neighbors, both ruffling and further spreading were found to be inhibited, preventing all but slight overlapping. Bell (using 3T3 fibroblasts) and Erickson (using BHK fibroblasts) did find somewhat more apparent overlapping (“crisscrossing”) between virus transformed fibroblasts than among untransformed ones, although with more nuclear overlapping between the transformed 3T3 cells than between the transformed BHK cells. In both cases, however, the increased overlaps seemed to be due to underlapping, and also to retraction clumping which occurs when one cell pulls loose from its substratum adhesions and retracts onto the surfaces of its neighbors. Erickson, for example, found not even a single case of true overlapping, in the sense of one cell crawling up onto and across the upper surface of another, using the other cell as a substratum, neither among the transformed or the untransformed BHK cells. Such true overlap does seem quite rare, but can occur, at least with very highly transformed fibroblasts; I have seen L cells and sarcoma- 180cells crawl across the upper surfaces of untransformed fibroblasts (see Figure 9 in Harris, 1982). Erickson also demonstrated, however, a very dramatic difference between the transformed and the untransformed BHK cells in the outcome of what one could call “front to side” collisions (where type A margins encounter type C margins). In the case of such encounters between untransformed cells, fully 87% of the contact events resulted in adhesion between the cells, with 64%changing or reversing direction, and some de-

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gree of underlapping in 36%. This was in contrast to the front to side encounters between her transformed BHK cells, in which only 9% resulted in adhesion, 7% resulted in a change in direction, and 93% resulting in underlapping! These results obviously cast a shadow over the naive but appealing view that transformed cells overlap more because of their reduced sensitivity to contact inhibition, and that this is part of the behavioral basis of their greater invasiveness inside the body. On the other hand, we should not be too quick to discard that view altogether. Even if transformed cells have not actually acquired an ability to use the surfaces of other cells as substrata, the work of both Bell and Erickson seems to show big increases in these cells’ ability to continue crawling forward in areas where they are closely underlapping and touching other cells. After all, equivalent nontransformed cells would quickly have undergone both contact paralysis and contact inhibition. The difference may lie only in a reduced ability to form new cell-cell adhesions, or perhaps in an increased ability to continue “type A” margins adjacent to cell-cell contacts, but these or other changes could also contribute to increased invasiveness inside the body. This is one of many examples where an increased understanding of the molecular nature of the events occurring along type A margins should soon provide a new perspective on a large body of older knowledge.

V.

GALVANOTAXIS: EFFECTS OF ELECTRIC FIELDS ON FIBROBLAST POLARITY

Fibroblasts react to voltage gradients by slowly elongating in the direction perpendicular to the axis of the gradient, and by reorienting in this direction if they had previously had a long axis in some other direction (Figure 14). This perpendicular response is in contrast to the behavior of at least some epithelial cells, which respond to electric fields by crawling toward the negative electrode, not to mention nerve cells, whose response is extension of growth cones toward the negative electrode. It is also in contrast to macrophages, which despite resembling miniature fibroblasts in shape and most behavior, have been shown to respond to voltage gradients by crawling directionally toward the positive electrode (Orida, 1980). Another cell type showing this type of directionally-reversed galvanotaxis is the osteoclast, which serves to break down bone; this is in contrast to the bone-depositing osteocytes, which migrate toward the cathode (Femer et al., 1986). It is not known whether these differences have anything to do with the opposing roles of these two cell types in skeletal morphogenesis, nor if there is any relation to the widelyhypothesized role of electric fields in bone growth. Note, however, that osteoclasts are believed to be formed by the fusion of macrophages with one another. It would be interesting to know how many of the different macrophage-like cells of the body, such as microglia in the nervous system and Kupfer cells in the liver, also move preferentially toward the positive ends of electric gradients.

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Chick heart fibroblasts, cultured on a silicone rubber substratum, that have oriented in response to a voltage gradient. The axis of the voltage gradient is perpendicular to the direction of cell alignment.

Figure 14.

It remains to be discovered exactly what differences in membrane or cytoskeletal components, or other differentiated cell characteristics, cause such differences in &rectionality of response. Besides fibroblasts, it has been found that neural crest cells, among others, respond by perpendicular elongation. To a variable degree, fibroblasts also crawl slowly toward the negative electrode. In my experience, however, the perpendicular orientation is by far the more dramatic and consistent effect. The fibroblasts’ contractility likewise becomes directed perpendicular to the voltage axis. The morphological reorientation occurs gradually over 20 to 30 minutes, and has now been observed in cultured fibroblasts from a wide variety of sources, including established cell lines and cells explanted from embryos. In most studies of this phenomenon, the steepness of the voltage gradients has been in the range of a volt per millimeter (equal to a millivolt per micrometer), which is considerably higher than is found within the tissues of developing embryos. Nevertheless, Nuccitelli andErickson (1983) have shown that fibroblasts can also orient in response to voltage gradients only a tenth this steep, and that is in the range of field strengths actually found in developing embryos. Their results thus suggest that galvanotaxis could be a functional mechanism of tissue morphogenesis. It is important to stress that these galvanotactic orientations depend on reasonably steady, long-lasting voltages, acting in a certain direction and not reversing polarity. Thus, while much larger transient voltages occur in many tissues, for example as a result of rapid stressing of cartilage, tendons and bone, fibroblasts are only found to reorient in response to sustained uni-directional (direct current) voltages. As is discussed below, fibroblasts do not seem to show any response to alternating current voltages, which has important implications not only for the possible functional significance of galvanotaxis. but also for its subcellular mechanism.

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Galvanotaxis by tissue culture cells, specifically neural growth cones, was first reported by S. Ingvar in 1920. However, Paul Weiss (1934) was unable to repeat those results, and pronounced that whatever effects Ingvar had observed must really have been secondary consequences of reorientation of extracellular fibers of plasma clot around the electrodes. Weiss proposed that contact guidance, rather than galvanotaxis, must have caused Ingvar’s cells to align along these extracellular fibers. Considerably later, Weiss and Scott (1963) reported their unsuccessful attempts to produce any changes in fibroblast locomotion by means of imposed voltages, even though these appear to have been well within the range of field strengths subsequently been shown to produce alignment. Such was Weiss’ influence, however, that the whole topic of tissue cell galvanotaxis went into suspended animation until the early 1980s,when it was more or less simultaneously rediscovered by several independent groups of researchers (Hinckle et al., 1981; Luther et al., 1983; Nuccitelli and Erickson, 1983; Cooper and Keller, 1984; Erickson and Nuccitelli, 1984; reviewed by Robinson, 1985; see also Soong et al., 1990). Among other things, these workers found that it is essential to keep the culture chamber quite thin; otherwise too much heating will be produced by the electrical current, thus killing the cells before they can reorient. This heating effect may explain Weiss’ inability to confirm what Ingvar had discovered. Fibroblast reorientation seems to occur predominantly by the withdrawal of any leading lamellae that are oriented parallel to the voltage gradient, followed by the extension of leading lamellae along the axis perpendicular to this gradient (Harris et al., 1990).Enlargement of preexisting leading lamellae oriented in this perpendicular direction also occurs. Variations are sometimes seen in the extent of this reorientation, however, sometimes for no very obvious reasons. I have seen whole cultures in which practically no orientation could be produced, as well as groups of recalcitrant cells within other cultures in which the remainder had all reoriented; my impression is that published papers in this field tend to understate the degree of such unexplained variations. Apparently, the cells’ locomotion has to be reasonably active for reorientation to occur. Unusually immobile cells, and especially quiescent cultures often won’t respond. I have also noted special insensitivity on the parts of those fibroblasts that happen to be adhering to one another more than to the glass or plastic substratum. Such unexplained variations are annoying, of course, but they may also serve as clues for the design of future experiments. As to the mechanism of fibroblast galvanotaxis, we need to ask two distinct classes of question. First, how does the cell “feel” the electric field: for example is it the electrophoresis of charged molecules in the plasma membrane, or is it the distortion of the transmembrane “resting potential” difference in voltage between the cytoplasm and the surrounding medium? The second type of question concerns the cells’ responses: for example, do the leading lamellae contract more strongly on the sides facing up and/or down the voltage gradient (and thus preferentially pull themselves loose from the substratum)? Alternatively, are the cell-substratum adhesions differentially weakened on the sides of the cell facing up and down this gradient?

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Or, as a third alternative, is the outward spreading of leading lamellae somehow favored on the sides of the cells not facing either pole of the electric field? It is essential to realize that these two classes of question are logically distinct. A cell’s preferential withdrawal of extension of lamellae could result from either electrophoretic effects or changes in transmembrane voltage. Likewise, it might be useful to reflect that differentiated cell types might differ from one another in the mechanisms of their responses to electric fields, or even that mixtures of effects may occur. Nerve cells might use one mechanism, and macrophages another. Only fibroblasts will be considered here, however. Note also that I am assuming that cells detect voltage gradients, rather than detecting the actual flow of electrical current that results from this voltage. Some who have written on galvanotaxis, Weiss in particular, seem to have been making the opposite assumption. But my reasoning is that the permeability of the plasma membrane to electric current is so much lower than that of either tissue culture medium or cytoplasm, that very little current can be expected to flow through the cell bodies. Likewise, when you feel the wind blowing past you, or when a house blows down in a hurricane, what is felt are the differences in air pressure (equivalent to voltage differences) and not the cubic volumes of air moving past (equivalent to electrical current amperage). In the case of a cell culture, the amount of current flowing has more to do with the electrical resistance of the surrounding medium (including its dimensions) than with any property of the cells. I do not believe it would be difficult to make tissue culture media whose electrical conductivities differed over a fairly wide range, so someone might wish to test the effects of different amounts of current with the same rate of change of voltage, as compared with the converse. It is also important to realize that both the electrophoresis of membrane components and also the biasing of the transmembrane potential will necessarily occur when you put a fibroblast in a voltage gradient. Neither effect can help occurring to some degree. Thus, the question is not which one occurs, but which one the cells are responding to when they reorient their axes. It is inevitable that any charged material on the cell’s surface will feel a displacement force proportional to its own charge and to the steepness of the voltage gradient being imposed. The extent to which this force will be resisted by the viscosity of the membrane, or overcome by other forces, are separate questions. But there will be a force. Likewise, because the electrical resistance of the plasma membrane is enormously larger than the resistances of either the cytoplasm or the surrounding medium, we can also be certain that only a negligible voltage gradient can be maintained between one end of a cell and the other, and similarly that very little current will flow there. This implies that we can discount the electrophoresis of any cytoplasmic materials. Just as in nerve and muscle cells, the transmembrane voltage of fibroblasts results primarily from the differences in the concentrations of potassium ions (high inside the cell and low outside the cell), combined with an appreciable permeability of the plasma membrane to potassium ions leaking through it out of the cell. Because none of the other ions are as free to leak though the membrane as is potassium,

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and because potassium ions are positively charged, the net result is a voltage (outside positive, inside negative) that is proportional in strength to the log of the inside/outside ratio of potassium concentrations. This voltage difference tends to be smaller in fibroblasts than in nerves or muscles; and is also not subject to active reversal by sodium-driven action potentials. In an externally-imposed voltage gradient, the much lower conductivity of the plasma membrane relative to either the medium or the cytoplasm means that the voltage gradient inside the cell will be much less steep than the one outside. This leads us to the important conclusion that the transmembrane voltage will have to vary from point to point along the cell surface, as you move along the axis of the external voltage gradient. Note that a fibroblast’s transmembrane potential of 30-40 millivolts or so is in the same range of magnitude as the voltage difference that is produced across the 30-40 micrometer width of a fully spread fibroblast by an imposed voltage gradient of about one millivolt per micrometer. On the other hand, it would be reassuring if someone were actually to measure the transmembrane voltages of different fibroblasts and fibroblast lines, to see how closely these match the steepnesses of the voltage gradients imposed on them. It seems obvious that when the voltage is initially lower inside a cell than outside, and an external voltage gradient is then imposed on the surroundings, but (because of the relative lack of permeability of the plasma membrane) the cytoplasmic voltage remains nearly constant throughout the cell, then the result must be for the end of the cell facing the positive electrode to become hyperpolarized, while the opposite side of the cell (in the direction facing the negative electrode) becomes depolarized (see diagram Figure 15). It would be useful for someone to use the fluorescent dye methods referred to above to confirm that the plasma membrane of a fibroblast subjected to a voltage

-

Outside cell Inside cell

........

Voltage

.:............... .:.:: . .:.:.:.:.:<->; j:i ...................................

:

-

. . .

...,!

Outside cell Irside cell In a gradient

figure 75. Diagram of the expected distribution of relativevoltages, inside and outside a fibroblast, when it is not in a voltage gradient (top), as compared with when a voltage gradient i s imposed on the culture medium around it (bottom). The point i s that the end of the cell toward the negative electrode will become depolarized while the end toward the positive electrode will become hyperpolarized.

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gradient really does vary according to this expected pattern. It would also be useful to know (from direct measurements, rather than theoretical calculations) which part of the cell remains at its original voltage, whether this is something that varies from one cell (or from one cell type) to another, and what kinds of ion fluxes occur as aresult of these imposed voltage gradients. For example, are we to suppose that potassium just continues to leak out through the depolarized parts of the plasma membrane, or that equal amounts of this ion are drawn in through the hyperpolarized areas? An especially important question is to what extent fibroblast membranes are like those of muscles in responding to depolarization by locally increased permeability to calcium ions. In a voltage gradient, this would be expected to occur at the end of the cell facing the negative electrode. Such localized changes in ion concentrations in the cytoplasm have been suggested as possible causes for morphological alignment. But although several laboratories have tested this possibility using calcium-sensitive probes, I believe with negative results, I am not aware that any such results have yet been published. One experimental approach to the question of whether the cells are responding to depolarization or to electrophoresis is to compare the effects of alternating currents, direct currents, and intermittent currents. Nuccitelli and Erickson (1983) discovered that cells pay no apparent attention to alternating current such as that from the power mains, which reverses its polarity 60 times per second. This approach was extended by Harris et al., (1990) who used an electronic timing device to reverse the direction of the voltage gradient at longer intervals: specifically once per second, once per ten seconds, and once per minute. Our idea was that 60 times per second might somehow be too fast for the cells to notice, or for cytoplasmic changes to take effect, but that longer durations would allow sufficient time for the opposite sides of the cells to complete their various hyperpolarizations, depolarizations and ion leakages (if any). Fibroblasts might then be expected to reorient their morphological axes, especially since this reorientation is essentially symmetrical with respect to the voltage axis. In other words, even though the cells would first be depolarized at one end, and then hyperpolarized at that same end and depolarized at the opposite one, again and again, time after time, they might still react in the same way as to adirect current, by withdrawing the leading lamellae at both the hyperpolarized and the depolarized sides. After all, if withdrawal is the response both to depolarization and to hyperpolarization, then why shouldn’t the response be the same to a regular alternation between these two states? In fact, what we found was that fibroblasts did not align in response to even these very slow alternating voltages. Instead, they continued to crawl about for 2 hours or more, with apparently normal morphology and behavior. These same cells then required only the usual 20 to 30 minutes to align once the reversals of polarity were stopped and the direction of the voltage was held constant. As a response to Cooper’s helpful suggestion that the ion pumps at each extremity of a fibroblast might effectively be “bailing out” whatever ions had leaked in during the periods of opposite polarity, we then did equivalent experiments with the same timer in which the volt-

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age was turned on for one minute, then off for the next minute, then on again with the same original polarity, then off again, and so forth. This would allow the “bailing out” to occur, even though subjecting the cell only to voltages of one of the two polarities. Nevertheless, the fibroblasts did reorient to these intermittent voltages, and in little more than their usual 30 minutes. Taken together, these results indicate that reversing the direction of a voltage has the consequence of undoing its previous effects (i.e. not just producing a different effect, but actually reversing that produced earlier by the voltage having the opposite polarity). This result, of course, is exactly what would be predicted by the hypothesis that the galvanotactic signal is electrophoretic in nature. Conversely, I do not see any obvious way to explain these responses in terms of ion leakages. If you had a ship and first let water leak in one side, then let water leak in the other side, and repeating this indefinitely, you could keep the ship on an even keel, but would eventually sink it. More research is needed, but the current evidence seems to indicate that at least the perpendicular orientation aspect of fibroblast galvanotaxis is most likely to result from the electrophoretic displacement of one or more membrane components into the ends of the cells that face the electrodes. These membrane components then somehow promote retraction of the cell extremities into which they have been accumulated. The second of our two classes of question concerns the physical nature of the fibroblast’s response. Three specific alternatives have been considered: (a) localized or directional strengthening to the contractility of leading lamellae, so that they pull loose from the substratum; (b) localized weakening of the adhesiveness of these leading lamellae to the substratum, so that their existing contractility becomes sufficient to retract them; or (c) favoring the protrusion of those leading lamellae that are oriented perpendicular to the voltage gradient. As a test of the first of these three possibilities (and to some extent also as a test of the third) fibroblasts were cultured on silicone rubber substrata, allowed to spread sufficiently to produce many compression wrinkles under themselves, and then subjected to a voltage gradient. Not only the wrinkles in the rubber, but also markers on its surface were observed to see whether any changes in the strength or directionality of cell contraction occurred in response to the imposed voltage. In the most sensitive of these experiments, tiny rectangular islands of silicone rubber were cut out of the sheet, with only one or a few cells per island, and left floating on the silicone fluid surface. The idea was that even the tiniest changes in direction or strength of cellular forces would be visible as changes in the shapes and dimensions of these islands. But the results were negative. The anticipated increase in contractility along the axis of the voltage gradient simply did not occur. Instead, there was a gradual weakening of the cells’ contractile force in this axis, beginning after 10 or 20 minutes and continuing until the cells had completely reoriented and were exerting little or no tension in this direction. Accompanying and following this slow reorientation, there was a gradual increase in substratum wrinkles running transverse to the cells’ new axis, indicating a gradual increase in the contractile force being exerted in the direction perpendicular to

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the voltage gradient. Because this increased force in the perpendicular direction developed so gradually, it was judged to be the effect, rather than the cause, of the morphological reorientation. The results of this experiment thus seemed to point to the second of the three hypotheses: achange in the spatial distribution of each cell’s adhesiveness. Interference reflection microscopy (Izzard and Lochner, 1976) offers a more direct approach to this question of whether fibroblasts respond to voltage gradients by changing the spatial distribution of their substratum adhesions. In fact, the study by Harris et al., (1990) originally included the use of this optical technique, with observations of cell-substratum contacts disappearing on the sides of fibroblasts facing the positive electrode and the apparent formation of broad dark contact areas on the sides of the fibroblasts facing the negative electrodes. In other words, the voltage gradients seemed to favor the formation of new adhesions on the negative side and to favor the breakage of adhesions on the positive side. This process was recorded by time-lapse video, and observed throughout entire cultures in several replicate experiments. Nevertheless, in later experiments, the formation of these broad, dark contact areas inexplicably failed to occur, even though the fibroblasts did undergo the usual alignment perpendicular to the voltage gradient. We were unable to determine what difference in the cells or the culture conditions was responsible and omitted the interference reflection observations from the final version of the paper. I mention this in the hope that some reader of this review may be stimulated to reexamine this phenomenon. As was for so long true of galvanotaxis itself, there is a tendency just to ignore the very existence of phenomena so long as some of the variables that control them remain unknown.

VI.

MICROTUBULES IN THE CONTROL OF FIBROBLAST POLARITY

Several lines of evidence indicate that microtubules must play some important role in governing the polarization of fibroblasts. For one thing, cytoplasmic microtubules are generally oriented along a fibroblast’s long axis, when it has one; and reorientation of a fibroblast’s axis (for example in response to voltage gradients, see Harris et al., 1990) is accompanied by reorientation of the microtubules. Furthermore, when fibroblasts are crawling in a definite direction, such as along the edge of a scraped area “wound” or other boundary between high and low population densities, their microtubule organizing centers (MTOCs) usually become positioned between the nucleus and the leading margin (Singer and Kupfer, 1986). The Golgi apparatus also becomes localized in this area. Likewise, there is evidence of preferential stabilization of those microtubules directed toward the cells’ leading margins (Gunderson and Bulinski, 1988). The question is whether these phenomena should be regarded as actual causes of the cells’ directionality and polarity, as opposed to being consequences or byproducts. In other words, does the end of the cell that

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faces toward the lower population density become the front end because the MTOC is located there? Or does the MTOC move there because this end has become the front, for example because of the new patterns of contractile forces within the cytoplasm? For some years, many people in this field seem to have tended to the former conclusion: that MTOC position actually caused directionality, rather than the other way around. More recently, however, observations on the time course of changes in MTOC position have led to a general reversal of opinion (Euteneuer and Schliwa, 1992). In careful studies of the time course of changes during reorientations of fibroblasts along the edges of scraped regions, it was found that the MTOCs do eventually move to the side of each cell between its leading edge and its nucleus; but this change in MTOC position often occurs after the change in direction of movement, implying that it more a result than a cause. In addition, it has been found that when fibroblasts are cultured in a collagen gel, then the position of the MTOC bears no particular geometric direction to the leading edge or direction of the cells’ movement, very much in contrast to the situation when equivalent cells are grown on flat glass or plastic (Schutze et al., 1991). This last observation is quite puzzling from any point of view, but certainly doesn’t support the idea of MTOCs as controlling directionality. Unfortunately, there doesn’t seem to be any way to do the converse types of experiments and force microtubules to reorient in a chosen direction. We might, for example, imagine putting an iron filing inside a MTOC, and then using a magnet to pull it around from one side of a fibroblast to another. Would this cause a corresponding redirection of the fibroblast’s direction of locomotion and morphological polarity? No one knows. On the other hand, it is not difficult to disrupt or even eliminate cytoplasmic microtubules using spindle poisons such as colchicine, vinblastine or nocodazole, and the long term morphological effects of these drugs can be quite dramatic. As Vasiliev and his colleagues first demonstrated, exposure to these drugs tends to reduce or eliminate fibroblast polarity ( et al., 1970; Vasiliev and Gelfand, 1976; see also Vasiliev and Gelfand, 1977; see also Vasiliev. 1991).Fibroblasts treated with these drugs sometimes lose their polarity so completely that they revert to a “fried egg” morphology, in which most or all of the cell margin switches to what we are calling the type A class of behavior, extending 360 degrees around the periphery. An example of such a cell is shown in Figure 16; before treatment with vinblastine this cell had been elongate and bipolar with a narrow leading lamella at each end. The implication of such morphological changes seems to be that fibroblasts need their microtubules to subdivide or polarize different parts of their margins into distinct types of behavior. This result is very much as if intact cytoplasmic microtubules were somehow required to maintain the category of margins we are calling type C (and perhaps also the type B margins). Sensitivity to this morphological effect varies widely among fibroblasts, with those from primary cultures (from explants) not being susceptible, in other words

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Figure 76. High magnification view of a single vinblastine-treated BHK fibroblast. Notice the cell’s nearly circular “fried egg” shape, which is very much in contrast to the usual elongate shape of BHK cells. Notice also the formation of a ring of small blebs, or lobopodia, all concentrated along the cell margin. A few of the characteristic vinblastine-induced crystals of microtubule protein are also visible in the cytoplasm surrounding the nucleus.

undergoing little or no morphological depolarization in response to any of the antimicrotubule drugs. No one knows why fibroblasts recently derived from explants should fail to undergo the morphological depolarization, nor why the effect is so dramatic in certain established lines (BHK cells are particularly good). One plausible hypothesis had been that it might reflect the possession (by insensitive cells) of larger sub-populations of especially long-lived and otherwise stable microtubules, for example those made of detyrosinated tubulin. Recent observations by Middleton et al. (1989) do not seem to support this type of explanation, in that fibroblasts from primary cultures were not found to contain any greater proportions of tubules containing these modified tubulins. In fact, given that fibroblasts from primary cultures continue not to undergo the morphological depolarization, even after prolonged exposures (of days) to substantial concentrations of any of the anti-microtubule drugs, the conclusion seems inescapable that the difference must lie in how the cells respond to their lack of microtubules, as opposed to how completely the cells are deprived of them. In other words, fibroblasts from primary culturesjust must not need microtubules to maintain their polarity. In contrast, fibroblasts from many established lines seem unable to maintain their polarity without microtubules. Maybe polarity is normally maintained by Some kind of combined effort of several cytoplasmic elements,one of which is microtubules, but the other elements are able to do the job by themselves in fibroblasts from primary cultures. Cell lines that have spent much longer in culture, becoming aneuploid and otherwise jaded, apparently lack these other contributors to polarity.

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What seems to be a related effect occurs when fibroblasts treated with antimicrotubule drugs are cultured on “haptotactic substrata”, in particular those substrata having adhesive metal-coated “islands” separated by lines where the substratum is less adhesive. Normal fibroblasts conform only very roughly to the dimensions and boundaries of the adhesive islands, extending lamellae beyond their edges in some places, while retracting well inside these boundaries in other places. But when the same fibroblasts are grown in media containing microtubule poisons, they change behavior so as to conform much more exactly to the adhesive boundaries (Harris, 1973a). Such cells often become exactly square or triangular, taking on the precise size and shape of the island they happen to be adhering to (see Figure 17). As with the “fried egg” morphology, it is as if fibroblasts deprived of their microtubules lack autonomy in controlling their shape. There are several different kinds of mechanisms by which cytoplasmicmicrotubules might contribute to fibroblast polarity, three of which will now be discussed. One such category of mechanism would simply be that the physical strength and resistance to bending of microtubules would provide enough mechanical support to favor elongation and polarity. This class of explanation is frequently encounteredin implicit form, but seems not to have been developed explicitly. A second class of possible mechanism is that polarization results from the transport functions known to be provided by microtubules,using kinesins or dyneins. The best evidence for the second of these possibilitiescomes from recent work by Rodionov et al. (1993) who injected antibodies specific for the motor domains of the protein kinesin. This pro-

Figure 77. Vinblastine-treated fibroblasts cultured on a series of “haptotactic islands” consisting of squares of vacuum-evaporated palladium metal overlying a nonwettable (and relatively nonadhesive) polystyrene culture dish. Notice how the shapes of these cells approximate the artificial shapes of the adhesive islands. This is in contrast to equivalent fibroblasts not treated with microtubule poisons, which also tend to be confined to the adhesive areas, but adopt their usual stellate shapes much more independently of the shapes of the islands themselves.

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tein, of course, is one of those that serves to transport materials along microtubules. The injection of these antibodies did not inhibit the development of normal arrays of cytoplasmic microtubules. But it did mimic some of the morphological effects of colchicine and other microtubule poisons, implying cell polarity does not depend SO much on the presence or absence of microtubules, but rather depends on whether materials can be transported along microtubules. For example, unspecified proteins might be transported to some regions of the cell periphery in preference to other regions, thereby causing these areas to behave differently in respect to contractility, adhesiveness or other properties. In previous work, Vasiliev’s group had studied the disruption of the actin cytoskeleton in response to phorbol ester tumor promoters, which act by overstimulating protein kinase C (Brown et al., 1989; Bershadsky et al., 1990) . A principal result of this over-stimulation is that actomyosin stress fibers break down, and what we here are calling the type A margins crawl forward actively, stretching the cell body out into thin cytoplasmic necks (Dugina et al., 1987). Among the other discoveries made by members of this group was that fibroblasts that had been treated with microtubule poisons became essentially immune to this phorbolinduced redistribution of their actin cytoskeleton (Lyass et al., 1988). Quite reasonably, they interpreted this in terms of the microtubules being needed in some way to transport or otherwise redistribute the actin or other cytoskeletal proteins to new locations and arrangements. Parallel studies by Danowski and Harris had shown that phorbol esters cause a rapid weakening of the tractional and contractile forces exerted by fibroblasts, as revealed by reductions in the wrinkling of rubber substrata on which the cells were cultured (Danowski and Harris, 1988). These studies also found that fibroblasts treated with phorbol esters become able to crawl onto relatively nonadhesive and hydrophobic substrata, to which their adhesions had previously been too weak to permit spreading. This apparently results from their weakened contractility rather than from any increase or other change in their adhesiveness per se. When Danowski (1989) studied whether these effects on contractility might likewise be inhibited by microtubule poisons, following up the work of Lyass et al. described above (1988), she found several very unexpected things. First she confirmed that treating fibroblasts with amicrotubule poison along with the phorbol ester will indeed prevent the usual effect of the phorbol: the actin cytoskeleton is not disrupted. But then she discovered that even if cells have already been treated with the phorbol, and their actin cytoskeletons have already been disrupted, if you then add colchicine or one of the other microtubule inhibitors to the medium, this results in a restoration of the actin cytoskeleton! The cells’ contractility, measured by distortions of rubber substrata, is also restored even though the phorbol ester is still present. In other words, microtubule poisons don’t just prevent the cytoskeletal effects of phorbol esters, they actually counteract and reverse these effects. Obviously, this is not at all what one would expect if the role of microtubules in the process were that of transporting cytoskeletal materials to their new locations. If

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that had been the explanation, then microtubule poisons ought to prevent restoration or normal cytoskeletal patterns, not promote it. An even more surprising discovery (Danowski, 1989) was that colchicine, vinblastine and nocodazole all cause large and rapid strengthening of fibroblast contractility, even in cells not treated with phorbol esters. Within a few minutes of treatment with any one of these three antimicrotubule drugs, fibroblasts increase their contractility to levels two or even three times as strong as they had been exerting just prior to treatment. Danowski discovered this using the (relatively nonquantitative) silicone rubber substratum technique; but by using electronic strain gauges Kolodney and Wysolmerski (1992) were able to confirm her conclusions quite precisely for the case of large numbers of fibroblasts pulling on collagen gels. Danowski also showed that these anti-microtubule drugs promote formation of actomyosin stress fibers, and that not only do these drugs prevent and reverse the disruption of stress fibers by phorbol esters, they also block and reverse the disruption of the cytoskeleton by quite unrelated drugs, including ones acting by altering cytoplasmic concentrations of cyclic AMP. As if all this weren’t surprising enough, when Danowski tried a wide range of concentrations of the anti-microtubule drugs, she discovered that their contractility and stress fiber-promoting effect occurs even at extremely low drug concentrations - so low that many microtubules are visible when the fibroblasts were stained with antibodies against microtubules, and so low that mitosis can still occur (note, the same effects also occur at higher concentrations of these drugs). What all this seems to imply is that what matters is not so much the continued existence of cytoplasmic microtubules, but something more subtle, like their state of dynamic equilibrium. Merely disturbing the microtubules, or their process of assembly, seems to be enough. It somehow favors assembly of actin into stress fibers, thereby magnifying the contractile forces that fibroblasts exert on their surroundings, and preventing (and reversing) the disruption of these stress fibers with TPA and other treatments. To explain the increased contractile force itself, the most obvious interpretation would be that microtubules normally exert a substantial pushing force, which is more than counterbalanced by the pull of the acto-myosin stress fibers. With the removal of these microtubules, the full strength of the actin-based contractility would be revealed. This type of explanation would fit in well with Donald Ingber’s (1993) recent attempts to explain cell mechanics in terms of “tensegrity.” Indeed, given that fibroblast contractile force doubles or even triples in response to colchicine or equivalent drugs, this pushing interpretation would imply that microtubules exert very strong forces indeed. They would have to push with forces of the order of at least 2 or 3 hundreths of a dyne per cell, that being the order of magnitude of the forces that fibroblasts ordinarily apply to rubber substrata (Harris et al., 1980). I doubt if any such direct pushing can be the correct interpretation, however. For one thing, it would not explain why disruption of microtubules is able to reverse the actin-disrupting effects of phorbol ester tumor promoters and poisons of cyclic

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AMP metabolism. Furthermore, Danowski also tried the effects of the antimicrotubule drug taxol, which acts by promoting microtubule assembly rather than inhibiting it. This drug has inconsistent effects on the exertion of contractile forces, but even though it caused all the microtubules to form a dense radial ball at one end of the cells, nevertheless when the taxol was washed out and replaced by medium containing nocodazole or equivalent drugs, the result was still a large increase in contractile force relative to what the cells had been exerting in the presence of the taxol. The point here is that the microtubules in the taxol-treated cells couldn’t have been doing any effective pushing along the cell’s lengths, since they were all concentrated into a ball at one end. These and other results strongly support Danowski’s conclusion that there must be some actual stimulation of actin fiber assembly produced by the disruption of microtubules. It is a puzzle why this should occur, either in terms of mechanism or functional significance. One class of possibilities is that one or more of the MAPS (microtubule associated proteins), being released into the cytoplasm by the disrupted microtubules, could promote assembly either of actin itself, or of one of the other components of stress fibers. Functionally, it could be part of the special mechanism for controlling locations of actin polymerization, with the dynamic instability of microtubules serving to control actin organization.To the extent that this control were inhibitory in nature; then it would make sense that any generalized depolymerization of cytoplasmic microtubules would, as a side effect, promote an equally generalized polymerization of cytoplasmic actin. One of the consequences would be the loss of directional polarity, while another would be the strengthening of contractility.

VII. QUESTIONS ABOUT THE AUTONOMY OF FIBROBLAST POLARITY Some of the more important questions one can ask about fibroblast polarity concern the continuity of differences in polarity over time, and the mechanisms by which variations in polarization can be passed by a cell to its mitotic progeny. To see dramatic evidence of such inheritance, one need only take fibroblasts of an established line, trypsinize them and plate them out on glass or plastic at low enough population densities to produce discrete, clonal colonies, and then wait long enough for 3 or 4 cell cycles to have been completed. When doing this in the course of other experiments, I have frequently been strongly impressed by “family resemblances” between the 8 to 16 or so cells making up each clone. You see a bunch of long straight ones here, a colony of circular ones there, and above it a clone with nearly all triangular cells. Any observant tissue culturist will have noticed such recurring patterns, although the effect is of course much less striking in more homogeneous cultures, such as primary explants and secondary cultures, since the cell morphologies are so much more uniform within the population as a whole.

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The systematic study of similarities in sibling cells, especially their locomotory behaviors, has somehow become the province of a single researcher, AlbrechtBuehler, whose work I will now attempt to summarize. He began with reports that mitotic sister cells tend to crawl in paths that are mirror images of one another (Albrecht-Buehler, 1977), including some degree of mirror image similarity between the actin cytoskeletons of sister cells; he then proposed that cell directionality is controlled by the 9+0 structure of the centriole, with this control being reflected in a tendency of fibroblasts to turn at angles preferentially close to some multiple of 40 degrees (Albrecht-Buehler, 1979,1981). In case the reason for 40 degrees is not obvious, it is 360 degrees divided by the 9 outer doublets of the typical centriole or basal body structure. Not only does he regard the centriole as “the brain of the cell”, but also proposes that it serves as a visual organ, by means of which fibroblasts are able to extend lamellae directionally toward sites of emission of near infrared light (Albrecht-Buehler, 1991, 1992), especially if this light flashes on and off a hundred or so times per second. This is suggested to be a mechanism of communication between cells, even on opposite sides of sheets of glass, by which they control one another’s directions of elongation. This same author has also proposed that the computational processes of the cellular “brain,” equivalent to synapses and transistors, are accomplished at the molecular level by assembly and disassembly of protein monomers, such as tubulin and actin, into cytoplasmic fibers (Albrecht-Buehler, 1985).The ideais that monomer assembly encodes information in a manner analogous to that of a digital computer. We can regard it as a credit to the openness of science to new and imaginative ideas that all these proposals have been published in the best journals. We also must ask whether the boundary has been crossed from the imaginative to the imaginary.

Figure 78. A pair of mitotic sister 3T3 fibroblasts, just after completing division. Notice how nearly they seem to be mirror images of one another. In fact, until seconds before the picture was taken, there had also been nearly a mirror image pattern of compression wrinkles beneath the cells in the silicone rubber membrane on which they are spreading.

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With regard to whether mitotic daughter cells are sometimes mirror images of each other, or whether they follow mirror image paths, the question is not whether such things sometimes occur, but whether mirror images occur significantly more frequently than would be expected to occur at random. We also need to ask what specific features of the cells’ movements need to be non-randomly correlated in order to produce a given type and frequency of apparent mirror images. Figure 18 shows a pair of mitotic sibling fibroblasts which not only were noticeably near to being mirror images of one another, and (up until just before the photograph was taken!) had even been producing wrinkling patterns in their rubber substrata that were also mirror images! Even the most negative critic should admit such things do occur, sometimes. But how often? There can be an awful lot of fibroblasts in one Petri dish. The proposal of mirror-image paths arose out of studies of the ability of fibroblasts to clear away small particles in their paths by means of the retrograde transport process discussed previously. Albrecht-Buehler (1977) showed that this phenomenon can be used to track cell paths, calling the cleared areas “phagokinetic tracks” (although phagocytosis plays only a minor role, if any; and the accumulation of too many particles on a cell’s surface was already known to alter its behavior). By first coating a substratum with closely spaced particles of colloidal gold, and then observing by dark field microscopy after the fibroblasts have had a chance to crawl about, one finds that the cells’ paths become dramatically visible by their relative darkness, compared with the brightness of the gold that has been left in place. One can easily observe thousands of cell paths, including the branched paths of cells that had undergone one or more mitotic divisions during the time since they were plated onto the gold covered surface, The illustrations in the paper show over a dozen such cases in which the paths of mitotic sibling cells seemed to be approximate mirror images of each other; and when the sibling cells were fluorescently stained for actin stress fibers, considerable degrees of similarity were found in their cytoskeletons as well. Albrecht-Buehler reported that approximately 60% of mitotic sister cells moved in paths that he classified either as approximate mirror images, or as “identical”. At this point, we need to ask ourselves how frequently such paths should occur randomly, i.e. if there were no greater correlation between the directions and speeds of sister cells than there is between unrelated cells. Intuitively, it probably doesn’t seem as if this random frequency could possibly be anywhere near as large as 60%. Figure 19 shows a series of computer-generated pairs of paths meant to be analogous to those of mitotic sistercells. How many of these pairs would you count as being “mirror images”? Many people might count only a, b and c. On the other hand, Albrecht-Buehler also counted pairs in which the turning directions were the same, rather than opposite; these are the pairs he classifies as “identical”; this means counting d and e, as well as a, b, and c. His reasoning was that one of the cells might have gotten turned upside down during mitosis, so that its path was really a mirror image. Thus all but the last (f) above, would be counted. In other words, if neither

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Five possible pairs of pathsof mitotic sister cells. In a, neither undergoes any major turns; in b, c, d, and e, each undergoes a single turn; and in f one turns and the other doesn’t. By Albrecht-Buehler’s criteria, all but f would be classified as mirror images.

Figure 19.

cell turns during the observation period, or if both turn only once (as is quite often the case, to judge from Albrecht-Buehler’s published illustrations, and also based on my own experience), then all possible paths would be counted. The only way to escape that fate is to make a different number of turns, or to turn more than once; and for the case of two turns each, no fewer than half the possible directional combinations would be counted as “mirror images”. The point is that daughter cells need only resemble one another in respect to how frequently they turn in order to produce quite a high percentage of “mirror images.” Perhaps what should surprise us is that Albrecht-Buehler did not find even more than the reported 60%, using such elastic criteria. The eye can often “see” patterns in what is actually randomness, for example in the case of the Rorschach (inkblot) test in psychology. Figure 20 shows a series of computer-generated random path-pairs, about half of which would be counted as approximate “mirror images” by the criteria used by Albrecht-Buehler. These are the first 10 paths generated by a program, written in Turbo-Pascal on a Macintosh computer, with “directions” and “speeds” being assigned by sequences of random numbers. For each of the pairs, two circles were started out at the same location; displacements in the X and Y directions were separately determined for each, based on the compiler’s own random number sequence. For each of the 40 recalculations per cell, its new speed in the X direction equals the old “X speed” plus a random number between -9 and +9; and its new speed in the Y direction likewise equals its old Y speed plus another random number in this same range. There is no connection at all between the calculations for one cell and its sibling. In addition, for each cell at each of the 40 cycles, there is a random one-tenth chance that its speeds in both the X and Y directions will be reset to zero, so as to produce the effect of a random turn. The percentage of apparent “mirror-images” can be increased further by such variations as making the turns occur every so many cycles. My conclusion is that there need not be any real correlations in the directions of cell turns to produce the types and frequencies of “mirror images” reported by Albrecht-Buehler. The eye can find pat-

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figure 20. Ten path-pairs that were generated by a computer program. Although all these paths are actually entirely random, the majority could easily be classified as ”mirror images.“ terns even in randomness; and the author will be glad to provide free copies of this Pascal program to anyone interested in trying it out. The reports of positive phototaxis to certain frequencies of infrared light (Albrecht-Buehler, 1991,1992) also deserve comment. There is certainly no reason to doubt the possibility of some such taxis; there are plenty of other physical stimuli to which fibroblasts are known to orient themselves, including several reviewed above. On the other hand, the reported need for the light’s intensity to oscillate at certain frequencies would certainly be a puzzle, and is also part of what makes independent repetition of this work very much more difficult than it seems at first. The use of a special sapphire lens is also part of this difficulty. As in earlier the “mirror image” work, these reports lack explicit criteria for comparing frequencies of extension of cellular processes toward the light source, as compared with what should occur randomly. Anyone knows, who has observed the particular type of fibroblasts used in these studies, that they are constantly and continually extending and retracting leading lamellae and lamellipodia of all sorts of sizes and durations of existences. This means you have to decide which protrusions are going to “count”. For example, you might decide to count only those protrusions of greater than, say, 5 micrometers in width and length, and of these only those that maintain at least these dimensions for longer than 10 minutes. Equally essential are criteria for the directionality of these protrusions. For example, you might draw an imaginary line from the center of the cell’s nucleus outward to the furthest tip of each cytoplasmic protrusion, or perhaps a line bisecting the two sides of the protrusion. With such criteria in hand, one could then count the frequency with which a fibroblast extends processes to within plus or minus 20 degrees of any given

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direction. Alternatively, you might subdivide the cell periphery into quadrants and count the frequency of protrusions into each one. What you can’t do is just look at the video recordings and say “that extension counts as being toward the light source; that other one doesn’t.” If the cells in question were like Amoebaproteus, with an actual front and arear, or even if they had the degree of directionality of leucocytes, than subjectivejudgments could perhaps be sufficient, especially if the directional response were marked. Nor is the situation aided by doing statistical analyses on one’s previous subjective estimates of which protrusions should “count,” especially when that observer is strongly committed to the hypothesis. As regards the demonstrable tendency of fibroblasts on opposite sides of very thin sheets of glass to orient perpendicular to one another, I would suggest that this is an example of contact guidance and reflects the slight curvature of the glass sheets. On the convex surface, fibroblasts tend to orient in the direction of minimum curvature; while on a slightly concave surface, their direction tends to be across the curvature, with the cells arranged like bow strings relative to the curve of the substratum. One might try flexing such a thin sheet slowly, so that one side is convex one day, with the other side becoming convex the next day. The point is that such phenomena often have fairly mundane explanations, without the need to postulate infrared Morse code being sent from one cell to another.

VIII. CONCLUSIONS AND PROSPECTS To summarize the current situation in this field: much information has been gathered and still is accumulating about the spatial distributions within crawling fibroblasts of the various cytoskeletal and membrane proteins apparently responsible for their locomotion (Beshadsky and Vasiliev, 1988).The force-exertion mechanism of this locomotion seems at last to have been identified, and consists of a continuous assembly of cytoplasmic actin localized along the cell’s leading margins, with actin fibers flowing centripetally from there just beneath the plasma membrane, pulling certain integral membrane proteins along with them, with these “tine proteins” serving to transmit quite strong shearing forces (traction) tangentially through the plasma membrane, from the inside to the outside (reviewed in Heath and Hollifield, 1991;Harris, 1990).On the other hand, this information is just beginning to be applied to questions of cell shape and polarity. Presumably, a fibroblast’s polarity reflects the localization of such processes as actin assembly, the formation of new cell-substratum adhesions, the aggregation of transmembrane adhesion proteins into focal plaques, the disruption of old adhesions, and so on, so that they occur in the correct spatial locations relative to one another. Each molecular and physical event of locomotion must somehow be concentrated at the right places, rather than occurring either randomly or homogeneously. We also have to presume that this localization is self-organizing as well as

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self- perpetuating; otherwise, trypsinized cells would not be able to re-establish polarity and maintain it for periods of a few hours or more. Here are some specific questions. By what means is actin assembly focused so precisely right along the outer edges of what we have been calling the “type A” margins? Perhaps the mechanisms are comparable to those found in leucocytes (Cassimeris et al., 1990). Why and how are the type I myosins concentrated at these edges? Do they somehow promote or localize actin assembly, in the sense that if they were relocated to what had been a type B margin, would actin assembly then begin there? Conversely, which of the special protein constituents of these type A margins would you have to take away in order to convert it to either a type B margin or a type C margin? Another sort of question would involve forces. What does it mean that the herniation of the plasma membrane into blebs is often localized right along the edge of type A margins? It could be that the resistance to outward pressure is either weakest or least stable there; or it could be that the outward forces themselves are concentrated there. Based on subjective impressions of watching too many time lapse films, I would suggest that an important controlling factor may be the state of tension in the cell body. Certainly, an generalized loss of tension will almost always lead to a reorganization of cell polarity. Of course, we already know that tension is created by the outward crawling and traction of the type A margins; but perhaps this tension itself promotes the subdivision of the cell margin into type A versus type C behaviors; in particular, the type C behavior would be promoted where the direction of stretching is approximately parallel to the edge, whereas type A behavior would be promoted where the stretching is directed perpendicular to the edge. If so, then one might predict that rotating the directions of maximum tension, either using flexible substrata or micromanipulator needles, would allow you to control which parts of a cell’s margins behaved in the “A,” the “B,” or the “C” patterns. Another promising direction for research in this field would seem to be that of reinterpreting directional control phenomena, such as contact inhibition and contact guidance, in terms of what we now know about “actin treadmilling” and “membrane raking.” Perhaps we will now also be able to explain how these forcegenerating processes get inactivated near sites of cell-cell contact, why some cells are more susceptible to this inactivation than others, and to what extent reductions in this susceptibility contribute to the greater invasiveness of such things as macrophages and cancer cells. Likewise, in the case of contact guidance or galvanotaxis, we need to ask which molecular processes are promoted or inhibited, and how external cues produce these effects. Also important and timely is the control of actin organization and contraction by microtubules, and specifically by their disassembly. Some sort of controlling function has long been suspected for microtubules in relation to cell polarity and actin organization; but no one expected the discovery by Danowski that disassembling microtubules causes increased assembly of actin, and even reverses disruption of actin by other treatments. Small and Rinnerthaler (1985) have noted that the forrna-

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tion of new adhesion plaques tends to occur near the tips of microtubules, no mechanism for this has been known (see also Geiger et al., 1984; also see Rinnerthaler et al., 1988). Possibly it is related to the formation of the actin stress fibers extending from the adhesion plaque. Certainly, in the case of cytokinesis, it has long been suspected that the tips of the aster microtubules may serve to signal to cortical actin where to form the contractile ring, and thereby the cleavage furrow. In that case, it has been debated whether the aster microtubules should inhibit cortical contraction or promote it; but no one seems to have asked if it is merely the presence of microtubules that cause the stimulation or inhibition, or whether these effects might actually be caused by the disassembly of the microtubule tips. Recent discoveries about the dynamic equilibrium state of cytoplasmic microtubules seem to raise this latter possibility. These are the types of questions we now need to be asking about the control of fibroblast polarity.

REFERENCES Abercrombie, M. (1967). Contact inhibition: the phenomenon and its biological implications. Natl. Cancer Inst. Monogr. 26,249-277. Abercrombie, M. (1970). Contact inhibition in tissue culture. In Vitro 6, 128-142. Abercrombie, M. & Ambrose, E. J. (1958). Interference reflectionmicroscopystudies ofcell contacts in tissue culture. Exp. Cell Res. 15, 332-345. Abercrombie, M. & Dunn, G. A. (1975). Adhesions of fibroblasts to substratum during contact inhibition observed by interference reflection microscopy. Exp. Cell Res. 92, 57-62. Abercrombie, M. & Heaysman, J. E. M. (1953). Observations on the social behaviour of cells in tissue culture. I. Speed of movement of chick heart fibroblasts in relation to their mutual contacts. Exp. Cell Res. 5 , 111-131. Abercrombie, M. & Heaysman, J. E. M. (1954). Observations on the social behaviour of cells in tissue culture. 11. Monolayering of fibroblasts. Exp. Cell Res. 6,293-306. Abercrombie, M., Heaysman, J. E. M. & Karthauser, H. M. (1957). Social behaviour of cells in tissue culture. 111. Mutual influence of sarcoma cells and fibroblasts. Exp. Cell Res. 13, 276-291. Abercrombie, M., Heaysman, J. E. M. & Pegrum, S. M. (1970a). The locomotion of fibroblasts in culture I. Movement of the leading edge. Exp. Cell Res. 59,393-398 Abercrombie, M., Heaysman, J.E.M. & Pegrum, S. M. (1970b). The locomotionoffibroblasts in culture 11. ‘Ruffling’ Exp. Cell Res. 60,437444. Abercrombie, M., Heaysman, J.E.M. & Pegrum, S. M. (1970~).The locomotionof fibroblasts in culture Ill. Movements of particles on the dorsal surface of the leading lamella. Exp. Cell Res. 62, 389-398. Abercrombie, M., Heaysman, J. E. M. & Pegrum, S. M. (1971). The locomotionoffibroblasts in culture IV. Electron microscopy of the leading lamella. Exp. Cell Res. 67,359-367. Abercrombie,M.,Heaysman, J. E. M. &Pegrum,S. M. (1972). Thelocomotionoffibroblastsinculture V. Surface marking with concanavalin A. Exp. Cell Res. 73,536-539 Albrecht-Buehler, G. (1977). Daughter 3T3 cells, are they mirror images ofeach other? J. Cell Biol. 72, 595-603 Albrecht-Buehler, G. (1979). The angular distribution of directional changes in guided 3T3 cells. J. Cell Biol. 80, 53-60.

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Albrecht-Buehler, G. (1981). Does the geometric design of centrioles imply their function? Cell Motility 1,237-245 Albrecht-Buehler,G. (1985). Iscytoplasmintelligenttoo?InCellandMuscleMotility. J. W. Shay(ed.), VOI. 6. pp. 1-21. Albrecht-Buehler, G. (1991). Surface extensions of 3T3 cells towards distant infrared light sources. J. Cell Biol. 114,493-502. Albrecht-Buehler, G. (1992). Rudimentary form of cellular “vision.” Proc. Natl. Acad. Sci. USA. 89, 8288-8292. Bell, P. B. Jr. (1977). Locomotory behavior, contact inhibition, and pattern formation of 3T3 and polyoma virus-transformed cells in culture. J. Cell Biol. 74, 963-982. Bershadsky, A. D. & Vasiliev, J.M. (1988). Cytoskeleton. Plenum Press, New York. Bershadsky, A. D., Ivanova, 0. Y., Lyass, L. A., Pletyushkina, 0. Y., Vasiliev, J. M., & Gelfand, I. M. (1990). Cytoskeletal reorganizations responsible for the phorbol ester-induced formation of cytoplasmic processes: Possible involvement of intermediate filaments. Proc. Nat. Acad. Sci. USA 87, 1884-1888. Blodgett, K. B. (1935). Films built by depositing successive monomolecular layers on a solid surface. J. Am Chem. SOC.57, 1007-1022. Britland, S. Clark, P. Connolly , P. & Moores, G.(1992). Micropatterned substratum adhesiveness, a model for morphogenetic cues controlling cell behavior. Exp. Cell Res. 198, 124-129 Brown, A. F. Dugina, V.; Dunn, G. A. & Vasiliev, J. M. (1989). A quantitative analysis ofalterations in the shape of cultured fibroblasts induced by tumour-promoting phorbol ester. Cell Biol. Int. Rep. 13,357-366. Brunette, D. M. (1986). Spreading and orientation of epithelial cells on grooved substrata. Exp. Cell Res. 167,203-217. Brunette, D. M. (1988). The effect of surface topography on cell migration and adhesion. In, Surface Charaterization of Biomaterials. Ratner, BD (ed.), pp.203-217. Elsevier Science Publishers, Amsterdam. Brunette, D. M., Kenner, G. S. & Gould, T. R. L. (1983). Grooved titanium surfaces orient growth and migration of cells from human gingival explants. J. Dent. Res. 62, 1045-1048. Boyde, A,, Grainger, F. & James, D. W. (1969). Scanning electron microscopic observations of chick embryo fibroblasts in vitro, with particular reference to the movement of cells under others. Z. Zellforsch. mikrosk. Anat. 94,4&55. Buck, R. C. (1980). Reorientation response of cells to repeated stretch and recoil ofthe substratum Exp. Cell Res. 127,47&474. Burridge, K. & Connell, L. (1983). A new protein of adhesion plaques and ruMing membranes. J. Cell Biol. 97,35%367. Burridge, K. & Fath, K. (1989). Focal contacts: transmembrane links between the extracellular matrix and the cytoskeleton. Bioessays 10, 104-108. Campbell, R. D. & Marcum, B. A. (1980). Nematocyte migration in Hydra: evidence for contact guidance in vivo. J. Cell Sci. 41,33-51. Cassimeris, L., McNeill, H. & Zigmond, S. H. (1990). Chemotactant-stimulated polymorphonuclear leucocytes contain two populations of actin filaments that differ in their spatial distribution and relative stabilities. J. Cell Biol. 110, 1067-1075. Chen, W-T. (1981). Mechanism of retraction of the trailing edge during fibroblast movement. J. Cell Biol. 90, 187-200. Clark, P. Connolly, P., Curtis, A. S. G., Dow, J. A. T. & Wilkinson, C. D. W. (1987). Topographical control of cell behavior: I . Simple step cues. Development 99,439448. Clark, P. Connolly, P., Curtis, A. S. C., Dow, J. A. T. & Wilkinson, C. D. W. (1990). Topographical control of cell behavior: 11. Multiple grooved substrata. Development 108,635464. Clark, P.Connolly, P., Curtis, A. S. G., Dow, J. A. T. & Wilkinson, C. D. W. (1991). Cell guidance by ultrafine topography in vitro. J. Cell Sci. 99,73-79.

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Clark, P., Connolly, P. & Moores, G. R. (1992). Cell guidance by micropatterned adhesiveness invitro. J. Cell Sci. 103,287-292. Cooper, M. S. & Keller, R. E. (1984). Perpendicular orientation and directional migration of amphibian neural crest cells in DC electrical fields. Proc. Natl. Acad. Sci. USA 81, 16G164. Curtis, A. S. G. &Clark, P. (1990). The effects oftopographic and mechanical properties of materials on cell behavior. Crit. Rev. Biocompat. 5,343-362. Danowski, B. A. (1989). Fibroblast contractility and actin organization are stimulated by microtubule inhibitors. J. Cell Sci. 93,255-266. Danowski, B. A. & Harris, A. K. (1988). Changes in fibroblast contractility, morphology and adhesion in responses to a phorbol ester tumor promoter. Exp. Cell Res. 177,47-59 DePasquale, J. A. & lzzard, C. S. (1987). Evidence for an actin-containingcytoplasmicprecursor ofthe focal contact and the timing of incorporation of vinculin at the focal contact. J.Cell Biol. 105, 2803-2809. DePasquale, J. A. & Izzard, C. S. (1991). Accumulation of talin in nodes at the edge of the lamellipodium and separate incorporation into adhesion plaques at focal contacts in fibroblasts. J. Cell Biol. 113, 1351-1359. DiPasquale, A. & Bell, P. B. (1974). The upper cell surface: its inability to support active cell movement in culture. J. Cell Biol. 62, 198-214. Dow, J. A. T., Clark, P., Connolly, P., Curtis, A. S. G. & Wilkinson, C. D. W. (1987). Novel methods for the guidance and monitoring of single cells and simple networks in culture. In, Cell Behavior: Shape, Adhesion and Motility. Heaysman, J. E. M., Middletion, C. A,, & Watt, F. M. (ed.) J. Cell Sci. Suppl. 8, 55-79. Dugina, V. B., Svitkina, T. M., Vasiliev, J. M. & Gelfand, I. M. (1987). Special type ofmorphological reorganization induced by phorbol ester: reversible partition of cell into motile and stable domains. Proc. Natl. Acad. Sci. USA 84,41224125. Dunn, G. A (1991). How do cells respond to ultrafine surface contours? Bioessays 13,541-543. Dunn, G. A. &Brown, A. F. (1986). Alignmentoffibroblastson groovedsurfaces described by asimple geometric transformation. J. Cell Sci. 83,313-340. Dunn, G. A. & Heath, J. P. (1976). A new hypothesis ofcontact guidance in tissue cells. Exp. Cell Res. 101, 1-14. Elsdale, T. R. (1968). Parallel orientation of fibroblasts in vitro. Exp. Cell Res. 51,439450. Erickson, C. A. (1978a). Analysis of the formation of parallel arrays by BHK cells in vitro. Exp. Cell Res. 115,303-315. Erickson, C. A. (1978b). Contact behaviour and pattern formation of BHK and polyoma virus-transformed BHK fibroblasts in culture. J. Cell Sci. 33, 53-84. be influenced Erickson, C. A. &Nuccitelli, R. (1984). Embryonicfibroblastmotilityandorientationcan by physiological electric fields. J. Cell Biol. 98,296-307. Euteneuer, U. & Schliwa, M. (1992). Mechanism ofcentrosome positioningduring the wound response in BSC-I cells. J. Cell Biol. 116, 1157-1 166. Ferrier, J., Ross, S. M., Kanehisa, J. & Aubin, J. E. (1986). Osteoclasts and osteoblasts migrate in opposite directions in response to a constant electrical field. J. Cell Physiol. 129,283-288. Fisher, G. W., Conrad, P. A,, DeBiasio, R. L. & Taylor, D. L. (1988). Centripetal transport of cytoplasm, actin, and the cell surface in lamellipodia of fibroblasts. Cell Motil. & Cytoskel. 11, 235-247. Forscher, P. & Smith, S. J. (1988). Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J. Cell Biol. 107, 1505-1516. Garber, B. (1952). Quantitative studies on the dependenceofcell morphology andmotility uponthe fine structure of the medium in tissue culture. Exp. Cell Res. 5, 132-146 Geiger, B., Avnur, Z., Rinnerthaler, G., Hinssen, H. & Small, J. V. (1984). Microfilament organizing centers in areas of cell contact: Cytoskeletal interactions during cell attachment and locomotion. J. Cell Biol. 99, 83-91,

250

ALBERT K. HARRIS

Gelfand, V. I., Glushankova,N. A,, Ivanova,O. Yu., Mittelman, L. A,, Pletyushkina,O. Yu. Vasiliev. J. M. & Gelfand. I . M. (1985). Polarization of cytoplasmic fragments microsurgically detached from mouse fibroblasts. Cell Biol. Int. Rep. 9,883-892. Guelstein, V.I., Ivanova, 0. Y., Margolis, L. B., Vasiliev, J. M. & Gelfand, I.M. (1973). Contact inhibition of movement in the cultures of transformed cells. Proc. Natl. Acad. Sci. USA 70, 201 1-2014. Gundersen, G . G. & Bulinski, J. C. (1988). Selective stabilization of microtubules oriented toward the direction of cell migration. Proc. Natl. Acad. Sci. USA 85, 59465950. Harris, A. K. (1973a). Behavior of cultured cells on substrata ofvariable adhesiveness. Exp. Cell Res. 77,285-297. Harris, A. K. (1973b). Location of cellular adhesions to solid substrata. Dev. Biol. 35, 97-1 14. Harris, A. K. (1974). Contact inhibition of cell locomotion. In Cell communication. R.P. Cox (ed.) John Wiley & Sons, New York, pp. 147-185. Harris, A. K. (1982). Traction and its relations to contraction in tissue cell locomotion. In Cell Behavior: a tribute to Michael Abercrombie. R. Bellairs, A. Curtis & Dunn, G. (eds.) Cambridge U. Press, Cambridge, pp. 109-134. Harris, A. K. (1990). Protrusive activity of the cell surface and the movements of tissue cells. In Biomechanics of Active Movements and Deformation of Cells. Akkas, N. (ed.), NATO AS1 Series H: Cell Biology, vol. 42. pp. 249-294. Springer-Verlag, New York. Harris, A. K. & Dunn G. A. (1972). Centripetal transport of attached particles on both surfaces of moving fibroblasts. Exp. Cell Res. 73, 519-523. Harris, A. K., Wild, P. & Stopak, D. (1980). Silicone rubber substrata: anew wrinkle in the study ofcell locomotion. Science 208, 177-179. Harris, A. K., Pryer, N. K. & Paydarfar, D. (1990). Effects of electric fields on fibroblast contractility and the cytoskeleton. J. Exp. Zool. 253, 163-176. Harrison,R. G. (1914). Thereactionofembryoniccells tosolidstructures. J. Exp. Zool. 17,521-544. Heaysman, J.E.M. (1978). Contact inhibitionoflocomotion:areappraisal. Int. Rev. Cytol. 55,4946. Heath, J. P. & Dunn, G. A. (1978). Cell to substratum contacts of chick fibroblasts and their relation to the microfilament system. A correlated interference-reflexion and high voltage electron-microscope study. J. Cell Sci. 29, 197-212. Heath, J. P. & Holifield, B. F. (1991). Cell locomotion: new research tests old ideas on membrane and cytoskeletal flow. Cell Motility and the Cytoskeleton 18, 245-257. Hinckle, L. C. D., Mc Caig, C. D. & Robinson, K. R. (1981). The direction of growth of differentiating neurons and myoblasts from frog embryos in an appiied electric field. J. Physiol. (Lond.) 314, 121-135. Holifield, B. F., Ishihara, A. & Jacobson, K. (1990). Comparative behavior of membrane protein-antibody complexes on motile fibroblasts: Implications for a mechanism of capping. J. Cell Biol. 111,2499-2512. Ingber, D. E. (1993). Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton. J. Cell Sci. 104, 613-627. Ingvar, S. (1920). Reactions of cells to the galvanic current in tissue cultures Proc. Am. SOC.Exp. & Med. 17, 198 Ishihara, A,, Holifield, B. & Jacobson, K. (1988). Analysis of lateral redistribution of a monoclonal antibody complex plasma membrane glycoprotein which occurs during cell locomotion. J. Cell Biol. 106,329-343. Izzard, C. S. (1988). A precursor ofthe focal contact in cultured fibroblasts. Cell Motil. Cytoskeleton. 10, 137-142. Izzard, S. L. & Izzard, C. S. (1987). Actin-associated proteins related to focal and close cell-substrate contacts in murine fibroblasts. Exp. Cell. Res. 170,214-227. Izzard, C. S. & Lochner, L. R. (1976). Cell to substrate contacts in living fibroblasts: an interference relexion study with an evaluation of the technique. J. Cell Sci. 21, 129-159.

Fibroblast Polarity

251

h a r d , C. S. & Lochner, L. R. (1980). Formationofcell-to-substratecontacts duringfibroblastmotility: an interference-reflexion study. J. Cell Sci. 42,81-116. Kolodney, M. S. & Wysolmerski, R. B. (1992). Isometric contraction by fibroblasts and endothelid cells in tissue culture; a quantitative study. J. Cell Biol. 117, 73-82. Luther, P.W., Peng, H. B. & Lin, J. J-C. (1983). Changes in cell shape and actin distribution induced by constant electric fields. Nature 303, 61-64. Lyass, L. A., Bershadsky, A. D., Vasiliev, J. M. & Gelfand, I. M. (1988). Microtubule-dependenteffect of phorbol ester on the contractility of cytoskeleton of cultured fibroblasts. Proc. Natl. Acad. Sci. USA. 85,9538-9541. Middleton, C. A., Brown, A. F., Brown, R. M., Karavanova, 1. D., Roberts, D. J. & Vasiliev, J. M. (1989). The polarization of fibroblasts in early primary cultures is independent of microtubule integrity. J. Cell Sci. 94,25-32. Nuccitelli, R. & Erickson, C. A. (1983). Embryonic cell motility can be guided by physiological electric fields. Cell Motil. 2,243-255. Oakley, C. & Brunette, D. M. (1993). The sequence of alignment of microtubules, focal contacts and actin filaments in fibroblasts spreadingonsmooth and grooved titanium substrata. J. Cell Sci. 106, 343-354. O’Hara, P. T. & Buck, R. C. (1979). Contact guidance in vitro. A light, transmission and scanning electron microscopic study. Exp. Cell Res. 121,235-249. Orida, N (1980). Directed lamellipodial protrusive activity in macrophages induced by extracellular electric fields J. Cell Biol. 87, 92a. Paddock, S. W. & Dunn, G. A. (1986). Analysingcollisionsbetween fibroblasts and fibrosarcomacells: fibrosarcomacells show an active invasionary response. J. Cell Sci. 81, 163-187. Rich, A. M. & Harris, A. K. (198 1). Anomalous preferences of cultured macrophages for hydrophobic and roughened surfaces. J. Cell Sci. 50, 1-7. Rinnerthaler, G., Geiger, B. & Small, J. V. (1988). Contact formation during fibroblast locomotion: Involvement of membrane ruffles and microtubules. J. Cell Biol. 106, 747-760. Robinson, K. R. (1985). The responses of cells to electrical fields: A review. J. Cell Biol. 101, 2023-2027. Rodionov, V. I., Gyoeva, F. K., Tanaka, E., Bershadsky, A. D., Vasiliev, J. M. & Gelfand, V. I. (1993). Microtubule-dependent control of cell shape and pseudopodial activity is inhibited by the antibody to kinesin motor domain. J. Cell Biol. 123, 181 1-1820. Rosenberg, M. ( I 962). Long-range interactions between cell and substratum. Proc Natl. Acad. Sci. USA 48, 1342-1349. Rosenberg, M. (1963). Cell guidance by alterations in monomolecular films. Science 139, 41 1-412. Astudy Rovensky, Y. A. & Slavnaja, 1. L (1974). Spreadingoffibroblast-likecellsongroovedsurfaces. by scanning electron microscopy. Exp. Cell Res. 84, 199-206. Schutze, K., Maniotis, A. & Schliwa, M. (1991). The position of the microtubule-organizing center in directionally migrating fibroblasts depends on the nature of the substratum. Proc. Natl. Acad. Sci. USA 88,8367-8371. Singer, S. J. & Kupfer, A. (1986). The directed migration of eukaryotic cells. Ann. Rev. Cell Biol. 2, 337-365. Small, J. V. (1981). Organization of actin in the leading edge of cultured cells: Influence of osmium tetroxide and dehydration on the ultrastructure of actin networks. J. Cell Biol. 91,695-705. Small, J. V. & Rinnerthaler, G. (1985). Cytostructural dynamics of contact formation during fibroblast locomotion in vitro. Exp. Bio. Med. 10, 54-68. Soong, H. K., Parkinson, W. C., SuIik, G. L. & Bafna, S. (1990). Effects of electric fields on cytoskeleton of corneal stromal fibroblasts. Cum. Eye Res. 9, 893-901. Stopak, D. & Harris, A. K. (1982). Connective tissue morphogenesis by fibroblast traction I . Tissue culture observations. Dev. Biol. 90, 385-398.

252

ALBERT K. HARRIS

Svitkina,T. M.,Neyfakh, A. A. Jr. & Bershadsky,A. D. (1986). Actincytoskeletonofspreadfibroblasts appears to assemble at the cell edges. J. Cell Sci. 82,235-248. Symons, M. H. & Mitchison, T. J. (1991). Control of actin permeabilization in live and permeabilized fibroblasts. J. Cell Biol. 114,503-513. Trinkaus, J. P. ,Betchaku, T. & Krulikowski, L. S. (1971). Local inhibition of ruffling during contact inhibition of cell movement. Exp. Cell Res. 64,291-300. Vasiliev, J. M. (1991). Polarization of pseudopodial activities: cytoskeletal mechanisms J. Cell Sci. 98, 1-4. Vasiliev, J. M. &Gelfand, I. M. (1976). Effectofcolcemidon morphogeneticprocessesand locomotion of fibroblasts. In Cell Motility, ed. Goldman, R., Pollard T. & Rosenbaum. J. (eds.), Cold Spring Harbor Conferences on Cell Proliferation, Vol. 3, Book A, pp. 279-304, New York, Cold Spring Harbor Laboratory. Vasiliev, J. M. & Gelfand, I. M. (1977). Mechanisms of morphogenesis in cell cultures. lnt. Rev. Cytol. 50, 159-274. Vasiliev, J. M., Gelfand, I. M.,Domnina,L. V., Ivanova,O. Y. Komm, S.G. & Olshevkaja,L. V. (1970). Effect of colcemid on the locomotory behavior of fibroblasts. J. Embryol. Exp. Morphol. 24, 625-640. Vasiliev, J. M., Gelfand, I. M., Domnina, L. V., Zacharova, 0. S. & Ljubimov, A.V. (1975). Contact inhibition of phagocytosis in epithelial cell sheets: Alterations of cell surface properties induced by cell-cell contact. Proc. Natl. Acad. Sci. USA 72, 719-722. Vasiliev, J. M., Gelfand,l.M., Domnina,L.V., Dorfman,N.A. & Pletyushkina,O.Y. (1976). Activecell edge and movements of concanavalin A receptors of the surface of epithelial and fibroblastic cells. Proc. Natl. Acad. Sci. USA 73,4085-4089. Wagner, M. C., Barylko, B. & Albanesi, J. P. (1992). Tissue distribution and subcellular localization of mammalian myosin I. J. Cell Biol. 119, 163-170. Wang, Y-L. (1985). Exchange ofactin subunits at the leading edge ofliving fibroblasts: Possible role of treadmilling. J. Cell Biol. 101,597-602. Weiss, P. A. (1934). In vitro experiments on the factors determining the course of the outgrowing nerve fiber. J. Exp. Zool. 68,393-448. Weiss, P. A. (1961). Guidingprinciples in cell locomotionand cell aggregation. Exp. Cell Res. Suppl. 8, 260-28 1. Weiss, P. A. & Garber, B. (1952). Shape and movement of mesenchyme cells as functions of the physical structure of the medium. Contributions to a quantitative cell morphology. Proc. Nat. Acad. Sci. USA 38,264-280. Weiss, P. A. & Scott, B. I. H. (1963). Polarizationofcell locomotion in vitro. ProcNatl. Acad. Sci. USA 50,33&336. Witkowski, J. A. & W. D. Broughton (1971). Stages of spreading of human diploid cells on glass surfaces. Exp. Cell Res. 68,372-380. Zand, M. S. & Albrecht-Buehler, G. (1989). What structures, besides adhesions prevent spread cells from rounding up? Cell Motility and the Cytoskeleton. 13,54-67. Zand, M. S. & Albrecht-Buehler, G. (1992). Mechanical perturbationofwebbededges in 3T3 cells. Cell Motility and the Cytoskeleton. 21, 195-21 1.

INDEX

al-and a2-subunits, 167-168 AbpIp, 10 Acetylcholine, 159 Acetylcholinereceptors (AChR), selective retention, 146-147 AChR (see Acetylcholine receptors) ACT I/ACT 2 genes, 7,49 Actin, 5, 24 adhesion, 76 Cdc42p localization, 19, 25 fibroblasts, 206,209, 212, 215,241, 245 filamentous, 136 genetically interacting genes, 15-16 Actin cytoskeleton (see also Cytoskeleton) budding components, 6-9,32-33,34, 41 disruption in fibroblasts, 238-239 mating projection components, 4950,52-53 Actin-binding proteins, 10-15, 21 (see also Proteins) dystrophin-gl ycoprotein complex, 175-176 genetically interacting genes, 15-16 localization to axons and dendrites, 136 Adaptin related molecules, sorting, 110

Adhesion blastomeres, 76-77 compaction, 68-70,70-73 desmosomes, 85-86 fibroblasts, 207-208, 215-216, 245 GPI-anchored proteins, 140-141 localization, 137 uvomorulin, 80 Ankyrinlspectrin cytoskeletal complexes, 176-177 selective retention, 145-146 Apical dominance, pseudohyphal budding, 42 Axon differential protein localization, 136138 neuronal polarity, 134-135, 147-149 P-subunit, 168 Bemlp, 16-17,51 Bem2p, 23 Bem3p, 20 BFA (see Fungal metabolite brefeldin A) BHK, fibroblasts, 224-227,236 Blastocyst formation, 68 tissue specificity, 85-86 Blastomere compaction, 68-70,72

253

254

division, 73-75 16-cell, 75-77 32-cell, 78-79 Blebs, fibroblasts, 207,223 Brefeldin A (see Fungal metabolite brefeldin A) Budding cell cycle and growth control, 33-36 component pathways, 24-26 cytoskeletal and secretory components, 4- 18,32-33,44-45 formation and growth, 40-41 haploid and diploid cells, 26-29 influenza virus, 102-103 organelles segregation, 37-40 Rho GTPase cycles, 18-24 signalling mechanisms, 36-37 site selection, 26-33 vesicular stomatitis virus, 102-103 Cadherins, 80,85 Calcium regulation, yeast budding, 36 Calmodulin (Cmdlp) budding, 16-17,25 Myo2p function regulation, 9- 10 Cancer cells, contact inhibition, 223224,226 Capping protein, 10 Caveolae skeletal muscle, 179-181 skeletal myofibrils, 164-165 CaveolinNIP2 1, neuronal polarity, 144 Cavitation, inhibition, 79 Cdc28p-Clnp, bud morphogenesis, 3334 Cdc42p, regulators, 19-21,51-52 Cdc42p GTPase budding, 19 model, 21 Chitin, in yeast cell, 5 Cingulin, synthesis, 84 Clotting, fibroblasts, 214 Cmd 1 p (see Calmodulin)

INDEX

Coated-pits localization, 102 targeting, 107-11 1 Cofilin, 10 Colchicine, 238, 239 Compaction, 68-75 Contact guidance, fibroblasts, 2 12-221, 245 Contact inhibition, fibroblasts, 221-227 Contact paralysis, fibroblasts, 221-227 Contact retraction, 225 COPS, targeting, 120 Costameres cytoskeletal complexes, 172-173 integrin-100A complex, 177-178 sarcolemmal protein markers assembly, 186-192 Cyclic AMP, 239-240 Cycloheximide, growth inhibition, 3637 Cytokinesis, yeast cell budding, 5-6, 29, 30-3 1,48-49 Cytoskeletal complexes costameres, 172-173 spectrin-ankyrin based, 176-177 surface sarcolemma, 170-172 Cytoskeleton (see also Actin cytoskeleton) axonal and dendritic, 135-136 budding components, 6- 18, 32-33 compaction, 68-70, 7 1 mating projection components, 4952 protein sorting, 101 &subunit, 167-168 Dendrite differential protein localization, 136138 mRNA localization, 145 neuronal polarity, 134-135, 147-149 Desmosomes, 79, 85-86

Index

DHPR (see 1,Cdihydropyridine receptor) Drosophila costameres, 188-189, 190 genetics, 121-123, 174 RNA localization, 145-146 Duchenne’s Muscular Dystrophy, 146147 Dystrophin, selective retention, 146147 Dystrophin-glycoprotein complex, skeletal muscle, 174-176, 191- 192 E-C coupling, T-tubules, 165, 179-181 Epithelial cells, 2, 80 differentiation, 75-77 GTPases, 115 membrane polarity, 96-99 and neuronal polarity, 138-139 polarized sorting and target machinery, 114-123 sorting signal, 102-114 Fibroblasts conclusions, 245-247 contact guidance, 2 12-221,245 contact inhibition, 221-227 contact paralysis, 22 1-227 galvanota-xis,227-234 introduction, 202-206 locomotion, 204-205,2 14-215, 2 17218,223,229,241,245-246 margins, 206-2 12 microtubules, 234-240 polarity, 240-245 Fimbrin (Sac6p), 10 FRT cell, cell-type specificity, 112-114 Fruit fly (see Drosophila) FtsZ, 17 Fungal metabolite brefeldin A (BFA) sorting and targeting, 1 19 vesicle bud formation, 1 19-121 Fus3p, 46

255

GABA, 137 Galvanotaxis, fibroblasts, 227-234 h-subunit, 168 GAP-43, 137-138 Glutamate, 137 Glycine, 137 Glycosphingolipids (GSLs), apical sorting, 111-112 Golgi cisternae, protein biosynthesis, 99-101 GPI-anchor, neurons, 140-141 GPI-linkages, sorting, 111- 112, 122 Growth control and morphogenesis, 35-36 Slt2p MAPK pathway, 34-35 Growth inhibition, yeast budding, 3637 GSLs (see Glycosphingolipids) GTP hydrolysis, 21 GTPases budding, 24, 31-32,41 epithelial polarity, 115 Hepatocytes, sorting and targeting, 100 Heterotrimeric G proteins, sorting, 121 Hippocampal system, neuronal polarity, 148 H,K-ATPase, stomach cells, 96-97, 106 Hoglp MAPK pathway, 37 Human placental alkaline phosphatase (PLAP), sorting, 122 ICM (see Inner cell mass) Influenza virus, budding, 102-103 Inner cell mass (ICM) desmosomes, 86 division, 75 formation, 68 stimulation, 79 tight junction, 80-82, 84 Integrin, costamere-specific, 188-189 Integrin-100A filament based complex, 177-178

256

Integrin-actin filament-based complex, 173-174, 187-191 Intracellular effector molecules, axonal and dendritic localization, 137-138 Intracellular targeting, neuronal polarity, 140-147 Invertase, actin secretion, 8 Ion channels, domain specific localization. 137 Kidney cells MDCK cell-type specificity, 112114 sodium absorption, 96-97 Kinesin heavy chain (KHC), neurons, 136 Lamellipodia contact inhibition, 223 fibroblast margins, 206-207 Laminin, 76,79 Langmuir trough, fibroblasts, 2 18 LDLR (see Low density lipoprotein receptor) Leading lamella, 203-204 Lipids epithelial cells, 101 segregation, 134 Locomotion, fibroblasts, 204-205,2 14215,217-218,223,229,241, 245-246 Low density lipoprotein receptor (LDLR), sorting signal, 105107 Lysosomes, targeting, 102 Macrophages contact guidance, 218 contact inhibition, 223, 224 MAPs (see Microtuble-associated proteins)

INDEX

Mating projection formation, 44-45, 52-53 components, 49-52 cytology, 45-46 pheromone signalling, 46-47 MDCK cell apical sorting, 11 1-112 sorting, 100,102-103,104,106,107, 122 targeting, 117, 120 tissue and cell-type specificity, 112114 Mechanocyte (see Fibroblasts) Mg*+-ATPase,muscular fiber, 170 Microtuble organizing centers (MTOCS), 234-235 Microtuble-associated proteins (MAPs), axonal and dendritic cytoskeletons, 135-136 Microtubules compaction, 68-70 fibroblast orientation, 217 fibroblast polarity, 234-240, 246 mating projection formation, 50-5 1, 52-53 yeast budding, 17-18,25-26,39 Mitochondria compaction, 69 polarization, 77 segregation, 39-40 Monclonal antibodies, 166 Mouse early development conclusions, 87 cytological aspects, 68-79 introduction, 68 molecular aspects, 80-86 mRNA costameres, 189-190 localization, I35 MSBI/MSBZ genes, 20-21 MTOCs (see Microtuble organizing centers)

Index

MY02P budding, 16, 18,25,49-50 function regulation, 9-10 Myofibrils skeletal muscle, 157-158 and surface sarcolemma, 187-191 Myosin adhesion, 76 budding components, 9 Myotendenousjunctions, 162 Myotubes, 179

257

1,Cdihydropyndine receptor (DHPR) protein markers, 166-170 TS28, 182-186 Osteoclasts, 227

Pertussis toxin, sorting inhibition, 121 Phdlp, 43 Pheromone, yeast cells, 3,43,46-49 Phorbol esters, 237-239 Phosphorylation compaction, 72-73 SBF, 35 sorting, 107 N-CAM, cell adhesion, 70 Phototaxis, fibroblasts, 244 N-ethlymaleimide Sensitive Factor pIgR (see Polymeric immunoglobulin (NSF), intracellular transport, receptor) 143-144 Pkclp, 21 Na+K+-ATPase PKC (see Protein kinase C) basolateral localization, 78-79 Placental alkaline phosphatase expression, 139 (PLAP), sorting, 105 sorting, 100, 106, 112-114 PLAP (see Human placental alkaline Nerve cells, 2 phosphatase; Placental alkaNeurofilaments, axons and dendrites, line phosphatase) 136 Pollen cells, 2 Neuronal polarity Polymeric immunoglobulin receptor axonal and dendritic cytoskeletons, (pIgR), sorting signals, 105135-136 107 axonal and dendritic differences, Polymorphonuclear leucocytes, 21 1 134-135 Polyoma virus, contact inhibition, 226 differential protein localization, 136- Polyribosomes, exclusion from axon, 138 145 and epithelial cells, 138-139 Potassium, galvanotaxis, 230-232 intrinsic factors, 147-149 Principle of Dynamic Polarization, 134 introduction, 134 Profilin, 10 mechanisms, 139-147 Protein kinase C (PKC) Neurons, domains, 134-135 compaction, 72-73 Nocodazole, 39,235,239 muscular fiber, 169 Nodes of Ranvier, protein localization, Proteinaceous signals, neuronal target134-135 ing, 141-142 NSF (see N-ethlymaleimide Sensitive Proteins (see also specific proteins) Factor) actin-binding, 10-15 cortical, 5 1 Nutrient sensing, yeast budding, 35cytokinesis tag, 30-3 1 36

258

GPI-anchored, 140-141 heterotrimeric G, 121 pseudohyphal growth, 43-44 putative neck filament, 17 segregation, 134 sorting, 103-105 T-tubules, 182-186 targeting, 102 Pseudohyphal growth, 3,41-42 inhibitors and enhancers, 42-44 Purkinje cells, dendritic differentiation, 148 Putative neck filament proteins, 17 Rabs intracellular targeting, 142-143 targeting and sorting, 115-118 Rholp, budding, 21,41,51-52 Rho3p/Rho4p, budding, 23 Rho GTPase cycles, budding, 18-24 RNA, localization, 145 Rsrlp, 31-32,41 Rugophobia, 2 18 Rvs167p, 35-36 Saccharomyces cerevisiae budding during vegetative growth, 441 conclusion, 53-54 mating projection formation, 44-53 overview, 2-4 pseudohyphal growth, 41-44 Sarcoplasmic reticulum (SR), skeletal muscle, 158, 163 SBF, phosphorylation, 35 Schmoo, 45-47 SEC, targeting and sorting, 115-1 18 Secretory apparatus budding components, 6- 18 budding cytology, 4-6 Selective retention localization, ankyridspectrin, 145-147

INDEX

6-dimethyl-aminopurine (6-DMAP), 73 Skeletal muscle cell surface cell surface polarization, 178-192 introduction, 157-159 organization in tissue, 159-160 polarity of domains, 161- 178 Skeletal myofibers, ultrastructure, 161165 SL (see Surface sarcolemma) Slt2p MAPK pathway, budding, 34-35, 52 Smylp, 18,25 SNAP, 118, 119, 143 SNARE proteins, 18, 143-144 epithelial polarity, 118 sorting process, 101 Sodium absorption, 96-97 Sorting (see also Targeting) apical, 111-112 epithelial cells, 98 heterotrimeric G proteins, 121 intracellular protein transport machinery, 114-123 polarized cells, 99-101 Sorting signal epithelial membrane polarity, 102114 paradigm, 101-102 Spa2p, budding, 16-17, 24, 25,34,4950, 5 1, 52-53 SPB (see Spindle pole body) Spectrin, selective retention, 145-146 Spectrin-ankyrin based cytoskeletal complexes, 176-177 Spindle pole body (SPB) mating projection formation, 50-51 yeast budding, 25-26,27,30, 38-40 SR (see Sarcoplasmic reticulum) Staurosporine, 73 Surface sarcolemma (SL) caveoiae, 164-165 costameres, 186-192

Index

protein composition, 165-178 protein markers, 170-178 skeletal muscle, 158-159, 179-181 skeletal myofibers, 161-165 Synaptic interactions, 134 Synaptic vesicle proteins, neuronal polarity, 143-144 T-tubules caveolae, 164-165 protein composition, 165-178 proteins, 182-186 skeletal muscle, 158-159, 179-181 skeletal myofibers, 161-165 Talin, 174 Targeting (see also Sorting) axonal protein, 138-139 basolateral determinants, 107-111 epithelial cells, 98-101 GTPase cycle proteins, 3 1-32 intracellular, 140-147 intracellular protein transport machinery, 114-123 proteins, 102 vesicular stomatitis virus, 138-139 TGN, sorting process, 100 Tight junction, 80-84 Tissue segregation, mouse early development, 68-79, 80-86 Transcytosis, 77, 79, 120 Trophectoderm, formation, 68, 71, 73, 76, 80, 82, 85

259

Tropomyosin, 10 TS28, 168-169 DHPR, 182-186 Tubulin, fibroblasts, 241 Type A margins, fibroblasts, 206-210, 214,227,246 Type B margins, fibroblasts, 2 10-211, 246 Type C margins, fibroblasts, 21 1-212, 226,235,246 Tyrosine, sorting, 109-110 Tyrosinel’light-turn” structure, targeting, 102 Uvomorulin, cell adhesion, 69-70, 7273,80 Vacuole, segregation, 39-40 Vesicular stomatitis virus (VSV) budding, 102-103 sorting, 104-105 targeting, 138-139 Vinblastine, 235, 239 Vinculin, costameres, 173-174, 188189 Viral spike glycoproteins, sorting signal, 103 VSV (see Vesicular stomatitis virus) ZO-1 motein. 81-82,83, 84 -

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Advances in Molecular and Cell Biology Edited by E. Edward Bittar, Departmenf of Physiology, University of Wisconsin, Madison Volume 25, Oxyradicals in Medical Biology 1997,242pp. $109.50/f69.50 ISBN 0-7623-0379-4 Edited by Joe M. McCord, Webb-Waring Lung Institute, University of Colorado, Denver

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CONTENTS: Preface, Joe M. McCord. An Overview of Oxyradicals in Medical Biology, Irwin Fridovich. Regulation of Gene Expression by Oxidative Stress, Klaus Schulze-Osthoff and Patrick A. Baeuerle. Oxyradicals and Malignant Transformation, Larry W Oberly and Terry D. Oberley Oxidative Stress and Human ImmunodeficiencyVirus, Sonia C. Flores and Joe M. McCord. Neutrophils and Ischemic/Reperfusion Injury, Norman R. Harris and D. Neil Granger. Oxyradicals and Acute Lung Injury, Jonathan A. Leff, Brooks M. Hyberston, and John E. Repine. Amyotrophic Lateral Sclerosis (ALS), Z. Rahmani, L. Fox, H. Warner, and David Patterson. Nitric Oxide Regulation of Superoxide and PeroxynitriteDependent Reactions, Homero Rubbo and Bruce Freeman. Iron, Oxygen Radicals,and Disease, Sally K. Nelson and Joe M. McCord. Index. Also Available: Volumes 1-14,16-21,24 Volumes 15, 22-23 (2 part sets)

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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 and E. Edward Bittar, Department of Physiology, University of Wisconsin,Madison Volume 2,The Aging Brain 1997,347pp. ISBN 0-7623-0265-8

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Edited by Mark P. Mattson and James W. Geddes, Sanders-BrownResearch Center on Aging, University of Kentucky; Lexington

CONTENTS: Preface, Mark P. Mattson and James W. Geddes. Toward a Cognitive Neuroscience of Normal Aging, Peter R. Rapp and Michela Gallagher. The Neuronal Cytoskeleton: Changes Associated with Age, Neurodegenerative Disease, and Neuronal Insult, James W. Geddes and Andrew 1. Matus. Structural Changes in the Aged Brain, Dennis W. Dickson. Cerebrovascular Changes in the Aging Brain, J.C. de la Torre. Metabolism of the Aging Brain, John P. Blass, Gary E. Gibson, and Siegfried Hoyer. Contribution of Mitochondrial Alterations to Brain Aging, Gianni Benziand Antonio Moretti. Protein Oxidation Processes in Aging Brain, D. Allan Butterfield and Earl R. Stadtman. Neuroendocrine Aspects of the Aging Brain, Phyllis M. Wise, James P. Herman, and Philip W. Landfield. Changes in Neurotransmitter Signal Transduction Pathways in the Aging Brain, Jeremiah F. Kelly and George S. Roth. Food Restriction and Brain Aging, Caleb E. Finch and Todd E. Morgan. Neurotrophic Factors and the Aging Brain, Mark P. Mattson and Olle Lindvall. Index. Also Available: Volume 1 (1996)

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Advances in Molecular and Cellular Endocrinology Edited by Derek LeRoith, Diabetes Branch, NIDDK, National lnstitufes of Health, Bethesda, Maryland Volume 2,1998,216 pp. ISBN: 0-7623-0292-5

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CONTENTS: Preface, Derek LeRoith. Molecular Aspects of Prolactinand Growth Hormone Receptors, VincentGoffin, f a tima ferrag, and Paul A. Kel/y. Molecular Aspects of Growth Hormone Action, Michael J. Thomas and Peter Rotwein. Leptin, Marc Reitman. Cellular Mechanisms of Signal Transductiuon for Growth Factors, Alan R. Saltiel. Tissue-Specific Expression of the CYP19 (Aromatase) Gene, Evan R. Simpson, M. Dodson Michael, Veena R. Aganval, Margaret M. Hinshelwood, Serdar E. Bulun, and ving Zhao. Molecular Aspects of PrecociousPuberty, Wai-YeeChan and Gordon B. Cutler, Jr. Two G e n e s a n e Disease: The Molecular Basis of Congenital Nephrogenic Diabetes Insipidus, Walter Rosenthal, Alexander Oksche, and Daniel G. Bichet. Mechanisms of Radiation-Induced Carcinogenesis: The Thyroid Model, Yuri E. Nikiforov and James A. Fagin. Index. Also Available: Volume 1 (1997)

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JAI PRESS INC. 100 Prospect Street, P.O. Box 811 Stamford, Connecticut 06904-081I Tel: (203)323-9606 Fax: (203)357-8446