Current Topics in Developmental Biology, Vol. 51 [1st ed.] 0121531511, 9780121531515, 9780080490953

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Current Topics in Developmental Biology

Volume 51

Series Editor Gerald P. Schatten Departments of Obstetrics–Gynecology and Cell and Developmental Biology Oregon Regional Primate Research Center Oregon Health Sciences University Beaverton, Oregon 97006-3499

Editorial Board ¨ Peter Gruss Max Planck Institute of Biophysical Chemistry ¨ Gottingen, Germany

Philip Ingham University of Sheffield, United Kingdom

Mary Lou King University of Miami, Florida

Story C. Landis National Institutes of Health/ National Institute of Neurological Disorders and Stroke Bethesda, Maryland

David R. McClay Duke University, Durham, North Carolina

Yoshitaka Nagahama National Institute for Basic Biology, Okazaki, Japan

Susan Strome Indiana University, Bloomington, Indiana

Virginia Walbot Stanford University, Palo Alto, California

Founding Editors A. A. Moscona Alberto Monroy

Current Topics in Developmental Biology Edited by

Gerald P. Schatten Departments of Obstetrics–Gynecology and Cell and Developmental Biology Oregon Regional Primate Research Center Oregon Health Sciences University Beaverton, Oregon

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∞ This book is printed on acid-free paper.  C 2001 by ACADEMIC PRESS Copyright 

All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2001 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0070-2153/01 $35.00 Explicit permission from Academic Press is not required to reproduce a maximum of two figures or tables from an Academic Press chapter in another scientific or research publication provided that the material has not been credited to another source and that full credit to the Academic Press chapter is given.

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Academic Press Harcourt Place, 32 Jamestown Road, London NW1 7BY, UK http://www.academicpress.com International Standard Book Number: 0-12-153151-1 PRINTED IN THE UNITED STATES OF AMERICA 00 01 02 03 04 05 EB 9 8 7 6

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Contents

Contributors

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1 Patterning and Lineage Specification in the Amphibian Embryo Agnes P. Chan and Laurence D. Etkin I. II. III. IV. V. VI. VII.

Introduction 2 Xenopus as a Model System for Studying Early Embryogenesis Dorsal–Ventral Specification 4 The Spemann Organizer 22 The Three Germ Layers 35 Developmental Pathways and Tumorigenesis 47 Perspectives 48 References 49

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2 Transcriptional Programs Regulating Vascular Smooth Muscle Cell Development and Differentiation Michael S. Parmacek I. II. III. IV. V. VI. VII.

Introduction 69 Embryology of the Vascular Smooth Muscle Cell Lineage(s) 70 Commitment to the Smooth Muscle Cell Lineage 73 Transcriptional Control of Vascular Smooth Muscle Cell Differentiation SRF: A Nuclear Sensor Regulating Growth and Differentiation 76 Modulation of VSMC Phenotype 78 Conclusions and Future Challenges 80 References 82

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3 Myofibroblasts: Molecular Crossdressers Gennyne A. Walker, Ivan A. Guerrero, and Leslie A. Leinwand I. Myofibroblasts: An Overview 91 II. Myofibroblast Origin and the Role of PDGF 92 III. Cytokines and Myofibroblast Phenotypes 94 v

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Contents “Muscle” Structural Protein Expression in Myofibroblasts 95 Mechanisms of “Muscle-Specific” Gene Regulation in Myofibroblasts Myofibroblast Contractility 100 Myofibroblasts and the Cell Cycle 101 Perspectives 104 References 104

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4 Checkpoint and DNA-Repair Proteins Are Associated with the Cores of Mammalian Meiotic Chromosomes Madalena Tarsounas and Peter B. Moens I. Introduction 110 II. Structural Characteristics of Meiotic Chromosomes during the Prophase of Meiosis I 111 III. Meiotic Checkpoint and Recombination Proteins Are Associated with the Cores of the Meiotic Chromosomes 117 IV. Conclusions and Perspectives 127 References 127

5 Cytoskeletal and Ca2+ Regulation of Hyphal Tip Growth and Initiation Sara Torralba and I. Brent Heath I. II. III. IV.

Introduction: Characteristics of Fungal Growth The Cytoskeleton and Apical Growth in Fungi Calcium and Apical Growth in Fungi 154 Conclusions 170 References 170

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6 Pattern Formation during C. elegans Vulval Induction Minqin Wang and Paul W. Sternberg I. II. III. IV. V. VI. VII. VIII.

Introduction 190 Spatial Regulation of VPC Competence 193 Temporal Regulation of VPC Competence and Commitment Downstream Events of RAS Signaling 203 Negative Regulation of RAS Signaling 206 Lateral Signaling 210 Evolutionary Implications 212 Conclusions and Future Directions 213 References 214

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7 A Molecular Clock Involved in Somite Segmentation Miguel Maroto and Olivier Pourqui´e I. II. III. IV. V. VI.

Introduction 222 Models for Somite Formation 224 A Molecular Clock Linked to Somitogenesis 225 Notch Signaling Pathway 229 Other Genes Implicated in Somitogenesis 236 Conservation of the Segmentation Clock in Evolution References 243

Index 249 Contents of Previous Volumes

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Contributors

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

Agnes P. Chan (1), Department of Molecular Genetics, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030 Laurence D. Etkin (1), Department of Molecular Genetics, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030 Ivan A. Guerrero (91), Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309 I. Brent Heath (135), Biology Department, York University, Toronto, Ontario, M3J 1P3 Canada Leslie A. Leinwand (91), Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309 Miguel Maroto (221), Laboratoire de G´en´etique et de Physiologie du D´eveloppement (LGPD), Developmental Biology Institute of Marseille (IBDM), CNRSINSERM-Universit´e de la Mediterran´ee-AP de Marseille, France Peter B. Moens (109), Department of Biology, York University, Toronto, Ontario, M3J 1P3 Canada Michael S. Parmacek (69), Department of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Olivier Pourqui´e (221), Labatoire de G´en´etique et de Physiologie du D´eveloppement (LGPD), Developmental Biology Institute of Marseille (IBDM), CNRSINSERM-Universit´e de la M´editerran´ee-AP de Marseille, France Paul W. Sternberg (189), Howard Hughes Medical Institute and Division of Biology, California Institute of Technology, Pasadena, California 91125 Madalena Tarsounas (109), Department of Biology, York University, Toronto, Ontario, M3J 1P3 Canada Sara Torralba (135), Biology Department, York University, Toronto, Ontario, M3J 1P3 Canada Gennyne A. Walker (91), Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309 Minqin Wang (189), Howard Hughes Medical Institute and Division of Biology, California Institute of Technology, Pasadena, California 91125

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1 Patterning and Lineage Specification in the Amphibian Embryo Agnes P. Chan and Laurence D. Etkin Department of Molecular Genetics The University of Texas M. D. Anderson Cancer Center Houston, Texas 77030

I. Introduction II. Xenopus as a Model System for Studying Early Embryogenesis III. Dorsal–Ventral Specification A. Cytoplasmic Determinants B. Cortical Rotation C. Nieuwkoop Center D. Vegetal Cortical Cytoplasm/␤-Catenin Signaling Pathway E. Molecular Nature of the Dorsal Determinant in Vegetal Cortical Cytoplasm F. Cooperation between TGF-␤ Signaling and Wnt Signaling G. TGF-␤ Receptors, Smads, and Target Genes IV. The Spemann Organizer A. Organizer Genes Expressed in the Dorsal Vegetal Region B. Organizer Genes Expressed in the Prechordal Mesoderm C. Organizer Genes Expressed in the Anterior Endomesoderm D. A Mammalian Structure Analogous to the Anterior Endomesoderm E. How Is the Organizer Formed after All? V. The Three Germ Layers A. Endoderm B. Mesoderm C. Ectoderm D. A Theoretical Model of Germ Layer Formation VI. Developmental Pathways and Tumorigenesis VII. Perspectives References

Xenopus has been widely used to study early embryogenesis because the embryos allow for efficient functional assays of gene products by the overexpression of RNA. The first asymmetry of the embryo is initiated during oogenesis and is manifested by the darkly pigmented animal hemisphere and lightly pigmented vegetal hemisphere. Upon fertilization a second asymmetry, the dorsal–ventral asymmetry, is established, with the sperm entry site defining the prospective ventral region. During the cleavage stage, a vegetal cortical cytoplasm Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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(VCC)/␤-catenin signaling pathway is differentially activated on the prospective dorsal side of the embryo. The overlapping of the VCC/␤-catenin and transforming growth factor beta (TGF-␤) pathways in the dorsal vegetal quadrant specifies dorsal–vental axis formation by regulating formation of the Spemann organizer, including the anterior endomesoderm. The organizer initiates gastrulation to form a triploblastic embryo in which the mesoderm layer is located between the ectoderm layer and the endoderm layer. The interplay between maternal and zygotic TGF-␤s and the T-box transcription factors in the vegetal hemisphere initiates the specification of germ-layer lineages. TGF-␤ signaling originating from the vegetal region induces mesoderm in the equatorial region, and initiates endoderm differentiation directly in the vegetal region. The ectoderm develops from the animal region, which does not come into contact with the vegetal TGF-␤ signals. A large number of the downstream components and transcriptional targets of early developmental pathways have been identified and characterized. This review gives an overview of recent advances in the understanding of the functional roles and interactions of the molecular players important for axis determination and germ-layer specification during early Xenopus embryogenesis.  2001 Academic Press. C

I. Introduction The transformation of a single-celled zygote into a highly organized adult organism requires precise regulation of cell growth and differentiation. The study of embryogenesis has been an intriguing subject for biologists. Experimental embryology has provided fundamental knowledge of early embryonic interactions by careful manipulation of embryos. Almost 80 years after the initial discovery of the amphibian gastrula organizer, our understanding of early development has grown from a cellular level to a molecular level. A number of developmental processes can now be explained in terms of activation or repression of gene expression. These data have provided a molecular basis for understanding the cascade of genetic regulation during early embryonic development. This chapter focuses on early embryonic development of Xenopus laevis. The setting up of the dorsal–ventral axis in embryos has a critical function in determining the future body plan. In Xenopus, dorsal–ventral polarity is established at the time of fertilization by the triggering of the translocation of vegetal cortical cytoplasm (VCC) to the prospective dorsal side of the embryo. Components of the VCC, via the ␤-catenin-dependent pathway, activate target gene expression in the dorsal vegetal region in blastula-stage embryos. The VCC/␤-catenin signaling pathway functions cooperatively with the transforming growth factor beta (TGF-␤) pathway to induce specific gene expression in the Spemann organizer in gastrula-stage embryos. Induction of the three germ layers takes place concomitantly with the establishment of the dorsal–ventral axis. The endodermal and mesodermal layers of the embryo are induced as a result of active TGF-␤ signaling initiated by maternally localized transcripts. Bone morphogenetic protein (BMP)

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and post-MBT Wnt signaling further pattern the dorsal–ventral polarity of the germ layers by specifying a ventral cell fate. Examples from other model systems are included throughout the chapter to try to provide additional data in specific areas of embryonic development. The use of different developmental model systems can complement the deficiencies of individual systems. Integration of information obtained from different systems can allow the elucidation of complex developmental pathways that are evolutionarily conserved. A better understanding of the mechanisms controlling growth and differentiation is a prerequisite to fulfilling the ultimate goal in unraveling the cause and control of malignancy during tumorigenesis. This review gives an overview of recent advances in the understanding of the functional roles and interactions of the molecular players important for patterning and lineage specification during early embryogenesis in Xenopus laevis.

II. Xenopus as a Model System for Studying Early Embryogenesis Xenopus laevis is one of several model systems used for studying early embryogenesis (Harland and Gerhart, 1997). Other developmental systems include Caenorhabditis elegans (Rose and Kemphues, 1998; Labouesse and Mango, 1999), Drosophila (Baek and Lee, 1999), zebrafish (Kodjabachian et al., 1999; Mullins, 1999), the chick (Bachvarova, 1999), and the mouse (Beddington and Robertson, 1999; Gardner, 2000). The advantages of using Xenopus as an experimental system for studying early embryonic development are the ease of maintenance of the animals, the availability of large quantities of embryos year-round, and the rapid development of the embryos in simple salt solution at ambient conditions. The embryos are easy to manipulate for studies involving tissue transplantation, recombination, and explant cultures. The relatively large size of the Xenopus embryo allows efficient isolation of specific regions of embryonic tissues and provides sufficient quantities of starting materials for the construction of cDNA libraries (Blumberg et al., 1991). Several genes have been isolated from the screening of a Xenopus dorsal-lip cDNA library (Cho et al., 1991; Sasai et al., 1994; Bouwmeester et al., 1996). These genes have subsequently been used to isolate homologs from other developmental systems. Furthermore, the capacity of Xenopus embryos for large injection volumes has made possible rapid functional screening using cDNA expression libraries (Smith and Harland, 1991; Smith et al., 1993; Lemaire et al., 1995). Overexpression of RNA in Xenopus embryos is an efficient way to assay for gain-of-function phenotypes. In addition to the wild-type gene products, dominantinterfering constructs of growth factors and receptors, and transcription factors fused with activation or repression domains have been overexpressed in embryos to assay for gene functions (Amaya et al., 1991; Conlon et al., 1996; Ryan et al.,

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1996; Horb and Thomsen, 1997). The use of hormone-inducible transcription factor fusion constructs has further aided in the identification of genes that are the immediate targets of transcription factors (Kolm and Sive, 1995; Tada et al., 1998; Melby et al., 1999; Saka et al., 2000). Knockout studies in Xenopus, performed with the use of antisense oligonucleotides or antisense RNA, are another method of assessing gene functions. Maternal RNA transcripts can be depleted using antisense oligonucleotides (Shuttleworth and Colman, 1988; Kloc et al., 1989; Heasman et al., 1991). Antisense RNA expression has been shown to repress zygotic gene expression and thus permit demonstration of gene functions during normal development (Steinbeisser et al., 1995). The introduction of transgenic techniques to Xenopus (Etkin and Pearman, 1987; Kroll and Gerhart, 1994; Chan and Gurdon, 1996; Kroll and Amaya, 1996; Fu et al., 1998; Amaya and Kroll, 1999; Marsh-Armstrong et al., 1999) has exploited the potential of this model system for use in studying the zygotic effects of transgene expression and in characterizing the regulatory sequences of promoters. A gene trap approach in which transgenic techniques were used to carry out mutagenesis in Xenopus has been successful (Bronchain et al., 1999). Introduction of mutations into the Xenopus genome is likely to lead to the identification of novel genes on the basis of the mutant phenotypes. Although there are several advantages to using Xenopus laevis as a model of embryogenesis, the system also suffers from pitfalls. A relatively long generation time of around 1 year is required for the animal to reach sexual maturity, making germline transmission of genetic alterations impractical. The pseudotetraploid nature of the animal does not favor genetic analysis. However, the introduction of Xenopus tropicalis, which is diploid and has a much shorter generation time of 5 months, is likely to circumvent some of the limitations of the Xenopus system (Amaya et al., 1998). On the other hand, model systems widely used for genetic studies include C. elegans, Drosophila, zebrafish, and the mouse. Both C. elegans and the mouse are reliable systems in which to study the effect of knocking out gene functions. RNA interference has been demonstrated in C. elegans (Fire et al., 1998), and gene targeting has been widely applied to manipulate genomic sequences of the mouse (Koller and Smithies, 1992).

III. Dorsal–Ventral Specification The basic body plan of an embryo is elaborated upon the dorsal–ventral axis established during early embryogenesis. In Xenopus, dorsal–ventral polarity is set up at the time of fertilization. In the fertilized egg, cytoplasmic determinants are translocated to the future dorsal side by a rotation of the egg cortex. The VCC/␤-catenin signaling pathway is activated on the prospective dorsal region as a consequence of the cortical rotation. This pathway functions cooperatively

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with TGF-␤ signaling to activate expression of organizer genes at the mid-blastula transition (MBT), when zygotic gene expression first commences. In the past few years, the knowledge of vegetally localized maternal determinants and molecular components of this signaling pathways has grown dramatically.

A. Cytoplasmic Determinants In some embryos, cell fate is determined by the response of a cell to preexisting cytoplasmic factors. Embryonic cells containing such cytoplasmic factors can undergo autonomous differentiation according to their normal fate in the absence of cell–cell interactions. In Xenopus, the existence of cytoplasmic determinants with dorsalizing activity has been demonstrated in the vegetal cortex of unfertilized eggs (Fujisue et al., 1993; Holowacz and Elinson, 1993). Vegetal deposition of determinants in Xenopus probably results from an asymmetric distribution of maternal components such as RNAs and proteins along the animal–vegetal axis during oogenesis. Wild-type Xenopus embryos exhibit prominent external polarity beginning at mid oogenesis (stage III), when pigment granules become more highly concentrated in the animal cortex than in the vegetal cortex. However, animal–vegetal polarity can be traced back to the stage when the secondary oogonium undergoes mitotic divisions to give rise to nests of 16 oocytes. The secondary oogonium contains a large aggregate of mitochondria on only one side of the nucleus. It has been suggested that this aggregate is the precursor of the mitochondrial cloud, also known as the Balbiani body (Al-Mukhtar and Webb, 1971; Coggins, 1973). Differential localization of maternal RNAs in Xenopus follows one of two pathways, the message transport organizer (METRO or early) pathway and the late pathway (Forristall et al., 1995; Kloc and Etkin, 1995; Kloc et al., 2000). RNAs that follow the METRO pathway first localize to the mitochondrial cloud in stage I oocytes. Between late stage I and early stage II, the localized RNAs translocate together with the mitochondrial cloud to the vegetal region and become localized to the cortex, where they remain throughout oogenesis. RNAs that follow the late pathway are excluded from the mitochondrial cloud and are found throughout the cytoplasm in stage I oocytes. Between late stage II and early stage III, late-pathway RNAs localize to specific domains of the vegetal hemisphere including a crescentshaped region in proximity to the nucleus (Chan et al., 1999), a wedge-shaped structure in the vegetal cytoplasm (Kloc and Etkin, 1995), and at the vegetal cortical region (Melton, 1987). The RNAs eventually localize to the vegetal cortex and occupy a broader region than do METRO-pathway RNAs. A subtle difference during the process of vegetal localization has been observed between two late-pathway RNAs, Vg1 and fatvg. Whereas Vg1 mRNA shows no association with the mitochondrial cloud during oogenesis (Kloc and Etkin, 1995; Chan et al., 1999), fatvg mRNA has been found to localize to the mitochondrial

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cloud transiently in stage II oocytes (Chan et al., 1999). fatvg mRNA also associates with the germ plasm during early embryogenesis, similarly to METRO-pathway RNAs (A. P. Chan and L. D. Etkin, unpublished observation). Although the functional role of fatvg mRNA in the germ plasm is yet to be determined, the association of a late-pathway RNA to the germ plasm suggests that the association of localized RNA with the germ plasm in embryos is not limited to METRO-pathway RNAs. Both Vg1 and VegT mRNAs are localized by the late pathway and have been identified as possible cytoplasmic determinants. Xenopus Bicaudal-C (xBic-C) mRNA has also been found to localize to the vegetal cortex and shares a similar time course of localization as Vg1 and Veg T (Wesseley and De Robertis, 2000) Vg1 belongs to the TGF-␤ superfamily. Processed Vg1 can induce dorsal mesoderm and secondary axis (Dale et al., 1993; Thomsen and Melton, 1993). It has been suggested that the processing of vegetally localized Vg1 mRNA is spatially regulated so that Vg1 is active only in the dorsal vegetal region of the embryo (Thomsen and Melton, 1993). VegT is a T-box transcription factor (Zhang and King, 1996). This factor is also known as Antipodean (Apod) (Stennard et al., 1996), Xombi (Lustig et al., 1996b), and Brat (Horb and Thomsen, 1997). Antisense oligonucleotide knockout experiments have indicated that VegT acts as a maternal determinant in the specification of both the endoderm and mesoderm lineages (Zhang and King, 1996; Kofron, et al., 1999). Several METRO-pathway RNAs have been identified as possible candidates of cytoplasmic determinants. These include the Xwnt-11, Xcat2, DEADSouth (formerly Xcat3), Xpat and Xdazl mRNAs, all of which follow the METRO pathway (Kloc and Etkin, 1995) and have been found to associate with the germ plasm in cleavage stage embryos. Xcat2 is a zinc-finger protein (Mosquera et al., 1993). High-resolution electron microscopic studies have shown that Xcat2 mRNA is associated with the germinal granules of the germ plasm (Kloc et al., 1998, 1999). The Xcat2 protein is related to the Drosophila morphogen Nanos, which is involved in germ cell development, including formation and migration (Kobayashi et al., 1996; Forbes and Lehmann, 1998). The DEADSouth protein is a DEAD-box RNA-dependent helicase (MacArthur et al., 2000) Xpat mRNA is expressed in the primordial germ cells until the cells enter the dorsal mesentery. The Xpat protein does not contain any identifiable functional domains (Hudson and Woodland, 1998) Xdazl is an RNA-binding protein required for spermatogenesis (Houston et al., 1998). Depletion of maternal Xdazl mRNA has resulted in defective germ cell migration within the endoderm during early differentiation (Houston and King, 2000). Xwnt-11 mRNA encodes a maternal Wnt molecule (Ku and Melton, 1993). Although the RNA is localized to the entire vegetal cortex, the protein differentially accumulates on the dorsal side of the embryo because of regulated translation of the localized RNA along the dorsal–ventral axis (Schroeder et al., 1999). The control of Xwnt-11 mRNA translation is mediated by differential polyadenylation. Xwnt-11 or other maternal Wnt molecules may be the cytoplasmic determinants required for the VCC/␤-catenin pathway that specifies the dorsal identity of the embryo, provided that the pathway is dependent on a maternal ligand.

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The presence of a dorsalizing activity in Xenopus embryos has been demonstrated by blastomere transplantation studies (Gimlich and Gerhart, 1984; Takasaki and Konishi, 1989; Kageura, 1990). The dorsal vegetal blastomeres of a 64-cell embryo can rescue axis formation in UV-irradiated embryos, which would otherwise develop with a ventralized phenotype lacking any axis (Gimlich and Gerhart, 1984). When the same blastomeres are transplanted to the ventral side of a recipient embryo, an ectopic secondary axis forms in addition to a normal primary axis. The transplanted dorsal vegetal cells therefore have a dorsalizing or axis-inducing activity. The spatial and temporal origins of the dorsalizing activity have been further determined by cytoplasmic-transfer and deletion experiments. Vegetal cytoplasm taken from activated eggs is capable of inducing an ectopic axis in recipient embryos (Fujisue et al., 1993; Holowacz and Elinson, 1993). Only the vegetal cortical cytoplasm—not the deep vegetal cytoplasm or the cortical cytoplasm from other regions—contributes to the dorsalizing activity. The cortical localization of a dorsalizing activity has also been demonstrated by the transplantation of cortical peels isolated from the vegetal region of activated eggs (Kageura, 1997). The segregation of dorsal determinants along the animal–vegetal axis has been investigated by removing cytoplasm from different regions by egg ligation and deletion experiments (Kikkawa et al., 1996; Sakai, 1996). When the vegetal region of an embryo is deleted just after fertilization, the embryo does not develop any dorsal identity. Vegetally deleted embryos can be rescued by injecting vegetal cytoplasm but not animal cytoplasm; again, this finding indicates the presence of a dorsalizing activity in the vegetal cytoplasm. The dorsalizing activity is present before oocyte maturation in prophase I oocytes (Elinson and Pasceri, 1989; Holowacz and Elinson, 1993, 1995). Exposure of prophase I oocytes to UV irradiation produces ventralized phenotypes even though cortical rotation has taken place. In these embryos, no dorsalizing activity can be detected after cytoplasmic transplantation (Elinson and Pasceri, 1989). This shows that dorsal determinants in the oocytes are destroyed by UV irradiation. UV irradiation of one-cell embryos also results in ventralized phenotypes, but the UV target is different from that in prophase I oocytes and is believed to be the microtubule array required for cortical rotation (Elinson and Pasceri, 1989). Studies in ascidian embryos have demonstrated the existence of prelocalized ooplasmic factors in different regions of the fertilized eggs. The animal, vegetal, and posterior regions of the embryo contain tissue-specific determinants for the development of epidermis, endoderm, and muscle, respectively (Nishida, 1997). Blastomeres isolated from the posterior region of ascidian embryos develop autonomously to form muscle (Deno et al., 1984; Nishida, 1992). A search for localized maternal RNAs in ascidian embryos led to the identification of such RNAs specifically localized to the myoplasm (Swalla and Jeffery, 1995) and ectoplasm (Swalla and Jeffery, 1996). A maternal transcript, pem-3, has also been shown to localize to the posterior-vegetal cytoplasm of the egg after fertilization (Satou, 1999). The protein product of pem-3 contains putative RNA-binding

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domains known as the KH-domain. These findings provide a molecular basis for studying cytoplasmic determinants in ascidians. The possible existence of cytoplasmic determinants in mouse embryos is still being vigorously investigated (Gardner, 1998, 1999). It has been shown that embryos subject to centrifugation or mechanical mixing of cytoplasm develop normally (Mulnard and Puissant, 1984; Evsikov et al., 1994). This finding argues against specific localization of components in the cytoplasm. Deletions of different regions of a one-cell mouse embryo have no effect on normal development (ZernickaGoetz, 1998). Again, this finding provides no evidence for there being early regional asymmetry in the mouse egg cytoplasm. In addition, extensive cell mixing that occurs in the mouse epiblast prior to gastrulation is inconsistent with an early segregation of cell lineages by the inheritance of cytoplasmic determinants (Beddington and Robertson, 1989). However, STAT3 and leptin have been shown to localize to the cortex of mouse and human oocytes, and potentially function to specify asymmetry during early cleavage (Antczak and Van Blerkom, 1997). The alignment of the animal–vegetal axis of the mouse zygote with the axis of bilateral symmetry in blastocysts and the proximal–distal axis in egg cylinders suggests that some degree of regional specification or polarity might already exist in the egg cytoplasm (Gardner, 1997; Weber et al., 1999).

B. Cortical Rotation A critical step in determining the prospective dorsal region of the embryo is triggered by cortical rotation. The mechanism and consequences of such movement has attracted considerable attention. An unfertilized Xenopus egg is radially symmetrical along the animal–vegetal axis. Upon fertilization, the dorsal–ventral axis is defined by the site of sperm entry in the animal region. The sperm entry site marks the future ventral side of the embryo and overlaps with the first cleavage plane, which divides the egg bilaterally into right and left halves. After fertilization, cortical rotation takes place one-third of the way through the first cell cycle (100 min), when the outer cortical layer of the fertilized egg rotates 30◦ with respect to a stationary core (Vincent et al., 1986; Vincent and Gerhart, 1987). However, this degree of rotation seems to be inconsistent with results from cytoplasmic transfer studies, which have demonstrated the presence of a dorsalizing activity around the equatorial region 90◦ away from the vegetal cortex (Yuge et al., 1990; Fujisue et al., 1993). In fact, axis-inducing activity has also been detected above the equatorial region in the animal dorsal sector (Gallagher et al., 1991; Hainski and Moody, 1992; Kageura, 1997). A possible explanation for the apparent discrepancy between the degree of cortical rotation and the localization of dorsal activity comes from studies of microtubule-dependent movement in the vegetal cortical region (Elinson and Rowning, 1988; Rowning et al., 1997). A set of parallel microtubules is found in a transport zone 4–8 ␮m below the cortex associated with the inner cytoplasmic

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core (Larabell et al., 1996). These multiple layers of microtubules align with the direction of rotation (Elinson and Rowning, 1988). Since the rotation movement precedes the formation of the microtubule arrays, it has been suggested that the microtubules are not responsible for the rotation of the cortex (Larabell et al., 1996). However, small organelles can be propelled along the parallel array of microtubules that function independently of the cortex. Small organelles may therefore move along the microtubules by motor molecules toward the plus-end to the dorsal side of the embryo. Evidence suggests that endogeneous organelles can be translocated 60–90◦ away from the vegetal pole (Rowning et al., 1997). Transport by microtubules thus accounts for the apparent differences in the localization of dorsalizing activity as predicted from the degree of rotation of the egg cortex. In keeping with the microtubule transport model, a downstream component of the Wnt pathway, Dishevelled (Dsh), has been shown to associate with small vesicle-like organelles that are translocated to the prospective dorsal side by microtubules during cortical rotation (Miller et al., 1999). Microtubule transport is not only specific for dorsal–ventral specification in Xenopus embryos. A dynamic distribution of microtubules has also been observed in the yolk cells of zebrafish embryos (Jesuthasan and Stahle, 1996). In zebrafish embryos, a set of parallel microtubules at the vegetal pole region is required for setting up initial asymmetry at the one-cell stage. At the eight-cell stage, microtubule tracks originating from the dorsal equatorial blastomeres extend toward the vegetal pole. These microtubule tracks may function to mediate directional transport of organelles or determinants required for dorsal development.

C. Nieuwkoop Center After cortical rotation takes place, it is thought that a signaling center—the Nieuwkoop center—is activated in the dorsal vegetal region that subsequently induces the formation of the organizer in the overlying cells in a non-cell-autonomous manner. In a series of tissue recombination experiments, different regions of the yolky vegetal mass of Urodele embryos were tested for their inductive capacity on animal caps (Boterenbrood and Nieuwkoop, 1973). The dorsal vegetal region induced dorsal axial structures, whereas the lateral and ventral vegetal regions induced only ventral structures. This tissue recombination assay has been referred to as the Nieuwkoop recombinant assay. The dorsal vegetal region carrying a dorsal endomesoderm-inducing property is commonly referred to as the Nieuwkoop center (Gerhart et al., 1989). This region is a signaling center required for specifying the dorsal–ventral axis. The inductive effect of the dorsal vegetal cells is active between the early cleavage stage and the late blastula stage (Boterenbrood and Nieuwkoop, 1973). Cell progenies from the Nieuwkoop center do not contribute to the dorsal lip or axial structures formed during gastrulation. The progenies are located vegetal to the dorsal lip, in the endoderm, and are fated to become part of the anterior gut endoderm, as shown by lineage labeling (Bauer et al., 1994;

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Vodicka and Gerhart, 1995). The proposed function of the Nieuwkoop center is to induce the cells immediately above in the equatorial region to form the organizer. However, the requirement of the dorsal vegetal cells for dorsal development has not been directly demonstrated by studies in which all tier C and tier D dorsal blastomeres are removed. In fact, all blastomeres in tier D can be removed without affecting axis formation (Gimlich, 1986). The finding that small organelles can translocate 60–90◦ away from the vegetal pole suggests that the cytoplasmic dorsal determinant that orginates in the vegetal cortical cytoplasm may translocate all the way to the prospective organizer region in the equatorial region (Rowning et al., 1997). Transplantations of cytoplasm, cortical peels, and blastomeres have demonstrated that dorsalizing activity is distributed broadly on the dorsal sector of the embryo, with the highest activities around the vegetal and equatorial regions before and after cortical rotation (Kageura, 1990; Yuge et al., 1990; Kageura, 1997). Thus, the Nieuwkoop center may overlap physically with the region where the prospective organizer forms. Studies of axis induction by transplantation of blastomeres from 32-cell embryos showed that both tier C and tier D dorsal blastomeres are active in the induction assay (Gimlich and Gerhart, 1984; Gimlich, 1986; Kageura, 1990). The dorsalizing activity of tier C dorsal blastomeres and their participation in organizer formation during normal development also support the idea that the region that produces the early dorsal inductive signal overlaps with the organizer. In summary, the Nieuwkoop center is an important concept in defining early inductive signaling during dorsal–ventral specification. The Nieuwkoop center has been regarded as a physical entity spatially and temporally distinct from the gastrula organizer. However, it is also possible that the cytoplasmic dorsal determinants, after translocation to the dorsal side by cortical rotation and interaction with components in the dorsal region, activate the Nieuwkoop center at the dorsal equatorial region during cleavage stage (Kodjabachian and Lemaire, 1998; Moon and Kimelman, 1998). The Nieuwkoop center in turn directly induces formation of the organizer at the equatorial region during the blastula stage (Kodjabachian and Lemaire, 1998; Moon and Kimelman, 1998). In this model, the Nieuwkoop center and the organizer essentially occupy the same region in the embryo but remain temporally distinct. Molecular analysis of the Nieuwkoop center will help to clearly define its role in inducing the formation of the gastrula organizer.

D. Vegetal Cortical Cytoplasm/␤-Catenin Signaling Pathway Since vegetal cortical cytoplasm contains a dorsal determinant with the ability to induce a complete secondary axis, zygotic gene products that can produce a similar effect when they are expressed ectopically are likely to act along the same pathway as the dorsal determinant. Siamois (Lemaire et al., 1995), ␤-catenin (Funayama et al., 1995; Guger and Gumbiner, 1995), Xwnt-8 (Smith and Harland, 1991), and

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some mouse Wnts (McMahon and Moon, 1989; Sokol et al., 1991) are able to generate a complete secondary axis containing most anterior structures. Siamois can be activated directly by Xwnt-8 and also by vegetal cortical cytoplasm (Carnac et al., 1996; Darras et al., 1997). Siamois and ␤-catenin are components of the VCC/␤-catenin signaling pathway involved in defining the dorsal identity of an embryo before the MBT (Brannon and Kimelman, 1996; Fagotto et al., 1997). On the other hand, Xwnt-8 is not maternally expressed and cannot be the maternal dorsal determinant. Endogenous Xwnt-8 is expressed in the ventral–lateral marginal zone after the MBT and is involved in a Wnt pathway required for patterning of the mesoderm (Christian and Moon, 1993; Hoppler et al., 1996). The axis-inducing effect of overexpressed Xwnt-8 is probably due to activation of components of the VCC/␤-catenin signaling pathway. Since the possible involvement of a Wnt ligand to initiate VCC/␤-catenin signaling has yet to be determined, “VCC/␤-catenin” signaling will be used to describe the endogenous signaling event occurring in the dorsal region in cleavage stage Xenopus embryos in the following discussion. The basic components of the Wnt pathway are largely conserved between different developmental processes found in C. elegans, Drosophila, Xenopus (Cadigan and Nusse, 1997) and sea urchins (Wikramanayake et al., 1998; Logan et al., 1999; Huang et al., 2000). The wingless pathway involved in epidermal cell differentiation in Drosophila has been characterized (Klingensmith and Nusse, 1994; Siegfried and Perrimon, 1994) and provides a basis for study of the Wnt pathway in other systems. In the absence of Wnt signaling, ␤-catenin is continuously phosphorylated and degraded by the glycogen synthase kinase-3 (GSK-3) complex via a ubiquitination-dependent pathway (Fig. 1). The resulting low level of ␤-catenin cannot induce target gene expression. In contrast, when a secreted Wnt molecule is recognized by a transmembrane receptor of the frizzled family, a cytoplasmic component, Dsh, is activated. Dsh in turn suppresses the inhibitory effect of GSK3 on ␤-catenin. This results in an accumulation of ␤-catenin in the cytoplasm. ␤-Catenin translocates into the nuclei and interacts with the high mobility group (HMG)-box transcription factor family Tcf/Lef1 to activate target gene expression (Cadigan and Nusse, 1997). In Xenopus, results from promoter studies have shown that direct transcriptional targets inducible by VCC/␤-catenin signaling include the genes encoding the paired-like homeobox transcription factors Siamois (Brannon et al., 1997; Fan et al., 1998) and Twin (Laurent et al., 1997) and the TGF-␤ factor Xnr-3 (McKendry et al., 1997). Expression of the multiple-growth-factor antagonist Cerberus (Cer) has also been shown to be inducible by ␤-catenin in the absence of protein synthesis (Nelson and Gumbiner, 1999). 1. ␤-Catenin The first functional role identified for ␤-catenin was its interaction with E-cadherin, a cell adhesion molecule required for homotypic cell interaction (Nagafuchi and Takeichi, 1989; Ozawa et al., 1989). The requirement of ␤-catenin in embryonic

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Figure 1 Components of the Wnt signaling pathway. In the absence of Wnt signaling, ␤-catenin is continuously phosphorylated and degraded by the GSK-3 complex through a ubiquitination pathway. APC, Axin, ␤-TrCP, and PP2A are components of the GSK-3 complex. In the presence of a Wnt ligand, the Wnt receptor transduces the signal through Dsh and CKI, resulting in inactivation of GSK-3. GBP can also inhibit GSK-3 activity but is not a linear component of the pathway. The suppression of GSK3 activity results in the accumulation of ␤-catenin, which activates the transcription of target genes, including siamois, twin, Xnr-3, and cerberus.

patterning has later been demonstrated both in Drosophila and Xenopus embryos. The Drosophila homolog of ␤-catenin, armadillo, is involved in epidermal differentiation (Klingensmith and Nusse, 1994; Siegfried and Perrimon, 1994). In Xenopus embryos injected with antibodies against ␤-catenin formed secondary axes (McCrea et al., 1993). Embryos depleted of maternal ␤-catenin mRNA showed defects for the establishment of the dorsal–ventral axis (Heasman et al., 1994). ␤-Catenin therefore plays roles in multiple processes, including cell adhesion and embryonic patterning. In Xenopus, ␤-catenin mRNA and ␤-catenin protein are present maternally (DeMarais and Moon, 1992). In cleavage-stage embryos, the ␤-catenin protein is preferentially enriched in the cytoplasm on the prospective dorsal side (Larabell et al., 1997). Nuclear accumulation of ␤-catenin specifically in dorsal blastomeres begins at the 16-cell stage and lasts until the mid-blastula stage. ␤-Catenin contains a putative GSK-3-dependent phosphorylation site, armadillo repeats, and a

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transactivation domain, but no nuclear localization signal. ␤-Catenin mutants with phosphorylation sites deleted at the N terminus have increased stability and are prevented from degradation (Yost et al., 1996). The N terminus is also required for ubiquitination-dependent degradation of ␤-catenin in the proteasome (Aberle et al., 1997; Orford et al., 1997). Since ␤-catenin contains no nuclear localization signal, it may enter the nucleus by association with other nuclear proteins or by an importin-␤-dependent nuclear transport mechanism (Fagotto et al., 1998). The armadillo repeats have been suggested to interact with upstream and downstream components, such as adenomatous polyposis coli (APC) and the Tcf/Lef1 proteins (Huber et al., 1997). ␤-Catenin interacts with the Tcf/Lef1 proteins and provides a transcription activation domain to activate expression of target genes (Molenaar et al., 1996). In addition to the Tcf/Lef1 proteins, other HMG-box transcription factors, such as Xsox17␣ and Xsox3, also associate with ␤-catenin to repress transcription of certain ␤-catenin target genes (Zorn et al., 1999a). This indicates that potential interaction of ␤-catenin with other members of the HMG-box family may also be involved in the regulation of transcription. 2. The Tcf/Lef1 Proteins The mouse proteins Tcf1 and Lef1 are closely related lymphoid transcription factors and were the first proteins discovered in the Tcf/Lef1 family (referred to hereafter as the Tcf proteins) (Clevers and van de Wetering, 1997; Eastman and Grosschedl, 1999). Tcf-1, -3, and -4 and Lef1 have been identified in both mouse and human. hTcf-4 has been implicated in colon cancer (Korinek et al., 1997). XTcf3 is the only frog homolog identified so far (Molenaar et al., 1996). Tcf proteins contain binding domains for ␤-catenin, CREB-binding protein (CBP), and Groucho. Tcf proteins also contain a HMG domain that recognizes a DNA consensus sequence within the regulatory sequence of target genes. Groucho and CBP are corepressors that associate with Tcf proteins to repress transcription (van de Wetering et al., 1997; Cavallo et al., 1998; Roose et al., 1998). Groucho has been suggested to function with a histone deacetylase complex to regulate histone acetylation on chromatin (Choi et al., 1999). In Drosophila, dCBP lowers the affinity of armadillo to dTcf by the acetylation of a conserved lysine residue in the armadillo-binding domain (Waltzer and Bienz, 1998). Tcf-3 and Tcf-4 contain two conserved sites at the C terminus for the binding of C-terminal binding protein, which is also involved in transcriptional repression. Studies with antimorphic protein have shown that the Xenopus homolog of C-terminal binding protein is involved in the regulation of dorsoanterior development (Brannon et al., 1999). The Tcf proteins have been proposed to be architectual factors controlling chromatin structure and transcription (van Houte et al., 1993; Love et al., 1995). The Tcf proteins have dual functions in that they normally act as a transcriptional repressor, and act as an activator in the presence of ␤-catenin. The Tcf binding sites in the siamois promoter are required for the activation of siamois expression in the

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dorsal region (Brannon et al., 1997; Fan et al., 1998). They are also necessary for the repression of siamois expression on the ventral side of the embryo, since removal of Tcf binding sites from the siamois regulatory sequence results in ectopic expression in the ventral side of the embryo (Brannon et al., 1997). 3. Functional Roles of ␤-Catenin and the Tcf Proteins ␤-Catenin and the Tcf proteins are involved in dorsal–ventral axis specification. Overexpression of ␤-catenin on the ventral side of an embryo can induce a complete axis (Funayama et al., 1995). Depletion of the maternal ␤-catenin mRNA by antisense oligonucleotide knockout prevents formation of dorsal structures (Heasman et al., 1994). ␤-catenin knockout mice display defects in mesoderm formation (Haegel et al., 1995). The nuclear function of ␤-catenin is thought to involve the interaction of ␤-catenin with Tcf proteins. Overexpression of a Tcf-3 mutant lacking the ␤-catenin binding site blocks endogenous axis formation as well as the secondary axis induced by ectopic expression of ␤-catenin (Molenaar et al., 1996). Mice deficient in both Tcf1 and Lef1 show defects in the formation of axial structures (Galceran et al., 1999). The concept that ␤-catenin functions in the nucleus to regulate transcription has been challenged by a study using a membrane-tethered form of plakaglobin, a protein related to ␤-catenin. Overexpression of a membrane-tethered form or a wild-type cytoplasmic form of plakaglobin can result in axis duplication (Merriam et al., 1997). On the basis of this result, it has been suggested that ␤-catenin, like plakaglobin, functions in the cytoplasm to keep an inhibitor of dorsal gene expression out of the nucleus. However, such a mechanism is not consistent with the proposed nuclear function of ␤-catenin as a transcriptional activator. The use of a membrane-tethered form of ␤-catenin suggests that the membrane-tethered forms of ␤-catenin or plakaglobin in fact interfere with the ␤-catenin signaling pathway by binding to APC, an endogenous component required for ␤-catenin degradation (Miller and Moon, 1997). The axis-inducing activity of the membrane-tethered forms is therefore likely to result from removal of the inhibitory effect of APC and accumulation of endogenous ␤-catenin in the nucleus. Another result that appears inconsistent with the nuclear function of ␤-catenin for axis duplication is that a secondary axis forms in embryos injected ventrally with antibodies directed against ␤-catenin (McCrea et al., 1993). However, it is possible that, instead of blocking the function of ␤-catenin, the injected anti-␤-catenin antibodies stabilize ␤-catenin and thus result in an increased level of ␤-catenin in injected blastomeres. 4. Glycogen Synthase Kinase-3 The component upstream of ␤-catenin in the Wnt signaling pathway is GSK-3, a serin/threonine kinase, which actively promotes ␤-catenin degradation (Yost et al., 1996). Activation of the Wnt pathway suppresses the activity of Xgsk-3 and results

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in the accumulation of high levels of ␤-catenin. This is demonstrated by the expression of a dominant-negative form of Xgsk-3 on the ventral side of Xenopus embryos, which blocks its endogenous function and subsequently stabilizes ␤-catenin, and induces secondary axis formation (Dominguez et al., 1995; He et al., 1995; Pierce and Kimelman, 1995). Overexpression of wild-type Xgsk-3 on the dorsal side of the embryo suppresses Wnt signaling and inhibits endogenous axis formation (He et al., 1995; Pierce and Kimelman, 1995). Two distinct modes of Xgsk-3 regulation have been reported (Dominguez and Green, 2000). It is shown that an endogenous mechanism exists on the dorsal side of Xenopus embryos to deplete Xgsk-3 protein. A second mode of regulation occurs as a consequence of Wnt or Dsh expression, which causes a reduction in Xgsk-3 activity rather than depletion. 5. Adenomatous Polyposis Coli, Axin, Beta-Transducin Repeat-Containing Protein, and Protein Phosphatase 2A It has been proposed that GSK-3 functions in a large complex containing additional regulators of the Wnt signaling pathway, including APC, Axin, beta-transducin repeat-containing protein (␤-TrCP), and protein phosphatase 2A (PP2A), all of which promote downregulation of ␤-catenin by GSK-3 (Wodarz and Nusse, 1998; Sokol, 1999). APC was originally identified as a tumor suppressor gene because loss of APC function was observed in colon cancer (Polakis, 1997). The APC protein contains armadillo repeats, ␤-catenin binding sites, and protein domains that interact with Axin and PP2A (Bienz, 1999). The C terminus of the APC protein also contains a discs large (DLG)-binding domain that interacts with the PDZ protein domain found in the PSD-95, Discs large, and ZO-1 proteins (Fanning and Anderson, 1999). Both GSK-3 and APC are required to phosphorylate ␤-catenin. GSK-3 may directly phosphorylate ␤-catenin in the presence of APC; alternatively, GSK-3 may first phosphorylate APC, which binds to ␤-catenin and leads to ␤-catenin phosphorylation (Rubinfeld et al., 1996; Yost et al., 1996). However, the role of APC may seem to be more complex because overexpression of APC mutants that degrade ␤catenin in cultured cells induce secondary axis formation when expressed in Xenopus embryos (Vleminckx et al., 1997). This result is in contrast to the expected function of APC, which is to lower the ␤-catenin level. It has been suggested that APC and ␤-catenin may be involved in an independent pathway to induce axis formation or that the mutant forms of APC stabilize ␤-catenin by an unknown mechanism. A mouse mutation, fused, produces a truncated gene product of axin (Zeng et al., 1997). Mutant embryos exhibit duplicate axis formation implicating Axin as a negative regulator of axis formation. Axin contains a DIX (Dishevelled/Axin) domain at the C terminus and RGS (regulator of G protein signaling) domain at the N terminus. The DIX domain, also found in Dsh, has been proposed to allow interaction between Dsh and Axin or oligomerization of Axin (Hsu et al., 1999; Sakanaka and Williams, 1999). However, the DIX domain is not required for the

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ventralization of endogenous axis by axin. The RGS domain has been found in regulators of G protein signaling (Koelle, 1997). Removal of the RGS domain results in a dominant negative form of Axin (Hsu et al., 1999). Similar to the case with the GSK-3 dominant negative mutant, overexpression of the RGS-deleted form of Axin leads to axis duplication. The presence of an RGS domain suggests a possible involvement of G proteins in the regulation of the Wnt pathway. Axin may function in the GSK-3 complex to stabilize the interaction between GSK-3 and ␤-catenin. ␤-TrCP is a vertebrate homolog of Slimp, a F-box/WD40 protein that functions in the ubiquitination of ␤-catenin (Jiang and Struhl, 1998). A mutant form of ␤-TrCP induces ectopic axis formation (Marikawa and Elinson, 1998; Lagna and Hemmati-Brivanlou, 1999; Liu et al., 1999a). In Xenopus, three different transcripts of ␤-TrCP have been isolated, two of which are localized to the vegetal cortex during oogenesis (Hudson et al., 1996). The regulatory and catalytic subunits of PP2A have been found to interact with APC and Axin, respectively. The B56 subunit interacts with APC, as shown in a yeast two-hybrid assay (Seeling et al., 1999). The catalytic subunit of PP2A can bind to Axin (Hsu et al., 1999). It is possible that PP2A mediates specific dephosphorylation of components of the APC complex to inhibit GSK-3-dependent phosphorylation of ␤-catenin. 6. Glycogen Synthase Kinase-3 Binding Protein GSK-3 binding protein (GBP), an upstream regulator of GSK-3 activity, was isolated by a yeast two-hybrid assay (Yost et al., 1998). GBP is related to the product of a T-cell proto-oncogene, Frat1 (Jonkers et al., 1997). GBP inhibits the phosphorylation function of GSK-3 by direct interaction with GSK-3. Overexpression of GBP stabilizes ␤-catenin and can induce a secondary axis (Yost et al., 1998). Antisense oligonucleotide knockout experiments have demonstrated that GBP is required for the establishment of the dorsal–ventral axis in Xenopus embryos (Yost et al., 1998). Evidence indicates that GBP can also induce depletion Xgsk-3 proteins, similar to the endogenous activity existed on the dorsal side of the embryo (Dominguez and Green, 2000). It appears that GBP is a regulator of Wnt signaling but is not a linear component of the pathway. 7. Casein Kinase I and II Overexpression of casein kinase I (CKI) induces an ectopic axis on the ventral side of embryos (Peters et al., 1999). CKI acts downstream of Dsh and upstream of GSK-3. CKI has been shown to phosphorylate Dsh. It has been suggested that such phosphorylation is not required for the functioning of CKI. CKII is also able to phosphorylate Dsh (Willert et al., 1997). Further studies on the regulation of Dsh by these two kinases will be required to determine their roles in transducing the Wnt signal.

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8. Dishevelled Dsh is upstream of GSK-3 and can inhibit the negative regulatory function of GSK3 in the presence of Wnt signaling. A double inhibition downstream of Dsh results in an increase in the ␤-catenin level when Dsh is activated. Dsh is phosphorylated by active Wnt signaling (Yanagawa et al., 1995). The Dsh protein contains a DIX domain, a PDZ domain, and a DEP (Dishevelled, egl-10, and pleckstrin) domain. Removal of the central PDZ domain produces a dominant negative form of Dsh (Sokol, 1996), indicating that the PDZ domain is required for Dsh functioning. Dsh has been shown to translocate from the vegetal pole toward the prospective dorsal side (Miller et al., 1999). This suggests a possible endogenous role of Dsh during VCC/␤-catenin signaling. However, depletion of the maternal gene products will be required to provide definitive evidence for a role of Dsh in the establishment of dorsal–ventral axis in Xenopus. 9. Frizzled Frizzled proteins are receptors for Wnt ligands (Bhanot et al., 1996). Members of the Frizzled family contain a cysteine-rich domain, seven transmembrane domains, and sometimes a PDZ-binding domain. Overexpression of Xfrizzled-8 (Xfz8) alone on the ventral side of an embryo induces a secondary axis in the absence of exogenously supplied Wnt ligands (Deardorff et al., 1998). However, since the endogenous expression of Xfz8 is zygotically activated in the organizer region during gastrulation, Xfz8 is unlikely to function as the endogenous Wnt receptor for the dorsal specification pathway during the early cleavage of Xenopus embryos. A maternally encoded frizzled protein, Xfz7, has recently been isolated (Sumanas et al., 2000). Embryos depleted of Xfz7 mRNA are deficient in dorsal mesoderm formation. This study provides experimental evidence for the functioning of the Frizzled proteins upstream of other components of the Wnt pathway in dorsalventral mesoderm specification. 10. Wnt Molecules If an endogenous Wnt ligand is involved in the activation of the VCC/␤-catenin signaling pathway, such a Wnt molecule has not yet been identified. Several Wnt ligands have been studied, and these can be divided into two classes according to their axis-inducing ability. Mouse Wnt-1 (McMahon and Moon, 1989; Sokol et al., 1991), and several Xenopus Wnts, including Xwnt-8 (Smith and Harland, 1991), Xwnt-8b (Cui et al., 1995), Xwnt-2b (Landesman and Sokol, 1997), and Xwnt3A (Wolda et al., 1993), can induce a secondary axis when they are expressed ectopically in the ventral region. In contrast, Xwnt-4 (Du et al., 1995), Xwnt5A (Moon et al., 1993), and Xwnt-11 (Ku and Melton, 1993) have weak or no axis-inducing ability but cause morphogenetic defects when they are expressed ectopically. However, the Wnt molecules that have strong axis-inducing activity

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are not expressed at the right time and right place to be the dorsalizing signal (Moon and Kimelman, 1998). Overexpression of Xwnt-8 can induce a complete secondary axis, but endogenous expression is not detected until after the MBT in the ventral-lateral mesoderm. Although Xwnt-8b is maternally expressed and has strong-axis-inducing activity, it is localized to the animal region of cleavage-stage embryos (Cui et al., 1995). Xwnt-11 mRNA is localized to the vegetal cortex of the oocyte (Kloc and Etkin, 1995) and the protein is differentially translated along the dorsal–ventral axis (Schroeder et al., 1999); the possible involvement of Xwnt-11 in the VCC/␤-catenin signaling has yet to be demonstrated. It has recently been shown that Xwnt-11 does not signal through the canonical Wnt pathway involving GSK-3 and ␤-catenin for the regulation of morphogenetic movements (Heisenberg et al., 2000; Tada and Smith, 2000). This result demonstrates the divergence of signal transduction cascades that Wnt molecules can elicit. 11. Integration of the VCC/␤-catenin Signaling Pathway with Other Pathways Some Wnt molecules have been shown to signal through pathways independent of the ␤-catenin pathway. Wnt-5A signaling can increase the intracellular Ca2+ level in zebrafish embryos through a sequential action of the rat frizzled-2 receptor, G proteins, and stimulation of the phosphatidylinositol cycle (Slusarski et al., 1997). It has also been demonstrated that protein kinase C is a downstream component of the Xwnt-5A/rat frizzled-2 receptor G-protein-dependent pathway (Sheldahl et al., 1999). Treatment of embryos with lithium chloride during early cleavage stage produces a dorsalized phenotype. It was originally suggested that the dorsalizing effect of lithium is due to an activation of the phosphatidylinositol cycle (Maslanski et al., 1992). However, it was later demonstrated that lithium chloride treatment can inhibit GSK-3 activity, and the dorsalizing effect of the lithium ion is mediated by an activation of the ␤-catenin pathway (Klein and Melton, 1996; Stambolic et al., 1996). The activity of GSK-3 is also regulated by the ribosomal S6 protein kinase p90(rsk) (Torres et al., 1999). p90(rsk) inhibits GSK-3 by phosphorylation, and results in an increase of the total amount of ␤-catenin. Fibroblast growth factor (FGF) signaling has been shown to inhibit GSK-3 activity and can also activate p90(rsk). FGF signaling and p90(rsk) may play a role in modulating the activities of GSK-3 and ␤-catenin during early development.

E. Molecular Nature of the Dorsal Determinant in Vegetal Cortical Cytoplasm Like overexpression of ␤-catenin and Wnts, overexpression of wild-type dsh induces ectopic axis formation. A dominant negative form of dsh, Xdd1, although

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able to block ectopic axis formation by the activation of Wnt signaling, has no effect on endogenous axis formation (Sokol, 1996). Similar results have been obtained for Wnt molecules in dominant negative forms (Hoppler et al., 1996), frizzled proteins (Xu et al., 1998), and Wnt antagonists (Glinka et al., 1998). One interpretation of these results is that the VCC/␤-catenin pathway is not triggered by the presence of a Wnt ligand, but that a component of the Wnt pathway initiates the signaling cascade. Evidence supporting this model came from the study in which Dsh is shown to translocate from the vegetal pole toward the prospective dorsal side (Miller et al., 1999). This finding provides a link between the dorsal activity localized to the vegetal cortical cytoplasm of the egg and the activation of the VCC/␤-catenin signaling pathway on the dorsal sector of the embryo. However, the possibility that translocation of dsh toward the future dorsal side is preceded by the signaling of a Wnt ligand at the vegetal cortex of the egg should not be ruled out. The effect of Xdd1 overexpression has also been examined in prospective ectoderm transplanted with VCC (Marikawa and Elinson, 1999). In keeping with the findings in embryos, overexpression of Xdd1 in prospective ectoderm has no effect on the activity of VCC, as demonstrated by the expression of target genes siamois and Xnr3. Overexpression of different components of the Wnt pathway has shown that the activity of VCC is not inhibited by Xdd1 or Xgsk-3 but is inhibited by Axin. On the basis of these findings, it has been proposed that VCC may in fact act on Axin instead of Xgsk-3 to mediate its dorsalizing activity. This model is distinct from the general concept that an inhibition of Xgsk-3 activity by the endogenous dorsalizing signal is a prerequisite for ␤-catenin-mediated gene expression in the dorsal region.

F. Cooperation between TGF-␤ Signaling and Wnt Signaling Activation of the VCC/␤-catenin pathway alone is not sufficient for specification of the dorsal–ventral axis. In UV-irradiated embryos, which lack any dorsal structures, the VCC/␤-catenin pathway is still activated in the vegetal pole and siamois, a transcriptional target of the Wnt pathway, is expressed in the vegetal pole region (Darras et al., 1997). Furthermore, transplantation of vegetal cortical cytoplasm, or overexpression of siamois, only induces ectopic axis formation when the recipient site is the equatorial region but not when the recipient site is the animal region (Kageura, 1997). Thus, the VCC/␤-catenin pathway has to synergize with other components in the equatorial region to specify axis formation. One distinct possibility is that there is a cooperation between the VCC/␤-catenin and TGF-␤ signaling pathways. Evidence for this includes the induction of notochord in prospective ectoderm by Xwnt-8 and the TGF-␤ signaling molecule, Vg1 (Cui et al., 1996). Overexpression of Vg1 alone induces dorsal mesoderm but no notochord, and Xwnt-8b alone does not induce mesoderm formation. Coexpressing components of the Wnt and TGF-␤ pathways enhances expression of both the Wnt-responsive gene siamois and the activin/TGF-␤-responsive genes goosecoid (gsc) and chordin (Crease et al., 1998). Both TGF-␤ response elements and Wnt

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response elements are present in the promoter of the organizer gene gsc. In addition, it is also shown that both Wnt and TGF-␤ signaling are required for specifying formation of anterior endoderm. All these findings further strengthen the idea of cooperativity between the Wnt and the TGF-␤ pathways (Watabe et al., 1995). In overexpression experiments, several TGF-␤s have been shown to induce dorsal development. For dorsal development to occur, the endogenous ligand has to be active in the embryo at the right time and place. Spatial restriction of Vg1 mRNA to the vegetal cortex is suggested to provide a localized source of maternal protein to act as a dorsal inducer (Thomsen and Melton, 1993). Overexpression of a mutant form of Vg1 cannot perturb axis formation, although dorsal mesoderm and endoderm formation are affected (Joseph and Melton, 1998). This finding indicates that Vg1 is required for dorsal development, but expression of Vg1 alone is not sufficient in specifying formation of the dorsal–ventral axis. Activin protein is present maternally and may be a candidate for the endogenous TGF-␤ signal involved in dorsal–ventral axis formation (Fukui et al., 1994). Results from the overexpression of a dominant negative activin type II receptor in Xenopus embryos, although they have demonstrated the requirement of TGF-␤ signaling, have not been conclusive because of the inhibition of additional TGF-␤ members (Hemmati-Brivanlou and Melton, 1992). The use of a specifically designed dominant negative activin type II receptor, containing only the extracellular domain and lacking the transmembrane domain and intracellular domain, has circumvented the problem of nonspecific interference with other receptors at the cell surface (Dyson and Gurdon, 1997). This dominant negative receptor selectively blocks the function of activin but not that of Vg1 and nodal-related factors Xnr1 and Xnr2, although BMP signaling is slightly inhibited. Overexpression of this dominant negative activin type II receptor in Xenopus embryos has demonstrated the requirement of activin for the development of dorsal structures and the initiation of mesoderm induction. Furthermore, although knockout mice deficient in different activin subunits also show no defects in early development (Matzuk et al., 1995a), a type I activin receptor knockout does produce a defect in gastrulation (Gu et al., 1998). Therefore, it is likely that an activin-like TGF-␤ is required for early development. The zygotically activated Nodal-related TGF-␤s are the best candidates for the endogenous TGF-␤ signals involved in mesoderm and endoderm formation.

G. TGF-␤ Receptors, Smads, and Target Genes Members of the TGF-␤ superfamily are involved in a wide variety of cellular processes, including cell growth and differentiation (Massague, 1998). Two major types of serine/threonine kinase receptors, types I and II, are required for TGF-␤ signaling. Upon ligand binding of TGF-␤ or activin, type II receptors recruit and phosphorylate type I receptors (Fig. 2). The signal is transduced from the activated type I receptors by Smad proteins, which shuttle between the cytoplasm and the

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Figure 2 Components of the TGF-␤ signaling pathway. TGF-␤ ligands are first recognized by the type II receptor, which recruits and phosphorylates the type I receptor upon ligand binding. The signal is transduced by the R-Smad, which is activated by the type I receptor. The R-Smad associates with a co-Smad and translocates into the nucleus. The R-Smad and co-Smad complex with additional transcriptional factors to activate expression of target genes such as mix2 and gsc.

nucleus to control target gene expression (Massague, 1998). Specific Smad proteins transduce signals from different subgroups of TGF-␤ molecules. Several Smads function as transcription factors. Smads proteins specific for TGF-␤ signaling can bind to activin-response elements in the promoter of activin responsive genes. Three classes of Smad proteins have been identified (Christian and Nakayama, 1999). Receptor Smads (R-Smads) are phosphorylated by the intracellular domain of type I receptor. The activated R-Smads escort a second class of Smads, the coactivator Smads (co-Smads), into the nucleus to control target gene expression. The third class of Smads, the inhibitory Smads, prevent the R-Smads from binding to the type I receptor or the co-Smads. The Smad proteins contain three domains: a MH1 domain, a linker region, and a MH2 domain. The activity of Smad proteins is under constitutive inhibition by interaction between the MH1 and MH2 domains (Hata et al., 1997). Such autoinhibition is released by the phosphorylation of the MH2 domain at a C-terminal motif of R-Smads. The MH1 domain is required for DNA binding, whereas the MH2 domain plays a role in transcriptional activation and interaction with type I receptors, binding with co-Smads and other DNA-binding

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proteins (Baker and Harland, 1996; Liu et al., 1996, 1997; Meersseman et al., 1997). Smad2 and Smad3 are the R-Smads downstream of TGF-␤1 and activin signaling (Baker and Harland, 1996; Chen et al., 1996b; Graff et al., 1996; Lagna et al., 1996; Nakao et al., 1997b). Smad1, Smad5, and Smad8 are downstream of BMP activation (Graff et al., 1996; Hoodless et al., 1996; Liu et al., 1996; Thomsen, 1996; Chen et al., 1997; Suzuki et al., 1997a). Smad4 is a co-Smad (Lagna et al., 1996). Smad6 is an inhibitory SMAD for BMP signaling (Imamura et al., 1997; Hata et al., 1998). Smad 7 is an inhibitory Smad for BMP and TGF-␤ signaling (Hayashi et al., 1997; Nakao et al., 1997a). In addition to the inhibitory Smads, other mechanisms regulating TGF-␤ signaling have been reported in Xenopus embryos. BAMBI, a transmembrane protein related to TGF-␤ type I receptors but lacking an intracellular kinase domain, has been identified as a pseudoreceptor (Onichtchouk et al., 1999). Expression of BAMBI is induced by BMP signaling. BAMBI has been shown to associate with TGF-␤ family receptors to inhibit BMP, activin, and TGF-␤ signaling. Smurf1, a ubiquitin ligase, can interact with and trigger ubiquitination and consequently the inactivation of Smad proteins specific for the BMP pathway (Zhu et al., 1999a). Overexpression of Smurf1 can enhance the cellular responses to Smad2, which mediates the activin/TGF-␤ pathway. Smurf1 may conrol the competence of cells to respond to different TGF-␤ signals by regulating the levels of Smad proteins specific for BMP signaling (Zhu et al., 1999a). A maternal forkhead domain DNA-binding protein, FAST-1, was first identified as a component of a transcriptional complex containing Smad2 and Smad4 involved in the activation of Mix.2, an immediate response gene of activin signaling (Chen et al., 1996a). The transcriptional complex that binds to the gsc promoter contains FAST-2 (Labbe et al., 1998). Overexpression of FAST-1 fusion constructs containing either a transcriptional activation domain or a repressor domain has shown that FAST-1 is in fact involved in the induction of a set of activin/Vg1 responsive mesodermal and endodermal genes. This finding suggests that FAST-1 is a key maternal regulator of transcriptional responses to mesoderm inducers (Watanabe and Whitman, 1999).

IV. The Spemann Organizer In amphibians, the organizer forms at the dorsal lip of the blastopore of the gastrula embryo. The organizer is known as the Spemann organizer because the axisinducing activity of this region was demonstrated for the first time in 1921 using urodelean amphibians by Hilda Mangold, a student of Hans Spemann (Hamburger, 1988). Mangold transplanted dorsal blastoporal lips of advanced gastrulae of unpigmented newts to the flanks of pigmented host newts at the same stage of development (Spemann and Mangold, 1924). In the most successful case, the

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resulting embryo had a secondary body axis containing anterior structures that included hindbrain and otic vesicles. In the secondary axis, the neural tube was composed entirely of cells that originated from the pigmented host embryo; the notochord contained cells that originated from the unpigmented transplanted tissue; and the somites were derived from a mixture of cells from both origins. The ability of the transplanted dorsal blastoporal lip to recruit cells from the host embryo to participate in the formation of the secondary axis demonstrated that the transplaned tissue had an organizing ability. The inability of the dorsal lip of advanced gastrula to generate a complete secondary axis including the head prompted Spemann to test the hypothesis that the head and trunk were induced by different regions of the mesodermal tissue of the organizer. Since the organizer is a highly dynamic structure and cells at the blastoporal lip continously undergo involution, the cell population at the blastoporal lip is distinct during different stages of gastrulation. These cells assume a progressively more posterior identity as gastrulation proceeds. By transplanting dorsal lip tissues to the ventral side of the blastocoel, Spemann demonstrated that blastoporal lip from the early gastrula induced head and brain structures while blastoporal lip from the late gastrula induced spinal cord and tail structures. Spemann introduced the terms “head organizer” and “trunk organizer” to specify the regional and temporal differences of the organizer (Spemann, 1927). Regional specification of organizer-derived tissue was further demonstrated by Otto Mangold (Mangold, 1933). O. Mangold showed that, at the neurula stage, the organizer-derived mesodermal tissue layer had assumed an anteroposterior identity and could be divided into regions including the anterior endomesoderm, the prechordal mesoderm, and the anterior and posterior chordamesoderm. Of these, only the prechordal mesoderm, fated to form the head mesenchyme, demonstrated head-inducing ability. The anterior endomesoderm, the most anterior region containing cells of endodermal and mesodermal origins and fated to give rise to the liver, showed little or no head-inducing activity. The anterior region of the chordamesoderm gave rise to the hindbrain and spinal cord, whereas the posterior region of the chordamesoderm formed only the spinal cord. In Xenopus, the organizer is formed in the dorsal vegetal region of the embryo as a consequence of signaling by TGF-␤ and VCC/␤-catenin (Harland and Gerhart, 1997; Heasman, 1997; Niehrs, 1999, 2000). Molecular characterization of the organizer has led to the identification of genes that are specifically expressed in the organizer and can induce secondary axes by acetopic expression. These organizer genes can be classified into three groups on the basis of the type of secondary axis generated. However, it should be noted that the organizer tissue is a dynamic structure. The organizer is not a constant population of cells, and considerable cell movement and rearrangement take place during gastrulation. The first group of organizer genes are transcriptional targets of the VCC/␤-catenin pathway, and some are expressed in the dorsal vegetal region well before the appearance of the dorsal lip during the blastula stage. The products of some of these genes can induce

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Figure 3 Expression domains of organizer genes in the head and trunk organizers. The organizer consists of four expression domains: anterior endomesoderm, prechordal mesoderm, superficial layer, and chordamesoderm. The head organizer is made up of the prechordal mesoderm and anterior endomesoderm. The chordamesoderm possesses trunk organizer activity. (modified from Harland and Gerhart, 1997; with permission, from the Annual Review of Cell and Developmental Biology, Volume c 13, 1997, by Annual Reviews www.AnnualReviews.org)

complete secondary axes containing head and trunk when the genes are overexpressed on the ventral side of the embryo. The second group of organizer genes are expressed in the prechordal mesoderm region of the head organizer at the gastrula stage and can generate ectopically incomplete secondary axes lacking anterior structures. The last group of organizer genes are expressed in the anterior endomesoderm of the head organizer at the gastrula stage. The product of some of these genes can produce an ectopic head without a trunk by overexpression. The expression domains of the organizer genes at the gastrula stage are summarized in Fig. 3.

A. Organizer Genes Expressed in the Dorsal Vegetal Region siamois and twin are paired-like homeobox genes that are transcriptional targets of VCC/␤-catenin signaling (Brannon et al., 1997; Laurent et al., 1997; Fan et al., 1998). Overexpression of either siamois or twin induces ectopic secondary axis with a complete head (Lemaire et al., 1995; Laurent et al., 1997). siamois is expressed in the dorsal vegetal region soon after the MBT. Vegetal cortical cytoplasm can induce siamois expression ectopically, indicating that siamois is a downstream target of the dorsal determinant in the vegetal cortical cytoplasm (Darras et al., 1997). Transplantation of vegetal cortical cytoplasm to the animal hemisphere activates the expression of chordin, whereas transplantation of vegetal cortical

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cytoplasm to the vegetal hemisphere activates the expression of cer. This shows that different regions along the animal–vegetal axis have diferent abilities to respond to the cytoplasmic dorsal determinant. Activation of cer expression in the vegetal hemisphere by siamois induces the formation of anterior endomesoderm, whereas the activation of chordin in the animal hemisphere induces the formation of the head organizer (Darras et al., 1997). siamois is therefore able to induce the formation of the anterior endomesoderm and the head and trunk organizers. Since overexpression of siamois mRNA in either the ventral vegetal region or the equatorial region results in ectopic axis formation (Laurent et al., 1997), the action of siamois may be mediated by a diffusible growth factor that instructs cells in the equatorial region to participate in axis formation. twin has axis-inducing properties similar to those of siamois. Like Siamois, Twin has been shown to bind to and activate the Wnt-responsive regulatory elements of the gsc promoter (Laurent et al., 1997). In zebrafish, a paired-like homeobox gene, nieuwkoid/dharma encoded by the bozozok locus, expressed in dorsal blastoderm and dorsal yolk syncytial layer has demonstrated organizer gene activity (Koos and Ho, 1998; Yamanaka et al., 1998; Fekany et al., 1999). The dorsal yolk syncytial layer forms at the blastula stage when cells in the deep marginal blastoderm release their nuclei into the yolk cells. bozozok mutants are deficient in activity and exhibit a loss of shield derivatives, equivalent of the Xenopus organizer, and anterior neural structures. Although sequence comparison does not suggest the existence of an ortholog for nieuwkoid/dharma, the involvement of a paired-like homeobox gene in organizer formation both in frogs and in teleofish indicates a potential conserved role of the paired-like homeobox transcription factor in other developmental systems.

B. Organizer Genes Expressed in the Prechordal Mesoderm Organizer genes expressed in the prechordal mesoderm include genes that code for transcription factors (Gsc, Xlim-1, Xanf-1, Xotx2), growth factor antagonists (noggin, chordin, follistatin, frzb, dickkopf-1 [dkk-1]), and growth factors (Xnr14 and anti-dorsalizing morphogenetic protein [ADMP]). Most of these genes are expressed above the dorsal lip in the prechordal mesoderm, which contributes to the head mesenchyme. dkk-1, Xnr-1, and Xnr-2 are also expressed in the anterior endomesoderm. chordin is expressed in the prechordal mesoderm, including the superficial layer. Xnr-3 is most strongly expressed in the superficial layer of the dorsal lip, which gives rise to the pharyngeal endoderm. ADMP is related to BMP3. Overexpression of ADMP inhibits dorsal mesodermal markers, including organizer genes, and induces ventral markers (Moos et al., 1995). It has also been shown that ADMP is expressed in the trunk organizer (Dosch and Niehrs, 2000). Since the function of ADMP cannot be blocked by a dominant-negative BMP receptor or other BMP antagonists except follistatin, it may function in the trunk organizer to antagonize head formation by a distinct pathway (Dosch and Niehrs, 2000).

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1. Genes Encoding Transcription Factors The transcription factor Gsc contains a paired class homeodomain and is able to generate an incomplete secondary embryonic axis when overexpressed (Cho et al., 1991; Steinbeisser et al., 1995). Dissociated Xenopus embryos express the same level of gsc as undissociated embryos, indicating that the expression of gsc is cell autonomous—that is, independent of cell–cell interaction (Lemaire and Gurdon, 1994). Analysis of isolated organizer explants has demonstrated that by the early gastrula stage, gsc expression is already limited to a discrete region of the organizer (Zoltewicz and Gerhart, 1997). gsc is expressed in the anterior half of isolated organizer explants. The gsc expression domain is spatially distinct from the Xnot expression domain in the posterior half, which normally gives rise to the chordamesoderm. In the mouse, gsc is expressed in the anterior primitive streak and the anterior mesoderm, which gives rise to the head process (Blum et al., 1992). gsc knockout mice have craniofacial and rib-cage defects, but their early development is largely unaffected (Rivera-Perez et al., 1995; Yamada et al., 1995). However, the neural-inducing strength of the mouse node from gsc knockout mice is impaired, as demonstrated by studies in which gsc-deficient mouse node was transplanted to chick embryos (Zhu et al., 1999b). The presence of gsc-related genes in different systems, including Drosophila (Goriely et al., 1996; Hahn and Jackle, 1996), chicken (Lemaire et al., 1997) and human (Gottlieb et al., 1997) suggests that a related family member may compensate for gsc function in gsc knockout mice. A hydra homolog of gsc, Cngsc, has also been identified (Broun et al., 1999). Cngsc is expressed in tissues with organizer activity and is able to induce a secondary axis when expressed in Xenopus embryos. This suggests that the function of the Gsc protein has been conserved during evolution. In Xenopus, the requirement of gsc function has been demonstrated by studies involving overexpression of antimorphic forms of gsc, containing transcription activation domain or multiple copies of the myc epitope at the N terminus (Ferreiro et al., 1998). The transcriptional activation effect of the myc epitope is not expected, and caution should be taken when the myc epitope is used in the context of a putative transcription factor (Ferreiro et al., 1998). Antimorphic gsc is expected to inhibit the function of endogenous Gsc as well as other closely related family members, if they do exist in Xenopus. When antimorphic gsc is expressed on the dorsal side of the embryo, ventral genes are activated ectopically, and embryos exhibit dorsoanterior defects. This finding shows that Gsc normally acts as a transcriptional repressor inhibiting ventral gene expression in the organizer region, and is consistent with the suggestion that Gsc functions as a transcriptional repressor to directly suppress the transcription of Xbra (Artinger et al., 1997; Latinkic et al., 1997). Overexpression of antimorphic Gsc also activates expression of endogenous gsc, indicating a possible self-repression of gsc in embryos. Like gsc, the mutant form of the LIM-domain-containing homeobox gene Xlim-1 (Taira et al., 1994) and the homeobox genes Xanf-1 (Zaraisky et al., 1995) and Xotx2 (Pannese et al., 1995) also produce an incomplete secondary embryonic

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axis lacking anterior structures when these genes are overexpressed in embryos. In addition, Xotx2 induces ectopic cement gland formation in embryos (Blitz and Cho, 1995; Pannese et al., 1995). Mice deficient in lim1 have anterior truncations, including a missing forebrain and midbrain (Shawlot and Behringer, 1995). A study of otx2 knockout mice demonstrated a loss of anterior neural structures in these animals (Ang et al., 1996). Xanf-1 is related to mouse Hesx-1/Rpx, which may be involved in anterior development (Hermesz et al., 1996; Thomas and Beddington, 1996). Gain-of-function studies in frogs and loss-of-function studies in mice together suggest an important role of Xlim-1, Xanf-1, and Xotx2 in development of the most anterior part of the head.

2. Genes Encoding Growth Factor Antagonists The organizer genes noggin (Smith et al., 1993), chordin (Sasai et al., 1994), and follistatin (Hemmati-Brivanlou et al., 1994) encode for protein factors originating from the organizer. When these genes are ectopically expressed on the ventral side of frog embryos, a secondary axis with an incomplete anterior structure is generated. The protein products of these genes are structurally distinct and have been shown to function through their ability to antagonize, and thus inhibit, BMP signaling by direct binding to BMP molecules. Noggin is a glycoprotein secreted as a homodimer. Mouse noggin (McMahon et al., 1998) is expressed in the node, which gives rise to prechordal mesoderm and participates in head formation. noggin knockout mice do not show the early developmental defects that would be expected when an organizer gene is disabled (Brunet et al., 1998; McMahon et al., 1998). noggin-deficient mice have defects in somites and neural tube formation. This suggests that although noggin is not required for neural induction, it is important for later events, including patterning of somites and neural tube. Three noggin genes have been isolated from zebrafish (Furthauer et al., 1999). noggin 1 and noggin 2 are expressed in the organizer and the notochord, respectively, whereas noggin 3 is involved in a later stage of development for chondrogenesis. Although only a single noggin gene has been reported so far in other species, the finding of multiple noggin genes in zebrafish suggests that the functional redundancy of related genes should be taken into consideration. Chordin contains cysteine-rich domains (CRs) and is also secreted. The CRs have been shown to be novel protein modules for BMP binding and therefore confer the biological activity of chordin (Larrain et al., 2000). The zebrafish chordino mutant, originally known as dino, displays a partially ventralized phenotype with reduced head formation and, often, lacks a notochord, indicating an effect of chordin in organizer function (Hammerschmidt et al., 1996). It is therefore informative to study mice deficient in either chordin or noggin or both of the genes to determine their requirement in specifying anterior development and a possible functional redundancy of these genes during early development. Double-homozygous mutant mice for chordin and noggin have been generated (Bachiller et al., 2000). These

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embryos showed severe defects in the development of the forebrain. Chordin and Noggin are indeed required for anterior development in the mouse. BMP signaling has been shown to play a role in embryonic patterning (Dale and Jones, 1999). Inhibition of BMP signaling by a cleavage mutant generates secondary axes (Hawley et al., 1995), whereas activation of the BMP pathway by overexpression of BMPs or constitutively active BMP receptors results in a ventralized phenotype (Jones et al., 1996b). bmp-2, -4, and -7 mRNAs are maternally expressed and are detected in the entire animal hemisphere. Only bmp-4 mRNA, but not bmp-4 or bmp-7 mRNA, is downregulated at the organizer region when the organizer forms. Chordin and Noggin have been shown to mediate a dorsalizing or axis-inducing effect by binding to BMPs and inhibiting BMP signaling in the organizer (Holley et al., 1996; Piccolo et al., 1996; Zimmerman et al., 1996). Chordin can bind BMP-2, BMP-4, and BMP-4/7 heterodimer but not TGF-␤. The antagonistic effect between Chordin and BMP is evolutionarily conserved, as demonstrated by the Drosophila homologs Sog and Dpp and the functional substitution of Chordin and BMP by Sog and Dpp in frog embryos (Sasai et al., 1995). Noggin can bind BMP-2 and BMP-4, and can bind BMP-7 less tightly, but is not able to bind TGF-␤. Noggin has a higher affinity to BMPs (K D = 20 pM) than does Chordin (K D = 300 pM, 1 nM for inducing neural response and dorsalization of mesoderm). The binding affinities between BMPs and their receptors are in the same range as the binding affinities between BMPs and Noggin or Chordin. An additional level of regulation of BMP signaling is carried out by a metalloprotease of the astacin family (Dumermuth et al., 1991), Tolloid, which can inhibit the antagonistic function of Chordin on BMP signaling. Genetic studies in Drosophila have shown that tolloid can potentiate the effects of Dpp (Shimell et al., 1991; Ferguson and Anderson, 1992). Xolloid and BMP-1, related proteins of the tolloid family, contain a metalloprotease domain that is followed by CUB (Cls and Clr/Uegf/BMP1) domains and EGF-like domains (Piccolo et al., 1997; Goodman et al., 1998; Wardle et al., 1999b). The CUB domains may be required for protein–protein interaction—for example, to interact with the substrate. Xolloid has been demonstrated to cleave Chordin within specific sites. This suggests that Xolloid can negate the inhibitory effects of chordin on BMP signaling. In Xenopus embryos, overexpression of Xolloid and XBMP-1 results in a ventralized phenotype as expected from the removal of Chordin from the embryo. Dominant negative forms of Xolloid and XBMP-1 produce embryos with a dorsalized phenoptype with enlarged heads and anterior structures, although the deletion mutant inhibits the activities of both Xolloid and XBMP-1. In zebrafish, Xolloid is encoded by the mini fin (mfn) gene (Connors et al., 1999). mfn mutants exhibit a loss of ventroposterior tissues, including the ventral fin, that is due to a dorsalization resulting from diffusion of Chordin to the most ventral marginal regions at the end of gastrulation. In wild-type embryos, the presence of Mfn may negatively regulate Chordin activity and promote BMP signaling in the ventral marginal cells. A related sea-urchin metalloprotease, SpAN, when expressed in Xenopus embryos, can block the dorsalizing activity of both noggin and chordin

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(Wardle et al., 1999a). Since SpAN does not cleave noggin or chordin directly, binding of Noggin or Chordin to BMPs may be a prerequisite for SpAN to function (Wardle et al., unpublished observations, 1999a). It has also been suggested that SpAN may trigger the release of BMPs from other BMP-binding proteins or the extracellular matrix. The mechanism by which SpAN inactivates the dorsalizing function of noggin and chordin has yet to be determined. A recently identified member of the BMP-4 synexpression group, Xenopus Twisted gastrulation (xTsg), encodes a secreted BMP-binding protein. xTsg functions as antagonist to provide a permissive signal for BMP signalling (Oelgeschlager et al., 2000). Follistatin was originally suggested to bind Activin directly and thereby control the amount of free Activin (Tashiro et al., 1991). However, in vitro data have demonstrated an affinity of Follistatin for BMP-4 (Fainsod et al., 1997; Iemura et al., 1998). The Follistatin/BMP-4 complex can bind to BMP receptors, suggesting that Follistatin regulates BMP signaling through a mechanism different from that by which Noggin and Chordin regulate BMP signaling (Iemura et al., 1998). The organizer function of Follistatin is further complicated by the fact that follistatin is not expressed in the equivalent of the organizer in zebrafish (Bauer et al., 1998) or mouse (Albano et al., 1994) and is only weakly expressed in chicken (Levin, 1998). In addition, no axial defects are detected in follistatin knockout mice (Matzuk et al., 1995b). This result argues against a requirement for follistatin during early development. The role of follistatin in the inhibition of BMP signaling and organizer activity requires further investigation. frzb, also known as frzb1, is expressed in the organizer region (Leyns et al., 1997; Wang et al., 1997). The expression pattern of frzb1 is complementary to endogenous Xwnt-8 expression in the ventral lateral mesoderm. Frzb functions as a growth factor antagonist by direct binding to Wnts and thus inhibition of Wnt signaling. Frzb belongs to a clas of proteins that are known as frizzled-related proteins (FRPs) because their structure is similar to that of the membrane-bound Wnt receptor of the frizzled family, except that FRPs lack the transmembrane domain. Overexpression of frzb generates a partial secondary axis at a low frequency, and overexpression of frzb in whole embryos causes dorsalization, with embryos showing enlarged heads and shortened body axes. Frzb also inhibits the effect of ectopic Xwnt-8 expression in a non-cell-autonomous manner, suggesting that Frzb functions extracellularly to suppress Wnt function. A related protein, FrzA, when overexpressed in embryos, shows a phenotype similar to that produced by overexpression of frzb in embryos (Xu et al., 1998). However, frzA is not involved in organizer function because expression commences at the neurula stage in the somitic mesoderm.

C. Organizer Genes Expressed in the Anterior Endomesoderm The identification of genes that are specifically expressed during gastrulation in the anterior endomesoderm has suggested a crucial role of this region as part of the head organizer for the generation of all the anterior structures of an axis.

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The anterior endomesoderm includes the deep dorsoanterior endodermal cells that do not undergo cell involution. The prechordal mesoderm, on the contrary, does involute when the dorsal lip forms. Several genes have been identified in the anterior endomesoderm of the Xenopus head organizer. Xnr-1, -2, and -4, Xblimp, and XHex can regulate specific gene expression in the anterior endomesoderm. The cer and dkk-1 genes encode growth factor antagonist expressed in the anterior endomesoderm and have been suggested to be involved in anterior development. 1. Genes That Regulate Anterior Endomesoderm Formation Xnr-1 and Xnr-2 are expressed in the anterior endomesoderm as well as the prechordal mesoderm (Jones et al., 1995). The involvement of the Xnrs in axis induction has been suggested by the observation that a complete secondary axis forms when Xnr-1 is coexpressed with noggin (Lustig et al., 1996a). noggin alone can only generate an incomplete secondary axis. Xnr-1 and Xnr-2 can induce the expression of anterior endomesoderm markers, including XHex, cer, frzb1, and Xsox17␤, in prospective ectodermal explants (Zorn et al., 1999b). The use of a cleavage mutant form of Xnr-2, cmXnr2, permits loss-of-function analysis of Xnr-2. Overexpression of cmXnr2 results in anterior truncation and delayed or suppressed expression of dorsoanterior endodermal genes (Osada and Wright, 1999). A similar phenotype has also been observed with the overexpression of a mutant Activin type II receptor containing the extracellular domain (Dyson and Gurdon, 1997) or a Smad2 dominant negative construct (Hoodless et al., 1999). In zebrafish, a similar phenotype is observed in the cyclops/squints and MZoep mutants (Feldman et al., 1998; Gritsman et al., 1999). cyclops and squint are nodal-related genes. MZoep is a mutant lacking both maternal and zygotic activities of oep, which is a member of the EGF-CFC (epidermal growth factor-Cripto/Frl-1/Cryptic) family. Oep is membrane-bound and is produced extracellularly in cells responsive to Nodal. Oep functions as an essential cofactor to facilitate Nodal signaling. The mutant phenotype of MZoep can be rescued by overexpressing activin, activated activin receptor, and Smad2. This suggests that Nodal signaling activates an activin-like pathway during early embryonic patterning. In the frog, a temporal and spatial regulation of Nodal signaling is required for the development of the most anterior structures. Maternal factors (Vg1, VegT, or both) may activate the expression of Xnrs, which induce the formation of the anterior endomesoderm through an activin-like pathway. It is thought that one of the functions of the anterior endomesoderm is then to create a Nodal-free zone, by the expression of growth factor antagonists, in the anterior endomesoderm within the head organizer for anterior development (Piccolo et al., 1999). The VCC/␤-catenin pathway is also involved in inducing expression of genes that regulate anterior endomesoderm formation. When a N-Tcf3 mutant is overexpressed in embryos, expression of XHex and cer is inhibited (Zorn et al., 1999b). Overexpression of a dominant negative Siamois mutant also represses cer

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expression in embryos (Darras et al., 1997). This indicates that a functional Wnt pathway is required for anterior endomesoderm formation. It is also shown that overexpression of bmp inhibits XHex and cer expression in embryos (Zorn et al., 1999b). Although overexpression of chordin or noggin cannot induce XHex or cer expression, they are required to suppress BMP signaling to maintain XHex and cer expression in the anterior endomesoderm. Xblimp1 encodes a zinc-finger transcription repressor and is similar to the mammalian PRDI-BF1/Blimp-1 gene (de Souza et al., 1999). Expression of Xblimp1 is detected in the anterior endomesoderm and prechordal mesoderm. Overexpression of Xblimp1 mRNA on the ventral side of the embryo induces dorsoanterior marker genes, including cer, gsc, and Xotx2, but not frzb or dkk-1. This shows that Xblimp is able to regulate expression of specific genes in the anterior endomesoderm. A complete secondary axis can be generated by the overexpression of Xblimp1 and the BMP-antagonist chordin in embryos. XHex is a transcription factor expressed in the anterior endomesoderm (Newman et al., 1997; Jones et al., 1999). The expression domain of XHex largely overlaps with that of cer in the deep dorsal marginal cells in blastula- and gastrula-stage embryos (Zorn et al., 1999b). Overexpression of XHex induces cer expression in explants derived from ventral endoderm. The mouse homolog Hex is expressed in the primitive endoderm of mouse blastocysts and later in the visceral endoderm at the distal tip of the egg cylinder (Thomas et al., 1998). Hex is one of the earliest markers of the anterior visceral endoderm (AVE) in mouse embryos and is initially detected in the primitive endoderm of blastocysts. The AVE is involved in setting up an early asymmetry of the mouse embryo before the node is formed and has been suggested to be analogous to the anterior endomesoderm in Xenopus (Bouwmeester and Leyns, 1997; Beddington and Robertson, 1998). In both mouse and frog embryos, expression of Hex is also detected in the angioblasts, which are precursors for the blood cells and endothelium during vasculogenesis (Newman et al., 1997; Thomas et al., 1998). It has been suggested that Hex could be a marker gene for stem cell populations of endodermal origin. 2. Growth Factor Antagonists Expressed in the Anterior Endomesoderm Cer is a growth factor antagonist expressed in the anterior endomesoderm of Xenopus embryos (Bouwmeester et al., 1996). Cer contains a single cysteine-rich domain containing conserved cysteine residues and is secreted. In Xenopus, although the anterior endomesoderm does not show any head-inducing property, overexpression of cer can generate ectopic head structures in the absence of a trunk. Such overexpression of cer may activate additional organizer genes that function in conjunction with cer to induce an ectopic head. Cer also demonstrates a neuralizing activity as shown by the induction of the forebrain marker Xotx2 and neural marker NCAM in prospective ectoderm (Bouwmeester et al., 1996). Several mammalian

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proteins have been identified that are related to Cer and the product of the tumor suppressor gene DAN (Ozaki and Sakiyama, 1993). These proteins form the Cerberus/DAN family, which is also known as the Can family (Pearce et al., 1999). A C. elegans homolog of the Can family, CeCan1, has also been identified, suggesting that the protein is highly conserved throughout evolution (Pearce et al., 1999). Overexpression of cer inhibits the functions of Xnr-1, BMP, and Xwnt-8 through direct binding of different regions of the Cer protein to these growth factors (Piccolo et al., 1999). A truncated form of the Cer protein, Cer-S, retains only the site for Nodal binding and can specifically inhibit Nodal signaling. The ventral marginal zone can be induced to show headlike properties by inhibiting BMP and Nodal signaling with dominant negative BMP receptors and Cer-S. It has been suggested that expression of cer in the anterior endomesoderm results in a zone free of Nodal, BMP, and Wnt molecules and is required for head development. However, ablation of cer-expressing tissue in the endoderm affects formation of the heart, but has no effect on head development (Schneider and Mercola, 1999). Furthermore, mouse deletion mutants with the cer1 locus deleted have been shown to develop normally without any defects in anterior patterning (Simpson et al., 1999). Mouse homozygous embryos deficient in a cerberus-like gene also showed the same result. One interpretation of these results is that functional redundant protein products of the same family may compensate for the requirement of Cer during early development (Belo et al., 2000). Dkk-1 is a Wnt antagonist expressed in the anterior endomesoderm, prechordal mesoderm, and anterior chordamesoderm (Glinka et al., 1998). Members of the Dkk protein family are secreted proteins containing two cysteine-rich regions with conserved cysteine residues in each region. Several related proteins have been identified in chicken and mouse (Monaghan et al., 1999). A family of Dkk-related proteins have been identified in human, including hDkk-1, -2, -3, and -4 and Soggy (Krupnik et al., 1999). Soggy is related to Dkk-3 but lacks the cysteine domains. hDkk-2 and hDkk-4 undergo processing resulting in removal of the second cysteine-rich region. Both hDkk-1 and hDkk-4 can suppress the secondary axis generated by ectopic expression of Wnt but not that generated by downstream components such as dsh and frizzled-8. This suggests that Dkk is likely to function upstream of the Wnt receptor to antagonize Wnt signaling. The frog homolog dkk-1 has been shown to antagonize the early axis-inducing ability and the late ventralizing effect of Xwnt8 (Glinka et al., 1998). dkk-1 cannot induce a secondary axis when it is expressed alone. It can do so only when BMP signaling is inhibited by a dominant-negative BMP receptor or BMP antagonists. This demonstrates that inhibition of Wnt and BMP activity alone is sufficient for head induction. However, this apparently conflicts with the requirement of the antagonistic function of Cer as well in inhibiting Nodal signaling during head formation. In fact, cer expression is induced by an inhibition of both BMP and Wnt signaling, thus indicating that Cer, and therefore a repression of Nodal activity in addition to that of BMP and Wnt, is involved in the development of anterior structures (Piccolo et al., 1999).

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Xfz8, a membrane-bound Wnt receptor, is also expressed in the anterior endomesoderm and dorsal lip region (Deardorff et al., 1998; Itoh et al., 1998). In late gastrula and neurula-stage embryos, Xfz8 is expressed in the most anterior ectoderm. Both wild-type Xfz8 and a dominant negative construct, ECD8, containing only the extracellular region, induce secondary axes when expressed ventrally in embryos (Itoh and Sokol, 1999). The in vivo function of Xfz8 has yet to be elucidated. Lineage labeling shows that cells expressing ECD8 mostly contribute to the head ectoderm, which induces the secondary notochord in a noncell-autonomous manner. ECD8 suppresses the activity of several Wnts, including ones that are not inhibited by other Wnt antagonists, such as Frzb. It has been suggested that a Wnt ligand required to suppress a dorsal cell fate in the ventral region is inhibited by ECD8. The mode of action of Xfz8 in the anterior endomesoderm requires further studies.

D. A Mammalian Structure Analogous to the Anterior Endomesoderm In frogs, the anterior endomesoderm is required for the formation of a complete axis including the most anterior structure, and anterior endomesoderm formation is regulated by TGF-␤, Wnt, and BMP signaling. Studies of the mouse AVE, which may be functionally analogous to the anterior endomesoderm in frog, have demonstrated the role of TGF-␤ signaling in specifying AVE formation during early embryonic development. In the mouse, the AVE is derived from the extraembryonic lineage and does not contribute to the formation of the embryo proper. During gastrulation, cells in the AVE are progressively displaced by the definitive endoderm that originates from the node (Lawson et al., 1991). The definitive endoderm in the AVE then differentiates into liver and gut endoderm. The generation of chimeric mouse embryos consisting of wild-type and mutant cells in specific cell lineages—such as the embryonic epiblast and the extraembryonic visceral endoderm—has provided a useful tool for determining the role of AVE during early patterning (Beddington and Robertson, 1999). Mice deficient in Nodal cannot gastrulate and show no anteroposterior specification (Conlon et al., 1991; Varlet et al., 1997). Introduction of wild-type cells into nodal-deficient blastocysts gives rise to chimeric embryos in which wild-type cells are included in the epiblast. In these embryos, the gastrulation defect but not the anterior defect is rescued, indicating that Nodal signaling is required in the epiblast for primitive streak formation and therefore gastrulation. Introduction of nodal-deficient cells into wild-type blastocysts gives rise to chimeric embryos in which the visceral endoderm is composed entirely of wild-type cells and the epiblast is composed of nodal-deficient cells and wild-type cells. Both the gastrulation defect and anterior–posterior patterning are rescued in these embryos. This finding suggests that Nodal signaling is required in the visceral endoderm during gastrulation for the specification of the anteroposterior axis of early embryos (Varlet et al., 1997).

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Studies in mice deficient in Smad2 have also demonstrated a role for Smad2 in early anteroposterior patterning and mesoderm formation (Nomura and Li, 1998; Waldrip et al., 1998; Weinstein et al., 1998). The extreme alleles show a complete lack of mesoderm formation. One of the studies demonstrated the formation of extraembryonic mesoderm, but more important, the study also showed the absence of anterior visceral marker expression, indicating a possible involvement of Smad signaling in anterior specification (Waldrip et al., 1998). The phenotype of Smad4 knockout mice is similar to the extreme phenotypes of Smad2 knockouts in terms of defective gastrulation and mesoderm formation (Sirard et al., 1998). Gastrulation is rescued in chimeric embryos containing wild-type extraembryonic tissue and defective Smad4 embryonic tissue. This indicates that Smad4 is required in the visceral endoderm to mediate gastrulation in the epiblast in a non-cell-autonomous manner. These findings substantiate the role of Nodal and Smad signaling in specifying the AVE during early axis formation. In Xenopus, some components of the Wnt pathway are required to specify the dorsal–ventral axis, but there is no evidence for the involvement of a maternal Wnt molecule. In contrast, studies in mouse has revealed early Wnt signaling during axis formation by the Wnt3 knockout study (Liu et al., 1999b). Mice deficient in Wnt3 do not gastrulate or form mesoderm. However, expression of the AVE marker genes tested is not affected. The involvement of a Wnt pathway in mesoderm formation in mouse embryos has also been demonstrated by ␤-catenin knockout mutants and Tcf1/Lef1 double mutants, which are defective in mesoderm formation (Haegel et al., 1995; Galceran et al., 1999). This indicates that Wnt signaling is required for mesoderm formation and gastrulation in mouse, but Wnt3 expression is not required for AVE formation. In the mouse embryo, it appears that mesoderm and primitive streak formation are regulated by both TGF-␤ and Wnt signaling and that anteroposterior specification of the visceral endoderm is mainly dependent on TGF-␤ signaling.

E. How Is the Organizer Formed after All? One possible model of organizer formation is the following: The vegetal hemisphere of the egg contains localized maternal components that are able to activate TGF-␤/Nodal signaling (Fig. 4). Fertilization, followed by cortical rotation, displaces a cytoplasmic dorsal determinant, an activator or a component of the Wnt pathway, to the dorsal vegetal region, where Wnt signaling interacts with the TGF␤ signaling pathway to induce the formation of the anterior endomesoderm and the organizer. TGF-␤/Nodal signaling alone is not sufficient for axis induction but requires interaction with the Wnt pathway to initiate early patterning. Thus, TGF␤ and Wnt signaling may be activated in parallel before the MBT. After the MBT, zygotic transcription provides an integration point when a combined effect from both pathways regulates expression of organizer genes and induces axis formation.

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Figure 4 VCC/␤-catenin signaling sets up dorsal–ventral differences. The sperm entry point (SEP) marks the future ventral side of the embryo. Upon fertilization, the egg cortex rotates toward the future dorsal side. The movement displaces cytoplasmic dorsal determinants, originally localized to the vegetal cortex, to the dorsal side of the embryo. The dorsal determinants activate the Wnt signaling pathway, resulting in the nuclear localization of ␤-catenin in dorsal cells. Wnt signaling and TGF-␤ signaling superimpose in the dorsal vegetal quadrant to initiate expression of organizer genes, such as gsc, during the MBT. Both Gsc and Vox are transcriptional repressors. gsc activates chordin expression indirectly through a double inhibition. The organizer expresses a number of growth factor antagonists to keep the region free of BMPs, Wnts, and Nodal signaling. These antagonists are also required to pattern the mesoderm and ectoderm, but it is not known if they are also required for endodermal patterning. BMP and Wnt signaling are required outside the organizer region for a ventral cell fate. Maternal TGF-␤s or VegT may be involved in the early phase of mesoderm induction. TGF-␤s are also zygotically activated in the vegetal region for the specification of mesoderm and endoderm.

V. The Three Germ Layers In almost all metazoans, the basic body plan is derived from three germ layers: the endoderm, mesoderm, and ectoderm. In Xenopus embryos, it has been suggested that the vegetal endoderm secretes a source of mesoderm-inducing signal that can direct neighboring ectodermal cells toward the mesodermal lineage. The mesoderm forms in the equatorial region in the presence of TGF-␤ signaling. The endoderm forms in the vegetal region, and formation of the endoderm requires a high level of TGF-␤ signaling. The ectoderm forms in the animal region in the absence of TGF-␤ stimulation from the vegetal region. A. Endoderm The endoderm arises from the yolky cells of tier D in the vegetal region of a 32-cell embryo. These vegetally derived blastomeres are fated to form the lining of the gastrointestinal and respiratory tracts and liver and pancreas (Chalmers

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and Slack, 1998; Wells and Melton, 1999). In tissue recombination experiments, Nieuwkoop demonstrated the induction of endodermal cell lineages in animal cap cells that had been in contact with vegetal regions (Nieuwkoop, 1997). Pharyngeal and dorsal endoderm were induced in the animal caps. It was suggested that the animal cap tissue contained two major layers, including an inner sensorial layer and an outer epithelial layer. The outer layer can be induced by the vegetal cells to form endoderm. Treatment of prospective ectodermal tissues with Activin or processed Vg1 induces expression of the XlHbox8 and IFABP marker genes, which are specifically expressed in the endodermal lineage (Wright et al., 1989; Gamer and Wright, 1995; Henry et al., 1996). Studies with a cleavage mutant of Vg1 have demonstrated that Vg1 is required for development of endoderm and dorsal mesoderm (Joseph and Melton, 1998). Treatment of prospective ectodermal tissues with bFGF does not induce XlHbox8 expression, although a functional FGF signaling pathway is required for endodermal differentiation. During the tailbud stage, XlHbox8, a homeodomain transcription factor, is expressed in cells that give rise to the pancreas. XlHbox8 is expressed in dorsal vegetal explants isolated from blastula embryos and is a marker of anterior endoderm. The response of XlHbox8 expression to lithium chloride treatment and UV irradiation is similar to what has been seen with organizer genes: Lithium chloride treatment increases XlHbox8 expression and UV irradiation abolishes XlHbox8 expression in whole embryos. IFABP, a cysteine-rich protein, is expressed in the epithelium of the small intestine. IFABP is a marker of general mesoderm since it is expressed in both dorsal and ventral vegetal explants. Endodermin (edd) has also been used as an endoderm differentiation marker (Sasai et al., 1996). In gastrula-stage embryos, edd expression is expressed in the endodermal mass and is also activated in precursor cells giving rise to the prechordal mesoderm and notochord and the superficial layer of the dorsal blastopore lip. By the tailbud stage, edd expression is restricted to the endoderm. edd encodes a novel member of the ␣2-macroglobulin protein family, a proposed function of which is to inactivate proteases. Both Mixer (Henry and Melton, 1998) and Xsox17␣ and ␤ (Hudson et al., 1997) have been shown to initiate endodermal differentiation in prospective ectoderm. Processed Vg1 induces expression of Mixer, which can activate the endodermal markers XlHbox8, IFABP, LFABP, and edd in prospective ectodermal tissues in the absence of mesoderm. The induction of endodermal marker genes by Vg1 is blocked by a Mixer-engrailed repressor fusion construct, Mixer-ENR, indicating that endoderm differentiation induced by the overexpression of Vg1 is mediated by Mixer. Whereas the mesodermal marker Xbra is induced in cells adjacent to Vg1-expressing cells, Mixer expression is activated in the same cells in which Vg1 is overexpressed. This is consistent with the idea that a high concentration of TGF␤ is required for endodermal development and a lower concentration of TGF-␤ is required for mesoderm induction. The HMG-box transcription factors Xsox17␣ and ␤ are specifically expressed in the endodermal mass at the early gastrula stage.

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Figure 5 The maternal transcription factor VegT is required for both endoderm and mesoderm formation. Maternal determinants, and to some extent VegT, can activate cell-autonomous expression of endoderm-specific genes and TGF-␤-related growth factors. The TGF-␤s activate further expression of endoderm-specific genes and reinforce the expression of TGF-␤s during endoderm formation in the vegetal region. TGF-␤s activated by maternal VegT are also required for mesoderm formation. VegT may activate its zygotic isoform to maintain TGF-␤ expression during mesoderm formation.

Overexpression of Xsox17␣ or ␤ in prospective ectoderm initiates endodermal differentiation by inducing the expression of XlHbox8 and IFABP. Expression of Mixer-ENR in vegetal explants inhibits the expression of Xsox17␣ and ␤, XlHbox8, and IFABP, but not edd. It appears that the induction of XlHbox8 and IFABP by Mixer is mediated through the action of Xsox17 and that Mixer regulates edd expression independently of Xsox17 by a different pathway. The vegetally localized maternal mRNA, xBic-C, has also been shown to lead to endoderm formation when over expressed in ectodermal explant (Wessely and De Robertis, 2000). This result demonstrate that endoderm formation in Xenopus embryos is also governed by maternal determinant. It has been suggested that endoderm formation takes place in two distinct steps involving maternal determinants and zygotically activated TGF-␤s (Yasuo and Lemaire, 1999). This is consistent with the demonstration that specification of the endodermal lineage occurs during the mid-blastula stage (Heasman et al., 1984). Some early genes that are expressed in the endoderm, such as the genes encoding the transcription factors Xsox17␣ and Mix.1 and the genes encoding the TGF-␤s Xnr-1, -2, and -4, Activin B, and Derriere, can be activated cell-autonomously by maternal determinants in dissociated embryonic cells (Clements et al., 1999; Yasuo and Lemaire, 1999). TGF-␤ signaling by zygotically expressed Xnr-1, Xnr-2, Derriere, and possibly other TGF-␤s subsequently activates the zygotic expression of Mixer and GATA-4 together with an upregulation of Xsox17␣, Mix.1, Xnr-1, and Xnr-2 expression (Fig. 5). VegT is a maternal determinant, one of its function is involved in endoderm differentiation (Lustig et al., 1996b; Stennard et al., 1996; Zhang and King, 1996; Horb and Thomsen, 1997). Maternal VegT mRNA is localized to the oocyte vegetal cortex, which is fated to give rise to the endodermal lineage. Endoderm differentiation is inhibited in embryos in which maternal VegT mRNA has been depleted

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by antisense oligonucleotides (Zhang et al., 1998a). These embryos express mesodermal and neural markers instead of endodermal markers in the vegetal region. Overexpression of VegT in prospective ectoderm activates early endodermal genes, including Xnr-2, Xsox17, and Mix.1, similar to the endogenous maternal determinants discussed previously (Yasuo and Lemaire, 1999). Activation of Xnr-1 expression may require other factors in addition to VegT (Yasuo and Lemaire, 1999). In zebrafish embryos, the molecular components of the TGF-␤ signaling pathway leading to endoderm formation is similar to that observed in Xenopus. During zebrafish endoderm development, the nodal-related growth factors Cyclops and Squint activate a type I TGF-␤ receptor TARAM-A in the presence of Oep activity (Renucci et al., 1996; Schier et al., 1997; Peyrieras et al., 1998; Zhang et al., 1998b). Activation of TARAM-A signaling induces Mixer expression, which acts through the casanova gene locus for the expression of sox17 (Alexander et al., 1999; Alexander and Stainier, 1999). Some of the molecular components involved in the formation of endoderm are therefore conserved between frogs and teleofish. This is a good example to demonstrate the advantages of studying different model systems for the elucidation of a signaling pathway. Ectopic expression of the organizer genes chordin and noggin also induce endoderm formation in prospective ectoderm (Sasai et al., 1996). Inhibition of BMP signaling by Chordin, Noggin, or a dominant negative BMP receptor can induce neural as well as endodermal gene expression. In prospective ectodermal explants, FGF treatment in addition to chordin or noggin overexpression induces neural markers with a posterior character. However, inhibition of FGF signaling in prospective ectodermal tissues overexpressing chordin or noggin shifts the induction from a neural toward an endodermal fate. In embryos, FGF signaling has been demonstrated in all three germ layers (LaBonne and Whitman, 1997). The endogenous requirement for FGF signaling in the induction of endoderm by chordin and noggin requires further analysis. It is conceivable that the organizer emits anti-BMP signals such as Chordin and Noggin to pattern the endodermal layer, similar to the processes of neural induction in the ectoderm and dorsalization in the mesoderm. Transcription factors such as Mix.1, milk, and Bix1-4 are involved in endoderm formation. Mix.1 (Rosa, 1989; Lemaire et al., 1998) and milk (Ecochard et al., 1998) are immediate-early genes expressed in response to activin treatment. The expression pattern and the function of milk are similar to those of Mix.1. Mix.1 is initially expressed in the entire vegetal hemisphere. The expression domains of Mix.1 and Xbra become mutually exclusive during gastrulation. Ectopic expression of Mix.1 in the marginal zone downregulates Xbra expression in the mesoderm. Mix.1 induces edd expression in prospective ectodermal tissues, but only activates the endodermal markers XlHbox8 and IFABP when Mix.1 is coexpressed with siamois. edd is also activated in milk- or Mix.1-expressing cells, indicating a cellautonomous activation of edd expression by these transcription factors. With the use of a mutant form of Mix.1, namely m11, it has been found that Mix.1 mediates the ventralization effect of bmp-4 during mesoderm formation

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(Mead et al., 1996). However, a different study suggested that bmp-4 and Mix.1 are expressed in different domains and that wild-type Mix. 1 represses both dorsal and ventral mesoderm formation (Lemaire et al., 1998). These suggestions argue against the function of Mix .1 as a ventralizing agent. One suggested explanation is that the mutation introduced into m11 may have altered its specificity to affect functions of genes other than Mix.1. The Bix genes (Bix1-4) were identified by searching for Xbra target genes using a hormone-inducible glucocorticoid receptor fusion construct, Xbra-GR (Tada et al., 1998; Casey et al., 1999). The Bix proteins are similar to the Mix proteins, including Mix.1, Mix2, Mixer, and Milk. The Bix2 protein is identical to milk. The Bix genes are expressed at the onset of gastrulation in the mesoderm and endoderm, and their expression precedes that of Xbra. Bix1 is also an immediateearly gene in response to Activin, Xbra, and VegT. A low level of Bix1 expression in the prospective ectoderm induces ventral mesoderm formation, and a high level of Bix1 expression induces endoderm. The T-box transcription factors Xbra and VegT can activate the promoter sequence of the Bix1 gene. The promoter of the Bix4 gene also contains three T-box binding sites. Through the use of transgenic embryos, it has been demonstrated that two T-box binding sites of the Bix4 promoter are sufficient for mesodermal and endodermal expression. Expression of Bix4 requires maternal VegT. In VegT-depleted embryos, expression of Bix4 rescues endodermal markers but not mesodermal markers or mesoderm-inducing activity, suggesting a role of Bix4 in endoderm formation.

B. Mesoderm The blastomeres from tier B and tier C of a 32-cell embryo contribute to the prospective mesoderm lineage, which is further patterned into dorsal, intermediate, and ventral mesodermal derivatives (Dale and Slack, 1987a). Fate mapping studies show that cells from the dorsal marginal zone contribute to dorsal mesoderm, including head mesoderm, notochord, and somites, whereas cells from the ventral marginal zone form somites and blood cells. Specification studies show that the lateral marginal zone gives rise to intermediate mesoderm, such as somites and pronephros, only as a result of interaction between the dorsal and ventral mesoderm, a process known as dorsalization (Dale and Slack, 1987b). By combining vegetal and animal regions from embryos at different developmental stages, it has been determined that the vegetal region emits mesoderminducing signals between stage 6 and stage 10.5 (Jones and Woodland, 1987). The responsiveness of the animal cap region to the mesoderm-inducing signal has a similar timing. This indicates that mesoderm induction can be initiated as early as the early cleavage stage before the MBT. By combining vegetal masses from early and late blastula embryos with prospective ectodermal tissues, it has been shown that post-MBT vegetal masses have a much stronger induction potential for both

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the mesoderm-inducing and endoderm-inducing activities than those derived from pre-MBT embryos (Wylie et al., 1996; Yasuo and Lemaire, 1999). The increase in inducing in activity of the vegetal mass is probably due to the zygotic activation of TGF-␤ molecules. These molecules are likely to produce amplified inducing signals after the MBT to reinforce mesoderm formation that may have begun before the MBT. Activins can induce the formation of different mesodermal derivatives in prospective ectodermal tissues, including both dorsal and ventral mesoderm, in a dose-dependent fashion (Smith, 1995). It has been suggested that a morphogen gradient is involved in the specification of mesodermal fates in developing embryos (Neumann and Cohen, 1997; Gurdon et al., 1998). Two different mechanisms have been suggested for the setting up of a TGF-␤ morphogen gradient in Xenopus blastula-stage embryos: sequential short-range signaling by a relay mechanism and long-range signaling by passive diffusion. In the relay model, secondary inducing signals are involved in the propagation of the TGF-␤1 signal across cell boundaries (Reilly and Melton, 1996). In the diffusion model, the Activin protein diffuses among cells and directly activates gene expression in a concentrationdependent manner (Gurdon et al., 1994, 1996). This long-range diffusion is a unique property of Activin and is not observed with other TGF-␤s such as Xnr-2 and BMP4, which evoke mesoderm gene expression in a cell-autonomous manner. Protein secretion and processing and the extracellular matrix are constraints affecting the diffusion of different TGF-␤ molecules (Jones et al., 1996a). This finding indicates that different mechanisms are involved in controlling the distribution of TGF-␤ molecules within embryonic tissues. Overexpression of constitutively active activin type I receptors activates cell-autonomous gene expression, again arguing against the relay mechanism and supporting a direct action of Activin in the responding cells (Jones et al., 1996a). It has also been shown that responding cells activate expression of different mesoderm genes, according to their position in the morphogen gradient, by sensing the absolute number of occupied receptors, but not the relative number of occupied versus unoccupied receptors (Dyson and Gurdon, 1998). Furthermore, cell– cell interaction is not necessary for cultured blastula cells to respond to the activin morphogen gradient in a concentrationdependent way (Gurdon et al., 1999). This finding suggests that individual blastula cells can respond to Activin, arguing against the requirement for interaction between neighboring cells to refine the response to a morphogen gradient. Whereas Activin is required for the activation of mesodermal gene expression, FGF is required for maintaining the expression of these genes (LaBonne and Whitman, 1994). FGFs induce the formation of mesodermal derivatives of ventral characters in prospective ectodermal tissues (Smith, 1995). FGF induces Xbra expression through the activation of a MAP kinase pathway (LaBonne et al., 1995; Umbhauer et al., 1995). A substantial increase in MAP kinase activity has been demonstrated in dissected embryos (LaBonne and Whitman, 1997). This suggests that wounding introduced by dissection may cause the release of growth factors and a subsequent activation of MAP kinase signaling. It should be noted that such

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artefactual activation of MAP kinase can be circumvented by performing dissection at a low temperature (LaBonne and Whitman, 1997). Overexpression of dominant negative receptor constructs for activin and FGF results in embryos that have no mesoderm and embryos that have mesodermal defects, respectively (Amaya et al., 1991, 1993; Hemmati-Brivanlou and Melton, 1992). These phenotypes suggest a role of TGF-␤ and FGF in mesoderm formation. Studies with Vg1 mutant have demonstrated the role of Vg1 in the formation of dorsal mesoderm (Joseph and Melton, 1998). The maternal store of Vg1 mRNA and Vg1 protein, the potent mesoderm-inducing activity of processed Vg1, and the requirement of Vg1 for dorsal mesoderm formation are all consistent with a role for Vg1 as a maternal TGF-␤ required for mesoderm induction. Members of the Nodal-related family, a class of proteins related to TGF-␤, are zygotically expressed in the marginal zone. Xnr-1, Xnr-2, and Xnr-4 are expressed in the marginal zone, with higher concentration in the dorsal mesoderm (Jones et al., 1995; Joseph and Melton, 1997). Overexpression of Xnr-1 and Xnr-2 induces both dorsal and ventral mesoderm in prospective ectodermal tissues and, like that of organizer genes such as noggin, chordin, and siamois, can dorsalize tissue explants of ventral marginal zone. A number of T-box transcription factors are involved in mesoderm formation (Stennard et al., 1997). Xbra is the prototype of the Xenopus T-box family, which also includes VegT and Eomesodermin (Eomes) (Smith et al., 1991; Ryan et al., 1996). Overexpression of Xbra or different FGFs induce ventral mesodermal cell types in prospective ectoderm. Although basic FGF (bFGF) has been used as an inducing source for animal cap assays, the endogenous source of FGF is likely to be eFGF (Isaacs et al., 1994). Embryonic FGF (eFGF), but not bFGF, is secreted and is expressed in the equatorial region of gastrula-stage embryos. eFGF and Xbra can activate the expression of each other in an autoregulatory loop. Xbra expression is suppressed in dissociated embryos, indicating that Xbra expression is non-cell-autonomous (Lemaire and Gurdon, 1994). The promoter of Xbra2, a pseudoallele of Xbra, is regulated by FGF and Activin (Latinkic et al., 1997). The Xbra promoter is also subject to regulation by gsc in response to different levels of Activin. A high level of Activin activates gsc expression, which in turn suppresses Xbra expression. A low level of activin activates Xbra but not gsc expression. The demarcation between the endoderm and mesoderm regions may be regulated in a similar manner, such that a high level of endogenous TGF-␤ induces Mix.1 expression, which represses Xbra expression in the endoderm. Studies involving the depletion of maternal VegT transcripts has demonstrated a role of VegT for the establishment of the germ layers in Xenopus embryos. An antisense oligonucleotide knockout study showed that maternal VegT mRNA is required for the formation of endoderm (Zhang et al., 1998a). With an increased efficiency of depletion by using an increased dose of antisense oligonucleotides and HPLC-purification of the oligonucleotides, maternal VegT has been demonstrated to be required for formation of 90% of the mesodermal tissues (Kofron et al., 1999). In VegT-depleted embryos, the expression of TGF-␤ factors, including Xnr-1,-2,

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and -4 and Derriere, is not detected. Overexpression of the same TGF-␤s in VegTdepleted embryos rescues mesoderm and body axis formation with the use of Cer-S, which can antagonize Xnr factors, the existence of an endogenous gradient of Xnrs in the endoderm has been demonstrated (Agius et al., 2000). Both studies demonstrated that TGF-␤s such as Xnr-1, -2, and -4 and derriere, which are activated by the maternal transcription factor VegT, are involved in mesoderm development. An isoform of maternal VegT is zygotically expressed not in the endoderm, but in the equatorial region. This isoform contains a different N-terminal region as a result of alternative splicing (Stennard et al., 1999). Unlike the maternal VegT isoform, expression of the zygotic VegT isoform can be activated by Activin and is non-cell-autonomous in dissociated embryos. In contrast, expression of the maternal VegT isoform is not responsive to Activin treatment. Overexpression of zygotic VegT induces both dorsal and ventral mesoderm in prospective ectodermal tissues in a dose-dependent manner. In whole embryos, overexpression of zygotic VegT induces ectopic dorsal lip formation and morphogenetic cell movement (Lustig et al., 1996b). Zygotic VegT is first expressed above the dorsal lip region; the expression domain subsequently extends laterally and ventrally in the marginal zone and marks the lateral and ventral mesoderm similar to the expression pattern of Xbra. Overexpression of zygotic VegT also upregulates Xbra expression in prospective ectodermal tissues. The expression of Xbra and zygotic VegT overlaps in the marginal zone, but not in the notochord, in which only Xbra is expressed. Another T-box gene, eomes, has an expression pattern similar to that of zygotic VegT in the early gastrula (Ryan et al., 1996). eomes is an early T-box gene expressed at the MBT. Expression of eomes in prospective ectodermal tissues can be induced by growth factors, including Activin and BMP-4, but not eFGF or Xwnt-8. Like overexpression of zygotic VegT, eomes overexpression can induce expression of dorsal and ventral mesodermal marker genes. A dominant-interfering construct containing eomes and the engrailed repressor domain produces a gastrulation defect that can be rescued by wild-type eomes. Zygotically expressed VegT, eFGF, and derriere, a novel member of the TGF␤-like family, regulate the expression of each other (Sun et al., 1999). derriere is expressed in the prospective endoderm and mesoderm during the blastula stage and is later restricted to the posterior mesoderm. The expression pattern of derriere is similar to that of zygotic VegT. derriere activates mesodermal markers of both dorsal and ventral character, as well as some ectodermal, neural, and endodermal marker genes. derriere can also activate zygotic VegT and eFGF expression. The expression of derriere can be induced by growth factors, including Activin, BVg1, bFGF, Derriere itself, and the transcription factors encoded by T-box genes such as Xbra and zygotic VegT. Ectopic expression of wild-type derriere in embryos induces a partial axis on the ventral side or microcephaly on the dorsal side, similar to the phenotypes produced by overexpression of the zygotic VegT isoform. Expression of a cleavage mutant of Derriere results in embryos with posterior

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truncation missing somites and tail. This mutant phenotype is distinct from the dorsoanterior defects produced by the Vg1 mutant and the activin cleavage mutant, thus demonstrating the requirement of different TGF-␤s in both dorsal and ventral mesoderm development. The similarity of the Derriere cleavage mutant phenotype to overexpression of dominant negative forms of FGFR, Xbra, and VegT further confirms the regulation between Derriere, FGF, and VegT. When does mesoderm induction occur? Tissue recombination assays have demonstrated an early mesoderm-inducing signal during cleavage stage. However, it has also been shown that a mesoderm-inducing signal is zygotically activated. Can these data be reconciled with each other? On the basis of the available information, it is possible that mesoderm induction occurs in two phases, one before and one after the MBT. During the early cleavage stage, TGF-␤ signaling is initiated, albeit at a low level. At the MBT, maternal VegT activates expression of transcriptional targets, such as Xnr-1, Xnr-2, and derriere, to amplify and reinforce TGF-␤ signaling in the prospective endoderm to initiate endodermal differentiation. In addition, these factors signal to cells in the marginal zone to induce the expression of the zygotic isoform of VegT, and other T-box transcription factors, such as Xbra and Eomes. Zygotic VegT specifies mesoderm formation by further activating the transcription of TGF-␤s and other T-box members in the marginal zone. For example, zygotic VegT can activate derriere, which is able to induce FGF expression. Zygotic VegT is also able to induce the expression of eomes and Xbra. In this model, maternal determinants and zygotic TGF-␤s are both required for mesoderm induction. An intricate regulation cascade between TGF-␤ and the T-box transcription factors therefore functions in the vegetal and marginal zones to bring about mesoderm formation. In addition to TGF-␤s and FGF, bmp overexpression induces ventral mesoderm formation in prospective ectodermal tissues, including blood and mesenchyme (Dale and Jones, 1999). It has been suggested that only heterodimers between BMP-2 or BMP -4 and BMP-7, but not homodimers, are potent inducers of ventral mesoderm (Nishimatsu and Thomsen, 1998). Homodimers are only able to ventralize mesoderm that has already been induced by activin. Inhibition of homoand heterodimers of BMPs by the BMP antagonists noggin and gremlin in the ventral marginal region shows an activation of organizer genes, but the expression of panmesodermal markers is unaffected (Eimon and Harland, 1999). This result suggests that BMP signaling is required for the patterning of mesoderm rather than primary mesoderm induction. A high level of BMP in the ventral marginal zone induces blood, the most ventral mesoderm, and the expression of both Xvent-1 (Gawantka et al., 1995) and Vox (also known as Xvent-2, Xom, and PV.1) (Ault et al., 1996; Ladher et al., 1996; Onichtchouk et al., 1996; Schmidt et al., 1996). A lower level of BMP in the lateral marginal zone leads to muscle formation and only induces the expression of Vox (Onichtchouk et al., 1998). Vox is expressed in the marginal zone and animal cap region excluded from the organizer and is later excluded from the

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notochord and neural plate. The expression pattern of Vox is complementary to that of chordin and Xnot (Schmidt et al., 1996). Studies using a dominant-interfering construct Vox have shown that Vox is a transcriptional repressor for a subset of organizer genes, including chordin, gsc, Xotx2, XFD’-1, and Xnr-1, but not noggin (Melby et al., 1999). Since gsc overexpression represses Vox (Ferreiro et al., 1998), the finding that Vox represses chordin explains the observation that gsc activates chordin expression. The activation of chordin by gsc is therefore indirect and is mediated by a suppression of the inhibitory effect of Vox. This demonstrates a repression of chordin by bmp-4 at the transcriptional level. The regulation between chordin and bmp-4 is not limited to protein–protein interaction. Xnf7 is a maternally expressed transcription factor that shows a differential cytoplasmic localization before and after the MBT (El-Hodiri et al., 1997). Overexpression of an engrailed-xnf7 fusion construct has been shown to induce expression of the BMP-4 antagonist chordin and the formation of incomplete secondary axes. This indicates the possible involvement of a maternal transcription factor for the regulation of BMP4 signaling in axial patterning (H. El-Hodiri and L. D. Etkin, unpublished observations). Members of the EGF-CFC family may be required for Nodal signaling. One of the family members, FRL-1, induces mesoderm and neural differentiation in prospective ectoderm (Kinoshita et al., 1995). Another family member, Cryptic, is expressed in mesoderm and midline neuroectoderm and is thought to be required for mesoderm and neural patterning (Shen et al., 1997). Similar to the case with the BMPs, Wnts are also involved in mesoderm development. However, Wnts are not mesoderm inducers, because they do not induce mesoderm formation in prospective ectoderm. Xwnt-8 is expressed in the ventral lateral marginal zone in gastrula-stage embryos. Wnt antagonists such as cer, dkk-1, and frzb are expressed in the dorsal mesoderm, whereas sizzled is expressed in the ventral mesoderm. frzb and sizzled contain the extracellular region of the Wnt receptor frizzled family. Wnt signaling is involved in patterning the mesodermal layer. Ectopic expression of Xwnt-8 after the MBT in the dorsal marginal zone exerts a ventralizing effect and increases the amount of somitic tissues formed at the expense of the notochord (Christian and Moon, 1993; Hoppler et al., 1996). In contrast, suppression of Wnt-signaling with a dominant negative Xwnt-8 construct or the Wnt antagonist frzb inhibits ventral gene expression (Hoppler et al., 1996; Leyns et al., 1997; Wang et al., 1997). It has also been suggested that Xwnt-8 is required to pattern and sharpen the boundary between notochord and somites in the dorsal and dorsolateral marginal zone (Hoppler and Moon, 1998). In the ventral marginal zone, the Wnt antagonist sizzled, which is activated by BMP-4, may restrict Wnt signaling to the lateral region (Salic et al., 1997; Marom et al., 1999). In the marginal zone, Wnt signaling is therefore eliminated in the dorsal and ventral mesoderm by different Wnt antagonists. As a consequence, Wnt signaling is active only in the lateral marginal region to specify formation of lateral mesodermal derivatives.

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C. Ectoderm Blastomeres from tier A of a 32-cell embryo give rise to the prospective ectoderm, which differentiates into neural and nonneural ectodermal lineages (Chang and Hemmati-Brivanlou, 1998). At the gastrula stage, the dorsal ectoderm in proximity to the organizer develops into the neural plate, and the ventral ectoderm gives rise to the epidermis. The cement gland, various placodes, and neural crest are formed at the boundary between the neural plate and the epidermal region. By embryonic cell dissociation experiments, it has been shown that BMP signaling mediates the decision between neural and epidermal cell fates (Wilson and Hemmati-Brivanlou, 1995). Whereas intact ectodermal explants develop into atypical epidermis and express epidermal keratin, dissociated ectodermal cells follow a neural fate and express neural markers. Epidermal cell fate can be rescued in dissociated animal cap cells by treatment with BMP-4 but not activin. BMP signaling is therefore required to repress neural development in the ectoderm. A number of organizer genes, including chordin, noggin, follistatin, Xnr-3, cer, and dkk-1, can induce prospective ectoderm to adopt a neural character by exerting a dorsalizing activity. Chordin, Noggin, Follistatin, Xnr-3, and Cer possess anti-BMP activities, whereas Cer and Dkk-1 have anti-Wnt activities. Therefore, a suppression of the BMP or Wnt pathway results in induction of a neural fate in the ectoderm. FGF signaling has been suggested to be involved in neural induction and anterior neural patterning. Studies using a dominant negative FGF receptor provide evidence that FGF signaling is not required for the early event of neural induction but is involved in the posterior development of the mesoderm and neuroectoderm layers (Kroll and Amaya, 1996; Holowacz and Sokol, 1999). However, in a different study, it has also been demonstrated that FGF signaling is involved in anterior neural induction (Hongo et al., 1999). Chordin and Noggin can both interact with BMPs physically and inhibit binding of BMPs to the receptor, albeit with different affinities (Piccolo et al., 1996; Zimmerman et al., 1996). Follistatin inhibits BMP function through a mechanism different from that of Noggin and Chordin because the Follistatin/BMP-4 complex is able to bind to BMP receptors (lemura et al., 1998). It has been suggested that follistatin may direct specific degradation of BMPs by mediating binding to receptors. Xnr-3 induces neural marker expression in prospective ectoderm (Hansen et al., 1997). Unlike Xnr-1, -2, and -4, Xnr-3 has no mesoderm-inducing activity, but can dorsalize ventral mesoderm. The activation of neural genes by Xnr-3 can be inhibited by overexpression of BMP-4 or an activated form of the BMP receptor. This suggests that Xnr-3 also inhibits BMP activity during neural induction. The mechanism by which Xnr-3 inhibits BMP functions is unclear. It has been proposed that Xnr-3 may dimerize with BMP-4 to produce a nonfunctional complex or act as a BMP receptor antagonist that competes with BMP for binding to the receptor. Cer also has neural-inducing activity in prospective ectoderm (Bouwmeester et al., 1996). Since the neuralizing effect of Cer can be blocked by overexpression of

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BMP-4, Cer also mediates its neurogenic effect by inhibiting BMP signaling. The demonstration of a BMP-4-binding site in the N terminus of the Cer protein further substantiates a physical interaction between Cer and BMPs. Xenopus brain factor 2 (XBF-2) has been shown to be a common downstream target of noggin and cer (Mariani and Harland, 1998). Expression of noggin or cer in prospective ectoderm induces XBF-2 expression, which acts as a transcriptional repressor that inhibits BMP-4 transcription. Thus, noggin and cer play a role in neural induction by antagonizing BMP signaling at both the protein level and the transcriptional level. The control of neural and nonneural cell fate provides the basis for neural and ectodermal differentiation. However, there is not yet any evidence for the diffusion of any of the secreted growth factors expressed in the organizer. It appears that additional positional information is required to further pattern the ectoderm. It has been proposed that different levels of BMP generate a morphogen gradient across the dorsal–ventral axis to specify different cell fates in the ectoderm (Dale and Wardle, 1999). A dose-dependent effect of BMP-4 overexpression in inducing neural and epidermal gene expression in dissociated animal cap cells has been observed (Wilson et al., 1997). A low level of BMP-4 induces neural marker expression, and intermediate and high doses activate cement gland and epidermal keratin expression, respectively. A similar dose-dependent effect has been observed with a dominant-negative BMP receptor, tBR, and the signal transducer of BMP signaling Smad1. It is not clear how such a gradient is established in embryos. The msx1 gene is a target of BMP-4 that can mediate epidermal induction (Suzuki et al., 1997b). Msx1 is a homeobox transcription factor and is an immediate early gene expressed in response to BMP-4 induction. Expression of msx1 is detected in ventral ectoderm and mesoderm. In whole embryos, overexpression of msx1 causes ventralization. Overexpression of msx1 rescues epidermal differentiation in dissociated ectoderm, which would otherwise follow a neural fate. It has also been shown that the Xmsx2 gene is involved in the anterior–posterior patterning of dorsal mesoderm in Xenopus embryos (Gong and Kiba, 1999). The induction of neural marker expression by the Wnt antagonist, dkk-1, suggested a possible involvement of Wnt signaling in the regulation of ectoderm differentiation (Glinka et al., 1998). It has been suggested that a Wnt signal may be required during the cleavage stage to suppress BMP signaling in the dorsal region of the embryo during neural induction. The dorsal ectoderm is consequently sensitized to respond to the neuralizing signals emanating from the organizer (Baker et al., 1999). The neural-inducing organizer genes display different expression domains during gastrulation. At mid-gastrula stage, cer is expressed in a broad area of the anterior endomesoderm, including the leading edge mesoderm. A gap of cer expression is observed along the prechordal plate region, where chordin and noggin are expressed (Bouwmeester et al., 1996). These results show that different neural inducers occupy distinct positions within the mesodermal layer. Therefore, a combination of differentially expressed neural inducers and neural-inducing activity in the mesodermal layer, and a spatial restriction of epidermal inducers in

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the ectoderm, may play a role in the specification of neural and nonneural regions within the ectoderm.

D. A Theoretical Model of Germ Layer Formation On the basis of what is known about the different signaling pathways active during germ layer formation, a picture of this process is emerging. In this model, the specification of the three germ layers is governed by the TGF-␤/Nodal signaling pathway, whereas the BMP and Wnt pathways pattern the germ layers along the dorsal–ventral axis. Vegetal cells are under the influence of a high level of TGF-␤/Nodal signaling and differentiate into prospective endoderm. Cells in the marginal zone are subject to a moderate level of TGF-␤/Nodal signaling and are induced to form the prospective mesoderm. The animal region develops into the prospective ectoderm, since it receives little or no TGF-␤/Nodal signal from the vegetal cells. Although a VCC/␤-catenin pathway is required before the MBT in the dorsal region to set up the dorsal–ventral axis, Wnt signaling is inhibited in the dorsal marginal zone after the MBT by Wnt antagonists. In addition, Nodal and BMP signaling are inhibited in the dorsal marginal zone by growth factor antagonists. The dorsal marginal zone, free of Wnt, Nodal, and BMP signaling, gives rise to the head organizer. Outside the organizer region, Wnt and BMP signaling are actively required to pattern the marginal zone into lateral and ventral mesoderm. In the ectoderm, Wnt and BMP signaling are required for specification of neural versus epidermal differentiation. The dorsal ectoderm follows a neural cell fate in the absence of Wnt and BMP signaling, and the ventral ectoderm exhibits active Wnt and BMP signaling and undergoes epidermal differentiation.

VI. Developmental Pathways and Tumorigenesis The study of embryogenesis in different developmental systems has provided a basic knowledge of the signaling pathways controlling normal growth and differentiation. A deregulation, or improper activation, of components such as ligands, receptors, intracellular components, and target genes sometimes results in the development of cancer. Mutations in APC and ␤-catenin have been reported in colon tumorigenesis (Kinzler and Vogelstein, 1996). APC is a tumor suppressor gene mutated in familial colon cancer and most cases of sporadic colon cancers. The mutation in APC results in a truncated form of the protein that lacks the ability to maintain a low ␤-catenin level by degradation. The mutated forms of ␤-catenin escape regulation by other components, such as GSK-3 phosphorylation, leading to increased stability. An increased level of ␤-catenin triggers cellular transformation in colon cells. Frat1 is a homolog of GBP that can suppress the inhibitory effect of the GSK-3 complex and

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results in the accumulation of ␤-catenin (Yost et al., 1998). Activation of the Frat1 gene has been reported to contribute to T-cell lymphomas (Jonkers et al., 1997). Smad4 functions together with receptor Smads to transduce downstream signals originating from members of the TGF-␤ superfamily. The DPC4 gene is the human homolog of Smad4. DPC4 is deleted in the majority of pancreatic carcinomas (Hahn et al., 1996). Loss of function of DPC4 has been reported in both pancreatic and colorectal carcinomas. Members of the EGF-CFC protein family, required for Nodal signaling, have been shown to have transformation potential. Overexpression of cripto can transform mammalian cell lines and stimulate proliferation of breast cancer cell lines (Ciccodicola et al., 1989; Normanno et al., 1994). Finally, DAN, a gene that encodes a growth factor antagonist belonging to the Cerberus/DAN family, has been identified as a potential tumor suppressor gene (Ozaki and Sakiyama, 1993). DAN suppresses cell growth in nontransformed cells, and expression of DAN is downregulated in transformed fibroblasts. Deregulation of the Wnt or TGF-␤ pathways has also been reported in other carcinomas that have not been discussed earlier. The elucidation of the basic components of signaling pathways, mechanisms of regulation, and target gene activation during early development will provide a key to understanding the molecular basis of human cancer. The more we understand the regulatory mechanisms of cell growth and differentiation, the more likely it is that we will be able to unravel and control the causes of malignancy.

VII. Perspectives Xenopus embryos offer an efficient system in which to unravel gene function by the overexpression of RNA. Both gain-of-function and dominant-interference assays have provided valuable information regarding the functional roles and requirement of genes for embryonic axis determination and pattern formation. However, more emphasis on protein distribution is needed to further substantiate the endogenous role of gene products during early development. The depletion of maternal transcripts by antisense oligonucleotides has demonstrated the requirement of maternal gene products for early specification of cell lineages. Additional roles of maternal genes in controlling cell fate determination and early development are likely to be elucidated through the use of antisense oligonucleotides knockout experiments. The cloning and characterization of the promoter sequence of genes activated during early development will shed light on the genetic pathways controlling embryogenesis. In addition, these characterized promoter sequences will increase the selection of regulatory sequences for use in controlling the spatial and temporal expression of transgenes in transgenic Xenopus embryos. Although Xenopus laevis does not favor genetic manipulation, such techniques are under rapid development with the use of Xenopus tropicalis. This will certainly provide an additional advantage for the use of Xenopus as a developmental

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system. In addition, genetics analyses have also been well established in other developmental models which provide additional experimental systems. For example, the mouse has been used for knocking out gene functions by gene targeting to determine the requirements for gene products during development. In studying early embryogenesis, the use of a combination of different developmental models is likely to complement the deficiency of individual systems. The integration of unlimited information from different developmental systems will definitely provide perspectives in the elucidation of developmental gene functions and aid in the understanding of early embryonic development.

Acknowledgments APC and LDE thank Drs. Patrick Lemaire, Fiona Stennard, Aaron Zorn, Malgozata Kloc, and Maki Wakamiya for helpful discussions, suggestions, and comments on the manuscript. Work from the author laboratory has been supported by grants from NSF, NIH and March of Dimes.

References Aberle, H., Bauer, A., Stappert, J., Kispert, A., and Kemler, R. (1997). beta-catenin is a target for the ubiquitin-proteasome pathway. EMBO J. 16, 3797–3804. Agius, E., Oelgeschlager, M., Wessely, O., Kemp, C., and De Robertis, E. M. (2000). Endodermal Nodal-related signals and mesoderm induction in Xenopus. Development 127, 1173–1183. Al-Mukhtar, K. A. K., and Webb, A. C. (1971). An ultrastructural study of primordial germ cells, oogonia and early oocytes in Xenopus laevis. J. Embryol. Exp. Morphol. 26, 195–217. Albano, R. M., Arkell, R., Beddington, R. S., and Smith, J. C. (1994). Expression of inhibin subunits and follistatin during postimplantation mouse development: decidual expression of activin and expression of follistatin in primitive streak, somites and hindbrain. Development 120, 803–813. Alexander, J., and Stainier, D. Y. (1999). A molecular pathway leading to endoderm formation in zebrafish. Curr. Biol. 9, 1147–1157. Alexander, J., Rothenberg, M., Henry, G. L., and Stainier, D. Y. (1999). Casanova plays an early and essential role in endoderm formation in zebrafish. Dev. Biol. 215, 343–357. Amaya, E., and Kroll, K. L. (1999). A method for generating transgenic frog embryos. Methods Mol. Biol. 97, 393–414. Amaya, E., Musci, T. J., and Kirschner, M. W. (1991). Expression of a dominant negative mutant of the FGF receptor disrupts mesoderm formation in Xenopus embryos. Cell 66, 257–270. Amaya, E., Stein, P. A., Musci, T. J., and Kirschner, M. W. (1993). FGF signalling in the early specification of mesoderm in Xenopus. Development 118, 477–487. Amaya, E., Offield, M. F., and Grainger, R. M. (1998). Frog genetics: Xenopus tropicalis jumps into the future. Trends Genet. 14, 253–255. Ang, S. L., Jin, O., Rhinn, M., Daigle, N., Stevenson, L., and Rossant, J. (1996). A targeted mouse Otx2 mutation leads to severe defects in gastrulation and formation of axial mesoderm and to deletion of rostral brain. Development 122, 243–252. Antczak, M., and Van Blerkom, J. (1997). Oocyte influences on early development: the regulatory proteins leptin and STAT3 are polarized in mouse and human oocytes and differentially distributed within the cells of the preimplantation stage embryo. Mol. Hum. Reprod. 3, 1067–1086.

50

Agnes P. Chan and Laurence D. Etkin

Artinger, M., Blitz, I., Inoue, K., Tran, U., and Cho, K. W. (1997). Interaction of goosecoid and brachyury in Xenopus mesoderm patterning. Mech. Dev. 65, 187–196. Ault, K. T., Dirksen, M. L., and Jamrich, M. (1996). A novel homeobox gene PV.1 mediates induction of ventral mesoderm in Xenopus embryos. Proc. Natl. Acad. Sci. USA 93, 6415–6420. Bachiller, D., Klingensmith, J., Kemp, C., Belo, J. A., Anderson, R. M., May, S. R., McMahon, J. A., McMahon, A. P., Harland, R. M., Rossant, J., and De Robertis, E. M. (2000). The organizer factors chordin and Noggin are required for mouse forebrain development. Nature 403, 658–661. Bachvarova, R. F. (1999). Establishment of anterior–posterior polarity in avian embryos. Curr. Opin. Genet. Dev. 9, 411–416. Baek, K. H., and Lee, K. Y. (1999). Signal transduction pathway for anterior–posterior development in Drosophila. J. Biomed. Sci. 6, 314–319. Baker, J. C., and Harland, R. M. (1996). A novel mesoderm inducer, Madr2, functions in the activin signal transduction pathway. Genes Dev. 10, 1880–1889. Baker, J. C., Beddington, R. S., and Harland, R. M. (1999). Wnt signaling in Xenopus embryos inhibits bmp4 expression and activates neural development. Genes Dev. 13, 3149–3159. Bauer, D. V., Huang, S., and Moody, S. A. (1994). The cleavage stage origin of Spemann’s Organizer: analysis of the movements of blastomere clones before and during gastrulation in Xenopus. Development 120, 1179–1189. Bauer, H., Meier, A., Hild, M., Stachel, S., Economides, A., Hazelett, D., Harland, R. M., and Hammerschmidt, M. (1998). Follistatin and noggin are excluded from the zebrafish organizer. Dev. Biol. 204, 488–507. Beddington, R. S., and Robertson, E. J. (1989). An assessment of the developmental potential of embryonic stem cells in the midgestation mouse embryo. Development 105, 733–737. Beddington, R. S., and Robertson, E. J. (1998). Anterior patterning in mouse. Trends Genet. 14, 277– 284. Beddington, R. S., and Robertson, E. J. (1999). Axis development and early asymmetry in mammals. Cell 96, 195–209. Belo, J. A., Bachiller, D., Agius, E., Kemp, C., Borges, A. C., Marques, S., Piccolo, S., and De Robertis, E. M. (2000). Cerberus-like is a secreted BMP and nodal antagonist not essential for mouse development. Genesis 26 (4), 265–270. Bhanot, P., Brink, M., Samos, C. H., Hsieh, J. C., Wang, Y., Macke, J. P., Andrew, D., Nathans, J., and Nusse, R. (1996). A new member of the frizzled family from Drosophila functions as a Wingless receptor. Nature 382, 225–230. Bienz, M. (1999). APC: the plot thickens. Curr. Opin. Genet. Dev. 9, 595–603. Blitz, I. L., and Cho, K. W. (1995). Anterior neuroectoderm is progressively induced during gastrulation: the role of the Xenopus homeobox gene orthodenticle. Development 121, 993–1004. Blum, M., Gaunt, S. J., Cho, K. W., Steinbeisser, H., Blumberg, B., Bittner, D., and De Robertis, E. M. (1992). Gastrulation in the mouse: the role of the homeobox gene goosecoid. Cell 69, 1097–1106. Blumberg, B., Wright, C. V., De Robertis, E. M., and Cho, K. W. (1991). Organizer-specific homeobox genes in Xenopus laevis embryos. Science 253, 194–196. Boterenbrood, E. C., and Nieuwkoop, P. D. (1973). The formation of the mesoderm in urodelean amphibians. V. Its regional induction by the endoderm. Wilhelm Roux’s Arch. 173, 319–332. Bouwmeester, T., and Leyns, L. (1997). Vertebrate head induction by anterior primitive endoderm. Bioessays 19, 855–863. Bouwmeester, T., Kim, S., Sasai, Y., Lu, B., and De Robertis, E. M. (1996). Cerberus is a head-inducing secreted factor expressed in the anterior endoderm of Spemann’s organizer. Nature 382, 595–601. Brannon, M., and Kimelman, D. (1996). Activation of Siamois by the Wnt pathway. Dev. Biol. 180, 344–347. Brannon, M., Gomperts, M., Sumoy, L., Moon, R. T., and Kimelman, D. (1997). A beta-catenin/XTcf-3 complex binds to the siamois promoter to regulate dorsal axis specification in Xenopus. Genes Dev. 11, 2359–2370.

1. Early Xenopus Development

51

Brannon, M., Brown, J. D., Bates, R., Kimelman, D., and Moon, R. T. (1999). XCtBP is a XTcf-3 co-repressor with roles throughout Xenopus development. Development 126, 3159–3170. Bronchain, O. J., Hartley, K. O., and Amaya, E. (1999). A gene trap approach in Xenopus. Curr. Biol. 9, 1195–1198. Broun, M., Sokol, S., and Bode, H. R. (1999). Cngsc, a homologue of goosecoid, participates in the patterning of the head, and is expressed in the organizer region of hydra. Development 126, 5245–5254. Brunet, L. J., McMahon, J. A., McMahon, A. P., and Harland, R. M. (1998). Noggin, cartilage morphogenesis, and joint formation in the mammalian skeleton. Science 280, 1455–1457. Cadigan, K. M., and Nusse, R. (1997). Wnt signaling: a common theme in animal development. Genes Dev. 11, 3286–3305. Carnac, G., Kodjabachian, L., Gurdon, J. B., and Lemaire, P. (1996). The homeobox gene Siamois is a target of the Wnt dorsalisation pathway and triggers organiser activity in the absence of mesoderm. Development 122, 3055–3065. Casey, E. S., Tada, M., Fairclough, L., Wylie, C. C., Heasman, J., and Smith, J. C. (1999). Bix4 is activated directly by VegT and mediates endoderm formation in Xenopus development. Development 126, 4193–4200. Cavallo, R. A., Cox, R. T., Moline, M. M., Roose, J., Polevoy, G. A., Clevers, H., Peifer, M., and Bejsovec, A. (1998). Drosophila Tcf and Groucho interact to repress Wingless signalling activity. Nature 395, 604–608. Chalmers, A. D., and Slack, J. M. (1998). Development of the gut in Xenopus laevis. Dev. Dyn. 212, 509–521. Chan, A. P., and Gurdon, J. B. (1996). Nuclear transplantation from stably transfected cultured cells of Xenopus. Int. J. Dev. Biol. 40, 441–451. Chan, A. P., Kloc, M., and Etkin, L. D. (1999). fatvg encodes a new localized RNA that uses a 25nucleotide element (FVLE1) to localize to the vegetal cortex of Xenopus oocytes. Development 126, 4943–4953. Chang, C., and Hemmati-Brivanlou, A. (1998). Cell fate determination in embryonic ectoderm. J. Neurobiol. 36, 128–151. Chen, X., Rubock, M. J., and Whitman, M. (1996a). A transcriptional partner for MAD proteins in TGF-beta signalling. Nature 383, 691–696. Chen, Y., Lebrun, J. J., and Vale, W. (1996b). Regulation of transforming growth factor beta- and activin-induced transcription by mammalian Mad proteins. Proc. Natl. Acad. Sci. USA 93, 12992– 12997. Chen, Y., Bhushan, A., and Vale, W. (1997). Smad8 mediates the signaling of the ALK-2. Proc. Natl. Acad. Sci. USA 94, 12938–12943. Cho, K. W., Blumberg, B., Steinbeisser, H., and De Robertis, E. M. (1991). Molecular nature of Spemann’s organizer: the role of the Xenopus homeobox gene goosecoid. Cell 67, 1111–1120. Choi, C. Y., Kim, Y. H., Kwon, H. J., and Kim, Y. (1999). The homeodomain protein NK-3 recruits Groucho and a histone deacetylase complex to repress transcription. J. Biol. Chem. 274, 33194– 33197. Christian, J. L., and Moon, R. T. (1993). Interactions between Xwnt-8 and Spemann organizer signaling pathways generate dorsoventral pattern in the embryonic mesoderm of Xenopus. Genes Dev. 7, 13– 28. Christian, J. L., and Nakayama, T. (1999). Can’t get no SMADisfaction: Smad proteins as positive and negative regulators of TGF-beta family signals. Bioessays 21, 382–390. Ciccodicola, A., Dono, R., Obici, S., Simeone, A., Zollo, M., and Persico, M. G. (1989). Molecular characterization of a gene of the ‘EGF family’ expressed in undifferentiated human NTERA2 teratocarcinoma cells. EMBO J. 8, 1987–1991. Clements, D., Friday, R. V., and Woodland, H. R. (1999). Mode of action of VegT in mesoderm and endoderm formation. Development 126, 4903–4911.

52

Agnes P. Chan and Laurence D. Etkin

Clevers, H., and van de Wetering, M. (1997). TCF/LEF factor earn their wings. Trends Genet. 13, 485–489. Coggins, L. W. (1973). An ultrastructural and radioautographic study of early oogenesis in the toad Xenopus laevis. J. Cell Sci. 12, 71–93. Conlon, F. L., Barth, K. S., and Robertson, E. J. (1991). A novel retrovirally induced embryonic lethal mutation in the mouse: assessment of the developmental fate of embryonic stem cells homozygous for the 413.d proviral integration. Development 111, 969–981. Conlon, F. L., Sedgwick, S. G., Weston, K. M., and Smith, J. C. (1996). Inhibition of Xbra transcription activation causes defects in mesodermal patterning and reveals autoregulation of Xbra in dorsal mesoderm. Development 122, 2427–2435. Connors, S. A., Trout, J., Ekker, M., and Mullins, M. C. (1999). The role of tolloid/mini fin in dorsoventral pattern formation of the zebrafish embryo. Development 126, 3119–3130. Crease, D. J., Dyson, S., and Gurdon, J. B. (1998). Cooperation between the activin and Wnt pathways in the spatial control of organizer gene expression. Proc. Natl. Acad. Sci. USA 95, 4398–4403. Cui, Y., Brown, J. D., Moon, R. T., and Christian, J. L. (1995). Xwnt-8b: a maternally expressed Xenopus Wnt gene with a potential role in establishing the dorsoventral axis. Development 121, 2177–2186. Cui, Y., Tian, Q., and Christian, J. L. (1996). Synergistic effects of Vg1 and Wnt signals in the specification of dorsal mesoderm and endoderm. Dev. Biol. 180, 22–34. Dale, L., and Jones, C. M. (1999). BMP signalling in early Xenopus development. Bioessays 21, 751–760. Dale, L., and Slack, J. M. (1987a). Fate map for the 32-cell stage of Xenopus laevis. Development 99, 527–551. Dale, L., and Slack, J. M. (1987b). Regional specification within the mesoderm of early embryos of Xenopus laevis. Development 100, 279–295. Dale, L., and Wardle, F. C. (1999). A gradient of BMP activity specifies dorsalventral fates in early Xenopus embryos. Semin. Cell Dev. Biol. 10, 319–326. Dale, L., Matthews, G., and Colman, A. (1993). Secretion and mesoderm-inducing activity of the TGF-beta-related domain of Xenopus Vg1. EMBO J. 12, 4471–4480. Darras, S., Marikawa, Y., Elinson, R. P., and Lemaire, P. (1997). Animal and vegetal pole cells of early Xenopus embryos respond differently to maternal dorsal determinants: implications for the patterning of the organiser. Development 124, 4275–4286. de Souza, F. S. J., Gawantka, V., Gomez A. P., Delius, H., Ang. S. L., and Niehrs, C. (1999). The zinc finger gene Xblimp1 controls anterior endomesodermal cell fate in Spemann’s organizer. EMBO J. 18, 6062–6072. Deardorff, M. A., Tan, C., Conrad, L. J., and Klein, P. S. (1998). Frizzled-8 is expressed in the Spemann organizer and plays a role in early morphogenesis. Development 125, 2687–2700. DeMarais, A. A., and Moon, R. T. (1992). The armadillo homologs beta-catenin and plakoglobin are differentially expressed during early development of Xenopus laevis. Dev. Biol. 153, 337–346. Deno, T., Nishida, H., and Satoh, N. (1984). Autonomous muscle cell differentiation in partial ascidian embryos according to the newly verified cell lineages. Dev. Biol. 104, 322–328. Dominguez, I., and Green, J. B. (2000). Dorsal downregulation of GSK3␤ by a non-Wnt-like mechanism is an early molecular consequence of cortical rotation in early Xenopus embryos. Development 127, 861–868. Dominguez, I., Itoh, K., and Sokol, S. Y. (1995). Role of glycogen synthase kinase 3 beta as a negative regulator of dorsoventral axis formation in Xenopus embryos. Proc. Natl. Acad. Sci. USA 92, 8498– 8502. Dosch, R., and Niehrs, C. (2000). Requirement for anti-dorsalizing morphogenetic protein in organizer patterning. Mech. Dev. 90, 195–203. Du, S. J., Purcell, S. M., Christian, J. L., McGrew, L. L., and Moon, R. T. (1995). Identification of distinct classes and functional domains of Wnts through expression of wild-type and chimeric proteins in Xenopus embryos. Mol. Cell Biol. 15, 2625–2634.

1. Early Xenopus Development

53

Dumermuth, E., Sterchi, E. E., Jiang, W. P., Wolz, R. L., Bond, J. S., Flannery, A. V., and Beynon, R. J. (1991). The astacin family of metalloendopeptidases. J. Biol. Chem. 266, 21381–21385. Dyson, S., and Gurdon, J. B. (1997). Activin signalling has a necessary function in Xenopus early development. Curr. Biol. 7, 81–84. Dyson, S., and Gurdon, J. B. (1998). The interpretation of position in a morphogen gradient as revealed by occupancy of activin receptors. Cell 93, 557–568. Eastman, Q., and Grosschedl, R. (1999). Regulation of LEF-1/TCF transcription factors by Wnt and other signals. Curr. Opin. Cell Biol. 11, 233–240. Ecochard, V., Cayrol, C., Rey, S., Foulquier, F., Caillol, D., Lemaire, P., and Duprat, A. M. (1998). A novel Xenopus mix-like gene milk involved in the control of the endomesodermal fates. Development 125, 2577–2585. Eimon, P. M., and Harland, R. M. (1999). In Xenopus embryos, BMP heterodimers are not required for mesoderm induction, but BMP activity is necessary for dorsal/ventral patterning. Dev. Biol. 216, 29–40. El-Hodiri, H. M., Shou, W., and Etkin, L. D. (1997). xnf7 functions in dorsal–ventral patterning of the Xenopus embryo. Dev. Biol. 190, 1–17. Elinson, R. P., and Pasceri, P. (1989). Two UV-sensitive targets in dorsoanterior specification of frog embryos. Development 106, 511–518. Elinson, R. P., and Rowning, B. (1988). A transient array of parallel microtubules in frog eggs: potential tracks for a cytoplasmic rotation that specifies the dorsoventral axis. Dev. Biol. 128, 185–197. Etkin, L. D., and Pearman, B. (1987). Distribution, expression and germ line transmission of exogenous DNA sequences following microinjection into Xenopus laevis eggs. Development 99, 15–23. Evsikov, S. V., Morozova, L. M., and Solomko, A. P. (1994). Role of ooplasmic segregation in mammalian development. Roux. Arch. Dev. Biol. 203, 199–204. Fagotto, F., Guger, K., and Gumbiner, B. M. (1997). Induction of the primary dorsalizing center in Xenopus by the Wnt/GSK/beta-catenin signaling pathway, but not by Vg1, Activin or Noggin. Development 124, 453–460. Fagotto, F., Gluck, U., and Gumbiner, B. M. (1998). Nuclear localization signal-independent and importin/karyopherin- independent nuclear import of beta-catenin. Curr. Biol. 8, 181–190. Fainsod, A., Deissler, K., Yelin, R., Marom, K., Epstein, M., Pillemer, G., Steinbeisser, H., and Blum, M. (1997). The dorsalizing and neural inducing gene follistatin is an antagonist of BMP-4. Mech. Dev. 63, 39–50. Fan, M. J., Gruning, W., Walz, G., and Sokol, S. Y. (1998). Wnt signaling and transcriptional control of Siamois in Xenopus embryos. Proc. Natl. Acad. Sci. USA 95, 5626–5631. Fanning, A. S., and Anderson, J. M. (1999). Protein modules as organizers of membrane structure. Curr. Opin. Cell Biol. 11, 432–439. Fekany, K., Yamanaka, Y., Leung, T., Sirotkin, H. I., Topczewski, J., Gates, M. A., Hibi, M., Renucci, A., Stemple, D., Redbill, A., Schier, A. F., Driever, W., Hirano, T., Talbot, W. S., and Solnica-Krezel, L. (1999). The zebrafish bozozok locus encodes Dharma, a homeodomain protein essential for induction of gastrula organizer and dorsoanterior embryonic structures. Development 126, 1427– 1438. Feldman, B., Gates, M. A., Egan, E. S., Dougan, S. T., Rennebeck, G., Sirotkin, H. I., Schier, A. F., and Talbot, W. S. (1998). Zebrafish organizer development and germlayer formation require nodal-related signals. Nature 395, 181–185. Ferguson, E. L., and Anderson, K. V. (1992). Localized enhancement and repression of the activity of the TGF-beta family member, decapentaplegic, is necessary for dorsal-ventral pattern formation in the Drosophila embryo. Development 114, 583–597. Ferreiro, B., Artinger, M., Cho, K., and Niehrs, C. (1998). Antimorphic goosecoids. Development 125, 1347–1359. Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811.

54

Agnes P. Chan and Laurence D. Etkin

Forbes, A., and Lehmann, R. (1998). Nanos and Pumilio have critical roles in the development and function of Drosophila germline stem cells. Development 125, 679–690. Forristall, C., Pondel, M., Chen, L., and King, M. L. (1995). Patterns of localization and cytoskeletal association of two vegetally localized RNAs, Vg1 and Xcat-2. Development 121, 201–208. Fu, Y., Wang, Y., and Evans, S. M. (1998). Viral sequences enable efficient and tissue-specific expression of transgenes in Xenopus. Nat. Biotechnol. 16, 253–257. Fujisue, M., Kobayakawa, Y., and Yamana, K. (1993). Occurrence of dorsal axis–inducing activity around the vegetal pole of an uncleaved Xenopus egg and displacement to the equatorial region by cortical rotation. Development 118, 163–170. Fukui, A., Nakamura, T., Uchiyama, H., Sugino, K., Sugino, H., and Asashima, M. (1994). Identification of Activins A, AB, and B and Follistatin proteins in Xenopus embryos. Dev. Biol. 163, 279–281. Funayama, N., Fagotto, F., McCrea, P., and Gumbiner, B. M. (1995). Embryonic axis induction by the armadillo repeat domain of beta-catenin: evidence for intracellular signaling. J. Cell Biol. 128, 959–968. Furthauer, M., Thisse, B., and Thisse, C. (1999). Three different noggin genes antagonize the activity of bone morphogenetic proteins in the zebrafish embryo. Dev. Biol. 214, 181–196. Galceran, J., Farinas, I., Depew, M. J., Clevers, H., and Grosschedl, R. (1999). Wnt3a−/−-like phenotype and limb deficiency in Lef1(−/−) Tcf1(−/−) mice. Genes Dev. 13, 709–717. Gallagher, B. C., Hainski, A. M., and Moody, S. A. (1991). Autonomous differentiation of dorsal axial structures from an animal cap cleavage stage blastomere in Xenopus. Development 112, 1103–1114. Gamer, L. W., and Wright, C. V. (1995). Autonomous endodermal determination in Xenopus: regulation of expression of the pancreatic gene XlHbox 8. Dev. Biol. 171, 240–251. Gardner, R. L. (1997). The early blastocyst is bilaterally symmetrical and its axis of symmetry is aligned with the animal–vegetal axis of the zygote in the mouse. Development 124, 289–301. Gardner, R. L. (1998). Axial relationships between egg and embryo in the mouse. Curr. Top. Dev. Biol. 39, 35–71. Gardner, R. L. (1999). Scrambled or bisected mouse eggs and the basis of patterning in mammals. Bioessays 21, 271–274. Gawantka, V., Delius, H., Hirschfeld, K., Blumenstock, C., and Niehrs, C. (1995). Antagonizing the Spemann organizer: role of the homeobox gene Xvent-1. EMBO J. 14, 6268–6279. Gerhart, J., Danilchik, M., Doniach, T., Roberts, S., Rowning, B., and Stewart, R. (1989). Cortical rotation of the Xenopus egg: consequences for the anteroposterior pattern of embryonic dorsal development. Development 107, 37–51. Gimlich, R. L. (1986). Acquisition of developmental autonomy in the equatorial region of the Xenopus embryo. Dev. Biol. 115, 340–352. Gimlich, R. L., and Gerhart, J. C. (1984). Early cellular interactions promote embryonic axis formation in Xenopus laevis. Dev. Biol. 104, 117–130. Glinka, A., Wu, W., Delius, H., Monaghan, A. P., Blumenstock, C., and Niehrs, C. (1998). Dickkopf-1 is a member of a new family of secreted proteins and functions in head induction. Nature 391, 357–362. Gong, S. G., and Kiba, A. (1999). The role of Xmsx-2 in the anterior–posterior patterning of the mesoderm in Xenopus laevis. Differentiation 65, 131–140. Goodman, S. A., Albano, R., Wardle, F. C., Matthews, G., Tannahill, D., and Dale, L. (1998). BMP1-related metalloproteinases promote the development of ventral mesoderm in early Xenopus embryos. Dev. Biol. 195, 144–157. Goriely, A., Stella, M., Coffinier, C., Kessler, D., Mailhos, C., Dessain, S., and Desplan, C. (1996). A functional homologue of goosecoid in Drosophila. Development 122, 1641–1650. Gottlieb, S., Emanuel, B. S., Driscoll, D. A., Sellinger, B., Wang, Z., Roe, B., and Budarf, M. L. (1997). The DiGeorge syndrome minimal critical region contains a goosecoid-like (GSCL) homeobox gene that is expressed early in human development. Am. J. Hum. Genet. 60, 1194–1201. Graff, J. M., Bansal, A., and Melton, D. A. (1996). Xenopus Mad proteins transduce distinct subsets of signals for the TGF beta superfamily. Cell 85, 479–487.

1. Early Xenopus Development

55

Gritsman, K., Zhang, J., Cheng, S., Heckscher, E., Talbot, W. S., and Schier, A. F. (1999). The EGF-CFC protein one-eyed pinhead is essential for nodal signaling. Cell 97, 121–132. Gu, Z., Nomura, M., Simpson, B. B., Lei, H., Feijen, A., van den Eijnden-van Raaij, J., Donahoe, P. K., and Li, E. (1998). The type I activin receptor ActRIB is required for egg cylinder organization and gastrulation in the mouse. Genes Dev. 12, 844–857. Guger, K. A., and Gumbiner, B. M. (1995). beta-catenin has Wnt-like activity and mimics the Nieuwkoop signaling center in Xenopus dorsal–ventral patterning. Dev. Biol. 172, 115–125. Gurdon, J. B., Harger, P., Mitchell, A., and Lemaire, P. (1994). Activin signalling and response to a morphogen gradient. Nature 371, 487–492. Gurdon, J. B., Ryan, K., Stennard, F., McDowell, N., Crease, D., Dyson, S., Zorn, A., Garrett, N., Mitchell, A., and Carnac, G. (1996). Long range signalling process in embryonic development. Int. J. Dev. Biol. Suppl, 57S–58S. Gurdon, J. B., Dyson, S., and St Johnston, D. (1998). Cells’ perception of position in a concentration gradient. Cell 95, 159–162. Gurdon, J. B., Standley, H., Dyson, S., Butler, K., Langon, T., Ryan, K., Stennard, F., Shimizu, K., and Zorn, A. (1999). Single cells can sense their position in a morphogen gradient. Development 126, 5309–5317. Haegel, H., Larue, L., Ohsugi, M., Fedorov, L., Herrenknecht, K., and Kemler, R. (1995). Lack of beta-catenin affects mouse development at gastrulation. Development 121, 3529–3537. Hahn, M., and Jackle, H. (1996). Drosophila goosecoid participates in neural development but not in body axis formation. EMBO J. 15, 3077–3084. Hahn, S. A., Schutte, M., Hoque, A. T., Moskaluk, C. A., da Costa, L. T., Rozenblum, E., Weinstein, C. L., Fischer, A., Yeo, C. J., Hruban, R. H., and Kern, S. E. (1996). DPC4, a candidate tumor suppressor gene at human chromosome 18q21.1. Science 271, 350–353. Hainski, A. M., and Moody, S. A. (1992). Xenopus maternal RNAs from a dorsal animal blastomere induce a secondary axis in host embryos. Development 116, 347–355. Hamburger, V. (1988). “The Heritage of Experimental Embryology: Hans Spemann and the Organizer.” Oxford University Press, New York. Hammerschmidt, M., Pelegri, F., Mullins, M. C., Kane, D. A., van Eeden, F. J., Granato, M., Brand, M., Furutani-Seiki, M., Haffter, P., Heisenberg, C. P., Jiang, Y. J., Kelsh, R. N., Odenthal, J., Warga, R. M., and Nusslein-Volhard, C. (1996). dino and mercedes, two genes regulating dorsal development of the zebrafish embryo. Development 123, 95–102. Hansen, C. S., Marion, C. D., Steele, K., George, S., and Smith, W. C. (1997). Direct neural induction and selective inhibition of mesoderm and epidermis inducers by Xnr3. Development 124, 483– 492. Harland, R., and Gerhart, J. (1997). Formation and function of Spemann’s organizer. Annu. Rev. Cell Dev. Biol. 13, 611–667. Hata, A., Lo, R. S., Wotton, D., Lagna, G., and Massague, J. (1997). Mutations increasing autoinhibition inactivate tumour suppressors Smad2 and Smad4. Nature 388, 82–87. Hata, A., Lagna, G., Massague, J., and Hemmati-Brivanlou, A. (1998). Smad6 inhibits BMP/Smad1 signaling by specifically competing with the Smad4 tumor suppressor. Genes Dev. 12, 186–197. Hawley, S. H., Wunnenberg-Stapleton, K., Hashimoto, C., Laurent, M. N., Watabe, T., Blumberg, B. W., and Cho, K. W. (1995). Disruption of BMP signals in embryonic Xenopus ectoderm leads to direct neural induction. Genes Dev. 9, 2923–2935. Hayashi, H., Abdollah, S., Qiu, Y., Cai, J., Xu, Y. Y., Grinnell, B. W., Richardson, M. A., Topper, J. N., Gimbrone, M. A., Jr., Wrana, J. L., and Falb, D. (1997). The MAD-related protein Smad7 associates with the TGF-beta receptor and functions as an antagonist of TGF-beta signaling. Cell 89, 1165–1173. He, X., Saint-Jeannet, J. P., Woodgett, J. R., Varmus, H. E., and Dawid, I. B. (1995). Glycogen synthase kinase-3 and dorsoventral patterning in Xenopus embryos. Nature 374, 617–622. Heasman, J. (1997). Patterning the Xenopus blastula. Development 124, 4179–4191.

56

Agnes P. Chan and Laurence D. Etkin

Heasman, J., Wylie, C. C., Hausen, P., and Smith, J. C. (1984). Fates and states of determination of single vegetal pole blastomeres of X. laevis. Cell 37, 185–194. Heasman, J., Holwill, S., and Wylie, C. C. (1991). Fertilization of cultured Xenopus oocytes and use in studies of maternally inherited molecules. Methods Cell Biol. 36, 213–230. Heasman, J., Crawford, A., Goldstone, K., Garner-Hamrick, P., Gumbiner, B., McCrea, P., Kintner, C., Noro, C. Y., and Wylie, C. (1994). Overexpression of cadherins and underexpression of beta-catenin inhibit dorsal mesoderm induction in early Xenopus embryos. Cell 79, 791–803. Heisenberg, C. P., Tada, M., Rauch, G. J., Saude, L., Concha, M. L., Geisler, R., Stemple, D. L., Smith, J. C., and Wilson, S. W. (2000). Silberblick/Wnt11 mediates convergent extension movements during zebrafish gastrulation. Nature 405, 76–81. Hemmati-Brivanlou, A., and Melton, D. A. (1992). A truncated activin receptor inhibits mesoderm induction and formation of axial structures in Xenopus embryos. Nature 359, 609–614. Hemmati-Brivanlou, A., Kelly, O. G., and Melton, D. A. (1994). Follistatin, an antagonist of activin, is expressed in the Spemann organizer and displays direct neuralizing activity. Cell 77, 283–295. Henry, G. L., and Melton, D. A. (1998). Mixer, a homeobox gene required for endoderm development. Science 281, 91–96. Henry, G. L., Brivanlou, I. H., Kessler, D. S., Hemmati-Brivanlou, A., and Melton, D. A. (1996). TGFbeta signals and a pattern in Xenopus laevis endodermal development. Development 122, 1007–1015. Hermesz, E., Mackem, S., and Mahon, K. A. (1996). Rpx: a novel anterior-restricted homeobox gene progressively activated in the prechordal plate, anterior neural plate and Rathke’s pouch of the mouse embryo. Development 122, 41–52. Holley, S. A., Neul, J. L., Attisano, L., Wrana, J. L., Sasai, Y., O’Connor, M. B., De Robertis, E. M., and Ferguson, E. L. (1996). The Xenopus dorsalizing factor noggin ventralizes Drosophila embryos by preventing DPP from activating its receptor. Cell 86, 607–617. Holowacz, T., and Elinson, R. P. (1993). Cortical cytoplasm, which induces dorsal axis formation in Xenopus, is inactivated by UV irradiation of the oocyte. Development 119, 277–285. Holowacz, T., and Elinson, R. P. (1995). Properties of the dorsal activity found in the vegetal cortical cytoplasm of Xenopus eggs. Development 121, 2789–2798. Holowacz, T., and Sokol, S. (1999). FGF is required for posterior neural patterning but not for neural induction. Dev. Biol. 205, 296–308. Homon, J. A., and Gong, S. G. (1999). A statistical analysis of the overexpression of the msx2 RNA in Xenopus laevis. Arch. Oral Biol. 44, 795–803. Hongo, I., Kengaku, M., and Okamoto, H. (1999). FGF signaling and the anterior neural induction in Xenopus. Dev. Biol. 216, 561–581. Hoodless, P. A., Haerry, T., Abdollah, S., Stapleton, M., O’Connor, M. B., Attisano, L., and Wrana, J. L. (1996). MADR1, a MAD-related protein that functions in BMP2 signaling pathways. Cell 85, 489–500. Hoodless, P. A., Tsukazaki, T., Nishimatsu, S., Attisano, L., Wrana, J. L., and Thomsen, G. H. (1999). Dominant-negative Smad2 mutants inhibit activin/Vg1 signaling and disrupt axis formation in Xenopus. Dev. Biol. 207, 364–379. Hoppler, S., and Moon, R. T. (1998). BMP-2/-4 and Wnt-8 cooperatively pattern the Xenopus mesoderm. Mech. Dev. 71, 119–129. Hoppler, S., Brown, J. D., and Moon, R. T. (1996). Expression of a dominant-negative Wnt blocks induction of MyoD in Xenopus embryos. Genes Dev. 10, 2805–2817. Horb, M. E., and Thomsen, G. H. (1997). A vegetally localized T-box transcription factor in Xenopus eggs specifies mesoderm and endoderm and is essential for embryonic mesoderm formation. Development 124, 1689–1698. Houston, D. W., and King, M. L. (2000). A critical role for Xdazl a germ plasm–localized RNA, in the differentiation of primordial germ cells in Xenopus. Development 127, 447–456. Houston, D. W., Zhang, J., Maines, J. Z., Wasserman, S. A., and King, M. L. (1998). A Xenopus DAZ-like gene encodes an RNA component of germ plasm and is a functional homologue of Drosophila boule. Development 125, 171–180.

1. Early Xenopus Development

57

Hsu, W., Zeng, L., and Costantini, F. (1999). Identification of a domain of Axin that binds to the serine/threonine protein phosphatase 2A and a self-binding domain. J. Biol. Chem. 274, 3439–3445. Huang, L., Li, X., El-Hodiri, H. M., Dayal, S., Wikramanayake, A. H., and Klein, W. H. (2000). Involvement of Tcf/Lef in establishing cell types along the animal–vegetal axis of sea urchins. Dev. Genes Evol. 210, 73–81. Huber, A. H., Nelson, W. J., and Weis, W. I. (1997). Three-dimensional structure of the armadillo repeat region of beta-catenin. Cell 90, 871–882. Hudson, J. W., Alarcon, V. B., and Elinson, R. P. (1996). Identification of new localized RNAs in the Xenopus oocyte by differential display PCR. Dev. Genet. 19, 190–198. Hudson, C., and Woodland, H. R. (1998). Xpat, a gene expressed specifically in germ plasm and primordial germ cells of Xenopus laevis. Mech. Dev. 73, 159–168. Hudson, C., Clements, D., Friday, R. V., Stott, D., and Woodland, H. R. (1997). Xsox17-alpha and -beta mediate endoderm formation in Xenopus. Cell 91, 397–405. Iemura, S., Yamamoto, T. S., Takagi, C., Uchiyama, H., Natsume, T., Shimasaki, S., Sugino, H., and Ueno, N. (1998). Direct binding of follistatin to a complex of bone-morphogenetic protein and its receptor inhibits ventral and epidermal cell fates in early Xenopus embryo. Proc. Natl. Acad. Sci. USA 95, 9337–9342. Imamura, T., Takase, M., Nishihara, A., Oeda, E., Hanai, J., Kawabata, M., and Miyazono, K. (1997). Smad6 inhibits signalling by the TGF-beta superfamily. Nature 389, 622–626. Isaacs, H. V., Pownall, M. E., and Slack, J. M. (1994). eFGF regulates Xbra expression during Xenopus gastrulation. EMBO J. 13, 4469–4481. Itoh, K., and Sokol, S. Y. (1999). Axis determination by inhibition of Wnt signaling in Xenopus. Genes Dev. 13, 2328–2336. Itoh, K., Jacob, J., and Sokol, S. Y. (1998). A role for Xenopus Frizzled 8 in dorsal development. Mech. Dev. 74, 145–157. Jesuthasan, S., and Stahle, U. (1996). Dynamic microtubules and specification of the zebrafish embryonic axis. Curr. Biol. 7, 31–42. Jiang, J., and Struhl, G. (1998). Regulation of the Hedgehog and Wingless signalling pathways by the F-box/WD40-repeat protein Slimb. Nature 391, 493–496. Jones, E. A., and Woodland, H. R. (1987). The development of animal cap cells in Xenopus: a measure of the start of animal cap competence to form mesoderm. Development 101, 557–563. Jones, C. M., Kuehn, M. R., Hogan, B. L., Smith, J. C., and Wright, C. V. (1995). Nodal-related signals induce axial mesoderm and dorsalize mesoderm during gastrulation. Development 121, 3651–3662. Jones, C. M., Armes, N., and Smith, J. C. (1996a). Signalling by TGF-beta family members: short-range effects of Xnr-2 and BMP-4 contrast with the long-range effects of activin. Curr. Biol. 6, 1468–1475. Jones, C. M., Dale, L., Hogan, B. L., Wright, C. V., and Smith, J. C. (1996b). Bone morphogenetic protein-4 (BMP-4) acts during gastrula stages to cause ventralization of Xenopus embryos. Development 122, 1545–1554. Jones, C. M., Broadbent, J., Thomas, P. Q., Smith, J. C., and Beddington, R. S. (1999). An anterior signalling centre in Xenopus revealed by the homeobox gene XHex. Curr. Biol. 9, 946–954. Jonkers, J., Korswagen, H. C., Acton, D., Breuer, M., and Berns, A. (1997). Activation of novel protooncogene, Frat1, contributes to progression of mouse T-cell lymphomas. EMBO J. 16, 441–450. Joseph, E. M., and Melton, D. A. (1997). Xnr4: a Xenopus nodal-related gene expressed in the Spemann organizer. Dev. Biol. 184, 367–372. Joseph, E. M., and Melton, D. A. (1998). Mutant Vg1 ligands disrupt endoderm and mesoderm formation in Xenopus embryos. Development 125, 2677–2685. Kageura, H. (1990). Spatial distribution of the capacity to initiate a secondary embryo in the 32-cell embryo of Xenopus laevis. Dev. Biol. 142, 432–438. Kageura, H. (1997). Activation of dorsal development by contact between the cortical dorsal determinant and the equatorial core cytoplasm in eggs of Xenopus laevis. Development 124, 1543–1551. Kikkawa, M., Takano, K., and Shinagawa, A. (1996). Location and behavior of dorsal determinants during first cell cycle in Xenopus eggs. Development 122, 3687–3696.

58

Agnes P. Chan and Laurence D. Etkin

Kinoshita, N., Minshull, J., and Kirschner, M. W. (1995). The identification of two novel ligands of the FGF receptor by a yeast screening method and their activity in Xenopus development. Cell 83, 621–630. Kinzler, K. W., and Vogelstein, B. (1996). Lessons from hereditary colorectal cancer. Cell 87, 159–170. Klein, P. S., and Melton, D. A. (1996). A molecular mechanism for the effect of lithium on development. Proc. Natl. Acad. Sci. USA 93, 8455–8459. Klingensmith, J., and Nusse, R. (1994). Signaling by wingless in Drosophila. Dev. Biol. 166, 396–414. Kloc, M., and Etkin, L. D. (1995). Two distinct pathways for the localization of RNAs at the vegetal cortex in Xenopus oocytes. Development 121, 287–297. Kloc, M., Miller, M., Carrasco, A. E., Eastman, E., and Etkin, L. (1989). The maternal store of the xlgv7 mRNA in full-grown oocytes is not required for normal development in Xenopus. Development 107, 899–907. Kloc, M., Larabell, C., Chan, A. P., and Etkin, L. D. (1998). Contribution of METRO pathway localized molecules to the organization of the germ cell lineage. Mech. Dev. 75, 81–93. Kloc, M., Bilinski, S., Chan, A. P., and Etkin, L. D. (1999). The targeting of Xcat2 mRNA to the germinal granules depends on a cis-acting germinal granule localization element within the 3’UTR. Dev. Biol. 217, 221–229. Kloc, M., Bilinski, S., Chan, A. P., Allen, L. H., Zearfoss, N. R., and Etkin, L. D. (2000). RNA localization and germ cell determination in Xenopus. Int. Rev. Cytol. 203, 63–91. Kobayashi, S., Yamada, M., Asaoka, M., and Kitamura, T. (1996). Essential role of the posterior morphogen nanos for germline development in Drosophila. Nature 380, 708–711. Kodjabachian, L., and Lemaire, P. (1998). Embryonic induction: is the Nieuwkoop centre a useful concept? Curr. Biol. 8, R918–921. Kodjabachian, L., Dawid, I. B., and Toyama, R. (1999). Gastrulation in Zebrafish: what mutants teach us. Dev. Biol. 213, 231–245. Koelle, M. R. (1997). A new family of G-protein regulators—the RGS proteins. Curr. Opin. Cell Biol. 9, 143–147. Kofron, M., Demel, T., Xanthos, J., Lohr, J., Sun, B., Sive, H., Osada, S., Wright, C., Wylie, C., and Heasman, J. (1999). Mesoderm induction in Xenopus is a zygotic event regulated by maternal VegT via TGF␤ growth factors. Development 126, 5759–5770. Koller, B. H., and Smithies, O. (1992). Altering genes in animals by gene targeting. Annu. Rev. Immunol. 10, 705–730. Kolm, P. J., and Sive, H. L. (1995). Efficient hormone-inducible protein function in Xenopus laevis. Dev. Biol. 171, 267–272. Koos, D. S., and Ho, R. K. (1998). The nieuwkoid gene characterizes and mediates a Nieuwkoopcenter-like activity in the zebrafish. Curr. Biol. 8, 1199–1206. Korinek, V., Barker, N., Morin, P. J., van Wichen, D., de Weger, R., Kinzler, K. W., Vogelstein, B., and Clevers, H. (1997). Constitutive transcriptional activation by a beta-catenin-Tcf complex in APC−/−colon carcinoma. Science 275, 1784–1787. Kroll, K. L., and Amaya, E. (1996). Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signaling requirements during gastrulation. Development 122, 3173–3183. Kroll, K. L., and Gerhart, J. C. (1994). Transgenic X. laevis embryos from eggs transplanted with nuclei of transfected cultured cells. Science 266, 650–653. Krupnik, V. E., Sharp, J. D., Jiang, C., Robison, K., Chickering, T. W., Amaravadi, L., Brown, D. E., Guyot, D., Mays, G., Leiby, K., Chang, B., Duong, T., Goodearl, A. D., Gearing, D. P., Sokol, S. Y., and McCarthy, S. A. (1999). Functional and structural diversity of the human Dickkopf gene family. Gene 238, 301–313. Ku, M., and Melton, D. A. (1993). Xwnt-11: a maternally expressed Xenopus Wnt gene. Development 119, 1161–1173. Labbe, E., Silvestri, C., Hoodless, P. A., Wrana, J. L., and Attisano, L. (1998). Smad2 and Smad3 positively and negatively regulate TGF beta-dependent transcription through the forkhead DNA-bindign protein FAST2. Mol. Cell 2, 109–120.

1. Early Xenopus Development

59

LaBonne, C., and Whitman, M. (1994). Mesoderm induction by activin requires FGF-mediated intracellular signals. Development 120, 463–472. LaBonne, C., and Whitman, M. (1997). Localization of MAP kinase activity in early Xenopus embryos: implications for endogenous FGF signaling. Dev. Biol. 183, 9–20. LaBonne, C., Burke, B., and Whitman, M. (1995). Role of MAP kinase in mesoderm induction and axial patterning during Xenopus development. Development 121, 1475–1486. Labouesse, M., and Mango, S. E. (1999). Patterning the C. elegans embryo: moving beyond the cell lineage. Trends Genet. 15, 307–313. Ladher, R., Mohun, T. J., Smith, J. C., and Snape, A. M. (1996). Xom: a Xenopus homeobox gene that mediates the early effects of BMP-4. Development 122, 2385–2394. Lagna, G., and Hemmati-Brivanlou, A. (1999). A molecular basis for Smad specificity. Dev. Dyn. 214, 269–277. Lagna, G., Hata, A., Hemmati-Brivanlou, A., and Massague, J. (1996). Partnership between DPC4 and SMAD proteins in TGF-beta signalling pathways. Nature 383, 832–836. Landesman, Y., and Sokol, S. Y. (1997). Xwnt-2b is a novel axis-inducing Xenopus Wnt, which is expressed in embryonic brain. Mech. Dev. 63, 199–209. Larabell, C. A., Rowning, B. A., Wells, J., Wu, M., and Gerhart, J. C. (1996). Confocal microscopy analysis of living Xenopus eggs and the mechanism of cortical rotation. Development 122, 1281– 1289. Larabell, C. A., Torres, M., Rowning, B. A., Yost, C., Miller, J. R., Wu, M., Kimelman, D., and Moon, R. T. (1997). Establishment of the dorso-ventral axis in Xenopus embryos is presaged by early asymmetries in beta-catenin that are modulated by the Wnt signaling pathway. J. Cell Biol. 136, 1123–1136. Larrain, J., Bachiller, D., Lu, B., Agius, E., Piccolo, S., and De Robertis, E. M. (2000). BMP-binding modules in chordin: a model for signalling regulation in the extracellular space. Development 127, 821–830. Latinkic, B. V., Umbhauer, M., Neal, K. A., Lerchner, W., Smith, J. C., and Cunliffe, V. (1997). The Xenopus Brachyury promoter is activated by FGF and low concentrations of activin and suppressed by high concentrations of activin and by paired-type homeodomain proteins. Genes Dev. 11, 3265–3276. Laurent, M. N., Blitz, I. L., Hashimoto, C., Rothbacher, U., and Cho, K. W. (1997). The Xenopus homeobox gene twin mediates Wnt induction of goosecoid in establishment of Spemann’s organizer. Development 124, 4905–4916. Lawson, K. A., Meneses, J. J., and Pedersen, R. A. (1991). Clonal analysis of epiblast fate during germ layer formation in the mouse embryo. Development 113, 891–911. Lemaire, P., and Gurdon, J. B. (1994). A role for cytoplasmic determinants in mesoderm patterning: cell-autonomous activation of the goosecoid and Xwnt-8 genes along the dorsoventral axis of early Xenopus embryos. Development 120, 1191–1199. Lemaire, P., Garrett, N., and Gurdon, J. B. (1995). Expression cloning of Siamois, a Xenopus homeobox gene expressed in dorsal–vegetal cells of blastulae and able to induce a complete secondary axis. Cell 81, 85–94. Lemaire, L., Roeser, T., Izpisua-Belmonte, J. C., and Kessel, M. (1997). Segregating expression domains of two goosecoid genes during the transition from gastrulation to neurulation in chick embryos. Development 124, 1443–1452. Lemaire, P., Darras, S., Caillol, D., and Kodjabachian, L. (1998). A role for the vegetally expressed Xenopus gene Mix.1 in endoderm formation and in the restriction of mesoderm to the marginal zone. Development 125, 2371–2380. Levin, M. (1998). The roles of activin and follistatin signaling in chick gastrulation. Int. J. Dev. Biol. 42, 553–559. Leyns, L., Bouwmeester, T., Kim, S. H., Piccolo, S., and De Robertis, E. M. (1997). Frzb-1 is a secreted antagonist of Wnt signaling expressed in the Spemann organizer. Cell 88, 747–756. Liu, F., Hata, A., Baker, J. C., Doody, J., Carcamo, J., Harland, R. M., and Massague, J. (1996). A human Mad protein acting as a BMP-regulated transcriptional activator. Nature 381, 620–623.

60

Agnes P. Chan and Laurence D. Etkin

Liu, F., Pouponnot, C., and Massague, J. (1997). Dual role of the Smad4/DPC4 tumor suppressor in TGF-beta-inducible transcriptional complexes. Genes Dev. 11, 3157–3167. Liu, C., Kato, Y., Zhang, Z., Do, V. M., Yankner, B. A., and He, X, (1999a). beta-Trcp couples beta-catenin phosphorylation-degradation and regulates Xenopus axis formation. Proc. Natl. Acad. Sci. USA 96, 6273–6278. Liu, P., Wakamiya, M., Shea, M. J., Albrecht, U., Behringer, R. R., and Bradley, A. (1999b). Requirement for Wnt3 in vertebrate axis formation. Nat. Genet. 22, 361–365. Logan, C. Y., Miller, J. R., Ferkowicz, M. J., and McClay, D. R. (1999). Nuclear beta-catenin is required to specify vegetal cell fates in the sea urchin embryo. Development 126, 345–357. Love, J. J., Li, X., Case, D. A., Giese, K., Grosschedl, R., and Wright, P. E. (1995). Structural basis for DNA bending by the architectural transcription factor LEF-1. Nature 376, 791–795. Lustig, K. D., Kroll, K., Sun, E., Ramos, R., Elmendorf, H., and Kirschner, M. W. (1996a). A Xenopus nodal-related gene that acts in synergy with noggin to induce complete secondary axis and notochord formation. Development 122, 3275–3282. Lustig, K. D., Kroll, K. L., Sun, E. E., and Kirschner, M. W. (1996b). Expression cloning of a Xenopus T-related gene (Xombi) involved in mesodermal patterning and blastopore lip formation. Development 122, 4001–4012. MacArthur, H., Houston, D. W., Bubunenko, M., Mosquera, L., and King, M. L. (2000). DEADSouth is a germ plasm specific DEAD-box RNA helicase in Xenopus related to eIF4A. Mech. Dev. 95, 291–295. Mangold, O. (1933). Uber die Induktionsfahigkeit der verschiedenen Bezirke der Neurula von Urodelen. Naturwissenschaften 21, 761–766. Mariani, F. V., and Harland, R. M. (1998). XBF-2 is a transcriptional repressor that converts ectoderm into neural tissue. Development 125, 5019–5031. Marikawa, Y., and Elinson, R. P. (1998). beta-TrCP is a negative regulator of Wnt/beta-catenin signaling pathway and dorsal axis formation in Xenopus embryos. Mech. Dev. 77, 75–80. Marikawa, Y., and Elinson, R. P. (1999). Relationship of vegetal cortical dorsal factors in the Xenopus egg with the Wnt/beta-catenin signaling pathway. Mech. Dev. 89, 93–102. Marom, K., Fainsod, A., and Steinbeisser, H. (1999). Patterning of the mesoderm involves several threshold responses to BMP-4 and Xwnt-8. Mech. Dev. 87, 33–44. Marsh-Armstrong, N., Huang, H., Berry, D. L., and Brown, D. D. (1999). Germ-line transmission of transgenes in Xenopus laevis. Proc. Natl. Acad. Sci. USA 96, 14389–14393. Maslanski, J. A., Leshko, L., and Busa, W. B. (1992). Lithium-sensitive production of inositol phosphates during amphibian embryonic mesoderm induction. Science 256, 243–245. Massague, J. (1998). TGF-beta signal transduction. Annu. Rev. Biochem. 67, 753–791. Matzuk, M. M., Kumar, T. R., Vassalli, A., Bickenbach, J. R., Roop, D. R., Jaenisch, R., and Bradley, A. (1995a). Functional analysis of activins during mammalian development. Nature 374, 354–356. Matzuk, M. M., Lu, N., Vogel, H., Sellheyer, K., Roop, D. R., and Bradley, A. (1995b). Multiple defects and perinatal death in mice deficient in follistatin. Nature 374, 360–363. McCrea, P. D., Brieher, W. M., and Gumbiner, B. M. (1993). Induction of a secondary body axis in Xenopus by antibodies to beta-catenin. J. Cell Biol. 123, 477–484. McKendry, R., Hsu, S. C., Harland, R. M., and Grosschedl, R. (1997). LEF-1/TCF proteins mediate wnt-inducible transcription from the Xenopus nodal-related 3 promoter. Dev. Biol. 192, 420–431. McMahon, A. P., and Moon, R. T. (1989). Ectopic expression of the proto-oncogene int-1 in Xenopus embryos leads to duplication of the embryonic axis. Cell 58, 1075–1084. McMahon, J. A., Takada, S., Zimmerman, L. B., Fan, C. M., Harland, R. M., and McMahon, A. P. (1998). Noggin-mediated antagonism of BMP signaling is required for growth and patterning of the neural tube and somite. Genes Dev. 12, 1438–1452. Mead, P. E., Brivanlou, I. H., Kelley, C. M., and Zon, L. I. (1996). BMP-4-responsive regulation of dorsal–ventral patterning by the homeobox protein Mix.1. Nature 382, 357–360. Meersseman, G., Verschueren, K., Nelles, L., Blumenstock, C., Kraft, H., Wuytens, G., Remacle, J., Kozak, C. A., Tylzanowski, P., Niehrs, C., and Huylebroeck, D. (1997). The C-terminal domain

1. Early Xenopus Development

61

of Mad-like signal transducers is sufficient for biological activity in the Xenopus embryo and transcriptional activation. Mech. Dev. 61, 127–140. Melby, A. E., Clements, W. K., and Kimelman, D. (1999). Regulation of dorsal gene expression in Xenopus by the ventralizing homeodomain gene Vox. Dev. Biol. 211, 293–305. Melton, D. A. (1987). Translocation of a localized maternal mRNA to the vegetal pole of Xenopus oocytes. Nature 328, 80–82. Merriam, J. M., Rubenstein, A. B., and Klymkowsky, M. W. (1997). Cytoplasmically anchored plakoglobin induces a WNT-like phenotype in Xenopus. Dev. Biol. 185, 67–81. Miller, J. R., and Moon, R. T. (1997). Analysis of the signaling activities of localization mutants of beta-catenin during axis specification in Xenopus. J. Cell Biol. 139, 229–243. Miller, J. R., Rowning, B. A., Larabell, C. A., Yang-Snyder, J. A., Bates, R. L., and Moon, R. T. (1999). Establishment of the dorsal-ventral axis in Xenopus embryos coincides with the dorsal enrichment of dishevelled that is dependent on cortical rotation. J. Cell Biol. 146, 427–437. Molenaar, M., van de Wetering, M., Oosterwegel, M., Peterson-Maduro, J., Godsave, S., Korinek, V., Roose, J., Destree, O., and Clevers, H. (1996). XTcf-3 transcription factor mediates beta-catenin-induced axis formation in Xenopus embryos. Cell 86, 391–399. Monaghan, A. P., Kioschis, P., Wu, W., Zuniga, A., Bock, D., Poustka, A., Delius, H., and Niehrs, C. (1999). Dickkopf genes are co-ordinately expressed in mesodermal lineages. Mech. Dev. 87, 45–56. Moon, R. T., and Kimelman, D. (1998). From cortical rotation to organizer gene expression: toward a molecular explanation of axis specification in Xenopus. Bioessays 20, 536–545. Moon, R. T., Campbell, R. M., Christian, J. L., McGrew, L. L., Shih, J., and Fraser, S. (1993). Xwnt-5A: a maternal Wnt that affects morphogenetic movements after overexpression in embryos of Xenopus laevis. Development 119, 97–111. Moos, M., Jr., Wang, S., and Krinks, M. (1995). Anti-dorsalizing morphogenetic protein is a novel TGF-beta homolog expressed in the Spemann organizer. Development 121, 4293–4301. Mosquera, L., Forristall, C., Zhou, Y., and King, M. L. (1993). A mRNA localized to the vegetal cortex of Xenopus oocytes encodes a protein with a nanos-like zinc finger domain. Development 117, 377–386. Mullins, M. C. (1999). Embryonic axis formation in the zebrafish. Methods Cell Biol. 59, 159–178. Mulnard, J. G., and Puissant, F. (1984). Development of mouse embryos after ultracentrifugation at the pronuclei stage. Arch. Biol. 97, 301–315. Nagafuchi, A., and Takeichi, M. (1989). Transmembrane control of cadherin-mediated cell adhesion: a 94 kDa protein functionally associated with a specific region of the cytoplasmic domain of E-cadherin. Cell Regul. 1, 37–44. Nakao, A., Afrakhte, M., Moren, A., Nakayama, T., Christian, J. L., Heuchel, R., Itoh, S., Kawabata, M., Heldin, N. E., Heldin, C. H., and ten Dijke, P. (1997a). Identification of Smad7, a TGF-beta-inducible antagonist of TGF-beta signalling. Nature 389, 631–635. Nakao, A., Roijer, E., Imamura, T., Souchelnytskyi, S., Stenman, G., Heldin, C. H., and ten Dijke, P. (1997b). Identification of Smad2, a human Mad-related protein in the transforming growth factor beta signaling pathway. J. Biol. Chem. 272, 2896–2900. Nelson, R. W., and Gumbiner, B. M. (1999). A cell-free assay system for beta-catenin signaling that recapitulates direct inductive events in the early Xenopus laevis embryo. J. Cell Biol. 147, 367–374. Neumann, C., and Cohen, S. (1997). Morphogens and pattern formation. Bioessays 19, 721–729. Newman, C. S., Chia, F., and Krieg, P. A. (1997). The XHex homeobox gene is expressed during development of the vascular endothelium: overexpression leads to an increase in vascular endothelial cell number. Mech. Dev. 66, 83–93. Niehrs, C. (1999). Head in the WNT: the molecular nature of Spemann’s head organizer. Trends Gen et. 15, 314–319. Niehrs, C. (2000), Amphibian organizer activity. Methods Mol. Biol. 137, 179–183. Nieuwkoop, P. D. (1997). Short historical survey of pattern formation in the endomesoderm and the neural anlage in the vertebrates: the role of vertical and planar inductive actions. Cell Mol. Life Sci. 53, 305–318.

62

Agnes P. Chan and Laurence D. Etkin

Nishida, H. (1992). Regionality of egg cytoplasm that promotes muscle differentiation in embryo of the ascidian, Halocynthia roretzi. Development 116, 521–529. Nishida, H. (1997). Cell fate specification by localized cytoplasmic determinants and cell interactions in ascidian embryos. Int. Rev. Cytol. 176, 245–306. Nishimatsu, S., and Thomsen, G. H. (1998). Ventral mesoderm induction and patterning by bone morphogenetic protein heterodimers in Xenopus embryos. Mech. Dev. 74, 75–88. Nomura, M., and Li, E. (1998). Smad2 role in mesoderm formation, left-right patterning and craniofacial development. Nature 393, 786–790. Normanno, N., Ciardiello, F., Brandt, R., and Salomon, D. S. (1994). Epidermal growth factor-related peptides in the pathogenesis of human breast cancer. Breast Cancer Res. Treat 29, 11–27. Oelgeschlager, M., Larrain, J., Geissert, D., and De Robertis, E. M. (2000). The evolutionarily conserved BMP-binding protein Twisted gastrulation promotes BMP signalling. Nature 405, 757–763. Onichtchouk, D., Gawantka, V., Dosch, R., Delius, H., Hirschfeld, K., Blumenstock, C., and Niehrs, C. (1996). The Xvent-2 homeobox gene is part of the BMP-4 signalling pathway controlling dorsoventral patterning of Xenopus mesoderm. Development 122, 3045–3053. Onichtchouk, D., Glinka, A., and Niehrs, C. (1998). Requirement for Xvent-1 and Xvent-2 gene function in dorsoventral patterning of Xenopus mesoderm. Development 125, 1447–1456. Onichtchouk, D., Chen, Y. G., Dosch, R., Gawantka, V., Delius, H., Massague, J., and Niehrs, C. (1999). Silencing of TGF-beta signalling by the pseudoreceptor BAMBI. Nature 401, 480–485. Orford, K., Crockett, C., Jensen, J. P., Weissman, A. M., and Byers, S. W. (1997). Serine phosphorylation-regulated ubiquitination and degradation of beta-catenin. J. Biol. Chem. 272, 24735–24738. Osada, S. I., and Wright, C. V. (1999). Xenopus nodal-related signaling is essential for mesendodermal patterning during early embryogenesis. Development 126, 3229–3240. Ozaki, T., and Sakiyama, S. (1993). Molecular cloning and characterization of a cDNA showing negative regulation in v-src-transformed 3Y1 rat fibroblasts. Proc. Natl. Acad. Sci. USA 90, 2593–2597. Ozawa, M., Baribault, H., and Kemler, R. (1989). The cytoplasmic domain of the cell adhesion molecule uvomorulin associates with three independent proteins structurally related in different species. EMBO J. 8, 1711–1717. Pannese, M., Polo, C., Andreazzoli, M., Vignali, R., Kablar, B., Barsacchi, G., and Boncinelli, E. (1995). The Xenopus homologue of Otx2 is a maternal homeobox gene that demarcates and specifies anterior body regions. Development 121, 707–720. Pearce, J. J., Penny, G., and Rossant, J. (1999). A mouse cerberus/Dan-related gene family. Dev. Biol. 209, 98–110. Peters, J. M., McKay, R. M., McKay, J. P., and Graff, J. M. (1999). Casein kinase I transduces Wnt signals. Nature 401, 345–350. Peyrieras, N., Strahle, U., and Rosa, F. (1998). Conversion of zebrafish blastomeres to an endodermal fate by TGF-beta- related signaling. Curr. Biol. 8, 783–786. Piccolo, S., Sasai, Y., Lu, B., and De Robertis, E. M. (1996). Dorsoventral patterning in Xenopus: inhibition of ventral signals by direct binding of chordin to BMP-4. Cell 86, 589–598. Piccolo, S., Agius, E., Lu, B., Goodman, S., Dale, L., and De Robertis, E. M. (1997). Cleavage of Chordin by Xolloid metalloprotease suggests a role for proteolytic processing in the regulation of Spemann organizer activity. Cell 91, 407–416. Piccolo, S., Agius, E., Leyns, L., Bhattacharyya, S., Grunz, H., Bouwmeester, T., and De Robertis, E. M. (1999). The head inducer Cerberus is a multifunctional antagonist of Nodal, BMP and Wnt signals. Nature 397, 707–710. Pierce, S. B., and Kimelman, D. (1995). Regulation of Spemann organizer formation by the intracellular kinase Xgsk-3. Development 121, 755–765. Polakis, P. (1997). The adenomatous polyposis coli (APC) tumor suppressor. Biochim. Biophys. Acta. 1332, F127–147. Reilly, K. M., and Melton, D. A. (1996). Short-range signaling by candidate morphogens of the TGF beta family and evidence for a relay mechanism of induction. Cell 86, 743–754.

1. Early Xenopus Development

63

Renucci, A., Lemarchandel, V., and Rosa, F. (1996). An activated form of type I serine/threonine kinase receptor TARAM-A reveals a specific signalling pathway involved in fish head organiser formation. Development 122, 3735–3743. Rivera-Perez, J. A., Mallo, M., Gendron-Maguire, M., Gridley, T., and Behringer, R. R. (1995). Goosecoid is not an essential component of the mouse gastrula organizer but is required for craniofacial and rib development. Development 121, 3005–3012. Roose, J., Molenaar, M., Peterson, J., Hurenkamp, J., Brantjes, H., Moerer, P., van de Wetering, M., Destree, O., and Clevers, H. (1998). The Xenopus Wnt effector XTcf-3 interacts with Groucho-related transcriptional repressors. Nature 395, 608–612. Rosa, F. M. (1989). Mix.1, a homeobox mRNA inducible by mesoderm inducers, is expressed mostly in the presumptive endodermal cells of Xenopus embryos. Cell 57, 965–974. Rose, L. S., and Kemphues, K. J. (1998). Early patterning of the C. elegans embryo. Annu. Rev. Genet. 32, 521–545. Rowning, B. A., Wells, J., Wu, M., Gerhart, J. C., Moon, R. T., and Larabell, C. A. (1997). Microtubule-mediated transport of organelles and localization of beta-catenin to the future dorsal side of Xenopus eggs. Proc. Natl. Acad. Sci. USA 94, 1224–1229. Rubinfeld, B., Albert, I., Porfiri, E., Fiol, C., Munemitsu, S., and Polakis, P. (1996). Binding of GSK3beta to the APC/beta-catenin complex and regulation of complex assembly. Science 272, 1023–1026. Ryan, K., Garrett, N., Mitchell, A., and Gurdon, J. B. (1996). Eomesodermin, a key early gene in Xenopus mesoderm differentiation. Cell 87, 989–1000. Saka, Y., Tada, M., and Smith, J. C. (2000). A screen for targets of the Xenopus T-box gene Xbra. Mech. Dev. 93, 27–39. Sakai, M. (1996). The vegetal determinants required for the Spemann organizer move equatorially during the first cell cycle. Development 122, 2207–2214. Sakanaka, C., and Williams, L. T. (1999). Functional domains of axin. Importance of the C terminus as an oligomerization domain. J. Biol. Chem. 274, 14090–14093. Salic, A. N., Kroll, K. L., Evans, L. M., and Kirschner, M. W. (1997). Sizzled: a secreted Xwnt8 antagonist expressed in the ventral marginal zone of Xenopus embryos. Development 124, 4739–4748. Sasai, Y., Lu, B., Steinbeisser, H., Geissert, D., Gont, L. K., and De Robertis, E. M. (1994). Xenopus chordin: a novel dorsalizing factor activated by organizer-specific homeobox genes. Cell 79, 779–790. Sasai, Y., Lu, B., Steinbeisser, H., and De Robertis, E. M. (1995). Regulation of neural induction by the Chd and Bmp-4 antagonistic patterning signals in Xenopus. Nature 376, 333–336. Sasai, Y., Lu, B., Piccolo, S., and De Robertis, E. M. (1996). Endoderm induction by the organizersecreted factors chordin and noggin in Xenopus animal caps. EMBO J. 15, 4547–4555. Satou, Y. (1999). posterior end mark 3 ( pem-3), an ascidian maternally expressed gene with localized mRNA encodes a protein with Caenorhabditis elegans MEX-3-like KH domains. Dev. Biol. 212, 337–350. Schier, A. F., Neuhauss, S. C., Helde, K. A., Talbot, W. S., and Driever, W. (1997). The one-eyed pinhead gene functions in mesoderm and endoderm formation in zebrafish and interacts with no tail. Development 124, 327–342. Schmidt, J. E., von Dassow, G., and Kimelman, D. (1996). Regulation of dorsal–ventral patterning: the ventralizing effects of the novel Xenopus homeobox gene Vox. Development 122, 1711–1721. Schneider, V. A., and Mercola, M. (1999). Spatially distinct head and heart inducers within the Xenopus organizer region. Curr. Biol. 9, 800–809. Schroeder, K. E., Condic, M. L., Eisenberg, L. M., and Yost, H. J. (1999). Spatially regulated translation in embryos: asymmetric expression of maternal Wnt-11 along the dorsal–ventral axis in Xenopus. Dev. Biol. 214, 288–297. Seeling, J. M., Miller, J. R., Gil, R., Moon, R. T., White, R., and Virshup, D. M. (1999). Regulation of beta-catenin signaling by the B56 subunit of protein phosphatase 2A. Science 283, 2089–2091. Shawlot, W., and Behringer, R. R. (1995). Requirement for Lim1 in head-organizer function. Nature 374, 425–430.

64

Agnes P. Chan and Laurence D. Etkin

Sheldahl, L. C., Park, M., Malbon, C. C., and Moon, R. T. (1999). Protein kinase C is differentially stimulated by Wnt and Frizzled homologs in a G-protein-dependent manner. Curr. Biol. 9, 695–698. Shen, M. M., Wang, H., and Leder, P. (1997). A differential display strategy identifies Cryptic, a novel EGF-related gene expressed in the axial and lateral mesoderm during mouse gastrulation. Development 124, 429–442. Shimell, M. J., Ferguson, E. L., Childs, S. R., and O’Connor, M. B. (1991). The Drosophila dorsal– ventral patterning gene tolloid is related to human bone morphogenetic protein 1. Cell 67, 469–481. Shuttleworth, J., and Colman, A. (1988). Antisense oligonucleotide-directed cleavage of mRNA in Xenopus oocytes and eggs. EMBO J. 7, 427–434. Siegfried, E., and Perrimon, N. (1994). Drosophila wingless: a paradigm for the function and mechanism of Wnt signaling. Bioessays 16, 395–404. Simpson, E. H., Johnson, D. K., Hunsicker, P, Suffolk, R., Jordan, S. A., and Jackson, I. J. (1999). The mouse Cer1 (Cerberus related or homologue) gene is not required for anterior pattern formation. Dev. Biol. 213, 202–206. Sirard, C., de la Pompa, J. L., Elia, A., Itie, A., Mirtsos, C., Cheung, A., Hahn, S., Wakeham, A., Schwartz, L., Kern, S. E., Rossant, J., and Mak, T. W. (1998). The tumor suppressor gene Smad4/DPC4 is required for gastrulation and later for anterior development of the mouse embryo. Genes Dev. 12, 107–119. Slusarski, D. C., Yang-Snyder, J., Busa, W. B., and Moon, R. T. (1997). Modulation of embryonic intracellular Ca2+ signaling by Wnt-5A. Dev. Biol. 182, 114–120. Smith, J. C. (1995). Mesoderm-inducing factors and mesodermal patterning. Curr. Opin. Cell Biol. 7, 856–861. Smith, W. C., and Harland, R. M. (1991). Injected Xwnt-8 RNA acts early in Xenopus embryos to promote formation of a vegetal dorsalizing center. Cell 67, 753–765. Smith, J. C., Price, B. M., Green, J. B., Weigel, D., and Herrmann, B. G. (1991). Expression of a Xenopus homolog of Brachyury (T ) is an immediate-early response to mesoderm induction. Cell 67, 79–87. Smith, W. C., Knecht, A. K., Wu, M., and Harland, R. M. (1993). Secreted noggin protein mimics the Spemann organizer in dorsalizing Xenopus mesoderm. Nature 361, 547–549. Sokol, S. Y. (1996). Analysis of Dishevelled signalling pathways during Xenopus development. Curr. Biol. 6, 1456–1467. Sokol, S. Y. (1999). Wnt signaling and dorso-ventral axis specification in vertebrates. Curr. Opin. Genet. Dev. 9, 405–410. Sokol, S., Christian, J. L., Moon, R. T., and Melton, D. A. (1991). Injected Wnt RNA induces a complete body axis in Xenopus embryos. Cell 67, 741–752. Spemann, H. (1927). Neue Arbeiten u¨ ber Organisatoren in der tierischen Entwicklung. Naturwissenschaften 15, 946–951. ¨ Spemann, H., and Mangold, H. (1924). Uber Induktion von Embryonalanlagen durch Implantation artfremder Organisatoren. Roux’ Arch. f Entw. mech. 100, 599–638. Stambolic, V., Ruel, L., and Woodgett, J. R. (1996). Lithium inhibits glycogen synthase kinase-3 activity and mimics wingless signalling in intact cells. Curr. Biol. 6, 1664–1668. Steinbeisser, H., Fainsod, A., Niehrs, C., Sasai, Y., and De Robertis, E. M. (1995). The role of gsc and BMP-4 in dorsal-ventral patterning of the marginal zone in Xenopus: a loss-of-function study using antisense RNA. EMBO J. 14, 5230–5243. Stennard, F., Carnac, G., and Gurdon, J. B. (1996). The Xenopus T-box gene, Antipodean, encodes a vegetally localised maternal mRNA and can trigger mesoderm formation. Development 122, 4179–4188. Stennard, F., Ryan, K., and Gurdon, J. B. (1997). Markers of vertebrate mesoderm induction. Curr. Opin. Genet. Dev. 7, 620–627. Stennard, F., Zorn, A. M., Ryan, K., Garrett, N., and Gurdon, J. B. (1999). Differential expression of VegT and Antipodean protein isoforms in Xenopus. Mech. Dev. 86, 87–98. Sumanas, S., Strege, P., Heasman, J., and Ekker, S. C. (2000). The putative wnt receptor Xenopus

1. Early Xenopus Development

65

frizzled-7 functions upstream of beta-catenin in vertebrate dorsoventral mesoderm patterning. Development 127, 1981–1990. Sun, B. I., Bush, S. M., Collins-Racie, L. A., LaVallie, E. R., DiBlasio-Smith, E. A., Wolfman, N. M., McCoy, J. M., and Sive, H. L. (1999). derriere: a TGF-beta family member required for posterior development in Xenopus. Development 126, 1467–1482. Suzuki, A., Chang, C., Yingling, J. M., Wang, X. F., and Hemmati-Brivanlou, A., (1997a). Smad5 induces ventral fates in Xenopus embryo. Dev. Biol. 184, 402–405. Suzuki, A., Ueno, N., and Hemmati-Brivanlou, A. (1997b). Xenopus msx1 mediates epidermal induction and neural inhibition by BMP4. Development 124, 3037–3044. Swalla, B. J., and Jeffery, W. R. (1995). A maternal RNA localized in the yellow crescent is segregated to the larval muscle cells during ascidian development. Dev. Biol. 170, 353–364. Swalla, B. J., and Jeffery, W. R. (1996). PCNA mRNA has a 3’UTR antisense to yellow crescent RNA and is localized in ascidian eggs and embryos. Dev. Biol. 178, 23–34. Tada, M., and Smith, J. C. (2000). Xwnt11 is a target of Xenopus Brachyury: regulation of gastrulation movements via Dishevelled, but not through the canonical Wnt pathway. Development 127, 2227–2238. Tada, M., Casey, E. S., Fairclough, L., and Smith, J. C. (1998). Bix1, a direct target of Xenopus T-box genes, causes formation of ventral mesoderm and endoderm. Development 125, 3997–4006. Taira, M., Otani, H., Saint-Jeannet, J. P., and Dawid, I. B. (1994). Role of the LIM class homeodomain protein Xlim-1 in neural and muscle induction by the Spemann organizer in Xenopus. Nature 372, 677–679. Takasaki, H., and Konishi, H. (1989). Dorsal blastomeres in the equatorial region of the 32-cell Xenopus embryo autonomously produce progeny committed to the organizer. Dev. Growth Diff. 31, 147–156. Tashiro, K., Yamada, R., Asano, M., Hashimoto, M., Muramatsu, M., and Shiokawa, K. (1991). Expression of mRNA for activin-binding protein (follistatin) during early Embryonic development of Xenopus laevis. Biochem. Biophys. Res. Commun. 174, 1022–1027. Thomas, P., and Beddington, R. (1996). Anterior primitive endoderm may be responsible for patterning the anterior neural plate in the mouse embryo. Curr. Biol. 6, 1487–1496. Thomas, P. Q., Brown, A., and Beddington, R. S. (1998). Hex: a homeobox gene revealing periimplantation asymmetry in the mouse embryo and an early transient marker of endothelial cell precursors. Development 125, 85–94. Thomsen, G. H. (1996). Xenopus mothers against decapentaplegic is an embryonic ventralizing agent that acts downstream of the BMP-2/4 receptor. Development 122, 2359–2366. Thomsen, G. H., and Melton, D. A. (1993). Processed Vg1 protein is an axial mesoderm inducer in Xenopus. Cell 74, 433–441. Torres, M. A., Eldar-Finkelman, H., Krebs, E. G., and Moon, R. T. (1999). Regulation of ribosomal S6 protein kinase-p90(rsk), glycogen synthase kinase 3, and beta-catenin in early Xenopus development. Mol. Cell Biol. 19, 1427–1437. Umbhauer, M., Marshall, C. J., Mason, C. S., Old, R. W., and Smith, J. C. (1995). Mesoderm induction in Xenopus caused by activation of MAP kinase. Nature 376, 58–62. van de Wetering, M., Cavallo, R., Dooijes, D., van Beest, M., van Es, J., Loureiro, J., Ypma, A., Hursh, D., Jones, T., Bejsovec, A., Peifer, M., Mortin, M., and Clevers, H. (1997). Armadillo coactivates transcription driven by the product of the Drosophila segment polarity gene dTCF. Cell 88, 789– 799. van Houte, L., van Oers, A., van de Wetering, M., Dooijes, D., Kaptein, R., and Clevers, H. (1993). The sequence-specific high mobility group 1 box of TCF-1 adopts a predominantly alpha-helical conformation in solution. J. Biol. Chem. 268, 18083–18087. Varlet, I., Collignon, J., and Robertson, E. J. (1997). nodal expression in the primitive endoderm is required for specification of the anterior axis during mouse gastrulation. Development 124, 1033–1044. Vincent, J. P., and Gerhart, J. C. (1987). Subcortical rotation in Xenopus eggs: an early step in embryonic axis specification. Dev. Biol. 123, 526-539.

66

Agnes P. Chan and Laurence D. Etkin

Vincent, J. P., Oster, G. F., and Gerhart, J. C. (1986). Kinetics of gray crescent formation in Xenopus eggs: the displacement of subcortical cytoplasm relative to the egg surface. Dev. Biol. 113, 484–500. Vleminckx, K., Wong, E., Guger, K., Rubinfeld, B., Polakis, P., and Gumbiner, B. M. (1997). Adenomatous polyposis coli tumor suppressor protein has signaling activity in Xenopus laevis embryos resulting in the induction of an ectopic dorsoanterior axis. J. Cell Biol. 136, 411–420. Vodicka, M. A., and Gerhart, J. C. (1995). Blastomere derivation and domains of gene expression in the Spemann Organizer of Xenopus laevis. Development 121, 3505–3518. Waldrip, W. R., Bikoff, E. K., Hoodless, P. A., Wrana, J. L., and Robertson, E. J. (1998). Smad2 signaling in extraembryonic tissues determines anterior–posterior polarity of the early mouse embryo. Cell 92, 797–808. Waltzer, L., and Bienz, M. (1998). Drosophila CBP represses the transcription factor TCF to antagonize Wingless signalling. Nature 395, 521–525. Wang, S., Krinks, M., Lin, K., Luyten, F. P., and Moos, M., Jr. (1997). Frzb, a secreted protein expressed in the Spemann organizer, binds and inhibits Wnt-8. Cell 88, 757–766. Wardle, F. C., Angerer, L. M., Angerer, R. C., and Dale, L. (1999a). Regulation of BMP signaling by the BMP1/TLD-related metalloprotease, SpAN. Dev. Biol. 206, 63–72. Wardle, F. C., Welch, J. V., and Dale, L. (1999b). Bone morphogenetic protein 1 regulates dorsal– ventral patterning in early Xenopus embryos by degrading chordin, a BMP4 antagonist. Mech. Dev. 86, 75–85. Watabe, T., Kim, S., Candia, A., Rothbacher, U., Hashimoto, C., Inoue, K., and Cho, K. W. (1995). Molecular mechanisms of Spemann’s organizer formation: conserved growth factor synergy between Xenopus and mouse. Genes Dev. 9, 3038–3050. Watanabe, M., and Whitman, M. (1999). FAST-1 is a key maternal effector of mesoderm inducers in the early Xenopus embryo. Development 126, 5621–5634. Weber, R. J., Pedersen, R. A., Wianny, F., Evans, M. J., and Zernicka-Goetz, M. (1999). Polarity of the mouse embryo is anticipated before implantation. Development 126, 5591–5598. Weinstein, M., Yang, X., Li, C., Xu, X., Gotay, J., and Deng, C. X. (1998). Failure of egg cylinder elongation and mesoderm induction in mouse embryos lacking the tumor suppressor smad2. Proc. Natl. Acad. Sci. USA 95, 9378–9383. Wells, J. M., and Melton, D. A. (1999). Vertebrate endoderm development. Annu. Rev. Cell Dev. Biol. 15, 393–410. Wessely, O., and De Robertis, E. M. (2000). The Xenopus homologue of Bicaudal-C is a localized maternal mRNA that can induce endoderm formation. Development 127, 2053–2062. Wikramanayake, A. H., Huang, L., and Klein, W. H. (1998). beta-Catenin is essential for patterning the maternally specified animal–vegetal axis in the sea urchin embryo. Proc. Natl. Acad. Sci. USA 95, 9343–9348. Willert, K., Brink, M., Wodarz, A., Varmus, H., and Nusse, R. (1997). Casein kinase 2 associates with and phosphorylates dishevelled. EMBO J. 16, 3089–3096. Wilson, P. A., and Hemmati-Brivanlou, A. (1995). Induction of epidermis and inhibition of neural fate by Bmp-4. Nature 376, 331–333. Wilson, P. A., Lagna, G., Suzuki, A., and Hemmati-Brivanlou, A. (1997). Concentration-dependent patterning of the Xenopus ectoderm by BMP4 and its signal transducer Smad1. Development 124, 3177–3184. Wodarz, A., and Nusse, R. (1998). Mechanisms of Wnt signaling in development. Annu. Rev. Cell Dev. Biol. 14, 59–88. Wolda, S. L., Moody, C. J., and Moon, R. T. (1993). Overlapping expression of Xwnt-3A and Xwnt-1 in neural tissue of Xenopus laevis embryos. Dev. Biol. 155, 46–57. Wright, C. V., Schnegelsberg, P., and De Robertis, E. M. (1989). XlHbox 8: a novel Xenopus homeo protein restricted to a narrow band of endoderm. Development 105, 787–794. Wylie, C., Kofron, M., Payne, C., Anderson, R., Hosobuchi, M., Joseph, E., and Heasman, J. (1996). Maternal beta-catenin establishes a ‘dorsal signal’ in early Xenopus embryos. Development 122, 2987–2996.

1. Early Xenopus Development

67

Xu, Q., D’Amore, P. A., and Sokol, S. Y. (1998). Functional and biochemical interactions of Wnts with FrzA, a secreted Wnt antagonist. Development 125, 4767–4776. Yamada, G., Mansouri, A., Torres, M., Stuart, E. T., Blum, M., Schultz, M., De Robertis, E. M., and Gruss, P. (1995). Targeted mutation of the murine goosecoid gene results in craniofacial defects and neonatal death. Development 121, 2917–2922. Yamanaka, Y., Mizuno, T., Sasai, Y., Kishi, M., Takeda, H., Kim, C. H., Hibi, M., and Hirano, T. (1998). A novel homeobox gene, dharma, can induce the organizer in a non-cell-autonomous manner. Genes Dev. 12, 2345–2353. Yanagawa, S., van Leeuwen, F., Wodarz, A., Klingensmith, J., and Nusse, R. (1995). The dishevelled protein is modified by wingless signaling in Drosophila. Genes Dev. 9, 1087–1097. Yasuo, H., and Lemaire, P. (1999). A two-step model for the fate determination of presumptive endodermal blastomeres in Xenopus embryos. Curr. Biol. 9, 869–879. Yost, C., Torres, M., Miller, J. R., Huang, E., Kimelman, D., and Moon, R. T. (1996). The axis-inducing activity, stability, and subcellular distribution of beta-catenin is regulated in Xenopus embryos by glycogen synthase kinase 3. Genes Dev. 10, 1443–1454. Yost, C., Farr, G. H., III, Pierce, S. B., Ferkey, D. M., Chen, M. M., and Kimelman, D. (1998). GBP, an inhibitor of GSK-3, is implicated in Xenopus development and oncogenesis. Cell 93, 1031–1041. Yuge, M., Kobayakawa, Y., Fujisue, M., and Yamana, K. (1990). A cytoplasmic determinant for dorsal axis formation in an early embryo of Xenopus laevis. Development 110, 1051–1056. Zaraisky, A. G., Ecochard, V., Kazanskaya, O. V., Lukyanov, S. A., Fesenko, I. V., and Duprat, A. M. (1995). The homeobox-containing gene XANF-1 may control development of the Spemann organizer. Development 121, 3839–3847. Zeng, L., Fagotto, F., Zhang, T., Hsu, W., Vasicek, T. J., Perry, W. L., III, Lee, J. J., Tilghman, S. M., Gumbiner, B. M., and Costantini, F. (1997). The mouse Fused locus encodes Axin, an inhibitor of the Wnt signaling pathway that regulates embryonic axis formation. Cell 90, 181–192. Zernicka-Goetz, M. (1998). Fertile offspring derived from mammalian eggs lacking either animal or vegetal poles. Development 125, 4803–4808. Zhang, J., and King, M. L. (1996). Xenopus VegT RNA is localized to the vegetal cortex during oogenesis and encodes a novel T-box transcription factor involved in mesodermal patterning. Development 122, 4119–4129. Zhang, J., Houston, D. W., King, M. L., Payne, C., Wylie, C., and Heasman, J. (1998a). The role of maternal VegT in establishing the primary germ layers in Xenopus embryos. Cell 94, 515–524. Zhang, J., Talbot, W. S., and Schier, A. F. (1998b). Positional cloning identifies zebrafish one-eyed pinhead as a permissive EGF-related ligand required during gastrulation. Cell 92, 241–251. Zhu, H., Kavsak, P., Abdollah, S., Wrana, J. L., and Thomsen, G. H. (1999a). A SMAD ubiquitin ligase targets the BMP pathway and affects embryonic pattern formation. Nature 400, 687–693. Zhu, L., Belo, J. A., De Robertis, E. M., and Stern, C. D. (1999b). Goosecoid regulates the neural inducing strength of the mouse node. Dev. Biol. 216, 276–281. Zimmerman, L. B., De Jesus-Escobar, J. M., and Harland, R. M. (1996). The Spemann organizer signal noggin binds and inactivates bone morphogenetic protein 4. Cell 86, 599–606. Zoltewicz, J. S., and Gerhart, J. C. (1997). The Spemann organizer of Xenopus is patterned along its anteroposterior axis at the earliest gastrula stage. Dev. Biol. 192, 482–491. Zorn, A. M., Barish, G. D., Williams, B. O., Lavender, P., Klymkowsky, M. W., and Varmus, H. E. (1999a). Regulation of Wnt signaling by Sox proteins: XSox17 alpha/beta and XSox3 physically interact with beta-catenin. Mol. Cell 4, 487–498. Zorn, A. M., Butler, K., and Gurdon, J. B. (1996b). Anterior endomesoderm specification in Xenopus by Wnt/beta-catenin and TGF-beta signalling pathways. Dev. Biol. 209, 282–297.

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2 Transcriptional Programs Regulating Vascular Smooth Muscle Cell Development and Differentiation Michael S. Parmacek Department of Medicine University of Pennsylvania Philadelphia, Pennsylvania 19104

I. Introduction II. Embryology of the Vascular Smooth Muscle Cell Lineage(s) A. Mesodermally Derived SMCs B. Neural Crest-Derived SMCs C. Coronary Vasculature D. Vascular Smooth Muscle Cell Heterogeneity III. Commitment to the Smooth Muscle Cell Lineage IV. Transcriptional Control of Vascular Smooth Muscle Cell Differentiation V. SRF: A Nuclear Sensor Regulating Growth and Differentiation VI. Modulation of VSMC Phenotype VII. Conclusions and Future Challenges References

I. Introduction Vascular smooth muscle cells (VSMCs) modulate arterial tone through expression of a unique set of genes encoding SMC lineage-restricted myofibrillar and cytoskeletal proteins and intracellular enzymes. During postnatal development, VSMCs, located in the tunica media, arrest in the G0/G1 phase of the cell cycle and express a set of genes encoding SMC-specific contractile proteins (for review see Owens, 1998). However, in response to vessel wall injury, VSMCs reenter the cell cycle, migrate to the arterial intima, and express a set of genes encoding extracellular matrix and adhesion molecules. The modulation of SMC phenotype has been implicated in the pathogenesis of vascular proliferative syndromes including Correspondence should be addressed to: Michael S. Parmacek, M.D., University of Pennsylvania, Department of Medicine, Cardiovascular Division, 9123 Founders Pavilion, 3400 Spruce Street, Philadelphia, PA 19104. Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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atherosclerosis, restenosis following balloon angioplasty, and transplant arteriopathy (Newby and Zaltsman, 1999; Ross, 1999; Schwartz, 1997; Schwartz and Murry, 1998). Interestingly, the phenotype of VSMCs following arterial injury closely resembles that observed during embryonic and neonatal angiogenesis (Shanahan and Weissberg, 1997). Therefore, understanding the molecular mechanisms that control SMC development and differentiation may provide fundamental insights into pathological processes including atherosclerosis. Despite the important roles VSMCs play in the cardiovascular system, relatively little is currently understood about the molecular mechanisms that control vascular smooth muscle cell development and differentiation. This is due, in part, to the complex, and poorly understood, embryological origins of VSMCs and the lack of definite VSMC markers. In other cell lineages, characterization of the transcriptional programs that regulate cell lineage–specific gene expression has provided fundamental insights into the molecular mechanisms that control cell fate decisions (for review see Orkin, 1992; Tapscott and Weintraub, 1991; Weintraub et al., 1991). For example, identification and characterization of the basic-helixloop-helix (bHLH) family of myogenic regulatory factors led to understanding of skeletal muscle specification and differentiation (Lassar et al., 1989). This, in turn, led to identification of novel upstream and downstream signaling pathways that control growth and adaptation of skeletal muscle cells (Chin et al., 1998). In this review, I have attempted to critically analyze our current understanding of the transcriptional programs that control SMC development and differentiation. Until recently, this field of investigation was limited, to some extent, by in vitro model systems that failed to fully recapitulate the complex and dynamic milieu that the VSMC experiences in the vessel wall (Chamley-Campbell et al., 1979; Ross, 1971). However, with the advent of new techniques, including cell fate mapping, viral-mediated gene transfer, production of transgenic mice, and creation of targeted gene mutations, understanding of SMC biology has increased tremendously. Because this is a relatively new field of investigation, this review may raise more questions than it answers. As such, it is my hope that it will stimulate new avenues of investigation in this important and exciting field. The readers are also referred to several excellent reviews in closely related areas (Campbell et al., 1988; Kirby and Waldo, 1995; Owens, 1998; Schwartz and Liaw, 1993; Shanahan and Weissberg, 1997; Sobue et al., 1999; Treisman, 1995).

II. Embryology of the Vascular Smooth Muscle Cell Lineage(s) A. Mesodermally Derived SMCs As shown in (Fig. 1, see color insert), vascular smooth muscle cells arise from two distinct embryological origins: lateral mesenchyme (mesoderm) and neural crest (ectoderm). The majority of VSMCs arise from lateral mesoderm-derived

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mesenchyme (Hungerford et al., 1996; McHugh, 1995; Miano et al., 1994). During embryonic angiogenesis, local mesenchyme is recruited to the primary capillary network and through poorly understood inductive interactions differentiates into pericytes and VSMCs. In the mouse embryo, VSMCs are first recognized in the dorsal aorta at embryonic day (E)9.5 (McHugh, 1995). Embryonic VSMCs proliferate and express low, but detectable, levels of SM-specific markers, including smooth muscle myosin heavy chain (SM-MHC), SM22␣, and calponin (Li et al., 1996a; Miano et al., 1994, 1996; Samaha et al., 1996). In the early neonatal period, VSMC proliferation decreases and vessel wall growth and stabilization continues through SMC hypertrophy and elaboration of extracellular matrix (Gerrity and Cliff, 1975; Olivetti et al., 1980). During this period, SMCs express high levels of genes encoding SMC contractile proteins as well as extraceullar matrix (Adam et al., 1993; Shanahan et al., 1998; Shanahan and Weissberg, 1997). Subsequently, in the mouse, at approximately 12 weeks of age, vessel wall growth slows and expression of genes encoding SMC-specific contractile proteins is downregulated to levels required to sustain contractile function. B. Neural Crest-Derived SMCs A second embryologically distinct population of VSMCs, termed ectomesenchymal SMCs, is derived from the cardiac neural crest (Kirby et al., 1983; Kirby and Waldo, 1995). These cells arise in the neural folds adjacent to the midotic placode and rostral somites and migrate to, and populate, the caudal three pharyngeal arches (Miyagawa-Tomita et al., 1991). They contribute eventually to the cardiac outflow tract and the tunica media of the proximal aorta, pulmonary trunk, brachiocephalic arteries, common carotid artery, and lateral subclavian artery (Duband et al., 1992; Hughes, 1943; Lelievre and Ledouarin, 1975; Miyagawa-Tomita et al., 1991; Romanoff, 1960; Rosenquist and Beall, 1990). It remains to be determined whether common, overlapping, or distinct molecular mechanisms regulate commitment of SMCs from mesenchyme and neural crest, respectively. Because they possess the capacity to differentiate from neural crest into SMCs in vitro, Monc1 cells may prove to be a valuable experimental reagent to examine the molecular mechanisms that regulate differentiation of VSMCs from the cardiac neural crest (Jain et al., 1998; Shah et al., 1996; Stemple and Anderson, 1992). Abnormal patterning of the cardiac neural crest is associated with a number of congenital cardiovascular malformations (for review, see Kirby and Waldo, 1990). Ablation of the cardiac neural crest in chicks results in conotruncal defects, including persistent truncus arteriosus, transposition of the great arteries, interrupted aortic arch, and coarctation of the aorta (Kirby and Waldo, 1990; Nishibatake et al., 1987). In humans, the relationship between conotruncal defects and neural crest is also compelling (Epstein, 1996). The DiGeorge syndrome features various abnormalities in neural crest–derived structures, including the aortic arch, thymus, parathyroid gland, and craniofacial structures. The DiGeorge syndrome

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locus has been mapped to a critical region of human chromosome 22q11 and several candidate genes have been identified (Lindsay et al., 1999; Yamagishi et al., 1999). The underlying genetic basis of other conotruncal defects are not well understood. One gene that has been implicated in neural crest patterning is the paired homeodomain factor, Pax3 (for review see Epstein, 1996). Splotch mice, harboring a null mutation in the gene encoding Pax3, exhibit defective septation of the truncus arteriosis and rarely other outflow tract defects (Epstein et al., 1993; Franz, 1989).

C. Coronary Vasculature Until recently, it was assumed that the coronary vasculature was established by sprouting from the aortic root and ingrowth into the heart (Aikawa and Kawano, 1982; Hutchins et al., 1988). However, elegant retroviral and vital dye tagging (Mikawa and Fischman, 1992; Mikawa and Gourdie, 1996) and neural crest ablation studies (Kirby and Waldo, 1995) refute this model. These studies demonstrate conclusively that the majority of VSMCs populating the coronary vasculature arise from the subepicardial mesenchyme that is derived from the proepicardial organ (Dettman et al., 1998). At mid-gestation, progenitor populations from the proepicardial organ migrate over the epicardial surface of the tubular heart, invade the supepicardium, and undergo epithelial to mesenchymal transformation into mature SMCs (Mikawa and Gourdie, 1996). The majority of experimental evidence suggests that coronary SMCs differentiate only after migration over the epicardial surface when they come in direct contact with endothelial cells (Mikawa and Fischman, 1992; Mikawa and Gourdie, 1996). This process appears to be dependent on the expression of the MADS box transcription factor, SRF (Landerholm et al., 1999). It is important to recognize that VSMCs populating the coronary arteries and the proximal aorta represent embryologically distinct populations.

D. Vascular Smooth Muscle Cell Heterogeneity There is some evidence that blood vessel ontogeny may play a role in the varying susceptibility of different component of the circulatory system to atherosclerosis (Hood and Rosenquist, 1992; Topouzis et al., 1992). VSMCs derived from lateral mesoderm and neural crest appear morphologically identical and express nearly identical levels of smooth muscle cell markers both in vitro and in vivo (Topouzis and Majesky, 1996). However, mesoderm-derived VSMCs cannot rescue the outflow tract defects observed in neural crest-ablated chicken embryos (Kirby, 1988; Rosenquist and Beall, 1990). Moreover, neural crest- and mesodermally derived VSMCs exhibit distinct growth and receptor-mediated transcriptional responses to the growth factor TGF-␤ (Topouzis and Majesky, 1996). These data suggest that embryologically distinct subpopulations of smooth muscle cells

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are not functionally equivalent. Whether these differences contribute to the heterogeneity of smooth muscle cells in response to vessel wall injury remains to be determined.

III. Commitment to the Smooth Muscle Cell Lineage The molecular mechanisms leading to commitment to the smooth muscle cell fate from undifferentiated mesenchyme and pleuripotent neural crest cells are not well understood. Environmental factors implicated in this process include epithelial– mesenchymal interactions (Cunha et al., 1992; Duluc et al., 1994; Hirschi et al., 1998; Yang et al., 1998b), growth factors (Bostrom et al., 1996; Hirschi et al., 1998; Holycross et al., 1992; Zhou et al., 1996), and hormones (Blank et al., 1995). In vitro heterotypic coculture experiments demonstrate that endothelial cells recruit undifferentiated mesenchymal cells via a PDGF-mediated signaling pathway (Hirschi et al., 1998). Consistent with this finding, PDGF-␣ receptor deficient Patch mice exhibit reduced numbers of VSMCs (Schatteman et al., 1995). Similarly, PDGF-B null mutant mice have reduced numbers of pericytes (Lindahl et al., 1997). In addition, SMC specification is regulated at least in part through a TGF␤ signaling pathway (Hirschi et al., 1998; Orlani et al., 1994). Antibodies that recognize TGF-␤ block the in vitro differentiation of SMCs from mesenchymally derived cell lines (Hirschi et al., 1998). Consistent with this finding, TGF-␤1 null mice exhibit abnormal yolk sac vasculogenesis (Dickson et al., 1995). Cytoskeletal organization, or cell shape, independently influences commitment and differentiation of SMCs (Schuger et al., 1997; Yang et al., 1998b, 1999). Expression of genes encoding SMC markers is preceded by a change in the shape of SMC precursors from round to elongated (Roman and McDonald, 1992). Undifferentiated mesenchymal cells spontaneously differentiate into SMCs only after they are induced to elongate (Yang et al., 1999). Of note, TGF-␤1 stimulates synthesis of SMC proteins in elongated cells, but is ineffective in round cells (Yang et al., 1999). Taken together, these data suggest a model wherein integrin-mediated reorganization of the cytoskeleton plays a permissive role in commitment of undifferentiated mesenchyme to the SMC lineage. Subsequently, growth factor–mediated signals (including TGF-␤1) stimulate differentiation of SMCs and expression of genes encoding SMC contractile proteins. Skeletal myogenesis is controlled by a family of dominant-acting transcription factors, the bHLH family of muscle regulatory factors, that possess the capacity to irreversibly commit an undifferentiated mesodermal cell to the skeletal muscle cell fate (Lassar et al., 1989). Despite intense investigation, SMC lineage-specific transcription factors and/or determination factors have not been identified. Some evidence suggests that the vertebrate MADS box transcription factor, MEF2, may be involved in regulating specification of SMCs (Belaguli et al., 1997; Browning et al., 1998; Firulli et al., 1996; Morrisey et al., 1996; Suzuki et al., 1995). Flies

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(Drosophila melanogaster) harboring a mutation in the single Drosophila MEF2 factor, D-MEF2, fail to develop visceral, cardiac, or skeletal muscle (Lilly et al., 1995). However, SMCs are observed in mice harboring a targeted mutation in the gene encoding MEF2C (though they do exhibit abnormal angiogenesis) (Lin et al., 1998). Because four MEF2 isoforms encoded by separate genes are expressed in mice (Firulli et al., 1996), it remains theoretically possible that MEF2 may play a role in specification of SMCs. However, it is noteworthy that MEF2 functions downstream of bHLH transcription factors in the transcriptional cascade, leading to skeletal muscle differentiation (for review, see Black and Olson, 1998; Reecy et al., 1999). Moreover, MEF2 isoforms are upregulated in response to vessel wall injury concomitant with downregulation of genes encoding SMC-specific contractile proteins (Firulli et al., 1996). Thus, it is relatively unlikely that MEF2 regulates commitment to the SMC fate. In any case, further studies are required to elucidate the transcriptional program(s) leading to specification of the SMC lineage.

IV. Transcriptional Control of Vascular Smooth Muscle Cell Differentiation One approach to understanding the molecular mechanisms that regulate SMC differentiation is to identify the transcriptional regulatory elements and transcription factors that regulate SMC-specific gene expression. This approach has fundamentally increased our understanding of skeletal and cardiac myocyte-specific gene expression and differentiation. Our group, and others, have focused on examining the transcriptional programs that control expression of SM22␣ because of its SMC lineage-restricted pattern of expression (Kim et al., 1997; Li et al., 1996a,b; Moessler et al., 1996; Osbourn et al., 1995; Solway et al., 1995). SM22␣ is expressed exclusively in vascular and visceral SMCs during postnatal development (Li et al., 1996; Solway et al., 1995). Like most other SMC markers, it is also transiently expressed in embryonic cardiac and skeletal muscle. The SM22␣ gene encodes a 22-kDa cytoskeletal protein with structural homology to the vertebrate thin filament myofibrillar protein calponin. Consistent with the pattern of genes encoding SMC contractile proteins, SM22␣ is downregulated when primary aortic SMCs modulate their phenotype in vitro and within atherosclerotic plaques (Shanahan et al., 1994). As shown in (Fig. 2, see color insert), the SM22␣ promoter restricts expression of a lacZ reporter gene to arterial smooth muscle cells, the myotomal component of the somites and the bulbocordis (future right ventricle) in the mouse embryo (Kim et al., 1997; Li et al., 1996b; Moessler et al., 1996). Surprisingly, in contrast to the endogenous pattern of SM22␣ gene expression, ␤-galactosidase expression is not observed in coronary arterial, venous, or visceral smooth muscle cells. In fact, up to

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Figure 3 Schematic representation of the murine SM22␣ promoter. DNase I footprint analyses performed with VSMC nuclear extracts revealed six nuclear protein binding sites, designated SME1-6, respectively, in the 441-bp SM22␣ promoter (clear rectangles). EMSAs revealed that SME-1 and SME4 bound specifically to SRF (shown as circles). Other transcription factors that bind specifically to the SM22␣ promoter, including Sp1, YY1, ATF-1, and CREB-1, are also shown. Unidentified nuclear protein complexes are also shown.

5 kb of SM22␣ 5′ flanking sequence is not sufficient to direct transgene expression to venous or visceral SMCs (Kim et al., 1997). Similarly, a transgene under the transcriptional control of the SM myosin heavy chain promoter was expressed selectively throughout the vasculature (Madsen et al., 1998). In contrast, the SM ␣-actin promoter and intragenic transcriptional enhancer restricted gene expression to visceral SMCs and almost all vascular beds, including the coronary, mesenteric, and renal vasculature in transgenic mice (Mack and Owens, 1999). Taken together, these data suggest that distinct transcriptional programs may distinguish previously unrecognized tissue-restricted smooth muscle cell sublineages. Analysis of the murine SM22␣ promoter revealed six nuclear protein binding sites (designated smooth muscle element (SME) 1–6, respectively), including two consensus SREs or CArG boxes (Kim et al., 1997) (Fig. 3). The MADS box transcription factor, SRF (for review see Treisman, 1995), binds specifically to CArG boxes conforming to the consensus sequence CC(A/T)6 GG. Mutations of the SM22␣ promoter that abolish binding of SRF totally abolish its activity in transgenic mice (Kim et al., 1997). Moreover, four copies of either SM22␣ CArG box linked to a minimal promoter is sufficient to direct SM22␣ gene expression to arterial SMCs in transgenic mice (Kim et al., 1997; Solway et al., 1995). Most importantly, CArG boxes have been identified in multiple other SMC-specific transcriptional regulatory elements, including the rat smooth muscle ␣-actin intragenic transcriptional enhancer (Mack and Owens, 1999), the rat smooth muscle myosin heavy chain promoter and intragenic transcriptional enhancer (Madsen et al., 1998; White and Low, 1996), the mouse smooth muscle ␥ -actin promoter (Qian et al., 1996), the rabbit telokin promoter (Herring and Smith, 1997), and the chicken caldesmon promoter (Yano et al., 1995) (see Table I). Taken together, these data demonstrate that SRF plays a critical role in regulating SMC-specific gene expression. Moreover, as each CArG box-dependent SMC gene is downregulated when VSMCs modulate their phenotype in response to vessel wall injury, these data suggest that SRF plays an important role in maintenance of the contractile SMC phenotype.

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Table I CArG Box-Dependent SMC Transcriptional Regulatory Elements Elementa SM22␣ promoter (SME-1) SM22␣ promoter (SME-4) SM ␣-actin (intron 1) SM MyHC (intron 1)

CArG boxb CCATAAAAGG CCAAATATGG CCTAATTAGG

Telokin promoter Caldesmon promoter (CArG1) Caldesmon promoter (CArG2) ␤-Tropomyosin promoter

CCAAAAAAGG CCAAAAAAGG CCTAAAAAGG

Transgene expressionc Arterial SMCs (only) VSMCs, visceral SMCs Selective vascular beds (-pulm, coronary veins) Visceral SMCs (mosaic pattern) Visceral SMCs (? vascular and bronchial) N.D. N.D.

a Column

indicates the SMC-restricted transcriptional regulatory element. shows the nucleotide sequence of the CArG Box. c Column indicates the pattern of transgene expression. b Column

V. SRF: A Nuclear Sensor Regulating Growth and Differentiation As discussed earlier, the MADS box transcription factor, SRF, plays an important role in regulating SMC differentiation. Every smooth muscle cell lineage–restricted transcriptional regulatory element characterized to date includes a functionally important CArG element that binds specifically to SRF. In addition, SRF gene expression occurs coincident with SMC-specific gene expression in vitro and in vivo. Dominant negative SRF constructs inhibit the ex vivo differentiation of smooth muscle cells (Landerholm et al., 1999). However, it remains to be determined how a ubiquitously expressed transcription factor, such as SRF, activates genes in an SMC lineage–restricted fashion. Moreover, it remains unclear how a single transcription factor activates multiple SMC genes that are expressed in distinct temporal and spatial patterns during embryonic development and in response to vessel wall injury (Li et al., 1996a; Miano et al., 1994; Owens, 1995; Solway et al., 1995). The answers to these questions require further understanding of the lineage relationships of smooth muscle cells, as well as the molecular mechanisms regulating activity of SRF in vivo. SRF is a member of the ancient MADS box family of transcription factors and has been shown to regulate serum- and growth factor–responsive gene expression (for review, see Treisman, 1995a,b). The serum response element (SRE), or CArG box, was characterized originally as an essential element within the c-fos promoter that mediates immediate-early gene expression (Treisman, 1995b). However, it is now appreciated that SRF and CArG boxes also play an important role in regulating skeletal and cardiac-specific gene expression (Bergsma et al., 1986; Chow and

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Schwartz, 1990; Grichnik et al., 1988; Minty and Kedes, 1986; Muscat et al., 1992). SRF is a 508-amino acid protein that binds DNA as a dimer (Mohun et al., 1991). Its DNA-binding activity is mediated through an evolutionarily conserved 90-amino-acid MADS box domain (Mueller and Nordheim, 1991). The MADS box domain also mediates heterodimerization with other transcription factors, including homeodomain proteins that are expressed in a lineage-restricted fashion (Grueneberg et al., 1992; Vershon and Johnson, 1993). In the developing vertebrate embryo, SRF is expressed preferentially in cell lineages derived from embryonic mesoderm, including within the myotomal component of the somites, the embryonic heart, and within the medial layer of developing arteries and veins (Belaguli et al., 1997; Croissant et al., 1996). In addition, the gene is expressed, at low levels, in cell types derived from neuroectoderm (Yu et al., 1992). The capacity of SRF to activate genes encoding skeletal and cardiac-specific contractile proteins has been hypothesized to occur through its capacity to heterodimerize with other transcriptional activators, coactivators, and repressors (Reecy et al., 1999). For example, SRF physically associates with the cardiac lineage–restricted transcription factors Nkx2.5 and GATA4 and synergistically activates expression of the cardiac-specific genes ␣-cardiac actin and ANF (Chen et al., 1996; Durocher et al., 1996; Sepulveda et al., 1998). Similarly, SRF physically associates, and functionally synergizes, with members of the bHLH family of myogenic regulatory factors (Sartorelli et al., 1990). Several laboratories are examining the functional significance of SMC lineage–restricted transcription factors that physically associate with SRF. At least two distinct signaling pathways regulate SRF transcriptional activity in response to growth factor stimulation. The classical pathway requires the physical association of SRF with a ternary complex factor (TCF) (for review, see Treisman, 1994). TCFs are encoded by at least three related genes in the Ets family of transcription factors (Sap1, Sap2/Net/Erp, and Elk1) (Price et al., 1995). TCF functional activity is dependent upon specific protein–DNA and protein–protein (with SRF) interactions. To date, functionally important TCF- or Ets-binding sites have not been identified adjacent to CArG boxes in SMC-specific transcriptional regulatory elements. Of note, TCFs contain a transcriptional activation domain that is activated by the Ras/Raf/mitogen-activated protein (MAP) kinase signaling cascade. As Ras activates genes associated with cellular proliferation and suppresses myogenic differentiation (Konieczny and Emerson, 1987), it will be important to determine what role, if any, this signaling cascade plays in the developing vasculature and in vascular proliferative syndromes. An alternative novel cytoskeletal-SRF signaling pathway has been described that may explain how cytoskeletal signals are transduced to the SMC nucleus (Sotiropoulos et al., 1999). Treisman and colleagues demonstrated that SRF activation by LIM kinase-1 (LIMK-1) is dependent upon its ability to regulate cytoskeletal actin treadmilling. LIMK-1 phosphorylates the cytoskeletal protein cofilin, resulting in the stabilization of filamentous actin (Arber et al., 1998;

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Carlier et al., 1997; Lappalainen and Drubin, 1997; Yang et al., 1998a). As the Ras-like GTPase RhoA also regulates cytoskeletal organization (for review, see Van Aelst and D’Souza-Schorey, 1997) and activates SRF (Carnac et al., 1998; Hill et al., 1995), this represents a convergence point for serum- and LIMK-1-induced signaling to SRF (Sotiropoulos et al., 1999). Taken together, these data are consistent with a model wherein RhoA-dependent and independent cytoskeletal signaling activates SRF-dependent transcription. These findings raise the intriguing possibility that pharmacological agents that affect cytoskeletal organization may prove to be therapeutically efficacious in the treatment of vascular proliferative syndromes.

VI. Modulation of VSMC Phenotype A distinguishing feature of the SMC lineage is its capacity to reversibly modulate its phenotype during postnatal development (Bochaton-Pillat et al., 1996; ChamleyCampbell et al., 1979). The modulation of VSMC phenotype has been implicated in the pathogenesis of vascular proliferative syndromes, including atherosclerosis, restenosis following angioplasty, and transplant arteriopathy (for review, see Newby and Zaltsman, 1999; Ross, 1999; Schwartz, 1997; Schwartz and Murry, 1998). It is now recognized that the paradigm of “contractile” versus “synthetic” SMC phenotypes that evolved from the analysis of primary VSMCs in culture (Chamley-Campbell et al., 1979) represents extremes of a spectrum of phenotypes that may exist in the blood vessel wall (for review, see Shanahan and Weissberg, 1997). Many laboratories have observed extensive SMC phenotypic heterogeneity in the blood vessel wall and within atherosclerotic plaques. The molecular basis for this heterogeneity is not well understood, though lineage relationships, genetic factors, and local environmental cues have all been implicated. It has been postulated that the molecular mechanisms that control SMC differentiation during embryonic development are recapitulated during blood vessel repair and in vascular proliferative syndromes (Shanahan et al., 1994). This hypothesis is supported by the observation that neointimal SMCs and neonatal SMCs appear morphologically similar and express common sets of genes encoding extracellular matrix and some markers of differentiation (Bochaton-Pillat et al., 1996; Kim et al., 1994; Majesky et al., 1992). Of note, histopathological analyses have shown that VSMCs within atherosclerotic plaques display remarkable phenotypic heterogeneity (Shanahan et al., 1994). VSMCs within the “fibrous cap” of atherosclerotic plaque express high levels of some SMC-restricted contractile proteins, but not others (Shanahan et al., 1994). For example, SM22␣ is expressed at high levels in medial SMCs, is not expressed in intimal thickenings, but is expressed within fibrous caps (Shanahan et al., 1994). In contrast, the thin-filament myofibrillar protein calponin is not expressed in intimal SMCs within early or late fibrotic atherosclerotic plaques (Shanahan et al., 1994). These data demonstrate that the phenotypes

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of SMCs observed in vivo are far more complex than those observed in cell culture and emphasize the necessity for studying VSMCs in the blood vessel wall. An ongoing controversy in the field is whether neointimal SMCs arise from medial SMCs that modulate their phenotype or a subpopulation of “immature” SMCs present in the vessel media and/or adventitia (Shi et al., 1996). The former view is supported by the finding that adult VSMCs grown in culture possess the capacity to reversibly modulate their phenotype in response to a variety of extracellular stimuli (Bochaton-Pillat et al., 1996; Chamley-Campbell et al., 1979). The latter hypothesis is supported by the monclonality of intimal SMCs in atherosclerotic plaques (Benditt and Benditt, 1973; Murry et al., 1997). In addition, VSMC clones have been identified that maintain distinct phenotypes in vitro (Bochaton-Pillat et al., 1996; Lemire et al., 1996). These hypotheses are not mutually exclusive and can be reconciled. In fact, it is theoretically possible that in atherosclerotic plaques, neointimal SMCs arise from clonal proliferation of a subpopulation of cells in the vessel wall, whereas response to balloon injury occurs via activation of a large subset of quiescent contractile medial SMCs. This hypothesis is supported by the distinct histopathologies and chronologies of atherosclerotic and restenotic lesions, respectively. Several lines of evidence suggest that atheroslerosis results from an inflammatory response within the arterial wall that is mediated, in part, through activation of the transcription factor nuclear factor-kappa B (NF-␬B) (Bochaton-Pillat et al., 1996; Bourcier et al., 1997; Clesham et al., 1998; Dechend et al., 1999; Krzesz et al., 1999; Landry et al., 1997; Lemire et al., 1996; Ross, 1999; Selzman et al., 1999; Welch et al., 1998). NF-␬B was first identified as a transcription factor in activated lymphocytes (for review, see Baeuerle and Henkel, 1994). Activated NF-␬B is a heterodimeric complex consisting of the p50 and p65 (RelA) subunits. In the inactive state, the NF-␬B dimer is present in the cytosol bound to the inhibitory protein I␬B (Baeuerle and Baltimore, 1988). Upon cell stimulation, I␬B is degraded and NF-␬B is translocated to the nucleus, where it binds DNA and activates transcription. Following experimental balloon injury, a dramatic induction of p50, p65, and c-Rel is observed in the arterial wall with a concomitant reduction in the inhibitory peptides I␬B (Landry et al., 1997). Expression of NF-␬B-regulated genes, including VCAM-1, MCP-1, and iNOS, is observed in SMCs within 4 hr vascular injury (Landry et al., 1997). Moreover, activated NF-␬B has been localized within SMCs of atherosclerotic plaques (Bourcier et al., 1997). Taken together, these data suggest strongly that NF-␬B signaling is involved in the activation of quiescent VSMCs. Other candidate regulators of SMC phenotype have been proposed, though their function(s) in vivo remains to be elucidated. The zinc finger transcription factor, GATA6, is expressed at high levels in medial VSMCs, but not in immortalized SMC lines (Morrisey et al., 1996). GATA6 is rapidly downregulated in the rat carotid artery following experimental balloon injury (Toshiaki et al., 1999). Adenoviral-mediated delivery of GATA6 to the injured artery partially reversed

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alterations in SMC gene expression that occur in response to injury (Toshiaki et al., 1999). Similarly, expression of Gax, a diverged homeobox gene, is downregulated when VSMCs are induced to proliferate (Gorski et al., 1993). Conversely, MEF2 factors are not expressed in medial SMCs, but MEF2A, MEF2B, and MEF2D gene expression is upregulated in neointimal SMCs following vessel wall injury (Firulli et al., 1996). Improved tissue-specific gene targeting strategies should help define the role that these, and other, transcription factors play in modulating SMC phenotype.

VII. Conclusions and Future Challenges SMCs subserve multiple important functions, including modulation of arterial tone, regulation of airway resistance, and control of gastrointestinal and genitourinary motility. However, we currently understand very little about the molecular mechanisms that regulate SMC development and differentiation. In contrast to cardiac and skeletal myocytes, which upon differentiation express a stable set of genes encoding muscle-specific contractile proteins, SMCs retain the capacity to modulate their phenotype, reenter the cell cycle, and proliferate (Chamley-Campbell et al., 1979; Owens, 1989; Sobue et al., 1999). In the vascular system, phenotypic plasticity is required during primary angiogenesis and for reparative processes during postnatal development. In addition, the capacity of SMCs to modulate their phenotype has been implicated in the pathogenesis of vascular proliferative syndromes, including atherosclerosis and restenosis following balloon angioplasty (Newby and Zaltsman, 1999; Ross, 1999; Schwartz, 1997; Schwartz and Murry, 1998). VSMCs arise from two embryological origins, lateral mesoderm and neural crest. However, it remains uncertain whether SMCs represent a single cell lineage that adapts its phenotype to local environmental cues, or whether SMCs are genetically preprogrammed to distinct tissue-restricted sublineages. Subtle differences in phenotype are observed between mesenchymal and neural crest–derived VSMCs in the proximal aorta (Landerholm et al., 1999). Transcriptional regulatory elements have been identified that restrict gene expression to tissue-restricted subsets of SMCs in transgenic mice (Kim et al., 1997; Li et al., 1996a,b; Madsen et al., 1998; Moessler et al., 1996). In addition, lineage-restricted transcription factors have been identified that are expressed only in tissue restricted subsets of SMCs (Kim et al., 1997; Morrisey et al., 1996). Thus, it remains possible that lineage-restricted subsets of SMCs are programmed at the level of transcription. Identification of transcription factors that regulate SMC differentiation and characterization of the signaling pathways that activate these factors should help define the lineage relationships of SMCs. Compelling evidence suggests that the MADS box transcription factor SRF plays an important role in regulating SMC-specific gene expression. Functionally important CArG boxes have been identified in every SMC-specific transcriptional

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regulatory element characterized to date (Herring and Smith, 1997; Kim et al., 1997; Li et al., 1996a,b; Mack and Owens, 1999; Madsen et al., 1998; Moessler et al., 1996). Multimerized copies of the SMC-specific CArG elements are sufficient to direct gene expression to SMCs in transgenic mice (Kim et al., 1997; Solway et al., 1995). Dominant-negative SRF constructs block the induction of SMCs from undifferentiated mesechyme (Landerholm et al., 1999). However, SMC-specific consensus nucleotide sequences within or flanking CArG motifs have not been identified. Moreover, SMC-restricted ternary complex factors (TCF), or SRF-binding partners, have not been identified. It is possible that SRF activity within SMCs is activated posttranscriptionally. In this regard, it is noteworthy that the related MADS box family member, MEF2, is regulated, in part, by its capacity to heterodimerize with transcriptional activators and coactivators in skeletal muscle. Alternatively, it is possible that SRF is activated in SMCs by a novel cytoskeletal signal mediated through actin treadmilling (Sotiropoulos et al., 1999). Elucidation of the molecular mechanisms regulating activity of SRF in SMCs should provide fundamental insights into SMC differentiation and possibly modulation of SMC phenotype in response to vessel wall injury. Current paradigms suggest that SMCs reversibly modulate their phenotype from a contractile cell to a synthetic cell (Owens, 1998; Owens et al., 1996). This model is based on the behavior of vascular smooth muscle cells in culture (ChamleyCampbell et al., 1979; Ross, 1971). It is now clear that this bimodal model actually represents extremes of a spectrum, and SMC phenotypic “shades of gray” exist in the vessel wall and in atherosclerotic plaques (Shanahan et al., 1998). These various phenotypes recapitulate embryonic and neonatal VSMC phenotypes (Shanahan et al., 1994). A great deal is understood about the environmental factors and signaling pathways that modulate SMC phenotype (for review, see Owens, 1998; Schwartz, 1997; Schwartz et al., 1986; Schwartz and Liaw, 1993). However, relatively little is understood about how these signals are transduced to the VSMC nucleus. NF-␬B signaling appears to play an imporant role in activating SMCs in response to vessel wall injury and inflammation (Lindner, 1998). It is also noteworthy that MEF2 and GATA factors that regulate growth responses in cardiac myocytes are differentially regulated in response to vessel wall injury (Firulli et al., 1996; Toshiaki et al., 1999). Elucidation of the mechanisms that regulate VSMC phenotype and proliferation is fundamentally important and could eventually lead to novel treatments for vascular proliferative syndromes, including atherosclerosis. In conclusion, despite the important role VSMCs play in modulating arterial tone and in the pathogenesis of vascular proliferative syndromes, relatively little is currently understood about the transcriptional programs that control SMC development and differentiation. The recent explosion of biological information and technologies, including experimental transgenesis and conditional gene knockouts, genomics, and stem cell biology, has catalyzed the move of SMC biology from the tissue culture plate to the arterial wall and atherosclerotic plaque. However, analysis of SMCs in vivo has served to highlight the complexity and heterogeneity

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of this muscle cell lineage. As such, elucidation of the transcriptional programs that control SMC development and differentiation may establish new paradigms in developmental biology that require integration of the concepts of cellular differentiation and functional heterogeneity.

Acknowledgments I thank Angelika Boyce for expert secretarial assistance. I thank Jon Epstein and Clayton Buck for their thoughtful review and comments on this manuscript. This manuscript was supported in part by NIH RO1 56915 to MSP.

References Adam, P. J., Weissberg, P. L., and Metcalfe, J. C. (1993). Polyubiquitin is a new phenotype marker of contractile vascular smooth muscle cells. Cardiovasc. Res. 33, 416–421. Aikawa, E., and Kawano, J. (1982). Formation of coronary arteries sprouting from the primitive aortic sinus wall of the chick embryo. Experientia 38, 816–818. Arber, S., Barbayannis, F. A., Hanser, H., Schneider, C., Stanyon, C. A., Bernard, O., and Caroni, P. (1998). Regulation of actin dynamics through phosphorylation of cofilin by LIM-kinase. Nature 393, 805–809. Baeuerle, P. A., and Baltimore, D. (1988). I kappa B: a specific inhibitor of the NF-kappa B transcription factor. Science 242, 540–546. Baeuerle, P. A., and Henkel, T. (1994). Function and activation of NF-kappa B in the immune system. Annu-Red. Immunol. 12, 141–179. Belaguli, N. S., Schildmeyer, L. A., and Schwartz, R. J. (1997). Organization and myogenic restricted expression of the murine serum response factor gene. J. Biol. Chem. 272, 18222–18231. Benditt, E. P., and Benditt, J. M. (1973). Evidence for the monoclonal orgin of human atheroslerotic plaques. Proc. Natl. Acad. Sci. USA 70, 1753–1756. Bergsma, D. J., Grichnik, J. M., Gossett, L. M., and Schwartz, R. J. (1986). Delimitation and characterization of cis-acting DNA sequences required for the regulated expression and transcriptional control of the chicken skeletak alpha-actin gene. Mol. Cell. Biol. 6, 2462–2475. Black, B. L., and Olson, E. N. (1998). Transcriptional control of muscle development by myocyte enhancer factor-2 (MEF2) proteins. Annu. Rev. Cell Dev. Biol. 14, 167–196. Blank, R. S., Swartz, E. A., Thompson, M. M., Olson, E. N., and Owens, G. K. (1995). A retinoic acid-induced clonal cell line derived from multipotential P19 embryonal carcinoma cells express smooth muscle characteristics. Circ. Res. 76, 742–749. Bochaton-Pillat, M.-L., Ropraz, P., Gabbiani, F., and Gabbiani, G. (1996). Phenotypic heterogeneity of rat arterial smooth muscle cell clones. Artherioscler. Thromb. Vasc. Biol. 16, 815–820. Bostrom, H., Willetts, K., Oekny, M., leveen, O., Lindahl, P., Hedstrand, H., Pekna, M., Hellstrom, M., Gebre-Medhin, S., Schalling, M., Nilsson, M., Kurland, S., Tormell, J., Heath, J., and Betsholtz, C. (1996). PDGF-A signaling is a critical event in lung alveolar myofibroblast development and alveogenesis, Cell 85, 863–873. Bourcier, T., Skhova, G., and Libby, P. (1997). The nuclear factor kappa-␤ signaling pathway participates in dysregulation of vascular muscle cells in vitro and in human atherosclerosis. J. Biol. Chem. 272, 15817–15824. Browning, C. L., Culberson, D. E., Aragon, I. V., Fillmore, R. A., Croissant, J. D., Schwartz, R. J., and Zimmer, W. E. (1998). The developmentally regulated expression of serum response factor plays a key role in the control of smooth muscle-specific genes. Dev. Biol. 194, 18–37.

2. Vascular Smooth Muscle Cell Development and Differentiation

83

Campbell, G. R., Campbell, J. H., Manderson, J. A., Horrigan, S., and Rennick, R. E. (1988). Arterial smooth muscle: a mutifunctional mesenchymal cell. Arch. Pathol. Lab. Med. 112, 977–986. Carlier, M. F., Laurent, V., Santolini, J., Meiki, R., Didry, D., Xia, G. X., Hong, Y., Chua, N. H., and Pantaloni, D. (1997). Actin depolymerizing factor (ADF/cofilin) enhances the rate of filament turnover: implication in actin-based motility. J. Cell. Biol. 136, 1307–1322. Carnac, G., Primig, M., Kitzman, M., Chafey, P., Tuil, D., Lamb, N., and Fernandez, A. (1998). RhoA GTPAase and serum response factor control selectively the expression of MyoD without affecting Myf5 in mouse myoblasts. Mol. Biol. Cell 9, 1891–1902. Chamley-Campbell, J., Campbell, G. R., and R. Ross, R. (1979). The smooth muscle cell in culture. Physiol. Rev. 59, 1–6. Chen, C. Y., Croissant, J., Majesky, M., Topouzis, S., McQuinn, T., Frankovsky, M. J., and Schwartz, R. J. (1996). Activation of the cardiac alpha-actin promoter depends upon serum response factor, Tinman homologue, Nkx-2.5, and intact serum response elements. Dev. Genet. 19, 119–130. Chin, E. R., Olsen, E. N., Richardson, J. A., Yang, Q., Humphries, C., Shelton, J. M., Wu, H., Zhu, W., Bassel-Duby, R., and Williams, R. S. (1998). A calcineurin-dependent transcriptional pathway controls skeletal muscle fiber type. Genes Dev. 12, 2499–2509. Chow, K. L., and Schwartz, R. J. (1990). A combination of closely associated positive and negative cis-acting promoter elements regulates transcription of the skeletal alpha-actin gene. Mol. Cell. Biol. 10, 528–538. Clesham, G. J., Adam, P. J., Proudfoot, D., Flynn, P. D., Efstathoiu, S., and Weissberg, P. L. (1998). High adenoviral loads stimulate NF kappa␤–dependent gene expression in human vascular smooth muscle cells. Gene Therapy 5, 174–180. Croissant, J. D., Kim, J.-H., Eichele, G., Goering, L., Lough, J., Prywes, R., and Schwartz, R. J. (1996). Avian serum response factor expression restricted to muscle cell lineages is required for a-actin gene transcription. Dev. Biol. 177, 250–264. Cunha, G. R., Battle, E., Young, P., Brody, J. R., Donjacour, A., Hayashi, N., and Kinbara, H. (1992). Role of epithelial–mesenchymal interactions in the differentiation and spatial organization of visceral smooth muscle. Epithelial Cell Biol. 1, 76–83. Dechend, R., Maass, M., Gieffers, J., Dietz, R., Scheidereit, C., Leutz, A., and Gulba, D. C. (1999). Chlamydia pneumoniae infection of vascular smooth muscle and endothelial cells activates NDkappaB and induces tissue factor and PAI-1 expression: A potential link to accelerated arteriosclerosis. Circulation 100, 1369–1373. Dettman, R., Denetclaw, W., Ordahl, C., and Bristow, J. (1998). Common epicardial origin of coronary vascular smooth muscle, perivascular fibroblasts and intermyocardial fibroblasts in the avian heart. Dev. Biol. 193, 169–181. Dickson, M. C., S., M. J., Cousins, F. M., Kulkarni, A. B., Karlsson, S., and Akhurst, R. J. (1995). Defective haematopoiesis and vasculogenesis in transforming growth factor beta1 knock out mice. Development 121, 1845–1854. Duband, J. L., Belkin, A. M., Syfrig, J., Thiery, J. P., and Koteliansky, V. E. (1992). Expression of alpha 1 integrin, a laminin-collagen receptor, during myogenesis and neurogenesis in the avian embryo. Development 116, 585–600. Duluc, I., Freund, J.-N., Lweberquier, C., and Kedinger, M. (1994). Fetal endoderm primarily holds the temporal and positional information required for mammalian intestinal development. J. Cell Biol. 126, 211–221. Durocher, D., Chen, C. Y., Ardati, A., Schwartz, R. J., and Nemer, M. (1996). The atrial natriuretic factor promoter is a downstream target for Nkx-2.5 in the myocardium. Mol. Cell. Biol. 16, 4648–4655. Epstein, J. A. (1996). Pax3, neural crest and cardiovascular development. Trends Cardiovasc. Med. 6, 255–261. Epstein, D. J., Vogan, k. J., Trasler, D. G., and Gros, P. (1993). A mutation within intron 3 of the pax-3 gene produces aberrantly spliced mRNA transcripts in the Splotch (Sp) mouse mutant. Proc. Natl. Acad. Sci. USA 90, 532–536.

84

Michael S. Parmacek

Firulli, A. B., Miano, J. M., Weizhen, B., Johnson, A. D., Casscells, W., Olson, E. N., and Schwarz, J. J. (1996). Myocyte enhancer binding factor-2 expression and activity in vascular smooth muscle cells. Circ. Res. 78, 196–204. Franz, T. (1989). Persistent truncus arteriosus in the Splotch mutant mouse. Anat. Embryol. (Berl.) 180, 457–464. Gerrity, R. G., and Cliff, W. J. (1975). The aortic tunica media of the developing rat, I: quantitative steroeologic and biochemical analysis. Lab. Invest. 32, 585–600. Gorski, D. H., Lepage, D. F., Patel, C. V., Copeland, N. G., Jenkins, N. A., and Walsh, K. (1993). Molecular cloning of a diverged homeobox gene that is rapidly down-regulated during the G0/G1 transition in vascular smooth muscle cells. Mol. Cell Biol. 13, 3722–3733. Grichnik, J. M., French, B. A., and Schwartz, R. J. (1988). The chicken skeletal alpha-actin gene promoter region exhibits partial dyad symmetry and a capacity to drive bidirectional transcription. Mol. Cell. Biol. 8, 4587–4597. Grueneberg, D. A., Natesan, S., Alexandre, C., and Gilman, M. Z. (1992). Human and Drosophila homeodomain proteins that enhance the DNA-binding activity of serum response factor. Science 257, 1089–1095. Herring, B. P., and Smith, A. F. (1997). Telokin expression in A10 smooth muscle cells requires serum response factor. A. J. Physiol 272, C1394–1404. Hill, C. S., Wynne, J., and Treisman, R. (1995). The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell 81, 1159–1170. Hirschi, K. J., Rohovsky, S. A., and D’Amore, P. A. (1998). PDGF, TGF-beta and heterotypic cell–cell interactions mediate endothelial cell-induced recruitment of 10T 1/2 cells and their differentiation to a smooth muscle fate. J. Cell Biol. 141, 805–814. Holycross, B. J., Blank, R. S., Thompson, M. M., Peach, M. J., and Owens, G. K. (1992). Plateletderived growth factor-BB-induced suppression of smooth muscle cell differentiation. Circ. Res. 71, 1525–1532. Hood, L. C., and Rosenquist, T. H. (1992). Coronary artery development in the chick: origin and deployment of smooth muscle cells, and the effects of neural crest ablation. Anat. Rec. 234, 291– 300. Hughes, A. F. W. (1943). The histogenesis of the arteries of the chick embryo. J. Anat. 77, 266–287. Hungerford, J. E., Owens, G. K., Argraves, W. S., and Little, C. D. (1996). Development of the aortic vessel wall as defined by vascular smooth muscle and extracellular matrix markers. Dev. Biol. 178, 375–392. Hutchins, G. M., Kessler-Hanna, A., and Moore, G. W. (1988). Development of the coronary arteries in the embryonic human heart. Circulation 77, 1250–1258. Jain, M. K., Layne, M. D., Watanabe, M., Chin, M. T., Feinberg, M. W., Sibinga, N. E. S., Hsieh, C.M., Yet, S.-F., Stemple, D. L., and Lee, M.-E. (1998). In vitro system for differentiating pluripotent neural crest cells into smooth muscle cells. J. Biol. Chem. 273, 5993–5996. Kim, D.-K., Zhang, L., Dzau, V. J., and Pratt, R. E. (1994). H19, a developmentally regulated gene, is reexpressed in rat vascular smooth muscle cells after injury. J. Clin. Invest. 93, 355–360. Kim, S., Ip, H. S., Lu, M. M., Clendenin, C., and Parmacek, M. S. (1997). A serum response factor– dependent transcriptional regulatory program identifies distinct smooth muscle cell sublineages. Mol. Cell. Biol. 17, 2266–2278. Kirby, M. L. (1988). Role of extracardiac factors in heart development. Experientia 44, 944–951. Kirby, M. L., and Waldo, K. L. (1990). Role of neural crest in congenital heart disease. Circulation 82, 332–340. Kirby, M. L., and Waldo, K. L. (1995). Neural crest and cardiovascular patterning. Circ. Res. 77, 211–215. Kirby, M. L., Gale, T. F., and Stewart, D. E. (1983). Neural crest cells contribute to normal aorticopulmonary septation. Science 220, 1059–1061. Konieczny, S. F., and Emerson, C. P., Jr. (1987). Complex regulation of the muscle-specific contractile protein (troponin I) gene. Mol. Cell. Biol. 7, 3065–3075.

2. Vascular Smooth Muscle Cell Development and Differentiation

85

Krzesz, R., Wagner, A. H., Cattaruzza, M., and Hecker, M. (1999). Cytokine-inducible CD40 gene expression in vascular smooth muscle cells is mediated by nuclear factor kappa␤ and signal transducer and activation of transcription-1. FEBS Lett. 453, 191–196. Landerholm, T. E., Dong, X.-R., Belaguli, N. S., Schwartz, R. J., and Majesky, M. W. (1999). A role for serum response factor in coronary smooth muscle differentiation from proepicardial cells. Development 126, 2053–2062. Landry, D. B., Couper, L. L., Bryant, S. R., and Lindner, V. (1997). Activation of the NF kappa ␤ and I kappa ␤ system in smooth muscle cells after rat arterial injury. Induction of vascular cell adhesion molecule-1 and monocyte chemoattractant protein 1. Am. J. Pathol. 151, 1085–1095. Lappalainen, P., and Drubin, D. G. (1997). Cofilin promotes rapid actin filament turnover in vivo. Nature 388, 78–82. Lassar, A. B., Buskin, J. N., Lockshon, D., Davis, R. L., Apone, S., Hauschka, S. D., and Weintraub, H. (1989). MyoD is a sequence-specific DNA binding protein requiring a region of myc homology to bind to the muscle creatine kinase enhancer. Cell 58, 823–831. Lelievre, C. S., and Ledouarin, N. M. (1975). Mesenchymal derivatives of the neural crest: Analysis of chimeric quail and chick embryos. J. Embryol. Exp. Morphol. 34, 125–154. Lemire, J. M., Potter-Perigo, S., Hall, K. L., Wight, T. N., and Schwartz, S. M. (1996). Distinct rat aortic smooth muscle cells differ in versican/PG-M expression. Arterioscler. Thromb. Vasc. Biol. 16, 821–829. Li, L., Miano, J. M., Cserjesi, P., and Olson, E. N. (1996a). SM22␣, a marker of adult smooth muscle, is expressed in multiple myogenic lineages during embryogenesis. Circ. Res. 78, 188–195. Li, L., Miano, J. M., Mercer, B., and Olson, E. N. (1996b). Expression of the SM22a promoter in transgenic mice provides evidence for distinct transcriptional regulatory programs in vascular and visceral smooth muscle cells. J. Cell Biol. 132, 849–859. Lilly, B., Zhao, B., Ranganayakulu, G., Paterson, B. M., Schulz, R. A., and Olson, E. N. (1995). Requirement of MADS domain transcription factor D-MEF2 for muscle formation in Drosophila. Science 267, 688–693. Lin, Q., Lu, J., Yanagisawa, H., Webb, R., Lyons, G. E., Richardson, J. A., and Olson, E. N. (1998). Requirement of the MADS-box transcription factor MEF2C for vascular development. Development 125, 4565–4574. Lindahl, P., Johansson, B. R., P., L., and Betsholtz, C. (1997). Pericyte loss and microaneurysm formation in PDGF-B-deficient mice. Science 277, 242–245. Lindner, V. (1998). The NF-kappa␤ and Ikappa␤ system in injured arteries. Pathobiology 66, 311–320. Lindsay, E. A., Botta, A., Jurecic, V., Carattini-Rivera, S., Cheah, Y. C., Rosenblatt, H. M., Bradley, A., and Baldini, A. (1999). Congenital heart disease in mice deficient for the DiGeorge syndrome region. Nature 401, 335–337. Mack, C. P., and Owens, G. K. (1999). Regulation of smooth muscle alpha-actin expression in vivo is dependent on CArG elements within the 5′ and first intron promoter regions. Circ. Res. 84, 852–861. Madsen, C. S., P., R. C., Hungerford, J. E., White, S. L., Manabe, I., and Owens, G. K. (1998). Smooth muscle-specific expression of the smooth muscle myosin heavy chain gene in transgenic mice requires 5′ -flanking and first intronic DNA sequence. Circ. Res. 82, 908–917. Majesky, M. W., Giachelli, C. M., Reidy, M. A., and Schwartz, S. M. (1992). Rat carotid neointimal smooth muscle cells reexpress a developmentally regulated mRNA phenotype during repair of arterial injury. Circ. Res. 71, 759–768. McHugh, K. M. (1995). Molecular analysis of smooth muscle development in the mouse. Devel. Dynam. 204, 278–290. Miano, J. M., Cserjesi, P., Ligon, K. L., Periasamy, M., and Olson, E. N. (1994). Smooth muscle myosin heavy chain exclusively marks the smooth muscle lineage during mouse embryogenesis. Circ. Res. 75, 803–812. Miano, J. M., Topouzis, S., Majesky, M. W., and Olson, E. N. (1996). Retinoid receptor expression and all-trans retinoic acid–mediated growth inhibition in vascular smooth muscle cells. Circulation 93, 1886–1895.

86

Michael S. Parmacek

Mikawa, T., and Fischman, D. A. (1992). Retroviral analysis of cardiac morphogenesis: Discontinuous formation of coronary vessels. Proc. Natl. Acad. Sci. USA 89, 9504–9508. Mikawa, T., and Gourdie, R. G. (1996). Pericardial mesoderm generates a population of coronary smooth muscle cells migrating into the heart along with ingrowth of the epicardial organ. Dev. Biol. 174, 221–232. Minty, A., and Kedes, L. (1986). Upstream regions of the human cardiac actin gene that modulate transcription in muscle cells: presence of an evolutionarily conserved repeated motif. Mol. Cell. Biol. 6, 2125–2136. Miyagawa-Tomita, S., Waldo, K., Tomita, H., and Kirby, M. L. (1991). Temporospatial study of the migration and distribution of cardiac neural crest in quail-chick chimeras. Am. J. Anat. 192, 79– 88. Moessler, H., Mericskay, M., Li, Z., Nagl, S., Paulin, D., and Small, J. V. (1996). The SM22 promoter directs tissue-specific expression in arterial but not in venous or visceral smooth muscle cells in transgenic mice. Development 122, 2415–2425. Mohun, T. J., Chambers, A. E., Towers, N., and Taylor, M. V. (1991). Expression of genes encoding the transcription factor SRF during early development of Xenopus laevis: identification of a CArG box-binding activity as SRF. EMBO J. 10, 933–940. Morrisey, E. E., Ip, H. S., Lu, M. M., and Parmacek, M. S. (1996). GATA-6: a zinc finger transcription factor that is expressed in multiple cell lineages derived from lateral mesoderm. Dev. Biol. 177, 309–322. Mueller, C. G. F., and Nordheim, A. (1991). A protein domain conserved between yeast MCM1 and human SRF directs ternary complex formation. EMBO J. 10, 4219–4229. Murry, C. E., Gipaya, C. T., Bartosek, T., Benditt, E. P., and Schwartz, S. M. (1997). Monoclonality of smooth muscle cells in human atherosclerosis. Am J. Pathol. 151, 697–706. Muscat, G. E. O., Perry, S., Prentice, H., and Kedes, L. (1992). The human skeletal ␣-actin gene is regulated by a muscle-specific enhancer that binds three nuclear factors. Gene Expression 2, 111–126. Newby, A. C., and Zaltsman, A. B. (1999). Fibrous cap formation or destruction — the critical importance of vascular smooth muscle cell proliferation, migration and matrix formation. Cardiovasc. Res. 41, 345–360. Nishibatake, M., Kirby, M. L., and Van Mierop, L. H. S. (1987). Pathogenesis of persistent truncus anteriosus and dextroposed aorta in the chick embryo after neural crest ablation. Circulation 75, 255–264. Olivetti, G., Anversa, P., Milissari, M., and Loud, A. V. (1980). Morphometric study of early postnatal development of the thoracic aorta in the rat. Circ. Res. 47, 417–424. Orkin, S. H. (1992). GATA-binding transcription factors in hematopoietic cells. Blood 80, 575–581. Orlani, A., Ropraz, P., and Gabbiani, G. (1994). Proliferative activity and alpha-smooth muscle actin expression in cultured rat aortic smooth muscle cells are differently modulated by transforming growth factor-␤1 and heparin. Exp. Cell Res. 214, 528–536. Osbourn, J. K., Weissberg, P. L., and Shanahan, C. M. (1995). A regulatory element downstream of the rat SM22 alpha gene transcription start point enhances reporter gene expression in vascular smooth muscle cells. Gene 154, 249–253. Owens, G. K. (1995). Regulation of differentiation of vascular smooth muscle cells. Phys. Rev. 75, 487–517. Owens, G. K. (1998). Molecular control of vascular smooth muscle cell differentiation. Acta Physiol. Scand. 164, 623–635. Owens, G. K., Vernon, S. M., and Madsen, C. S. (1996). Molecular regulation of smooth muscle cell differentiation. J. Hypertension 14, S55–S64. Price, M. A., Rogers, A. E., and Treisman, R. (1995). Comparative analysis of the ternary complex factors Elk-1, SAP-1a, and SAP-2. EMBO J. 14, 2589–2601. Qian, J., Kumar, A., Szuscik, J. C., and Lessard, J. L. (1996). Tissue and developmental specific

2. Vascular Smooth Muscle Cell Development and Differentiation

87

expression of murine smooth muscle ␥ -actin fusion genes in transgenic mice. Dev. Dynam. 207, 135–144. Reecy, J., Belaguli, N., and Schwartz, R. (1999). Serum response factor-NK homeodmain factor interactions, role in cardiac development. In “Heart Development” (R. Harvey and N. Rosenthal, eds.), pp. 273–287. Academic Press, San Diego. Roman, J., and McDonald, J. A. (1992). Expression of fibronectin, the integrin alpha5, and alphasmooth muscle actin in heart and lung development. Am. J. Respir. Mol. Biol 6, 472–480. Romanoff, A. L. (1960). “The Avian Embryo: Structural and Functional Development.” MacMillan Co., New York. Rosenquist, T. H., and Beall, A. C. (1990). Elastogenic cells in the developing cardiovascular system. Ann. N. Y. Acad. Sci. 588, 106–119. Ross, R. (1971). The smooth muscle cell. II. Growth of smooth muscle in culture and formation of elastic fibers. H. Cell Biol. 50, 172–186. Ross, R. (1999). Mechanisms of disease: atherosclerosis—an inflammatory disease. N. Engl. J. Med. 340, 115–126. Samaha, F. F., Ip, H. S., Morrisey, E. E., Seltzer, J., Tang, Z., Solway, J., and Parmacek, M. S. (1996). Developmental pattern of expression and genomic organization of the calponin-h1 gene; a contractile smooth muscle cell marker. J. Biol. Chem. 271, 395–403. Sartorelli, V., Webster, K. A., and Kedes, L. (1990). Muscle-specific expression of the cardiac ␣-actin gene requires MyoD1, CArG-box binding factor, and Sp1. Genes Dev. 4, 1811–1822. Schatteman, G. C., Motley, S. T., Effmann, E. L., Bowen-Pope, D. F. (1995). Platelet-derived growth factor receptor alpha subunit deleted patch mouse exhibits severe cardiovascular dysmorphogenesis. Teratol. 51, 351–366. Schuger, L., Skubitz, A. P. N., Zhang, J., Sorokin, L., and He, L. (1997). Laminin alpha1 chain synthesis in the mouse developing lung: requirement for epithelial–mesenchymal contact and possible role in bronchial smooth muscle development. J. Cell. Biol. 139, 553–562. Schwartz, S. M. (1997). Smooth muscle migration in atherosclerosis and restenosis. J. Clin. Invest. 100, S87–S89. Schwartz, S. M., and Liaw, L. (1993). Growth control and morphogenesis in the development and pathology of arteries. J. Cardiovasc. Pharm. 21, S31–S49. Schwartz, S. M., and Murry, C. E. (1998). Proliferation and the monoclonal origins of atherosclerotic lesions. Ann. Rev. Med. 49, 437–460. Schwartz, S. M., Campbell, G. R., and Campbell, J. H. (1986). Replication of smooth muscle cells in vascular disease. Circ. Res. 58, 427–444. Selzman, C. H., Shames, B. D., Reznikov, L. L., Miller, S. A., Meng, X., Barton, H. A., Werman, A., Harken, A. H., Dinarello, C. A., and Banerjee, A. (1999). Liposomal delivery of purified inhibitorykappa␤ alpha inhibits tumor necrosis factor-alpha-induced human vascular smooth muscle proliferation. Circ. Res. 84, 867–875. Sepulveda, J. L., Belaguli, N., Nigam, V., Chen, C. Y., Nemer, M., and Schwartz, R. J. (1998). GATA-4 and Nkx-2.5 coactivate Knx-2 DNA binding targets: role for regulating early cardiac gene expression. Mol. Cell. Biol. 18, 3405–3415. Shah, N. M., Groves, A. K., and Anderson, D. J. (1996). Alternative neural crest cell fates are instructively promoted by TGFbeta superfamily members. Cell 85, 331–343. Shanahan, C. M., and Weissberg, P. L. (1997). Smooth muscle cell heterogeneity: Patterns of gene expression in vascular smooth muscle cells in vitro and in vivo. Arteriosclerosis, Thromb. Vas. Biol. 183, 333–338. Shanahan, C. M., Cary, N. R. B., Metcalfe, J. C., and Weissberg, P. L. (1994). High expression of genes for calcification-regulating proteins in human atherosclerotic plaques. J. Clin. Invest. 93, 2393–2402. Shanahan, C. M., Proudfoot, D., Farzaneh-Far, A., and Weissberg, P. L. (1998). The role of G1a proteins in vascular calcification. Crit. Rev. Eukaryotic Gene Expression 8, 357–375.

88

Michael S. Parmacek

Shi, Y., O’Brien, J. E., Fard, A., Mannion, J. d., Wang, D., and Zalewski, A. (1996). Adventitial myofibroblasts contribute to neointimal formation in injured porcine coronary arteries. Circulation 94, 1655–1664. Sobue, K., Hayashi, K., and Nishida, W. (1999). Expressional regulation of smooth muscle cell–specific genes in association with phenotypic modulation. Mol. Cell. Biochem. 190, 105–118. Solway, J., Seltzer, J., Samaha, F. F., Kim, S., Alger, L. E., Niu, Q., Morrisey, E. E., Ip, H. S., and Parmacek, M. S. (1995). Structure and expression of a smooth muscle cell–specific gene, SM22a. J. Biol. Chem. 270, 13460–13469. Sotiropoulos, A., Dziugas, G., J., C., and Treisman, R. (1999). Signal-regulated activation of serum response factor is mediated by changes in actin dynamics. Cell 98, 159–169. Stemple, D. L., and Anderson, D. J. (1992). Isolation of a stem cell for neurons and glia from the mammalian neural crest. Cell 71, 973–985. Suzuki, E., Guo, K., Kolman, M., Yu, Y.-T., and Walsh, K. (1995). Serum induction of MEF2/RSRF expression in vascular myocytes is mediated at the level of translation. Mol. Cell. Biol. 15, 3415– 3425. Tapscott, S. J., and Weintraub, H. (1991). MyoD and the regulation of myogenesis by helixloop-helix proteins. J. Clin. Invest. 87, 1133–1138. Topouzis, S., and Majesky, M. W. (1996). Smooth muscle lineage diversity in the chick embryo. Two types of aortic smooth muscle cell differ in growth and receptor-mediated transcriptional responses to transforming growth factor-␤. Dev. Biol. 178, 430–445. Topouzis, S., Catravas, J. D., Ryan, J. W., and Rosenqust, T. H. (1992). Influence of vascular smooth muscle heterogeneity on angiotensin converting enzyme activity in chicken embryonic aorta and in endothelial cells in culture. Circ. Res. 71, 923–931. Toshiaki, M., Zhengyu, L., Malendowicz, S. L., Evans, T., and Walsh, K. (1999). Reversal of GATA-6 downregulation promotes smooth muscle cell differentiation and inhibits intimal hyperplasia in balloon injured rat carotid arteries. Circ. Res. 84, 647–654. Treisman, R. (1994). Ternary complex factors: growth factor regulated transcriptional activators. Curr. Opin. Genet. Dev. 4, 96–101. Triesman, R. (1995a). Inside the MADS box. Nature 376, 468–469. Treisman, R. (1995b). Journey to the surface of the cell: Fos regulation and the SRE. EMBO J. 14, 4905–4913. Van Aelst, L., and D’Souza-Schorey, C. (1997). Rho GTPases and signaling networks. Genes Dev. 11, 2295–2322. Vershon, A. K., and Johnson, A. D. (1993). A short, disordered protein region mediates interactions between the homeodomain of the yeast alpha 2 protein and the MCM1 protein. Cell 72, 105–112. Weintraub, H., Davis, R., Tapscott, S., Thayer, M., Krause, M., Benezra, R., Blackwell, T. K., Turner, D., Rupp, R., and Hollenberg, S. (1991). The myoD gene family: nodal point during specification of the muscle cell lineage. Science 251, 761–766. Welch, G. N., Upchurch, G. R., Jr., Farivar, R. S., Pigazzi, A., Vu, K., Brecher, P., Keaney, J. F. J., and Loscalzo, J. (1998). Homocysteine-induced nitric oxide production in vascular smooth-muscle cells by NF-kappa B-dependent transcriptional activation of Nos-2. Proc. Assoc. Am. Physicians 110, 22–31. White, S. L., and Low, R. B. (1996). Identification of promoter elements involved in cell-specific regulation of rat smooth muscle myocin heavy chain gene transcription. J. Biol. Chem. 271, 15008– 15017. Yamagishi, H., Garg, V., Matsuoka, R., Thomas, T., and Srivastava, D. (1999). A molecular pathway revealing a genetic basis for human cardiac and craniofacial defects. Science 283, 1158– 1161. Yang, N., Higuchi, O., Ohashi, K., Nagata, K., Wada, A., Kangwa, K., Nishida, E., and Mizuno, K. (1998a). Cofilin phosphorylation by LIM-kinase 1 and its role in Rac-mediated actin reorganization. Nature 393, 809–812.

2. Vascular Smooth Muscle Cell Development and Differentiation

89

Yang, Y., palmer, K. C., Relan, N., Diglio, C., and Schuger, L. (1998b). Role of laminin polymerization at the epithelial mesenchymal interface in bronchial myogenesis. Development 125, 2621–2629. Yang, Y., Relan, N. K., Przywara, D. A., and Schuger, L. (1999). Embryonic mesenchymal cells share the potential for smooth muscle differentiation: myogenesis is controlled by the cell’s shape. Development 126, 3027–3033. Yano, H., Hayashi, K., T., M., Haruna, M., and Sobue, K. (1995). Transcriptional regulation of the chicken caldesmon gene. J. Biol. Chem. 270, 23661–23666. Yu, Y.-T., Breitbart, R. E., Smoot, L. B., Lee, Y., Mahdavi, V., and Nadal-Ginard, B. (1992). Human myocyte-specific enhancer factor 2 comprises a group of tissue-restricted MADS box transcription factors. Genes Dev. 6, 1783–1798. Zhou, L., Dey, C. R., Wert, S. E., and Whitsett, J. A. (1996). Arrested morphogenesis in transgenic mice bearing an SP-C-TGF-beta-1 chimeric gene. Dev. Biol. 1, 227–238.

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3 Myofibroblasts: Molecular Crossdressers Gennyne A. Walker, Ivan A. Guerrero, and Leslie A. Leinwand Department of Molecular, Cellular and Developmental Biology University of Colorado, Boulder 80309

I. Myofibroblasts: An Overview II. Myofibroblast Origin and the Role of PDGF III. Cytokines and Myofibroblast Phenotypes IV. “Muscle” Structural Protein Expression in Myofibroblasts V. VI. VII. VIII.

Mechanisms of “Muscle-Specific” Gene Regulation in Myofibroblasts Myofibroblast Contractility Myofibroblasts and the Cell Cycle Perspectives References

Myofibroblasts are unique mesenchymal cells with properties inherent to both muscle and nonmuscle cells. They are widely distributed in embryos, are essential for the formation of functional adult tissues, and are intimately involved in tissue homeostasis and wound healing. Cytoskeletal protein expression and contractile properties distinguish them from other cell types. Myofibroblasts also express skeletal muscle structural and regulatory proteins, including sarcomeric myosin heavy chain and MyoD. Despite the presence of such myogenic regulatory proteins, these cells do not terminally differentiate into skeletal muscle. This article focuses on the interesting biology of myofibroblasts, their origin, and the molecular mechanisms that allow these cells to maintain a state intermediate between muscle and nonmuscle cells.  2001 Academic Press. C

I. Myofibroblasts: An Overview Myofibroblasts are mesenchymal cells that have the ultrastructural properties of both muscle and nonmuscle cells. Originally described as “modified” fibroblasts Correspondence should be addressed to: Leslie A. Leinwand, Department of Molecular, Cellular and Developmental Biology, University of Colorado at Boulder, Campus Box 347, Boulder CO, 803090347. Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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located in granulation tissue (Gabbiani et al., 1971) they have subsequently been documented in a diverse array of tissues (reviewed in Powell et al., 1999). There are numerous cell types that have been characterized as myofibroblasts, including stromal cells, pericytes, stellate cells, interstitial cells, mesangial cells, and granulation tissue fibroblasts. They are intriguing cells that have been described for decades, but their molecular, cellular, and developmental properties have not been well elucidated. They are highly plastic and diverse in their phenotypes, depending on their tissue of origin and whether the tissue is normal or pathological. Common features include the expression of both muscle and nonmuscle structural and regulatory proteins, contractile properties, and secretion of extracellular matrix (ECM). Myofibroblasts have been shown to be essential for normal development and tissue homeostasis. They exhibit contractile properties absent in nonmuscle cells, and this contraction is thought to be essential in regulating blood flow (Forbes et al., 1999). In addition to their normal cellular functions, myofibroblasts are also consistently implicated in wound repair. However, if they proliferate and persist at the site of the wound, fibrosis may result. Their persistence has implicated them in fibrosis in various tissues, such as liver, heart, and kidney (reviewed in Schurch et al., 1998). During tissue injury resident quiescent mesenchymal cells transform into what is called an activated state (Fig. 1). Myofibroblast activation is characterized by the induction of a unique set of muscle genes, morphological change, contractility, and an increase in the production of extracellular matrix (ECM) (reviewed in Schurch et al., 1998). This in vivo activation can also be reproduced in cell culture by treatment with numerous cytokines, notably endothelin 1 (ET-1), angiotensin II (Ang II), platelet-derived growth factor (PDGF), and transforming growth factor-␤ (TGF-␤) (Desmouliere et al., 1993). A more extensive discussion of activation follows.

II. Myofibroblast Origin and the Role of PDGF The origin and differentiation pathways of myofibroblasts in different tissues have not been well characterized. Myofibroblasts are present during normal development and have been detected at 14.5dpc in developing kidney, brain, heart, lung, and brown adipose tissue. Their presence during development requires PDGF, but the myofibroblasts of different tissues have different PDGF requirements, as elegantly delineated by genetic inactivation studies in mice (Lindahl et al., 1998). PDGFs are homo- or heterodimers of A and B chains that interact with two distinct receptor tyrosine kinases, PDGFR-␣ and -␤, which appear to elicit very distinct downstream biological responses (reviewed in Rosenkranz and Kazlauskas, 1999). Mice deficient for PDGF-A exhibit lung defects and die either during embryogenesis or just after birth. These defects are caused by a lack of alveolar septation due to the absence of alveolar myofibroblasts (Bostrom et al., 1996). Conversely,

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Figure 1 Diagram of the myofibroblast activation process. Myofibroblasts may derive from pluripotent stem cells during embryogenesis or from mesenchymal cells (such as local fibroblasts) in adult tissue. Some myofibroblasts, such as liver stellate cells, exist in two morphologically distinct states, activated and stellate. Several cytokines and growth factors contribute to these processes, including PDGF (platelet-derived growth factor), TGF-␤ (transforming growth factor ␤), EGF (epidermal growth factor), bFGF (basic fibroblast growth factor), IL-1 (interlukin-1), cAMP (cyclic adenosine monophosphate), and cholera toxin.

animals lacking either PDGF-B or PDGF-R␤ die of hemorrhage and edema in late embryogenesis or early neonatal life, and demonstrate an absence of kidney mesangial cells and myofibroblasts in brain, heart, lung, and brown adipose tissue (Lindahl et al., 1997). Further analysis of these animals identified abnormal capillary structures, indicating that pericytes regulate microvessel structure and that the absence of pericytes/myofibroblasts causes rupture when blood pressure increases just after birth. The differences between the PDGF-A and PDGF-B/ PDGFR-␤ null phenotypes demonstrate that PDGFs differentially regulate the generation of specific populations of myofibroblasts during development. While myofibroblasts are necessary for normal lung development, their continued proliferation has been implicated in the pathogenesis of pulmonary fibrosis. Tyrosine kinase inhibitors of the tyrphostin class, which specifically block autophosphorylation of the PDGF receptors or epidermal growth factor receptor (EGF-R), have been successfully used to suppress myofibroblast proliferation in a rat model of pulmonary fibrosis (Rice et al., 1999). This provides additional evidence that the PDGF receptors are involved in the proliferation and activation of myofibroblasts.

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The different requirements of myofibroblasts for PDGF suggest that myofibroblasts from different tissues are specialized mesenchymal cellular isoforms that may share a common ancestor and can transform into different or similar myofibroblastic phenotypes, given the particular microenvironment. Characterization of kidney and liver myofibroblast cell lines in our laboratory has demonstrated distinct patterns of gene expressions between these two myofibroblast types (Mayer and Leinwand, 1997).

III. Cytokines and Myofibroblast Phenotypes Although the molecular mediators of myofibroblast activation are not defined in great detail, both TGF-␤ and ET-1 have been implicated in the activation of myofibroblasts at sites of tissue damage (Desmouliere et al., 1993). ET-1 is a potent vasoconstrictive agent expressed in many models of tissue injury. It has been demonstrated that addition of ET-1 to cultured myofibroblasts results in a plethora of effects, including increased cytosolic free calcium concentration (Rockey, 1997), increased expression of sarcomeric proteins (Mayer and Leinwand, 1997), and contraction (Rockey and Chung, 1996). It is noteworthy that after tissue injury, there are both localized and systemic increases in ET-1 levels. In a rodent myocardial infarction model, blocking ET-1 function with a receptor antagonist can reduce collagen secretion and improve survival rate of the animals, implicating ET-1 as a causative agent for myofibroblast activation in vivo (Sakai et al., 1996). Ang II is also thought to be a “player” in myofibroblast activation. An interesting hypothesis is that intracardiac Ang II generation at the site of tissue repair involves the activation of myofibroblasts (reviewed in Weber, 1997). With respect to fibrosis in other tissues, accumulation and proliferation of macrophages and myofibroblasts in renal fibrosis has been reported to be inhibited by the blockade of Ang II activity (Wu et al., 1997). Like ET-1, Ang II is a potent vasoconstrictor, and also like ET-1, administration of Ang II receptor antagonists can result in the regression of injurious phenotypes such as hypertension, nephropathy, and cardiac hypertrophy (Madeddu et al., 2000). Ang II is commonly found at sites of tissue damage, inflammation, and repair. It has been reported to stimulate autocrine release of endogenous growth factors such as TGF-␤ (Campbell and Katwa, 1997) and ET-1 (Gray et al., 1998) from smooth muscle cells, cardiac fibroblasts, and myofibroblasts. Cultured myofibroblasts from rat subcutaneous and cardiac scar tissue express components necessary for Ang II autocrine stimulation, such as the precursor angiotensinogen, angiotensin converting enzyme (ACE), and Ang II receptors (AT1 and AT2 ) (Sun, 1997). Since myofibroblasts secrete ECM proteins, this activation is thought to represent an autocrine/paracrine model for collagen turnover in injured tissues (Weber, 1997). The cellular origin of Ang II at the site of myocardial infarction remains unknown. The expression of genes required for

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Ang II synthesis make myofibroblasts excellent candidates for Ang II production in myocardial infarction.

IV. “Muscle” Structural Protein Expression in Myofibroblasts Myofibroblasts have the unusual property of expressing proteins otherwise found only in smooth and skeletal muscle, but are not themselves muscle cells. Their presence in numerous tissue types and their apparent inability to morphologically differentiate into muscle or to withdraw from the cell cycle (discussed in Section VII) distinguish them from the well-studied skeletal muscle myoblast. The levels of expression and the number of muscle genes expressed are dependent upon the microenvironment (be it a pathological or normal setting) and tissue type. Included in the genes that can be expressed by myofibroblasts are ␣-smooth muscle actin (ASMA) and desmin, as well as both smooth and sarcomeric myosin heavy chain (MyHC) isoforms. Desmin is a muscle-specific intermediate filament found in all three muscle tissue types (smooth, skeletal, and cardiac) (Steinert and Roop, 1988) and is one of the earliest known myogenic markers of skeletal and cardiac myogenisis (reviewed in Capetanaki et al., 1997). ASMA is initially expressed during development in all three muscle types (smooth, cardiac, and skeletal), but disappears from cardiac and skeletal muscle after birth (Woodcock-Mitchell et al., 1988). In addition to smooth muscle myosin, myofibroblasts can express several sarcomeric MyHC isoforms, as depicted in Table I. Cultured kidney myofibroblasts (baby hamster kidney, BHK-21) constitutively express six skeletal MyHCs, including both developmental and adult fast isoforms (Mayer and Leinwand, 1997). Rat liver myofibroblasts (stellate cells) express skeletal muscle IIa, IId, and perinatal MyHC genes (Mayer and Leinwand, 1997). Analysis of liver sections with a sarcomeric MyHC specific antibody found that expression of skeletal MyHC also occurs in vivo in liver stellate cells (Mayer and Leinwand, 1997), demonstrating that sarcomeric MyHC expression is not a cell culture artifact. Sarcomeric MyHC expression is constitutive and appears to be limited to the skeletal muscle program since the cardiac-specific ␣-MyHC isoform is not expressed. The normal developmental cues for regulating MyHC gene expression appear to be altered or completely absent in myofibroblasts (Mayer and Leinwand, 1997). In BHK-21 and liver stellate cells, both developmental and adult isoforms of sarcomeric genes are expressed simultaneously rather than sequentially, as they are during skeletal muscle development. The existence of distinct gene expression profiles in different myofibroblasts also suggests that the entire program of muscle gene expression is not recapitulated and may be diverse in different myofibroblasts. In liver stellate cells, skeletal myosin has thus far been the only sarcomeric protein observed. On the other hand, BHK-21 cells express the thin filament sarcomeric

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Table I Muscle Protein Expression in Liver and Kidney Myofibroblastsa

Myofibroblast markers

Sarcomeric thick filament

Sarcomeric thin filament

Transcription factors

Liver stellate

BHK-21

References

Desmin

+

+

␣-Smooth muscle actin

+

+

Smooth muscle myosin Embryonic MyHC Perinatal MyHC Skeletal IIa MyHC Skeletal IIb MyHC Skeletal IId MyHC ␤-MyHC ␣-Cardiac MyHC Sarcomeric myosin light chain Titin

+ − + + − + − − −

+ + + + + + + − +

Yokoi et al., 1984; Tuszynski et al., 1979 Ramadori et al., 1990; Schaart et al., 1991 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997

−

+

Myosin binding protein C Sarcomeric actin Sarcomeric tropomyosin Sarcomeric troponin T Nebulin MyoD

n/d − − − − +

+ + + + + +

Myogenin MEF2 Myf5

+ n/d −

+ + −

Mayer and Leinwand, 1997; Schaart et al., 1991 van der Ven and Furst, 1998 Koffer and Dickens, 1987 Mayer and Leinwand, 1997 Mayer and Leinwand, 1997 van der Ven and Furst, 1998 Mayer and Leinwand, 1997; Redfield et al., 1997 Mayer and Leinwand, 1997 Redfield et al., 1997 Mayer and Leinwand, 1997

a Summary of muscle specific protein expression in liver and kidney myofibroblasts, as determined by immunohistochemical methods. The presence (+) or absence (−) of a specific protein is specified; (n/d) indicates that protein expression has not been determined.

proteins troponin, tropomyosin, and skeletal actin (Koffer and Dickens, 1987), as well as the sarcomeric scaffolding proteins titin, myosin binding protein C, and nebulin (van der Ven and Furst, 1998). These proteins are abundantly expressed in striated muscle and interact with each other to form the highly ordered contractile unit known as the sarcomere. Although both liver stellate and BHK-21 cells have contractile properties (discussed in Section VI), the unit of contraction in these cells has not been determined. The absence of thin filament proteins in liver stellate cells suggests that neither the traditional sarcomere nor the full components of the sarcomere are necessary for myofibroblast contractile function. The involvement of sarcomeric proteins in myofibroblast contractility is discussed in Section VI. The differences in sarcomeric gene expression between the liver stellate and BHK-21 cells and between myofibroblasts and skeletal muscle myotubes provide

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compelling evidence that a unique mechanism of muscle gene regulation may exist in myofibroblasts. Although it has not been demonstrated that differences occur among myofibroblasts in vivo, it is tempting to speculate that such differences are inherent to the function of myofibroblasts within the context of a specific tissue.

V. Mechanisms of “Muscle-Specific” Gene Regulation in Myofibroblasts One question that arises from the observation of “muscle-specific” protein expression in myofibroblasts is whether the same gene regulatory mechanisms that exist in skeletal muscle are active in myofibroblasts. Skeletal muscle gene expression is controlled by a complex group of regulators that derive from one of two major families of transcription factors. The first is the MyoD family of myogenic regulatory factors (MRFs) consisting of MyoD, Myf5, myogenin, and MRF4 (herculin or myf6) (reviewed in Molkentin and Olson, 1996). This family is characterized by each member’s ability to convert nonmuscle cells into skeletal muscle fibers through activation of the complete muscle differentiation program. MRFs heterodimerize with the ubiquitously expressed family of E-proteins (Lassar et al., 1991) and activate transcription by binding to conserved E-box elements (CANNTG) often found in muscle regulatory sequences. The second family of transcriptional activators is the myocyte enhancer factor 2 (MEF2) group of MADS box regulators (reviewed in Black and Olson, 1998), which is composed of four genes: Mef2a, Mef2b, Mef2c, and Mef2d. MEF2 proteins bind to AT-rich elements that are also located in numerous cardiac and skeletal muscle regulatory regions (Black et al., 1998). Members of both families are essential to the process of cell determination, muscle gene expression, and differentiation. Although MRF expression is limited primarily to cells of skeletal lineage, expression of MyoD and myogenin have also been detected in myoid cells of the thymus (Grounds et al., 1992) and more recently in myofibroblasts (Mayer and Leinwand, 1997). MyoD and myogenin proteins have been detected in both liver stellate and BHK-21 cell lines (Redfield et al., 1997) and are localized to the nucleus. Myf-5 was not detected in either cell line (Mayer and Leinwand, 1997), whereas attempts to detect MRF4 in myofibroblasts have not been described. The MEF2 proteins are more widely distributed than the MRFs and are predominantly found in all three types of muscle (skeletal, cardiac, and smooth) as well as in neuronal tissue (Leifer et al., 1994). MEF2 protein expression, detected with an antibody that recognizes all four gene products, has been reported in BHK-21 cells (Redfield et al., 1997); no attempts to identify MEF2 expression in other myofibroblast cells have been reported. It is well documented that most cell types terminally differentiate when any one of the MRFs is exogenously expressed. For example, overexpression of MyoD

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induces the transformation of both primary fibroblasts and differentiated melanoma, liver, and adipocyte cells into muscle (Crescenzi et al., 1990), demonstrating its pivotal and dominant role in the muscle differentiation pathway. Interestingly, despite the endogenous expression of both MyoD and myogenin, myofibroblast cell lines do not morphologically differentiate into multinucleated myotubes, nor do they withdraw from the cell cycle (Mayer and Leinwand, 1997). What may render myofibroblasts resistant to cell cycle withdrawal is discussed in Section VII. The presence of MRFs does not necessarily mean that they are active. For example, MyoD is present in BC3H1 cells (derived from a rhabdosarcoma) but is inactive (Tapscott et al., 1993). However, myogenic transcription factor activity is present in myofibroblasts (Mayer and Leinwand, 1997). A muscle-specific promoter linked to a reporter gene (Wentworth et al., 1991) was transfected into BHK-21 or liver stellate cells. The level of activity of this construct in myofibroblasts was comparable to that of differentiated C2 C12 skeletal muscle myotubes. This construct contains the myosin light chain 1,3 promoter and downstream muscle specific enhancer linked to a chloramphenicol acetyltransferase reporter gene (Rosenthal et al., 1989). In skeletal muscle, maximal activation of this construct depends upon three intact E-boxes and a MEF2 binding site located in the enhancer region (Rao et al., 1996). Constructs in which the E-boxes have been mutated are inactive in both myofibroblasts and C2 C12 myotubes, suggesting that at least some aspects of the sarcomeric muscle gene program in myofibroblasts are functional. However, it has not been determined to what extent each of the three muscle transcription factors (MyoD, myogenin, and MEF2) participates in myofibroblast muscle gene expression in vitro or in vivo. It is interesting that of the four MRFs only MyoD and myogenin are known to be expressed in myofibroblasts, as each has been assigned a separate and nonredundant function in myogenesis. Myf-5 and MyoD are thought to function redundantly in muscle lineage determination because, although inactivation of either gene individually in transgenic mice results in no gross skeletal muscle defects, the double null mouse is completely devoid of skeletal muscle (Rudnicki et al., 1993). On the other hand, myogenin and MRF4 are not necessary for cell determination but are believed to be intimately involved with the morphological and biochemical differentiation of a determined cell (Nabeshima et al., 1993), and with adult myogenesis (Olson et al., 1996). Although the MRF4 null mice display varied and complicated phenotypes (Olson et al., 1996), MRF4 is the predominant MRF expressed in adult muscle (Rhodes and Konieczny, 1989), implicating it in the later stages of muscle development. Myogenin null mice are capable of developing myoblasts, but subsequent fusion of these cells into myotubes does not occur in vivo (Nabeshima et al., 1993), even though these myoblasts are able to fuse when placed into culture (Nabeshima et al., 1993). Genetic inactivation of the individual MEFs are just beginning to allow elucidaton of their functions: Mef2C null mice have severe defects in cardiogenesis (Bi et al., 1999), whereas Mef2A and Mef2B null mice are viable (E. Olson, personal communication).

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It is still unclear how the MRFs and MEFs coordinate with other transcription factors and gene regulatory elements to activate the full program of muscle specific gene expression, even in skeletal muscle. Expression of the muscle creatine kinase and myosin light chain genes has been shown to be directly activated by MRFs (Wentworth et al., 1991) and MEFs (Cserjesi et al., 1994) via E-boxes and AT-rich elements, respectively. These same motifs are located in the MyHC IIb promoter of rat and mouse, but utilization of these regulatory elements differs between the two species (Wheeler et al., 1999). This is interesting since MyHC IIb is expressed in the hamster renal myofibroblast line, but not in rat liver myofibroblasts. Although such species variation in muscle gene regulation may or may not contribute to the differences in myofibroblast MyHC isoform expression, it does exemplify the difficulty in assessing the contribution of each myogenic transcription factor to MyHC expression within the context of the myofibroblast. Complicating the issue further is the discovery that the MRFs and MEF2 proteins can also function synergistically at a single DNA site (Molkentin et al., 1995). Such synergy is mediated through interactions between the bHLH and MADs box domains of these proteins and does not appear to require the direct binding of DNA by MEF2. The MRFs and MEFs appear to function in a combinatorial manner at common regulatory elements to control the spatial and temporal expression patterns of skeletal muscle genes. Which regulatory mechanisms are in place in myofibroblasts remains to be elucidated, although the constitutive expression of both developmental and adult MyHC isoforms suggests that a unique combination of myogenic factors may be interacting at muscle-specific regulatory elements. Expression of desmin and ␣-smooth muscle actin (ASMA) is also regulated by the myogenic transcription factors. The desmin regulatory regions identified thus far have two E-boxes and a MEF2 site that work cooperatively to activate muscle-specific transcription (Li and Capetanaki, 1994; van de Klundert et al., 1994). E-boxes are also found in the regulatory region of the ASMA gene. It has been reported that an ASMA promoter construct can be activated in skeletal muscle myotubes via direct interactions between these E-boxes and myogenin (Johnson and Owens, 1999). This finding is not unexpected, since ASMA is expressed during early cardiogenesis and skeletal myogenesis (WoodcockMitchell et al., 1988) in addition to developing and adult smooth muscle. What is surprising is that these E-boxes are also involved in smooth muscle cell–specific activation but are bound by a different bHLH protein, USF (Johnson and Owens, 1999). The smooth muscle MyHC regulatory region also contains several CArG and E-box elements, as well as a MEF2-like element (White and Low, 1996); however the participation of these elements in smooth muscle MyHC expression has not been analyzed in a skeletal muscle cell context. The utilization of these regulatory elements by two different muscle cell types raises an interesting question: What are the relative contributions of the skeletal and smooth muscle gene regulatory systems to muscle gene expression in myofibroblasts?

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Obviously, the discovery of skeletal muscle structural and regulatory proteins in myofibroblasts has generated more speculation than explanation regarding the control of sarcomeric gene expression, the molecular basis of contractility in these cells, and even their cellular origin. Determining whether or not the MRFs and MEFs are singularly responsible for muscle gene expression in liver and kidney myofibroblasts will be significant to both myofibroblast and skeletal muscle research. In addition, it will be important to find out if myofibroblasts from other tissues express these muscle-specific genes, and to assess whether or not a correlation between muscle gene expression and tissue-specific functions may exist.

VI. Myofibroblast Contractility The observation of sarcomeric protein expression and the demonstration that myofibroblasts have contractile properties led to the attractive hypothesis that sarcomeric proteins participate in, or are required for myofibroblast contractility. In this section, we summarize what is known about myofibroblast contractility. Large numbers of myofibroblasts appear during wound contraction and disappear after normal wound healing. Myofibroblasts from cultured cell lines and isolated as primary cultures from injured tissues have been shown to be able to contract a collagen matrix in response to cytokine treatment. This contraction is accompanied by an increase in expression of muscle contractile proteins such as ASMA and MyHC (Mayer and Leinwand, 1997). When myofibroblasts are treated with ET-1, there is a general induction of all sarcomeric proteins that are already expressed in untreated cells, but no induction of additional sarcomeric proteins (Mayer and Leinwand, in preparation). This induction appears to be specific to the skeletal muscle proteins, since there is no increase in total protein synthesis or in expression of nonmuscle or smooth muscle MyHC. It is quite reasonable that a cell involved in remodeling would have a well-defined cytoskeletal network to provide support during the intense stress forces incurred in myocardial or tissue remodeling; proteins that could provide such a cytoskeletal network are expressed in myofibroblasts. However, the organization of the cytoskeleton and myofibroblast contractility raises an intriguing question. The traditional structural unit for the contractile proteins of striated muscle is the sarcomere. Even though myofibroblasts express a wide variety of muscle contractile proteins, there are no sarcomeres present. Electron microscopic analysis of sarcomeric MyHC in myofibroblasts has identified large filaments that are distinct from those formed by nonmuscle MyHC, but higher order structures are not apparent (Mayer and Leinwand, 1997). Despite this observation, sarcomeric MyHC does appear to play a role in myofibroblast contractility. We have shown that contraction of myofibroblasts is accompanied by an increase in sarcomeric MyHC expression and that inhibition of sarcomeric MyHC assembly blocks

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contraction (Mayer and Leinwand, in preparation). It remains to be determined what the contractile unit of myofibroblasts is and what interactions between nonmuscle and muscle contractile proteins result in the contractility of these cells.

VII. Myofibroblasts and the Cell Cycle Myofibroblasts pose a unique conundrum in the world of muscle differentiation. Of particular interest is their ability to express functional MyoD and myogenin and yet remain active in the cell cycle. The fact that myofibroblasts express functional MRFs but continue cell division is an interesting exception to many experimental observations of forced MRF expression (Table II). In most cases, expression of MRFs results in cell cycle withdrawal and morphological differentiation (Cserjesi et al., 1994). Since this aspect of myofibroblast biology has not been investigated, the following is a summary of the issues that may be at play in the resistance of myofibroblasts to MRF-induced cell cycle withdrawal. It has been shown that cell cycle withdrawal and terminal differentiation are temporally separable events in myogenesis. This process was elucidated in C2 C12 myoblasts, where it was determined that myogenin expression preceded the expression of the cyclin-dependent kinase inhibitor (CKI) p21 (Andres and Walsh, 1996).

Table II Ability of Nonmuscle Cells to be Converted to Muscle by Ectopic Expression of MyoDa Cell line 10 T1/2 Swiss 3T3 NIH 3T3 L cells BALB/3T3 B78 B16 BNL B50 P19 Ca(Co2) GH3 MEL P3881 CV1 a Not

Cell type

Species

MyHC

Desmin

Stable fibroblast line Stable fibroblast line Stable fibroblast line Stable fibroblast line Stable fibroblast line Melanoma Melanoma Liver cells Neuroblastoma Embryonal carcinoma Colon carcinoma Pituitary gland Erythroleukemia Macrophage Kidney

Mouse Mouse Mouse Mouse Mouse Mouse Mouse Mouse Rat Mouse Human Rat Mouse Mouse Primate

+ + + + + + + + + + − − − − −

+ + + + + + + + + + − − − − −

all nonmuscle cells are susceptible to myogenic conversion by MyoD. The listed cells were stably transfected with a MyoD expressing retrovirus and assayed for MyHC, desmin, and MyoD by immunostaining and Northern analysis (Weintraub et al., 1989; Davis et al., 1987).

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Figure 2 Cell cycle withdrawal and terminal differentiation are depicted here as temporally separable events. Proliferating myoblasts express myogenin upon serum withdrawal and exit the cell cycle. Myogenin positive cells then induce p21 expression, activate muscle specific gene expression, and differentiate into myotubes.

These effects are illustrated in Fig. 2. Proliferating myoblasts express myogenin upon serum withdrawal, and subsequently exit from the cell cycle. At this point, p21 expression is induced, sarcomeric genes such as MyHC are activated, and the cells biochemically and morphologically differentiate into myotubes (Walsh and Perlman, 1997). Despite the presence of myogenin in liver and kidney myofibroblasts, the withdrawal of serum does not appear to activate this sequence of events (unpublished observations). Myoblasts transfected with p21 expression constructs irreversibly withdraw from the cell cycle, even upon serum restimulation (Guo et al., 1995). Although the ability of p21 to initiate G1 arrest and terminal differentiation has been well documented in nonmuscle cell lines, it is apparently not necessary for maintenance of G1 arrest. In human diploid fibroblasts, p21 levels decline once senescence is reached, indicating that p21 is nonessential once a postmitotic state is reached in these cells (Di Cunto et al., 1998). In contrast, p21 expression in terminally differentiated muscle cells remains constant and increases when nonmuscle 10T1/2 fibroblasts stably expressing MyoD are induced to differentiate (Halevy et al., 1995). C2 C12 skeletal myoblasts that fail to express p21 after myogenin induction become sensitive to apoptosis because p21 offers protection against apoptotic cell death (Wang and Walsh, 1996). How myofibroblasts can express MyoD and myogenin and proliferate without any signs of apoptosis is a mystery. Although p21 expression has not been analyzed in myofibroblasts, their ability to proliferate in a myogenin-positive background would suggest that some mechanism is protecting them from apoptotic events and inhibiting cell cycle arrest (Fig. 3). If p21 is expressed, why do myogenin positive myofibroblasts exit the cell cycle like

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Figure 3 Myofibroblasts prosses myogenic activators and muscle-specific molecular markers, indicating a strong potential for terminal differentiation. Forced induction of cyclin-dependent kinase inhibitors such as p21 may lead to cell cycle arrest, ordered arrangement of muscle structural proteins, and myotube formation.

myogenin positive myoblasts? If p21 is not expressed, then what mechanism is acting to protect myofibroblasts from apoptosis? Cell cycle withdrawal in muscle differentiation may involve pathways that are distinct from those in other systems. Andrew Lassar’s group has shown that MyoD expression correlates with p21 expression and terminal differentiation, but that it appears to be independent of p53 expression. Since p53 is a major transcriptional regulator of p21 in response to DNA damage and cellular stress, this suggests that muscle terminal differentiation proceeds via a different pathway (Halevy et al., 1995). The active or hypophosphorylated retinoblastoma protein (pRB) has also been shown to be required for maintenance of postmitotic muscle cells. When pRB is phosphorylated, it releases transcription factors such as E2F, which then activate late G1/early S phase genes. Stem cells and myotubes isolated from pRB −/− mice are able to synthesize DNA upon serum restimulation (Martelli et al., 1994). There is also a higher frequency of apoptosis during the muscle differentiation program in these mice (Wang et al., 1997). It will be interesting to examine the expression and phosphorylation state of pRB in myofibroblasts, to determine its role in myofibroblast cell cycle regulation. Myofibroblasts appear to have functional MyoD and myogenin proteins, as evidenced by the MRF-dependent activity of a reporter gene driven by the MLC 1,3 promoter in transfected BHK and liver stellate cells (Mayer and Leinwand, 1997). It is possible that proteins that bind the MRFs and inhibit terminal differentiation, such as Id1 and Id3 (but not Id2), Twist, and Mist-1 (Lemercier et al., 1998; Leifer et al., 1994), are expressed but unable to fully repress MRF activity. Interestingly,

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Id2 has been shown to bind and inactivate pRb (Lasorella et al., 1996), suppressing its ability to terminally arrest cells. If Id2 is active in myofibroblasts, it might help explain why the MRFs are active, yet continue to proliferate. The question remains: What is keeping myofibroblasts from terminally differentiating? Much remains to be determined in these cells, although it would appear that myofibroblasts in vivo do have a regulatory mechanism that calls them to action during injury, and turns them off when their work is done. It will be interesting to see how this regulation occurs, as it may redefine our ideas regarding myogenisis and help in the treatment of excessive fibrosis during tissue damage.

VIII. Perspectives In summary, it seems that myofibroblasts use unusual molecular mechanisms to control gene expression, cell proliferation, and contractility. Further study of these cells is likely to challenge several well-established paradigms of these processes. Because of their vital role in wound healing and their damaging role in fibrosis, understanding the origin and regulation of myofibroblasts is a worthwhile and important goal.

Acknowledgments A portion of the work described here has been supported by NIH GM29090 to L.A.L. We thank Ghislaine Mayer for sharing unpublished data and Lauren Millette for input into the initial phases of this article.

References Andres, V., and Walsh, K. (1996). Myogenin expression, cell cycle withdrawal, and phenotypic differentiation are temporally separable events that precede cell fusion upon myogenesis. J. Cell Biol. 132, 657–666. Bi, W., Drake, C. J., and Schwarz, J. J. (1999). The transcription factor MEF2C-null mouse exhibits complex vascular malformations and reduced cardiac expression of angiopoietin 1 and VEGF. Dev. Biol. 211, 255–267. Black, B. L., and Olson, E. N. (1998). Transcriptional control of muscle development by myocyte enhancer factor-2 (MEF2) proteins. Annu. Rev. Cell Dev. Biol. 14, 167–196. Black, B. L., Molkentin, J. D., and Olson, E. N. (1998). Multiple roles for the MyoD basic region in transmission of transcriptional activation signals and interaction with MEF2. Mol. Cell Biol. 18, 69–77. Bostrom, H., Willetts, K., Pekny, M., Leveen, P., Lindahl, P., Hedstrand, H., Pekna, M., Hellstrom, M., Gebre-Medhin, S., Schalling, M., Nilsson, M., Kurland, S., Tornell, J., Heath, J. K., and Betsholtz, C. (1996). PDGF-A signaling is a critical event in lung alveolar myofibroblast development and alveogenesis. Cell 85, 863–873.

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Campbell, S. E., and Katwa, L. C. (1997). Angiotensin II stimulated expression of transforming growth factor-beta1 in cardiac fibroblasts and myofibroblasts. J. Mol. Cell Cardiol. 29, 1947–1958. Capetanaki, Y., Milner, D. J., and Weitzer, G. (1997). Desmin in muscle formation and maintenance: knockouts and consequences. Cell Struct. Funct. 22, 103–116. Crescenzi, M., Fleming, T. P., Lassar, A. B., Weintraub, H., and Aaronson, S. A. (1990). MyoD induces growth arrest independent of differentiation in normal and transformed cells. Proc. Natl. Acad. Sci. USA 87, 8442–8446. Cserjesi, P., Lilly, B., Hinkley, C., Perry, M., and Olson, E. N. (1994). Homeodomain protein MHox and MADS protein myocyte enhancer-binding factor-2 converge on a common element in the muscle creatine kinase enhancer. J. Biol. Chem. 269, 16740 –16745. Davis, D. L., Weintraub, H., and Lassar, A. B. (1987). Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell 51, 987–1000. Desmouliere, A., Geinoz, A., Gabbiani, F., and Gabbiani, G. (1993). Transforming growth factor-beta 1 induces alpha-smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J. Cell Biol. 122, 103–111. Di Cunto, F., Topley, G., Calautti, E., Hsiao, J., Ong, L., Seth, P. K., and Dotto, G. P. (1998). Inhibitory function of p21Cip1/WAF1 in differentiation of primary mouse keratinocytes independent of cell cycle control [see comments]. Science 280, 1069–1072. Forbes, J. M., Leaker, B., Hewitson, T. D., Becker, G. J., and Jones, C. L. (1999). Macrophage and myofibroblast involvement in ischemic acute renal failure is attenuated by endothelin receptor antagonists. Kidney Int. 55, 198–208. Gabbiani, G., Ryan, G. B., and Majne, G. (1971). Presence of modified fibroblasts in granulation tissue and their possible role in wound contraction. Experientia 27, 549–550. Gray, M. O., Long, C. S., Kalinyak, J. E., Li, H. T., and Karliner, J. S. (1998). Angiotensin II stimulates cardiac myocyte hypertrophy via paracrine release of TGF-beta 1 and endothelin-1 from fibroblasts. Cardiovasc. Res. 40, 352–363. Grounds, M. D., Garrett, K. L., and Beilharz, M. W. (1992). The transcription of MyoD1 and myogenin genes in thymic cells in vivo. Exp. Cell Res. 198, 357–361. Guo, K., Wang, J., Andres, V., Smith, R. C., and Walsh, K. (1995). MyoD-induced expression of p21 inhibits cyclin-dependent kinase activity upon myocyte terminal differentiation. Mol. Cell Biol. 15, 3823–3829. Halevy, O., Novitch, B. G., Spicer, D. B., Skapek, S. X., Rhee, J., Hannon, G. J., Beach, D., and Lassar, A. B. (1995). Correlation of terminal cell cycle arrest of skeletal muscle with induction of p21 by MyoD [see comments]. Science 267, 1018–1021. Johnson, A. D., and Owens, G. K. (1999). Differential activation of the SMalphaA promoter in smooth vs. skeletal muscle cells by bHLH factors. Am. J. Physiol. 276, C1420–C1431. Koffer, A., and Dickens, M. J. (1987). Isolation and characterization of actin from cultured BHK cells. J. Muscle Res. Cell Motil. 8, 397–406. Lasorella, A., Iavarone, A., and Israel, M. A. (1996). Id2 specifically alters regulation of the cell cycle by tumor suppressor proteins. Mol. Cell Biol. 16, 2570–2578. Lassar, A. B., Davis, R. L., Wright, W. E., Kadesch, T., Murre, C., Voronova, A., Baltimore, D., and Weintraub, H. (1991). Functional activity of myogenic HLH proteins requires heterooligomerization with E12/E47-like proteins in vivo. Cell 66, 305–315. Leifer, D., Golden, J., and Kowall, N. W. (1994). Myocyte-specific enhancer binding factor 2C expression in human brain development. Neuroscience 63, 1067–1079. Lemercier, C., To, R. Q., Carrasco, R. A., and Konieczny, S. F. (1998). The basic helix-loop-helix transcription factor Mist1 functions as a transcriptional repressor of myoD. EMBO J. 17, 1412– 1422. Li, H., and Capetanaki, Y. (1994). An E box in the desmin promoter cooperates with the E box and MEF-2 sites of a distal enhancer to direct muscle-specific transcription. EMBO J. 13, 3580–3589.

106

Gennyne A. Walker et al.

Lindahl, P., Johansson, B. R., Leveen, P., and Betsholtz, C. (1997). Pericyte loss and microaneurysm formation in PDGF-B-deficient mice. Science 277, 242–245. Lindahl, P., Hellstrom, M., Kalen, M., Karlsson, L., Pekny, M., Pekna, M., Soriano, P., and Betsholtz, C. (1998). Paracrine PDGF-B/PDGF-Rbeta signaling controls mesangial cell development in kidney glomeruli. Development 125, 3313–3322. Madeddu, P., Emanueli, C., Maestri, R., Salis, M. B., Minasi, A., Capogrossi, M. C., and Olivetti, G. (2000). Angiotensin II type 1 receptor blockade prevents cardiac remodeling in bradykinin B(2) receptor knockout mice. Hypertension 35, 391–396. Martelli, F., Cenciarelli, C., Santarelli, G., Polikar, B., Felsani, A., and Caruso, M. (1994). MyoD induces retinoblastoma gene expression during myogenic differentiation. Oncogene 9, 3579– 3590. Mayer, D. C., and Leinwand, L. A. (1997). Sarcomeric gene expression and contractility in myofibroblasts. J. Cell Biol. 139, 1477–1484. Molkentin, J. D., and Olson, E. N. (1996). Combinatorial control of muscle development by basic helix-loop-helix and MADS-box transcription factors. Proc. Natl. Acad. Sci. USA 93, 9366–9373. Molkentin, J. D., Black, B. L., Martin, J. F., and Olson, E. N. (1995). Cooperative activation of muscle gene expression by MEF2 and myogenic bHLH proteins. Cell 83, 1125–1136. Nabeshima, Y., Hanaoka, K., Hayasaka, M., Esumi, E., Li, S., Nonaka, I., and Nabeshima, Y. (1993). Myogenin gene disruption results in perinatal lethality because of severe muscle defect [see comments]. Nature 364, 532–535. Olson, E. N., Arnold, H. H., Rigby, P. W., and Wold, B. J. (1996). Know your neighbors: three phenotypes in null mutants of the myogenic bHLH gene MRF4. Cell 85, 1–4. Powell, D. W., Mifflin, R. C., Valentich, J. D., Crowe, S. E., Saada, J. I., and West, A. B. (1999). Myofibroblasts. I. Paracrine cells important in health and disease. Am. J. Physiol. 277, C1–C9. Ramadori, G., Veit, T., Schwogler, S., Dienes, H. P., Knittel, T., Rieder, H., and Meyer zum Buschenfelde, K. H. (1990). Expression of the gene of the alpha-smooth muscle-actin isoform in rat liver and in fat-storing (ITO) cells. Virchows Arch. B Cell Pathol. Incl. Mol. Pathol. 59, 349–357. Rao, M. V., Donoghue, M. J., Merlie, J. P., and Sanes, J. R. (1996). Distinct regulatory elements control muscle-specific, fiber-type-selective, and axially graded expression of a myosin light-chain gene in transgenic mice. Mol. Cell Biol. 16, 3909–3922. Redfield, A., Nieman, M. T., and Knudsen, K. A. (1997). Cadherins promote skeletal muscle differentiation in three-dimensional cultures. J. Cell Biol. 138, 1323–1331. Rhodes, S. J., and Konieczny, S. F. (1989). Identification of MRF4: a new member of the muscle regulatory factor gene family. Genes Dev. 3, 2050–2061. Rice, A. B., Moomaw, C. R., Morgan, D. L., and Bonner, J. C. (1999). Specific inhibitors of plateletderived growth factor or epidermal growth factor receptor tyrosine kinase reduce pulmonary fibrosis in rats. Am. J. Pathol. 155, 213–221. Rockey, D. (1997). The cellular pathogenesis of portal hypertension: stellate cell contractility, endothelin, and nitric oxide. Hepatology 25, 2–5. Rockey, D. C., and Chung, J. J. (1996). Endothelin antagonism in experimental hepatic fibrosis. Implications for endothelin in the pathogenesis of wound healing. J. Clin. Invest. 98, 1381–1388. Rosenkranz, S., and Kazlauskas, A. (1999). Evidence for distinct signaling properties and biological responses induced by the PDGF receptor alpha and beta subtypes. Growth Factors 16, 201–216. Rosenthal, N., Kornhauser, J. M., Donoghue, M., Rosen, K. M., and Merlie, J. P. (1989). Myosin light chain enhancer activates muscle-specific, developmentally regulated gene expression in transgenic mice. Proc. Natl. Acad. Sci. USA 86, 7780–7784. Rudnicki, M. A., Schnegelsberg, P. N., Stead, R. H., Braun, T., Arnold, H. H., and Jaenisch, R. (1993). MyoD or Myf-5 is required for the formation of skeletal muscle. Cell 75, 1351–1359. Sakai, S., Miyauchi, T., Kobayashi, M., Yamaguchi, I., Goto, K., and Sugishita, Y. (1996). Inhibition of myocardial endothelin pathway improves long-term survival in heart failure. Nature 384, 353–355.

3. Myofibroblasts

107

Schaart, G., Pieper, F. R., Kuijpers, H. J., Bloemendal, H., and Ramaekers, F. C. (1991). Baby hamster kidney (BHK-21/C13) cells can express striated muscle type proteins. Differentiation 46, 105–115. Schurch, W., Seemayer, T. A., and Gabbiani, G. (1998). The myofibroblast: a quarter century after its discovery [editorial]. Am. J. Surg. Pathol. 22, 141–147. Steinert, P. M., and Roop, D. R. (1988). Molecular and cellular biology of intermediate filaments. Annu. Rev. Biochem. 57, 593–625. Sun, Y. (1997). Local angiotensin II and myocardial fibrosis. Adv. Exp. Med. Biol. 432, 55–61. Tapscott, S. J., Thayer, M. J., Weintraub, H. (1993). Deficiency in rhabdomyosarcomas of a factor required for MyoD activity and myogenesis. Science 259, 1450–1453. Tuszynski, G. P., Frank, E. D., Damsky, C. H., Buck, CA., and Warren, L. (1979). The detection of smooth muscle desmin-like protein in BHK21/C13 fibroblasts. J. Biol. Chem. 254, 6138–6143. van de Klundert, F. A., Jansen, H. J., and Bloemendal, H. (1994). A proximal promoter element in the hamster desmin upstream regulatory region is responsible for activation by myogenic determination factors. J. Biol. Chem. 269, 220–225. van der Ven, P. F., and Furst, D. O. (1998). Expression of sarcomeric proteins and assembly of myofibrils in the putative myofibroblast cell line BHK-21/C13. J. Muscle Res. Cell Motil. 19, 767–775. Walsh, K., and Perlman, H. (1997). Cell cycle exit upon myogenic differentiation. Curr. Opin. Genet. Dev. 7, 597–602. Wang, J., and Walsh, K. (1996). Resistance to apoptosis conferred by Cdk inhibitors during myocyte differentiation. Science 273, 359–361. Wang, J., Guo, K., Wills, K. N., and Walsh, K. (1997). Rb functions to inhibit apoptosis during myocyte differentiation. Cancer Res. 57, 351–354. Weber, K. T. (1997). Metabolic responses of extracellular matrix in tissue repair. Ann. Med. 29, 333– 338. Weintraub, H., Tapscott, S. J., Davis, R. L., Thayer, M. J., Adam, M. A., Lassar, A. B., and Miller, A. D. (1989). Activation of muscle-specific genes in pigment, nerve, fat, liver, and fibroblast cell lines by forced expression of MyoD. Proc. Natl. Acad. Sci. USA 86, 5434–5438. Wentworth, B. M., Donoghue, M., Engert, J. C., Berglund, E. B., and Rosenthal, N. (1991). Paired MyoD-binding sites regulate myosin light chain gene expression. Proc. Natl. Acad. Sci. USA 88, 1242–1246. Wheeler, M. T., Snyder, E. C., Patterson, M. N., and Swoap, S. J. (1999). An E-box within the MHC IIB gene is bound by MyoD and is required for gene expression in fast muscle. Am. J. Physiol. 276, C1069–C1078. White, S. L., and Low, R. B. (1996). Identification of promoter elements involved in cell-specific regulation of rat smooth muscle myosin heavy chain gene transcription. J. Biol. Chem. 271, 15008– 15017. Woodcock-Mitchell, J., Mitchell, J. J., Low, R. B., Kieny, M., Sengel, P., Rubbia, L., Skalli, O., Jackson, B., and Gabbiani, G. (1988). Alpha-smooth muscle actin is transiently expressed in embryonic rat cardiac and skeletal muscles. Differentiation 39, 161–166. Wu, L. L., Yang, N., Roe, C. J., Cooper, M. E., Gilbert, R. E., Atkins, R. C., and Lan, H. Y. (1997). Macrophage and myofibroblast proliferation in remnant kidney: role of angiotensin II. Kidney Int. Suppl. 63, S221–S225. Yokoi, Y., Namihisa, T., Kuroda, H., Komatsu, I., Miyazaki, A., Watanabe, S., and Usui, K. (1984). Immunocytochemical detection of desmin in fat-storing cells. (Ito cells). Hepatology 4, 709–714.

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4 Checkpoint and DNA-Repair Proteins Are Associated with the Cores of Mammalian Meiotic Chromosomes Madalena Tarsounas ∗ and Peter B. Moens Department of Biology York University Toronto, Ontario, M3J 1P3 Canada I. Introduction II. Structural Characteristics of Meiotic Chromosomes during the Prophase of Meiosis I A. Core Formation B. Meiotic Chromosome Synapsis C. SC Dissolution III. Meiotic Checkpoint and Recombination Proteins Are Associated with the Cores of the Meiotic Chromosomes A. The HR Model B. Recombination Proteins C. Checkpoint Proteins IV. Conclusions and Perspectives References

Meiotic checkpoints are manifested through protein complexes capable of detecting an abnormality in chromosome metabolism and signaling it to effector molecules that subsequently delay or arrest the progression of meiosis. Some checkpoints act during the first meiotic prophase to monitor the repair of chromosomal DSBs, predominantly by meiotic recombination, or to ensure the correct establishment of synapsis and its well-timed dissolution. In mammals, a number of checkpoint and repair proteins localize to the meiotic chromosomal cores, sometimes in the context of the synaptonemal complex (SC).1 Here we discuss possible functions of these proteins in the accomplishment of meiotic recombination and normal progression of the meiotic pathway. Also, we present arguments for a structural role of cores and SCs in the assembly of the repair and checkpoint protein complexes on the chromosomes.  2001 Academic Press. C

∗ Present

address: Imperial Cancer Research Fund, Clare Hall Laboratories, South Mimms, Herts, EN6 3LD, England. 1 Abbreviations used in the text: dsDNA, double stranded DNA; DSB, double stranded break; EM, electron microscopy; HR, homologous recombination; IR, ionizing radiation; LE, lateral element; NHEJ, nonhomologous end joining; SC, synaptonemal complex; SMC, structural maintenance of the chromosomes; ssDNA, single-stranded DNA; TF, transverse filament. Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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I. Introduction Meiosis is a special type of cell division in which two rounds of chromosome segregation follow one round of chromosome replication. Premeiotic DNA replication creates two copies of each parental chromosome (Fig. 1, see color insert). Each copy, termed homologous chromosome or homolog, consists of a pair of sister chromatids. During the prophase of meiosis I, connections are established between homologs and the resulting structure is referred to as a bivalent. At this stage, the synaptonemal complex, an assembly of nucleic acids and proteins positioned along the length of the homologs, brings the maternal and paternal chromosomes together, facilitating their recombination. The process of meiotic recombination accounts for the genetic variability in the germline and the phenotypic diversity of the progeny. The first meiotic division is reductional, that is, the two chromosomes in a bivalent, each consisting of a pair of sister chromatids, migrate to opposite poles. Sister chromatid and sister kinetochore cohesion are essential at this stage. At meiosis II (the equational division) the two sister chromatids in each chromosome separate and segregate to opposite poles. Overall, four haploid cells are generated from one diploid parental cell. When they become capable of undergoing fertilization, these haploid cells are identified as gametes. During sexual reproduction, fusion of two gametes of opposite mating types restores diploidy. The changes in chromosome morphology accompanying the progression through the meiotic pathway are usually referred to as the meiotic chromosome metabolism. Among them, recombination of meiotic chromosomes has a unique evolutionary significance, as it creates novel gene combinations in the progeny. Through natural selection, the genes that translate into successful survival and reproduction traits tend to spread within populations, ultimately leading to the appearance of new species. Therefore, meiosis can be viewed as the process that fueled evolution. In this context, it is obvious that evolutionary pressure for an error-free meiotic chromosome metabolism must have led to the development of rigorous surveillance mechanisms that monitor the correct accomplishment of each step of meiosis. Homologous recombination (HR) of the meiotic chromosomes is initiated during early meiotic prophase I with the formation of double stranded breaks (DSBs) in the chromosomes, which are repaired synchronously with the exchange of genetic information between homologous chromosomes. Elaborate activities for the repair of DSBs by HR have evolved, along with stringent checkpoint mechanisms that monitor repair and suppress errors. In most species, the role of these mechanisms is mainly to delay the transition from prophase I to subsequent meiotic stages (in mammals, the prophase I is the lengthiest stage of meiosis), in order to allow correct processing of the HR intermediates into the final recombination products. If an error exists, the checkpoints can arrest the progression of meiosis at the stage where the defect is detected.

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Many of the meiotic recombination and checkpoint activities are accomplished by proteins originating in somatic cells, from which they have probably been recruited during evolution into the meiotic process. Supporting this view is the fact that mutations in these genes cause genomic instability, massive cell death, and cancer due to DSB repair failure, and frequently, sterility due to impaired repair of meiotic DSBs. We summarize here some of the recent research on the structure of meiotic chromosomes during prophase I, as well as on the meiotic chromosome-associated proteins participating in the repair meiotic DSBs, or in the surveillance of meiotic progression. The focus of this review is on the structure of mammalian meiotic chromosomes and on the proteins associated with them that play repair and surveillance roles in the maintenance of genomic integrity.

II. Structural Characteristics of Meiotic Chromosomes during the Prophase of Meiosis I Following DNA replication, a genome-wide search for homology occurs and, as a result, homologous chromosomes align with each other side-by-side, a configuration that is stabilized by transient paranemic DNA–DNA interactions (Weiner and Kleckner, 1994). This process is termed homolog pairing and it precedes synapsis, the stage at which the homologs are intimately associated in the context of the SC. In most organisms, pairing is initiated at leptotene, the first structurally defined stage of prophase I (Fig. 2, see color insert), but despite intensive investigation, very little molecular detail has been accumulated on this process (for reviews, see Roeder, 1997; Zickler and Kleckner, 1998). The presence of certain proteins such as Saccharomyces cerevisiae Hop2 has been shown to be required for homolog pairing: In its absence, normal levels of synapsis are detected, but this is entirely established between nonhomologous chromosomes (Leu et al., 1998). This phenotype indicates that ectopic DNA interactions may occur during prophase I, and specialized proteins are required to prevent pairing and synapsis between nonhomologous chromosomes. In mammals, such a role may be played by the recombination protein DMC1, as the spermatocytes of Dmc1−/− mice sometimes exhibit nonhomologous synapsis (Yoshida et al., 1998).

A. Core Formation Simultaneously with chromosome pairing, proteinaceous cores (also termed axial elements or chromosome axes) form along each homolog. In rodents and probably other eukaryotes, the cores represent anchorage sites for the chromatin loops (Heng

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et al., 1994, 1996; Moens et al., 1998). Sequencing of the DNA present in the rodent SCs has demonstrated an enrichment in LINE and SINE fragments (Pearlman et al., 1992). Biochemical evidence for the DNA-binding role of the cores was obtained from recent studies in yeast indicating the ability of the core component Hop1 (see below) to bind duplex DNA in vivo and in vitro (Kironmai et al., 1998). At synapsis, the cores form the lateral elements (LEs) of the SC (Fig. 2). It is postulated that they may represent the structures to which the transverse filaments (TFs) attach in order to effect synapsis. Two protein components of the cores are known in the yeast S. cerevisiae: Hop1 and Red1 (Hollingsworth and Byers, 1989; Rockmill and Roeder, 1990). They colocalize on the prophase I chromosomes, where they are found in the unsynapsed cores and mature SCs. Red1 and Hop1 dissociate from the chromosomes as the SC disassembles (Smith and Roeder, 1997). Red1 is required for LE assembly (Rockmill and Roeder, 1990), as well as for the assembly of Hop1 on the chromosomes (Smith and Roeder, 1997). Therefore it is postulated that Red1 nucleates the formation of axial elements in yeast (Roeder, 1997). red1 and hop1 mutants lack SCs and exhibit residual levels of recombination and chromosome nondisjunction, which causes severe spore inviability (Hollingsworth and Byers, 1989; Rockmill and Roeder, 1990; Mao-Draayer et al., 1996). This points to a major role for the correct core formation in recombination and the successful completion of meiosis. Remarkably, the core components in budding yeast have no similarity at the amino acid level with potential functional homologs from other eukaryotes. This conclusion has been based initially on the observed lack of cross-reactivity of polyclonal antibodies between eukaryotic species (S. cerevisiae, Drosophila, Xenopus; P. B. Moens, unpublished) and was subsequently confirmed by the identification of core protein components in a number of different species and comparison of their amino acid sequences. Determining the protein composition of the cores in rodents has represented a challenge for quite some time because of difficulties in the isolation of these chromosome-associated proteins. This work was pioneered by Heyting et al. (1985) with the isolation of SCs from rat spermatocytes. The SCs have been injected in mice and rabbits, and the antibodies generated recognized the cores and the SC when tested by indirect immunofluorescence (Moens et al., 1987; Heyting et al., 1988; Moens et al., 1992). In immunological screenings of expression libraries from rodent testis, the cDNAs encoding the core components COR1(SCP3)2 (Dobson et al., 1994; Lammers et al., 1994) and SCP2 (Offenberg et al., 1998), as well as the synaptic protein SYN1(SCP1) (Dobson et al., 1994), were isolated. SCP2 was also isolated in a two-hybrid screen of a testis cDNA 2 This protein component of the cores is termed COR1 in hamster and SCP3 in rat. Similarly, the synaptic protein is termed SYN1 in hamster and SCP1 in rat. Here we use a compound name that includes both denominations for each protein.

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library with COR1(SCP3) as a bait (Tarsounas et al., 1999b). These three proteins are expressed exclusively during meiosis. The distribution of COR1(SCP3) in the chromosome cores and LEs (Fig. 2) was studied with specific antibodies raised against the full-length protein (Dobson et al., 1994; Lammers et al., 1994). COR1(SCP3), a Mr 30,000/33,000 protein, appears as short segments on the chromosomes at leptotene. The staining becomes continuous at zygotene when it clearly marks the cores in the process of alignment. At pachytene, the two homologous cores are in close juxtaposition and the COR1(SCP3) staining identifies bivalents, rather than individual homologs at the light microscopy level of resolution. At diplotene, anti-COR1(SCP3) antibodies stain the separating cores and clearly mark chiasmatic configurations. Consistent with an essential role for COR1(SCP3) in forming the cores of the meiotic chromosomes, the Cor1(Scp3)−/− mice lack the potential to assemble fully developed SCs (Yuan et al., 2000). The continuous presence of COR1(SCP3) along the chromosomes at all prophase I stages and its disappearance at the first meiotic division indicate a possible role in sister chromatid cohesion. In addition, COR1(SCP3) is detected at anaphase II in the space between the separating kinetochores (Moens and Spyropoulos, 1995). Its persistence at the kinetochores until anaphase II suggests a possible role in establishing and maintaining sister kinetochore cohesion until sister chromatid separation, a role similar to that of Drosophila Mei-S332 (Kerrebrock et al., 1995; Moore et al., 1998) and Schizosaccharomyces pombe cohesin comples (Tanaka et al., 1999). COR1(SCP3) participates in homotypic interactions in a two-hybrid system and in vitro that may be mediated by coiled-coil formation (Tarsounas et al., 1997). This observation suggested that COR1(SCP3) may assemble multimeric aggregates on the chromosomes. This seems indeed to be the case, as Yuan et al. (1998) have shown that the mouse homolog of COR1(SCP3), termed SCP3, assembles multistranded fibers in the nucleus and cytoplasm of fibroblasts in culture. A region with coiled-coil forming potential is required for the assembly of COR1(SCP3) fibers, which confirmed the importance of this region for the COR1(SCP3) homotypic interactions (Tarsounas et al., 1997). These fibers are structurally related to the intermediate filaments; hence the conclusion that COR1(SCP3) assembly occurs with the formation of filamentous structures along the chromosomes. COR1(SCP3) assembly most likely does not require the presence of DNA because COR1(SCP3) filaments can form in the cytoplasm of the transfected cells (Yuan et al., 1998) and cytoplasmic poly-SC complexes are detected in several organisms (Goldstein, 1987). However, it does not exclude the possibility that these filaments have DNA binding ability, as the chromatin loops appear embedded in the chromosomal cores at the level of resolution of immunofluorescence microscopy (Heng et al., 1997). Studies of chromatin organization in the Cor1(Scp3)−/− mice are currently in progress.

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A second component of the chromosome cores in rat spermatocytes has been characterized (Offenberg et al., 1998). This large protein, termed SCP2 (Mr 190,000), has a distribution pattern on the chromosomes similar to COR1(SCP3) (Offenberg et al., 1998; Schalk et al., 1998). The C-terminal fragment of the hamster homolog of the SCP2 protein has also been isolated in a two-hybrid screen of a hamster cDNA library using COR1(SCP3) as a bait (Tarsounas et al., 1999b). This fragment possesses coiled-coil forming ability, which demonstrates that this motif is important for the interaction between the COR1(SCP3) and SCP2 proteins as well as for homotypic COR1(SCP3) interactions. It is possible therefore that the two proteins form a structural complex of the meiotic chromosome cores. SCP2 and COR1(SCP3) may form individual filaments that assemble into a higher order structure. Alternatively, they may form mixed filaments in which SCP2 and COR1(SCP3) monomers assemble into the cores lining the chromosomes, possibly with a defined stoichiometry of the two monomers. The second scenario seems more probable, in the light of the observation that Cor1(Scp3)−/− mice do not form axial elements (Yuan et al., 2000). Studying the distribution of the SCP2 protein in these knockout mice will bring further insight into this matter. Similarly to COR1(SCP3) and SCP2 proteins, the yeast proteins Red1 and Hop1, which are both structural components of the yeast chromosome cores (Hollingsworth and Byers, 1989; Rockmill and Roeder, 1990; Mao-Draayer et al., 1996), interact with themselves and with each other in the axial elements (Hollingsworth and Ponte, 1997). Moreover, overexpression of Red1 has a negative effect on spore viability because a strict Red1/Hop1 stoichiometry is required for the axial element assembly and normal chromosome segregation at meiosis (Friedman et al., 1994). In the model proposed by Hollingsworth and Ponte (1997), the chromosomal cores assemble from Red1 and Hop1 monomers, and phosphorylation by Mek1 kinase is required for maintaining the structural stoichiometry. We entertain the possibility that although individual components do not share sequence similarity, the overall structure of the cores/LEs is evolutionarily conserved from yeast to mammals. A new dimension in the structure of meiotic chromosomes has been introduced by the discovery of structural maintenance of chromosomes (SMC) proteins in association with meiotic chromosomes. SMCs represent a family of heterodimeric proteins with established mitotic roles in chromosome condensation and gene dosage compensation (SMC2 and SMC4), and sister chromatid cohesion (SMC1 and SMC3; reviewed by Strunnikov, 1998; Hirano, 1999). In the yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe, the protein complex required for sister chromatid cohesion during mitosis, termed cohesin, is also functional at meiosis, where it plays a role in sister chromatid cohesion, core formation, and recombination, but at least one of its components, Rec8 is meiosis-specific (reviewed by Orr-Weaver, 1999). Eijpe et al. (2000) have found that the heterodimer SMC1/SMC3 is associated with the cores/SCs of the rat meiotic chromosomes, supporting the role previously

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proposed for cores/SCs in sister chromatid cohesion throughout prophase I of meiosis (Bickel and Orr-Weaver, 1996). The direct interactions determined between SMC1/SMC3 and core/SC components SCP1 and SCP2 (Eijpe et al., 2000) suggest that SMCs are required for the core assembly. Also, mammalian SMCs may bind chromatin directly and anchor it to the axial element of the meiotic chromosomes, similarly to their proposed mitotic function (Hirano, 1999). A role for mammalian SMCs in meiotic recombination is expected by analogy with yeast (Eijpe et al., 2000).

B. Meiotic Chromosome Synapsis Synapsis is accomplished when the axial elements delineating each homolog come in close proximity and become connected by the transverse filaments located in the central region of the SC (Fig. 2). The SYN1(SCP1) protein in rodents, also called the synaptic protein, has a critical role in establishment of chromosome synapsis (Meuwissen et al., 1992; Dobson et al., 1994; Schmekel et al., 1996). SYN1(SCP1) is thought to be the molecular component of the TFs that closes as a zipper at synapsis (Fig. 2). SC assembly initiates at zygotene, synchronously with the nuclear appearance of SYN1(SCP1) in regions where the homologs are intimately connected. These regions, which are positioned randomly along the length of rodent chromosomes, will extend to form fully synapsed chromosomes at the pachytene stage (Meuwissen et al., 1992; Dobson et al., 1994). Epitope mapping of the Mr 125,000 SYN1(SCP1) with electron microscopy (EM) revealed that its C terminus is located in the LE and the N terminus in the center of the SC (Fig. 2; Dobson et al., 1994; Schmekel et al., 1996; Liu et al., 1996). A similar configuration has been demonstrated for Zip1, a possible functional homolog of SYN1(SCP1) in yeast (Tung and Roeder, 1998). This supports the assumption that the TFs, cytologically positioned in the central region and perpendicular to the LE, may be formed by individual SYN1(SCP1) or Zip1 molecules. The N-terminal 100 amino acids of the SYN1(SCP1) protein, which lack the potential of forming coiled-coil motifs, self-interact in a two-hybrid system (Liu et al., 1996). Regions mapping to the middle and C-terminus of the SYN1(SCP1) molecule with high probability of forming coiled-coils do not participate in homotypic interactions (Tarsounas et al., 1997). This distribution supports the idea that SYN1(SCP1) forms the TFs in the central region of the SC, positioned perpendicularly to the LEs and connected in the centre of the SC by direct interactions between the N termini of molecules positioned opposite each other (Fig. 2). Detection of such interactions in a two-hybrid system, however, does not exclude the possibility that other proteins may act as adapters between the N-terminal regions of SYN1(SCP1) in the central region of the SC. A two-hybrid interaction was detected between full-length mouse SYN1(SCP1) molecules, but not between full-length SYN1(SCP1) and COR1(SCP3) (Tarsounas

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et al., 1997), which suggests that most likely COR1(SCP3) is unlikely to be required for the anchorage of SYN1(SCP1)-formed TFs to the lateral regions of the SC. As Cor1(Scp3)−/− spermatocytes are incapable of assembling SYN1(SCP1) and the central region correctly (Yuan et al., 2000), it is possible that the severe general defect in core formation in these cells also affect SYN1(SCP1) positioning on the chromosomes. An as-yet unknown component of the SC may act as an adapter between the TFs and the lateral region of the SC, and its localization may be dependent upon COR1(SCP3) filament formation. In budding yeast, the Zip1 protein has been shown to be a central region component (Sym et al., 1993, 1995) with similar meiotic functions as SYN1(SCP1). In addition, a second protein termed Zip2 is required for establishment of synapsis in yeast. Analysis of Zip1 and Zip2 distributions in the zip2 and zip1 mutants, respectively, suggest that Zip2 plays a structural role, being required for the assembly of Zip1 on the chromosomes (Chua and Roeder, 1998). Zip2 has been shown to localize to the “axial associations,” which are bridgelike structures connecting the LE at a few sites along their length before synapsis, and may also represent recombinogenic sites as suggested by the presence of Rad50 protein (Chua and Roeder, 1998). This finding supports the assumption that recombination intermediates are required for initiation of synapsis. Consistent with this, a spo11 mutant with a DSB formation defect does not assemble SCs (Cha et al., 2000). In mammals, only a limited number of SC components have been identified thus far, and no protein sharing the structural features of Zip2 is yet known. The direct interaction detected between RAD51 and SYN1(SCP1), as well as the presence of the RAD51/DMC1 complexes between the homologous axes before synapsis (Tarsounas et al., 1999b), suggest that RAD51 may recruit SYN1(SCP1) at the synapsis initiation sites, from where synapsis subsequently extends along the chromosomes.

C. SC Dissolution SC disassembly starts during late pachytene with the gradual removal of SYN1(SCP1) from the separating chromosome cores (Tarsounas et al., 1999a). At early diplotene, only short stretches of SYN1(SCP1) remain visible along the chromosomes, and they disappear by late diplotene (Moens and Spyropoulos, 1995). The COR1(SCP3) protein remains in the axes of the post-prophase I chromosomes until the first meiotic division, and it is lost thereafter from this location. COR1(SCP3) is last detected at the separating anaphase II kinetochores (Moens and Spyropoulos, 1995). The nature of the signal that triggers the dissolution of the SC has not yet been established. It is very likely that the molecules that monitor meiotic progression will transmit a signal to the structural blocks that form the SCs once meiotic recombination has been successfully completed. This may cause SC components to

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gradually dissociate from the chromosomes and probably be targeted for degradation thereafter. One possibility is that phosphorylation triggers this series of events. Several potential protein kinase target sites have been revealed by analysis of the amino acid sequences of the three mammalian SC components COR1(SCP3), SYN1(SCP1), and SCP2. Direct evidence that the SC components COR1(SCP3) and SYN1(SCP1) are phosphoproteins in pachytene spermatocyte extracts has been reported by Lammers et al. (1995) and Tarsounas et al. (1999a). Alternative mechanisms such as ubiquitination (Tarsounas et al., 1997) should also be investigated.

III. Meiotic Checkpoint and Recombination Proteins Are Associated with the Cores of the Meiotic Chromosomes A concrete structural function for the SC during meiosis has not yet been established in mammalian cells. One hypothesis addressed in a review by Moens et al. (1998) is that the cores represent attachment sites for the chromatin loops, thus contributing to the general organization of the meiotic chromosomes. Following premeiotic replication, the bases of the loops of the two sister chromatids are attached to the protein cores (Fig. 2). The central SC region, which assembles later in meiosis, then establishes the physical connection between the homologs, possibly creating a steric environment favorable for recombination events to take place (reviewed in Roeder, 1997). Increasing evidence supports a novel role of the cores/SCs in HR by providing the structural matrix for the assembly of DSB repair complexes. Meiotic recombination involves the formation of DSBs along the length of the chromosomes and their repair with concomitant exchange of genetic information between the maternal and paternal chromosomes. Much of what we currently know about meiotic HR actually comes from studies in the budding yeast, S. cerevisiae (reviewed by Smith and Nicolas, 1998). Yeast is a very popular model, mainly because it offers the facility to induce meiosis (upon removal of essential nutrients) and to obtain synchronous populations of cells at various meiotic stages in which HR can be monitored both biochemically and cytologically (Padmore et al., 1991). Generation of single and multiple mutants in HR genes helps in gaining insight into their function (Kleckner, 1996; Roeder, 1997). In yeast, chromosome synapsis was assessed in recombination- or sporulation-deficient mutants, and vice versa, and a correlation has been established between the two. In most eukaryotes, chromosome synapsis is required for normal levels of recombination. Conversely, SC formation may require at least some of the recombination steps to be correctly completed (Bishop, 1994; Rockmill et al., 1995; Nairz and Klein, 1997). The fission yeast S. pombe represents an exception to this rule because it lacks SCs, but exhibits normal levels of meiotic recombination. However, structures resembling the cores have been detected, and these may be sufficient for the meiotic recombination to

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occur (Scherthan et al., 1994). The lack of synapsis correlates with the absence of crossover interference in this organism (Kohli and Bahler, 1994). Similar functional studies in mammals involve generation of knockout mice and are very laborious. In addition, some knockouts are embryonic lethal (e.g., Rad51−/−); therefore, a meiotic phenotype is not available. Indirect evidence comes from biochemical studies of mammalian enzymatic activities in vitro, protein–protein interactions, and the patterns of immunolocalization of these proteins on the meiotic chromosomes. Recent data suggest that proteins directly involved in recombination and in monitoring repair of meiotic DSBs by HR are associated with the meiotic chromosomes. These are primarily proteins involved in DNA damage repair and damage-induced checkpoint control during the cell cycle. These proteins are presumably recruited to perform similar repair and surveillance functions during meiosis. It is particularly interesting that their association with meiotic chromosomes does not occur randomly: These proteins are specifically positioned on the cores of meiotic chromosomes, where the chromatin loops also attach. This led to the speculation that crucial meiotic events such as HR occur within the context of the SC, between DNA sequences located at the base of the chromatin loops. The recently reported phenotype of Cor1(Scp3)−/− mice supports this hypothesis. The absence of COR1(SCP3) changes the nuclear distribution of repair and recombination proteins such RAD51 and RP-A (Yuan et al., 2000), suggesting a structural role for the axial elements/SCs in HR. We present here the immunolocalization patterns on the mammalian meiotic chromosomes reported for some recombination and checkpoint proteins, and the functional aspects inferred from them. It is worth mentioning that although the data summarized here have been obtained exclusively from examination of spermatocyte nuclei, it is generally assumed that similar events governed by similar proteins occur during female meiosis.

A. The HR Model The working model for the succession of recombination events in eukaryotes predicts the formation of DSBs on one of the homologous chromosomes (Szostak et al., 1983; Fig. 3, see color insert). In experiments using the well-characterized HIS4-LEU2 DSB “hot-spot” (Cao et al., 1990) in S. cerevisiae, Padmore et al. (1991) showed the correlation between chromosome synapsis and the succession of recombination events leading to the repair of meiotic DSBs. Evidence for the generation of meiotic DSBs in organisms other than yeast is currently lacking. Despite this limitation, it is assumed that the meiotic recombination model based on DSBs is valid among all eukaryotes. DSBs appear prior to or concomitant with the initiation of axial core assembly on the chromosomes at the leptotene stage of prophase I, and they persist until pachytene when the SC is fully assembled on the chromosomes. DSBs represent

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substrates for an exonuclease activity that resects the DSBs leaving 3 singlestranded DNA (ssDNA) tails. These ssDNA tails, which represent binding sites for RecA homologs, may have the ability to assess sequence identity or similarity in another DNA molecule and to invade an intact DNA duplex on a homologous chromosome, leading to the formation of a Holliday junction. This may coincide with initiation of chromosome synapsis. Following branch migration, this intermediate structure is resolved by cleavage and ligation activities, resulting in a crossover or noncrossover product. These recombinant products are cytologically visible at the end of the pachytene stage when desynapsis initiates, as chiasmata, sites of constriction along the length of the chromosomes.

B. Recombination Proteins 1. SPO11 The catalytic subunit of the meiosis-specific enzymatic activity that produces DSBs was originally identified in yeast as Spo11, a member of a conserved topoisomerase family represented in fission yeast, nematodes, and archaebacteria (Keeney et al., 1997; Bergerat et al., 1997; Dernburg et al., 1998). Identification of Spo11 as the meiotic DNA cleavage activity strongly suggests that meiotic DSBs occur by a topoisomerase-like transesterification reaction, rather than by endonucleolytic hydrolysis. This observation has important consequences for the meiotic progression, a major one being that, since topoisomerase-mediated breaks can be reversed, it is possible to reverse meiotic DSBs in the absence of a suitable recombination partner (Keeney et al., 1997). This reversibility may also explain why only a small fraction of meiotic DSBs lead to successful crossover events. The human and mouse homologs of Spo11 have been identified and found to be transcribed and expressed from the early stages of meiosis (leptotene/zygotene) to mid-pachytene (Keeney et al., 1999; Shannon et al., 1999; Romanienko and Camerini-Otero, 1999), which suggests a role for this protein during the late stages of prophase I as well. Antibodies against mammalian SPO11 preferentially stain the SCs along their length from mid- to late pachytene (P. Baudat and S. Keeney, personal communication). This pattern coincides with the one reported for mammalian topoisomerase II by Moens and Earnshaw (1989). The presence of topoisomerases on the SC at this stage indicate either that DSBs are made in higher eukaryotes at a later stage than in yeast, that is, after synapsis is completed (Shannon et al., 1999), or that topoisomerases play a structural as well as enzymatic role during meiosis. 2. MRE11/RAD50/NSB1 These human proteins have been identified based on their structural (MRE11/ RAD50) and functional (NBS1) homology to the well-characterized yeast Mre11/

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Rad50/Xrs2 repair complex. The three proteins interact with each other forming IR-induced DNA repair complexes, visualized as immunofluorescent foci in the nuclei of somatic cells (Maser et al., 1997). The enzymatic activity of these complexes is mediated by MRE11 (reviewed by Haber, 1998), a nuclease that is essential for genomic stability and cell viability in human cells (Yamaguchi-Iwai et al., 1999). While RAD50 enhances the enzymatic activity of MRE11 (Paull and Gellert, 1998), no concrete function has been assigned to NBS1, except for a putative structural requirement in the assembly of RAD50/MRE11 at sites of DNA damage (Carney et al., 1998). The NBS1 gene has been shown to be mutated in the Nijmegen breakage syndrome (NBS) (Carney et al., 1998; Varon et al., 1998), a chromosomal-instability syndrome whose symptoms include increased cancer incidence, cell cycle checkpoint defects, and ionizing radiation sensitivity; this is the first implication of this protein in a DNA repair pathway. The MRE11/RAD50/NBS1 complex is required for the repair of DSBs by both the HR and the NHEJ pathway in both meiotic and mitotic cells (reviewed by Haber, 1998; Dasika et al., 1999). The nuclease activity of this complex creates 3′ overhangs that may function as binding sites for the proteins involved in strandexchange reactions (Fig. 3). The yeast complex has been proposed to be also active in remodeling the chromatin into a structure that facilitates DSB formation (Usui et al., 1998). At meiosis, mammalian RAD50 and MRE11 show identical temporal and spatial localization patterns, with abundant diffuse nuclear staining at leptotene/zygotene (Goedecke et al., 1999). At this stage both DSB formation and resection are assumed to occur, confirming the assigned role in DSB processing. The fact that the MRE11/RAD50 complexes are not specifically positioned on the meiotic chromosome cores is consistent with the notion that at this stage DSBs occur randomly throughout the nuclear chromatin. The persistence of these complexes to mid-pachytene in association with specialized spermatocyte domains, such as the sex vesicle (Goedecke et al., 1999), has an unknown biological significance. 3. RAD51 and DMC1 These mammalian RecA homologs share a high degree of identity at the amino acid level and exhibit similar catalytic activities in vitro, including the ability to promote homologous DNA pairing and strand transfer reactions (Baumann et al., 1996; Baumann and West, 1997; Li et al., 1997; Masson et al., 1999). RAD51 is expressed in both somatic and meiotic tissues and its deficiency is lethal early in embryogenesis (Lim and Hasty, 1996; Tsuzuki et al., 1996), suggesting an essential function for cell viability. In somatic cells in culture, it is postulated that repair complexes are assembled at the sites of DSBs upon DNA-damage induction, although methods for direct visualization of the DSBs in these cells are not currently available. RAD51 is an essential component of the initial repair complexes, and its multimers can be detected as nuclear foci by indirect immunofluorescence. Many other proteins involved in DNA repair by HR, such as RAD52, RP-A, RAD54,

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BRCA1, and BRCA2, are also recruited to these sites (Dasika et al., 1999). The Dmc1 gene is specifically expressed in meiosis, and its null-mutation in mice causes meiotic arrest at the zygotene stage without homolog synapsis (Pittman et al., 1998) or with occasional synapsis between nonhomologs (Yoshida et al., 1998). This characteristic phenotype substantiates the role of DMC1 in promoting interactions between homologous chromosomes, which appears to be conserved from yeast to mammals (Schwacha and Kleckner, 1997). During the early stages of meiosis, RAD51 and DMC1 proteins have been shown in various species to form complexes visualized by immunofluorescence microscopy as discrete foci along the chromosome cores (Bishop, 1994; Terasawa et al., 1995; Dresser et al., 1997; Anderson et al., 1997; Moens et al., 1997; Barlow et al., 1997a; Tarsounas et al., 1999b). Immunogold EM with antibodies that recognize specifically each protein showed that the two are always present in mixed complexes on the chromosomes. These complexes are most likely formed by direct protein–protein interactions between the two recombinases (and possibly other proteins), as shown in vitro and in a two-hybrid system (Tarsounas et al., 1999b; Masson et al., 1999). Because of their ability to form multimeric filaments on ssDNA in vitro, the RAD51/DMC1 complexes have been proposed to coincide with the sites of resected DSBs (Bishop, 1994). Supporting this assumption is the decrease in the numbers of RAD51/DMC1 foci from leptotene/zygotene to pachytene that correlates with the disappearance of DSB. Therefore, in the early steps of meiotic recombination RAD51 and DMC1 may act directly in the formation of joint molecules by strand-exchange reactions (Fig. 3), consistent with their potential demonstrated in vitro. 4. RP-A Replication protein A (RP-A) is a ssDNA binding protein involved in DNA replication, repair, and recombination in mitotic cells (reviewed by Wold, 1997). The biochemical properties of the heterotrimeric RP-A protein include the potential to enhance efficiency of strand transfer reactions promoted by RecA homologs (reviewed by Baumann and West, 1998). This synergistic action involves RP-A binding to the ssDNA and removal of secondary structures, thus allowing RAD51 to form continuous filaments. Consistent with these biochemical data, RP-A and RAD51 colocalize extensively at discrete foci in meiotic cells and mitotic cells exposed to genotoxic treatments (Fig. 4, see color insert; Plug et al., 1998; Golub et al., 1998). However, in rodent spermatocytes there is a significant number of sites at which the two proteins do not colocalize when visualized by immunofluorescence microscopy (Figs. 4A and 4B) and, more obviously, by immunogold labeling in EM preparations (Figs. 4C and 4D). The most striking example is perhaps the X chromosome of the XY pair in which RAD51/DMC1 complexes are abundant (Fig. 4D), but RP-A cannot be detected. Also, the pseudoautosomal region of the

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XY pair, which is postulated to contain an obligatory crossover and contains at least one site of RAD51/DMC1 accumulation, lacks RP-A staining (Fig. 4D). This differential positioning may be explained by the fact that, at the early pachytene stage, RP-A is abundant on the cores of the meiotic chromosomes and the SCs, practically coating them along their entire length (Fig. 4A). Consequently, RP-A probably competes with RAD51 for the potential ssDNA binding sites, and either displaces RAD51 or delays its assembly. The sites where the two proteins colocalize are probably the recombinogenic sites where RP-A enhances the ability of RAD51 to promote strand exchange, as predicted by the biochemical data. This example of RAD51 and RP-A immunolocalization on meiotic chromosome axes is a striking example of the enhanced resolution obtained when gold grains are used for antigen detection by EM, as opposed to the regular immunostaining techniques. 5. MLH1 A role for homologs of the Escherichia coli mismatch repair apparatus has been demonstrated in the late steps of meiotic recombination, primarily in the correction of mismatches in heteroduplex DNA (Stahl, 1996; Roeder, 1997). Three genes, mutS, mutL, and mutH, are essential for the correction of replication errors in E. coli: MutS recognizes the mismatch and MutH protein acts in a complex with MutS and MutL as an endonuclease that nicks the newly synthesized strand (for a review see Modrich, 1991). In humans, alterations in the products of the mutS homologs, msh2, msh4, and msh5, as well as the mutL homologs mlh1 and pms2 are associated with various forms of cancer (Baker et al., 1995, 1996). The idea that mutations in these mismatch repair genes could have consequences in the germline through perturbation of meiotic events such as genetic recombination and homolog synapsis, was confirmed by gene targeting experiments in mice. Occasional sterility (Mlh1) accompanied by pairing defects, fragmentation of the axial elements of the SC, and nonhomologous synapsis represent the most common abnormalities of the mice carrying null mutations in Pms2 (Baker et al., 1995), Mlh1 (Edelmann et al., 1996; Baker et al., 1996), and Msh5 (Edelmann et al., 1999). Probably the most likely candidate for a protein directly involved in the late steps of meiotic recombination is MLH1, which is the only mismatch repair protein that has been shown to localize on the mouse and human meiotic chromosomes at discrete foci representing late recombination nodules (Baker et al., 1996; Barlow and Hulten, 1998). The late recombination nodules are electron-dense structures, positioned along the SCs and thought to harbor the enzymatic activities required for the late steps of meiotic recombination (Roeder, 1997). These late recombination nodules evolve into chiasmata. In mouse spermatocytes, the MLH1 protein immunolocalizes to an average of 1.2 foci per SC at pachytene (Hassold, 1996), which corresponds to the expected number of chiasmata per bivalent. In addition,

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air-dried human nuclear preparations for light microscopy revealed that the total number of MLH1 foci per nucleus at the mid- to late pachytene stage is comparable to the total number of chiasmata per nucleus (Barlow and Hulten, 1998). The distribution pattern of MLH1 in mouse pachytene spermatocytes is dependent on the SC length and shows crossover interference (Anderson et al., 1999). These observations indicate that MLH1 may indeed mark the sites of crossing over on the meiotic chromosomes. Therefore, MLH1 is thought to promote HR and act synergistically with the recombination machinery of the cell, possibly by suppressing insertion/deletion mispairs introduced during Holliday junction processing. 6. BLM and WRN Whereas RAD51 and DMC1, the eukaryotic homologs of the RecA strand exchange protein of bacteria, have been thoroughly investigated, very little is known about the proteins involved in the late steps of HR, including branch migration and resolution. The BLM and WRN proteins are encoded by genes mutated in the Bloom and Werner syndrome, respectively, two genetic diseases that result in similar cellular and clinical phenotypes, including genomic instability and a high incidence of cancer (German, 1993; Yu et al., 1996). Both proteins have helicase domains and are able to complement the mutation of sgs1 helicase gene in S. cerevisiae (Yamagata et al., 1998), which makes them potential candidates for enzymes involved in the branch migration step of recombination. To date, the WRN protein has not been reported to be associated with the progression of meiosis. Antibodies against this protein do not stain any discernible structures on rodent meiotic chromosomes (P. B. Moens, unpublished data). In mammalian mitotic cells WRN is found in the nucleolus (Marciniak et al., 1998). Two reports (Walpita et al., 1999; Moens et al., 2000) have shown the BLM protein to play a role during meiosis. From the data accumulated thus far, it is not clear whether this role is in promoting or suppressing HR. On meiotic chromosomes, BLM is concentrated in foci that, at the level of resolution of EM, colocalize significantly with the RAD51/DMC1 foci at the zygotene/early pachytene stages of prophase I (Moens et al., 2000). This observation may suggest that BLM acts synergistically to RAD51/DMC1 during HR. In addition, mouse spermatocytes show an excess of BLM protein in the pseudoautosomal region of the XY pair that is highly recombinogenic. Arguments for the role of BLM in suppressing, rather than promoting, HR are based on the hyperrecombination phenotype of BS cells (Watt et al., 1996), which suggests that BLM may prevent heteroduplex formation. The number of DSBs made during early prophase I exceeds the number of recombination products estimated from the number of chiasmata. Therefore, it is possible that helicases such as BLM will act at the noncrossover sites to reverse branch migration and suppress HR. These nonrecombinogenic DSBs can be repaired by alternative pathways,

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such as nonhomologous end joining (NHEJ; reviewed by Critchlow and Jackson, 1998). 7. BRCA1/BRCA2 BRCA1 and BRCA2 are primarily known because germline mutations in the corresponding genes confer a high risk of breast and ovarian cancer in the affected individuals (for a review, see Welcsh et al., 2000). BRCA1 and BRCA2 encode large nuclear proteins with no detectable amino acid sequence homology to each other. However, the two proteins may have synergistic functions in DNA damage repair pathways. Their involvement in DNA repair was primarily suggested by the high sensitivity of BRCA1−/− and BRCA2−/− cells to various DNA-damaging agents (Connor et al., 1997; Sharan et al., 1997; Gowen et al., 1998; Shen et al., 1998; Chen et al., 1998; Patel et al., 1998; Abbott et al., 1999). These cells also exhibit aneuploidy and other chromosomal aberrations (Marcus et al., 1996; Patel et al., 1998) indicative of a role of BRCA1 and BRCA2 in the maintenance of genomic stability and DSB repair (Moynahan et al., 1999). Furthermore, BRCA1 and BRCA2 colocalize with each other upon induction of DNA damage in mitotic cells (Chen et al., 1998) and interact directly with RAD51 (Scully et al., 1997; Sharan et al., 1997; Wong et al., 1997). Both proteins are expressed at high levels in meiotic cells (Zabludoff et al., 1996; Rajan et al., 1997) and colocalize on the unsynapsed cores of meiotic chromosomes during zygotene and on the SCs during pachytene (Chen et al., 1998). The mixed BRCA1/BRCA2 foci in general coincide with RAD51 complexes, suggesting that the three proteins act together in the repair of DSBs during meiotic recombination, but their precise mechanism of action is not known.

C. Checkpoint Proteins 1. ATM and ATR ATM and ATR belong to the PI3 -kinase-related (PIK) protein kinase superfamily (reviewed by Hoekstra, 1997) and are thought to participate in a meiotic and mitotic surveillance mechanism. Mutation in the ATM gene causes the human genetic disorder ataxia telangiectasia (A-T), characterized by chromosomal instability, radiosensitivity and defective cell cycle checkpoint activation. DSBs persist in A-T cells after irradiation, because of impaired HR-mediated DSB repair (Morrison et al., 2000). The best-characterized substrate for the kinase activities of ATM and ATR in mitotic cells is p53 (Canman et al., 1998; Banin et al., 1998; Tibbetts et al., 1999; Lakin and Jackson, 1999), which can be phosphorylated by either kinase in response to different types of DNA damage (ionizing radiation or UV-induced damage, respectively). ATM interaction with dsDNA in vitro is

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enhanced by IR (Suzuki et al., 1999). Subsequently, a model has been proposed in which ATM is capable of detecting the DNA damage directly and signaling it to cell cycle suppressor proteins, such as p53. Other substrates for ATM include RP-A and NBS1, both involved in DSB repair (Dasika et al., 1999), as well as c-Abl, which most likely phosphorylates RAD51 in vivo, inducing its recruitment to DSBs upon IR-induced DNA damage (Chen et al., 1999). The functional significance of these phosphorylation reactions for the cellular response to DNA damage is not yet understood. Atm−/− mice have both mitotic and meiotic defects (Barlow et al., 1996; Xu et al., 1996). These mice exhibit striking defects in both spermatogenesis and oogenesis, which result in infertility. Keegan et al. (1996) reported that ATM is localized specifically to the synapsed chromosomes, a result that could not be duplicated by other laboratories (Barlow et al., 1998). Further experimentation is required to settle these contradictory results. A constant concern in immunocytological work is the reproducibility of staining patterns obtained with different antibodies and by different groups. Polyclonal antibodies tend to have individual specificities that sometimes are not conserved from one bleed to another. To minimize these effects, it is generally recommended that more than one antibody against the same antigen be used. Also, the antibody specificity should be determined in Western blots of cell extracts of the same type as the ones used in immunocytology, and in blocking experiments in which preincubation with the antigen should block the staining pattern generated by the antibody. The absence of ATM produces severe meiotic disruption in mice, with mislocalization of the RAD51/DMC1 and ATR complexes to the chromatin loops (Barlow et al., 1998), and leads to an arrest at the leptotene stage, which is partially reversed in Atm−/− p53−/− double mutant mice (Barlow et al., 1997b). Therefore, ATM may provide a checkpoint function necessary to monitor the events preceding RAD51 assembly on the chromosomes (e.g., the correct level of DSB formation and their resection) to ensure that they are correctly completed before RAD51 assembly is attempted (Barlow et al., 1998). Upon detection of DSBs, an ATM-dependent signal to prevent or delay meiotic progression is transmitted to downstream effector proteins such as p53 (Levine, 1997), probably by direct phosphorylation, and the meiotic progression is halted. In the absence of p53 protein (i.e., Atm−/− p53−/− double knockout), defective meiosis is probably allowed to proceed to a downstream checkpoint where an alternative, p53-independent surveillance pathway becomes functional. The spermatocytes of double knockout mice are thus arrested at pachytene. ATR is an essential component of surveillance mechanisms as indicated by its requirement for survival in human cells following DNA damage, and by the fact that Atr inactivation in mice results in early embryonic lethality (Dasika et al., 1999). The kinase activity of ATR is required during the S-phase of the cellcycle when replication is halted by hydroxyurea treatment, and at the G2/M transition to prevent mitosis in the presence of DNA damage (Cliby et al., 1998).

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Unlike ATM mutant cells, wild-type cells respond to ionizing radiation by inhibiting DNA synthesis. Overexpression of ATR can restore radiation-induced DNA synthesis in ATM−/− cells, arguing that ATR and AMT can substitute for each other in the DNA damage response (Cliby et al., 1998). It has been reported that ATR phosphorylates p53 in vivo and in vitro at a serine residue critical for the function of p53 as a tumor suppressor (Tibbetts et al., 1999). It is possible, therefore, that ATR signals the presence of DNA damage to the tumor suppressor p53, which then blocks cell cycle progression until DNA repair has been completed. During meiosis, ATR localizes to discrete foci along the chromosomal cores during early prophase I stages and it is still present on the SCs at pachytene (Moens et al., 1999). The majority of the ATR localization sites are distinct at the EM level from the RAD51 foci, implying that even if the two proteins may be involved in the same pathway of DNA repair, they do not interact directly. Interestingly, ATR foci are most abundant on the unpaired sections of the homologous chromosomes in the process of synapsis, and decrease in numbers on the synapsed cores where it persists until mid-pachytene. The unsynapsed regions presumably contain the unprocessed DSBs that did not yet interact with the homologous chromosome, and therefore may also identify chromosomal regions with delayed synapsis. This immunolocalization pattern suggests a role for ATR in monitoring not only the early steps of HR, but also the correct establishment of synapsis. Its function may be mediated by signals indirectly received from the RAD51/DMC1 complexes, given that these proteins identify the sites of yet unrepaired DSBs. ATR probably transduces these signals to effector proteins, such as p53, that have the ability to block meiotic progression if required. 2. RAD1 Human RAD1 (HRAD1) has been proposed to be a component of a DNA-damage checkpoint (Freire et al., 1998), based on its homology to the S. cerevisiae RAD17 and S. pombe Rad1+ genes (Lydall and Weinert, 1995; Al-Khodairy and Carr, 1992). Recent observations indicate that hRAD1 protein localizes in foci to both synapsed and unsynapsed chromosome cores, being most abundant during the early stages of prophase I, and that the number of foci decreases from leptotene to pachytene. The overall dynamics is similar to that of RAD51/DMC1, suggesting that the hRAD1-dependent checkpoint monitors the correct fulfillment of the early meiotic DSB processing, including the assembly of the strand exchange complexes. As in the case of ATR, at the EM level of resolution, the hRAD1 and RAD51/DMC1 proteins appear on distinct locations along the chromosome cores (Freire et al., 1998). This observation most likely implies that other proteins are required to bridge the gap between the site of a DSB (presumably identified by RAD51/DMC1 complexes) and the position of the sensor protein (hRAD1). Similarly to ATR, RAD1 may signal the status of DSB repair to effector molecules (Dasika et al., 1999) that delay or arrest meiotic progression.

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IV. Conclusions and Perspectives Repair and checkpoint proteins are not randomly dispersed in the chromatin loops, but form complexes specifically localized along the chromosome cores or SCs. This observation supports the hypothesis that SCs are required to stabilize the assembly of these proteins in close proximity to the DNA or even to facilitate their direct binding to the DNA. It is possible that the SCs are required to maintain the physical connections between the components of these complexes with a dynamic protein composition. As illustrated in Fig. 3, most of the proteins currently known to participate directly in meiotic HR or to monitor HR are abundant on the meiotic chromosomes during the early stages of prophase I (leptptene/zygotene), when DSB are created and the first steps in their repair are thought to occur (Padmore et al., 1991). The presence of checkpoint proteins (e.g., ATR and RAD1) in association with the meiotic chromosomes at these stages, as well as the early meiotic arrest in mouse spermatocytes lacking the surveillance protein ATM, suggests that the first steps in DSB processing are critical for their correct repair. The extensive molecular detail available for these early stages of meiotic HR is not found in the late steps of recombination, which include branch migration and resolution events. Many other genes are probably required for successful completion of meiotic recombination events in mammalian cells. Based on knowledge from the DNA repair field, it is expected that proteins found to be active in DNA damage repair in mammalian mitotic cells will also play some role in meiotic recombination. Meiosis-specific proteins acting directly in meiotic recombination or in its surveillance are also expected to be identified. Possible methods for their identification include complementation analyses with known meiotic yeast mutants, and twohybrid screens of testis or ovary cDNA expression libraries using proteins known to act in meiotic HR as baits. Characterization of the biochemical and cellular functions of these proteins, complemented with the definition of the complex functional interactions among them, represents the next step toward a full understanding of meiotic recombination mechanisms in mammalian cells.

References Abbott, D. W., Thompson, M. E., Robinson-Benion, C., Tomlinson, G., Jensen, R. A., and Holt, J. T. (1999). BRCA1 expression restores radiation resistance in BRCA1-defective cancer cells through enhancement of transcription-coupled DNA repair. J. Biol. Chem. 274, 18808–18812. Al-Khodairy, F., and Carr, A. M. (1992). DNA repair mutants defining G2 checkpoint pathways in Schizosaccharomyces pombe. EMBO J. 11, 1343–1350. Anderson, L. K., Offenbeg, H. H., Verkuijlen, W. M. H. C., and Heyting, C. (1997). RecA proteins are components of early meiotic nodules in lily. Proc. Natl. Acad. Sci. USA 94, 6868–6873. Anderson, L. K., Reeves, A., Webb, L. M., and Ashley, T. (1999). Distribution of crossing over on mouse synaptonemal complexes using immunofluorescent localization of MLH1 protein. Genetics 151, 1569–1579.

128

Madalena Tarsounas and Peter B. Moens

Baker, S. M., Bronner, C. E., Zhang, L., Plug, A. W., Robatzek, M., Warren, G., Elliott, E. A., Yu, J., Ashley, T., and Arnheim, N. (1995). Male mice defective in the DNA mismatch repair gene PMS2 exhibit abnormal chromosome synapsis in meiosis. Cell 82, 309–319. Baker, S. M., Plug, A. W., Prolla, T. A., Bronner, C. E., Harris, A. C., Yao, X., Christie, D. M., Monell, C., Arnheim, N., Bradley, A., Ashley, T., and Liskay, R. M. (1996). Involvement of mouse Mlh1 in DNA mismatch repair and meiotic crossing over. Nature Genet. 13, 336–342. Banin, S., Moyal, L., Shieh, S., Taya, Y., Anderson, C. W., Chessa, L., Smorodinsky, N. I., Prives, C., Reiss, Y., Shiloh, Y., and Ziv, Y. (1998). Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science 281, 1674–1677. Barlow, A. L., and Hulten, M. A. (1998). Crossing over analysis at pachytene in man. Eur. J. Hum. Genet. 6, 350–358. Barlow, C., Hirotsune, S., Paylor, R., Liyanage, M., Eckhaus, M., Collins, F., Shiloh, Y., Crawley, J. N., Ried, T., Tagle, D., and Wynshaw-Boris, A. (1996). Atm-deficient mice: a paradigm of ataxia telangiectasia. Cell 86, 159–171. Barlow, A. L., Benson, F. E., West, S. C., and Hulten, M. A. (1997a). Distribution of the Rad51 recombinase in human and mouse spermatocytes. EMBO J. 16, 5207–5215. Barlow, C., Liyanage, M., Moens, P. B., Deng, C. X., Ried, T., and Wynshaw-Boris, A. (1997b). Partial rescue of the prophase I defects of Atm-deficient mice by p53 and p21 null alleles. Nature Genet. 17, 462–466. Barlow, C., Liyanage, M., Moens, P. B., Tarsounas, M., Nagashima, K., Brown, K., Rottinghaus, S., Jackson, S. P., Tagle, D., Ried, T., and Wynshaw-Boris, A. (1998). Atm deficiency results in severe meiotic disruption as early as leptonema of prophase I. Development 125, 4007–4017. Baumann, P., and West, S. C. (1997). The human RAD51 protein: polarity of strand transfer and stimulation by hRP-A. EMBO J. 16, 5198–5206. Baumann, P., and West, S. C. (1998). Role of the human RAD51 protein in homologous recombination and double-stranded-break repair. Trends Biochem. Sci. 23, 247–251. Baumann, P., Benson, F. E., and West, S. C. (1996). Human Rad51 protein promotes ATP-dependent homologous pairing and strand transfer reactions in vitro. Cell 87, 757–766. Bergerat, A., de Massy, B., Gadelle, D., Varoutas, P.-C., Nicolas, A., and Forterre, P. (1997). An atypical topoisomerase II from archea with implications for meiotic recombination. Nature 386, 414– 417. Bickel, S. E., and Orr-Weaver, T. L. (1996). Holding chromatids together to ensure they go their separate ways. Bioessays 18, 293–300. Bishop, D. K. (1994). RecA homologs Dmc1 and Rad51 interact to form multiple nuclear complexes prior to meiotic chromosome synapsis. Cell 79, 1081–1092. Canman, C. E., Lim, D. S., Cimprich, K. A., Taya, Y., Tamai, K., Sakaguchi, K., Appella, E., Kastan, M. B., Siliciano, J. D. (1998). Activation of the ATM kinase by ionizing radiation and Phosphorylation of p53. Science 281, 1677–1679. Cao, L., Alani, E., and Kleckner, N. (1990). A pathway for generation and processing of double stranded breaks during meiotic recombination in S. cerevisiae. Cell 61, 1089–1101. Carney, J. P., Maser, R. S., Olivares, H., Davis, E. M., Le Beau, M., Yates, J. R. 3rd, Hays, L., Morgan, W. F., and Petrini, J. H. (1998). The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response. Cell 93, 477–486. Cha, R. S., Weiner, B. M., Keeney, S., Dekker, J., and Kleckner, N. (2000). Progression of meiotic DNA replication is modulated by interchromosomal proteins, negatively by Spo11p and positively by Rec8p. Genes Dev. 14, 493–503. Chen, J., Silver, D. P., Walpita, D., Cantor, S. B., Gazdar, A. F., Tomlinson G, Couch FJ, Weber BL, Ashley T, Livingston, D. M., and Scully R. (1998). Stable interaction between the products of the BRCA1 and BRCA2 tumor suppressor genes in mitotic and meiotic cells. Mol. Cell 2, 317–328. Chen, G., Yuan, S. S., Liu, W., Xu, Y., Trujillo, K., Song, B., Cong, F., Goff, S. P., Wu, Y., Arlinghaus, R.,

4. DNA Repair and Checkpoint Proteins at Meiosis

129

Baltimore, D., Gasser, P. J., Park, M. S., Sung, P., and Lee, E. Y. (1999). Radiation-induced assembly of Rad51 and Rad52 recombination complex requires ATM and c-Abl. J. Biol. Chem. 274, 12748– 12752. Chua, P. R., and Roeder, G. S. (1998). Zip2, a meiosis-specific protein required for the initiation of chromosome synapsis. Cell 93, 349–359. Cliby, W. A., Roberts, C. J., Cimprich, K. A., Stringer, C. M., Lamb, J. R., Schreiber, S. L., and Friend, S. H. (1998). Overexpression of a kinase-inactive ATR protein causes sensitivity to DNAdamaging agents and defects in cell cycle checkpoints. EMBO J. 17, 159–169. Connor, F., Bertwistle, D., Mee, P. J., Ross, G. M., Swift, S., Grigorieva, E., Tybulewicz, V. L., and Ashworth, A. (1997). Tumorigenesis and a DNA repair defect in mice with a truncating Brca2 mutation. Nature Genet. 17, 423–430. Critchlow, S. E., and Jackson, S. P. (1998). DNA end-joining: from yeast to man. Trends Biochem. Sci. 23, 394–398. Dasika, G. K., Lin, S. C., Zhao, S., Sung, P., Tomkinson, A., and Lee, E. Y. (1999). DNA damage– induced cell cycle checkpoints and DNA strand break repair in development and tumorigenesis. Oncogene 18, 7883–7899. Dernburg, A. F., McDonald, K., Moulder, G., Barstead, R., Dresser, M., and Villeneuve, A. M. (1998). Meiotic recombination in C. elegans initiates by a conserved mechanism and is dispensable for homologous chromosome synapsis. Cell 94, 387–398. Dobson, M. J., Pearlman, R. E., Karaiskakis, A., Spyropoulos, B., and Moens, P. B. (1994). Synaptonemal complex proteins: occurrence, epitope mapping and chromosome disjunction. J. Cell Sci. 107, 2749–2760. Dresser, M. E., D. J. Ewing, M. N. Conrad, A. M. Dominguez, R. Barstead, H. Jiang, and T. Kodadek. (1997). DMC1 functions in a Saccharomyces cerevisiae meiotic pathway that is largely independent of the RAD51 pathway. Genetics 147, 533–544. Edelmann, W., Cohen, P. E., Kane, M., Lau, K., Morrow, B., Bennett, S., Umar, A., Kunkel, T., Cattoretti, G., Chaganti, R., Pollard, J.W., Kolodner, R. D., and Kucherlapati, R. (1996). Meiotic pachytene arrest in MLH1-deficient mice. Cell 85, 1125–1134. Edelmann, W., Cohen, P. E., Kneitz, B., Winand, N., Lia, M., Heyer, J., Kolodner, R., Pollard, J. W., and Kucherlapati, R. (1999). Mammalian MutS homologue 5 is required for chromosome pairing in meiosis. Nature Genet. 21, 123–127. Eijpe, M., Heyting, C., Gross, B., and Jessberger, R. (2000). Association of mammalian SMC1 and SMC3 proteins with meiotic chromosomes and synaptonemal complexes. J. Cell Sci. 113, 673–682. Freire, R., Murguia, J. R., Tarsounas, M., Lowndes, N. F., Moens, P. B. and Jackson, S. P. (1998). Human and mouse homologs of Schizosaccharomyces pombe rad1(+) and Saccharomyces cerevisiae RAD17: linkage to checkpoint control and mammalian meiosis. Genes Dev. 12, 2560–2573. Friedman, D. B., Hollingsworth, N. M., and Byers, B. (1994). Insertional mutations in the yeast HOP1 gene: evidence for multimeric assembly in meiosis. Genetics 136, 449–464. German, J. (1993). Bloom syndrome: a Mendelian prototype of somatic mutational disease. Medicine (Baltimore) 72, 393–406. Goedecke, W., Eijpe, M., Offenberg, H. H., van Aalderen, M., and Heyting, C. (1999). Mre11 and Ku70 interact in somatic cells, but are differentially expressed in early meiosis. Nature Genet. 23, 194–198. Goldstein, P. (1987). Multiple synaptonemal complexes (polycomplexes): origin, structure and function. Cell. Biol. Int. Rep. 11, 759–796. Golub, E. I., Gupta, R. C., Haaf, T., Wold, M. S., and Radding, C. M. (1998). Interaction of human rad51 recombination protein with single-stranded DNA binding protein, RP-A. Nucleic Acids Res. 26, 5388–5393. Gowen, L. C., Avrutskaya, A. V., Latour, A. M., Koller, B. H., and Leadon, S. A. (1998). BRCA1 required for transcription-coupled repair of oxidative DNA damage. Science 281, 1009–1012. Haber, J. E. (1998). The many interfaces of Mre11. Cell 95, 583–586.

130

Madalena Tarsounas and Peter B. Moens

Hassold, T. J. (1996). Mismatch repair goes meiotic. Nature Genet. 13, 261–262. Heng, H. H., Tsui, L. C., and Moens, P. B. (1994). Organization of heterologous DNA inserts on the mouse meiotic chromosome core. Chromosoma 103, 401–407. Heng, H. H., Chamberlain, J. W., Shi, X. M., Spyropoulos, B., Tsui, L. C., and Moens, P. B. (1996). Regulation of meiotic chromatin loop size by chromosomal position. Proc. Natl. Acad. Sci. USA 93, 2795–2800. Heng, H. H., Spyropoulos, B., and Moens, P. B. (1997). FISH technology in chromosome and genome research. Bioessays 19, 75–84. Heyting, C., Dietrich, A. J., Redeke,r E. J., and Vink, A. C. (1985). Structure and composition of synaptonemal complexes, isolated from rat spermatocytes. Eur. J. Cell Biol. 36, 307–314. Heyting, C., Dettmers, R. J., Dietrich, A. J., Redeker, E. J., and Vink, A. C. (1988). Two major components of synaptonemal complexes are specific for meiotic prophase nuclei. Chromosoma 96, 325–332. Hirano, T. (1999). SMC-mediated chromosome mechanics: a conserved scheme from bacteria to vertebrates? Genes Dev. 13, 11–19. Hoekstra, M. F. (1997). Responses to DNA damage and regulation of cell cycle checkpoints by the ATM protein kinase family. Curr. Opin. Genet. Dev. 7, 170–175. Hollingsworth, N. M., and Byers, B. (1989). HOP1: a yeast meiotic pairing gene. Genetics 121, 445– 462. Hollingsworth, N. M., and Ponte, L. (1997). Genetic interactions between HOP1, RED1 and MEK1 suggest that MEK1 regulates assembly of axial element components during meiosis in the yeast Saccharomyces cerevisiae. Genetics 147, 33–42. Keegan, K. S., Holtzman, D. A., Plug, A. W., Christenson, E. R., Brainerd, E. E., Flaggs, G., Bentley, N. J., Taylor, E. M., Meyn, M. S., Moss, S. B., Carr, A. M., Ashley, T., and Hoekstra, M. F. (1996). The ATR and ATM protein kinases associate with different sites along meiotically pairing chromosomes. Genes Dev. 10, 2423–2437. Keeney, S., Giroux, C. N., and Kleckner, N. (1997). Meiosis-specific DNA double-strand breaks are catalyzed by Spo11, a member of a widely conserved protein family. Cell 88, 375–384. Keeney, S., Baudat, F., Angeles, M., Zhou, Z. H., Copeland, N. G., Jenkins, N. A., Manova, K., Jasin, M. (1999). A mouse homolog of the Saccharomyces cerevisiae meiotic recombination DNA transesterase Spo11p. Genomics 61, 170–182. Kerrebrock, A. W., Moore, D. P., Wu, J. S., and Orr-Weaver, T. L. (1995). Mei-S332, a Drosophila protein required for sister-chromatid cohesion, can localize to meiotic centromere regions. Cell 83, 247–256. Kironmai, K. M., Muniyappa, K., Friedman, D. B., Hollingsworth, N. M., and Byers, B. (1998). DNA-binding activities of Hop1 protein, a synaptonemal complex component from Saccharomyces cerevisiae. Mol. Cell Biol. 18, 1424–1435. Kleckner, N. (1996). Meiosis: how could it work? Proc. Natl. Acad. Sci. USA 93, 8167–8174. Kohli, J., and Bahler, J. (1994). Homologous recombination in fission yeast: absence of crossover interference and synaptonemal complex. Experientia 50, 295–306. Lakin, N. D., and Jackson, S. P. (1999). Regulation of p53 in response to DNA damage. Oncogene 18, 7644–7655. Lammers, J. H. M., Offenberg, H. H., van Aalderen, M., Vink, A. C. G., Dietrich, A. J. J., and Heyting, C. (1994). The gene encoding a major component of the lateral elements of synaptonemal complexes of the rat is related to X-linked lymphocyte-regulated genes. Mol. Cell Biol. 14, 1137–1146. Lammers, J. H. M., van Aalderen, M., Peters, A. H. F. M., van Pelt, A. A. M., Gaemers, I. C., de Rooij, D. G., de Boer, P., Offenberg, H. H., Dietrich, A. J. J., and Heyting, C. (1995). A change in the phosphorylation pattern of the 30,000–33,000 Mr synaptonemal complex proteins of the rat between early and mid-pachytene. Chromosoma 104, 154–163. Leu, J. Y., Chua, P. R., and Roeder, G. S. (1998). The meiosis-specific Hop2 protein of S. cerevisiae ensures synapsis between homologous chromosomes. Cell 94, 375–386.

4. DNA Repair and Checkpoint Proteins at Meiosis

131

Levine, A. J. (1997). p53, the cellular gatekeeper for growth and division. Cell 88, 323–331. Li, Z., Golub, E. I., Gupta, R., and Radding, C. M. (1997). Recombination activities of HsDmc1 protein, the meiotic human homolog of RecA protein. Proc. Natl. Acad. Sci. USA 94, 11221–11226. Lim, D. S., and Hasty, P. (1996). A mutation in mouse RAD51 results in an early embryonic lethal that is suppressed by a mutation in p53. Mol. Cell. Biol. 16, 7133–7143. Liu, J. G., Yuan, L., Brundell, E., Bjorkroth, B., Daneholt, B., and Hoog, C. (1996). Localization of the N-terminus of SCP1 to the central element of the synaptonemal complex and evidence for direct interactions between the N-termini of SCP1 molecules organized head-to-head. Exp. Cell Res. 226, 11–19. Lydall, D., and Weinert, T. (1995). Yeast checkpoint genes in DNA damage processing: implications for repair and arrest. Science 270, 1488–1491. Mao-Draayer, Y., Galbraith, A. M., Pittman, D. L., Cool, M., and Malone, R. E. (1996). Analysis of meiotic recombination pathways in the yeast Saccharomyces cerevisiae. Genetics 144, 71–86. Marciniak, R. A., Lombard, D. B., Johnson, F. B., and Guarente, L. (1998). Nucleolar localization of the Werner syndrome protein in human cells. Proc. Natl. Acad. Sci. USA 95, 6887–6892. Marcus, J. N., Watson, P., Page, D. L., Narod, S. A., Lenoir, G. M., Tonin, P., Linder-Stephenson, L., Salerno, G., Conway, T. A., and Lynch, H. T. (1996). Hereditary breast cancer: pathobiology, prognosis, and BRCA1 and BRCA2 gene linkage. Cancer 77, 697–709. Maser, R. S., Monsen, K. J., Nelms, B. E., Petrini, J. H. (1997). hMre11 and hRad50 nuclear foci are induced during the normal cellular response to DNA double-strand breaks. Mol. Cell. Biol. 17, 6087–6096. Masson, J. Y., Davies, A. A., Hajibagheri, N., Van Dyck E., Benson, F. E., Stasiak, A. Z., Stasiak, A., and West, S. C. (1999). The meiosis-specific recombinase hDmc1 forms ring structures and interacts with hRad51. EMBO J. 18, 6552–6560. Meuwissen, R. L. J., Offenberg, H. H., Dietrich, A. J. J., Riesewijk, A. van Iersel, M., and Heyting, C. (1992). A coiled-coil related protein specific for synapsed regions of meiotic prophase chromosomes. EMBO J. 11, 5091–5100. Modrich, P. (1991). Mechanisms and biological effects of mismatch repair. Annu. Rev. Genet. 25, 229–253. Moens, P. B., and Earnshaw, W. C. (1989). Anti-topoisomerase II recognizes meiotic chromosome cores. Chromosoma 98, 317–322. Moens, P. B., and Spyropoulos, B. (1995). Immunocytology of chiasmata and chromosomal disjunction at mouse meiosis. Chromosoma 104, 175–182. Moens, P. B., Heyting, C., Dietrich, A. J., van Raamsdonk, W., and Chen, Q. (1987). Synaptonemal complex antigen location and conservation. J. Cell Biol. 105, 93–103. Moens, P. B., Spyropoulos, B., Dobson, M., Karaiskakis, A., and Pearlman, R. E. (1992). Searching for synaptonemal complex proteins and their genes. Dev. Genet. 13, 435–439. Moens, P. B., Chen, D. J., Shen, Z., Kolas, N., Tarsounas, M., Heng, H., and Spyropoulos, B. (1997). RAD51 immunocytology in rat and mouse spermatocytes and oocytes. Chromosoma 106, 207– 215. Moens, P. B., Pearlman, R. E., Heng, H. H., and Traut, W. (1998). Chromosome cores and chromatin at meiotic prophase. Curr. Top. Dev. Biol. 37, 241–262. Moens, P. B., Tarsounas, M., Morita, T., Rottinghaus, S., Jackson, S., Barlow, C., and Wynshaw-Boris, A. (1999). The association of the ATR protein with meiotic chromosome cores. Chromosoma 108, 95–102. Moens, P. B., Freire, R., Tarsounas, M., Spyropoulos, B., and Jackson, S. P. (2000). Expression and nuclear localization of BLM, a chromosome stability protein mutated in Bloom’s syndrome, suggest a role in recombination during meiotic prophase. J. Cell Sci. 113, 663–672. Moore, D. P., Page, A. W., Tang, T. T., Kerrebrock, A. W., and Orr-Weaver, T. L. (1998). The cohesion protein MEI-S332 localizes to condensed meiotic and mitotic centromeres until sister chromatids separate. J. Cell Biol. 140, 1003–1012.

132

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Morrison, C., Sonoda, E., Takao, N., Shinohara, A., Yamamoto, K., and Takeda, S. (2000). The controlling role of ATM in homologous recombinational repair of DNA damage. EMBO J. 19, 463–471. Moynahan, M. E., Chiu, J. W., Koller, B. H., and Jasin, M. (1999). Brca1 controls homology-directed DNA repair. Mol. Cell 4, 511–518. Nairz, K., and Klein, F. (1997). mre11S—a yeast mutation that blocks double-strand-break processing and permits nonhomologous synapsis in meiosis. Genes Dev. 11, 2272–2290. Offenberg, H. H., Schalk, J. A., Meuwissen, R. L., van Aalderen, M., Kester, H. A., Dietrich, A. J., and Heyting, C. (1998). SCP2: a major protein component of the axial elements of synaptonemal complexes of the rat. Nucleic Acids Res. 26, 2572–2579. Orr-Weaver, T. L. (1999). The ties that bind: localization of the sister-chromatid cohesin complex on yeast chromosomes. Cell 99, 1–4. Padmore, R., Cao, L., and Kleckner, N. (1991). Temporal comparison of recombination and synaptonemal complex formation during meiosis in S. cerevisiae. Cell 66, 1239–1256. Patel, K. J., Vu, V. P., Lee, H., Corcoran, A., Thistlethwaite, F. C., Evans, M. J., Colledge, W. H., Friedman, L. S., Ponder, B. A., and Venkitaraman, A. R. (1998). Involvement of Brca2 in DNA repair. Mol. Cell 1, 347–357. Paull, T. T., and Gellert, M. (1998). The 3′ to 5′ exonuclease activity of Mre 11 facilitates repair of DNA double-strand breaks. Mol. Cell 1, 969–979. Pearlman, R. E., Tsao, N., and Moens, P. B. (1992). Synaptonemal complexes from DNase-treated rat pachytene chromosomes contain (GT)n and LINE/SINE sequences. Genetics 130, 865–872. Pittman, D. L., Cobb, J., Schimenti, K. L., Wilson, L. A., Cooper, D. M., Brignull, E., Handel, M. A., and Schimenti, J. C. (1998). Meiotic prophase arrest with failure of chromosome synapsis in mice deficient for Dmc1, a germline-specific RecA homolog. Mol. Cell 1, 697–705. Plug, A. W., Peters, A. H., Keegan, K. S., Hoekstra, M. F., de Boer, P., and Ashley, T. (1998). Changes in protein composition of meiotic nodules during mammalian meiosis. J. Cell Sci. 111, 413–423. Rajan, J. V., Marquis, S. T., Gardner, H. P., and Chodosh, L. A. (1997). Developmental expression of Brca2 co-localizes with Brca1 and is associated with proliferation and differentiation in multiple tissues. Dev. Biol. 184, 385–401. Rockmill, B., and Roeder, G. S. (1990). Meiosis in asynaptic yeast. Genetics 126, 563–574. Rockmill, B., Sym, M., Scherthan, H., and Roeder, G. S. (1995). Roles for two RecA homologs in promoting meiotic chromosome synapsis. Genes Dev. 9, 2684–2695. Roeder, G. S. (1997). Meiotic chromosomes: it takes two to tango. Genes Dev. 11, 2600–2621. Romanienko, P. J., and Camerini-Otero, R. D. (1999). Cloning, characterization, and localization of mouse and human SPO11. Genomics 61,156–169. Schalk, J. A. C., Dietrich, A. J. J., Vink, A. C. G., Offenberg, H. H., van Aalderen, M., and Heyting, C. (1998). Localization of SCP2 and SCP3 protein molecules within synaptonemal complexes of the rat. Chromosoma 107, 540–548. Scherthan, H., Bahler, J., and Kohli, J. (1994). Dynamics of chromosome organization and pairing during meiotic prophase in fission yeast. J. Cell Biol. 127, 273–285. Schmekel, K., Meuwissen, R. L. J., Dietrich, A. J. J., Vink, A. C. G., van Marle, J., van Veen, H., and Heyting, C. (1996). Organization of SCP1 protein molecules within synaptonemal complexes of the rat. Exp. Cell Res. 226, 20–30. Schwacha, A., and Kleckner, N. (1997). Interhomolog bias during meiotic recombination: meiotic functions promote a highly differentiated interhomolog-only pathway. Cell 90, 1123–1135. Scully, R., Chen, J., Plug, A., Xiao, Y., Weaver, D., Feunteun, J., Ashley, T., and Livingston, D. M. (1997). Association of BRCA1 with Rad51 in mitotic and meiotic cells. Cell 88, 265–275. Shannon, M., Richardson, L., Christian, A., Handel, M. A., and Thelen, M. P. (1999). Differential gene expression of mammalian SPO11/TOP6A homologs during meiosis. FEBS Lett. 462, 329–334. Sharan, S. K., Morimatsu, M., Albrecht, U., Lim, D. S., Regel, E., Dinh, C., Sands, A., Eichele, G., Hasty, P., and Bradley, A. (1997). Embryonic lethality and radiation hypersensitivity mediated by Rad51 in mice lacking Brca2. Nature 386, 804–810.

4. DNA Repair and Checkpoint Proteins at Meiosis

133

Shen, S. X., Weaver, Z., Xu, X., Li, C., Weinstein, M., Chen, L., Guan, X. Y., Ried, T., and Deng, C. X. A. (1998). Targeted disruption of the murine Brca1 gene causes gamma-irradiation hypersensitivity and genetic instability. Oncogene 17, 3115–3124. Smith, K. N., and Nicolas, A. (1998). Recombination at work for meiosis. Curr. Opin. Genet. Dev. 8, 200–211. Smith, A. V., and Roeder, G. S. (1997). The yeast Red1 protein localizes to the cores of meiotic chromosomes. J. Cell Biol. 136, 957–967. Stahl, F. (1996). Meiotic recombination in yeast: coronation of the double-strand-break repair model. Cell 87, 965–968. Strunnikov, A. V. (1998). SMC proteins and chromosome structure. Trends Cell Biol. 8, 454–459. Suzuki, K., Kodama, S., and Watanabe, M. (1999). Recruitment of ATM protein to double strand DNA irradiated with ionizing radiation. J. Biol. Chem. 274, 25571–25575. Sym, M., and Roeder, G. S. (1995). Zip1-induced changes in synaptonemal complex structure and polycomplex assembly. J. Cell Biol. 128, 455–466. Sym, M., Engebrecht, J. A., and Roeder, G. S. (1993). Zip1 is a synaptonemal complex protein required for meiotic chromosome synapsis. Cell 72, 365–378. Szostak, J. W., Orr-Weaver, T. L., Rothstein, R. J., and Stahl, F. W. (1983). The double-strand-break repair model for recombination. Cell 33, 25–35. Tanaka, T., Cosma, M. P., Wirth, K., and Nasmyth, K. (1999). Identification of cohesin association sites at centromeres and along chromosome arms. Cell 98, 847–858. Tarsounas, M., Pearlman, R. E., Gasser, P. J., Park, M. S., and Moens, P. B. (1997). Protein–protein interactions in the synaptonemal complex. Mol. Biol. Cell 8, 1405–1414. Tarsounas, M., Pearlman, R. E., and Moens, P. B. (1999a). Meiotic activation of rat pachytene spermatocytes with okadaic acid: the behaviour of synaptonemal complex components SYN1/SCP1 and COR1/SCP3. J. Cell Sci. 112, 423–434. Tarsounas, M., Morita, T., Pearlman, R. E., and Moens, P. B. (1999b). RAD51 and DMC1 form mixed complexes associated with mouse meiotic chromosome cores and synaptonemal complexes. J. Cell Biol. 147, 207–220. Terasawa, M., Shinohara, A., Hotta, Y., Ogawa, H., and Ogawa, T. (1995). Localization of RecA-like recombination proteins on chromosomes of the lily at various meiotic stages. Genes Dev. 9, 925–934. Tibbetts, R. S., Brumbaugh, K. M., Williams, J. M., Sarkaria, J. N., Cliby, W. A., Shieh, S. Y., Taya, Y., Prives, C., and Abraham, R. T. (1999). A role for ATR in the DNA damage-induced phosphorylation of p53. Genes Dev. 13, 152–157. Tsuzuki, T., Fujii, Y., Sakumi, K., Tominaga, Y., Nakao, K., Sekiguchi, M., Matsushiro, A., Yoshimura, Y., and Morita, T. (1996). Targeted disruption of the Rad51 gene leads to lethality in embryonic mice. Proc. Natl. Acad. Sci. USA 93, 6236–6240. Tung, K. S., and Roeder, G. S. (1998). Meiotic chromosome morphology and behaviour in zip1 mutants of Saccharomyces cerevisiae. Genetics 149, 817–832. Usui, T., Ohta, T., Oshiumi, H., Tomizawa, J., Ogawa, H., and Ogawa, T. (1998). Complex formation and functional versatility of Mre11 of budding yeast in recombination. Cell 95, 705–716. Varon, R., Vissinga, C., Platzer, M., Cerosaletti, K. M., Chrzanowska, K. H., Saar, K., Beckmann, G., Seemanova, E., Cooper, P. R., Nowak, N. J., Stumm, M., Weemaes, C. M., Gatti, R. A., Wilson, R. K., Digweed, M., Rosenthal, A., Sperling, K., Concannon, P., and Reis, A. (1998). Nibrin, a novel DNA double-strand break repair protein, is mutated in Nijmegen breakage syndrome. Cell 93, 467–476. Walpita, D., Plug, A. W., Neff, N. F., German, J., and Ashley, T. (1999). Bloom’s syndrome protein, BLM, colocalizes with replication protein A in meiotic prophase nuclei of mammalian spermatocytes. Proc. Natl. Acad. Sci. USA 96, 5622–6527. Watt, P. M., Hickson, I. D., Borts, R. H., and Louis, E. J. (1996). SGS1, a homologue of the Bloom’s and Werner’s syndrome genes, is required for maintenance of genome stability in Saccharomyces cerevisiae. Genetics 144, 935–945.

134

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Weiner, B. M., and Kleckner, N. (1994). Chromosome pairing via multiple interstitial interactions before and during meiosis in yeast. Cell 77, 977–991. Welcsh, P. L., Owens, K. N., and King, M. C. (2000). Insights into the functions of BRCA1 and BRCA2. Trends Genet. 16, 69–74. Wold, M. S. (1997). Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu. Rev. Biochem. 66, 61–92. Wong, A. K. C., Pero, R., Ormonde, P. A., Tavtigian, S. V., and Bartel, P. L. (1997). RAD51 interacts with the evolutionarily conserved BRC motifs in the human breast cancer susceptibility gene Brca2. J. Biol. Chem. 272, 31941–31944. Yamagata, K., Kato, J., Shimamoto, A., Goto, M., Furuichi, Y., and Ikeda, H. (1998). Bloom’s and Werner’s syndrome genes suppress hyperrecombination in yeast sgs1 mutant: implication for genomic instability in human diseases. Proc. Natl. Acad. Sci. USA 95, 8733–8738. Yamaguchi-Iwai, Y., Sonoda, E., Sasaki, M. S., Morrison, C., Haraguchi, T., Hiraoka, Y., Yamashita, Y. M., Yagi, T., Takata, M., Price, C., Kakazu, N., and Takeda, S. (1999). Mre11 is essential for the maintenance of chromosomal DNA in vertebrate cells. EMBO J. 18, 6619–6629. Yoshida, K., Kondoh, G., Matsuda, Y., Habu, T., Nishimune, Y., and Morita, T. (1998). The mouse RecA-like gene Dmc1 is required for homologous chromosome synapsis during meiosis. Mol. Cell 1, 707–718. Yu, C. E., Oshima, J., Fu, Y. H., Wijsman, E. M., Hisama, F., Alisch, R., Matthews, S., Nakura, J., Miki, T., Ouais, S., Martin, G. M., Mulligan, J., and Schellenberg, G. D. (1996). Positional cloning of the Werner’s syndrome gene. Science 272, 258–262. Yuan, L., Pelttari, J., Brundell, E., Bjorkroth, B., Zhao, J., Liu, J. G., Brismar, H., Daneholt, B., and Hoog, C. (1998). The synaptonemal complex protein SCP3 can form multistranded, cross striated fibers in vivo. J. Cell Biol. 142, 331–339. Yuan, L., Liu, J. G., Zhao, J., Brundell, E., Daneholt, B., and Hoog, C. (2000). The murine Scp3 gene is required for synaptonemal complex assembly, chromosome synapsis, and male fertility. Mol. Cell 5, 73–83. Xu, Y., Ashley, T., Brainerd, E. E., Bronson, R. T., Meyn, M. S., and Baltimore, D. (1996). Targeted disruption of Atm leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma. Genes Dev. 10, 2411–2422. Zabludoff, S. D., Wright, W. W., Harshman, K., and Wold, B. J. (1996). BRCA1 mRNA is expressed highly during meiosis and spermiogenesis but not during mitosis of male germ cells. Oncogene 13, 649–653. Zickler, D., and Kleckner, N. (1998). The leptotene-zygotene transition of meiosis. Annu. Rev. Genet. 32, 619–697.

5

Cytoskeletal and Ca2+ Regulation of Hyphal Tip Growth and Initiation

Sara Torralba and I. Brent Heath Biology Department York University Toronto, Ontario, M3J 1P3 Canada

I. Introduction: Characteristics of Fungal Growth II. The Cytoskeleton and Apical Growth in Fungi A. Introduction B. Actin Distribution in Hyphae C. Actin-Binding Proteins D. Role of Actin in Tip Growth III. Calcium and Apical Growth in Fungi A. Internal Ca2+ Gradients in Hyphae: Methods for Their Detection B. Cytosolic Ca2+ Homeostasis in Fungi: Roles of Plasma Membrane Transport and Intracellular Sequestration of Ca2+ C. Roles of Ca2+ Gradients in Tip Growth IV. Conclusions References

Hyphal tip growth is a complex process involving finely regulated interactions between the synthesis and expansion of cell wall and plasma membrane, diverse intracellular movements, and turgor regulation. F-actin is a major regulator and integrator of these processes. It directly contributes to (a) tip morphogenesis, most likely by participation in an apical membrane skeleton that reinforces the apical plasma membrane, (b) the transport and exocytosis of vesicles that contribute plasma membrane and cell wall material to the hyphal tips, (c) the localization of plasma membrane proteins in the tips, and (d) cytoplasmic and organelle migration and positioning. The pattern of reorganization of F-actin prior to formation of new tips during branch initiation also indicates a critical role in early stages of assembly of the tip apparatus. One of the universal characterisitics of all critically examined tip-growing cells, including fungal hyphae, is the obligatory presence of a tip-high gradient of cytoplasmic Ca2+ that probably regulates both actin and nonactin components of the apparatus, and the formation of which may also initiate new tips. This review discusses the diversity of evidence behind these concepts.  2001 Academic Press. C

Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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I. Introduction: Characteristics of Fungal Growth Hyphal cells of filamentous fungi exhibit a polarized mode of growth that occurs through tip expansion. This method of cell growth gives rise to tube-shaped cells. Budding or fission yeasts show determinate tip growth and have been used as model systems for genetic and molecular analysis of the process (reviewed by Drubin and Nelson, 1996; Mata and Nurse, 1997). Tip growth is also fundamental to plant reproduction and nutrition and alga development, being the basis of growth of pollen tubes, root hairs, and alga rhizoids (Steer and Steer, 1989; Kropf, 1994; Kropf et al., 1998). Despite their taxonomic diversity, tip-growing cells share an important characteristic: They are not motile, suggesting that tip growth is a consistent solution for exploring the environment through directed growth. In fungi, the regulation of apical growth provides mechanisms for colonization of substrates, secretion of hydrolytic enzymes, absorption of nutrients, regulation of morphogenesis, and recognition of environmental signals (Gooday, 1983; Wessels, 1986; Heath, 1990). Reinhardt (1892) introduced the idea that fungi are tube-dwelling amoebae, a concept that is still viable and has recently been reviewed (Heath and Steinberg, 1999). The essential feature of the idea is that the cytoplasm migrates through the tubular hyphae as an animal cell migrates over a solid substrate. During apical growth the hyphal tip is protruded out into the environment from the subapical tube. The protrusive process enables the hypha to explore and penetrate its environment. Tip growth in filamentous fungi has been studied for over a century because they have a strong impact on human life, both prejudicial and otherwise. These organisms can cause diseases or decay. The mode of nutrient acquisition requires that filamentous fungi secrete a large amount of enzymes into their environment in order to break down extracellular macromolecules into units suitable for uptake into the cell. Some of the substances secreted by fungi are the foundation of major industries, whereas others are essential for nutrient recycling. Protein secretion is related to fungal growth. Saccharomyces mutant strains specifically blocked in secretion are also defective in cell-surface expansion (Novick and Sheckman, 1983). In filamentous fungi, secretion is a polarized process mainly restricted to the growing hyphal tips where there are large pores in the young wall through which the enzyme may readily diffuse (Chang and Trevithick, 1974) or move together with nascent wall polymers (Wessels, 1990). Antibodies against pectinesterase, glucoamylase, and lignin peroxidase have been used to demonstrate that secretion of these degradative enzymes occurs at the growing hyphal apices of Phytophthora, Aspergillus, and Phanerochaete, respectively (F¨orster and Mendgen, 1987; W¨osten et al., 1991; Moukha et al., 1993). In filamentous fungi, the extreme polarization of hyphae applies not only to their growth pattern but also to their organelle positioning and cytoskeletal and ion distributions. A tip-focused gradient of Ca2+ is a general feature of growing hyphae, which is thought to play a role in apical morphology, morphogenesis, and growth. To date, all microscopic studies of filamentous fungi describe a polarized

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organization of the hyphal cytoplasm. The hyphal apex is a highly specialized region of the fungal cell where accumulation of numerous vesicles takes place. These wall vesicles are assumed to be involved in the incorporation of new membrane and cell wall material at the tip (Harold, 1997), and therefore they can be called wall vesicles (Heath et al., 1985). Below the apex, in the subapical region of the hyphae, a polarized organization of organelles such as mitochondria, Golgi body– like structures, endoplasmic reticulum (ER), and nuclei occurs (McClure et al., 1968; Girbardt, 1969; Grove and Bracker, 1970; Howard and Aist, 1979; Howard, 1981; Rupeˇs et al., 1995). A quantitative analysis of the organelle organization in the oomycete Saprolegnia revealed a well-defined pattern of both radial and longitudinal distribution of all organelles in the hyphae, possibly controlled by the cytoskeleton (Heath and Kaminskyj, 1989). Oomycetes are not real fungi or eufungi (Bhattacharya et al., 1992; Money, 1998), but they are included in this review because they are typically considered by mycologists and show hyphal tip growth. Apical growth is a complex process because it involves the highly localized synthesis and expansion of cell wall and plasma membrane (PM), diverse aspects of intracellular movements, turgor control, and the fine regulation of the interaction of all these processes in order to generate the characteristic tube shape of the hyphae. A Ca2+ -regulated F-actin skeleton has been postulated to contribute to a number of these tip growth processes (Heath, 1995), and evidence exists for its involvement in tip morphogenesis (Jackson and Heath, 1993; Gupta and Heath, 1997; Hyde and Heath, 1997), vesicle transport (Heath, 1990; McGoldrick et al., 1995), branching (Bachewich and Heath, 1998; Grinberg and Heath, 1997), and cytoplasmic migration (reviewed in Kaminskyj and Heath, 1996).

II. The Cytoskeleton and Apical Growth in Fungi A. Introduction The question of how fungal hyphae sustain polarized growth while maintaining a highly organized cytoplasm have been often addressed, and although many advances have been made since the early observations of Reinhardt (1892), little is understood about the mechanisms involved in the process. Published reports (Howard and Aist, 1977; Grove and Sweigard, 1980; Herr and Heath, 1982; Heath, 1987; Heath and Kaminskyj, 1989; Temperli et al., 1990; Heath, 1994; Heath and Steinberg, 1999; etc.) have implicated cytoskeletal systems in the maintenance and regulation of the cellular machinery required for hyphal elongation. 1. Cytoskeletal Components Eukaryotic cells are characterized by three major intracellular networks, the tubulin, the actin, and the intermediate filament networks, which serve structural and several other functions. In hyphae, predominant components of the cytoskeleton

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are microtubules (MTs) and actin microfilaments (Heath, 1990, 1994). Intermediate filaments have not been detected yet in filamentous fungi (May and Hyams, 1998), although an intermediate filament-like protein has been found in Saccharomyces (McConnell and Yaffe, 1992). A straightforward approach to determining the role of cytoskeletal components is to disrupt them with specific inhibitors and to examine the effects of such treatments. The fungicide Benomyl and related compounds such as methyl benzimidazole-2-yl carbamate (MBC) and nocodazole are anti-microtubule drugs (Clemons and Sisler, 1969; Davidse and Flach, 1978) that have been used to study the roles of MTs in hyphae (Howard and Aist, 1980; Howard, 1981; Herr and Heath, 1982; Caesar-Ton That et al., 1988; DeLucas et al., 1993; Harris et al., 1994; Raudaskoski et al., 1994; Pedregosa et al., 1995; Torralba et al., 1996, 1998b; Hyde et al., 1999). Cytochalasins and Latrunculins are known for their ability to inhibit polymerization of actin and offer the possibility of studying the functions of the actin cytoskeleton (Oliver, 1973; Allen et al., 1980; Grove and Sweigard, 1980; Srinivasan et al., 1996; Gupta and Heath, 1997; Torralba et al., 1998a; Bachewich and Heath, 1999). The organization and functions of the cytoskeletal polymers are determined by MT-associated proteins (Olmsted, 1986, 1991) and actin-binding proteins (ABPs) (St¨ossel, 1993; Hartwig and Kwiatkowski, 1991), respectively. The ABPs and MTassociated proteins are essentially of two types, those that form static connections between the polymers and therefore determine the pattern of the polymers (e.g., form bundles or meshworks) and those that are mechanochemical force generators (e.g., myosin sliding along actin filaments in muscle cells). Motor proteins are able to migrate along filamentous tracks, and if the motor protein is coupled to a cargo, such as a vesicle or a nucleus, the organelles are transported correspondingly. They are very important in filamentous fungi, since these organisms show prominent intracellular traffic along F-actin and/or MTs over long distances (reviewed in Heath, 1995; Gow, 1995). Three major classes of motor proteins have been described: kinesins and dyneins, both of which are MT-dependent motors, and myosins, which are actin-dependent motors. Fungal representatives of cytoplasmic dynein in Neurospora (Plamann et al., 1994) and Aspergillus (Xiang et al., 1994), of the myosin I family in Aspergillus (McGoldrick et al., 1995), and of conventional kinesin from Neurospora (Steinberg and Schliwa, 1995), Ustilago (Lehmler et al., 1997), and Nectria (Wu et al., 1998) have been described (overview in Yamashita and May, 1998). Our knowledge about motor proteins, however, is mainly based on studies of the yeast Saccharomyces (review by Steinberg, 1998). The role of these molecular motors in nuclear migration, organelle positioning, and exocytosis during hyphal growth is currently under intensive study. 2. Functions of the Fungal Cytoskeleton MTs are well known for their function in mitosis, where they are crucial for chromosome segregation (Morris and Enos, 1992; Thaler and Haimo, 1996; Jung

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et al., 1998). In addition to intranuclear MTs, astral MTs emanate from the spindle pole body into the cytoplasm, being implicated in mitotic nuclear division (Aist and Berns, 1981; Kaminskyj et al., 1989; Palmer et al., 1992). After mitosis, cytoplasmic MTs are found as long filaments oriented longitudinally within fungal hyphae. Experiments with antimicrotubular drugs have pointed out the importance of cytoplasmic MTs in polarized growth (Howard and Aist, 1980; Howard, 1981; Herr and Heath, 1982; Peterbauer et al., 1992; Pedregosa et al., 1995; Torralba et al., 1996, 1998b). However, all these studies showed that treatment with MT-depolymerizing agents does not stop elongation, indicating that MTs are not essential for tip growth. Some researchers have therefore questioned the involvement of MTs in the apical growth of filamentous fungi, suggesting that MTs could play an indirect role in the process (Heath, 1990; Srinivasan et al., 1996), perhaps by supporting the actin network at the apex (Torralba et al., 1998b). Even so, these inhibitor studies indicate that MTs help determine tip morphology, position nuclei, and maintain cytoplasmic organization in the hyphae. As discussed earlier, MT-based motors have been identified in fungi. These proteins have been implicated in organelle motility, mitotic spindle formation, chromosome separation, nuclear migration, and nuclear distribution (Hackney, 1995). In mammalian cells, the actin cytoskeleton is required for cell motility and surface remodeling. It mediates cell shape changes during mitosis; it is essential for several contractile activities, such as muscle contraction or the separation of daughter cells by the contractile ring during cytokinesis; it controls cell–cell and cell–substrate interactions together with adhesion molecules; and it participates in transmembrane signaling, endocytosis, and secretion (Luna and Hitt, 1992; St¨ossel, 1993; Gottlieb et al., 1993; Bretscher, 1993; Juliano and Haskill, 1993; Nobes and Hall, 1995). In tip-growing cells, apical elongation is governed mainly by F-actin, having several putative functions, including structural reinforcement of the cell cortex at the position of weakest cell wall, force generation for wall expansion under conditions of low turgor, vesicle transport and docking within the apical dome, localization of PM-associated proteins, cytoplasmic migration, and marking of the position of outgrowth during tip establishment (Heath, 1995; Kropf, 1997). The functions of actin in fungal growth are reviewed in more detail in Section II.D. B. Actin Distribution in Hyphae 1. Cytoplasmic Distribution of Actin Labeling of cytoskeletal elements, MTs and actin, has provided much information on the role of these elements in tip growth. Excellent overviews of technical aspects of labeling are available (e.g., Bendayan, 1984; Harris, 1994; Hardham and Mitchell, 1998) and are not included in this article. Actin in filamentous fungi has been localized with fluorescence microscopy both indirectly using immunocytochemistry and, in certain fungi, directly using

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phallotoxins (reviewed by Heath, 1990). Different patterns of actin have been described in fungal cells using light microscopic methods: individual actin plaques, thin actin filaments, coarse actin fibers, diffuse–amorphous cytoplasmic actin, Spitzenk¨orper staining, and subapical fluorescent cross-bands (Hoch and Staples, 1983; Heath, 1987; Salo et al., 1989; Jackson and Heath, 1990b; Temperli et al., 1990; Tiburzy et al., 1990; Raudaskoski et al., 1991; Hasek and Bartnicki-Garcia, 1994; Kaminskyj and Heath, 1994; Srinivasan et al., 1996; Gupta and Heath, 1997; Torralba et al., 1998a). Not all of these arrays are visualized in any single study, and it seems likely that difficulty in preserving actin microfilaments means that often the complete actin array is not preserved by any one technique (see Heath, 1987; Harold and Harold, 1992; Kaminskyj and Heath, 1994). Clearly, innovative approaches are needed in order to obtain more information on the processes occurring during hyphal growth. Perhaps studies similar to that of Doyle and Botstein (1996), in which GFP-actin has been used to visualize movement of cortical actin patches in living Saccharomyces cells, could shed light on the actin distribution during hyphal growth. In oomycetes and true fungi (or eufungi), actin is concentrated in growing apices and in regions of septum formation (Hoch and Staples, 1983; Runeberg et al., 1986; Heath, 1987; Butt and Heath, 1988; Salo et al., 1989; Jackson and Heath, 1990b; Raudaskoski et al., 1991; Roberson, 1992; Harris et al., 1994; McGoldrick et al., 1995; Srinivasan et al., 1996; Torralba et al., 1998a). It seems reasonable to assume that the accumulation of actin at sites of wall growth is indicative of a functional role. However, the way in which it contributes may differ between the species because aspects of its organization vary. At the apex, the most obvious difference is between oomycetes, which have a netlike “cap” of filaments adjacent to the apical PM (Heath, 1987; Temperli et al., 1990; Harold and Harold, 1992), and eufungi, which contain many small peripheral plaques or patches in a similar location. There is also apparent diversity in the eufungi. In the majority of eufungi (Candida [Anderson and Soll, 1986; Akashi et al., 1994], Coprinus [Tsukamoto et al., 1996], Heterobasidion, Paxillus, Suillus [Salo et al., 1989], Neozygites [Butt and Heath, 1988], Neurospora [Tinsley et al., 1996], Saccharomyces [Adams and Pringle, 1984], Schizosaccharomyces [May et al., 1998], and Schizophyllum [Runeberg et al., 1986]), the plaques are tip-high, but in other species (Allomyces [Srinivasan et al., 1996], Aspergillus [Torralba et al., 1998a], Sclerotium [Roberson, 1992], and Uromyces [Hoch and Staples, 1985; Corrˆea and Hoch, 1993]) the plaques are most abundant about 5 ␮m behind the tips. The identity of the plaques is unclear. It has been proposed that the plaques represented the equivalent of focal contacts at which F-actin was attached to the plasmic membrane (Hoch and Staples, 1983; Adams and Pringle, 1984; Kaminskyj and Heath, 1996; Bachewich and Heath, 1997). In fern protonema and algal rhizoids, which lack the dense F-actin meshwork in the apex, there is a transverse ring of cortical F-actin at the base of the apical dome; the cortical ring is associated with tight adhesion of the PM to the cell wall (Kagawa

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et al., 1992; Henry et al., 1996). In Saccharomyces, the plaques appear to be F-actin coiled around tubular invaginations of the PM (Mulholland et al., 1994; Kaminskyj and Heath, 1996). Such invaginations are absent from the tips of hyphal species, where instead at least some plaques appear to coincide with structures termed filasomes (Howard, 1981; Bourett and Howard, 1991; Roberson, 1992; Corrˆea and Hoch, 1993). Filasomes are a class of coated vesicles composed of a microvesicle core surrounded by a dense fibrillar meshwork. Some studies of immunoelectron microscopy supported that actin is a component of the fibrillar meshwork associated with filasomes and that filasomes represent the ultrastructural equivalent of actin plaques visualized with fluorescence microscopy (Bourett and Howard, 1991; Roberson, 1992; Czymmek et al., 1995; Srinivasan et al., 1996). The function of filasomes remains a mystery, although they have been suggested to be related to endocytosis (Mulholland et al., 1994). A fibrillar meshwork similar to that seen associated with filasomes has been observed by electron microscopy along the inner surface of the invaginating PM during septum formation in Fusarium (Howard, 1981), Laetisaria (Hoch and Howard, 1980), Trametes (Girbardt, 1979) and Sclerotium (Roberson, 1992). Whether this fibrillar meshwork is composed of actin and represents the ultrastructural equivalent of the fluorescent cross-band seen by immunofluorescence microscopy at sites of septum formation is unclear, since labeled actin associated with septa has not been observed at the ultrastructural level. At the apex, a network of microfilaments have been observed between the wall vesicles in some ascomycetes (Howard, 1981; Roberson and Fuller, 1988; Torralba et al., 1998b) and in the basidiomycete Sclerotium (Roberson, 1992). In Aspergillus, microfilaments connect wall vesicles to MTs extending up to the apex (Torralba et al., 1998b). These filaments could be composed of actin, since Spitzenk¨orper labeling using fluorescence microscopy has been reported in Allomyces (Srinivasan et al., 1996), Magnaporthe, Trichoderma (Czymmek et al., 1995), Geotrichum (Fig. 1), and Neurospora (Degous´ee et al., 2000). Binding of the actin antibodies to the Spitzenk¨orper core was also reported using immunoelectron microscopy in Magnaporthe (Bourett and Howard, 1991) and Trichoderma (Czymmek et al., 1995), but not in Sclerotium (Roberson, 1992), possibly because of the extreme lability of this structure, which is very sensitive to fixation conditions. 2. Membrane Interactions with the Actin Cytoskeleton A membrane skeleton similar to that in animal cells, minimally containing F-actin, spectrin, and integrin, has been proposed to play a crucial role in the tip growth of hyphae (Heath, 1995). The role of this membrane skeleton would be to reinforce the PM at the tip, which then becomes the site of regulation of tip extensibility. Supporting that actin is associated with the plasmalemma, Degouse´e et al. (2000) have shown that isolated PM of Neurospora contains F-actin, and also spectrin

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and integrin homologs. Coisolation of actin, spectrin, and tubulin has been also described in diverse plant tissues (Abe and Davies, 1995; Faraday and Spanswick, 1993; Ito et al., 1994), emphasizing the membrane skeleton as a general component of cells. Evidence for the existence of a membrane skeleton was first based in the observation of an F-actin cap specifically adjacent to the apical PM in growing tips of Saprolegnia (Heath, 1987), which reinforced the apical PM and protected it against lysis (Jackson and Heath, 1990a). This array of F-actin seems to be universal in the oomycetes (Temperli et al., 1990; Harold and Harold, 1992). However, as discussed before, actin is usually not observed in the form of a filamentous cap in eufungi; instead, it forms plaques in the vicinity of the PM (reviewed by Heath, 1990). It has been proposed that the plaques represented the equivalent of focal contacts at which F-actin was attached to the PM (Hoch and Staples, 1983; Adams and Pringle, 1984). In hyphae, these plaques have been related to filasomes (Howard, 1981; Bourett and Howard, 1991; Roberson, 1992; Czymmek et al., 1995; Srinivasan et al., 1996), whose function is unknown but does not exclude an adhesive-type role. Spectrin, together with actin, is the most common and quantitatively dominant membrane-skeleton molecule. A protein with similarities to animal spectrin is located adjacent to the PM and had a tip-high gradient in hyphae of both Saprolegnia (Kaminskyj and Heath, 1995, 1996) and Neurospora (Degous´ee et al., 2000). In Neurospora, spectrin forms a very tightly concentrated “cap” in the tips of growing hyphae (Degous´ee et al., 2000), whereas in the oomycete Saprolegnia the spectrin gradients were shallower (Kaminskyj and Heath, 1995). The differences between Saprolegnia and Neurospora could indicate a different contribution of these two proteins, actin and spectrin, in the apical membrane stability during extension in these organisms. It demonstrates that in species where the apical actin is less obviously “caplike” and less abundant than it is in Oomycetes, spectrin is more prominent. Spectrin may be the major expansion-resisting part of the morphogenic system in Neurospora, whereas actin may have this function in Saprolegnia, suggesting the possibility of functional complementation between these two membrane-skeleton components. It is possible that part of the strength of the PM derives from an interaction of the membrane skeleton with cell wall polymers. Focal contacts in animal cells contain integrins that mediate the connection between the cytoskeleton, the PM, and the extracellular matrix (Hynes, 1992; Sastry and Horwitz, 1993; Williams and Solomkin, 1999). Integrin homologs are suggested to have a similar function in plant and fungal cells where the cell wall would be the equivalent of the extracellular matrix (Kropf et al., 1988; Kropf, 1992; Lord and Sanders, 1992; Wyatt and Carpita, 1993; Heath, 1995). The presence of integrin in walled cells, including fungi, is based on immunoblot cross reactivity with anti-integrin antisera (Marcantonio and Hynes, 1988; Schindler et al., 1989; Quatrano et al., 1991; Kaminskyj

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Figure 1 Rhodamine-labeled phalloidin stained actin in hyphal tips of Geotrichum showing apical Spitzenk¨orper labeling and subapical peripheral plaques. Micrograph courtesy of Michael Bonham. Bar: 5 ␮m.

and Heath, 1995; Gale et al., 1996; Bachewich and Heath, 1997; Degouse´e et al., 2000) and cloning and functional analysis of the integrin gene INT1 in the human pathogen Candida (Bendel and Hostetter, 1993; Gale et al., 1996, 1998). Furthermore, peptides that interfere with integrin–extracellular matrix interactions also disrupt hyphal differentiation in the plant pathogen Uromyces (Corrˆea et al., 1996). C. Actin-Binding Proteins The actin cytoskeleton contains proteins other than actin itself. Associated with microfilaments are a wide spectrum of accessory proteins, the ABPs, which regulate the form, functions and properties of actin (Bray, 1992). The ABPs are essentially of two types, those that regulate actin assembly and those that are mechanochemical force generators and thus move one polymer relative to the other (e.g., myosin sliding along actin filaments in muscle cells). A large number of ABPs regulate actin assembly by controlling filament formation and crosslinking of the actin network (Pollard and Cooper, 1986; St¨ossel, 1993; Welch et al., 1997; Schafer and Cooper, 1995; Sun et al., 1995). The activities of these proteins are often modulated by signaling molecules such as Ca2+

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or phosphorylated phosphoinositides (Janmey, 1994). Proteins that interact with actin and alter the assembly state of the actin microfilaments in several models have been described in some fungi, principally Saccharomyces (Botstein et al., 1997; Schmidt and Hall, 1998). The phenotypes of mutants defective in the genes specifying these actin-associated proteins tend to be similar to at least some aspects of the actin mutant phenotypes, especially with respect to loss of cell polarity (Novick and Botstein, 1985; Gabriel and Kopecka, 1995; Goodson et al., 1996; Ayscough et al., 1997; Mulholland et al., 1997; McGoldrick et al., 1995). The interaction of filamentous actin with myosin forms the basis for contractile movement, although the actin cytoskeleton is able to generate motility without the participation of myosin. Conventional (class II) myosins were the first molecular motors to be discovered and have been studied for many years because of their role in muscle contraction. In fungi, they seem implicated in septation (Steinberg, 1998; Fischer, 1999). Unconventional or mini-myosins (classes I, III– XII) (Goodson and Spudich, 1993) have been studied for a relatively short time. The class I myosins appear to be involved in a variety of cellular processes, including cell motility, endocytosis and exocytosis, cytoskeletal structure, and organelle movement (Mermall et al., 1998). Class I myosins have been identified in Saccharomyces (Goodson and Spudich, 1995) and Aspergillus (McGoldrick et al., 1995), where they seem to have a role in apical growth. In Saccharomyces, there are two highly related classic type I myosin isoforms, MYO3 and MYO5, which appear to have overlapping activities (Goodson et al., 1996). Deletion of both MYO3 and MYO5 results in severe defects in actin organization and phenotypes associated with actin cytoskeletal dysfunction, including slow growth, accumulation of intracellular membranes and vesicles, cell rounding, random bud site selection, and impaired secretion and endocytosis (Geli and Riezman, 1996; Goodson et al., 1996). In Aspergillus, a gene encoding a class I myosin motor was identified using PCR based on the high sequence conservation of known myosins from different species (McGoldrick et al., 1995). This enzyme was localized at the tips of growing hyphae where it is essential for polarized tip growth. Furthermore, tip growth in at least Schizosaccharomyces is inhibited by BDM, which is known to inhibit myosin activity (May et al., 1998; Steinberg and McIntosh, 1998). However, both yeast buds and hyphal tips are rich in myosin Vs (summarized in Steinberg, 1998). These myosins are probably involved in vesicle or organelle transport and are also inhibited by BDM; thus, it is not easy to interpret BDM inhibition results.

D. Role of Actin in Tip Growth Picton and Steer (1982) were the first to formulate a model drawn for research with pollen tubes that assigns actin filaments a prominent role in regulating tip extension. Cytological and genetic studies on yeast and filamentous fungi leave no doubt that actin and ABPs perform obligatory functions in polarized growth. In

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filamentous fungi, a variety of genes affecting growth polarity have been identified. These include components of the actin cytoskeleton, such as myosin (McGoldrick et al., 1995). Some of the genes affecting cell polarity also affect patterns of actin distribution and/or septation. For example, mutations in the Neurospora regulatory subunit of protein kinase A (mcb-1) cause major defects in growth polarity, actin localization, and septation (Bruno et al., 1996). Cell wall deposition during septum formation is similar to hyphal tip growth in that it also requires recruitment of actin to specific sites (Gow, 1995). Several studies have demonstrated that a transient actin ring forms at the incipient division site and that loss of functional microfilaments prevents septum formation (Harris et al., 1994; Torralba et al., 1998a). In Aspergillus several genes required for growth polarity and septation have been characterized and are related to the cytoskeleton, such as sepA and podB (Harris et al., 1997; Kaminskyj and Hamer, 1998). SEPA and PODB are required for the formation of actin rings during septation in Aspergillus (Harris et al., 1997) and also to organize the actin cytoskeleton at the tips (Harris et al., 1999) in this fungus. Mutations in these genes alter the normal pattern of polarized morphogenesis, suggesting an underlying role of the actin cytoskeleton (Harris et al., 1999). The budding yeast Saccharomyces shifts between isotropic and polarized growth modes (Lew and Reed, 1995; Roemer et al., 1996), and actin is concentrated in areas of growth. Bud growth requires numerous gene products involved in cytoskeletal and secretory functions. These include actin, septins, and products of the BUD genes (Govindan et al., 1995). The fission yeast Schizosaccharomyces also switches between two distinct growth patterns, unipolar and bipolar, and, as is true for Saccharomyces, actin is concentrated in the regions of new cell wall addition (Marks and Hyams, 1985). Recently, two genes important for proper morphogenesis in Schizosaccharomyces have been identified: tea1 and pom1, which play a role in localization of the growth machinery (Mata and Nurse, 1997; Bahler and Pringle, 1998). Like Saccharomyces, filamentous fungi employ two distinct growth modes. When spores of filamentous fungi break dormancy, they first grow isotropically, adding new cell wall material uniformly in every direction. Later they switch to polarized growth, with new cell wall material forming an emerging germ tube. The critical difference in polarity establishment between yeast and filamentous fungi is that the axis of polarity in yeast reorients, but, once established, the axis of polarity in filamentous fungi remains fixed. Studies of swo mutants of Aspergillus suggest that polarity establishment and polarity maintenance are genetically separate events (Momany et al., 1999). Actin seems to have a role in both processes: F-actin is involved in maintaining tip growth; it participates in the maintenance of tip shape (Jackson and Heath, 1990a), stabilization of the tip localization of elements required for tip growth at hyphal apices (Heath, 1995), and the transport and exocytosis of wall vesicles (Novick and Botstein, 1985; Ayscough et al., 1997; Botstein et al., 1997). F-actin is also involved in establishing tip growth. In diverse fungi, F-actin disruption prevents germ tube emergence (Grove and Sweigard,

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1980; Tucker et al., 1986; Vargas et al., 1993; Torralba et al., 1998a). In addition, localized changes in actin organization (radial actin arrays forming) have been detected prior to branching and spore germination in Saprolegnia (Bachewich and Heath, 1998). Similarly, in the initial stages of fission and budding yeast elongation (Marks and Hyams, 1985; Anderson and Soll, 1986), moss protonema side branch formation (Quadar and Schnepf, 1989), and fucoid zygote germination (Brawley and Robinson, 1985), F-actin is observed in diverse patterns at the site of initiation. 1. Morphogenesis A critical aspect of apical growth is the regulation of the extensibility of the cell surface, so that the shape of the hyphae can be generated. There have been many attempts to develop models that will predict and explain apical growth. The traditional view is that morphogenesis in tip growth results from the balance between turgor pressure and the regulated yield of the tip cell wall. The steadystate model of tip growth (Robertson, 1965; Wessels, 1990) invokes changes in the physical properties of the cell wall to explain tip growth. At the tip, the wall is plastic, to permit expansion; a gradient of subapically declining wall plasticity generates the tubular form (e.g., Bartnicki-Garcia and Lippman, 1972; Wessels, 1990). Since the tip is deformable, turgor has been proposed to determine both tip extension rate and hyphal morphology (Wessels, 1988). The validity of these models has been questioned (Money, 1997; Heath and Steinberg, 1999). The wall is important in determining the properties of the hyphal tip, but a poor correlation between growth rates and turgor pressure (Kaminskyj et al., 1992a) and normal growth with little or no turgor pressure (Money and Harold, 1992, 1993; Money, 1997) indicate that turgor pressure may not be obligatory. Working with pollen tubes, Picton and Steer (1982) proposed a model suggesting that an F-actin containing network could be the morphogenic factor. Picton and Steer’s (1982) model envisaged an F-actin cap associated with the tip and anchored to the subapical part of the cell. The extensibility of the cap, or its anchors, would regulate tip extension. In hyphae, the extensibility of the plastic surface of the tip may be partially determined by the membrane skeleton, composed of F-actin and spectrin (see Section II.B.2.; Fig. 2, see color insert). This skeleton restrains or protrudes the hyphal tips under the differing turgor conditions:

r Under normal turgor pressure, the function of the membrane skeleton may be resisting turgor pressure. It has been shown that actin mutants show an increased osmotic sensitivity (Novick and Botstein, 1985; Chowdhury et al., 1992) and antiactin drugs induced tip swelling and/or growth acceleration (Allen et al., 1980; Grove and Sweigard, 1980; Srinivasan et al., 1996; Gupta and Heath, 1997; Torralba et al., 1998a). Swelling and accelerated growth would be in response to turgor in the absence of the normal restraint by actin.

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r Under low-turgor conditions, the apical actin arrays function in a manner comparable to that in animal cell pseudopodia; they protrude the tips (Heath, 1990; Money, 1995; Harold et al., 1995; Heath, 1995; Heath and Steinberg, 1999). F-actin disruption under low turgor pressure resulted in loss of tip protrusion (Gupta and Heath, 1997). This is consistent with the amoeboid model, which compares hyphal tip growth with amoeboid movement inside a tube. The idea is that the hyphal cytoplasm migrates through the tubular hypha as an animal cell migrates over a solid substrate. In the amoeboid movement of animal cells, the cytoplasm protrusions must interact with a rigid substrate that is comparable to the rigid subapical cell wall of fungi. Similarities between hyphal tip growth and amoeboid movement have been reviewed by Heath and Steinberg (1999). The amoeboid model explains many observations concerning tip growth, including an easy explanation for normal tip growth in the absence of turgor pressure. Actin could exert mechanical force by the assembly of a microfilament meshwork from monomers, exerting direct force upon the PM, or it may be credited to actin filaments translocated by myosin I. Myosin is concentrated in both Saccharomyces buds (Goodson et al., 1996) and hyphal tips of Aspergillus (McGoldrick et al., 1995). Irrespective of the regulation of extensibility, it is clear that the morphogenesis of hyphae demands, at least in part, the delivery and exocytosis of wall vesicles that add new membrane and wall material to tip-growing regions. Thus precise regulation of exocytosis in tip-growing cells is a critical part of tip growth. Theoretical models (Heath and Janse van Rensburg, 1996; Bartnicki-Garcia et al., 1989) show that appropriate gradients of exocytosis can generate the hyphal shape. One proposal for the regulation of the apical exocytotic gradient invokes a “vesicle supply center” (VSC) (Bartnicki-Garcia et al., 1989), which distributes vesicles in all directions. Its location in fungal hyphae is coincident with a structure known as the Spitzenk¨orper. The Spitzenk¨orper is recognized as a dense accumulation of vesicles in hyphal tips of actively growing hyphae that appears as a dark sphere when viewed with phase contrast optics (Girbardt, 1957; Howard and Aist, 1977). As the hypha elongates, the linear advance of the VSC would automatically set up a gradient of exocytosis that determines the shape of the apical dome as well as the subapical tube (Bartnicki-Garcia et al., 1989, 1995). It has been suggested that the Spitzenk¨orper position is controlled by the cytoskeleton (Bartnicki-Garcia, 1995; Riquelme et al., 1998). The model implies that the site of exocytosis is determined by the site of impact of the wall vesicles at the end of their trajectories from the VSC. However, Heath and Janse van Rensburg (1996) observed that both vesicle and large organelle trajectories in hyphae of Saprolegnia are contrary to predictions of the VSC model, indicating that the model does not apply to this organism, and thus cannot be universal for tip growing cells. These authors proposed a new model in which a gradient (highest at the apex) of vesicle-fusion sites alone is sufficient to generate the hyphal shape (Heath and Janse van Rensburg, 1996).

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The tip-high gradient distribution and/or activation of membrane proteins known as SNAREs can meet the requirement for the generation of such an exocytotic gradient. These proteins are highly conserved integral membrane proteins that are an essential component of the regulation of vesicle fusions and that are believed to confer specificity to vesicle docking and fusion reactions on the basis of protein– protein recognition (Ferro-Novick and Jahn, 1994). Gupta and Heath (2000) have provided the first evidence for the presence of a PM t-SNARE homolog in fungal hyphae. Its activity and tip-high gradient distribution could provide the necessary spatial/temporal control of exocytosis that is needed for hyphal tip formation. It has been suggested that the distribution of t-SNAREs may be retained by F-actin associated with the PM (Drubin and Nelson, 1996). 2. Branch Initiation Hyphal branching makes possible mycelium development. Although the kinetics of branching have been extensively studied in several fungi (Trinci, 1984), the mechanisms of branch initiation are not yet clear. Branch formation is reflected by a redistribution of cytoskeletal elements. Localized changes in actin organization have been detected prior to branching in tip growing cells, suggesting an actin role in initiating tip formation. Bachewich and Heath (1998) demonstrated that radial actin arrays form in Saprolegnia in regions with no detectable surface protrusion. Similar radial arrays preceded germ tube formation in asexual spores. Longitudinally oriented actin arrays focused on the future site of septation in a fission yeast–like fungus (Butt and Heath, 1988), and radial arrays occurred in regenerating pollen tube protoplasts (Rutten and Derkson, 1990), but in the latter, the patterns were variable and did not predict the site of outgrowth. In other organisms F-actin patterns preceding polar growth do not include radial organization. For example, a membrane-associated ring of filaments precedes branch formation in moss protomemata (Quadar and Schnepf, 1989), but its functions are unknown. The localized movement and reorganization of membrane-bound actin, as shown by the production of the radial F-actin arrays, could induce changes both in the cell wall and in the PM necessary for branch growth. At developing branch sites, the mature wall must be softened for extension to begin (Mullins, 1979). It has been proposed that the wall is plasticized by lytic enzymes transported in the wall vesicles, so that the relative frequency of vesicle insertion determines wall deformability (Bartnicki-Garcia, 1990). Bracker et al. (1997) showed that the Spitzenk¨orper can be moved in live fungal cells, during manipulation with a laser beam (optical tweezers). When the Spitzenk¨orper was forced away from its usual position in the apex, it would start deforming any wall adjacent to it: It could make a bump on the subapical wall, change the direction of growth of the hyphal tube, or even start a new branch, confirming that the wall vesicles contain enzymes able to soften the cell wall. One of actin’s functions in initiating branching is probably directing the delivery of wall vesicles containing wall lytic enzymes to the points

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of new branch formation at the presumably inextensible, subapical walls. Actin could also induce changes in the localization and anchorage of Ca2+ channels, glucan synthases, and vesicle docking proteins in the PM (see Section II.D.3.). Colocalization of all these components would lead to accelerating, self-sustaining hyphal tip growth (Fig. 3, see color insert). Elevations in Ca2+ concentration influence the site of branch formation, which could affect the gradient of F-actin polymerization (discussed in Sections III.C.2. and III.C.3). ABPs are probably also implicated: In root hairs, profilin, which is known as a potent regulator of actin dynamics in both plant and animal cells (Sun et al., 1995), is present in the bulge of outgrowing root hairs (Braun et al., 1999). 3. Membrane Protein Distribution Hyphal tips contain PM proteins are likely to show a tip-high gradient of abundance: enzymes responsible for the synthesis of the fibrillar cell wall polymers (e.g., chitin and cellulose), ion channels, and SNARE proteins, which are essential for the regulation of vesicle fusions. Several observations are consistent with the notion that F-actin can serve to localize these elements required for tip growth at hyphal apices. There are two basic mechanisms by which gradients of PM proteins can be generated and maintained: dynamic localized apical insertion and subapical removal or static linkage to a skeletal system at the apex. Because the vesicle transport system in the tips utilizes the cytoskeleton (see Section II.D.4.) and because the plasmalemma-associated F-actin system is well situated to play the skeletal role (see Section II.B.2.), the cytoskeleton is likely to be responsible for the gradients by either model. Since the PM is constantly being synthesized at the hyphal tips by exocytosis of the wall vesicles, apical insertion is very likely. Presumably general membrane turnover, or possibly endocytosis (Hoffmann and Mendgen, 1998), would account for subapical removal. However, there is evidence that, at least in Saprolegnia, the tip-high gradient of mechanosensitive Ca2+ channels is maintained by action of the actin-containing membrane skeleton (Levina et al., 1994). In addition, in Neurospora, the distribution of the spectrin–actin network at the apex can be superimposed upon the t-SNARE derived exocytotic gradients, as would be expected if this membrane skeleton was responsible for tethering proteins involved in vesicle docking and fusion to the apical domain (Gupta and Heath, 2000). F-actin also seems to regulate wall fibril synthetic enzyme distribution. For example, actin disruption induces delocalization of wall synthetic enzymes (Novick and Botstein, 1985; Gabriel and Kopecka, 1995; Chiu et al., 1997), which could produce the subapical wall thickenings observed in hyphae treated with latrunculin (Bachewich and Heath, 1998) and cytochalasins (Allen et al., 1980; Grove and Sweigard, 1980). Fungal mutants containing disrupted actin showed similar fibril-enriched wall thickenings (Chiu et al., 1997; Gabriel and Kopecka, 1995) and delocalization of chitin synthesis (Novick and Botstein, 1985). Thus, it is likely

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that F-actin restrains wall synthetic complexes to the apex, as it does for membrane channel proteins (Levina et al., 1994). 4. Vesicle Transport and Exocytosis The process of apical growth in fungi involves the net migration of wall vesicles transporting raw materials for cell growth from the Golgi bodies or equivalents to the hyphal tip, where they are ultimately exocytosed. Localized exocytosis is the key to forming a tube that extends at its tip. Therefore, the mechanisms transporting these vesicles to the correct destination at the tip and then ensuring their accurate site and rate of exocytosis are clearly fundamental to the process of tip growth. The cytoskeleton plays a major role in these processes. The transport mechanism proposed by Regalado and Sleeman (1999) predicts that vesicle migration is a diffusion-driven process, with cytoskeletal viscoelastic forces representing the necessary bias. However, this mechanism for vesicle motion implies no intimate contact between vesicles and cytoskeleton filaments, and therefore motor proteins, such as kinesin, dynein, and myosin, would not be required. This model disagrees with experimental observations that confirm a central role of these mechanoenzymes in organelle transport, polarized growth, and secretion in fungi (discussed in Sections II.A.2 and II.C). By analogy with the functionally comparable neuronal system (Bray, 1992), MTs were proposed to function as tracks, and vesicles would be coated with mechanochemical effectors such as cytoplasmic dynein or kinesin so that they slide along the MTs to the hyphal tip. There are data to support aspects of this prediction. Cytoplasmic MTs run parallel to the long apex of the hypha and thus to the direction of transport (Roberson and Fuller, 1988; Heath and Kaminskyj, 1989; Temperli et al., 1991; Raudaskoski et al., 1991; Torralba et al., 1998b). Research on kinesins in fungi shows a possible role of these proteins in the delivery of wall vesicles to the hyphal tip (Steinberg and Schliwa, 1993, 1996; Steinberg, 1997; Lehmler et al., 1997; Seiler et al., 1997). However, there are data contrary to a role of the MTs system in apical vesicle transport. At least some kinesin mutations (Enos and Morris, 1990; Yamashita and May, 1998) and other mutations (Oakley and Rinehart, 1985; Osmani et al., 1990) or drugs (Herr and Heath, 1982) that inhibit MT-based processes do not disrupt tip growth. Contrary to the situation of the neurons, wall vesicles do not show any special morphological association with MTs (Heath, 1995). In addition, disruption of MTs does not always induce the disassembly of the apical accumulation of wall vesicles as observed by Howard and Aist (1980); it has been reported that the apical vesicle accumulation remains intact (Herr and Heath, 1982), changes its position (Rupeˇs et al., 1995), or even increases in number of wall vesicles (Torralba et al., 1998b) in the presence of antiMT drugs. An indirect role of MTs in wall vesicle transport has been suggested, perhaps by supporting the essential organization of the actin at the tips (Heath, 1990; Srinivasan et al., 1996; Torralba et al., 1998b).

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It is established that vesicles travel along actin microfilament tracks in conjunction with myosin, in tip growing cells of plants (reviewed by Kropf et al., 1998). Actin could also have a primary role in the movement and fusion of wall vesicles during apical growth in fungi (Heath, 1990; Johnston et al., 1991). Indeed, electron microscopic observations have shown vesicles in close association with microfilaments (Howard, 1981; Heath and Kaminskyj, 1989; Vargas et al., 1993). In Saccharomyces, mutants defective in actin or actin-associated proteins display characteristic defects in cell morphology and protein secretion, and accumulate vesicles (reviewed by Welch et al., 1997; Botstein et al., 1997). However, MTs are not essential (Huffaker et al., 1988). It appears that myosin motors, translocating on actin cables, direct at least some types of vesicles to the vicinity of the budding or mating site in Saccharomyces (Govindan et al., 1995; Finger and Novick, 1998; Pruyne et al., 1998). In Aspergillus, the myosin I motor was localized at the tip of growing hyphae, where it is essential for polarized tip extension and seems implicated in the movement of wall vesicles to the apex (McGoldrick et al., 1995). Moreover, secretion of acid phosphatase, an exoenzyme localized in wall vesicles of filamentous fungi (Hill and Mullins, 1980), was strongly reduced in mutants with deletion of myosin. 5. Organelle Movements and Positioning In fungi, the correct distribution of their organelles is vital to their efficient physiology, growth, development, and reproduction. Since the distribution of organelles is maintained as the colonies grow, a prerequisite for polarized hyphal growth is multiplication and the transport of all cell organelles as hyphae elongate to keep the polarized organization of the hyphae (Suelmann et al., 1997, 1998; Fischer, 1999). Organelle migration requires cytoskeletal components, mechanochemical motor proteins, and regulatory proteins to control movement both spatially and temporally. As discussed before, a class I myosin motor in Aspergillus is essential for polarized tip extension and seems implicated in the movement of wall vesicles to the apex (McGoldrick et al., 1995). Whether the actin–myosin system is also involved in the movement of other organelles, or the MT–dynein or MT–kinesin systems are also implicated, is not known. Nuclear movement and positioning are essential processes in fungal development (reviewed in Aist, 1995; Morris et al., 1995; Fischer, 1999). In hyphae, nuclei keep up with the growing tip and appear stationary compared to the growing apex. In addition to this positioning effect, nuclei show active movement at rates up to several micrometers per second (Steinberg and Schliwa, 1993; Suelmann et al., 1997; reviewed in McKerracher and Heath, 1987). Nuclear migration is dependent on MTs and MT-dependent motor proteins (Oakley and Morris, 1980; Heath, 1982; McKerracher and Heath, 1986a; Hoch et al., 1987; That et al., 1988; Heath and Kaminskyj, 1989; Hyde and Hardham, 1992; Steinberg and Schliwa,

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1993; Eshel et al., 1993; Xiang et al., 1994, 1995, 1997; Fischer, 1999). Cytoplasmic dynein is thought to be the major motor for nuclear migration in filamentous fungi (reviewed in Morris et al., 1995). Besides dynein, kinesin has been implicated in nuclear migration in Neurospora (Seiler et al., 1997; Henningsen and Schliwa, 1997). Because mutations of this motor protein resulted in a clustering of nuclei, the function could be required for nuclear positioning rather than for nuclear migration itself. Surprisingly, some studies in Saccharomyces implicate the actomyosin system in nuclear migration (Watts et al., 1987; Palmer et al., 1992), suggesting that both the actin and MT systems are necessary. In many fungi, mitochondria are thought to move along MTs (Herr and Heath, 1982; Aist and Bayles, 1991; Steinberg and Schliwa, 1993; reviewed in Heath, 1995); in other fungi, however, a MT-independent mechanism has been described (Huffaker et al., 1988; Oakley and Reinhart, 1985; reviewed in Heath, 1995). The actin cytoskeleton and intermediate filaments seem to be important for mitochondrial movement in Saccharomyces (Simon et al., 1995; Hermann et al., 1997; McConnell and Yaffe, 1992; Fisk and Yaffe, 1997; Berger et al., 1997; Boldogh et al., 1998). The knowledge of movement and the dynamics of other organellar systems, such as the Golgi apparatus, ER, and vacuoles, is very limited. Vacuole migrations have been associated with MTs (Herr and Heath, 1982; Shepherd et al., 1993; Hyde et al., 1999), but the shape of vacuoles in Saprolegnia appears to be F-actin imposed (Bachewich and Heath, 1998). Actin colocalization with vacuole membranes and a role of actin–myosin in directed vacuole movement have been demonstrated in Saccharomyces (Hill et al., 1996). The effects of experimental perturbations are puzzling and make it difficult to draw any absolute conclusions regarding the precise role of actin and MTs. It is possible that different cytoskeletal elements are responsible for movement and positioning of different organelles and/or that different cytoskeletal components direct a given process in different organisms. The action of both MT and actin microfilaments may be required. These two systems have been shown to have a close interrelationship (see Section II.D.7), and studies have demonstrated the movement of the same organelles on both types of filament (Langford, 1995). It may also be that the simple actin and/or MT dichotomy is inadequate. Characterization of MDM mutants of Saccharomyces has indicated that nuclear and mitochondria transmission to daughter cells depends on Mdm1p. This protein is able to form filaments in vitro and was compared to intermediate filaments (McConnell and Yaffe, 1992; Fisk and Yaffe, 1997; Berger et al., 1997). 6. Cytoplasmic Migration As fungal hyphae grow via wall deposition and expansion at their tips, the cytoplasm of the cell continually relocates with respect to the lateral walls in order to remain near the tip. The force for this migration seems to be intrinsic to the

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apical cytoplasm, which moves forward actively to maintain its position as the tip extends, in an amoeboid way (Kaminskyj and Heath, 1996). The cytoplasmic migration during tip growth and its similarities with amoebal movement have been reviewed (Heath and Steinberg, 1999). Thus, cytoplasmic migration in hyphae is an active process, likely involving contractions similar to those of amoeboid cells. Consistent with this, fungal cytoplasm can show extensive active contraction (McKerracher and Heath, 1986b, 1987; Heath, 1990; Kaminskyj et al., 1992b; Jackson and Heath, 1992; L´opez-Franco and Bracker, 1996; Reynaga-Pe˜na et al., 1996; Reynaga-Pe˜na and BartnickiGarcia, 1997). In common with pseudopodia, these contractions are (a) apparently Ca2+ mediated (McKerracher and Heath, 1986b; Jackson and Heath, 1992; Kaminskyj et al., 1992b), (b) rich in F-actin (McKerracher and Heath, 1987; Heath, 1990; Jackson and Heath, 1992), and (c) predominantly unidirectional toward the tips (McKerracher and Heath, 1986b; Heath, 1990; Jackson and Heath, 1992; Kaminskyj et al., 1992b; L´opez-Franco and Bracker, 1996), thus indicating an underlying cytoskeletal organization capable of forward protrusion. Amoeboid cells require cytoskeleton/substrate attachments, such as focal contacts, in order to anchor the force necessary for movement. These contain integrins that mediate the connection between the cytoskeleton, the PM, and the extracellular matrix (Hynes, 1992; Sastry and Horwitz, 1993; Williams and Solomkin, 1999). A similar function has been proposed in plant and fungal cells where the cell wall is the equivalent of the extracellular matrix (Kropf et al., 1988; Kropf, 1992; Lord and Sanders, 1992; Wyatt and Carpita, 1993; Kaminskyj and Heath, 1995, 1996; Bachewich and Heath, 1997). Candida (Marcantonio and Hynes, 1988; Bendel and Hostetter, 1993), Saprolegnia (Kaminskyj and Heath, 1995), and Neurospora (Degouse´e et al., 2000) have immunoreactive integrin homologs, whose distribution and behavior suggest that they are important for anchoring the cytoplasm to the cell wall. 7. Relationships with Microtubules: Integration of Cytoskeletal Activity Although actin plays the major role, MTs are also required for optimal tip growth. Since a close association between microfilaments and MTs has been demonstrated in hyphae (Heath and Heath, 1978; Hoch and Staples, 1983; Heath and Kaminskyj, 1989; Vargas et al., 1993; Srinivasan et al., 1996; Torralba et al., 1998b) and plant cells (Lancelle and Hepler, 1991), we assume that MTs more likely play an indirect role in apical growth. In Aspergillus, the antimicrotubule MBC is known to cause alterations of growth and reduction of enzyme secretion (Jochov`a et al., 1993; DeLucas et al., 1993; Torralba et al., 1996, 1998b). The microscopical observation of MBC-treated hyphae indicate that the disassembly of cytoplasmic MTs by the drug-induced changes in the pattern of actin at the hyphal tips (Torralba et al., 1998b). In the yeast Candida, actin distribution has also been reported to change

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after depolymerization of MTs with benomyl (Akashi et al., 1994). These results suggest that in fungal hyphae, MTs organize the actin at the tip. Cytoplasmic MTs could thus play an indirect role in growth and secretion by supporting the actin network at the apex. However, in Saprolegnia F-actin arrays remain normal in nocodazole-treated hyphae (Gupta and Heath, unpublished results), whereas latrunculin B induces concomitant alteration in MT and actin organization (Gupta and Heath, 1997). Therefore, F-actin could stabilize MT organization in this organism. Since similar effects result from incubation in latrunculin with nocodazole and in latrunculin alone, the MT rearrangements are a consequence of the alterations in the actin cytoskeleton induced by latrunculin B. Although there have been many hints through the years of synergy between MTs and actin filaments in eukaryotic cells (Gavin, 1997), there has been little solid evidence about how they might interact. A direct physical interaction between motors would be a very straightforward way of mediating coordination. Fath et al. (1994) found evidence for cooperation between a class I myosin and dynein: These motors copurified with Golgi-derived vesicles from a polarized epithelial cell, and a given vesicle could bind to both actin filaments and MTs. It has become clear that several different transport events, previously thought to be based solely on actin (pigment transport in mouse melanocytes and fish retinal pigment epithelium, budding in yeast) or MT (axonal transport, pigment transport in melanophores), instead involve (or may involve) different balances of cooperative interactions between the two systems. Most of these events employ class V myosin as well as kinesins and dynein (Brown, 1999). The organization and functions of the cytoskeletal polymers can be determined by Ca2+ , which therefore seems to have a role in apical growth; Ca2+ is also related to exocytotic processes in eukaryotic cells (Douglas, 1968; Katz, 1969; Hepler and Wayne, 1985; Gomperts, 1986; Hille et al., 1999), and exocytosis is a fundamental process in apical growth. Therefore, investigations of the ionic regulation of hyphal growth have focused most attention on the role of Ca2+ (Jackson and Heath, 1993, and references therein).

III. Calcium and Apical Growth in Fungi Calcium is of central importance in eukaryotic cell signaling and is involved in the regulation of a wide range of biological activities. Cytoplasmic Ca2+ has emerged as an intracellular regulator of central importance in transducing external stimuli in animals (reviewed by Berridge et al., 1999) and in plants (Tretyn, 1999; Knight, 2000). In fungi, intracellular Ca2+ probably participates in multiple regulatory functions, including branching, sporulation, tip growth, hyphal reorientation toward localized stimuli, cyst germination, blue light–induced conidiation, regulation of dimorphism, zoospore motility, pheromone-mediated sexual reproduction, cell cycle control, and cytokinesis (see references in reviews by Pitt and Ugalde,

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1984; Jackson and Heath, 1993; Gadd, 1994; also, Hyde and Heath, 1995; Jackson and Hardham, 1996; Grinberg and Heath, 1997). Several lines of argument indicate a central, coordinating role for cytoplasmic Ca2+ in tip growth. Ca2+ may be important to the polarized transport of wall vesicles and their fusion during apical growth, and to influence the structure and rigidity of the actin network in the hyphal tip. Ca2+ is well suited to such a role, given its ubiquitous functions as second messenger in eukaryotic cells, where Ca2+ regulation of the cytoskeleton and vesicle fusion rates is well known (Douglas, 1968; Katz, 1969; Hepler and Wayne, 1985; Gomperts, 1986; Jackson and Heath, 1993; Hille et al., 1999). A. Internal Ca2+ Gradients in Hyphae: Methods for Their Detection The most striking feature of cytosolic Ca2+ distribution in fungi is the presence of a tip-focused gradient of cytosolic free Ca2+ as a general feature of growing fungal hyphae. Similarly, Ca2+ gradients are also associated with cell polarity in other highly polarized cells such as zygotes and rhizoids of algae (Brownlee and Wood, 1986; Berger and Brownlee, 1993; Brownlee and Bouget, 1998), pollen tubes and root hairs of higher plants (Clarkson et al., 1988; Miller et al., 1992; Malho et al., 1994; Pierson et al., 1994; Felle and Hepler, 1997; Hermmann and Felle, 1995; Bibikova et al., 1997, 1999), and neurites of animal cells (Connor, 1986; Mattson, 1999). All of these tip-growing cells have a higher concentration of Ca2+ in their apices than in their subapical regions. No apical growth has been observed in the absence of Ca2+ gradients. The tip-high gradient of Ca2+ distribution is thought to play a role in establishing and maintaining apical organization, morphogenesis, and growth. In this section we review what is known about Ca2+ distribution in growing hyphae. The surprising paucity of imaging studies of Ca2+ in fungal hyphae may be explained by the technical complexity of imaging Ca2+ (Gadd, 1994; Knight et al., 1993). Therefore in this section we critically discuss the techniques for imaging Ca2+ . These methodologies are driving continual technical development, thus providing new means of visualizing Ca2+ and improved methods of detection. The typical tip-to-base Ca2+ gradient in tip-growing cells can be detected by all available methods that show the distribution of intracellular Ca2+ , independent of whether the methods indicated cytosolic free, membrane-associated, and organellestored Ca2+ or the total calcium content. 1. Free Ca2+ Several methods that have been used to localize intracellular ionized Ca2+ rely primarily on chelating agents such as Fluo-3, Fura-2, Calcium Green, and more recently on aequorin-based probes (see review by Hyde, 1998). The necessity of

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using a probe introduces the first potentially serious but unavoidable biological perturbation that is faced in every Ca2+ imaging study: Ca2+ bound by a probe molecule is not available for cellular metabolism. Thus, all imaging studies must proceed under the assumption that at probe concentrations that provide sufficient fluorescence for visualization, Ca2+ buffering is not severe enough to interfere with normal cellular activities. Fortunately, with fast tip growing cells, there is a very sensitive indicator of normal physiology. Any problems with the dye-loaded hyphae are indicated by abnormalities in hyphal morphology or growth rates (Jackson and Heath, 1993). An additional problem with most Ca2+ probes is the need for light stimulation in the excitation wavelength/s of the probe, which can also affect normal functioning of the hyphae. An alternative strategy involves using organisms genetically transformed with the aequorin gene (Knight et al., 1991). Aequorin is a luminiscent Ca2+ -sensitive photoprotein that emits light on binding to free Ca2+ in a dose dependent manner. Unfortunately, the light output from this dye has so far only been suitable for use with mycelia, and not individual cells (Parton and Read, 1998). The literature is replete with studies in which Ca2+ has been successfully localized in mammalian cells; however, for plant cells, and in particular for fungal cells, accurate localization of Ca2+ has been more difficult to achieve. For fungi, even what is typically the very first step of Ca2+ imaging study, dye loading, has been problematic. Loading of Ca2+ probes per se is not the problem in fungi, but postloading sequestration: Too often the ionized Ca2+ -measuring probes become sequestered in various organelles in these organisms, making it difficult to interpret the cytosolic levels of calcium (Knight et al., 1993; Read et al., 1992). In this decade there have been only seven reports of success (Garrill et al., 1993; Jackson and Heath, 1993; Levina et al., 1995; Yuan et al., 1995; Jackson and Hardham, 1996; Hyde and Heath, 1997; Silverman-Gavrila and Lew, 2000). Most of these studies were of oomycetes, with only two reports of success with a true fungus, Neurospora (Levina et al., 1995; Silverman-Gavrila and Lew, 2000). In other attempts to visualize Ca2+ in true fungi, dyes were sequestered into cytoplasmic organelles (Knight et al., 1993; Chandra et al., 1999). Hyde and Heath (1997) were able to demonstrate apparent cytosolic free Ca2+ distribution with little intracellular sequestration of the dye in hyphal apices of Saprolegnia using lower concentrations of the fluorescent dye. More recent studies by Read et al. (1998) have shown that pressure microinjection of a 10 kDa dextran conjugate of Oregon Green-1 shows promise in overcoming some aspects of this problem. Interestingly, loading of hyphae of oomycetes and true fungi with the pH dye carboxy-SNARF has not been so problematic (Parton and Read, 1998; Bachewich and Heath, 1997), for unknown reasons. The distribution of cytoplasmic free Ca2+ in hyphae has been studied by using two types of fluorescent dyes:

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(a) Single-wavelength dyes such as Fluo-3 or Calcium Green change only fluorescence intensity when bound to Ca2+ . Misinterpretation of the Ca2+ concentration in the cell can occur because the fluorescence intensity of these dyes depends on both dye and Ca2+ concentrations; therefore, regions of cells with a larger accumulation of dye will fluoresce more intensely, regardless of the Ca2+ concentration. With single-wavelength dyes, fluorescence also depends on organelle density, with higher signals emerging from the regions with fewer organelles (e.g., see Hyde and Heath, 1997). However, these problems can be solved by using cells dual-loaded with Ca2+ -sensitive and Ca2+ -insensitive dyes (“dual dye ratio technique”). Variations in the ratio between the intensities of the two dyes, the Ca2+ -sensitive emissions, and the Ca2+ -independent emissions, should indicate variations in Ca2+ concentrations, assuming that there are no differences in concentrations of the two dyes. (b) Radiometric dyes shift either their excitation (Fura-2) or their emission (Indo-1) spectra upon binding Ca2+ . By measuring the ratio between the intensity of fluorescence at the Ca2+ -bound and Ca2+ -free wavelengths, it is possible to determine the concentration of free Ca2+ , independently of possible variations in dye concentration in different parts of the cell. The radio dyes would be thus more suitable for detecting a Ca2+ gradient. Unfortunately, their affinity for Ca2+ is affected by fluctuations in cellular pH, protein concentration, and cytoplasmic viscosity (Lattanzio and Bartschat, 1991; Haugland, 1996). In addition, these probes require UV excitation, and UV photons are high energy and may damage cell components (Konig et al., 1996). This phototoxicity often severely limits how long one can view a cell in a fluorescence study. Interestingly, UV damage can result in Ca2+ release, which has been used as an experimental tool (Hyde and Heath, 1997; Grinberg and Heath, 1997). Another problem is that UV wavelengths, and shorter wavelengths in general, are also most likely to excite “autofluorescence,” that is, fluorescence from cellular components. A possible solution is the use of two-photon laser scanning microscopy, which appears to be very promising: Using high-energy pulsed lasers, many probes can be triggered by light of double their normal excitation wavelength (Szmacinski et al., 1993). Near-infrared light is much less problematic than UV, and also penetrates much deeper. Studies on the Ca2+ distribution in hyphae are rare, no doubt because of the problem of sequestration of the dyes discussed earlier. At present there are only two genera, Neurospora (Levina et al., 1995; Silverman-Gavrila and Lew, 2000) and notably the oomycete Saprolegnia (Garrill et al., 1993; Jackson and Heath, 1993; Yuan et al., 1995; Hyde and Heath, 1997), where Ca2+ imaging has come to significant conclusions concerning the nature of the gradients and their role in tip growth. In these studies, the observation of hyphae with normal morphology and growing at normal rates revealed a general trend: that the highest concentrations of the indicator probe–Ca2+ complex were visualized predominately in the apical

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regions of the cells, although there are differences reported as to whether the extreme apex contains the highest concentrations. These differences seem to depend not only on the species studied but also on the method used. In Saprolegnia, use of the radiometric dye Indo-1 indicates that [Ca2+ ] is highest at the very tip of the hypha (Garrill et al., 1993), whereas the fluorescence of the nonradiometric Ca2+ dye, Fluo-3, peaks 3–9 ␮m back (Jackson and Heath, 1993). However, the subapical position of the Ca2+ peak suggested by Fluo-3 seems to be artifactual, resulting from a high cytoplasm to organelle ratio in this region (Jackson and Heath, 1993). Indeed, when Fluo-3 was ratioed against SNARF-1, the Ca2+ peaks at the extreme tip in Saprolegnia hyphae (Hyde and Heath, 1997). In contrast, use of the dual dye ratio technique showed that Neurospora contains a tip-high gradient that peaks 3 ␮m behind the tip (Levina et al., 1995; Silverman-Gavrila and Lew, 2000). 2. Membrane-Associated Ca2+ A simple method to visualize calcium is the use of chlortetracycline (CTC) fluorescence. CTC readily enters cells and its fluorescence demonstrates the presence of membrane-associated Ca2+ , although its absence may not always represent the reverse (Caswell, 1979; Blinks et al., 1982; Kauss, 1987). In contrast to the aforementioned indicators, CTC is a highly effective antibiotic, causing distinct effects after longer application (Reiss and Herth, 1979). Therefore, its use as a vital probe is only possible within the first few minutes. Other restrictions are the ionic composition as well as the pH of the medium, which can lead to different fluorescence patterns (Tang and Beeler, 1990; Jackson and Heath, 1993). The Ca2+ form a complex with CTC, which subsequently associates with adjacent membranes, its fluorescence simultaneously increasing significantly. Tip-high gradients of CTC-membrane-associated Ca2+ have been observed in the fungus Neurospora (Schmid and Harold, 1988; Prokisch et al., 1997) and in Saprolegnia (Jackson and Heath, 1989; Yuan and Heath, 1991; Yuan et al., 1995). The steepness of the intracellular gradient is influenced by variations of the concentration of exogenous Ca2+ in Saprolegnia (Jackson and Heath, 1989; Yuan et al., 1995). Assuming that Ca2+ in the complex is in equilibrium with that in the cytosol, CTC fluorescence could give some indication of the distribution of the total intracellular Ca2+ (Schmid and Harold, 1988). However, it is somewhat questionable whether CTC fluorescence indicates any physiologically relevant Ca2+ distribution or only the distribution of membranes or certain organelles. 3. Total Calcium Intracellular ionized Ca2+ concentrations can be directly influenced by the release and/or uptake of cytosolic Ca2+ by intracellular stores under physiologically normal and pathological situations. To elucidate the function of calcium in

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apical growth, the measurement of both ionized calcium and stored total calcium is necessary. Most of the techniques for localizing total calcium can get down to the TEM resolution, and ultrastructural knowledge of calcium distribution may indicate which organelles are actively involved in sequestering calcium or functioning as calcium sinks during tip growth. Information regarding the localization of total calcium in identified subcellular compartments in all eukaryotic cells is still fragmentary, primarily because of (a) inadequacy of preparative procedures for the maintenance in the specimens of the original elemental distribution and/or (b) limitations of analytical techniques. a. Preparative Procedures. The methods to localize total calcium are not suitable for live cell analysis. Ca2+ is very small and soluble; therefore, a careful sample preparation is required to fix the cells and immobilize the ions in their native state for subsequent analysis. Otherwise, most free Ca2+ is washed out or redistributed during preparation of specimens. Different procedures have attempted to minimize the diffusion of the element:

r Classical chemical fixation is not suitable for localizing calcium within the cells, since loss/redistribution of Ca2+ take place during exposure to aqueous fixatives and dehydration solutions. In some animal (Chan et al., 1984; Probst, 1986; Blanco and Villela, 1998) and plant (Van Iren et al., 1979; Wick and Hepler, 1982) cells, the preservation of the native distribution of Ca2+ was attempted by addition to fixatives of Ca2+ precipitating anions. However, artifactual accumulations and/or displacements before the element is ultimately immobilized cannot be excluded. Subsequent elemental analysis of the precipitates is necessary to confirm the presence of Ca2+ , since most of the anions used in these studies are not specific for Ca2+ . We have used this method, chemical fixation and precipitation with the anion pyroantimoniate, to localize Ca2+ in hyphae by electron microscopy for the first time. Precipitates of calcium pyroantimoniate are observed associated to wall vesicles in the apices of Neurospora (Fig. 5). These deposits were confirmed to contain Ca2+ by X-ray microanalysis (not shown). r Quick-freezing and freeze-substitution can also cause redistribution and loss of cell components, including ions, since the extraction of water from the samples requires exposure to organic solvents in which ions are soluble. A method to decrease the washout of ions can be freeze substitution with a nonpolar substitution fluid such as ether, in which ion solubility is lower (Harvey et al., 1976; Harvey, 1982; Hodson and Sangster, 1989). This method, freeze-substitution with a nonpolar solvent, has not been used to localize calcium in fungal hyphae. r A physical approach that grants ionic preservation is long-term freeze-drying, after quick-freezing. The quick-freezing–freeze-drying

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approach has been exploited only to a limited extent to investigate cell ultrastructure and composition in animal cells (Jorgensen and McGuffee, 1987; Grohovaz et al., 1996; Pezzati and Grohovaz, 1999). This method has been used in only one fungus, Uromyces, to preserve fungal cells for subsequent calcium mapping using ion microscopy (Chandra et al., 1999). As pointed out by Hodson and Sangster (1989), even if material is prepared in anhydrous conditions during fixation and embedding, soluble ions are likely to be lost during sectioning, since sections must be cut onto water. The solution could be the use of cryosections, but this implies the use of thick preparations, limiting the choice of the subsequent analytical technique and reducing the sensitivity. b. Analytical Techniques. Different analytical methods can be used to identify and localize the calcium in the samples where diffusion of the element has been minimized.

r Until now mapping of elements within the cell was obtained primarily by X-ray microanalysis, using scanning probes that collect X-rays inside the transmission electron microscope (reviews by Hodson, 1995; Morgan et al., 1999). In these studies, the use of cryosections is certainly advantageous and has been shown to preserve the distribution of diffusible ions to be analyzed. Because of the physical basis of the method, thinner sections produce higher spatial resolution, but with lower detection sensitivity. In practice, the best assays (spatial resolution ∼100 nm) have been carried out on 100 to 200 nm thick, freeze-dried cryosections, cut from quick-frozen samples (Thibaut and Ansel, 1973, 1976; Somlyo et al., 1977). Most often, thicker sections were employed (up to 1 ␮m), with higher elemental sensitivity but lower spatial resolution, which is only appropriate for large structures inside the cell. X-ray microanalysis has been used to detect calcium in polyphosphate granules in chemically fixed and sectioned fungal specimens (Doonan et al., 1979). However, chemical fixation is likely to have distorted this distribution. Calcium was analyzed by X-ray microanalysis in freeze-quenched hyphae of Chaetomium showing the highest concentrations in the apical regions, with significantly lower concentrations with increasing distance from the apex (Galpin et al., 1978). r Electron energy loss microanalysis can be employed to detect ions within the cells in the spectrum mode (electron energy loss spectroscopy; EELS) and/or in the image mode (electron spectroscopic imaging; ESI). ESI/EELS can reveal selected elements with high sensitivity and spatial resolution, but only when sections are substantially thinner than the mean free path of inelastic collisions. This implies that, at 85–100 kV, only sections thinner than 25 nm can be profitably used. Although the use of ESI instrumentation with higher accelerating voltage should, in principle, allow the use of thicker sections, this approach has not been widely exploited yet. The need to use

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Figure 5 Ultrastructural localization of Ca2+ in hyphae of Neurospora. Electron micrograph of the apical region of Neurospora hyphae prepared by classical chemical fixation where the distribution of Ca2+ is showed by precipitation with the anion pyroantimoniate. Wall vesicles (v) have dense precipitates of calcium pyroantimoniate. Some deposits are also associated with the plasma membrane. Mitochondria (mit) do not contain deposits. Bar = 1 ␮m.

very thin specimens has represented a severe limitation and in particular, the use of cryosections proved problematical. Another aspect to be considered is sensitivity. Although in absolute terms the elemental detection is higher for ESI/EELS than for X-ray microanalysis, because of the required thinness of the analyzed specimens and the large contribution of the carbon peak, only relatively high calcium levels can be detected (Ottensmeyer and Andrew, 1980). Although the detection sensitivity of EELS has been analyzed in nonbiological standards (Shuman and Somlyo, 1987; Door et al., 1997), in cellular samples it has not been precisely established yet. In standards obtained by deposition or evaporation of calcium on different substrates, samples containing 100 mmol/liter and above calcium showed detectable calcium signals (Grohovaz et al., 1996).

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r The technique of ion microscopy, based on secondary ion mass spectrometry, is capable of detecting any element from H to U via isotopic detection based on its mass-to-charge ratio with detection limits in the parts-per-million to parts-per-billion range (Morrison and Slodzian, 1975; Linton and Goldsmith, 1992). Ion microscopy is capable of multielemental imaging, and the high sensitivity of the technique is very well suited for imaging of elemental distributions in hyphae. Unfortunately, this technique does not get down to electron microscopy resolution, and therefore it is often not useful for localizing calcium into identified subcellular compartments. However, it can provide information about the overall distribution of calcium and other ions. Uromyces germlings examined for total calcium distribution by ion microscopy showed the highest total calcium in the apical region, between the nuclei and the germling tip, whereas the apical 20 ␮m appeared to be slightly lower in total calcium (Chandra et al., 1999). B. Cytosolic Ca2+ Homeostasis in Fungi: Roles of Plasma Membrane Transport and Intracellular Sequestration of Ca2+ 1. Homeostasis of Ca2+ Ca2+ homeostasis is tightly controlled in all eukaryotic cells, since intracellular Ca2+ plays important roles in signal transduction and in the regulation of many proteins, and also because Ca2+ is cytotoxic at high levels. In eukaryotic cells in general, Ca2+ enters the cytoplasm passively (moving down its concentration gradient) either through Ca2+ channels in the PM or from Ca2+ -sequestering organelles such as the ER, mitochondria, and vacuoles. Cytosolic Ca2+ is maintained at low levels, either by pumping Ca2+ out of the cell through the PM or by sequestering them into organelles (Chattopadhyay et al., 1997; Williams, 1998). 2. Influx of Extracellular Calcium: Fungal Ca2+ Channels The origin of the cytoplasmic Ca2+ gradient in the fungal hyphae could be the entrance of Ca2+ from the extracellular medium through Ca2+ channels on the PM. Although there has been considerable research on channel location and clustering in animal neuronal cells (reviewed by Whatley and Harris, 1996), very few studies have been performed in fungi. Ca2+ stretch-activated channels have been identified and characterized by patch clamping in Saccharomyces (Gustin et al., 1986, 1988), Uromyces (Zhou et al., 1991), Saprolegnia (Garrill et al., 1992, 1993; Levina et al., 1994), and Neurospora (Levina et al., 1995) cells. The patch clamp method offers the ability to directly measure ion channel activity and to analyze ion channel distributions. The technique has been described in detail elsewhere (Sakmann and Neher, 1995), and its use in fungal species has been reviewed (Garrill and Davies,

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1994). Garrill et al. (1992, 1993) devised a method of preparing protoplasts that permitted the study of channels in protoplasts derived from different regions of the hyphae. Therefore, with the patch clamp technique, it is possible to map ion channel locations to determine whether they are asymmetrically located along the hyphae, play a role in asymmetric ion currents, and are part of the architectural requirements of tip growth (Garrill et al., 1992; Garrill and Davies, 1994; Levina et al., 1994, 1995). Intuitively, if Ca2+ channels were explicitly required for the maintenance of the Ca2+ gradient during the process of tip growth, one would expect them to be localized at the tip-growing region of the hypha (Fig. 2). Zhou et al. (1991) reported the presence of a cation-selective mechanosensitive ion channel in germinating urediospores of the rust fungus Uromyces, although questions have been raised about the source of the membrane being patch clamped (Garrill et al., 1992). Gadolinium inhibits channel activity and, in vivo, germ tube growth and differentiation, suggesting that the channel may function in sensing leaf topography. In the patch clamp experiments using Uromyces, the distribution of the channels along the germ tube was not examined and would certainly shed additional light on their possible physiological roles. Using patch clamp techniques, the ion channel locations have been characterized and mapped along the hyphae of the oomycete Saprolegnia (Garrill et al., 1992, 1993; Levina et al., 1994). Two distinct channel types are observed: stretchactivated channels that are permeable to Ca2+ and spontaneous channels that are K+ permeable. Along the hyphae, there are tip-high gradients of spontaneous K+ permeable channels and stretch-activated Ca2+ permeable channels. Interestingly, actin disruption with cytochalasin E causes both tip-high gradients to disappear, suggesting a role of actin in the positioning of the channels at the apex (Levina et al., 1994). Gadolinium inhibits the stretch-activated channels, inhibits growth, and abolishes a tip-high intracellular Ca2+ gradient observed during growth. These results suggest that, in Saprolegnia, tip-localized entrance of Ca2+ through the Ca2+ channels may cause the elevated cytosolic Ca2+ concentration at the apex, being the origin of the tip-high Ca2+ gradient necessary to grow. This is confirmed by using ion-selective vibrating probes, where a net Ca2+ influx is observed at the tip in the growing hyphae of Saprolegnia (Lew, 1999). This situation is similar to growing pollen tubes (Miller et al., 1992, Pierson et al., 1994) and root hairs (Schifelbein et al., 1992; Felle and Hepler, 1997; Wymer et al., 1997) where a tiplocalized Ca2+ influx through Ca2+ channels may also cause the elevated cytosolic Ca2+ concentration at the tip. Neurospora has two basic channel types similar to those found in Saprolegnia: stretch-activated channels permeable to Ca2+ and spontaneous K+ permeable channels (Levina et al., 1995). However, in Neurospora, there is no indication of a tip-focused gradient of either the K+ permeable channels or stretchactivated channels. The absence of a tip-high gradient of Ca2+ channels implies that they do not have a function in tip expansion. Indeed, the inhibition of the stretch-activated channels with gadolinium only transiently inhibits growth. In

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addition, the ion-selective vibrating probes technique has shown that there is not a net influx of Ca2+ at the tips of growing hyphae of Neurospora (Lew, 1999). Therefore, the tip-high Ca2+ gradient is not produced by the apical entrance of external Ca2+ through Ca2+ channels (Levina et al., 1995; Lew, 1999), as it happens in Saprolegnia. The Ca2+ gradient seems to be maintained by an internal mechanism involving internal stores and recycling of the ions, a process described as “bootstrapping” (Jackson and Heath, 1993). Wall vesicles could have a role in Ca2+ shuttling to the growing tip, since Ca2+ has been localized in the wall vesicles of Neurospora (Fig. 5). At the tip, wall vesicles fuse with the PM, thus depositing their contents into the periplasmic space. The localized release of Ca2+ at the apex may induce an area with high concentration of this ion in the apical periplasmic space; the Ca2+ could re-enter the cell here maintaining the cytoplasmic tip-high Ca2+ gradient (Fig. 4, see color insert). The differences between the water mold Saprolegnia and the terrestrial Neurospora could be a result of evolution in two very different environments. In air, the fungal hyphal tip does not have an assured supply of extracellular Ca2+ . As Neurospora lives in an environment with variable Ca2+ availability, the mechanism of Ca2+ “bootstrapping” described above is probably necessary to maintain the Ca2+ gradient. By contrast, evolution in an aquatic environment means that Ca2+ is more accessible to the growing tip. Hence, Saprolegnia would have no difficulty utilizing external ions and Ca2+ channels in the tip growth process. In this organism, Ca2+ enter the cell preferentially in the hyphal apex thanks to the channels that are present there, causing the elevated cytosolic Ca2+ concentration at the apex, necessary to grow. However, Saprolegnia can grow and maintain the Ca2+ gradient in Ca2+ -free medium for 12–24 h (Jackson and Heath, 1989). Ca2+ recycling, which functions in normal conditions in Neurospora, must be important in Saprolegnia in situations of Ca2+ depletion (Jackson and Heath, 1993). Internal recycling is unlikely to last indefinitely in this organism. This may explain why Saprolegnia grown in Ca2+ -free media cease growing after 12–24 h (Jackson and Heath, 1989). The slower growth of Saprolegnia hyphae in Ca2+ -free medium also suggests that the reutilization of internal Ca2+ may not be as effective as influx via PM channels (Jackson and Heath, 1989). The ability of some other tip-growing systems to grow in Ca2+ -free media (e.g., fucoid zygotes; Kropf, 1992) might also be attributable to recycling from internal stores. Actin seems to be involved in generating and maintaining the Ca2+ gradient in the hyphae irrespective of whether it is produced by the localized entrance of external Ca2+ or by Ca2+ recycling and Ca2+ shuttling in wall vesicles to the apex. The F-actin network is important for establishing and/or maintaining gradients of mechanosensitive Ca2+ channels at hyphal tips (see Section II.D.3) and for the vesicle transport system to the growing tips (see Section II.4.D). Thus, the actin cytoskeleton is likely to be responsible for the gradients by either model.

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3. Ca

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Ca2+ stores are likely to be important in setting up and maintaining the Ca2+ gradient by sequestering/releasing Ca2+ in a manner directly essential to tip growth. Sequestration of Ca2+ by organelles, such as ER (Miller et al., 1992), mitochondria (Yuan and Heath, 1991; Pitt and Barnes, 1993), or vacuoles (Allaway et al., 1997), has been hypothesized to play a role in generating the Ca2+ gradient. In Saprolegnia, the Ca2+ flows preferentially into the hyphal tip thanks to the Ca2+ channels that are abundant there. The organelles located behind the tip could remove Ca2+ from the cytoplasm, confining the zone of high concentration of Ca2+ to the tip itself. At present, information regarding the distribution of intracellular stores of Ca2+ in filamentous fungal cells is lacking, although vacuoles, ER, and/or mitochondria are good Candidates (Jackson and Heath, 1993; Allaway et al., 1997). UV irradiation of Saprolegnia hyphae elevated Ca2+ concentration in the cytoplasm, very likely by releasing Ca2+ from internal stores (Hyde and Heath, 1997). The increase in the Ca2+ concentration occurred about 10 ␮m behind the tip rather than at the very tip. This region of the hyphae is enriched in mitochondria (Heath and Kaminskyj, 1989), ER (Yuan and Heath, 1991), and tubular vacuoles (Allaway et al., 1997), all of which are likely Ca2+ stores (Jackson and Heath, 1993; Allaway et al., 1997). In animal cells, the ER has long been known as a major intracellular calcium store under the primary control of inositol triphosphate (IP3 ) (Streb et al., 1983; Somlyo et al., 1985). More recently, the Golgi apparatus was recognized as an active calcium store (Chandra et al., 1991; Zha et al., 1995). Mitochondria may also play a significant role in sequestering calcium, especially under conditions of stress to maintain Ca2+ concentrations below cytosolic toxic levels (Somlyo et al., 1979). The key question is whether there is any special regulatory system that controls the release of Ca2+ from internal stores in a manner directly essential for growth. The most likely Candidate for such a system is the phosphoinositide system since IP3 is effective in various contexts at releasing sequestered Ca2+ (Streb et al., 1983; Csutora et al., 1999). Phosphoinositides have been implicated in a variety of cellular processes, including Ca2+ regulation, actin rearrangement, vesicle trafficking, cell survival and mitogenesis in animal cells (Tolias and Cantley, 1999) and the occurrence of polyphosphoinositides in fungi is established (Bowman et al.,1987). IP3 releases Ca2+ from Neuropora vacuoles (Cornelius and Nakashima, 1987) but not from internal stores in either Saccharomyces (Csutora et al., 1999) or Penicillium (Pitt and Barnes, 1993). Lithium, which blocks the phosphoinositide cycle (Alberts et al., 1994; Nahorski et al., 1992; Fauroux and Freeman, 1999), decreases the rate of tip extension and increases branching in Neurospora (Hanson, 1991), suggesting that IP3 -mediated Ca2+ release, possibly from the vacuoles, may be important in the regulation of hyphal growth. However, it is no clear whether

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these effects are exerted directly on Ca release from sequestration or are more indirect. IP3 is essential for cell division in Saccharomyces (Uno et al., 1988) and possibly in other fungi. In addition, the activities of actin binding proteins are often modulated by phosphoinositides (Janmey, 1994; Schmidt and Hall, 1998) and therefore IP3 inhibitors may disturb the actin network, thus affecting fungal growth.

C. Roles of Ca2+ Gradients in Tip Growth 1. Ca2+ and Hyphal Extension One of the universal characteristics of all tip growing cells, including all hyphae critically examined, is the presence of a tip-high gradient of Ca2+ (Garrill et al., 1993; Hyde and Heath, 1997; Jackson and Heath, 1993; Levina et al., 1995; Silverman-Gavrila and Lew, 2000). This gradient is present in all growing tips, is dissipated in nongrowing tips (Garrill et al., 1993; Hyde and Heath, 1997; Levina et al., 1995; Silverman-Gavrila and Lew, 2000), and forms early during the initiation of branches (Hyde and Heath, 1997). Experimental evidence shows that the polarized distribution of Ca2+ is essential for hyphal growth. Several studies have reported morphological effects in fungi, specifically modifications of the apical extension and branching patterns in response to disturbances of external Ca2+ levels brought about either by withholding Ca2+ from media or by the use of Ca2+ ionophores (Reissig and Kinney, 1983; Harold and Harold, 1986; Schmid and Harold, 1988; McGillivray and Gow, 1987). In general, conditions of decreased Ca2+ influx caused by ion chelation, use of channel blockers, or lowering the concentration of exogenous Ca2+ lead to a reduction in hyphal extension rate and increased branching. However, high extracellular Ca2+ inhibits branching and induces hyphal elongation (reviewed by Jackson and Heath, 1993). These effects are thought to be related to the influence that exogenous Ca2+ has on the steepness of the intracellular gradient (Jackson and Heath, 1989; Hyde and Heath, 1997). In Neurospora, even when external Ca2+ and stretch-activated Ca2+ channels in the PM are not required for normal hyphal growth, the intracellular tip-high Ca2+ gradient is essential, since the dissipation of the Ca2+ gradient induced by BAPTA injection in Neurospora hyphae induces the cessation of growth (Silverman-Gavrila and Lew, 2000). Similar BAPTA effects were reported for tip-growing pollen tubes and root hairs (Pierson et al., 1994; Felle and Hepler, 1997). There is evidence supporting a primary role of Ca2+ in apical growth, such as some studies where imposed external gradients of Ca2+ or the Ca2+ -transporting ionophore A23187 orient the polarized extension of fucoid rhizoids (Robinson and Cone, 1980; Robinson and Jaffe, 1975), neuronal growth cones (Gundersen and Barrett, 1980), and hyphae of the oomycete Saprolegnia (Hyde and Heath, 1995). Besides, in the fucoid system, the site of rhizoidal

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emergence is predicted by the presence of elevated levels of Ca within the initially spherical zygote (Berger and Brownlee, 1993). Localized vesicular fusion is an undisputed essential common feature of all tip systems and Ca2+ is known to promote the fusion of exocytotic vesicles with the PM. More than 30 years ago, Douglas (Douglas and Rubin, 1961; Douglas, 1968) proposed that intracellular Ca2+ controls stimulus–secretion coupling in endocrine cells, and Katz and Miledi (1967; Katz, 1969) proposed that Ca2+ controls the rapid release of neurotransmitters from synapses. The high cytoplasmic concentration of Ca2+ subjacent to the apical membrane in filamentous fungi could promote the exocytosis of wall vesicles, thus localizing growth to the tip by ensuring that new cell wall and PM is deposited at the apex. The wall vesicles would perpetuate the growing tip by supplying new calcium channels and/or Ca2+ from internal stores (Fig. 2). Ca2+ -dependent modulation of cellular processes may occur via intracellular Ca2+ -binding proteins, of which calmodulin is one of the best characterized (Means, 1994). Several cellular processes in filamentous fungi have been shown to be Ca2+ -calmodulin dependent, such as conidiation in Penicillium (Pitt and Barnes, 1993), cell cycle control in Aspergillus (Rasmussen et al., 1994; Nanthakumar et al., 1996; Dayton and Means, 1996), dimorphism in Candida (Paranjape et al., 1990) and in Ophiostoma (Brunton and Gadd, 1991), lipase formation in Fusarium (Hoshino et al., 1991), secretion of xylanases in Trichoderma (Mach et al., 1998), appressorium formation of Magnaporthe (Lee and Lee, 1998), and polarized hyphal growth in Neurospora (Ortega Perez and Turian, 1987). In this fungus, calmodulin is known to activate chitin synthase (Suresh and Subramanyam, 1997) and interacts with actin (Capelli et al., 1997). In addition, inhibitors of calmodulin increase the frequency of branching and slow tip growth in Neurospora (Ortega Perez and Turian, 1987). Calmodulin, in its Ca2+ activated form, is able to activate or inactivate a number of enzymes, including protein kinases and at least one phosphatase called calcineurin (Klee et al., 1988). The evidence shows that protein phosphorylation by kinases potentiates Ca2+ -dependent exocytosis in mammalian cells (Hille et al., 1999), and may be the physiological trigger of exocytosis in fungi. Kinases and phosphatases in fungi are likely to be important mediators of fungal proliferation and development as well as signal transduction and infection-related morphogenesis (reviewed by Dickman and Yarden, 1999). Calcineurin is a Ca2+ - and calmodulin-dependent protein that is likely to be involved in hyphal growth. Calcineurin is a heterodimeric protein composed of a catalytic subunit (calcineurin A), and a regulatory subunit (calcineurin B). Activation of calcineurin occurs when calcineurin B binds Ca2+ , and calmodulin binds to a region within calcineurin A (Aitken et al., 1984; Stemmer and Klee, 1994). Calcineurin is present in fungi, and several works suggest that this phosphatase could participate in the establishment of the Ca2+ gradient, regulating hyphal growth and morphology. The catalytic subunit calcineurin A, encoded by can-1,

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is essential for growth of Neurospora (Prokisch et al., 1997) and is located at high concentrations at the tip (Kincaid, 1993). Decreased expression of can-1 caused growth arrest, preceded by an increase in hyphal branching, changes in hyphal morphology, and concomitant loss of the tip-high Ca2+ gradient (CTCmembrane) (Prokisch et al., 1997). The same effects are observed after treatment of Neurospora with the calcineurin inhibitors cyclosporin A or FK506 (Prokisch et al., 1997). Exogenous Ca2+ cannot restore the Ca2+ gradient in hyphae with inactivated CAN1, indicating that the loss of the gradient may be due to a failure to maintain the gradient rather than changes in the avilability of exogenous Ca2+ . These results indicate that ion transport across the PM of the growing tip of Neurospora is not essential for growth as discussed in Section III.B.3. Calcineurin B is also required for normal vegetative growth in Neurospora and is related to conidiation (Kothe and Free, 1998). Mutants affected in the gene encoding a calcineurin B homolog lose their ability to repress entry into macroconidiation and are unable to generate vegetative hyphae, producing chains of septated cells. This suggests that the regulated functioning of calcineurin is required for vegetative hyphal growth and plays a role in defining the morphology and developmental state of Neurospora. Calcineurin has also been implicated in controlling the morphology of Schizosaccharomyces (Yoshida et al., 1994) and nerve cells (Chang et al., 1995). 2. Ca2+ and Branching Increase of cytoplasmic Ca2+ by different methods induces branches in tip-growing cells, suggesting a role of Ca2+ in branching (Reissig and Kinney, 1985; Grinberg and Heath, 1997; Silverman-Gavrila and Lew, 2000). In studies of the cytological events that take place during hyphal branching, it has been shown that a momentary cytoplasmic contraction, which produces the retraction of the Spitzenk¨orper, precedes the initiation of a branch (Reynaga-Pe˜na et al., 1996; Reynaga-Pe˜na and Bartnicki-Garcia, 1997). Ca2+ -dependent cytoplasmic contractions have been described in hyphae and can be induced by UV irradiation (McKerracher and Heath, 1986b; Jackson and Heath, 1992) or ionophores (Kaminskyj et al., 1992b). In Saprolegnia, UV irradiation increases cytoplasmic Ca2+ concentration and induces the formation of one or more branches near the irradiation site (Grinberg and Heath, 1997). The sequence of events following UV irradiation, that is, cessation of growth after 2–2.5 min accompanied by a change in hyphal morphology and followed by resumption of growth, is almost identical to that observed preceding branch initiation (Hyde and Heath, 1997; Reynaga-Pe˜na and Bartnicki-Garcia, 1997). In both cases, cytoplasmic contractions have been observed, further suggesting that the contractions that precede branch initiation may be due to a Ca2+ signal. A model of Ca2+ regulation of branch formation is shown in Fig. 3. A gradient of Ca2+ forms as a branch develops, but not prior to branch appearance (Hyde and

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Heath, 1997), indicating that either an unknown initiator or possibly subdetectable Ca2+ concentrations are required to initiate the branch (Hyde and Heath, 1997). A transient release of sequestered Ca2+ from subapical cytoplasmic organelles or the entry of external Ca2+ through Ca2+ channels in the subapical PM may induce an elevation of cytoplasmic Ca2+ , but at a concentration too low to be detected by current methods. One effect of this increase of Ca2+ concentration may be the localized changes in actin organization that have been detected prior to branching in tip growing cells. Bachewich and Heath (1998) demonstrated that radial actin arrays form in Saprolegnia in regions with no yet-detectable surface protrusion. One of actin’s functions in initiating branching is probably directing the transport of wall vesicles to the points of new branch formation, and thus delivering mechanosensitive Ca2+ channels or Ca2+ from internal stores. It is known that the F-actin network is important for establishing and/or maintaining gradients of Ca2+ channels at hyphal tips (Section II.D.3) and the vesicle transport system utilizes actin in fungi and other tip-growing cells (Section II.D.4). This could induce an increase in the concentration of Ca2+ in the branch initials, detected by fluorescence methods (Hyde and Heath, 1997). The actin cap characteristic of tips replaces the radial arrays in the new branches. Colocalization of all these components (Ca2+ , mechanosensitive channels, actin) at the new apex would lead to an accelerating, self-sustaining, growing hyphal tip. 3. Ca2+ Regulation of the Actin Network Actin cytoskeleton assembly is regulated by Ca2+ at multiple levels and variations in the Ca2+ concentration translate into changes in mechanical status of the actin network (Nakamura and Kohama, 1999; Richelme et al., 2000). Ca2+ can bind to actin directly and affect monomer conformation (Bertazzon and Tsong, 1990; Miki, 1990) or modulate the activities of a large number of ABPs (Janmey, 1994; Schmidt and Hall, 1998). Many of these proteins regulate actin assembly by controlling filament formation and cross-linking of the actin network (Pollard and Cooper, 1986; St¨ossel, 1993; Schafer and Cooper, 1995; Sun et al., 1995; Welch et al., 1997). Since the polymerization of actin filaments is controlled by Ca2+ , the main function of Ca2+ may be the regulation of the structure and rigidity of the actin network at the apex. Regalado (1998) proposed a mathematical model where the Ca2+ status at the tip may be responsible for the apical aggregation of wall vesicles, based on the Ca2+ regulation of the mechanical properties (elasticity and viscosity) of the cytoskeletal network. In this model, the viscoelastic properties of the cytoskeleton are strictly controlled by the Ca2+ concentration, giving importance to the Ca2+ gradient for hyphal growth in contradistinction to the mere presence or absence of Ca2+ . In the absence of a gradient, there would be a homogeneous distribution of vesicles and absence of growth, whereas steeper gradients would favor polarized growth. The relationship between growth rates and both apical concentration and the steepness of its gradient has been experimentally demonstrated in

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Saprolegnia (Hyde and Heath, 1997). However, the model of Regalado (1998) does not consider the extensive known direct actin–myosin and actin–ABP interactions or describe the molecules involved in the postulated cytoplasm properties. The internal concentration of Ca2+ also initiates cytoskeletal contractions (Gollnick et al., 1991), which may be explained in terms of “gel–sol” transitions of the cytoskeleton as a consequence of variations in Ca2+ concentration (Taylor and Condeelis, 1979). Cytoplasmic contractions have been seen in fungi (see Section II.D.6) and are thought to be responsible for the tipward movement of the cytoplasm in growing hyphae (Jackson and Heath, 1992), allowing the cytoplasm to keep pace with the growth of the hyphal tip (McKerracher and Heath, 1986b, 1987). Cytoplasmic contractions have been also related to the initiation of branching (see Sections II.D.2/III.C.2).

IV. Conclusions Initiation and maintenance of tip growth are processes regulated by the actin cytoskeleton and by Ca2+ . A membrane skeleton similar to that in animal cells, minimally containing F-actin, spectrin, and integrin, seems to play a crucial role in the tip growth of hyphae. This skeleton restrains or protrudes the hyphal tips under the differing turgor conditions. Other functions of the actin cytoskeleton include cytoplasmic and organelle motility, vesicle transport and exocytosis, and localization of elements required for tip growth, such as Ca2+ channels at the hyphal apices. Ca2+ may influence the extensibility of the membrane skeleton and regulates exocytosis of wall vesicles at the apex, necessary for tip extension. The tip-focused gradient of cytosolic Ca2+ is a general feature of growing fungal hyphae, the concentration of which is influenced by actin, irrespective of whether it is produced by the localized entrance of external Ca2+ or by Ca2+ recycling and shuttling in wall vesicles to the apex, suggesting a possible feedback regulation mechanism.

Acknowledgment The preparation of this work was supported by gratefully acknowledged grants from NSERC.

References Abe, S., and Davies, E. (1995) Methods for isolation and analysis of the cytoskeleton. Methods Cell Biol. 50, 223–236. Adams, A. E., and Pringle, J. R. (1984). Relationship of actin and tubulin distribution to bud growth in wild-type and morphogenetic-mutant Saccharomyces cerevisiae. J. Cell Biol. 98, 934– 945.

5. Actin and Ca2+ in Hyphal Tip Growth

171

Aist, J. R. (1995). Independent nuclear motility and hyphal tip growth. Can. J. Bot. 73 (Suppl. 1), S122–S125. Aist, J. R., and Bayles, C. J. (1991). Ultrastructural basis of mitosis in the fungus Nectria hematococca (sexual stage of Fusarium solani). I. Asters. Protoplasma 161, 111–122. Aist, J. R., and Berns, M. W. (1981). Mechanics of chromosome separation during mitosis in Fusarium (Fungi imperfecti): new evidence from ultrastructural and laser microbeam experiments. J. Cell Biol. 91, 446–458. Aitken, A., Klee, C. B., Cohen, P. (1984). The structure of the B subunit of calcineurin. Eur. J. Biochem. 139, 663–671. Akashi, T., Kanbe, T., and Tanaka, K. (1994). The role of the cytoskeleton in polarized growth of the germ tube in Candida albicans. Microbiology 140, 271–280. Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K., and Watson, J. D. (1994). “Molecular Biology of the Cell,” 3rd ed. Garland Publishing Inc., New York and London. Allaway, W. G., Ashford, A. E., Heath, I. B., and Hardham., A. R. (1997). Vacuolar reticulum in oomycete hyphal tips: an additional component of the Ca2+ regulatory system? Fung. Genet. Biol. 22, 209–220. Allen, E. D., Aiuto, R., and Susman, A. S. (1980). Effects of cytochalasins on Neurospora crassa. I. Growth and ultrastructure. Protoplasma 102, 63–75. Anderson, J. M., and Soll, D. R. (1986). Differences in actin localization during bud and hypha formation in the yeast Candida albicans. J. Gen. Microbiol. 132, 2035–2047. Ayscough, K. R., Stryker, J., Pokala, N., Sanders, M., Crews, P., and Drubin, D. G. (1997). High rates of actin filament turnover in budding yeast and roles for actin in establishment and maintenance of cell polarity revealed using the actin inhibitor latrunculin-A. J. Cell Biol. 137, 399–416. Bachewich, C. L., and Heath, I. B. (1997). The cytoplasmic pH influences hyphal tip growth and cytoskeleton-related organization. Fung. Genet. Biol. 21, 76–91. Bachewich, C. L., and Heath, I. B. (1998). Radial F-actin arrays precede new hypha formation in Saprolegnia: Implications for establishing polar growth and regulating tip morphogenesis. J. Cell Sci. 111, 2005–2016. Bachewich, C. L., and Heath, I. B. (1999). Cytoplasmic migrations and vacuolation are associated with growth recovery in hyphae of Saprolegnia, and are dependent on the cytoskeleton. Mycol. Res. 103, 849–858. Bahler, J., and Pringle, J. R. (1998) Pom1p, a fission yeast protein kinase that provides positional information for both polarized growth and cytokinesis. Genes Dev. 12, 1356–1370. Bartnicki-Garcia, S. (1990). Role of vesicles in apical growth and a new mathematical model of hyphal morphogenesis. In: “Tip Growth in Plant and Fungal Cells” (I. B. Heath, Ed.), pp. 211–232. Academic Press, San Diego. Bartnicki-Garcia, S., and Lippman, E. (1972). The bursting tendency of hyphal tips of fungi: Presumptive evidence for a delicate balance between wall synthesis and wall lysis in apical growth. J. Gen. Microbiol. 73, 487–500. Bartnicki-Garcia, S., Hergert, F., and Gierz, G. (1989). Computer simulation of fungal morphogenesis and the mathematical basis of hyphal (tip) growth. Protoplasma 153, 46–57. Bartnicki-Garcia, S., Bartnicki, D. D., and Gierz, G. (1995). Determinants of fungal cell wall morphology: The vesicle supply center. Can. J. Bot. 73 (Suppl. 1), S372–S378. Bendayan, M. (1984). Enzyme-gold electron microscopic cytochemistry: A new affinity approach for the ultrastructural localization of macromolecules. J. Electron. Microsc. Tech. 1, 349–372. Bendel, C. M., and Hostetter, M. K. (1993). Distinct mechanisms of epithelial adhesion for Candida albicans and Candida tropicalis: Identification of the participating ligands and development of inhibitory peptides. J. Clin. Invest. 92, 1850–1849. Berger, F., and Brownlee, C. (1993). Ratio confocal imaging of free cytoplasmic calcium gradients in polarising and polarised Fucus zygotes. Zygote 1, 9–15. Berger, K. H., Sogo, L. F., and Yaffe, M. P. (1997). Mdm12p, a component required for mitochondrial inheritance that is conserved between budding and fission yeast. J. Cell Biol. 136, 545–553.

172

Sara Torralba and I. Brent Heath

Berridge, M., Lipp, P., and Bootman, M. (1999). Calcium signalling. Curr. Biol. 9, R157–R159. Bertazzon, A., and Tsong, T. Y. (1990). Study of effects of pH on the stability of domains in myosin rod by high-resolution differential scanning calorimetry. Biochemistry 29, 6455–6461. Bhattacharya, D., Medlin, L., Wainwright, P., Ariztia, E., Bibeau, C., Stickel, S., and Sogin, M. (1992). Algae containing chlorophylls a 1 c are paraphyletic: Molecular evolutionary analysis of the chromophyta. Evolution 46, 1801–1817. Bibikova, T. N., Zhigilei, A., and Gilroy, S. (1997). Root hair growth in Arabidopsis thaliana is directed by calcium and an endogenous polarity. Planta 203, 495–505. Bibikova, T. N., Blancaflor, E. B., and Gilroy, S. (1999). Microtubules regulate tip growth and orientation in root hairs of Arabidopsis thaliana. Plant J. 7, 657–65. Blanco, A. M., and Villela, L. C. (1998). Ultrastructural localization of calcium in peripheral nerve fibres undergoing Wallerian degeneration: an oxalate–pyroantimoniate and X-ray microanalysis study. J. Submicrosc. Cytol. Pathol. 30, 451–458. Blinks, J. R., Wier, W. G., Hess, P., and Prendergast, F. G. (1982). Measurement of Ca2+ concentrations in living cells. Prog. Biophys. Mol. Biol. 40, 1–114. Boldogh, I., Vojtov, N., Karmon, S., and Pon, L. A. (1998). Interaction between mitochondria and the actin cytoskeleton in budding yeast requires two integral mitochondrial outer membrane proteins, Mmm1p and Mdm10p. J. Cell Biol. 141, 1371–1381. Botstein, D., Chervitz, S. A., and Cherry, J. M. (1997). Yeast as a model organism. Science 277, 1259–1260. Bourett, T. M., and Howard, R. J. (1991). Ultrastructural immunolocalization of actin in a fungus. Protoplasma 163, 199–202. Bowman, B. J., Borgeson, C. E., and Bowman, E. J. (1987). Composition of Neurospora crassa vacuolar membranes and comparison to endoplasmic reticulum, plasma membranes and mitochondrial membranes. Exp. Mycol. 11, 197–205. Bracker, C. E., Murphy, D. J., and Lopez-Franco, R. (1997). Laser microbeam manipulation of cell morphogenesis in growing fungal hyphae. SPIE Int. Sym. Proc. Photonics West’97 2983, 67–80. Braun, M., Baluska, F., von Witsch, M., and Menzel, D. (1999). Redistribution of actin, profilin and phosphatidylinositol-4,5-bisphosphate in growing and maturing root hairs. Planta 209, 435– 443. Brawley, S. H., and Robinson, K. R. (1985). Cytochalasin treatment disrupts the endogenous currents associated with cell polarization in fucoid zygotes: studies of the role of F-actin in embryogenesis. J. Cell Biol. 100, 1173–1184. Bray, D. (1992). “Cell Movements.” Garland Publishing, Inc., New York and London. Bretscher, A. (1993). Microfilaments and membranes. Curr. Opin. Cell Biol. 5, 653–60. Brown, S. S. (1999). Cooperation between microtubule- and actin-based motor proteins. Annu. Rev. Cell Dev. Biol. 15, 63–80. Brownlee, C., and Bouget, F. Y. (1998). Polarity determination in Fucus: from zygote to multicellular embryo. Semin. Cell Dev. Biol. 9, 179–185. Brownlee, C., and Wood, J. W. (1986). A gradient of cytoplasmic free calcium in growing rizhoid cells of Fucus serratus. Nature (London) 320, 624–626. Bruno, K. S., Aramayo, R., Minke, P. F., Metzenberg, R. L., and Plamann, M. (1996). Loss of growth polarity and mislocalization of septa in a Neurospora mutant altered in the regulatory subunit of cAMP-dependent protein kinase. EMBO J. 15, 5772–5782. Brunton, A. H., and Gadd, G. M. (1991). Evidence for an inositol lipid signal pathway in the yeast– mycelium transition of Ophiostoma ulmi, the Dutch elm disease fungus. Mycol. Res. 95, 484– 491. Butt, T. M., and Heath, I. B. (1988). The changing distribution of actin and nuclear behavior during the cell cycle of the mite-pathogenic fungus Neozygites sp. Eur. J. Cell Biol. 46, 499–505. Caesar Ton That, C. T., Rossier, C., Barja, F., Turian, G., and Roos, U. P. (1988). Induction of multiple germ tubes in Neurospora crassa by antitubulin agents. Eur. J. Cell Biol. 46, 68–79.

5. Actin and Ca2+ in Hyphal Tip Growth

173

Capelli, N., Barja, F., van Tuinen, D., Monnat, J., Turian, G., Ortega Perez, R. (1997). Purification of a 47-kDa calmodulin-binding polypeptide as an actin-binding protein from Neurospora crassa. FEMS Microbiol. Lett. 147, 215–20. Caswell, A. H. (1979). Methods of measuring intracellular calcium. Int. Rev. Cytol. 56, 145–181. Chan, S. Y., Ochs, S., and Jersild, R. A., Jr. (1984). Localization of calcium in nerve fibers. J. Neurobiol. 15, 89–108. Chandra, S., Kable, E. P. W., Morrison, G. H., and Webb, W. W. (1991). Calcium sequestration in the Golgi apparatus of cultured mammalian cells revealed by laser scanning confocal microscopy and ion microscopy. J. Cell Sci. 100, 747–752. Chandra, S., Leinhos, G. M., Morrison, G. H., and Hoch HC. (1999). Imaging of total calcium in urediospore germlings of Uromyces by ion microscopy. Fung. Genet. Biol. 27, 77–87. Chang, H. Y., Takei, K., Sydor, A. M., Born, T., Rusnak, F., and Jay, D. G. (1995). Asymmetric retraction of growth cone filopodia following focal inactivation of calcineurin. Nature 376, 686–690. Chang, P. L. Y., and Trevithick, J. R. (1974). How important is secretion of exoenzymes through apical cell walls of fungi. Arch. Microbiol. 101, 281–293. Chattopadhyay, N., Vassilev, P. M., and Brown, E. M. (1997). Calcium-sensing receptor: roles in and beyond systemic calcium homeostasis. Biol. Chem. 378, 759–768. Chiu, Y. H., Xiang, X., Dawe, A. L., and Morris, N. R. (1997). Deletion of nudC, a nuclear migration gene of Aspergillus nidulans, causes morphological and cell wall abnormalities and is lethal. Mol. Biol. Cell. 8, 1735–1749. Chowdhury, S., Smith, K. W., and Gustin, M. C. (1992). Osmotic stress and the yeast cytoskeleton: phenotype-specific suppression of an actin mutation. J. Cell Biol. 118, 561–571. Clarkson, D. T., Brownlee, C., and Ayling, S. M. (1988). Cytoplasmic calcium measurements in intact higher plant cells: results from fluorescence ratio imaging of Fura-2. J. Cell. Sci. 91, 71–80. Clemons, G. P., and Sisler, H. D. (1969). Formation of a fungitoxic derivate from Benlate. Phytopathology 59, 705–706. Connor, J. A. (1986). Digital imaging of free calcium changes and of spatial gradients in growing processes in single mammalian central nervous system cells. Proc. Natl. Acad. Sci. USA 83, 6179– 6183. Cornelius, G., and Nakashima, H. (1987). Vacuoles play a decisive role in calcium homeostasis in Neurospora crassa. J. Gen. Microbiol. 133, 2341–2347. Corrˆea, A., and Hoch, H. C. (1993). Microinjection of urediospore germlings of Uromyces appendiculatus. Exp. Mycol. 17, 241–252. Corrˆea, A., Staples, R. C., and Hoch, H. C. (1996). Inhibition of thigmostimulated cell differentiation with RGD-peptides in Uromyces germlings. Protoplasma 194, 91–102. Csutora, P., Su, Z., Kim, H. Y., Bugrim, A., Cunningham, K. W., Nuccitelli, R., Keizer, J. E., Hanley, M. R., Blalock, J. E., and Marchase, R. B. (1999). Calcium influx factor is synthesized by yeast and mammalian cells depleted of organellar calcium stores. Proc. Natl. Acad. Sci. USA 96, 121–126. Czymmek, K. J., Bourettt, T. M., and Howrad, R. J. (1995). Immunolocalization of tubulin and actin in thick-sectioned fungal hyphae after freeze-substitution fixation and methacrilate de-embedment. J. Microsc. 181, 153–161. Davidse, L. C., and Flach, W. (1978). Interaction of thiabendazole with fungal tubulin. Biochim. Biophys. Acta 543, 82–90. Dayton, J. S., and Means, A. R. (1996). Ca2+ /calmodulin-dependent kinase is essential for both growth and nuclear division in Aspergillus nidulans. Mol. Biol. Cell. 7, 1511–1519. De Lucas, J. R., Monistrol, I. F., and Laborda, F. (1993). Effect of antimicrotubular drugs on the secretion process of extracellular proteins in Aspergillus nidulans. Mycol. Res. 97, 961–966. Degouse´e N., Gupta, G. D., Lew, R. R., and Heath, I. B. (2000). A putative spectrin-containing membrane skeleton in hyphal tips of Neurospora crassa. Fung. Genet. Biol. 30, 33–44. Dickman, M. B., and Yarden O. (1999). Serine/threonine protein kinases and phosphatases in filamentous fungi. Fung. Genet. Biol. 26, 99–117.

174

Sara Torralba and I. Brent Heath

Doonan, B. B., Crang, R. E., Jensen, T. E., and Baxter, M. (1979). In situ X-ray microanalysis of polyphosphate bodies in Aureobasidium pullulans. J. Ultrastruct. Res 69, 232–238. Door, R., Breitig, D., and Martin, R. (1997). Videodensitometric analysis of electron spectroscopic micrographs, a tool for detection of biologically relevant elements with high resolution. J. Microsc. 187, 170–183. Douglas, W. W. (1968). Stimulus–secretion coupling: the concept and clues from chromaffin and other cells. Brit. J. Pharmacol. 34, 453–474 Douglas, W. W., and Rubin, R. P. (1961). The role of calcium in the secondary response of the adrenal medulla to acetylcholine. J. Physiol. 159, 40–47. Doyle, T., and Botstein, D. (1996). Movement of yeast cortical actin cytoskeleton visualized in vivo. Proc. Natl. Acad. Sci. USA 93, 3886–3891. Drubin, D., and Nelson, W. (1996). Origins of cell polarity. Cell. 84, 335–344. Enos, A. P., and Morris, N. R. (1990). Mutation of a gene that encodes a kinesin-like protein blocks nuclear division in Aspergillus nidulans. Cell 60, 1019–1027. Eshel, D., Urrestarazu, L. A., Vissers, S., Jauniaux, J. C., Van Vliet-Reedijk, J. C., Planta, R. J., and Gibbons, I. R. (1993). Cytoplasmic dynein is required for normal nuclear segregation in yeast. Proc. Natl. Acad. Sci. USA 90, 11172–11176. Faraday, C. D., and Spanswick, R. M. (1993). Evidence for a membrane skeleton in higher plants: a spectrin-like polypeptide co-isolates with rice root plasma membranes. FEBS Lett. 318, 313–316. Fath, K. R., Trimbur, G. M., and Burgess, D. R. (1994). Molecular motors are differentially distributed on Golgi membranes from polarized epithelial cells. J. Cell Biol. 126, 661–675. Fauroux, C. M., and Freeman, S. (1999). Inhibitors of inositol monophosphatase. J. Enzym. Inhib. 14, 97–108. Felle, H. H., and Hepler, P. K. (1997). The cytosolic Ca2+ concentration gradient of Sinapis alba root hairs as revealed by Ca2+ selective microelectrode tests and fura-dextran ratio imaging. Plant Physiol. 114, 39–45. Ferro-Novick, S., and Jahn, R. (1994). Vesicle fusion from yeast to man. Nature 370, 191–193. Finger, F., and Novick, P. (1998). Spatial regulation of exocytosis: lessons from yeast. J. Cell Biol. 142, 609–612. Fischer, R. (1999). Nuclear movement in filamentous fungi. FEMS Microbiol. Rev. 23, 39–68. Fisk, H. A., and Yaffe, M. P. (1997). Mutational analysis of Mdm1p function in nuclear and mitochondrial inheritance. J. Cell Biol. 138, 485–494. F¨orster, H., and Mendgen, K. (1987). Immunocytochemical localization of pectinesterases in hyphae of Phytophthora infestans. Can. J. Bot. 65, 2607–2613. Gabriel, M., and Kopecka, M. (1995). Disruption of the actin cytoskeleton in budding yeast results in formation of an aberrant cell wall. Microbiology 141, 891–899. Gadd, G. M. (1994). Signal transduction in fungi. In “The Growing Fungus” (N. A. R. Gow and G. M. Gadd, Eds.), Chapman and Hall, London. Gale, C., Finkel, D., Tao, N., Meinke, M., McClellan, M., Olson, J., Kendrick, K., and Hostetter, M. (1996). Cloning and expression of a gene encoding an integrin-like protein in Candida albicans. Proc. Natl. Acad. Sci. USA 93, 357–361. Gale C. A., Bendel C. M., McClellan M., Hauser M., Becker J. M., Berman J., and Hostetter M. K. (1998). Linkage of adhesion, filamentous growth, and virulence in Candida albicans to a single gene, INT1. Science 279, 1355–1358. Galpin, M. F. J., Jennings, D. H., Oates, K., and Hobot, J. A. (1978). Localization by X-ray microanalysis of soluble ions, particularly potassium and sodium, in fungal hyphae. Exp. Mycol. 2, 258–269. Garrill, A., and Davies, J. M. (1994). Patch clamping fungal membranes: A new perspective on ion transport. Mycol. Res. 98, 256–263. Garrill, A., Lew, R. R., and Heath, I. B. (1992). Stretch-activated Ca2+ and Ca2+ -activated K+ channels in the hyphal tip plasma membrane of the oomycete Saprolegnia ferax. J. Cell Sci. 101, 721– 730.

5. Actin and Ca2+ in Hyphal Tip Growth

175

Garrill, A., Jackson, S. L., Lew, R. R., and Heath, I. B. (1993). Ion channel activity and tip growth, Tip-localized stretch-activated channels generate an essential Ca2+ gradient in the oomycete, Saprolegnia ferax. Eur. J. Cell Biol. 60, 358–365. Gavin, R. H. (1997). Microtubule-microfilament synergy in the cytoskeleton. Int. Rev. Cytol. 173, 207–242. Geli, M. I., and Riezman, H. (1996). Role of type I myosins in receptor-mediated endocytosis in yeast. Science 272, 533–535. Girbardt, M. (1957). Der Spitzenk¨orper von Polystictus versicolor (L). Planta 50, 47–59. Girbardt, M. (1969). Die Ultrastruktur der Apikalregion von Pilzhyphen. Protoplasma 67, 413–441. Girbardt, M. (1979). A microfilamentous septal belt (FSB) during induction of cytokinesis in Trametes versicolor (L. ex. Fr.). Exp. Mycol. 3, 215–228. Gollnick, F., Meyer, R., and Stockem, W. (1991). Visualization and measurement of calcium transients in Amoeba proteus by fura-2 fluorescence. Eur. J. Cell Biol. 55, 262–271. Gomperts, B. D. (1986). Calcium shares the limelight in stimulus-secretion coupling. Trends Biol. Sci. 11, 290–292. Gooday, G. W. (1983). The hyphal tip. In “A contemporary Synthesis” (L. E. Smith Ed.), pp. 315–356. Dekker, New York. Goodson, H. V., and Spudich, J. A. (1993). Molecular evolution of the myosin family: relationships derived from comparisons of amino acid sequences. Proc. Natl. Acad. Sci. USA 90, 659– 663. Goodson, H. V., and Spudich, J. A. (1995). Identification and molecular characterization of a yeast myosin I. Cell Motil. Cytoskele. 30, 73–84. Goodson, H. V., Anderson, B. L., Warrick, H. M., Pon, L. A., and Spudich J. A. (1996). Synthetic lethality screen identifies a novel yeast myosin I gene (MYO5): myosin I proteins are required for polarization of the actin cytoskeleton. J. Cell Biol. 133, 1277–1291. Gottlieb, T. A., Ivanov, I. E., Adesnik, M., and Sabatini, D. D. (1993). Actin microfilaments play a critical role in endocytosis at the apical but not the basolateral surface of polarized epithelial cells. J. Cell Biol. 120, 695–710. Govindan, B., Bowser, R., and Novick, P. (1995). The role of Myo2, a yeast class V myosin, in vesicular transport. J. Cell Biol. 128, 1055–1068. Gow, N. A. R. (1995). Tip growth and polarity. In “The Growing Fungus.” (N. A. R. Gow, and G. M. Gadd, Eds.), pp. 277–299. Chapman and Hall, London. Grinberg, A., and Heath, I. B. (1997). Direct evidence for Ca2+ regulation of hyphal branch induction. Fung. Genet. Biol. 22, 127–139. Grohovaz, F., Bossi, M., Pezzati, R., Meldolesi, J., and Tarelli, F. T. (1996). High resolution ultrastructural mapping of total calcium: electron spectroscopic imaging/electron energy loss spectroscopy analysis of a physically/chemically processed nerve-muscle preparation. Proc. Natl. Acad. Sci. USA 93, 4799–4803. Grove, S. N., and Bracker, C. E. (1970). Protoplasmic organization of hyphal tips among fungi: vesicles and Spitzenk¨orper. J. Bacteriol. 104, 989–1009. Grove, S. N., and Sweigard, J. A. (1980). Cytochalasin A inhibits spore germination and hyphal tip growth in Gilbertella periscaria. Exp. Mycol. 4, 239–250. Gundersen, R. W., and Barrett, J. N. (1980). Characterization of the turning response of dorsal root neurites toward nerve growth factor. J. Cell Biol. 87, 546–554. Gupta, G., and Heath, I. B. (1997). Actin disruption by latrunculin B causes turgor-related changes in tip growth of Saprolegnia hyphae. Fung. Genet. Biol. 21, 64–75. Gupta, G. D., and Heath, I. B. (2000). A tip-high gradient of a qutative plasma membrane SNARE approximates the exacytotic gradient in hyphal apices of the fungus Neurospora crassa. Fung. Genet. Biol. 29, 187–199. Gustin, M. C., Martinac, B., Saimi, Y., Culbertson, M. R., and Kung, C. (1986). Ion channels in yeast. Science 233, 1195–1197.

176

Sara Torralba and I. Brent Heath

Gustin, M. C., Zhou, X. L., Martinac, B., and Kung, C. (1988). A mechanosensitive ion channel in yeast plasma membrane. Science 242, 762–765. Hackney, D. D. (1995). Motor proteins. Polar explorations. Nature 376, 215–216. Hanson, B. A. (1991). The effects of lithium on phosphoinositides and inositol phosphates of Neurospora crassa. Exp. Mycol. 15, 76–90. Hardham, R. H., and Mitchell, H. J. (1998). Use of molecular cytology to study the structure and biology of phytopathogenic and mycorrhizal fungi. Fung. Genet. Biol. 24, 252–284. Harold, F. M. (1997). How hyphae grow: morphogenesis explained? Protoplasma 197, 137–147. Harold, R. L., and Harold, F. M. (1986). Ionophores and cytochalasins modulate branching in Achlya bisexualis. J. Gen. Microbiol. 132, 213–219. Harold, R. L., and Harold, F. M. (1992). Configuration of actin microfilaments during sporangium development in Achlya bisexualis: comparison of two staining protocols. Protoplasma 171, 110– 116. Harold, F. M., Harold, R. L., and Money, N. P. (1995). What forces drive cell wall expansion? Can. J. Bot. 73 (Suppl. 1), S379–S383. Harris, N. (1994). Immunocytochemistry for Light and Electron Microscopy. In “Plant Cell Biology: A Practical Approach” (N. Harris, and K. J. Oparka, Eds.), pp. 157–176. IRL Press, Oxford Univ. Press, Oxford. Harris, S. D., Morrell, J. L., and Hamer, J. E. (1994). Identification and characterization of Aspergillus nidulans mutants defective in cytokinesis. Genetics 136, 517–532. Harris, S. D., Hofmann, A. F., Tedford, H. W., and Lee, M. P. (1997). Identification and characterization of genes required for hyphal morphogenesis in the filamentous fungus Aspergillus nidulans. Genetics 151, 1015–1025. Harris, S. D., Hamer, L., Sharpless, K. E., and Hamer, J. E. (1999). The Aspergillus nidulans sepA gene encodes an FH1/2 protein involved in cytokinesis and the maintenance of cellular polarity. EMBO J. 16, 3474–3483. Hartwig, J. H., and Kwiatkowski, D. J. (1991). Actin-binding proteins. Curr. Opin. Cell Biol. 3, 87–97. Harvey, D. M. R. (1982). Freeze-substitution. J. Microsc. 127, 209–221. Harvey, D. M. R., Hall, J. R., and Flowers, T. J. (1976). The use of freeze-substitution in the preparation of plant tissues for ion localization studies. J. Microsc. 107, 189. Hasek, J., and Bartnicki-Garcia, S. (1994). The arrangement of F-actin and microtubules during germination of Mucor rouxii. Arch. Microbiol. 161, 363–369. Haugland, R. P. (1996). “Handbook of Fluorescent Probes and Research Chemicals.” Molecular Probes, Eugene. Heath, I. B. (1982). The effect of nocodazole on the growth and ultrastructure of the fungus Saprolegnia ferax. Evidence against a simple mode of action. In “Microtubules in Microrganisms” (P. Capuccinelli and N.R. Morris, Eds.), pp. 275–311. Dekker, New York/Basel. Heath, I. B. (1987). Preservation of a labile cortical array of actin filaments in growing hyphal tips of the fungus Saprolegnia ferax. Eur. J. Cell Biol. 44, 10–16. Heath, I. B. (1990). The roles of actin in tip growth of fungi. Int. Rev. Cytol. 123, 95–127. Heath, I. B. (1994). The cytoskeleton in hyphal growth, organelle movements, and mitosis. In “The Mycota I Growth, Differentiation and Sexuality” (J. G. H. Wessels, and F. Meinhardt, Eds.), pp. 43–65. Springer Verlag, Berlin. Heath, I. B. (1995). Integration and regulation of hyphal tip growth. Can. J. Bot. 73(Suppl. 1), S131– S139. Heath, I. B., and Heath, M. C. (1978). Microtubules and organelle movements in the rust fungus Uromyces phaseoli var. vignae. Cytobiologie 16, 393–411. Heath, I. B., and Kaminskyj, S. G. W. (1989). The organization of tip-growth-related organelles and microtubules revealed by quantitative analysis of freeze-substituted o¨omycete hyphae J. Cell Sci. 93, 41–52.

5. Actin and Ca2+ in Hyphal Tip Growth

177

Heath, I. B., and Steinberg, G. (1999). Mechanisms of hyphal tip growth: tube dwelling amebae revisited. Fung. Genet. Biol. 28, 79–93. Heath, I. B., and van Rensburg, E. J. (1996). Critical evaluation of the VSC model for tip growth. Mycoscience 37, 1–10. Heath I. B., Rethoret, K., Arsenault, A. L. and Ottensmeyer, F. P. (1985). Improved preservation of the form and the content of wall vesicles and the Golgi apparatus in freeze substituted hyphae of Saprolegnia. Protoplasma 128, 81–93. Henningsen, U., and Schliwa, M. (1997). Reversal in the direction of movement of a molecular motor. Nature 389, 93–96. Henry, C. A., Jordon, J. R., and Kropf, D. L. (1996). Localized membrane-wall adhesions in Pelvetia zygotes. Protoplasma, 190, 39–52. Hepler, P. K., and Wayne, R. O. (1985). Calcium and plant development. Annu. Rev. Plant Physiol. 36, 397–439. Hermann, G. J., King, E. J., and Shaw, J. M. (1997). The yeast gene, MDM20, is necessary for mitochondrial inheritance and organization of the actin cytoskeleton. J. Cell Biol. 137, 141–153. Herr, F. B., and Heath, M. C. (1982). The effects of antimicrotubule agents on organelle positioning in the cowpea rust fungus, Uromyces phaseoli var. vignae. Exp. Mycol. 6, 15–24. Hill, T. W., and Mullins, J. T. (1980). Hyphal tip growth in Achyla. Can. J. Microbiol. 26, 1132–1140. Hill, K. L. Catlett, N. L., and Weisman, L. S. (1996). Actin and myosin function in directed vacuole movement during cell division in Saccharomyces cerevisiae. J. Cell Biol. 135, 1535–1549. Hille, B., Billiard, J., Babcock, D. F., Nguyen, T., and Koh, D. S. (1999). Stimulation of exocytosis without a calcium signal. J. Physiol. (Lond). 520, 23–31. Hoch, H. C., and Howard, R. J. (1980). Ultrastructure of freeze-substituted hyphae of the basidiomycete Laetisaria arvalis. Protoplasma 103, 281–297. Hoch, H. C., and Staples, R. C. (1983). Ultrastructural organization of the non-differentiated uredospore germling of Uromyces phaseoli. Mycologia 75, 795–824. Hoch, H. C., and Staples, R. C. (1985). The microtubule cytoskeleton in hyphae of Uromyces phaseoli germlings: Its relationship to the region of nucleation and to the F-actin cytoskeleton. Protoplasma 124, 112–122. Hoch, H. C., Staples, R. C., Whitehead, B., Comeau, J., and Wolf, E. D. (1987). Signaling for growth orientation and cell differentiation by surface topography in Uromyces. Science 235, 1659–1662. Hodson, M. J. (1995). Ion localization and X-ray microanalysis. Methods Cell Biol. 49, 21–31. Hodson, M. J., and Sangster, A. G. (1989). Subcellular localization of mineral deposits in the roots of wheat (Triticum aestivum L.). Protoplasma 151, 19–32. Hoffmann, J., and Mendgen, K. (1998). Endocytosis and membrane turnover in the germ tube of Uromyces fabae. Fung. Genet. Biol. 24, 77–85. Hoshino, T., Mizutani, A., Shimizu, S., Hidaka, H., and Yamane, T. (1991). Calcium ion regulates the release of lipase of Fusarium oxysporum. J. Biochem. (Tokyo) 110, 457–461. Howard, R. J. (1981). Ultrastructural analysis of hyphal tip cell growth in fungi: Spitzenk¨orper, cytoskeleton and endomembranes after freeze-substitution. J. Cell Sci. 48, 89–103. Howard, R. J., and Aist, J. R. (1977). Effects of MBC on hyphal tip organization, growth, and mitosis of Fusarium acuminatum. Protoplasma. 92, 195–210. Howard, R. J., and Aist, J. R. (1979). Hyphal tip cell ultrastructure of the fungus Fusarium: improved preservation by freeze-substitution. J. Ultrastruct. Res. 66, 224–234. Howard, R. J., and Aist, J. R. (1980). Cytoplasmic microtubules and fungal morphogenesis: ultrastructural effects of methyl benzimidazole-2-yl-carbamate determinated by freeze-substitution of hyphal tip cells. J. Cell Biol. 87, 55–64. Huffaker, T. C., Thomas, J. H., and Botstein, D. (1988). Diverse effects of tubulin mutations on microtubule formation and function. J. Cell Biol. 106, 1997–2010. Hyde, G. J. (1998). Calcium imaging: A primer for mycologists. Fung. Genet. Biol. 24, 14–23.

178

Sara Torralba and I. Brent Heath

Hyde, G. J., and Hardham, A. R. (1992). Confocal microscopy of microtubule arrays in cryosectioned sporangia of Phytophthora cinnamomi. Exp. Mycol. 16, 207–218. Hyde, G. J., and Heath, I. B. (1995). Ca2+ -dependent polarization of axis establishment in the tipgrowing organism, Saprolegnia ferax, by gradients of the ionophore A23187. Eur. J. Cell Biol. 67, 356–362. Hyde, G. J., and Heath, I. B. (1997). Ca2+ gradients in hyphae and branches of Saprolegnia ferax. Fung. Genet. Biol. 21, 238–251. Hyde, G. J., Davies, D., Perasso, L., Cole, L., and Ashford, A. E. (1999). Microtubules, but not actin microfilaments, regulate vacuole motility and morphology in hyphae of Pisolithus tinctorius. Cell Motil. Cytoskeleton 42, 114–124. Hynes, R. O. (1992). Integrins: versatility, modulation, and signaling in cell adhesion. Cell 69, 11–25. Ito, Y., Abe, S., and Davies, E. (1994). Co-localization of cytoskeleton proteins and polysomes with a membrane fraction from peas. J. Exp. Botany 45, 253–259. Jackson, S. L., and Hardham, A. R. (1996). A transient rise in cytoplasmic free calcium is required to induce cytokinesis in zoosporangia of Phytophthora cinnamomi. Eur. J. Cell Biol. 69, 180–188. Jackson, S. L., and Heath, I. B. (1989). Effects of exogenous calcium ions on tip growth, intracellular Ca2+ concentration, and actin arrays in hyphae of the fungus Saprolegnia ferax. Exp. Mycol. 13, 1–12. Jackson, S. L., and Heath, I. B. (1990a). Evidence that actin reinforces the extensible hyphal apex of the oomycete Saprolegnia ferax. Protoplasma 157, 144–153. Jackson, S. L., and Heath, I. B. (1990b). Visualization of actin arrays in growing hyphae of the fungus Saprolegnia ferax. Protoplasma 154, 66–70. Jackson, S. L., and Heath, I. B. (1992). UV microirradiations elicit Ca2+ dependent apex-directed cytoplasmic contractions in hyphae. Protoplasma 170, 46–52. Jackson, S. L., and Heath, I. B. (1993). Roles of calcium ions in hyphal tip growth. Microbiol. Rev. 57, 367–382. Janmey, P. A. (1994). Phosphoinositides and calcium as regulators of cellular actin assembly and disassembly. Annu. Rev. Physiol. 56, 169–191. Jochov`a, J., Rupeˇs, I., and Peberdy, J.F. (1993). Effect of the microtubule inhibitor benomyl on protein secretion in Aspergillus nidulans. Mycol. Res. 97, 22–27. Johnston, G. C., Prendergast, J. A., and Singer, R. A. (1991). The Saccharomyces cerevisiae MYO2 gene encodes an essential myosin for vectorial transport of vesicles. J. Cell Biol. 113, 539– 551. Jorgensen, A. O., and McGuffee, L. J. (1987). Immunoelectron microscopic localization of sarcoplasmic reticulum proteins in cryofixed, freeze-dried, and low temperature-embedded tissue. J. Histochem. Cytochem. 35, 723–732. Juliano, R. L., and Haskill, S. (1993). Signal transduction from the extracellular matrix. J. Cell Biol. 120, 577–585 Jung, M. K., May, G. S., and Oakley, B. R. (1998). Mitosis in wild-type and beta-tubulin mutant strains of Aspergillus nidulans. Fung. Genet. Biol. 24, 146–160. Kagawa, T., Kadota, A., and Wada, M. (1992). The junction between the plasma membrane and the cell wall in fern protonemal cells, as visualized after plasmolysis, and its dependence on arrays of cortical microtubules. Protoplasma 170, 186–190. Kaminskyj, S. G., and Hamer J. E. (1998). Hyp loci control cell pattern formation in the vegetative mycelium of Aspergillus nidulans. Genetics 148, 669–680. Kaminskyj, S. G. W., and Heath, I. B. (1994). A comparison of techniques for localizing actin and tubulin in hyphae of Saprolegnia ferax . J. Histochem. Cytochem. 42, 523–530. Kaminskyj, S. W. G., and Heath, I. B. (1995). Integrin and spectrin homologues, and cytoplasm–wall adhesion in tip growth. J. Cell Sci. 108, 849–856. Kaminskyj, S. G. W., and Heath, I. B. (1996). Studies on Saprolegnia ferax suggest the general importance of the cytoplasm in determining hyphal morphology. Mycologia 88, 20–37.

5. Actin and Ca2+ in Hyphal Tip Growth

179

Kaminskyj, S. W. G., Garrill, A., and Heath, I. B. (1992a). The relation between turgor and tip growth in Saprolegnia ferax, Turgor is necessary, but not sufficient to explain apical extension rates. Exp. Mycol. 16, 64–75. Kaminskyj, S. W. G., Yoon, K. S., and Heath, I. B. (1992b). Fixation induces differential polarized translocations of organelles in hyphae of Saprolegnia ferax. J. Microsc. 167, 153–168. Katz, B. (1969). In “The Release of Neural Transmitter Substances” (Charles Thomas, Ed.). Springfield, IL. Katz, B., and Miledi, R. (1967). The timing of calcium action during neuromuscular transmission. J. Physiol. 189, 535–544. Kauss, H. (1987). Some aspects of calcium-dependent regulation in plant metabolism. Ann. Rev. Plant Physiol. 38, 47–72. Kincaid, R. (1993). Calmodulin-dependent protein phosphatases from microorganisms to man. In “Advances in Second Messengers and Phosphoprotein Research” Vol. 10, (S. Shenolikar and A. C. Nairn Eds.), pp. 1–23. Raven Press, New York. Klee, C. B., Draetta, G. F., and Hubbard, M. J. (1988). Calcineurin. Adv. Enzymol. 61, 149–200. Knight, H. (2000). Calcium signaling during abiotic stress in plants. Int. Rev. Cytol. 195, 269–324. Knight, M. R., Campbell, A. K., Smith, S. M., and Trewavas, A. J. (1991). Transgenic plant aequorin reports the effects of touch and cold-shock and elicitors on cytoplasmic calcium. Nature 352, 524–526. Knight, H., Trewavas, A. J., and Read, N. D. (1993). Confocal microscopy of living fungal hyphae microinjected with Ca2+ -sensitive fluorescent dyes. Mycol. Res. 97, 1505–1515. Konig, K., So, P. T., Mantulin, W. W., Tromberg, B. J., and Gratton, E. (1996). Two-photon excited lifetime imaging of autofluorescence in cells during UVA and NIR photostress. J. Microsc. 183, 197–204. Kothe, G. O., and Free, S. J. (1998). The isolation and characterization of nrc-1 and nrc-2, two genes encoding protein kinases that control growth and development in Neurospora crassa. Genetics 149, 117–130. Kropf, D. L. (1992). Establishment and expression of cellular polarity in fucoid zygotes. Microbiol. Rev. 56, 316–339. Kropf, D. L. (1994). Cytoskeletal control of cell polarity in a plant zygote. Dev. Biol. 165, 361–371. Kropf, D.L. (1997). Induction of polarity in fucoid zygotes. Plant Cell 9, 1011–1020. Kropf, D. L., Kloareg, B., and Quatrano, R. S. (1988). Cell wall is required for fixation of the embryonic axis in Fucus zygotes. Science 239, 187–190. Kropf, D. L., Bisgrove, S. R., and Hable, W. E (1998). Cytoskeletal control of polar growth in plant cells. Current Opin. Cell Biol. 10, 117–122. Lancelle, S. A., and Hepler, P. K. (1991). Association of actin with cortical microtubules revealed by immunogold localization in Nicotiana pollen tubes. Protoplasma 165, 167–172. Langford, G. M. (1995). Actin- and microtubule-dependent organelle motors: interrelationships between the two motility systems. Curr. Opin. Cell Biol. 7, 82–88. Lattanzio, F. A., Jr., and Bartschat, D. K. (1991). The effect of pH on rate constants, ion selectivity and thermodynamic properties of fluorescent calcium and magnesium indicators. Biochem. Biophys. Res. Commun. 177, 184–191. Lee, S. C., and Lee, Y. H. (1998). Calcium/calmodulin-dependent signaling for appressorium formation in the plant pathogenic fungus Magnaporthe grisea. Mol. Cells. 8, 698–704. Lehmler, C., Steinberg, G., Snetselaar, K. M., Schliwa, M., Kahmann, R., and Boelker, M. (1997). Identification of a motor protein required for filamentous growth in Ustilago maydis. EMBO J. 16, 3464–3473. Levina, N. N., Lew, R. R., and Heath, I. B. (1994). Cytoskeletal regulation of ion channel distribution in the tip-growing organism Saprolegnia ferax. J. Cell Sci. 107, 127–134. Levina, N. N., Lew, R. R., Hyde, G. J., and Heath, I. B. (1995). The roles of Ca2+ and plasma membrane ion channels in hyphal tip growth of Neurospora crassa. J. Cell Sci. 108, 3405–3417.

180

Sara Torralba and I. Brent Heath Ca2+

H+

Lew, R.R. (1999). Comparative analysis of and flux magnitude and location along growing hyphae of Saprolegnia ferax and Neurospora crassa. Eur. J. Cell Biol. 78, 892–902. Lew, D. J., and Reed, S. I. (1995). Cell cycle control of morphogenesis in budding yeast. Curr. Opin. Genet. Dev. 5, 17–23. Linton, R. W., and Goldsmith, J. G. (1992). The role of secondary ion mass spectrometry (SIMS) in biological microanalysis. Technique comparisons and prospects. Biol. Cell 74, 147–160. L´opez-Franco, R., and Bracker, C. E. (1996). Diversity and dynamics of the Spitzenk¨orper in growing hyphal tips of higher fungi. Protoplasma 195, 90–111. Lord, E. M., and Sanders, L. C. (1992). Roles for the extracellular matrix in plant development and pollination: a special case of cell movement in plants. Dev. Biol. 153, 16–28. Luna, E. J., and Hitt, A. L. (1992). Cytoskeleton-plasma membrane interactions. Science 258, 955–964. Mach, R. L., Zeilinger, S., Kristufek, D., and Kubicek, C. P. (1998). Ca2+ -calmodulin antagonists interfere with xylanase formation and secretion in Trichoderma reesei. Biochim. Biophys. Acta. 1403, 281–289. Malh´o, R., Read, N. D., Pais, M. S., and Trewavas, A. J. (1994). Role of cytosolic free calcium in the orientation of pollen tube growth. Plant J. 5, 331–341. Marcantonio, E. E., and Hynes, R. O. (1988). Antibodies to the conserved cytoplasmic domain of the integrin beta1-subunit react with proteins in vertebrates, invertebrates and fungi. J. Cell Biol. 106, 1765–1772. Marks, J., and Hyams, J. S. (1985). Localization of F-actin through the cell division cycle of Schizosaccharomyces pombe. Eur. J. Cell Biol. 39, 27–32. Mata, J., and Nurse, P. (1997). Tea1 and the microtubule cytoskeleton are important for generating global spatial order within the fission yeast cell. Cell 89, 939–949. Mattson, M. P. (1999). Establishment and plasticity of neuronal polarity. J. Neurosci Res. 57, 577– 589. May, K., Wheatly, S., Amin, V., and Hyams, J. (1998). The myosin ATPase inhibitor 2,3-butanedione2-monoxime (BDM) inhibits tip growth and cytokinesis in the fission yeast, Schizosaccharomyces pombe. Cell Motil. Cytoskel. 41, 117–125. May, K. M., and Hyams, J. S. (1998). The yeast cytoskeleton: the closer we look, the more we see. Fung. Genet. Biol. 24, 110–122. McClure, W. K., Park, D., and Robinson, P. M. (1968). Apical organization in the somatic hyphae of fungi. J. Gen. Microbiol. 50, 177–182. McConnell, S. J., and Yaffe, M. P. (1992). Nuclear and mitochondrial inheritance in yeast depends on novel cytoplasmic structures defined by the MDM1 protein. J. Cell Biol. 118, 385–395. McGillviray, A. M., and Gow, N. A. R. (1987). The transhyphal electrical current of Neurospora crassa is carried principally by protons. J. Gen. Microbiol. 133, 2875–2881. McGoldrick, C. A., Gruver, C., and May, G. S. (1995). myoA of Aspergillus nidulans encodes an essential myosin I required for secretion and polarized growth. J. Cell Biol. 128, 577–587. McKerracher, L. J., and Heath, I. B. (1986). Polarized cytoplasmic movement and inhibition of saltations induced by calcium-mediated effects of microbeams in fungal hyphae. Cell Motil. Cytoskel. 6, 136– 145. McKerracher, L. J., and Heath, I. B. (1987). Cytoplasmic migration and intracellular organelle movements during tip growth of fungal hyphae. Exp. Mycol. 11, 79–100. Means, A. R. (1994). Calcium, calmodulin and cell cycle regulation. FEBS Lett. 347, 1–4. Mermall, V., Post, P. L., and Mooseker, M. S. (1998). Unconventional myosins in cell movement, membrane traffic and signal transduction. Science 279, 527–533. Miki, M. (1990). Resonance energy transfer between points in a reconstituted skeletal muscle thin filament: a conformational change of the thin filament in response to a change in calcium concentration. Eur. J. Biochem. 187, 155–162. Miller, D. D., Callaham, D. A., Gross, D. J., and Hepler, P. K. (1992). Free Ca2+ gradient in growing pollen tubes of Lilium. J. Cell Sci. 101, 7–12.

5. Actin and Ca2+ in Hyphal Tip Growth

181

Momany, M., Westfall, P. J., and Abramowsky, G. (1999). Aspergillus nidulans swo mutants show defects in polarity establishment, polarity maintenance and hyphal morphogenesis. Genetics 151, 557–567. Money, N. P. (1995) Turgor pressure and the mechanics of fungal penetration. Can. J. Bot. 73(Suppl. 1), S96–S102. Money, N. P. (1997). Wishful thinking of turgor revisited: the mechanics of fungal growth. Fung. Genet. Biol. 21, 173–187. Money, N. (1998). Why oomycetes have not stopped being fungi. Mycol. Res. 102, 767–768. Money, N. P., and Harold, F. M. (1992). Extension growth of the water mold Achlya: interplay of turgor and wall strength. Proc. Natl. Acad. Sci. USA 89, 4245–4249. Money, N. P., and Harold, F. M. (1993). Two water molds can grow without measurable turgor pressure. Planta 190, 426–430. Morgan, A. J., Winters, C., and Sturzenbaum, S. (1999). X-ray microanalysis techniques. Methods Mol. Biol. 117, 245–276. Morris, N. R., and Enos, A. P. (1992). Mitotic gold in a mold: Aspergillus genetics and the biology of mitosis. Trends Genet. 8, 32–37. Morris, N. R., Xiang, X., and Beckwith, S. M. (1995). Nuclear migration advances in fungi. Trends. Cell Biol. 5, 278–282. Morrison, G. H., and Slodzian, G. (1975). Ion microscopy. Anal. Chem. 47, 932A–943A. Moukha, S. M., W¨osten, H. A. B., Asther, M., and Wessels, J. G. H. (1993). In situ localization of the secretion of lignin peroxidases in colonies of Phanerochaete chrysosporium using a sandwiched mode of cultive. J. Gen. Microbiol. 139, 969–978. Mulholland, J., Preuss, D., Moon, A., Wong, A., Drubin, D., and Botstein, D. (1994). Ultrastructure of the yeast actin cytoskeleton and its association with the plasma membrane. J. Cell Biol. 125, 381–391. Mulholland, J., Wesp, A., Riezman, H., and Botstein, D. (1997). Yeast actin cytoskeleton mutants accumulate a new class of Golgi-derived secretary vesicle. Mol. Biol. Cell 8, 1481– 1499. Mullins, J. T. (1979). A freeze-fracture study of hormone-induced branching in the fungus Achlya. Tissue Cell. 11, 585–595. Nahorski, S. R., Jenkinson, S., and Challiss, R. A. (1992). Disruption of phosphoinositide signalling by lithium. Biochem. Soc. Trans. 20, 430–434. Nakamura, A., and Kohama, K. (1999). Calcium regulation of the actin-myosin interaction of Physarum polycephalum. Int. Rev. Cytol. 191, 53–98. Nanthakumar, N. N., Dayton, J. S., and Means, A. R. (1996). Role of Ca2+ /calmodulin binding proteins in Aspergillus nidulans cell cycle regulation. Prog. Cell. Cycle Res. 2, 217–228. Nobes, C. D., and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81, 53–62. Novick, P., and Botstein, D. (1985). Phenotypic analysis of temperature-sensitive yeast actin mutants. Cell 40, 405–416. Novick, P., and Sheckman, R. (1983). Export of major cell surface proteins is blocked in yeast secretory mutants. J. Cell Biol. 96, 541–547. Oakley, B. R., and Morris, N. R. (1980). Nuclear movement is tubulin dependent in Aspergillus nidulans. Cell 19, 255–262. Oakley, B. R., and Rinehart, J. E. (1985). Mitochondria and nuclei move by different mechanisms in Aspergillus nidulans. J. Cell Biol. 101, 2392–2397. Oliver, P. T. P. (1973). Influence of cytochalasin B on hyphal morphogenesis of Aspergillus nidulans. Protoplasma 76, 279–281. Olmsted, J. B. (1986). Microtubule-associated proteins. Annu. Rev. Cell Biol. 2, 421–457. Olmsted, J. B. (1991). Non-motor microtubule-associated proteins. Curr. Opin. Cell Biol. 3, 52–58.

182

Sara Torralba and I. Brent Heath

Ortega Perez, R., and Turian., G. (1987). Cytomorphological defects produced by anti-calmodulin agents in outgrowing germ tubes and elongating hyphae of Neurospora crassa. Cytobios 49, 137– 145. Osmani, A. H., Osmani, S. A., and Morris, N. R. (1990). The molecular cloning and identification of a gene product specifically required for nuclear movement in Aspergillus nidulans. J. Cell Biol. 111, 543–551. Ottensmeyer, F. P., and Andrew, J. W. (1980). High resolution microanalysis of biological specimens by electron energy loss spectroscopy and by electron spectroscopy imaging. J. Ultrastruct. Res. 72, 336–348. Palmer, R. E., Sullivan, D. S., Huffaker T., and Koshland, D. (1992). Role of astral microtubules and actin in spindle orientation and migration in the budding yeast, Saccharomyces cerevisiae. J. Cell Biol. 119, 583–593. Paranjape, V., Roy, B. G., and Datta, A. (1990). Involvement of calcium, calmodulin and protein phosphorylation in morphogenesis of Candida albicans. J. Gen. Microbiol. 136, 2149–2154. Parton, R. M., and Read, N. D. (1998). Calcium and pH imaging in living cells. In “Light Microscopy in Biology—A Practical Approach” (A.J. Lacey, Ed.). IRL Press, Oxford. Pedregosa, A. M., R´ıos, S., Monistrol, I. F., and Laborda, F. (1995). Effect of the microtubule inhibitor methyl benzimidazol-2-yl carbamate (MBC) on protein secretion and microtubule distribution in Cladosporium cucumerinum. Mycol. Res. 99, 43–48. Peterbauer, C. K., Heidenreich, E., Baker, R. T., and Kubicek, C. P. (1992). Effect of benomyl and benomyl resistance on cellulase formation by Trichoderma reesei and Trichoderma harzianum. Can. J. Microbiol. 38, 1292–1297. Pezzati, R., and Grohovaz, F. (1999). The frog neuromuscular junction revisited after quick-freezingfreeze-drying: ultrastructure, immunogold labelling and high resolution calcium mapping. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 354, 373–378. Picton, J. M., and Steer, M. W. (1982). A model for the mechanism of tip extension in pollen tubes. J. Theor. Biol. 98, 15–20. Pierson, E. S., Miller, D. D., Callaham, D. A., Shipley, A. M., Rivers, B. A., Cresti, M., and Hepler, P. K. (1994). Pollen tube growth is coupled to the extracellular calcium ion flux and the intracellular calcium gradient—Effect of BAPTA-type buffers and hypertonic media. Plant Cell 2, 1815–1828. Pitt, D., and Barnes, J. C. (1993). Calcium homeostasis, signalling and protein phosphorylation during calcium-induced conidiation in Penicillium notatum. J. Gen. Microbiol. 139, 3053–3063. Pitt, D., and Ugalde, U. O. (1984). Calcium in fungi. Plant. Cell Envir. 7, 467–475 Plamann, M., Minke, P. E., Tinsley, J. H., and Bruno, K. S. (1994). Cytoplasmic dynein and actinrelated protein Arp1 are required for normal nuclear distribution in filamentous fungi. J. Cell. Biol. 127, 139–149. Pollard, T. D., and Cooper, J. A. (1986). Actin and actin-binding proteins. A critical evaluation of mechanisms and functions. Annu. Rev. Biochem. 55, 987–1035. Probst, W. (1986). Ultrastructural localization of calcium in the CNS of vertebrates. Histochemistry 85, 231–239. Prokisch, H., Yarden, O., Mischler, M., Tropschug, M., and Barthelmess, I. B. (1997). Impairing calcineurin of Neurospora crassa reveals its essential role for hyphal growth, morphology and maintenance of the apical Ca2+ -gradient. Mol. Gen. Genet. 256, 104–114. Pruyne, D. W., Schott, D. H., and Bretscher, A. (1998). Tropomyosin-containing actin cables direct the Myo2p-dependent polarized delivery of secretory vesicles in budding yeast. J. Cell Biol. 143, 1931–45. Quader, H., and Schnepf, E. (1989). Actin filament array during side branch initiation in protoneme cells of the moss Funaria hygrometrica: an actin organizing center at the plasma membrane. Protoplasma 151, 167–170. Quatrano, R. S., Brian, L., Aldridge, J., and Schultz, T. (1991). Polar axis fixation in Fucus zygotes: components of the cytoskeleton and extracellular matrix. Development Suppl. 1, 11–16.

5. Actin and Ca2+ in Hyphal Tip Growth

183

Rasmussen, C., Garen, C., Brining, S., Kincaid, R. L., Means, R. L., and Means, A. R. (1994). The calmodulin-dependent protein phosphatase catalytic subunit (calcineurin A) is an essential gene in Aspergillus nidulans. EMBO J. 15, 3917–3924. Raudaskoski, M., Rupeˇs, I., and Timonen, S. (1991). Inmunofluorescence microscopy of the cytoskeleton in filamentous fungi after quick-freezing and low-temperature fixation. Exp. Mycol. 15, 167–173. Raudaskoski, M., Mao, W., and Yli-Mattila, T. (1994). Microtubule cytoskeleton in hyphal growth. Response to nocodazol in a sensitive and a tolerant strain of the homobasidiomycete Schizophillum commune. Eur. J. Cell Biol. 64, 131–141. Read, N. D., Allan, W. T. G., Knight, H., Knight, M. R., Malh´o, R., Russell, A., Shacklock, P. S., and Trewavas, A. J. (1992). Imaging and measurement of cytosolic free calcium in plants and fungal cells. J. Microsc. (Oxford) 166, 57–86. Read, N. D., Fischer, S., and Parton, R. M. (1998). Imaging Sptizenk¨orper, pH and calcium dynamics in growing fungal hyphae. Pestic. Sci. 54, 179–181. Regalado, C. M. (1998). Roles of calcium gradients in hyphal tip growth: a mathematical model. Microbiology 144, 2771–2782. Regalado, C. M., and Sleeman, B. D. (1999). Aggregation and collapse in a mechanical model of fungal tip growth. J. Math. Biol. 39, 109–38. Reinhardt, M. O. (1892). Das Wachsthum der Pilzhyphen. Jahrb. Wiss. Bot. 23, 479–566. Reiss, H. D., and Herth, W. (1979). Calcium gradients in tip growing plant cells visualized by CTCfluorescence. Planta 146, 615–621. Reissig, J. L., and Kinney, S. G. (1983). Calcium as a branching signal in Neurospora crassa. J. Bacteriol. 154, 1397–1402. Reynaga-Pe˜na, C. G., and Bartnicki-Garcia, S. (1997). Apical branching in a temperature sensitive mutant of Aspergillus niger. Fung. Genet. Biol. 22, 153–167. Reynaga-Pe˜na, C. G., L´opez-Franco, R., and Bartnicki-Garcia, S. (1996). Cytoplasmic contractions in fungal hyphae and their morphogenetic consequences. Inoculum 47, 27. Richelme, F., Benoliel, A. M., and Bongrand, P. (2000). Dynamic study of cell mechanical and structural responses to rapid changes of calcium level. Cell Motil. Cytoskeleton 45, 93–105. Riquelme, M., Reynaga-Pe˜na, C. G., Gierz, G., and Bartnicki-Garcia, S. (1998). What determines growth direction in fungal hyphae? Fung. Genet. Biol. 24, 101–109. Roberson, R. W. (1992). The actin cytoskeleton in hyphal cells of Sclerotium rolfsii. Mycologia 84, 41–51. Roberson, R. W., and Fuller, M. S. (1988). Ultrastructural aspects of the hyphal tip of Sclerotium rolfsii preserved by freeze substitution. Protoplasma 146, 143–149. Robertson, N. F. (1965). Presidential address: the fungal hypha. Trans. Brit. Mycol. Soc. 48, 1–8. Robinson, K. R., and Cone, R. (1980). Polarization of fucoid eggs drive a calcium gradient ionophore gradient. Science 207, 77–78. Robinson, K. R., and Jaffe, L. F. (1975). Polarizing fucoid eggs drive a calcium current through themselves. Science 187, 70–72. Roemer, T., Madden, K., Chang, J., and Snyder, M. (1996). Selection of axial growth sites in yeast requires Axl2p, a novel plasma membrane glycoprotein. Genes Dev. 10, 777–793. Runeberg, P., Raudaskoski, M., and Virtanen, I. (1986). Cytoskeletal elements in the hyphae of the homobasidiomycete Schizophyllum commune visualized with indirect immunofluorescence and NBD-phallacidin. Eur. J. Cell Biol. 41, 25–32. Rupeˇs, I., Mao, W., Astrom, H., and Raudaskoski, M. (1995). Effects of nocodazole and brefeldin A on microtubule cytoskeleton and membrane organization in the homobasidiomycete Schizophyllum commune. Protoplasma 185, 212–221. Rutten, T. L. M., and Derkson, J. (1990). Organization of actin filaments in regenerating and outgrowing subprotoplasts from pollen tubes of Nicotiana tabacum L. Planta 180, 471–479 Sakmann, B., and Neher, E. (1995). “Single-Channel recording,” 2nd ed. Plenum, New York/London. Salo, V., Niini, S. S., Virtanen, I., and Raudaskoski, M. (1989). Comparative immunocytochemistry

184

Sara Torralba and I. Brent Heath

of the cytoskeleton of filamentous fungi with dikaryotic and multinucleate hyphae. J. Cell Sci. 94, 11–24. Sastry, S. K., and Horwitz, A. F. (1993). Integrin cytoplasmic domains: mediators of cytoskeletal linkages and extra- and intracellular initiated transmembrane signalling. Curr. Opin. Cell Biol. 5, 819–831. Schafer, D. A., and Cooper J. A. (1995). Control of actin assembly at filament ends. Annu. Rev. Cell Dev. Biol. 11, 497–518. Schifelbein, J. W., Shipley, A., and Rowse, P. (1992). Calcium influx at the tip of growing root-hair cells of Arabidopsis thaliana. Planta 187, 455–459. Schindler, M., Meiners, S., and Cheresh, D. A. (1989). RGD-dependent linkage between plant cell wall and plasma membrane: consequences for growth J. Cell Biol. 108, 1955–1965. Schmid, J., and Harold, F. M. (1988). Dual roles for calcium ions in apical growth of Neurospora crassa. J. Gen. Microbiol. 134, 2623–2631. Schmidt, A., and Hall, M. N. (1998). Signaling to the actin cytoskeleton. Annu. Rev. Cell Dev. Biol. 14, 305–338. Seiler, S., Nargang, F. E., Steinberg, G., and Schliwa, M. (1997). Kinesin is essential for cell morphogenesis and polarized secretion in Neurospora crassa. EMBO J. 16, 3025–3034. Shepherd, V. A., Orlovich, D. A., and Ashford, A. E. (1993). A dynamic continuum of pleiomorphic tubules and vacuoles in growing hyphae of a fungus. J. Cell Sci. 104, 495–507. Shuman, H., and Somlyo, A. P. (1987). Electron energy loss analysis of near-trace-element concentrations of calcium. Ultramicroscopy 21, 23–32. Silverman-Gavrila, L. B., and Lew, R. R. (2000). Calcium and tip growth in Neurospora crassa. Protoplasma, in press. Simon, V. R., Swayne, T. C., and Pon, L. A. (1995). Actin-dependent mitochondrial motility in mitotic yeast and cell-free systems: identification of a motor activity on the mitochondrial surface. J. Cell Biol. 130, 345–354. Smith, J., and Harold, J. M. (1988). Dual roles for calcium ions in apical growth of Neurospora crassa. J. Gen. Microbiol. 134, 2623–2631. Somlyo, A. V., Shuman, H., and Somlyo, A. P. (1977). Elemental distribution in striated muscle and the effects of hypertonicity. Electron probe analysis of cryosections. J. Cell Biol. 74, 828–57. Somlyo, A. P., Somlyo, A. V., and Shuman, H. (1979). Electron probe analysis of vascular smooth muscle. Composition of mitochondria, nuclei, and cytoplasm. J. Cell Biol. 81, 316–335. Somlyo, A. P., Bond, M., and Somlyo, A. V. (1985). Calcium content of mitochondria and endoplasmic reticulum in liver frozen rapidly in vivo. Nature (London) 314, 622–625. Srinivasan, S., Vargas, M. M., and Roberson, R. W. (1996). Functional, organizational, and biochemical analysis of actin in hyphal tip cells of Allomyces macrogynus. Mycologia 88, 57–70. Steer, M. W., and Steer, J. M. (1989). Pollen tube tip growth. New Phytol. 111, 323–358. Steinberg, G. (1997). A kinesin-like mechanoenzyme from the zygomycete Syncephalastrum racemosum shares biochemical similarities with conventional kinesin from Neurospora crassa. Eur. J. Cell Biol. 73, 124–131. Steinberg, G. (1998). Organelle transport and molecular motors in fungi. Fung. Genet. Biol. 24, 161– 177. Steinberg, G., and McIntosh, J. R. (1998). Effects of the myosin inhibitor 2,3-butanedione monoxime on the physiology of fission yeast. Eur. J. Cell Biol. 77, 284–293. Steinberg, G., and Schliwa, M. (1993). Organelle movements in the wild type and wall-less fz;sg;os-1 mutants of Neurospora crassa are mediated by cytoplasmic microtubules. J. Cell Sci. 106, 555– 564. Steinberg, G., and Schliwa, M. (1995). The Neurospora organelle motor: a distant relative of conventional kinesin with unconventional properties. Mol. Biol. Cell 6, 1605–1618. Steinberg, G., and Schliwa, M. (1996). Characterization of the biophysical and motility properties of kinesin from the fungus Neurospora crassa. J. Biol. Chem. 271, 7516–7521.

5. Actin and Ca2+ in Hyphal Tip Growth

185

Stemmer, P. M., and Klee C. B. (1994). Dual calcium ion regulation of calcineurin by calmodulin and calcineurin B. Biochemistry 33, 6859–6866. St¨ossel, T. P. (1993). On the crawling of animal cells. Science 260, 1086–1094. Streb, H., Irvine, R. F., Berridge, M. J., and Schulz, I. (1983). Release of Ca2+ from a non-mitochondrial intracellular store in pancreatic acinar cells by inositol 1,4,5-trisphosphate. Nature 306, 67–69. Suelmann, R., Sievers, N., and Fischer, R. (1997). Nuclear traffic in fungal hyphae: In vivo Aspergillus nidulans. Mol. Microbiol. 25, 757–769. Suelmann, R., Sievers, N., Galetzka, D., Robertson, L., Timberlake, W.E., and Fischer, R. (1998). Increased nuclear traffic chaos in hyphae of Aspergillus nidulans: molecular characterization of apsB and in vivo observation of nuclear behaviour. Mol. Microbiol. 30, 831–842. Sun, H. Q., Kwiatkowska K., and Yin H. L. (1995). Actin monomer binding proteins. Curr. Opin. Cell Biol. 7, 102–110. Suresh, K., and Subramanyam, C. (1997). A putative role for calmodulin in the activation of Neurospora crassa chitin synthase. FEMS Microbiol. Lett. 150, 95–100. Szmacinski, H., Gryczynski, I., and Lakowicz, J. R. (1993). Calcium-dependent fluorescence lifetimes of Indo-1 for one- and two-photon excitation of fluorescence. Photochem. Photobiol. 58, 341–345. Tang, S., and Beeler, T. (1990). Optical response of the indicator chlortetracycline to membrane potential. Cell Calcium 11, 425–429. Taylor, D. L., and Condeelis, J. S. (1979). Cytoplasmic structure and contractility in amoeboid cells. Int. Rev. Cytol. 56, 57–144. Temperli, E., Roos, U. P., and Hohl, H. R. (1990). Actin and tubulin cytoskeletons in germlings of the oomycete fungus Phytophthora infestans. Eur. J. Cell Biol. 53, 75–88. Temperli, E., Roos, U. P., and Hohl, H. R. (1991). Germ tube growth and the microtubule cytoskeleton in Phytophthora infestans. Mycol. Res. 95, 611–617. Thaler, C. D., and Haimo, L. T. (1996). Microtubules and microtubule motors: mechanisms of regulation. Int. Rev. Cytol. 164, 269–327. That, T. C. C. T., Rossier, C., Barja, F., Turian, G., and Roos, U. P. (1988) Induction of multiple germ tubes in Neurospora crassa. Eur. J. Cell Biol. 46, 68–79. Thibaut, M., and Ansel, M. (1973). Mycological applications of X-ray microanalysis. J. Bacteriol. 116, 1181–1184. Thibaut, M., and Ansel, M. (1976). Chemical analysis of five species of Aspergillus by combined scanning electron microscopy and X-ray spectrometry. Trans. Am. Microsc. Soc. 95, 210–214. Tiburzy, R., Hoch, H. C., and Staples, R. C. (1990). Isolation and identification of actin from the phytopathogenic filamentous fungus Uromyces appendiculatus. Eur. J. Cell Biol. 53, 364– 372. Tinsley, J. H., Minke, P. F., Bruno, K. S., and Plamann, M. (1996). p150 glued , the largest subunit of the dynactin complex, is nonessential in Neurospora but required for nuclear distribution. Mol. Biol. Cell. 7, 731–742. Tolias, K. F., and Cantley, L. C. (1999). Pathways for phosphoinositide synthesis. Chem. Phys. Lipids 98, 69–77. Torralba, S., Pedregosa , A. M., De Lucas, J. R., D´ıaz, M. S., Monistrol, I. F., and Laborda, F. (1996). Effect of the microtubule inhibitor methyl benzimidazol-2-yl carbamate (MBC) on production and secretion of enzymes in Aspergillus nidulans. Mycol. Research. 100, 1375–1382. Torralba, S., Raudaskoski, M., Pedregosa, A. M., and Laborda, F. (1998a). Effect of cytochalasin A on apical growth, actin cytoskeleton organization and enzyme secretion in Aspergillus nidulans. Microbiology 144, 45–53. Torralba, S., Raudaskoski, M., and Pedregosa, A. (1998b). Effects of methyl benzimidazole-2-yl carbamate on microtubule and actin cytoskeleton in Aspergillus nidulans. Protoplasma 202, 54– 64. Tretyn, A. (1999). Calcium-dependent signal transduction pathways in plants—phytochrome mechanism of action as an example. Pol. J. Pharmacol. 51, 145–151.

186

Sara Torralba and I. Brent Heath

Trinci, A. P. J. (1984). Regulation of hyphal branching and hyphal orientation. In “The Ecology and Physiology of the Fungal Mycelium” (D.H. Jennings and A.D.M. Rayner, Eds.), pp. 23–52. Cambridge Univ. Press, Cambridge, U.K. Tsukamoto, M., Yamamoto, M., Tanabe, S., and Kamada, T. (1996). Cytochalasin-E-resistant mutants of Coprinus cinereus: Isolation and genetic, biochemical, and cytological analyses. Fung. Genet. Biol. 20, 52–58. Tucker, B. E., Hoch, H. C., and Staples, R. C. (1986). The involvement of F-actin in Uromyces cell differentiation: the effects of cytochalasin E and phalloidin. Protoplasma 176, 43–52. Uno, I., Fukami, K., Kato., H., Takenawa, T., and Ishikawa, T. (1988). Essential role for phosphatidylinositol 4,5-triphosphate in yeast cell proliferation. Nature (London) 333, 188–190. Van Iren, F., Van Essen-Joolen, L., Van der Duyn Schouten, P., Boers-Van der Sluijs, P., and de Bruijn, W. C. (1979). Sodium and calcium localization in cells and tissues by precipitation with antimonate: a quantitative study. Histochemistry 63, 273–294. Vargas, M. M., Aronson, J. M., and Roberson, R. W. (1993). The cytoplasmic organization of hyphal tip cells in the fungus Allomyces macrogynus. Protoplasma 176, 43–52. Verkhusha, V. V., Tsukita, S., and Oda, H. (1999). Actin dynamics in lamellipodia of migrating border cells in the Drosophila ovary revealed by a GFP-actin fusion protein. FEBA Lett. 445, 395–401. Watts, F. Z., Shiels, G., and Orr, E. (1987). The yeast MYO1 gene encoding a myosin-like protein required for cell division. EMBO J. 6, 3499–3505. Welch, M. D., Mallavarapu, A., Rosenblatt, J., and Mitchison, T. J. (1997). Actin dynamics in vivo. Curr. Opin. Cell Biol. 9, 54–61. Wessels, J. G. H. (1986). Cell wall synthesis in apical hyphal growth. Int. Rev. Cytol. 104, 37–79. Wessels, J. G. H. (1988). A steady-state model for apical wall growth in fungi. Acta Bot. Neerl. 37, 3–16. Wessels, J. G. H. (1990). Role of cell wall architecture in fungal tip growth generation. In “Tip Growth in Plant and Fungal Cells” (I. B. Heath, Ed.), pp. 1–29. Academic Press, San Diego. Whatley, V. L., and Harris, R. A. (1996). The cytoskeleton and neurotransmitter receptors. Int. Rev. Neurobiol. 39, 113–143. Wick, S. M., and Hepler, P. K. (1982). Selective localization of intracellular calcium with potassium antimoniate. J. Histochem. Cytochem. 30, 1190–1204. Williams, R. J. (1998). Calcium: outside/inside homeostosis and signalling. Biochim. Biophus. Acta. 1448, 153–165. Williams, M. A., and Solomkin, J. S. (1999). Integrin-mediated signaling in human neutrophil functioning. J. Leukoc. Biol. 65, 725–736. W¨osten, H. A. B., Moukha, S. M., Sietsma, J. H., and Wessels, J. G. H. (1991). Localization of growth and secretion of proteins in Aspergillus niger. J. Gen. Microbiol. 137, 2017–2023. Wu, Q., Sandrock, T. M., Turgeon, B. G., Yoder, O. C., Wirsel, S. G., and Aist, J. R. (1998). A fungal kinesin required for organelle motility, hyphal growth, and morphogenesis. Mol. Biol. Cell. 9, 89–101. Wyatt, S. E., and Carpita, N. C. (1993). The plant cytoskeleton-cell-wall continuum. Trends Cell Biol. 3, 413–417. Wymer, C. L., Bibikova, T. N., and Gilroy, S. (1997). Cytoplasmic free calcium distributions during the development of root hairs of Arabidopsis thaliana. Plan J. 12, 427–439. Xiang, X., Beckwith, S. M., and Morris, N. R. (1994). Cytoplasmic dynein is involved in nuclear migration in Aspergillus nidulans Proc. Natl. Acad. Sci. USA 91, 2100–2104. Xiang, X., Roghi, C., and Morris, N. R. (1995). Characterization and localization of the cytoplasmic dynein heavy chain in Aspergillus nidulans. Proc. Natl. Acad. Sci. USA 92, 9890–9894. Xiang, X., Zhuo, W., and Morris, N. R. (1997). Dynein localization in living cells of Aspergillus nidulans. Mol. Biol. Cell 8, 161a. Yamashita, R. A., and May, G. S. (1998). Motoring along the hyphae: Molecular motors and the fungal cytoskeleton. Curr. Opin. Cell Biol. 10, 74–79.

5. Actin and Ca2+ in Hyphal Tip Growth

187

Yoshida,T., Toda, T., and Yanagida, M. (1994). A calcineurin-like gene ppb1+ in fission yeast: mutant defects in cytokinesis, cell polarity, mating and spindle pole body positioning. J. Cell Sci. 107, 1725–1735. Yuan, S., and Heath, I. B. (1991). Chlortetracycline Staining patterns of growing hyphal tips of the oomycete Saprolegmia ferax. Exp. Mycol. 15, 91–102. Yuan, S., Du, K. H., Qung, H. L., Lu, L., and Xue, S. B. (1995). Distribution of free Ca2+ and membrane bound Ca2+ in the cell of growing hyphal tip of Saprolegnia ferax. Acta Biol. Exp. Sinica 28, 263– 271. Zha, X., Chandra, S., Ridsdale, A. J., and Morrison, G. H. (1995). Golgi apparatus is involved in intracellular Ca2+ regulation in epithelial LLC-PK1 cells. Am. J. Physiol. 268, C1133–C1140. Zhou, X. L., Stumpf, M. A., Hoch, H. C., and Kung, C. (1991). A mechanosensitive channel in whole cells and in membrane patches of the fungus Uromyces. Science 253, 1415–1417.

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6 Pattern Formation during C. elegans Vulval Induction Minqin Wang and Paul W. Sternberg Howard Hughes Medical Institute and Division of Biology California Institute of Technology Pasadena, California 91125

I. Introduction II. Spatial Regulation of VPC Competence A. lin-39 B. mab-5 III. Temporal Regulation of VPC Competence and Commitment A. The Heterochronic Pathway B. Cell Cycle Regulation of VPC Competence and Commitment C. The Identity of VPCs and VPC Daughters IV. Downstream Events of RAS Signaling A. The RAS Signaling Pathway B. Changes of Gene Expression upon Activation of RAS C. Specificity of RAS Signaling V. Negative Regulation of RAS Signaling A. Inhibition of the Basal Activity of the LET-23 Signaling Pathway B. Downregulation of the Ligand-Induced Activity of the LET-23 Signaling Pathway VI. Lateral Signaling A. 2◦ Fate Specification B. Regulators of the Lateral Signaling Pathway VII. Evolutionary Implications VIII. Conclusions and Future Directions References

Studies of C. elegans vulval development provide insights into the process of pattern formation during animal development. The invariant pattern of vulval precursor cell fates is specified by the integration of at least two signaling systems. Recent findings suggest that multiple, partially redundant mechanisms are involved in patterning the vulval precursor cells. The inductive signal activates the LET-60/RAS signaling pathway and induces the 1◦ fate, whereas the lateral signal mediated by LIN-12/Notch is required for specification of the 2◦ fate. Several regulatory pathways antagonize the RAS signaling pathway and specify the non-vulval 3◦ fate in the absence of induction. The temporal and spatial regulation of VPC

Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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competence and production of the inductive and the lateral signal are precisely coordinated to ensure the wild-type vulval pattern.  2001 Academic Press. C

I. Introduction The development of C. elegans vulva exemplifies the fundamental mechanisms of pattern formation during organogenesis. It has been chosen as a model system because of its well-defined and fixed cell lineage, amenability to analyses at a single-cell level, and availability of genetic, cellular and molecular tools (Horvitz and Sternberg, 1991). One basic feature of vulval pattern formation in C. elegans is its precision. C. elegans proceeds through four larval stages (L1–L4) before adulthood. At the L1 stage, 12 ectoblasts, P1.p–P12.p, are aligned in an anterior to posterior row along the ventral midline of the hermaphrodite body (Sulston and Horvitz, 1977). The six central ectoblasts, P3.p–P8.p, form the multipotential group of vulval precursor cells (VPCs). Although they are all competent to choose among three fates (1◦ , 2◦ , and 3◦ ) (Sulston and White, 1980; Kimble, 1981; Sternberg and Horvitz, 1986), a pattern of 3◦ -3◦ -2◦ -1◦ -2◦ -3◦ is always established in wildtype animals (Fig. 1). All six VPCs divide once about 4 h after the L2 molt. The daughters of P3.p, P4.p, and P8.p, which adopt the nonvulval 3◦ fate, fuse with the hyp7 epidermal syncytium shortly after they are born. The daughters of P6.p, which adopts the 1◦ vulval fate, divide two more times to generate a lineage of eight descendants, while the daughters of the flanking P5.p and P7.p, which

gonad

AC P3.p ο

P4.p ο

3 or 4

fuse with hyp7

3

ο

P5.p 2

ο

inductive signal

P6.p 1

ο

vulva

P7.p lateral signal

ο

2

P8.p 3

ο

fuse with hyp7

Figure 1 C. elegans vulval development. Lateral view of an L3 hermaphrodite. Ventral is down; anterior is to the left. The anchor cell (AC) is part of the somatic gonad and is dorsal to P6.p. Six vulval precursor cells (VPCs), P3.p–P8.p, lie along the ventral midline in the central body region, surrounded by the hypodermal syncytium (hyp7). At least two signals function during patterning of VPC fates. The AC produces the inductive signal (straight arrows) and induces the 1◦ fate at a high level and the 2◦ fate at a low level. The closest VPC to the AC, P6.p, adopts the 1◦ fate and signals P5.p and P7.p laterally (curved arrows) to cause them to adopt the 2◦ fate. It is not clear where the inhibitory signal comes from.

191

6. Pattern Formation during C. elegans Vulval Induction P3

P4

P5

P6

P7

P8

0 hr

L1 10

L2

20

L3

30 or

L4

S

S

S

S

ο

ο

ο

S

L

L

T

A B 1 /B2 C 3

4

3

ο

2

N

T

T

D E

F

T F ο

1

T E

N

T

L

L

D

C B2 /B 1 A

S

ο

2

Figure 2 VPC lineages during wild-type development. The axis at left is marked in hours after hatching. Each larval stage ends with a molt, including lethargus, when the animal is inactive and ceases pumping and locomotion (checked boxes), and ecdysis, when the old cuticle is shed and the animal resumes pumping and locomotion (short horizontal lines). VPC fates are characterized by the division pattern and the morphogenetic behavior of the VPC descendants. Horizontal lines indicate cell divisions and vertical lines represent individual cells. L, longitudinal division; underlining, strong adherence to the ventral cuticle. T, transverse division. N, division did not occur and nucleus compact. S, VPC daughters did not divide and fuse with the epidermis. In wild-type animals, P3.p could either fuse at late-L2, or divide once and fuse with the epidermis at mid-L3. A, B1, B2, C, D, E, and F indicate seven types of vulval descendants.

assume the 2◦ vulval fate, give rise to a lineage of seven progeny (Fig. 2). In this review, we refer to Pn.p daughters as Pn.px, Pn.p granddaughters as Pn.pxx, and Pn.p great-granddaughters as Pn.pxxx. At the L4 stage, the 22 descendants of P5.p, P6.p, and P7.p can be classified as seven distinct types (A, B1, B2, C, D, E, and F), as they migrate dorsally and undergo cell fusion to form an epithelial tube consisting of seven toroidal cells (A to F). This structure connects to the uterus and eventually everts at late-L4 to form a mature vulva (Sharma-Kishore et al., 1999). Besides the canonical 1◦ and 2◦ lineages observed in wild-type animals, two kinds of aberrant VPC lineages, hybrid (sometimes called half-vulval) and intermediate, are observed when vulval patterning is disturbed. In hybrid lineages, one VPC daughter cell fuses with hyp7, while its sister behaves like a 1◦ or 2◦ VPC daughter. Intermediate lineages exhibit both 1◦ and 2◦ features (Katz et al., 1995).

S

ο

3

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At least two different intercellular signaling pathways are involved in patterning VPC fates (Fig. 1; reviewed in Eisenmann and Kim, 1994; Sundaram and Han, 1996; Greenwald, 1997; Kimble and Simpson, 1997; Kornfeld, 1997; Sternberg and Han, 1998). The inductive signal LIN-3, an epidermal growth factor (EGF)like molecule produced by the anchor cell (AC) in the somatic gonad, induces P6.p to adopt the 1◦ fate (Kimble, 1981; Hill and Sternberg, 1992; Katz et al., 1995). This signal is transduced by the receptor tyrosine kinase LET-23 and a conserved signaling pathway that activates LET-60/RAS (Aroian et al., 1990; Beitel et al., 1990; Han and Sternberg, 1990; Katz et al., 1996). Overexpression of lin-3 or let-60, or constitutive activation of let-23 or let-60, causes P3.p, P4.p, or P8.p to adopt vulval fates in addition to P5.p–P7.p (a multivulva or Muv phenotype). Reduction-of-function mutations in genes of this pathway can cause P5.p–P7.p to assume the nonvulval 3◦ fate (a vulvaless or Vul phenotype) (Hill and Sternberg, 1992; Aroian et al., 1990; Han et al., 1990; Katz et al., 1995). Lateral signaling between P6.p and its neighbors P5.p and P7.p utilizes a Notch/LIN-12 family receptor encoded by lin-12 (Yochem et al., 1988). All six VPCs can adopt a 2◦ fate when LIN-12 is activated by a gain-of-function mutation, whereas none of them adopt a 2◦ fate in a lin-12 loss-of-function mutant background (Greenwald et al., 1983; Sternberg, 1988; Sternberg and Horvitz, 1989). A third signal has been proposed to come from the surrounding hyp7 epidermal syncytium (Herman and Hedgecock, 1990; Hedgecock and Herman, 1995), although more recent findings suggest that it may also function autonomously within the VPCs, as discussed later (Lu and Horvitz, 1998; Thomas and Horvitz, 1999; A. Gonzalez-Serricchio and P. Sternberg, in preparation). It negatively regulates the basal activity of let-23 and ensures that no induction will occur in the absence of LIN-3 (Clark et al., 1994; Huang et al., 1994). Two distinct, but not mutually exclusive, models have been proposed to explain VPC fate patterning. The morphogen model is based on evidence demonstrating that VPCs can be induced to adopt the 1◦ or 2◦ fate depending on the level of LIN-3 signal they receive (Sternberg and Horvitz, 1986; Katz et al., 1995). Since P6.p is closer to the AC compared with flanking P5.p and P7.p, a graded signal from the AC would be able to directly induce both 1◦ and 2◦ fates in a dose-dependant manner. A null or subthreshold dose of LIN-3 leads to the 3◦ fate, whereas a low and a higher dose of LIN-3 induces the 2◦ and 1◦ fates, respectively. The sequential induction model is supported by results of genetic mosaic analysis of let-23 (Koga and Ohshima, 1995; Simske and Kim, 1995). In particular, mosaic animals that have LET-23 in P6.p, but not in P5.p and P7.p, typically have wild-type vulvae. Thus, LET-23 signaling may not be required cell-autonomously in P5.p and P7.p to adopt the 2◦ fate, at least in most animals. According to this model, the AC only induces P6.p to become 1◦ , which subsequently produces a lateral signal to induce P5.p and P7.p to become 2◦ through LIN-12. It is unclear whether either mechanism is sufficient to form the invariant pattern of VPC fates. The morphogen model requires either of two postulations to explain

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the role of LIN-12: Either there is autocrine activation of a LIN-12-dependent pathway, or LIN-12 is not necessary for the 2◦ fate with a low LIN-3 level. In the sequential model, the infrequent failure to induce LET-23 deficient neighbors to adopt the 2◦ fate by a wild-type P6.p suggests that P5.p and P7.p might normally be induced directly by the AC. It is possible that both mechanisms are used during normal development and are partially redundant. Additionally, detailed analysis of the patterning of VPC fates has indicated that almost all aspects of this process are strictly controlled to ensure that a functional vulva is always formed. In this review, we describe the latest advances in our understanding of the integration of multiple mechanisms to achieve precise pattern formation.

II. Spatial Regulation of VPC Competence To establish the pattern of VPC fates, both the production of inductive signal and the response of VPCs must be spatially and temporally regulated (Table I). The production of LIN-3 is spatially controlled by restricting its expression to the AC (Hill and Sternberg, 1992). Consequently, the closest VPC to the AC, P6.p, always receives the highest level of LIN-3. One closely related issue is the relative positioning of the VPCs to the AC, which appears to involve the homeotic gene mab-5 (Clandinin et al., 1997). In some mab-5(lf) mutants, the relative positioning of the VPCs to the AC is shifted so that P7.p, instead of P6.p, is the closest VPC to the AC. Also, the amount of LIN-3 expressed by the AC needs to be controlled to limit its potentially graded distribution, since excessive secretion can induce ectopic vulval fates. For example, transgenic animals bearing multiple copies of the lin-3 gene or heat shocked animals carrying a transgene expressing LIN-3 EGF domain under the control of a heat shock promoter (hs-LIN-3) display a multivulva (Muv) phenotype (Hill and Sternberg, 1992; Katz et al., 1995). Besides the limited production and potentially graded spatial distribution of LIN-3, spatially restricted VPC competence is another important mechanism to ensure the precision of vulval pattern formation.

A. lin-39 Of the epidermal Pn.p cells, P1.p–P11.p, the anterior P1.p–P2.p and the posterior P9.p–P.11p adopt the 4◦ (sometimes called F) fate (i.e., they fuse with hyp7 at the L1 stage; Sulston and Horvitz, 1977). P3.p–P8.p remain unfused and form the group of vulval precursor cells (VPCs) competent to generate the vulva (Sulston and White, 1980; Kimble, 1981; Sternberg and Horvitz, 1986). Two genes in the homeotic cluster (HOM-C) of C. elegans, lin-39 and mab-5, are expressed in all VPCs and the posterior two VPCs, respectively (Wang et al., 1993; Clark et al., 1993). lin-39 is the ortholog of the Drosophila gene Sex combs reduced/deformed/proboscipedia and

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Table I Genes involved in patterning of VPC fates VPC fates

Function

Genes

Predicted protein

Spatial competence

VPC specification and competence

lin-39 mab-5 bar-1

Scr homeodomain Antp homeodomain ␤-catenin

Temporal competence

The heterochronic pathway

1◦

Regulatory target of heterochronic genes Ligand Receptor Signal transducers

lin-4 lin-14 lin-28 cki-1

let-60 lin-45 mek-2 mpk-1 lin-2 lin-7 lin-10

Regulatory RNA Novel nuclear protein RNA binding protein Cyclin-dependent kinase inhibitor Epidermal growth factor Receptor tyrosine kinase SH3-SH2-SH3 adaptor Guanine nucleotide releasing protein RAS RAF MAP kinase kinase MAP kinase PDZ PDZ PDZ

ptp-2 ksr-1 sur-6 sur-8/soc-2 lin-25 sur-2 lin-1 lin-31

Tyrosine phosphatase SHP2 Novel protein kinase Protein phosphatase 2A-B Leucine-rich repeat Novel, nuclear Novel, nuclear ETS Winged helix

gap-1 unc-101 sli-1 ark-1 sur-5

GTPase activating protein AP47, clathrin adaptin CBL Tyrosine kinase Novel

lin-15A lin-8 lin-38 lin-56 egr-1 egl-27 lin-15B lin-9 lin-35 lin-36 lin-37

Novel Not cloned Not cloned Not cloned MTA1 MTA1 Novel Novel Rb Novel Not cloned

Localization of LET-23

Positive regulators

Transcription factors 3◦

Negative regulators of the ligand-induced activity of LET-23 Negative regulator of RAS Class A synMuv Negative regulators of the basal activity of LET-23

Class B synMuv

lin-3 let-23 sem-5 let-341

(Continues)

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6. Pattern Formation during C. elegans Vulval Induction Table I (Continued) VPC fates

Function

Genes

Class A and B synMuv NURD complex

2◦

Receptor Positive regulators Negative regulators

Predicted protein

lin-51 lin-52 lin-54 lin-55 tam-1

Not cloned Not cloned Not cloned Not cloned RING finger/B box nuclear protein

lin-53 hda-1 rba-1 chd-3 chd-4 lin-12 sel-12 sup-17

Rb-associated protein Histon deacetylase Rb-associated protein Highly similar chromodomain helicase protein Notch Presenilin ADAM metalloprotease

sel-1 sel-10

Extracellular protein CDC4

is required for VPC specification. In a lin-39 loss-of-function background, P3.p– P8.p fail to become VPCs and fuse with hyp7, as do the more anterior and posterior Pn.p cells (Clark et al., 1993). Thus, spatial limit of lin-39 expression in P3.p–P8.p might restrict the number of VPCs capable of generating vulval lineages (Fig. 3). LIN-39 activity is required not only in the L1 stage, but later as well, to keep VPCs unfused and maintain their identity. In wild-type animals, P3.p sometimes adopts the 4◦ fate, although it fuses with hyp7 at late-L2 rather than L1. The bar-1 gene, which encodes a ␤-catenin/Armadillo homolog in the Wnt signaling pathway, has been proposed to control the late 4◦ versus VPC fate decision in P3.p–P8.p by regulating lin-39 expression (Eisenmann et al., 1998). In bar-1 loss-of-function mutants, VPCs including P3.p adopt the 4◦ fate at late-L2 more

AC

P1.p

P2.p

VPC

VPC VPC

P3.p

P4.p

P5.p

VPC VPC VPC P6.p

P7.p

P8.p

P9.p

P10.p

P11.p

P12.pa

LIN-39 MAB-5 Figure 3 Spatial limitation of VPC competence. The VPC multipotential group consists of the six central Pn.p cells. lin-39 is expressed in P3.p–P8.p, while mab-5 is expressed in posterior Pn.p cells, including P7.p and P8.p. LIN-39 activity is required to specify P3.p–P8.p to be VPCs and promote their competence to respond to the inductive signal LIN-3. MAB-5 activity reduces the competence of P7.p and P8.p.

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frequently, presumably because of insufficient LIN-39 expression in VPCs. This late function of LIN-39 extends through VPC induction, where LIN-39 plays a critical role in regulating VPC response to inductive signal (Clandinin et al., 1997; Maloof and Kenyon, 1998; also see later discussion). B. mab-5 Genetic studies on the Antennapedia homolog mab-5 suggest that MAB-5 reduces the competence of P7.p and P8.p, thus forming a nonuniform pattern of VPC competence (Clandinin et al., 1997). Isolated P7.p and P8.p cells are less sensitive and have a reduced ability to respond to low levels of inductive signal compared with other anterior Pn.p cells, such as P6.p (Katz et al., 1995). Loss of mab-5 activity increases the sensitivity of P8.p to respond to both hs-LIN-3 and the cell-autonomous activation of the inductive pathway by a let-23 gain-of-function mutation (Clandinin et al., 1997). MAB-5 is expressed in P7.p–P11.p, but not in P3.p–P6.p (Fig. 3; Salser et al., 1993). Misexpression of MAB-5 in P3.p–P6.p (Salser and Kenyon, 1992) reduces their responsiveness to inductive signal in sensitized backgrounds (Clandinin et al., 1997). The competence of P6.p to respond to the AC signal is likely initially distinct from that of P7.p and P8.p, and the positional information provided by the HOM-C genes accounts for at least some of this difference. It is not known whether more anteriorly expressed homeotic genes, such as ceh-13 (Wang et al., 1993; Wittmann et al., 1997; Brunschwig et al., 1999), have similar functions to lin-39 or mab-5 to regulate the responsiveness of P3.p–P5.p. Overall, P6.p may be predisposed to be the one that perceives the highest level of inductive signal LIN-3.

III. Temporal Regulation of VPC Competence and Commitment Unlike the strict spatial limitation of LIN-3 expression, the temporal expression of LIN-3 in the AC appears to span from early L2 to mid L4 according to a lin-3::lacZ reporter gene (Hill and Sternberg, 1992; Euling and Ambros, 1996a). Although such reporter genes do not necessarily reflect the native gene expression in detail, we consider the issues that arise if LIN-3 is expressed for a longer period than is apparently necessary. Establishing windows of VPC competence to control timing of vulval induction is likely of importance, unless the release of LIN-3 is temporally regulated. In wild-type animals, the window of VPC competence is limited to late L2 and early L3, which is the normal time of inductive and lateral signaling. This makes sense in that induced VPCs need to be coordinated in their cell divisions and morphogenesis of their descendants. Furthermore, it is critical for a functional vulva to properly attach to the sex musculature, which is aligned by the gonad

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during the L3 stage (Sulston and Horvitz, 1977; Stern and DeVore, 1994). Proper attachment to the AC, which organizes the uterine–vulval connection throughout the L3 and L4 stages, is also essential (Newman et al., 1996). Problems that arise during vulval development caused by heterochronic mutations, which alter the timing of postembryonic development of most tissues including the vulva, but not the gonad, are consistent with these ideas. As in wild-type animals, vulval fates adopted by precociously dividing VPCs are dependent on the AC signal (Euling and Ambros, 1996a). This strongly supports that the AC signal is present earlier than needed and uncontrolled acquisition of VPC competence will result in untimely vulval development with deleterious consequences. As a result, these heterochronic mutants are defective in egg-laying because of abnormal vulval lineages and abnormal interaction between the vulva and neighboring tissues. In general, two basic mechanisms can regulate the timing of inductive responses in development: cell cycle and absolute time. Inductive signaling is often related to cell division history of responding cells, and cells can measure time progression by counting cell cycles (e.g., Raff et al., 1985). For example, cell cycle stage biases the decision of Dictyostelium cells to become either prespore or prestalk after starvation (Gomer and Firtel, 1987). In the mammalian cerebral cortex, S phase cortical progenitors are multipotent, whereas post-S progenitors are restricted in their competence (McConnell and Kaznowski, 1991). The other method of chronometry involves an internal clock of absolute time, illustrated by muscle formation in Xenopus (reviewed in Cooke and Smith, 1990). Also, cell cycle progression is not required for cell differentiation in patterning of the epidermal cells in Drosophila and neuronal induction in Xenopus (Hartenstein and Posakony, 1990; Edgar and O’Farrel, 1990; Harris and Hartenstein, 1991).

A. The Heterochronic Pathway Several studies have begun to elucidate the complex mechanisms that regulate the timing of VPC competence and choices of cell fates (Euling and Ambros, 1996a, 1996b; Ambros, 1999; Wang and Sternberg, 1999). In wild-type animals, the 20-h VPC cell cycle starts from mid-L1 to mid-L3 (Fig. 4). Based on measurement of DNA content and hydroxyurea sensitivity, VPC S phase occurs approximately 3 h before mitosis (Fig. 4; Euling and Ambros, 1996a). Previous results indicate that VPCs respond to inductive and lateral signals during the last 4 h of the VPC cell cycle, around the late L2 to early L3 stage (Kimble, 1981; Greenwald et al., 1983; Sternberg and Horvitz, 1986; Ferguson et al., 1987). This last 4 h of the VPC cell cycle corresponds to the end of the G1 phase, as well as S and G2 phases. Analysis of the heterochronic pathway has demonstrated coordination between VPC cell cycle progression and developmental signals in wild-type animals. Heterochronic genes regulate the timing of postembryonic development, and perhaps, in the case of VPC development, by affecting cell cycle progression of specific

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cells. Mutations in heterochronic genes have been shown to cause either precocious or delayed vulval development (Ambros and Horvitz, 1984). Specifically, heterochronic genes control the completion of the G1 phase of the VPC cell cycle without altering the time that VPCs are born, the cell cycle of VPC progeny, or the timing of inductive signal production. Loss-of-function mutations in lin-14 (a novel nuclear protein) or lin-28 (a cytoplasmic RNA binding protein), which cause precocious vulval development, specifically shorten the G1 phase of the VPC cell cycle (Ambros and Horvitz, 1984, 1987; Wightman et al., 1991; Euling and Ambros, 1996a; Moss et al., 1997). In contrast, lin-14(gf) or lin-4(lf) mutations cause the opposite phenotype: delayed or blocked VPC divisions (Euling and Ambros, 1996a; Ambros and Horvitz, 1987). In summary, heterochronic genes control both the acquisition of VPC competence and the completion of G1, although it has not been directly demonstrated whether these two events are causally related. Recent studies have identified a new gene, cki-1, as one of the regulatory targets of the heterochronic genes (Hong et al., 1999). CKI-1 belongs to the CIP/KIP

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family of cyclin-dependent kinase inhibitors, and regulates G1 progression of blast cells, including VPCs. In addition to its expression in cell-cycle arrested cells, cki-1::GFP is expressed during the G1 phase of the cell cycle in differentiating cells such as VPCs. Expression of CKI-1 specifically in P6.p under the control of the egl-17 promoter (Burdine et al., 1998) blocks the division of P6.p. In contrast, blocking cki-1 activity by RNA-mediated interference (RNAi) causes the six central Pn.p cells to undergo precocious cell division in the L2 stage and produce 12 VPCs. These data coupled to the observation that cki-1::GFP expression is reduced in VPCs in a lin-14(lf) mutant background suggest that lin-14 controls VPC cell cycle progression at least in part by regulating cki-1 (Hong et al., 1999).

B. Cell Cycle Regulation of VPC Competence and Commitment Although the heterochronic gene pathway affects both VPC cell cycle and competence, it is not clear whether VPC cell cycle progression is directly coupled to VPC competence or specification of cell fates. Ambros (1999) has shown that different phases of VPC cell cycle are linked to decisions of cell fates, using activation of egl-17::GFP expression (Burdine et al., 1998) and the downregulation of LIN-12::GFP (Levitan and Greenwald, 1998a) as two markers for differentiation of the 1◦ fate. A P6.p cell arrested in S phase by hydroxyurea treatment nonetheless exhibits features of a specified 1◦ VPC, including activation of egl-17::GFP expression, decreasing of lin-12::GFP expression, and inhibition of egl-17::GFP expression in its neighbors in a Muv mutant background. Thus, induction and differentiation of the 1◦ fate can occur prior to completion of the S phase of the VPC cell cycle. Temperature-shift experiments using a temperature sensitive gain-offunction allele of lin-12 suggest that the 1◦ fate decision is difficult to reverse to the 2◦ fate after a VPC has traversed the S phase of the cell cycle. In contrast, the 3◦ fate is readily convertible to the 2◦ fate after the S phase has passed. Therefore, specification of VPC fates is temporally coordinated so that specification of the 1◦ fate occurs prior to the completion of the S phase and therefore specification of the 2◦ or 3◦ fate, which may require the S phase (Fig. 4; Ambros, 1999). Complementary results have been obtained using a different approach, which addresses commitment rather than specification (Wang and Sternberg, 1999). In a sensitized background of reduced inductive signaling, the AC signal is required after the VPC division. By establishing a heat shock inducible LIN-3 system, one is able to turn on LIN-3 expression at different times. When challenged by LIN-3, presumptive 3◦ VPC (P3.p, P4.p and P8.p) daughters, 2◦ VPC (P5.p and P7.p) daughters with wild-type LIN-12 activity, and 2◦ VPC (P3.p–P8.p) daughters with activated LIN-12 are all competent to respond to LIN-3 and can be converted to the 1◦ fate. Therefore, specified 2◦ or 3◦ VPCs are not yet committed to their fates, and can later be switched to the 1◦ fate if LIN-3 is received by their daughters (Fig. 4; Wang and Sternberg, 1999).

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A pivotal step in vulval patterning is 1 fate specification and commitment. The 1◦ vulval lineage is a critical component of a functional vulva, whereas the 2◦ or 3◦ lineages are sometimes dispensable (Horvitz and Sulston, 1980; Sulston and White, 1980; M. Barr and P. Sternberg, unpublished). Moreover, the 1◦ VPC is required to laterally signal its neighbors to become 2◦ after its specification, in order to produce the wild-type vulval pattern. Combined mechanisms of establishing the sequence of specification of VPC fates, maintaining the competence of VPC daughters, and irreversibility of 1◦ fate decision are essential to achieve the prioritization of 1◦ over 2◦ and 3◦ fates as well as the temporal coordination of inductive and lateral signaling events (Fig. 4). A sequence of the 1◦ fate specification prior to the 2◦ fate biases the 1◦ fate decision and is consistent with the sequence of the inductive and lateral signaling process. Expanding the window of VPC competence to adopt the 1◦ fate to VPC daughters provides a greater window of time to maximize the possibility of induction of a 1◦ fate, which is especially important when the inductive signal is not sufficient. The irreversibility of the 1◦ fate decision, combined with reversibility of the decisions to become 2◦ and 3◦ , ensures that specification to be 1◦ overcomes a prior decision to be 2◦ or 3◦ and a 1◦ fate is specified regardless of when it happens. Since low levels of LIN-3 also appear to induce the 2◦ fate (Katz et al., 1995), different VPC fates can be considered as representing different states of VPCs that interpret different amounts of LIN-3 received over time and “ratchet” toward the final 1◦ fate. Similar models have been proposed in amphibia and Drosophila (Fig. 5A; Gurdon et al., 1995; Freeman, 1997). When the extent of vulval differentiation is reduced by reduction of signaling in the inductive signaling pathway, 2◦ and other noncanonical VPC fates are observed at the expense of the 1◦ fate. The “ratchet” model can explain hybrid or intermediate vulval lineages that arise under various conditions (Sulston and Horvitz, 1981; Sternberg and Horvitz, 1986; Ferguson et al., 1987; Sternberg and Horvitz, 1989; Thomas et al., 1990; Aroian and Sternberg, 1991; Han et al., 1993; Miller et al., 1993; Lackner et al., 1994; Tuck and Greenwald, 1995; Beitel et al., 1995; Katz et al., 1995; Koga and Oshima, 1995; Simske and Kim, 1995; Katz et al., 1996; Hajnal et al., 1997; Miller et al., 1996; Wang and Sternberg, 1999). In this model, different levels of inductive signaling and length of exposure to the inductive signal might generate a series of diverse responses and different states, which can be translated into a series of VPC fates (Fig. 5A). Thus, various fates of the VPCs can result from VPCs going through a common program and exiting at different times. In wildtype animals, the 3◦ , 2◦ , and 1◦ fates constitute the three basic states that VPCs go through when the level of the inductive signal is none, low, or high, respectively. Hybrid lineages with both 3◦ and 2◦ or 3◦ and 1◦ features (3◦ /2◦ and 3◦ /1◦ ), as well as intermediate lineages with both 2◦ and 1◦ features (1◦ /2◦ ), could be the result of intermediate states between 3◦ and 1◦ . VPCs can go through these states when the level of, or the exposure time to, the inductive signal LIN-3 is not sufficient.

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One interesting feature of hybrid lineages is that the VPC daughter closer to the signal source (the AC or the 1◦ VPC) is always the one generating the half vulval lineage, whereas the more distal daughter adopts the half 3◦ fate. Although this can be explained by polarized VPCs in the “ratchet” model, an alternative way of generating hybrid and intermediate lineages may be that daughters of uncommitted

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VPCs respond to later inductive or lateral signal independently (Fig. 5B). According to this scenario, daughters of a presumptive 3◦ VPC might be reprogrammed to generate hybrid lineages of 3◦ /2◦ or 3◦ /1◦ , or intermediate lineages of 2◦ /1◦ . Inducing daughters of a presumptive 2◦ VPC might lead to intermediate lineages of 2◦ /1◦ . It is probable that the combined action of both “ratchet” and responding VPC daughter mechanisms is responsible for hybrid and intermediate lineages.

C. The Identity of VPCs and VPC Daughters In wild-type animals, the six central Pn.p cells form multipotential VPCs, and Pn.px cells generated after one round of cell division become VPC daughters. One intriguing issue is what makes Pn.px cells different from their parents. Both cell cycle dependent and cell cycle independent mechanisms may be involved. The cell cycle related mechanism counts the numbers of cell cycles Pn.p cells have undergone instead of real time. For example, hydroxyurea arrested Pn.p cells exposed to LIN-3 at a time when they are chronologically older than normal VPC daughters still respond as VPCs rather than VPC daughters (Wang and Sternberg, 1999). Furthermore, precocious VPCs in a lin-14(lf) or lin-28(lf) background acquire competence to express vulval fates and produce precociously differentiated VPC daughters, presumably by shortening G1 of the VPC cell cycle (Euling and Ambros, 1996a). In addition, Pn.px and Pn.pxx cells, but not Pn.pxxx cells, in lin-28(lf) or lin-14(lf) mutants can be reprogrammed to the multipotential VPC states upon post-dauer development (Euling and Ambros, 1996b), indicating the importance of the cell division history of Pn.p cells and their descendants. Meanwhile, cell cycle independent mechanisms may exist to monitor the chronological age of central Pn.p cells and maintain their VPC identity until mid-L3, regardless of the cell division history of Pn.p cells or their progeny. Genes in the heterochronic pathway are probably involved. First, retarded cell divisions in a lin-4(lf) or lin-14(gf) background result in abnormal Pn.px cell identity (Euling and Ambros, 1996a), suggesting that they are not equivalent to retarded cell divisions resulting from hydroxyurea treatment, which are apparently normal. Second, although shortening the Pn.p cell cycle by lin-14 or lin-28 mutations shares similar properties with blocking cki-1 activity by RNAi, the results are distinct. In both cases, divisions of the central Pn.p cells occur precociously at the L2 stage, distinct from Pn.p divisions occurring at mid-L3 in wild-type animals. However, unlike Pn.px cells from precocious Pn.p divisions in lin-14(lf) or lin-28(lf) mutants, which produce precocious VPC daughters, Pn.px cells in cki-1 RNAi animals retain the developmental potential of VPCs after Pn.p cells enter the cell cycle and divide once (Hong et al., 1999). These VPCs specified by Pn.px cells are similar to the extra VPCs caused by rare precocious divisions of Pn.p cells in lin-31 and lin-25 mutants (Ferguson et al., 1987; Miller et al., 1993). It is worth noting that the absolute timing of Pn.p divisions, rather than the potential of cells to undergo more than

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two rounds of cell divisions, is likely to play an important role in determining the identity of Pn.p and Pn.px cells. For example, when the cullin CUL-1 is mutated, although the Pn.p cells fail to exit cell cycle and divide excessively, the divisions occur at mid-L3 as in wild-type animals, and no extra VPCs are produced (Kipreos et al., 1996).

IV. Downstream Events of RAS Signaling The inductive signal LIN-3 produced by the AC activates the RAS signaling pathway in responding VPCs. Extensive genetic screens and molecular approaches have identified numerous components in this pathway. However, relatively little is known about the downstream targets of the inductive signaling pathway, and how activation of RAS is translated into vulval fates.

A. The RAS Signaling Pathway According to the current model, lin-3 encodes an EGF-like growth factor, the inductive signal produced by the AC (Hill and Sternberg, 1992). LET-23, a receptor tyrosine kinase, is the receptor of the inductive signal (Aroian et al., 1990). Consistent with this model, LET-23 is not only expressed in all VPCs, but also localized on the basolateral surface of VPCs, the side facing the AC (Simske et al., 1996). As positive regulators of LET-23-mediated RAS activation, LIN-2, LIN-7, and LIN-10 have been identified as PDZ domain–containing proteins that are required to localize LET-23 basolaterally in VPCs (Simske et al., 1996; Hoskins et al., 1996; Kaech et al., 1998; Whitfield et al., 1999). Biochemical experiments have shown that LIN-2, LIN-7, and LIN-10 form a complex and bind to the C terminus of LET-23 via their PDZ domains. Loss-of-function mutations in lin-2, lin-7, or lin-10 result in mislocalization of LET-23 in the apical domain of the VPCs, which correlates with an incompletely penetrant Vul phenotype. In addition to allowing better access to LIN-3 from the AC, the basolateral clustering of LET-23 might also allow amplification or maximum activation of the RAS signaling pathway, which is independent of ligand binding. For example, multivulva mutants that might be independent of LIN-3 inductive signal have more VPCs adopting the vulval fates than do Muv animals with additional loss-of-function mutations in lin-2, lin-7, or lin-10 (Ferguson et al., 1987; Katz et al., 1996; Lu and Horvitz, 1998; Thomas and Horvitz, 1999). After the LIN-3 ligand activates its receptor LET-23, the signal is transduced by a series of factors. The essential genes in the pathway include sem-5 (GRB2), let-341 (SOS), let-60 (RAS), lin-45 (RAF), mek-2 (MAP kinase kinase), and mpk-1/sur-1 (MAP kinase) (Beitel et al., 1990; Han and Sternberg, 1990; Clark et al., 1992a, 1992b; Han et al., 1993; Lackner et al., 1994; Wu and Han, 1994;

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Kornfeld et al., 1995a; Wu et al., 1995; C. Chang, N. Hopper, and P. Sternberg, in preparation). ptp-2 (an SH2-containing protein tyrosine phosphatase SHP2), ksr-1 (a novel protein kinase), sur-6 (a regulatory B subunit of protein phosphatase 2A PP2A-B), and sur-8/soc-2 (a novel leucine-rich repeat containing protein) have been identified as positive regulators downstream or parallel of let-60 ras (Kornfeld et al., 1995b; Sundaram and Han, 1995; Gutch et al., 1998; Sieburth et al., 1998, 1999). lin-25 and sur-2 encode two novel protein that are positive factors downstream of MPK-1 (Singh and Han, 1995; Tuck and Greenwald, 1995). LIN-31 (a winged helix transcription factor) and LIN-1 (a ETS domain transcription factor) are two downstream effectors that are regulated by MAP kinase phosphorylation (Miller et al., 1993; Beitel et al., 1995; Tan et al., 1998). It is known in other organisms that activation of RAS signaling directly and indirectly leads to changes of gene transcription. However, exactly what happens in the VPC nucleus and how the 1◦ fate is programmed after RAS activation remains a mystery and largely unexplored. The phenotypes of lin-25, sur-2, lin-31, and lin-1 mutants are complex and epistasis analysis fails to place the transcription factors involved in a linear pathway (Miller et al., 1993; Beitel et al., 1995; Singh and Han, 1995; Tuck and Greenwald, 1995). The molecular switch determining the 1◦ fate may be the accumulation of transcription factors upon activation of RAS, which, once a certain threshold is reached, may turn on a whole set of downstream genes needed to execute the 1◦ fate. These genes may include those required for immediate and later responses to RAS activation, such as initiation of lateral signaling, inhibition of cell fusion, and promotion of cell division. This conceivably irreversible process could result in VPC commitment toward the final 1◦ fate.

B. Changes of Gene Expression upon Activation of RAS Several reports have addressed the issue of changes of gene expression in P6.p that manifest the differentiation of the 1◦ fate (Simske et al., 1996; Clandinin et al., 1997; Eisenmann et al., 1998; Burdine et al., 1998; Levitan and Greenwald, 1998a; Maloof and Kenyon, 1998). Currently, the known events following the activation of the RAS signaling pathway include upregulation of lin-39 and let-23 expression, downregulation of lin-12 expression, and activation of egl-17 expression. Besides its early function to prevent fusion of P3.p–P8.p in L1 and specify the VPCs, the HOM-C gene lin-39 is required later for VPCs to respond to the inductive signal from late L2 to early L3 (Clandinin et al., 1997; Maloof and Kenyon, 1998). First, immunocytochemical experiments using LIN-39 antibodies reveal that lin-39 expression is increased in P6.p during the time of vulval induction. Second, epistasis analysis indicates that LIN-39 functions downstream of the RAS signaling pathway. A reduction-of-function mutation of lin-39 results in defective

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vulval lineages and suppresses the Muv phenotype caused by let-60(gf). Furthermore, when provided with early LIN-39 activity, but no LIN-39 activity at the time of vulval induction, unfused VPCs in a lin-39(lf) background adopt nonvulval fates. Along with the temperature-sensitive periods of LIN-39, these results offer convincing evidence that LIN-39 activity is required again later during the time of vulval induction. LIN-39 may be functionally important for the responsiveness of VPCs to inductive signal, since lin-39(rf) reduces the induction level in animals with a sensitized background of a Vul mutation or low levels of lin-3 expression (Clandinin et al., 1997; Maloof and Kenyon, 1998). When mutated, bar-1, acting in VPCs to maintain the expression of lin-39 during vulval induction, also causes defects in 1◦ and 2◦ fate specification similar to those in a lin-39(rf) background (Eisenmann et al., 1998). Adoption of the 4◦ fate at late-L2 by P3.p in some of the wild-type animals and by some VPCs in bar-1(lf) mutants is reminiscent of the early function of lin-39 at L1. However, adoption of the 4◦ fate at late-L2 differs from Pn.p cell fusion at L1 as to whether the 4◦ fate can be suppressed by activation of the RAS pathway. Constitutive activation of the RAS pathway by Muv mutations inhibits P3.p from adopting the 4◦ fate (Ferguson et al., 1987; Clandinin et al., 1997). In addition, in bar-1(lf) mutants, P5.p–P7.p adopt the 4◦ fate significantly less frequently than P3.p, P4.p, and P8.p (Eisenmann et al., 1998). The rescue of the late 4◦ fate appears to be a result of positive regulation of lin-39 expression by activation of the RAS pathway, which does not take place in L1. It is therefore likely that both BAR-1 and the RAS pathway regulate LIN-39 in P5.p–P7.p, whereas BAR-1 alone regulates LIN-39 in P3.p, P4.p, and P8.p. Besides lin-39, the expression of let-23 is also upregulated by RAS signaling (Simske et al., 1996). As the receptor for the inductive signal LIN-3, LET-23 is apparently required for the responsiveness of VPCs together with LIN-39, whose mechanism of regulation of VPCs’ response is less clear. It is conceivable that increases in lin-39 and let-23 expression in P6.p following the initial inductive signaling could provide the basis for an amplified response of the presumptive 1◦ VPC, thereby locking the commitment toward the 1◦ fate. Increase of let-23 expression in P6.p could also serve to sequester more LIN-3 molecules on P6.p to prevent excess induction of distal VPCs (also see later discussion). According to a lin-12::GFP reporter construct, the level of LIN-12 is downregulated by RAS signaling (Levitan and Greenwald, 1998a). This is consistent with genetic epistasis results that LIN-12 acts downstream of LET-60 RAS (Han et al., 1990), and is certainly expected if the specification of the 1◦ fate occurs prior to that of the 2◦ fate (Ambros, 1999). Activation of the RAS pathway may turn on or upregulate the expression of a ligand in P6.p that laterally signals P5.p and P7.p, and subsequently, ligand-induced downregulation of LIN-12 receptor expression in P6.p. Such a mechanism would not only predispose P6.p to the 1◦ fate and enhance the difference between P6.p and its neighbors, it would also facilitate the coordination of inductive and lateral signals. Similar mechanisms may be used in

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other development systems, such as planar polarity establishment in Drosophila eye development (Tomlinson and Struhl, 1999). In this case, a polarizing signal from the equator mediated by the seven-transmembrane receptor Frizzled, followed by interaction between the photoreceptors mediated by Notch, ensures that the cells closer to the equator become the R3 photoreceptors and the polar cells adopt the R4 fate.

C. Specificity of RAS Signaling It has been known for a long time that mutations of genes in the LET-60/RAS pathway affect multiple processes in addition to vulval patterning, including viability, male spicule development, and P12 neuroectoblast fate specification (Fixsen et al., 1985; Aroian and Sternberg, 1991; Chamberlin and Sternberg, 1994; Jiang and Sternberg, 1998). How does activation of RAS in VPCs lead to adoption of vulval fates by the VPCs? The HOM-C gene lin-39 may be one of the factors required for specificity of the RAS signaling pathway (Maloof and Kenyon, 1998). Whereas lin-39 is expressed in hermaphrodite VPCs, mab-5 is expressed in male P9.p–P11.p, a posterior equivalence group similar to hermaphrodite VPCs, which divide three times to generate the hook structure in male. Ectopic expression of mab-5 in VPCs lacking LIN-39 activity causes the central Pn.p cells in the hermaphrodite to adopt features of the male posterior equivalence group. In contrast, replacing MAB-5 with misexpressed LIN-39 under control of a heat shock promoter results in precursors in the male posterior equivalence group displaying vulval characteristics. Another tissue-specific effector that specifies the outcome of the LET-60/RAS signaling is the winged helix transcription factor LIN-31 (Tan et al., 1998). lin-31 is expressed specifically in the VPCs during induction and apparently affects only vulval development when mutated. LIN-31 appears to prevent vulval induction by forming a complex with LIN-1, a general effector of activated RAS. RAS signaling induces vulval fates through the dissociation of LIN-31/LIN-1 complex upon phosphorylation of LIN-31 by MPK-1, a signal transducer downstream of RAS. Also, ectopic expression of lin-31 in another RAS responsive cell (P12) is sufficient to cause P12 to express a vulval specific marker, increased LET-23 expression.

V. Negative Regulation of RAS Signaling As discussed earlier, the system of VPC fate patterning utilizes mechanisms to impose a bias on P6.p to adopt the 1◦ fate, and P5.p and P7.p to become 2◦ . Since the number of VPCs competent to adopt the vulval fates exceeds the three central VPCs that actually do so in wild-type animals, it is crucial that additional

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mechanisms serve to prevent the anterior P3.p and P4.p and the posterior P8.p from being induced. Besides restricted spatial expression and level of LIN-3, the inductive signaling pathway is negatively regulated at two distinct levels.

A. Inhibition of the Basal Activity of the LET-23 Signaling Pathway Two functionally redundant pathways of synthetic multivulva (synMuv) antagonize the LET-23-mediated signal transduction (Horvitz and Sulston, 1980; Ferguson and Horvitz, 1985, 1989; Beitel, 1994; Clark et al., 1994; Huang et al., 1994; Lu and Horvitz, 1998; Hsieh et al., 1999; Thomas and Horvitz, 1999; Solari and Ahringer, 2000). These synMuv genes fall into two classes, A and B. So far six genes (lin-8, lin-15A, lin-38, lin-56, egr-1, and egl-27) have been found to belong to class A and 10 genes (lin-9, lin-15B, lin-35, lin-36, lin-37, lin-51, lin-52, lin-54, lin-55, and tam-1) have been placed into class B. Single or double mutants of genes belonging to the same class have a wild-type vulva, whereas double mutants of both class A and B genes display excessive induction of P3.p, P4.p, and P8.p (a Muv phenotype). It has been found that some synMuv genes (lin-53, hda-1, rba-1, chd-3, and chd-4) function in both synMuvA and synMuvB pathways (Lu and Horvitz, 1998; Shi and Mello, 1998; Solari et al., 1999; Solari and Ahringer, 2000). Several of the synMuv genes, including lin-15A, lin-15B, lin-36, and lin-9, have been cloned and shown to encode novel proteins (Clark et al., 1994; Huang et al., 1994; Beitel, 1994; Thomas and Horvitz, 1999). lin-35 and lin-53 have been shown to encode proteins similar to the tumor suppressor Rb and its binding protein RbAp48 (Lu and Horvitz, 1998). Two class A synMuv gene products, EGR-1 and EGL-27, are similar to the human metastasis tumor associated protein MTA1, a member of the nucleosome remodeling and histone deacetylase (NURD) complex (Solari et al., 1999; Solari and Ahringer, 2000). Detailed analysis on the question of site of action of synMuv genes suggests that some of them function autonomously in VPCs and others nonautonomously in the hypodermal syncytium hyp7. Genetic mosaic analysis studies of lin-15 (A and B) and lin-37 (a class B gene) suggest that both genes function nonautonomously in hyp7 (Herman and Hedgecock, 1990; Hedgecock and Herman, 1995). However, data on lin-35 (class B), lin-53 (class A and B), and lin-36 (class B) are consistent with the hypothesis that they function autonomously within the VPCs. Antibody staining demonstrates that LIN-35 protein is present in VPCs, but not in hyp7, whereas a GFP::lin-53 reporter gene is expressed in both (Lu and Horvitz, 1998). Results of both genetic mosaics and reporter gene expression of lin-36 support that this particular class B gene acts autonomously in VPC nuclei (Thomas and Horvitz, 1999). How do the synMuv pathways work to negatively regulate the RAS signaling pathway? The Muv phenotype caused by mutations in synMuv genes requires functional let-23 and genes downstream of the let-23 receptor, but does not require the

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Figure 6 Negative regulation of RAS signaling. (A) The RAS signaling output depends on both the ligand-independent and the ligand-dependent activities of LET-23. In all VPCs, genes in the synthetic multivulva pathway antagonize the basal activity of LET-23-mediated inductive pathway to prevent vulval induction when LIN-3 is absent. In P6.p, LIN-3 produced by the AC induces LET-23 to activate RAS signaling, while P3.p, P4.p and P8.p do not receive LIN-3 and adopt the nonvulval 3◦ fate. (B) LET-23 functions antagonistically in addition to activate the RAS pathway. LET-23 in P6.p sequesters LIN-3 and prevents the diffusion of LIN-3 molecules. Upon induction by LIN-3, LET-23 increases its expression level in P6.p, thereby limiting the amount of unbound LIN-3 molecules to reach distal VPCs.

gonad, the source of the LIN-3 ligand (Ferguson and Horvitz, 1989; Huang et al., 1994). Therefore, the synMuv pathways may inhibit the basal, ligand-independent activity of LET-23-mediated signaling pathway, which can be elevated by binding of the LIN-3 ligand to the LET-23 receptor (Fig. 6A). Alternatively, they may function to limit LIN-3 expression in the AC. For example, LIN-3 could be ectopically expressed in synMuv mutants, thereby inducing ectopic vulval fates even in the absence of the gonad. Finally, it is also possible that the synMuv pathways inhibit the RAS pathway by blocking a second, nongonadal signal, since three EGFlike molecules in addition to LIN-3 have been identified in the worm genome as possible candidates of EGF receptor ligands (Plowman et al., 1999). To understand the molecular mechanism by which the synMuv pathways antagonize RAS signaling, one needs to define the point at which the synMuv pathways interface with the Ras pathway, which could be at the level of the LET-23

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receptor, the nucleus, or both. In vitro binding experiments suggest interactions among LIN-35, LIN-53, and HDA-1 (a histone deacetylase involved in remodeling chromatin structure), as well as among LIN-53, LIN-36, LIN-37, and LIN-15A (Lu and Horvitz, 1998; Walhout et al., 2000). These data support the notion that LIN-35 Rb-mediated class B synMuv pathway may function by repressing transcription of target genes. The discovery of another synMuv B gene tam-1, which mediates context-dependent gene silencing, is consistent with a mechanism of transcriptional repression, possibly by chromatin remodeling (Hsieh et al., 1999). Furthermore, recent findings of components (LIN-53, HDA-1, RBA-1, CHD-3, and CHD-4) in the nucleosome remodeling and histone deacetylase (NURD) complex functioning in both synMuvA and synMuvB pathways suggest that these two pathways might redundantly recruit a core NURD complex and repress gene transcription by chromatin remodeling (Solari and Ahringer, 2000). Besides the chromatin remodeling model, it is also tempting to speculate that the class B synMuv pathway regulates cell fate specification and commitment of VPCs by Rb-associated G1/S cell cycle progression, thereby negatively regulating vulval induction.

B. Downregulation of the Ligand-Induced Activity of the LET-23 Signaling Pathway Genes that function at another level of negative regulation have been isolated as suppressors of the Vul phenotype of let-23(rf) or lin-10(rf) mutants. These include unc-101, sli-1, and gap-1 (Lee et al., 1994; Jongeward et al., 1995; Yoon et al., 1995; Hajnal et al., 1997). Other negative regulators, such as ark-1 and sur-5, have been recovered in genetic screens to look for Muv mutants in a sli-1 background and suppressors of dominant negative let-60 ras (Gu et al., 1998; N. Hopper, J. Lee, and P. Sternberg, unpublished). These negative regulators have been identified to encode a homolog of medium chain of the trans-Golgi clathrinassociated adaptin complex, the oncoprotein c-CBL, a GTPase activating protein for RAS, a tyrosine kinase, and a novel protein, respectively. Similar to single mutations in synMuv pathway genes, single mutations in these genes do not cause any vulval phenotype. However, unlike mutations in synMuv pathways, mutations of these negative regulators do not synergize greatly with synMuv mutations of either class. Also, double mutants of these negative regulators are largely or wholly dependent on the LIN-3 inductive signal in the gonad to display the Muv phenotype (Sternberg et al., 1994). Although each negative regulator may interface with the RAS signaling pathway by different underlying mechanisms, studies on these genes converge and reveal an antagonistic function mediated by LET-23 (Fig. 6B). This model was first proposed based on the hyperinduced (Hin) phenotype, defined as higher than wild-type vulval induction, caused by let-23(n1045) at 25◦ C (Aroian and Sternberg, 1991). The Hin phenotype is fundamentally different from the Muv phenotype caused

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by mutations in synMuv genes in that the excess induction is centered around the AC, is dependent on the AC signal, and is caused by a decrease of LET-23 activity (Aroian and Sternberg, 1991). Double mutants of certain reduction-offunction mutation of let-23, lin-2, lin-7, or lin-10 and sli-1, unc-101, or gap-1 also exhibit a Hin phenotype (Lee et al., 1994; Jongeward et al., 1995; Yoon et al., 1995; Hajnal et al., 1997). Mosaic analysis of let-23; gap-1 animals has shed light on the underlying mechanisms of a cell-nonautonomous inhibitory function of LET-23 (Hajnal et al., 1997). In short, the LET-23 receptors presented by the central VPCs might bind and sequester the LIN-3 molecules secreted by the AC, thereby reducing the amount of LIN-3 the distal VPCs (P3.p, P4.p, and P8.p) may receive (Fig. 6B). Increase of let-23 expression in P6.p upon activation of RAS may also facilitate the antagonistic role of LET-23 in addition to its role to activate RAS. In Hin animals, the LET-23 receptor might have reduced ability to bind and sequester LIN-3 because of mutations in the receptor itself or mislocalization caused by lin-2, lin-7, or lin-10. This leads to expanded spatial distribution of LIN-3 to reach the distal VPCs, which might induce the distal VPCs when their sensitivity is enhanced by reduction-of-function mutations in sli-1, unc-101, gap-1, or the n1045 allele of let-23.

VI. Lateral Signaling A. 2◦ Fate Specification Whereas the RAS pathway is required for the 1◦ fate, LIN-12 activity is required for the 2◦ fate. In the absence of LIN-12, the receptor for the lateral signal, VPCs never adopt the 2◦ fate (Greenwald et al., 1983; Sternberg and Horvitz, 1989). Although an isolated VPC receiving low levels of inductive signal can become 2◦ (Sternberg and Horvitz, 1986; Thomas et al., 1990; Katz et al., 1995), this has been proposed to depend on the autocrine activity of LIN-12 (e.g., Sternberg and Horvitz, 1989). Does lateral signaling prevent adjacent VPCs from both adopting the 1◦ fate or induce neighbors of a 1◦ VPC to adopt the 2◦ fate? In lin-15 mutants, all VPCs adopt vulval fates, but they display an alternating 1◦ -2◦ pattern (Sternberg, 1988). In particular, an isolated VPC lacking neighboring VPCs always adopts the 1◦ fate, whereas two isolated VPCs display a 1◦ -2◦ (or 2◦ -1◦ ) pattern. However, lateral signaling also functions in an inductive mode rather than an inhibitory mode. In let-23 mosaic animals, when P6.p has LET-23 and is induced to become 1◦ , P5.p and P7.p lacking LET-23 can still adopt the 2◦ fate (Koga and Ohshima, 1995; Simske and Kim, 1995). Strikingly, although never in wild-type animals, P4.p can become 2◦ when its neighbor P5.p adopts the 1◦ fate. Therefore, lateral signaling can induce the 2◦ fate in VPCs that do not receive the inductive signal. In wild-type animals, the induced 1◦ VPC, P6.p, signals laterally and might specify P5.p and P7.p to become 2◦ .

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One can infer from these results that the amount of LIN-3 received by P5.p and P7.p is controlled so that lateral signal always overrides inductive signal. When lin-3 is overexpressed, inductive signal can override lateral signaling to induce adjacent 1◦ fates (Katz et al., 1995). On the other hand, a wild-type vulval pattern can nevertheless be established even though the absolute level of LIN-3 produced by the AC appears to be flexible within a certain range. In some let23 mosaic animals where P6.p lacks LET-23 and therefore cannot sequester the LIN-3 molecules, LIN-3 from the AC is able to induce P5.p or P7.p to adopt the 1◦ fate (Koga and Ohshima, 1995; Simske and Kim, 1995). In dig-1 mutants in which the gonad is dorsally displaced, the AC often induces the 1◦ fate with 2◦ neighbors, although the AC is at a great distance from all VPCs (Thomas et al., 1990). Most surprisingly, some animals are Hin despite the lack of AC proximity. Therefore, the VPCs most likely interact with each other and compare the relative level rather than perceive the absolute level of the AC signal. In wild-type animals, the positioning of the AC and VPCs ensures that P6.p is biased toward the 1◦ fate, even when the intensity of the AC signal is not precisely controlled. It is not clear whether a morphogen mechanism or a sequential induction mechanism alone is sufficient to pattern the 2◦ fates invariantly. With respect to the morphogen model, although low levels of LIN-3 are capable of inducing VPCs to adopt the 2◦ fate (Katz et al., 1995), it is not necessary for 2◦ fate specification. Activated LIN-12 can cause all VPCs to adopt the 2◦ fate even when LIN-3 is not available (Sternberg and Horvitz, 1989; Greenwald and Seydoux, 1990). It is also conceivably difficult to solely rely on the distance between the AC and the VPCs to make the 1◦ versus 2◦ decision correctly every time. As to the sequential induction model, lateral signaling from a 1◦ VPC is not always sufficient to induce the 2◦ fate. In a small percentage of LET-23 mosaic animals with no LET-23 activity in P5.p or P7.p, a 1◦ P6.p can have a hybrid neighbor (Koga and Ohshima, 1995). Further clarification of how a low level of LIN-3 induces the 2◦ fate will likely provide better understanding of both mechanisms. One unique feature of the 2◦ lineage is its polarity, with D and C descendants closer to the 1◦ lineage and A and B descendants more further away. This polarity may be unrelated to LIN-12 function and require a signal from the gonad, as well as two receptors, lin-17 (a Wnt receptor) and lin-18 (a receptor tyrosine kinase) (Ferguson et al., 1987; Sternberg and Horvitz, 1988; Sawa et al., 1996; W. Katz and P. Sternberg, unpublished).

B. Regulators of the Lateral Signaling Pathway Although the identity of the lateral signal acting among the VPCs has yet to be revealed, screens for extragenic suppressors of the egg-laying defects in lin12 dominant gain-of-function mutants or lin-12 reduction-of-function mutants have identified additional components in the lateral signaling pathway (Sundaram

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and Greenwald, 1993; Levitan and Greenwald, 1995; Tax et al., 1997). Some cloned suppressors include sel-1 (an extracellular protein), sel-10 (an F-box/WD40 repeat-containing protein of the CDC4 family), sel-12 (presenilin), and sup-17 (a metalloprotease of the ADAM family) (Levitan and Greenwald, 1995; Grant and Greenwald, 1996; Hubbard et al., 1997; Wen et al., 1997). The two negative regulators of lin-12 activity, sel-1 and sel-10, may be involved in LIN-12 turnover. Specifically, SEL-10 may target LIN-12 for ubiquitin-mediated degradation, which is consistent with its cell-autonomous effect and its physical association with SKP-1 (a component of E3 complex for ubiquitin-dependent degradation) and the intracellular domain of LIN-12 (King et al., 1996; Hubbard et al., 1997; Walhout et al., 2000). The positive regulator sup-17 is homologous to Drosophila kuzbanian and is probably involved in processing the extracellular domain of LIN-12/Notch during its maturation before activation upon ligand binding (Pan and Rubin, 1997; Wen et al., 1997). SEL-12, a C. elegans presenilin, specifically affects LIN-12 accumulation in the VPC plasma membrane, and presenilin has been shown to physically interact with Notch (Levitan and Greenwald, 1998b; Ray et al., 1999). Since the reduction-of-function mutation in sel-12 only suppresses the activity of full-length LIN-12, but not constitutively activated, truncated LIN-12 possessing only the intracellular domain, sel-12 is likely involved in trafficking and cleavage of LIN-12 during its maturation before ligand binds to it. A physical association between SEL10 and SEL-12 demonstrated in vitro and in yeast cells further implies that SEL-12 may also regulate SEL-10 level in the VPCs (Wu et al., 1998; Walhout et al., 2000). The level of LIN-12 activity may be regulated by closely related processes, such as proteolytic processing, intracellular trafficking, and protein degradation. Studies of the foregoing regulators of the lateral signaling pathway raise the intriguing possibility that they are involved in regulating dynamic changes in LIN-12 activity, which is a critical element in patterning VPC fates (Levitan and Greenwald, 1998a). For example, RAS-dependent downregulation of LIN-12 in P6.p could be the result of LIN-3-induced degradation of LIN-12 that reinforces the difference between P6.p and its neighbors.

VII. Evolutionary Implications Comparative developmental studies in several nematode species within the same order as C. elegans have revealed that similar patterning of vulval fates can be accomplished by combination of a wide diversity of patterning mechanisms and cell signaling networks. These studies have brought us insight into the evolution of developmental processes, as well as deepened our understanding of C. elegans vulval development. First, the role of a single gene product can vary significantly between different species. In C. elegans, lin-39 functions both early and later in vulval cell fate specification (Clandinin et al., 1997; Maloof and Kenyon, 1998). Although its later

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role is obviously instructive, it is unknown whether the early function of LIN-39 to specify the multipotential VPC group is achieved by preventing cell fusion of the central Pn.p cells or promoting vulval fates. In Pristionchus pacificus, VPCs lacking LIN-39 activity die by programmed cell death rather than fuse with the epidermal syncytium hyp7. Double mutants with loss-of-function mutations in lin-39 and ced-3 (a gene required for apoptosis) have a functional vulva (Eizinger and Sommer, 1997; Sommer et al., 1998). Therefore, the lin-39 homolog Ppa-lin39 functions permissively to specify VPCs by inhibiting programmed cell death and does not actively participate in later vulval induction. Second, the mechanism by which the AC specifies a similar centered vulval pattern can differ. Whereas a one-step AC induction before VPC division is sufficient in wild-type C. elegans, several other nematode species (e.g., Panagrolaimus, Oscheius) utilize a two-step gonadal induction to pattern the VPC fates (F´elix and Sternberg, 1997). In C. elegans wild-type animals, the AC is no longer needed after VPC division to induce the 2◦ -1◦ -2◦ pattern. In Oscheius, ablation of the AC after VPC division results in all VPCs including P6.p adopting the same outer vulval fate normally only adopted by P5.p and P7.p. This observation suggests that two successive inductions are required for specification of the inner fates. The first induction of VPCs to adopt vulval fates in these species might correspond to an initially shared common program of 1◦ and 2◦ , perhaps a transiently induced early state in C. elegans (Fig. 5A). The second induction of VPC daughters to specify inner versus outer vulval fates might correspond to the specification of 1◦ versus 2◦ fates in C. elegans. Finally, the network of cell interactions may use an overlapping set of developmental mechanisms, but may change the relative contribution of redundant mechanisms to elicit similar outcomes. For instance, in C. elegans, differences of developmental potential among VPCs play a minor role compared with precisely controlled inductive and lateral signaling to generate the invariant vulval pattern. However, in Mesorhabditis, the gonad is not required for vulval induction after hatching (Sommer and Sternberg, 1994). In Pristionchus, there are only three VPCs in the vulval multipotential group (Sommer and Sternberg, 1996; Sommer, 1997). In these two species, positional differences among VPCs appear to be a major mechanism, probably more important than the gonadal induction. Nonetheless, use of multiple and even redundant mechanisms is one central idea to achieve a high degree of precision in patterning of vulval fates in C. elegans and other nematode species.

VIII. Conclusions and Future Directions There has been significant progress in understanding the mechanisms that are responsible for formation of the invariantly patterned C. elegans vulva. A comprehensive picture is emerging in which all aspects of cell interaction in this patterning process are under precise control. Meanwhile, our deepened knowledge raises more questions.

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While we continue to identify and functionally analyze new genes to describe the pathways involved, especially the lateral and inhibitory pathways, in greater detail, much effort is needed to understand the integration of these signaling pathways. We do not know whether any of the changes in the expression of egl-17, let-23, lin-39, and lin-12 within the 1◦ VPC are immediate steps following activation of RAS, or whether they are well downstream of RAS activation. Complex phenotypes displayed by lin-1 and lin-31 mutants make genetic analysis at the transcription factor level difficult (Miller et al., 1993; Beitel et al., 1995). Careful analysis of transcriptional regulatory regions of these genes and definitive identification of the transcription factors that bind to them should provide important insight into the signaling network that guides the expression of these genes. Moreover, the molecular link between the inductive and lateral signaling pathways and the identity of the lateral signal remain obscure, although lin-25 has been described to affect both pathways (Tuck and Greenwald, 1995). Neither do we know about the target genes of synMuv pathways and how they relate to the RAS pathway. Although the concepts of cell fate specification and commitment have been used for a long time, their precise molecular meaning is not clear. Further understanding in specification and commitment of the 1◦ and 2◦ fates will help elucidate the mechanisms underlying the bias of the 1◦ fate determination. Detailed studies of the molecular meaning of VPC competence to respond to both inductive and lateral signal, as well as its relationship to the cell-cycle machinery, will address the directly related questions of VPC commitment. In summary, future investigations of C. elegans VPC fate patterning using genetic and molecular tools will provide further comprehension of complex signaling networks and the mechanisms underlying precise pattern formation in development.

Acknowledgments We thank Tom Clandinin, Rene Garcia, Nadeem Moghal, and David Sherwood for helpful comments on the manuscript. P.W.S. is an Investigator with the Howard Hughes Medical Institute.

References Ambros, V. (1999). Cell cycle-dependent sequencing of cell fate decisions in Caenorhabditis elegans vulval precursor cells. Development 126, 1947–1956. Ambros, V., and Horvitz, H. R. (1984). Heterochronic mutants of the nematode Caenorhabditis elegans. Science 226, 409–416. Ambros, V., and Horvitz, H. R. (1987). The lin-14 locus of Caenorhabditis elegans controls the time of expression of specific postembryonic developmental events. Genes Dev. 1, 398–414. Aroian, R. V., and Sternberg, P. W. (1991). Multiple functions of let-23, a Caenorhabditis elegans receptor tyrosine kinase gene required for vulval induction. Genetics 128, 251–267.

6. Pattern Formation during C. elegans Vulval Induction

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Aroian, R. V., Koga, M., Mendel, J. E., Ohshima, Y., and Sternberg, P. W. (1990). The let-23 gene necessary for Caenorhabditis elegans vulval induction encodes a tyrosine kinase of the EGF receptor subfamily. Nature 348, 693–699. Beitel, G. J. (1994). Genetic and molecular analysis of let-60 ras, lin-1 and lin-9: genes that function in C. elegans vulval induction. Ph.D. thesis, Massachusetts Institute of Technology, Cambridge, Massachusetts. Beitel, G. J., Clark, S. G., and Horvitz, H. R. (1990). Caenorhabditis elegans ras gene let-60 acts as a switch in the pathway of vulval induction. Nature 348, 503–509. Beitel, G. J., Tuck, S., Greenwald, I., and Horvitz, H. R. (1995). The Caenorhabditis elegans gene lin-1 encodes an ETS-domain protein and defines a branch in the vulval induction pathway. Genes Dev. 9, 3149–3162. Brunschwig, K., Wittmann, C., Schnabel, R., B¨urglin, T. R., Tobler, H., and M¨uller, F. (1999). Anterior organization of the Caenorhabditis elegans embryo by the labial-like Hox gene ceh-13. Development 126, 1537–1546. Burdine, R. D., Branda, C. S., and Stern, M. J. (1998). EGF-17(FGF) expression coordinates the attraction of the migration sex myoblasts with vulval induction in C. elegans. Development 125, 1083–1093. Chamberlin, H. M., and Sternberg, P. W. (1994). The lin-3/let-23 pathway mediates inductive signaling during male spicule development in Caenorhabditis elegans. Development 120, 2713– 2721. Chang, C., Hopper, N. A., and Sternberg, P. W. (2000). C. elegans SOS-1 is required for multiple RAS-dependent events and cooperates with a SOS-1-independent pathway during vulval induction. Embo J. 19, 3283–3293. Clandinin, T. R., Katz, W. S., and Sternberg, P. W. (1997). Caenorhabditis elegans HOM-C genes regulate the response of vulval precursor cells to inductive signal. Dev. Biol. 182, 150–161. Clark, S. G., Stern, M. J., and Horvitz, H. R. (1992a). C. elegans cell-signaling gene sem-5 encodes a protein with SH2 and SH3 domains. Nature 356, 340–344. Clark, S. G., Stern, M. J., and Horvitz, H. R. (1992b). Genes involved in two Caenorhabditis elegans cell-signaling pathways. Cold Spring Harb. Symp. Quant. Biol. 57, 363–73. Clark, S. G., Chisholm, A. D., and Horvitz, H. R. (1993). Control of cell fates in the central body region of C. elegans by the homeobox gene lin-39. Cell 74, 43–55. Clark, S. G., Lu, X., and Horvitz, H. R. (1994). The Caenorhabditis elegans locus lin-15, a negative regulator of a tyrosine kinase signaling pathway, encodes two different proteins. Genetics 137, 987–997. Cooke, J., and Smith, J. C. (1990) Measurement of developmental time by cells of early embryos. Cell 60, 891–894. Edgar, B. A., and O’Farrel, P. H. (1990). The three postblastoderm cell cycles of Drosophila embryogenesis are regulated in G2 by string. Cell 62, 469–480. Eisenmann, D. M., and Kim, S. K. (1994). Signal transduction and cell fate specification during Caenorhabditis elegans vulval development. Curr. Opin. Genet. Dev. 4, 508–516. Eisenmann, D. M., Maloof, J. N., Simske, J, S., Kenyon, C., and Kim, S. K. (1998). The ␤-catenin homolog BAR-1 and LET-60 Ras coordinately regulate the Hox gene lin-39 during Caenorhabditis elegans vulval development. Development 125, 3667–3680. Eizinger, A., and Sommer, R. J. (1997). The homeotic gene lin-39 and the evolution of nematode epidermal cell fates. Science 278, 452–455. Euling, S., and Ambros, V. (1996a). Heterochronic genes control cell cycle progress and developmental competence of C. elegans vulval precursor cells. Cell 84, 667–676. Euling, S., and Ambros, V. (1996b). Reversal of cell fate determination in Caenorhabditis elegans vulval development. Development 122, 2507–2515. F´elix, M.-A., and Sternberg, P. W. (1997). Two nested gonadal inductions of the vulva in nematodes. Development 124, 253–259.

216

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Ferguson, E. L., and Horvitz, H. R. (1985). Identification and genetic characterization of 22 genes that affect the vulval lineages of the nematode Caenorhabditis elegans. Genetics 110, 17–72. Ferguson, E. L., and Horvitz, H. R. (1989). The multivulva phenotype of certain Caenorhabditis elegans mutants results from defects in two functionally redundant pathways. Genetics 123, 109–121. Ferguson, E. L., Sternberg, P. W., and Horvitz, H. R. (1987). A genetic pathway for the specification of the vulval cell lineages of Caenorhabditis elegans. Nature 326, 259–267. Fixsen, W., Sternberg, P. W., Ellis, H., and Horvitz, H. R. (1985). Genes that affect cell fates during the development of Caenorhabditis elegans. Cold Spring Harbor Symp. Quant. Biol. 50, 99–104. Freeman, M. (1997). Cell determination strategies in the Drosophila eye. Development 124, 261–270. Gomer, R. H., and Firtel, R. A. (1987). Cell-autonomous determination of cell-type choice in Dictyostelium development by cell-cycle phase. Science 237, 758–762. Grant, B., and Greenwald, I. (1996). The Caenorhabditis elegans sel-1 gene, a negative regulator of lin-12 and glp-1, encodes a predicted extracellular protein. Genetics 143, 237–247. Greenwald, I. (1997). Development of the vulva. In “C. elegans II”(D. Riddle, T. Blumenthal, B. Meyer, and J. Priess, Eds.), pp. 519–541. Cold Spring Harbor Laboratory Press, New York. Greenwald, I., and Seydoux, G. (1990). Analysis of gain-of-function mutations of the lin-12 gene of Caenorhabditis elegans. Nature 346, 197–199. Greenwald, I. S., Sternberg, P. W., and Horvitz, H. R. (1983). The lin-12 locus specifies cell fates in Caenorhabditis elegans. Cell 34, 435–444. Gu, T., Orita, S., and Han, M. (1998). Caenorhabditis elegans SUR-5, a novel but conserved protein, negatively regulates LET-60 Ras activity during vulval induction. Mol. Cell. Biol. 18, 4556–4564. Gurdon, J. B., Mitchell, A., and Mahony, D. (1985). Direct and continuous assessment by cells of their position in a morphogen gradient. Nature 376, 520–521. Gutch, M. J., Flint, A. J., Keller, J., Tonks, N. K., Hengartner, M. O. (1998). The Caenorhabditis elegans SH2 domain-containing protein tyrosine phosphatase PTP-2 participates in signal transduction during oogenesis and vulval development. Genes Dev. 12, 571–585. Hajnal, A., Whitfield, C. W., and Kim, S. K. (1997). Inhibition of Caenorhabditis elegans vulval induction by gap-1 and by let-23 receptor tyrosine kinase. Genes Dev. 11, 2715–2728. Han, M., and Sternberg, P. W. (1990). let-60, a gene that specifies cell fates during C. elegans vulval induction, encodes a ras protein. Cell 63, 921–931. Han, M., Aroian, R., and Sternberg, P. W. (1990). The let-60 locus controls the switch between vulval and non-vulval cell types in C. elegans. Genetics 126, 899–913. Han, M., Golden, A., Han, Y., and Sternberg, P. W. (1993). C. elegans lin-45 raf gene participates in let-60 ras-mediated vulval differentiation. Nature 363, 133–140. Harris, W. A., and Hartenstein, V. (1991). Neuronal induction without cell division in Xenopus embryos. Neuron 6, 499–515. Hartentein, V., and Posakony, J. W. (1990). Sensillum development in the absence of cell division: the sensillum phenotype of the Drosophila mutant string. Dev. Biol. 138, 147–158. Hedgecock, E. M., and Herman, R. K. (1995). The ncl-1 gene and genetic mosaics of Caenorhabditis elegans. Genetics 141, 989–1006. Herman, R. K., and Hedgecock, E. M. (1990). Limitations of the size of the vulval primordium of Caenorhabditis elegans by lin-15 expression in surrounding hypodermis. Nature 348, 169–171. Hill, R. J., and Sternberg, P. W. (1992). The gene lin-3 encodes an inductive signal for vulval development for C. elegans. Nature 358, 470–476. Hong, Y., Roy, R., and Ambros, V. (1999). Developmental regulation of a cyclin-dependent kinase inhibitor controls postembryonic cell cycle progression in Caenorhabditis elegans. Development 125, 3585–3597. Hopper, N. A., Lee, J., and Sternberg, P. W. (2000). ARK-1 inhibits EGFR signaling in C. elegans. Molec. Cell 6, 65–75. Horvitz, H. R., and Sternberg, P. W. (1991). Multiple intercellular signaling systems control the development of the Caenorhabditis elegans vulva. Nature 351, 535–541.

6. Pattern Formation during C. elegans Vulval Induction

217

Horvitz, H. R., and Sulston, J. E. (1980). Isolation and genetic characterization of cell-lineage mutants of the nematode Caenorhabditis elegans. Genetics 96, 435–454. Hoskins, R., Hajnal, A. F., Harp, S., and Kim, S. K. (1996). The C. elegans vulval induction gene lin-2 encodes a member of the MAGUK family of cell junction proteins. Development 122, 97– 111. Hsieh, J., Liu, J., Kostas, S. A., Chang, C., Sternberg, P. W., and Fire, A. (1999). The RING finger/B-Box factor TAM-1 and a retinoblastoma-like protein LIN-35 modulate context-dependent gene silencing in Caenorhabditis elegans. Genes Dev. 13, 2958–2970. Huang, L. S., Tzou, P., and Sternberg, P. W. (1994). The lin-15 locus encodes two negative regulators of Caenorhabditis elegans vulval development. Mol. Biol. Cell 5, 395–412. Hubbard, E. J. A., Wu, G., Kitajewski, J., and Greenwald, I. (1997). sel-10, a negative regulator of lin-12 activity in Caenorhabditis elegans, encodes a member of the CDC4 family of proteins. Genes Dev. 11, 3182–3193. Jiang, L. I., and Sternberg, P. W. (1998). Interaction of EGF, Wnt and HOM-C genes specify the P12 neuroectoblast fate in C. elegans. Development 125, 2337–2347. Jongeward, G. D., Clandinin, T. R., and Sternberg, P. W. (1995). sli-1, a negative regulator of let-23mediated signaling in C. elegans. Genetics 139, 1553–1566. Kaech, S. M., Whitfield, C. W., Kim, S. K. (1998). The LIN-2/LIN-7/LIN-10 complex mediates basolateral membrane localization of the C. elegans EGF receptor LET-23 in vulval epithelial cells. Cell 94, 761–771. Katz, W. S., Hill, R. J., Clandinin, T. R., and Sternberg, P. W. (1995). Different levels of the C. elegans growth factor LIN-3 promote distinct vulval precursor fates. Cell 82, 297–307. Katz, W. S., Lesa, G. M., Yannoukakos, D., Clandinin, T. R., Schlessinger, J., and Sternberg, P. W. (1996). A point mutation in the extracellular domain activates LET-23, the Caenorhabditis elegans Epidermal Growth Receptor homolog. Mol. Cell. Biol. 16, 529–537. Kimble, J. (1981). Alterations in cell lineage following laser ablation of cells in the somatic gonad of Caenorhabditis elegans. Dev. Biol. 87, 286–300. Kimble, J., and Simpson, P. (1997). The LIN-12/Notch signaling pathway and its regulation. Ann. Rev. Cell Dev. Biol. 13, 333–361. King, R. W., Deshaies, R. J., Peteres, J.-M., and Kirschner, M. W. (1996). How proteolysis drives the cell cycle. Science 274, 1652–1658. Kipreos, E. T., Lander, L. E., Wing, J. P., He, W. W., Hedgecock, E. M. (1996). cul-1 is required for cell cycle exit in C. elegans and identifies a novel gene family. Cell 85, 829–839. Koga, M., and Ohshima, Y. (1995). Mosaic analysis of the let-23 gene function in vulval induction of Caenorhabditis elegans. Development 121, 2655–2666. Kornfeld, K. (1997). Vulval development in Caenorhabditis elegans. Trends Genet. 13, 55–61. Kornfeld, K., Guan, K.-L., and Horvitz, H. R. (1995a). The Caenorhabditis elegans gene mek-2 is required for vulval induction and encodes a protein similar to the protein kinase MEK. Genes Dev. 9,756–768 Kornfeld, K., Hom, D. B., and Horvitz, H. R. (1995b). The ksr-1 gene encodes a novel protein kinase involved in ras-mediated signaling in Caenorhabditis elegans. Cell 83, 903–913. Lackner, M. R., Kornfeld, K., Miller, L. M., Horvitz, H. R., and Kim, S. K. (1994). A MAP kinase homologue, mpk-1, is involved in ras mediated induction of vulval cell fates in Caenorhabditis elegans. Genes Dev. 8, 160–173. Lee, J., Jongeward, G. D., and Sternberg, P. W. (1994). unc-101, a gene required for many aspects of Caenorhabditis elegans development and behavior, encoded a clathrin-associated protein. Genes Dev. 8, 60–73. Levitan, D., and Greenwald, I. (1995). Facilitation of lin-12-mediated signaling by sel-12, a Caenorhabditis elegans S182 Alzheimer’s disease gene. Nature 377, 351–354. Levitan, D., and Greenwald, I. (1998a). LIN-12 protein expression and localization during vulval development in C. elegans. Development 125, 3101–3109.

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Minqin Wang and Paul W. Sternberg

Levitan, D., and Greenwald, I. (1998b). Effects of SEL-12 presenilin on LIN-12 localization and function in Caenorhabditis elegans. Development 125, 3599–3606. Lu, X., and Horvitz, H. R. (1998). lin-35 and lin-53, two genes that antaganize a C. elegans Ras pathway, encode proteins similar to Rb and its binding protein RbAp48. Cell 95, 981–991. Maloof, J. N., and Kenyon, C. (1998). The Hox gene lin-39 is required during C. elegans vulval induction to select the outcome of Ras signaling. Development 125, 181–190. McConnell, S. K., and Kaznowski, C. E. (1991). Cell cycle dependence of laminar determination in developing neocortex. Science 254, 282–285. Miller, L. M., Gallegos, M. E., Morisseau, B. A., and Kim, S. K. (1993). lin-31, a Caenorhabditis elegans HNF-3/fork head transcription factor homolog, specifies three alternative cell fates in vulval development. Genes Dev. 7, 933–947. Miller, L. M., Waring, D. A., and Kim, S. K. (1996). Mosaic analysis using a ncl-1(+) extrachromosomal array reveals that lin-31 acts in the Pn.p cells during Caenorhabditis elegans vulval development. Genetics 143, 1181–1191. Moss, E. G., Lee, R. C., and Ambros, V. (1997). Control of developmental timing in C. elegans by the cold chock domain protein LIN-28 and its regulation by the lin-4 RNA. Cell 88, 637–646. Newman, A. P., White, J. G., and Sternberg, P. W. (1996). Morphogenesis of the C. elegans hermaphrodite uterus. Development 122, 3617–3626. Pan, D., and Rubin, G. M. (1997). Kuzbaninian controls proteolytic processing of Notch and mediates lateral inhibition during Drosophila and vertebrate neurogenesis. Cell 90, 271–280. Plowman, G. D., Sundarsanam, S., Bingham, J., Whyte, D., and Hunter, T. (1999). The protein kinases of Caenorhabditis elegans: a model for signal transduction in multicellular organisms. Proc. Natl. Acad. Sci. USA 96, 13603–13610. Raff, M. C., Abney, E. R., and Fok-Seang, J. (1985). Reconstitution of a developmental clock in vitro: a critical role for astrocytes in the timing of oligodendrocyte differentiation. Cell 42, 61– 69. Ray, W. J., Yao, M., Nowotny, P., Mumm, J., Zhang, W., Wu, J. Y., Kopan, R., and Goate, A. M. (1999). Evidence for physical interaction between presenilin and Notch. Proc. Natl. Acad. Sci. USA 96, 3263–3268. Salser, S. J., and Kenyon, C. (1992). Activation of a C. elegans Antennapedia homologue in migrating cells controls their direction of migration. Nature 355, 255–258. Salser, S. J., Loer, C. M., and Kenyon, C. (1993). Multiple HOM-C gene interactions specify cell fates in the nematode central nervous system. Genes Dev. 7, 1714–1724. Sawa, H., Lobel, L. and Horvitz, H. R. (1996). The Caenorhabditis elegans gene lin-17, which is required for certain asymmetric cell divisions, encodes a putative seven-transmembrane protein similar to the Drosophila Frizzled protein. Genes Dev. 10, 2189–2197. Sharma-Kishore, R., White, J. G., Southgate, E., and Podbilewicz, B. (1999). Formation of the vulva in Caenorhabditis elegans: a paradigm for organogenesis. Development 126, 691–699. Shi, Y., and Mello, C. (1998). A CBP/p300 homolog specifies multiple differentiation pathways in Caenorhabditis elegans. Genes Dev. 12, 943–955. Sieburth, D. S., Sun, Q., and Han, M. (1998). SUR-8, a conserved Ras-binding protein with leucine-rich repeats, positively regulates Ras-mediated signaling in C. elegans. Cell 94, 119–130. Sieburth, D. S., Sundaram, M., Howard, R. M., and Han, M. (1999). A PP2A regulatory subunit positively regulates Ras-mediated signaling during Caenorhabditis elegans vulval induction. Genes Dev. 13, 2562–2569. Simske, J. S., and Kim, S. K. (1995). Sequential signaling during Caenorhabditis elegans vulval induction. Nature 375, 142–146. Simske, J. S., Kaech, S. M., Harp, S. A., and Kim, S. K. (1996). LET-23 receptor localization by the cell junction protein LIN-7 during C. elegans vulval induction. Cell 85, 195–204. Singh, N., and Han, M. (1995). sur-2, a novel gene, functions late in the let-60 ras-mediated signaling pathway during Caenorhabditis elegans vulval induction. Genes Dev. 9, 2251–2265.

6. Pattern Formation during C. elegans Vulval Induction

219

Solari, F., and Ahringer, J. (2000). NURD-complex genes antagonise Ras-induced vulval development in Caenorhabditis elegans. Curr. Biol. 10, 223–226. Solari, F., Bateman, A., and Ahringer, J. (1998). The Caenorhabditis elegans genes egl-27 and egr-1 are similar to MTA1, a member of a chromatin regulatory complex, and are redundantly required for embryonic patterning. Development 126, 2483–2494. Sommer, R. J. (1997). Evolutionary changes of developmental mechanisms in the absence of cell lineage alterations during vulval formation in the Diplogastridae (Nematoda). Development 124, 243–251. Sommer, R. J., and Sternberg, P. W. (1994). Changes of induction and competence during the evolution of vulval development in nematodes. Science 265, 114–118. Sommer, R. J., and Sternberg, P. W. (1996). Apoptosis and change of competence limit the size of the vulva equivalence group in Pristionchus pacificus: a genetic analysis. Current Biology 6, 52–59. Sommer, R. J., Eizinger, A., Lee, K.-Z., Jungblut, B., Bubeck, A., and Schlak, I. (1998). The Pristionchus HOX gene Ppa-lin-39 inibits programmed cell death to specify the vulva equivalence group and is not required during vulval induction. Development 125, 3865–3873. Stern, M. J., and DeVore, D. L. (1994). Extending and connecting signaling pathways in C. elegans. Dev. Biol. 166, 443–459. Sternberg, P. W. (1988). Lateral inhibition during vulval induction in Caenorhabditis-elegans. Nature 335, 551–554. Sternberg, P. W., and Han, M. (1998). Genetics of RAS signaling in C. elegans. Trends Genet. 14, 466–452. Sternberg, P. W., and Horvitz, H. R. (1986). Pattern formation during vulval induction in C. elegans. Cell 44, 761–772. Sternberg, P. W., and Horvitz, H. R. (1988). lin-17 mutations of Caenorhabditis elegans disrupt certain asymmetric cell divisions. Dev. Biol. 130, 67–73. Sternberg, P. W., and Horvitz, H. R. (1989). The combined action of two intercellular signaling pathways specifies three cell fates during vulval induction in C. elegans. Cell 58, 679–693. Sternberg, P. W., Yoon, C. H., Lee, J., Jongeward, G. D., Kayne, P. S., Katz, W. S., Lesa, G., Liu, J., Golden, A., Huang, L. S., and Chamberlin, H. M. (1994). Molecular genetics of proto-oncogenes and candidate tumor suppressors in Caenorhabditis elegans. Cold Spring Harb. Symp. Quant. Biol. 59, 155–163. Sulston, J. E., and Horvitz, H. R. (1977). Postembryonic cell lineages of the nematode Caenorhabditis elegans. Dev. Biol. 56, 110–156. Sulston, J. E., and Horvitz, H. R. (1981). Abnormal cell lineages in mutants of the nematode Caenorhabditis elegans. Dev. Biol. 82, 41–55. Sulston, J. E., and White, J. G. (1980). Regulation and cell autonomy during postembryonic development of Caenorhabditis elegans. Dev. Biol. 78, 577–597. Sundaram, M., and Greenwald, I. (1993). Suppressor of a lin-12 hypomorph define genes that interact with both lin-12 and glp-1 in Caenorhabditis elegans. Genetics 135, 765–783. Sundaram, M., and Han, M. (1995). The Caenorhabditis elegans ksr-1 gene encodes a novel Raf-related kinase involved in Ras-mediated signal transduction. Cell 83, 889–901. Sundaram, M., and Han, M. (1996). Control and integration of cell signaling pathways during C. elegans vulval development. Bioessays 18, 473–480. Tan, P. B., Lackner, M. R., and Kim, S. K. (1998). MAP kinase signaling specificity mediated by the LIN-1 Ets/LIN-31 WH transcription factor complex during C. elegans vulval induction. Cell 93, 569–580. Tax, F. E., Thomas, J. H., Ferguson, E. L., and Horvitz, H. R. (1997). Identification and characterization of genes that interact with lin-12 in Caenorhabditis elegans. Genetics 147, 1675–1695. Thomas, J. H., and Horvitz, H. R. (1999). The C. elegans gene lin-36 acts cell autonomously in the lin-35 Rb pathway. Development 126, 3449–3459.

220

Minqin Wang and Paul W. Sternberg

Thomas, J. H., Stern, M. J., and Horvitz, H. R. (1990). Cell interactions coordinate the development of the C. elegans egg-laying system. Cell 62, 1041–1052. Tomlinson, A., and Struhl, G. (1999). Decoding vectorial information from a gradient: sequential roles of the receptors Frizzled and Notch in establishing planar polarity in the Drosophila eye. Development 126, 5725–5738. Tuck, S., and Greenwald, I. (1995). lin-25, a gene required in vulval induction in Caenorhabditis elegans. Genes Dev. 9, 341–357. Walhout, A. J. M., Sordella, R., Lu, X., Hartley, J. L., Temple, G. F., Brasch, M. A., Thierry-Mieg, N., and Vidal, M. (2000). Protein interaction mapping in C. elegans using proteins involved in vulval development. Science 287, 116–122. Wang, M., and Sternberg, P. W. (1999). Competence and commitment of Caenorhabditis elegans vulval precursor cells. Dev. Biol. 212, 12–24. Wang, B. B., Muller-Immergluck, M. M., Austin, J., Robinson, N. T., Chisholm, A., and Kenyon, C. (1993). A homeotic gene cluster patterns the anteroposterior body axis of C. elegans. Cell 74, 29–42. Wen, C., Metzstein, M. M., and Greenwald, I. (1997). SUP-17, a Caenorhabditis elegans ADAM protein related to Drosophila KUZBANIAN, and its role in LIN-12/NOTCH signaling. Development 124, 4759–4767. Whitfield, C. W., Benard, C., Barnes, T., Hekimi, S., and Kim, S. K. (1999). Basolateral localization of the Caenorhabditis elegans epidermal growth factor receptor in epithelial cells by the PDZ protein LIN-10. Mol. Biol. Cell 10, 2087–2100. Wightman, B., Burglin, T. R., Gatto, J., Arasu, P., and Ruvkun, G. (1991). Negative regulatory sequences in the lin-14 3’-untranslated region are necessary to generate a temporal switch during Caenorhabditis elegans development. Genes Dev. 5, 1813–1824. Wittmann, C., Bossinger, O., Goldstein, B., Fleischmann, M., Kohler, R., Brunschwig, K., Tobler, H., and M¨uller, F. (1997). The expression of the C. elegans labial-like Hox gene ceh-13 during early embryogenesis relies on cell fate and on anteroposterior cell polarity. Development 124, 4193–4200. Wu, Y., and Han, M. (1994). Suppression of activated Let-60 Ras defines a role of Caenorhabditis elegans sur-1 MAP kinase in vulval differentiation. Genes Dev. 8, 147–159. Wu, Y., Han, M., and Guan, K.-L. (1995). MEK-2, a Caenorhabditis elegans MAP kinase kinase, functions in Ras-mediated vulval induction and other developmental events. Genes Dev. 9, 742– 755. Wu, G., Hubbard, E. J. A., Kitajewski, J. K., and Greenwald, I. (1998). Evidence for functional and physical association between Caenorhabditis elegans SEL-10, a Cdc4p-related protein, and SEL-12 presenilin. Proc. Natl. Acad. Sci. USA 95, 15787–15791. Yochem, J., Weston, K., and Greenwald, I. (1988). The Caenorhabditis elegans lin-12 gene encodes a transmembrane protein with overall similarity to Drosophila Notch. Nature 335, 547–550. Yoon, C. H., Lee, J., Jongeward, G. D., and Sternberg, P. W. (1995). Similarity of sli-1, a regulator of vulval development in C. elegans, to the mammalian proto-oncogene c-cbl. Science 269, 1102–1105.

7 A Molecular Clock Involved in Somite Segmentation Miguel Maroto and Olivier Pourqui´e Laboratoire de G´en´etique et de Physiologie du D´eveloppement (LGPD) Developmental Biology Institute of Marseille (IBDM) CNRS-INSERM-Universit´e de la M´editerran´ee-AP de Marseille, France I. Introduction II. Models for Somite Formation III. A Molecular Clock Linked to Somitogenesis A. Dynamic and Periodic Expression of c-hairy1 B. Other hairy-Related Genes C. The hairy-Related Proteins and the Molecular Clock IV. Notch Signaling Pathway A. Background B. Notch Signaling and Somitogenesis C. Notch Signaling and the Molecular Clock V. Other Genes Implicated in Somitogenesis A. The MesP Family B. EphrinB2 and EphA4 C. Paraxis D. Fibronectin and N-cadherin E. Wnt Signaling Pathway VI. Conservation of the Segmentation Clock in Evolution References

Somites are transient embryonic structures that are formed from the unsegmented presomitic mesoderm (PSM) in a highly regulated process called somitogenesis. Somite, formation can be considered as the result of several sequential processes: generation of a basic metameric pattern, specification of the antero-posterior identity of each somite, and, finally, formation of the somitic border. Evidence for the existence of a molecular clock or oscillator linked to somitogenesis has been provided by the discovery of the rhythmic and dynamic expression in the PSM of c-hairy1 and lunatic fringe, two genes potentially related to the Notch signaling pathway. These oscillating expression patterns suggest that an important role of the molecular clock could reside in the temporal control of periodic Notch activation, ultimately resulting in the regular array of the somites. We discuss both the importance of the Notch signaling pathway in the molecular events of somitogenesis and its relationship with the molecular clock, and, finally, in that context we review a number of other genes known to play a role in somitogenesis.  2001 Academic Press. C

Current Topics in Developmental Biology, Vol. 51 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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I. Introduction Somites are transient embryonic structures that are the precursors of several segmented organs such as the axial skeleton, body skeletal muscles, and part of the dermis. They are formed in a highly regulated process called somitogenesis from the unsegmented presomitic mesoderm (PSM) (Christ and Ordahl, 1995; Gossler and Hrabe de Angelis, 1998; Keynes and Stern, 1988; Tam and Trainor, 1994). The PSM, which is organized in two bilateral rods of mesenchymal cells that flank the caudal neural tube, matures gradually in an anterior to posterior direction. During the formation of somites, the most mature PSM cells, which are located at the most rostral end of each PSM rod, bud off as an epithelial sphere of cells to form the somite. Somites later differentiate to give rise dorsally to the dermomyotome, which will yield the skeletal muscles and the dorsal dermis, and ventrally to the sclerotome, which will form the vertebral column. The generation of somites occurs simultaneously with the recruitment of new progenitor mesenchymal cells from the primitive streak /tail bud as they become part of the caudal region of the PSM (Catala et al., 1995; Psychoyos and Stern, 1996). The pace of somitogenesis and the total number of somites formed are species-specific phenomena and these parameters vary widely among vertebrate species (Richardson et al., 1998). Fate-mapping studies have shown that the cell population destined to form the PSM derives from two symmetrical territories of the epiblast located laterally to the forming primitive streak in the chick and mouse (Hatada and Stern, 1994; Lawson et al., 1991; Tam et al., 2000). Cell lineage analyses in these species have demonstrated that during gastrulation, these epiblast cells are recruited to the primitive streak where they become a resident population of somitogenic stem cells (Nicolas et al., 1996; Stern et al., 1992). The current model assumes that the stem cell population divides asymmetrically to generate a stem cell daughter that will stay in the primitive streak, and a progenitor cell that will become part of the PSM (Nicolas et al., 1996; Psychoyos and Stern, 1996; Stern et al., 1992). In the frog and fish, the somites derive from two territories of the marginal zone located either side of the Spemann organizer or shield, respectively (Holley and NussleinVolhard, 2000; Keller, 2000). During gastrulation, these territories involute at the blastopore, level. In amphibian and fish embryos, the structural homolog of the primitive streak is the blastopore, which is commonly thought to correspond to a region where populations of cells are continually transiting to become internalized. Therefore, the existence of resident stem cells at the blastopore level such as those reported in amniotes is not a widely accepted concept in amphibian and fish. However, lineage studies have provided arguments in favor of the existence of somitic stem cells in these species (Kimmel and Warga, 1987a, 1987b; ZernickaGoetz et al., 1996). In this review somite development is considered as the result of several sequential processes: generation of a basic metameric pattern, specification of the antero-posterior identity of each somite, and finally, formation of the somitic border

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that separates the rostral end of the PSM and the caudal region of the new somite, as discussed later. It has been shown that at any one time the entire PSM produces a consistent number of somites, 5–6 in mouse and 10–12 in chick (Packard and Meier, 1983; Tam and Beddington, 1986). In addition, cell labeling experiments in chick and mouse indicate that cells undergo little movement in the PSM (Stern et al., 1988; Tam and Beddington, 1986). These observations suggest that within the PSM, cells might be roughly allocated to their prospective somite. The existence of a segmental prepattern in the PSM has been proposed to be reflected by the somitomeres, which are patterned arrays of loose mesenchymal cells, only detected by stereo scanning electron microscopy (Meier, 1984). Such a concept implicating a segmental prepattern of the PSM would suggest that the generation of the basic metameric pattern occurs concomitantly with PSM formation in the caudal region of the embryo. However, apart from its rostralmost aspect, where several genes are expressed in a striped fashion, no further evidence for a prepattern of segmentation has been observed for the PSM tissue. Once they are formed, somites are already subdivided into rostral and caudal compartments, which express different sets of genes and which go on to exhibit different functional properties. The anterior part of the sclerotome in the differentiated ventral somite is permissive to motoneuron axon and neural crest cell migration, whereas the posterior part is refractory to their migration (Keynes and Stern, 1984; Rickmann et al., 1985). This property underlies the segmentation of the peripheral nervous system. Timing of the determination of antero-posterior compartment identity has been addressed by experiments of in vitro culture of isolated PSM explants or by PSM inversion in the chick embryo (Aoyama and Asamoto, 1988; Bronner-Fraser and Stern, 1991; Palmeirim et al., 1998). These experiments demonstrated that antero-posterior determination takes place at the level of the rostral region of the PSM, in contrast to dorso-ventral or medio-lateral somite specification, which occur only after the somite has formed (Keynes and Stern, 1988). That somite formation and antero-posterior identity determination are differentially regulated and temporally distinct events is further supported by analyses of some of the Notch pathway homozygous null mutant mice that are able to form some somites, and these lack a clear antero-posterior organization (Conlon et al., 1995; Hrabe de Angelis et al., 1997; Kusumi et al., 1998; Oka et al., 1995). These results indicate that formation of epithelial somites and determination of antero-posterior compartment identity are clearly two different processes. Finally, with regard to the formation of a somitic border, a number of experiments described later demonstrate that border formation is not an essential requisite for metamerism. Firstly, epithelialization or somite formation is blocked when isolated chick PSM is cultured without overlying ectoderm, although this tissue can achieve a segmental pattern as evidenced by the striped expression of c-Delta-1, a marker of the caudal somite (Palmeirim et al., 1998). Secondly, mice embryos homozygous null for the gene that encode the bHLH transcription factor paraxis (discussed

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later) do not form clear somite boundaries, even though the tissue displays a normal antero-posterior identity and the embryos retain a segmented organisation (Burgess et al., 1996). Taken together, these results indicate that segmentation and determination of antero-posterior compartment identity are processes that can be uncoupled from somitic border formation.

II. Models for Somite Formation The previous section introduced the concepts of metamerism and antero-posterior somitic compartmentalization, and their independence from border formation, in addition to where each of these events might initiate. This section discusses a number of models for how these processes might occur. The “clock and wavefront” model proposed by Cooke and Zeeman (1976) is one of a series of models aimed at explaining the different aspects of somitogenesis. Briefly, in this model the authors postulate the existence of an intracellular clock or oscillator in PSM cells that is synchronized between a group of neighboring cells. This clock would operate in conjunction with a wavefront traveling along the embryonic antero-posterior axis, reflecting the rostro-caudal differentiation gradient of the embryo. When a synchronized group of cells in a specific “permissive state” of the oscillatory cycle is reached by the wavefront, then it will form a somite. A second model was proposed by Meinhardt (1986), in which he also postulated the existence in PSM cells of an oscillatory mechanism, whereby the cells of the PSM fluctuate between two mutually exclusive states, anterior or posterior. In this model the fluctuations between the two states are initiated by the cells attaining specific threshold concentrations of an informational gradient that extends rostrocaudally along the PSM. At the level of the rostral PSM these states would become stabilized by a reaction-diffusion mechanism, such that a group of cells in the anterior PSM would influence each other locally to maintain alternative states corresponding to the prospective anterior and posterior somitic compartments. A third model was proposed by Stern and colleagues (Primmett et al., 1989; Stern et al., 1988). Experiments using a single heat-shock treatment in chick embryos and more recently in fish produced repeated abnormalities at 5 somite intervals in the fish and 6–7 in the chick, along the rostro-caudal axis (Primmett et al., 1988; Roy et al., 1999). These results suggest the existence within the PSM of a certain periodicity, such that spatially distinct groups of cells along the PSM share a degree of synchrony with respect to their susceptibility to heat-shock treatment. In addition, the authors have also shown that treatment of chick embryos with cell cycle inhibitors produces similar repeated abnormalities at 6–7 somite intervals (Primmett et al., 1989). Based on their results in chick embryos, the authors proposed a model that correlates somitogenesis with cell cycle division. The model proposes that somite formation always takes place at the same time point of the cell cycle, restricting the possibility of such a transition to a specific group of

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competent cells. Furthermore, the model holds that the process of somitogenesis would take place following a fixed number of cell cycles after the cell population becomes part of the PSM tissue. Each of these three models accounts in part for different aspects of somitogenesis. A more complete explanation for somitogenesis would require the global integration of each of them. In this integrated model the ideas proposed by Cooke and Zeeman and by Stern and colleagues would be worthy in order to explain the existence in the PSM of a basic metameric pattern, Meinhardt’s model would account for the anterior-posterior specification, whereas all three models provide a possible explanation for the generation of the somitic border.

III. A Molecular Clock Linked to Somitogenesis A. Dynamic and Periodic Expression of c-hairy1 The first molecular evidence for the existence of an intrinsic clock or oscillatory behavior in PSM cells prior to segmentation was provided by the discovery of the dynamic expression of c-hairy1 in the chick PSM (Fig. 1) (Palmeirim et al., 1997). This gene is a vertebrate homolog of the Drosophila hairy gene that encodes a bHLH transcription factor. Extensive genetic screens have led to the identification of a large number of bHLH protein-encoding genes that are implicated in the development of the Drosophila peripheral nervous system (Brunet and Ghysen, 1999; Campos-Ortega, 1995; Fisher and Caudy, 1998). Most of these genes are involved in the formation of the sensory organ precursors. At the genetic level, the genes have been grouped as transcriptional activators or repressors, depending on their role during the formation of the sensory organ precursors. Thus, the homozygous null mutant hairy flies display an increase in the number of sensory hairs, indicating that the protein encoded by the hairy gene functions as a repressor of neurogenesis (Ohsako et al., 1994; Rushlow et al., 1989; Van Doren et al., 1991). Furthermore, it has been shown that hairy also functions as a pair-rule gene during segmentation (Ish-Horowicz et al., 1985; Rushlow et al., 1989). In addition to the hairy gene, this family of transcriptional repressors includes the seven bHLH genes of the Enhancer of Split complex (E(spl)) and deadpan. All hairy/E(spl)-related genes encode proteins with a bHLH DNA-binding domain containing a proline residue at a specific position in the basic region and a 4-amino acid motif, WRPW, located at the C terminus of the protein (Bier et al., 1992; Delidakis and Artavanis-Tsakonas, 1992; Knust et al., 1992; Rushlow et al., 1989). The WRPW motif is necessary and sufficient to bind and recruit a nonHLH corepressor protein called Groucho (Fisher et al., 1996; Paroush et al., 1994). hairy/E(spl)-related proteins in different species are expected to interact with Groucho homologs and to thereby function as active transcriptional repressors.

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Figure 1 Schematic representation of c-hairy1 expression in the PSM, with respect to an individual presomitic cell. (A) These diagrams of the segmental plate during the formation of the next somite show the c-hairy1 expression wave that sweeps across the PSM caudo-rostrally every 90 min. Expression is initiated in a broad caudal domain that narrows as it progresses anteriorly and is finally restricted to the caudal domain of the next somite to be formed. Thus, an individual PSM cell (black dot) will experience a “c-hairy1 on” phase and a “c-hairy1 off ” phase during each oscillation. S0; forming somite. SI; newly formed somite. SII; last but one somite. (B) History of a presomitic cell (black dot) in the PSM: from the time it exits the domain of self-renewing stem cells in the primitive streak/tailbud and becomes resident in the PSM (0H), until it is incorporated into a somite (18H). This time interval corresponds to the formation of 12 somites, which is the number of prospective somites in the PSM tissue. Thus, during the time it resides in the PSM, each cell will undergo 12 cycles of c-hairy1 expression. These oscillations therefore define a clock linked to both somite segmentation and possibly to regionalization of the somites along the A-P body axis.

Thus, the chick c-hairy1 gene, whose expression is dynamic in the chick PSM, encodes a protein that contains the characteristic bHLH domain and the WRPW motif (Palmeirim et al., 1997). This dynamic expression of c-hairy1 in the chick PSM cycles with the same periodicity as that of somite formation, namely 90 min. These oscillations are synchronized in the two rods of PSM tissue and initially appear as broad dynamic caudal-rostral waves that sweep across the PSM and then narrow at its rostral end. Finally, the expression of c-hairy1 becomes fixed in a halfsomite domain that gives rise to the caudal or posterior half of the forming somite,

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where the expression is then maintained. This dynamic expression is independent of protein synthesis and cell movements, and it also seems to be independent of instructive signals from the surrounding structures. These findings are reminiscent of the model proposed by Meinhardt: c-hairy1, which is a marker for the posterior half-somite after it forms, shows oscillatory expression in the cells of the PSM with an identical period to that required to form one somite. The oscillation stops when the boundary is made and c-hairy1 expression becomes stabilized in the posterior half of the new somite.

B. Other hairy-Related Genes Since the description of c-hairy1 expression, other dynamic patterns of expression have been described for several hairy/E(spl)-related genes in both chick and mouse. Jouve et al. (2000) have described the dynamic expression of avian c-hairy2 and murine HES-1. c-hairy2 is a second chick hairy-related gene highly related to c-hairy1. Its expression oscillates synchronously with c-hairy1 in most of the PSM. At the most rostral end of the PSM and in the newly formed somite, c-hairy1 and c-hairy2 are detected in complementary domains. While c-hairy1 is maintained in the posterior part of the presumptive next somite and in the newly formed somite, expression of c-hairy2 is maintained in the anterior domain. On the other hand, HES-1 encodes a murine bHLH transcription factor that on the basis of sequence similarity seems to be more closely related to c-hairy2 (Fig. 2). However, its expression pattern in the PSM is more similar to c-hairy1. Thus, it remains unclear if the mammalian gene is the true ortholog of either of these two avian genes. Another murine hairy/E(spl)-related bHLH transcription factor is encoded for by the HES-5 gene (Akazawa et al., 1992), which is also expressed in the primitive streak and the rostral PSM (de la Pompa et al., 1997). To date, there are no data suggesting that HES-5 expression could be dynamic in the PSM. In addition to the HES genes, several groups have independently reported the isolation of other hairy/E(spl)-related murine genes that are expressed in the PSM, each group ascribing them with a different nomenclature, namely Hey, Hesr, and HRT (Kokubo et al., 1999; Leimeister et al., 1999; Nakagawa et al., 1999). Based on the variable patterns of expression displayed by these hairy/E(spl)-related genes in the PSM, Nakagawa et al. (1999) propose that their expression could be dynamic. These Hey/Hesr/HRT proteins form a new hairy/E(spl)-related subfamily of putative bHLH factors that carry a YRPW motif that is slightly different from the WRPW Groucho-binding motif described for the subfamily of c-hairy1, c-hairy2, and HES-1. Furthermore, the invariant proline residue within the basic domain that is implicated in the DNA binding specificity of the hairy/E(spl)-related factors is changed to glycine in the Hey/Hesr/HRT proteins.

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Figure 2 Dynamic expression of HES-1 is expressed in the mouse PSM. (Top) Dorsal view of the tail region of E10.5 mouse embryos hybridized with the HES-1 probe and arranged in the order of the wave progression. In each panel, the caudal boundary of the last formed somite (SI) is marked by a white arrow. Expression in the presomitic mesoderm (PSM) appears as a broad caudal domain, which progressively moves anteriorly, while narrowing as the caudal cleft of somite 0 (S0) forms. HES-1 expression in the rostral PSM becomes restricted to the caudal domain of the prospective somite. Rostral to the top. (Bottom) Schematic representation of the correlation between HES-1 expression in the PSM and the progression of somite formation.

C. The hairy-Related Proteins and the Molecular Clock The dynamic expression of c-hairy1 and other hairy-related genes in the chick PSM suggests that these transcription factors could be important components of the molecular clock machinery that may be responsible for the proper specification of antero-posterior identity. By extension, the clock could also be linked to the formation of boundaries between somites. However, the possible contribution of c-hairy1 and other hairy-related genes, if any, to the molecular clock mechanism involved in somitogenesis is unknown. Because of their similarity to Drosophila hairy, it is likely that the chicken hairy-related gene products are

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also transcriptional repressors. Moreover, in addition to regulating expression of other genes, it is possible that hairy-related proteins bind to and repress their own promoters through a negative feedback mechanism. Consistent with this idea is the fact that the murine HES-1 gene encodes a bHLH transcriptional repressor that is able to bind and repress its own promoter (Takebayashi et al., 1994). This auto-repression by HES-1 is modulated by phosphorylation of the DNAbinding domain (Strom et al., 1997). Thus, as part of the clock mechanism, it is possible that c-hairy1 or HES-1 regulate their own transcription by a similar negative feedback mechanism during the oscillatory cycle. It is important to point out that the molecular clock linked to somitogenesis remains functional in the HES-1 null mutant mice, and that the embryos do not exhibit segmental abnormalities (Ishibashi et al., 1995; Jouve et al., 2000). However, it is possible that another hairy-related protein present in the null mutant embryos, such as Hey/Hesr/HRT, is compensating for the loss of HES-1. Nevertheless, the idea of autorepression is not consistent with the finding that inhibition of protein synthesis by cycloheximide treatment does not prevent the oscillation of c-hairy1 RNA expression, at least over a 60-min culture period (Palmeirim et al., 1997). One interpretation of this result would be to consider that c-hairy1 oscillating expression is an output of the molecular clock rather than a key regulator of this oscillation. However, one cannot exclude that a more intricate interpretation might exist, in which the system would have a certain degree of oscillating autonomy over short time periods, and because of that the result of the cycloheximide treatment might be different if a longer time period (i.e., two cycles) was to be considered. Indeed, when explants are cultured for 90–120 min, expression of c-hairy1 in treated and control sides was found to differ in 50% of the cases (Palmeirim et al., 1997).

IV. Notch Signaling Pathway A. Background The phenotypic analyses of different mutant mice have shown that the Notch pathway is an essential mediator of the somitogenesis process. The Notch signaling pathway (Fig. 3) is an evolutionarily conserved mechanism that functions in multiple cell determination processes during metazoan development. The system allows neighboring cells to communicate with each other through local cell–cell interactions, amplifying and consolidating molecular differences that, eventually, become manifest as different cell fates (Artavanis-Tsakonas et al., 1999; Kopan and Turner, 1996). The Notch protein is a single-pass transmembrane receptor able to interact through its extracellular domain with the transmembrane ligands Delta and Serrate/Jagged (Nye and Kopan, 1995). The ligands can function by a nonautonomous and short-range mechanism, restricting their effect to the adjacent cells

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Figure 3 Schematic representation of the Notch signaling pathway within the responding cell. The three cleavage sites on the Notch protein are illustrated by the scissors. Refer to text for the description of the Notch signaling pathway.

that express the receptor (Heitzler and Simpson, 1991). Moreover, genetic analyses during neurogenesis stages of Drosophila development and in vitro experiments with mammalian neuroblastoma cells have revealed that the ligand Delta can also exert cell-autonomous effects via Notch signaling (Franklin et al., 1999; Jacobsen et al., 1998). The existence of such cell-autonomous regulation is consistent with studies in cell culture showing colocalization of the Notch receptor and the Delta ligand in individual cells (Fehon et al., 1991). Interestingly, the level of ligand expressed by a cell can influence its ability to activate the Notch receptor and thus, depending upon the level of its expression its activity may be either agonistic or antagonistic (Franklin et al., 1999; Jacobsen et al., 1998; Micchelli et al., 1997; Sun and Artavanis-Tsakonas, 1996). The current model for Notch signaling assumes that, as a consequence of ligand binding, the Notch receptor becomes subject to proteolytic processing at the membrane level, followed by the translocation of its cleaved intracellular domain

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into the nucleus (Kidd et al., 1998; Kopan et al., 1996; Lecourtois and Schweisguth, 1998; Struhl and Adachi, 1998). Once there, the Notch intracellular fragment interacts with the transcription factor Su(H)/RBP-J␬ (Jarriault et al., 1995; Lu and Lux, 1996), and this complex activates transcription of downstream targets such as the E(spl)/HES family of genes, which in turn regulate transcription of other downstream genes (Bailey and Posakony, 1995; Lecourtois and Schweisguth, 1995; Ohtsuka et al., 1999). At the extracellular level the interaction between Delta and Serrate/Jagged ligands and the Notch receptor can be influenced by the presence of the Fringe protein (Irvine and Wieschaus, 1994). The Fringe gene encodes a putative secreted protein that is structurally related to glycosyltransferases (Yuan et al., 1997). During development of the fly wing, Fringe has been implicated in the specification of the wing margin, which separates the dorsal and ventral compartments through differential modulation of Notch signaling, by preventing the cells from responding to Serrate and potentiating their capability to respond to Delta (Fleming et al., 1997; Micchelli and Blair, 1999; Panin et al., 1997; Rauskolb et al., 1999). The dorsal compartment coexpresses Fringe, Notch, and Serrate, whereas the ventral compartment expresses Notch and Delta. Thus, the presence of Fringe in the dorsal compartment prevents those cells from responding to the Serrate ligand, whereas the cells without Fringe in the ventral compartment remain receptive to the signal from both ligands. As a result of this Fringe activity, a potentiated and localized Notch activation at the dorso-ventral interface is produced by interaction between Fringe positive and negative compartments (Fleming et al., 1997; Micchelli and Blair, 1999; Panin et al., 1997; Rauskolb et al., 1999). The activity of Notch and its ligands seems to be largely regulated by posttranslational proteolytic processing. Initially, the Notch protein is synthesized as a 300-kDa precursor molecule that is constitutively cleaved in the trans-Golgi network by a furin-like convertase (Blaumueller et al., 1997; Logeat et al., 1998). This cleavage occurs as the receptor traffics towards the plasma membrane, where it is exposed as a heterodimeric protein. The furin-like cleavage site of Notch lies within its extracellular domain. A second extracellular cleavage by the metalloprotease TACE occurs in response to ligand binding (Brou et al., 2000; Mumm et al., 2000), which has been proposed to trigger a third intracellular/ transmembrane cleavage to release the intracellular fragment of Notch (De Strooper et al., 1999; Ray et al., 1999; Schroeter et al., 1998). There are obvious similarities between the proposed means of processing the Notch protein and that of the ␤-amyloid precursor protein (APP), the protein precursor of the amyloid-␤-peptide that is the main component of the amyloid plaques found in Alzheimer’s disease patients (Haass and De Strooper, 1999; Hardy, 1997; Selkoe, 1994; Wolfe et al., 1999). A ␥ -secretase activity is responsible for the third intracellular cleavage of both Notch and APP, and this activity is closely associated to presenilin (De Strooper et al., 1999; De Strooper et al., 1998; Struhl and Greenwald, 1999; Wolfe et al., 1999; Ye et al., 1999). It is not yet clear what

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the specific function of presenilin is, but there are some data to support the idea that it may indeed correspond to the aspartyl-protease responsible for the ␥ -secretase activity (Ray et al., 1999; Wolfe et al., 1999). Finally, another processing event mediated by Kuzbanian, a member of the ADAM family of metalloproteases, has been shown to be required for Notch signaling (Pan and Rubin, 1997). Kuzbanian appears to be responsible for, at least, the processing of the ligand Delta at the cell surface, thus releasing the extracellular fragment, which could then act potentially as a diffusible ligand (Qi et al., 1999).

B. Notch Signaling and Somitogenesis Analyses of homozygous null mutant mice for different Notch pathway components have strongly implicated this pathway in somitogenesis. In situ analyses of mouse embryos show that the expression of some of these components, such as Delta1, Delta3, or RBP-J␬, is strong and uniform along the PSM, whereas the genes that encode for the Notch receptors have a more restricted PSM expression pattern, suggesting that Notch signaling could act differentially along the rostrocaudal axis of the PSM (Conlon et al., 1995; del Barco Barrantes et al., 1999; Hrabe de Angelis et al., 1997; Kusumi et al., 1998; Oka et al., 1995; Williams et al., 1995). The Notch1 gene is expressed in the rostral region of the PSM with its strongest expression domain located just caudal to the most recently formed somite (del Amo et al., 1992; Reaume et al., 1992). In the somitic region, Notch1 expression is downregulated and becomes concentrated to the more medial and dorsal areas of the somites. Null mutant embryos for Notch1 die between 10 and 11 dpc (Conlon et al., 1995; Swiatek et al., 1994). Somitogenesis in the mutant embryos is delayed and poorly coordinated between the two sides of the embryo. The mutants form fewer somites when compared with wild-type embryos, and these are of irregular size. However, the somites exhibit normal anterior–posterior identity as shown by the normal expression of N-myc and Mox-1, markers of the anterior and posterior somitic compartments, respectively. These results suggest that Notch1 could play a role in coordinating somitogenesis or in the timing of boundary formation, but not in the determination of the antero-posterior identity process or the generation of metamerism. It is, however, possible that the presence of other members of the Notch family of receptors provides some functional redundancy. In support of this notion, at least two other Notch genes are expressed in the murine PSM, Notch2 and Notch3 (Williams et al., 1995). However, their contribution to somitogenesis remains unclear. In terms of segmentation, Notch2 null mutant mice are normal, although this phenotype may also reflect functional redundancy (Hamada et al., 1999). However, this is unlikely to be the case with Notch3, since this gene has been reported to repress Notch1-mediated activation (Beatus et al., 1999).

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Several genes encoding for Delta ligands have been identified in different vertebrate species, including Xenopus, zebrafish, and mouse (Bettenhausen et al., 1995; Chitnis et al., 1995; Dunwoodie et al., 1997; Haddon et al., 1998; Jen et al., 1997; Smithers et al., 2000). Murine Delta1 (Dll1) is widely expressed along the PSM and subsequently in the posterior half of the somites (Bettenhausen et al., 1995). Homozygous null mutant mice embryos for Dll1 exhibit severe somitogenesis defects, although a basic metameric pattern is established (Hrabe de Angelis et al., 1997). The mutant embryos form only the first few rostral somites. The rostrocaudal organization of the somites is severely disrupted and expression of posterior somitic compartment markers such as uncx4.1 is lost (del Barco Barrantes et al., 1999). Accordingly, analysis of the peripheral nervous system indicates that the dorsal root ganglia are fused, implying the prior loss or disruption of the caudal sclerotome (Hrabe de Angelis et al., 1997). Surprisingly, no ectopic expression of anterior markers such as mCer1 is observed, suggesting that although cells of the caudal somite lose their identity, they do not switch fate to adopt a rostral identity. In fact, mCer1 expression is reduced and poorly defined in the PSM and the somites (del Barco Barrantes et al., 1999). This phenotype suggests that Dll1 could be critical for establishing and/or maintaining antero-posterior compartment identity. This is further supported by the disturbed expression of the antero-posterior identity marker Mesp2 (discussed later) in Dll1(−/−) embryos (del Barco Barrantes et al., 1999). A mutation in the second murine Delta gene, Delta3 (Dll3), has been identified as responsible for the pudgy mutation (Kusumi et al., 1998). Dll3 is normally expressed at high levels along the PSM and subsequently in the rostral halves of the newly formed somites (Dunwoodie et al., 1997). In pudgy/Dll3 mutant embryos the somitic boundaries are not correctly formed, as indicated by the diffuse expression of Pax-1. Somites appear irregular in shape and size. Anterior–posterior compartment identity is disrupted in mutant embryos as manifested by abnormal expression of Meox-1/Mox-1, a marker of the caudal somitic compartment, and the subsequent irregular development of spinal nerves and ganglia through the rostral compartment (Kusumi et al., 1998). Interestingly, whereas in Dll3 mutant embryos the expression of Dll1 is lost in the posterior compartment of each somite (Kusumi et al., 1998), in Dll1 mutant embryos the expression of Dll3 is upregulated in the PSM (del Barco Barrantes et al., 1999). The phenotypes of Notch1, Dll1, and Dll3 mutant mice, having somites that are bilaterally desynchronized and irregular in size and shape, support a model in which Notch signaling may be directly implicated in the coordination of boundary formation and furthermore in the establishment or maintaining of antero-posterior identity. However, it would appear Notch signaling is not an essential component of either the process of border formation or the definition of the basic metameric pattern. The phenotype of the homozygous null mutant mice for RBP-J␬ is also consistent with this idea (Oka et al., 1995) (Fig 4, see color insert). Briefly, the vertebrate RBP-J␬ protein is a transcriptional repressor that is homologous to Drosophila (Su(H)) (Amakawa et al., 1993; Dou et al., 1994; Furukawa et al.,

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1992) and that has been reported to be a key element in the Notch signaling pathway (Bailey and Posakony, 1995; Fortini and Artavanis-Tsakonas, 1994; Lecourtois and Schweisguth, 1995). As mentioned earlier, (Su(H))/RBP-J␬ is able to interact with the intracellular Notch fragment in the nucleus, and the complex then turns into a transcriptional activator of downstream genes (Jarriault et al., 1995; Tamura et al., 1995). Since there is no evidence to date suggesting the existence of another RBP protein expressed during mouse development (Oka et al., 1995), this knockout is likely to reflect a Notch-signaling null. Supporting this notion, the RBP-J␬(−/−) mutant mice display the most severe phenotype of the Notch pathway component mutants. Thus, the null mutant embryos show severe growth retardation and form only 4–5 disorganized and irregularly shaped somites (Oka et al., 1995). In addition, the disturbed expression of uncx4.1 in the posterior halfsomite and EphA4 in the anterior domain indicate that the antero-posterior identity is poorly defined (del Barco Barrantes et al., 1999). As mentioned earlier, one possibility for the wide variation of the phenotype seen in mice lacking different components of the Notch pathway is the possible functional redundancy provided by other members of the pathway. Supporting evidence that this could be the case stems from the analysis of the presenilin mutants. The null mutant embryos for presenilin-1 (PS1) exhibit relatively mild segmental defects. They only form the rostralmost somites, and these display irregular size and shape and the formation of somite boundaries is uncoordinated (Shen et al., 1997; Wong et al., 1997). The expression of Dll1 and Notch1 is downregulated in the PSM of these PS1(−/−) embryos (Wong et al., 1997). In contrast, presenilin-2 (PS2) null mutant mice do not exhibit any somite abnormalities (Donoviel et al., 1999). However, the embryos of the double homozygous null mutant mice lacking both PS1 and PS2 do not form somites at all. These mutant embryos suffer similar morphological defects to those observed for the RBP-J␬(−/−) embryos. In these PS1(−/−)/PS2(−/−) embryos the expression of Dll1 and Uncx4.1 is undetectable in the PSM and the somites (Donoviel et al., 1999). The phenotype of these double mutant mice indicates that PS1 and PS2 have different, but partially overlapping, activities. Because of the phenotype of the presenilin mutants it seems that both presenilin proteins play an important role in the Notch pathway and, by extension, in somitogenesis. In addition to the phenotypic analyses of null mutant mice, the importance of Notch signaling in somitogenesis has been demonstrated in Xenopus and zebrafish by ectopic expression of several components of the pathway. Thus, the expression of a dominant-negative form of x-Delta2 or x-Su(H) in frog embryos disrupts the organisation of the myotomes and the normal segmental pattern of x-Delta2 and Hairy2A in the rostral PSM (Jen et al., 1997). Similar somite defects can also be produced by overexpression of deltaD in zebrafish (Dornseifer et al., 1997). Additional studies in Xenopus show that periodic repression of ESR-4 and ESR-5 is required during segmentation (Jen et al., 1999). These are two hairy/E(spl)related genes acting downstream of Notch signaling. The repression occurs in a

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posterior half-segment domain in the rostral PSM and requires an ESR-5-mediated negative feedback loop. Once this striped pattern is established, ESR-5 mediates a positive feedback loop implicated in maintaining expression of members of the pathway (Jen et al., 1999). Thus, these results indicate that in frog and zebrafish, proper somite formation and establishment and/or maintenance of a segmental antero-posterior identity is also likely to be regulated through periodic regulation of Notch signalling.

C. Notch Signaling and the Molecular Clock The existence of a connection between the Notch pathway and the molecular clock involved in somitogenesis became clear following the description of the dynamic expression of lunatic fringe in the PSM. In mouse and chick embryos, this gene oscillates in a spatiotemporal pattern that closely resembles that reported for c-hairy1 (Aulehla and Johnson, 1999; Forsberg et al., 1998; McGrew et al., 1998). The expression of lunatic fringe appears as a wave that sweeps across the PSM and narrows as it moves anteriorly. The phenotype of the null mutant mice for lunatic fringe resembles that of other Notch pathway components (Evrard et al., 1998; Zhang and Gridley, 1998), consistent with its potential role in modulating this pathway. The homozygous mutant embryos exhibit severe segmentation defects, since they form somites that display irregular size and poorly defined anterior– posterior expression domains of Notch1 and Dll1. They also exhibit a drastic downregulation of the known Notch target gene HES-5 in the PSM (de la Pompa et al., 1997; Evrard et al., 1998). Thus, based on these results lunatic fringe could act genetically upstream of the Notch pathway to position and delimit its activity during antero-posterior compartment determination and somitic border formation. As mentioned earlier, in Drosophila, Fringe has been characterized as a modulator of Notch signaling. Thus, one of the possible functions of the molecular clock would be the rhythmic control of lunatic fringe expression and by extension the periodic control of Notch signaling. This control in turn would lead to the periodic and local arrangement of the molecular and cellular properties responsible for the establishment of antero-posterior identity, the proper formation of a boundary, and the subsequent generation of a new somite. The links between the Notch pathway and the molecular clock have been further reinforced by the observation of the dynamic expression of HES-1 in the mouse PSM (Jouve et al., 2000). HES-1 is a hairy/E(spl)-like downstream target of Notch signaling, as has been shown in several systems (Jarriault et al., 1995, 1998; Kageyama and Ohtsuka, 1999; Tomita et al., 1999). This suggests that the dynamic expression of HES-1 in the PSM could result from periodic Notch pathway activation. This would be in agreement with the observation that HES-1 expression is lost in the Dll1(−/−) mutant mice (Jouve et al., 2000). Furthermore, expression of lunatic fringe is impaired in the mouse mutants for Dll1 and RBP-J␬ (del Barco

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Barrantes et al., 1999). These results argue strongly in favor of a role for Notch signaling in the molecular clock. Thus, in the mouse, the Notch pathway could, in principle, play a role either in the generation or maintenance of the oscillations, and/or in the readout of these oscillations, controlling the expression of the cycling genes. With respect to somite boundaries, it has been proposed that these are generated as a result of the interaction between lunatic fringe positive and negative domains, through a similar mechanism to that which has been proposed to establish the position of the dorso-ventral boundary of the developing wing blade in Drosophila (del Barco Barrantes et al., 1999). However, there are obvious differences between the expression domains of these factors during vertebrate somitogenesis and Drosophila wing development. In chick and mouse, Serrate/Jagged and Delta are not expressed in mutually exclusive domains in the PSM (Bettenhausen et al., 1995; del Barco Barrantes et al., 1999; Lindsell et al., 1995). Moreover, Notch signaling activity appears to occur in a large region of the PSM, as evidenced by the expression domains of downstream target genes that are not restricted to those cells adjacent to the somitic border (del Barco Barrantes et al., 1999). Furthermore, it is noteworthy that the pattern of expression of lunatic fringe in the rostral PSM and somites is different in mouse and chick; in mouse it is expressed in the posterior somite compartment, whereas in chick it is expressed in the anterior compartment (Forsberg et al., 1998; Johnston et al., 1997; McGrew et al., 1998). Thus, any potential model that attempts to explain border formation in terms of the interface between lunatic fringe positive and negative domains should also integrate this species-specific difference in the expression domain of lunatic fringe.

V. Other Genes Implicated in Somitogenesis In addition to the Notch pathway components, a wider range of genes that display specific regional expression in the PSM have been demonstrated to play a key role in somitogenesis (Fig. 5). As discussed earlier, the waves of c-hairy1, c-hairy2, and lunatic fringe gradually narrow as they reach the anterior PSM to finally become fixed in a half-somite domain. With respect to a specific cycle, the fixed expression of the cycling genes in the most anterior PSM could be interpreted as the final arrest of the molecular clock within these cells. Some of the genes expressed specifically in this region are either known members of the Notch pathway, such as Notch2 and HES-5, or at least seem to impinge upon the pathway, such as Mesp1 and Mesp2 in mouse, cMeso1 in chick, or Thylacine1 in frog. There is, however, a subset of genes that has been implicated in somitogenesis and that has not been documented to show any obvious link with the Notch pathway, such as fibronectin or N-cadherin.

Cell-matrix interaction Epithelialisation

Cell-cell interaction

?

Wnt signalling

paraxis Eph signalling

Somite formation

? A/P identity

metamerism

periodic on/off Notch signalling

(coordination of the somite formation)

Mesp signalling

Cycling genes (c-hairy1, lunatic fringe)

?

periodic Notch signalling

(generation and/or maintenance of the oscillations)

PSM

Molecular clock or oscillator

Gastrulation

Figure 5 Summary of the main events and molecular players in somitogenesis. Refer to text.

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A. The MesP Family MesP1 and MesP2 are two murine bHLH transcription factors that are specifically expressed in the anterior PSM. They are absent in the newly formed somite. Mesp1 and Mesp2 are coexpressed in the rostral PSM at the level of prospective somite-I, with a one-somite-width space separation from the newly formed somite (Saga et al., 1996, 1997). The homozygous Mesp2 null mutant mice exhibit severe skeletal malformations attributable to abnormal segmentation. The mutant embryos produce epithelial somites at the cervical region, but the properties of anterior and posterior somitic compartments are altered as evidenced at the sclerotomal level by widespread expression of caudal sclerotomal markers, such as Mox-1 and Pax-1, throughout the somite (Saga et al., 1997). In the prospective thoracolumbar region of the Mesp2(−/−) embryos, not only the rostro-caudal polarity of the sclerotome, but also the somitic boundary is disturbed or absent. These somitic defects are reminiscent to those described for some members of the Notch pathway. In the homozygous Mesp2(−/−) mice the expression of Notch1 and Notch2 in the PSM is drastically downregulated, whereas Dll1 is not affected. Moreover, in the Dll1 and RBP-J␬ homozygous null mutant mice the expression of Mesp2 is lost, proving that Mesp2 could interact with the Notch pathway via a feedback loop. The nature of this regulatory loop between Mesp2 and the Notch pathway and its relevance during segmentation is not yet clear. The normal Mesp2 expression domain at the level of somite -I appears to be necessary for the proper expression of Notch1, even though Notch1 has a much broader expression domain that extends caudal to that of Mesp2. This remarkable result could be interpreted as a rostral-to-caudal flow of information. Interestingly, the lack of expression of FGFR-1 in the anterior portion of the PSM and the newly formed somite suggests that Mesp2 could also interact with the FGF signaling pathway. Although the expression of Mesp1, which overlaps completely with the Mesp2 expression domain in the rostral PSM, appears to be normal in the Mesp2(−/−) mice, budding of the epithelial somites is not observed. This observation is consistent with the analysis of the Mesp1 null mutant mice. These mice, though embryonic lethal, in contrast to the Mesp2 mutants, do not exhibit a strong segmentation defect (Saga, 1998). Expression of the genes of the Notch pathway is not affected in the Mesp1(−/−) mice. Interestingly, when the Mesp1 gene is introduced into the Mesp2 locus by a knockin strategy, it rescues the Mesp2(−/−) segmentation defects in a dose dependent fashion. Thus, two copies of the ectopic Mesp1 gene are able to restore the skeletal defects and the expression of Notch1, Notch2, and FGFR-1 in the PSM to almost normal levels. Even if the two proteins perform some different roles during development, the results suggest that MesP1 and MesP2 may also have some redundant functions that play an important role during somitogenesis via interaction with and regulation of Notch and FGF signaling. The chick c-Meso1 gene encodes a bHLH transcription factor closely related to the mouse Mesp proteins. This gene is also expressed in the region of prospective

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somite-I immediately posterior to the region of the next prospective somite. Loss of function of c-Meso1 brought about by antisense RNA or antisense oligonucleotide treatment produces attenuation of somitogenesis, supporting its possible role in segmentation (Buchberger et al., 1998). A third gene related to the MesP family called Thylacine1 has been described in Xenopus. This gene is expressed only in the anterior half of prospective somites in the rostral PSM (Sparrow et al., 1998). Thylacine1 is expressed in a pattern very similar to that reported for x-Delta2, Hairy2A, and ESR-5. In vitro assays have shown that Thylacine1 is a potent transcriptional activator and that ectopic expression of Thylacine1 by RNA injection causes segmentation defects. This ectopic expression of Thylacine1 altered the normal segmental expression of x-Delta2 and ESR-5. Interestingly, alteration of the Notch signaling pathway by injection of x-Su(H) also affected Thylacine1 expression. Thus, Thylacine1 seems to interact with the Notch pathway via a feedback loop, similar to the situation described for MesP2 in mouse.

B. EphrinB2 and EphA4 The Eph tyrosine kinase receptors are a large family of cell surface molecules, some of which are expressed in the rostral PSM and in the somites of a number of vertebrate species. In particular, the specific PSM expression profile of the EphrinB2 ligand and the EphA4 ephrin receptor in the PSM is conserved among zebrafish, chick and mouse (Bergemann et al., 1995; Flenniken et al., 1996; Gale et al., 1996; Nieto et al., 1992). Experiments in zebrafish show that blocking Eph signaling leads to abnormal somite boundary formation, although the antero-posterior patterning within the somites is not affected (Durbin et al., 1998). This disruption of Eph signaling in zebrafish affects the regulation and normal switching off of deltaD and her1, a hairy/E(spl)-related gene, in the rostral PSM. Interestingly, EphA4 expression is severely downregulated in several Notch pathway null mutant mice such as Dll1 or RBP-J␬, suggesting the existence of another feedback loop between the Eph and Notch signaling pathways (del Barco Barrantes et al., 1999). However, it is surprising that the homozygous null mutant mice for EphA4 do not exhibit somite abnormalities (Dottori et al., 1998). One possibility for this lack of a somitogenic phenotype could be the functional redundancy of Eph signaling in the mouse PSM by other, as yet unidentified, Eph members.

C. Paraxis Paraxis is a bHLH transcription factor that is expressed in the rostral PSM and newly formed somites. As the somite matures, paraxis expression becomes restricted to the dermomyotome in mouse and chick (Blanar et al., 1995; Burgess et al.,

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1995; Quertermous et al., 1994). The importance of paraxis during the process of epithelialization was revealed through the analysis of the paraxis(−/−) mice that fail to form epithelial somites (Burgess et al., 1996). The mutant embryos exhibit a normal segmented pattern of paraxial mesoderm derivatives, such as axial skeleton, skeletal muscle, or dermis. The implication of paraxis in somite formation is corroborated by experiments using antisense oligonucleotide treatment against paraxis mRNA in chick embryos, which also inhibited normal somite epithelialization (Barnes et al., 1997). These results clearly indicate that the process of epithelialization can be separated from the processes of metamerism. These results suggest that paraxis function is required for the formation of epithelial somites. However, it has since been reported that isolated chick PSM is able to maintain paraxis expression after 4 h of incubation, although the explants do not form somite borders (Palmeirim et al., 1998), suggesting that paraxis may be a necessary but not a sufficient component in the generation of somitic boundaries. Interestingly, paraxis expression seems to be affected when the Eph or Notch signaling pathways are altered. Thus, disruption of Eph signaling in the fish affects the normal downregulation of the zebrafish paraxis homolog par1 in the anterior domain of the somites (Durbin et al., 1998). In principle, it is possible that this downregulation of par1 is a direct effect of the Eph signaling. Moreover, the expression of paraxis is reduced in Dll1(−/−) mice (Hrabe de Angelis et al., 1997), raising the possibility that normal paraxis expression is also a target of the Notch signaling pathway.

D. Fibronectin and N-cadherin Intercellular signaling is one of the key processes that underlies the final events of somitogenesis. There is clear evidence for the important role played by local adhesive interactions during epithelialization and somite formation; both cell-matrix (fibronectin and integrin) and cell–cell interactions (cadherin) (Duband et al., 1987; Tam and Trainor, 1994). However, the connection between periodic Notch signaling activity and these intercellular interactions is not obvious. Fibronectin is an adhesion molecule that interacts with the integrin cell surface receptor to mediate cell–extracellular matrix adhesions (George et al., 1993; Lash et al., 1987, 1984; Lash and Yamada, 1986; Ostrovsky et al., 1983). As evidenced by the phenotype of the null mutant mice, the functions of both integrin and fibronectin proteins are required during somitogenesis. The fibronectin(−/−) embryos die at 8.5 dpc and their phenotype suggests there is a quantitative deficit in mesoderm formation, as manifested by the lack of notochord and somites (George et al., 1993; Georges-Labouesse et al., 1996). A less severe phenotype is observed in the ␣5-integrin(−/−) embryos, which fail to produce epithelial somites even if the paraxial mesoderm shows segmented blocks of mesenchymal cells (Yang et al.,

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1999). This milder phenotype for the null mutant of ␣5-integrin compared to that observed in fibronectin(−/−) embryos could be due to functional redundancy provided by another fibronectin receptor expressed in the PSM, such as a different member of the integrin family. Similarly, there is also evidence that cadherins play a role in modulating cell– cell adhesion during the course of somite formation. The null mutant mice for N-cadherin have somite abnormalities that resemble those observed after anti-Ncadherin antibody treatment of chick embryos (Linask et al., 1998), suggesting that cadherin-mediated cell–cell adhesion operates in the maintenance of the epithelial somite in both mouse and chick. Moreover, addition of RGD peptides that contain the minimal specific adhesion recognition signal of fibronectin stimulated N-cadherin synthesis during somitogenesis (Linask et al., 1998). This result implies that N-cadherin mediated cell–cell events are coordinated with fibronectinassociated cell–substratum adhesion. It is also interesting to point out that isolated chick PSM explants treated with RGD peptides are able to segment and make somites (Lash and Yamada, 1986), similar to what is observed if the PSM is cultured together with overlying ectoderm (Palmeirim et al., 1998).

E. Wnt Signaling Pathway The cell–cell adhesion molecule cadherin interacts via its cytoplasmic domain with ␤-catenin, which, in turn, binds ␣-catenin, thereby linking the cadherins, with the cytoskeleton (Wheelock and Knudsen, 1991). In addition to its interaction with cadherin, in which it plays a role in cytoskeletal interactions in the cytoplasm, ␤-catenin is also implicated in transcriptional regulation in the nucleus, where it binds and activates the Lef-1/Tcf-1 transcription factors (Behrens et al., 1996; Molenaar et al., 1996). In principle, one cannot rule out the possibility that molecules that mediate cell–matrix interactions could also influence cell–cell interactions, which could, in turn, also be implicated in the activity of some transcription factors. However, to date there is no clear data supporting such interconnection. Members of the Lef-1/Tcf-1 family have been identified as downstream components in the transduction pathway of a subgroup of Wnt signals (Hsu et al., 1998; Korinek et al., 1998; van de Wetering et al., 1997). The possible involvement of Lef-1 and/or Tcf-1 in somitogenesis is supported by the abnormalities observed in chick embryos after LiCl treatment (Linask et al., 1998), which has been described as mimicking Wnt signaling (Hedgepeth et al., 1997; Klein and Melton, 1996; Stambolic et al., 1996). This implication is even more evident from analyses of the phenotype of double null mutant mice for Lef-1 and Tcf-1 (Galceran et al., 1999). These mutant embryos suffer severe defects in somite formation. In the region anterior to the forelimb level, abnormal somitic structures are formed that lack the somite boundaries. Moreover, the mutant embryos do not form somites

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in the caudal region. They appear even to lack paraxial mesoderm in the region posterior to the forelimb level, where they form ectopic neural tubes (Galceran et al., 1999). This paraxial mesoderm phenotype of the Lef-1(−/−)/Tcf-1(−/−) mutant embryos is very similar to that observed in the Wnt3A(−/−) mutant embryos (Takada et al., 1994; Yoshikawa et al., 1997), consistent with a function of Lef-1 and Tcf-1 downstream of Wnt3A. Interestingly, Notch1 expression is lost in the PSM of the Lef-1(−/−)/Tcf-1(−/−) mutant embryos (Galceran et al., 1999). The existence of an interaction between the Notch and Wnt signaling pathways has been extensively described in Drosophila (Dierick and Bejsovec, 1999; Hing et al., 1994; Johnston and Edgar, 1998; Klein and Arias, 1998; Wesley, 1999). This connection introduces a new level of complexity to be incorporated into the understanding of the mechanism of the molecular clock involved in somitogenesis. Moreover, a direct interaction between PS1 and members of the catenin family has been reported (Zhou et al., 1997). The results show that PS1 functions to increase ␤-catenin stability (Zhang et al., 1998), which is crucial for signaling through Wnt and perhaps other signal-transduction pathways.

VI. Conservation of the Segmentation Clock in Evolution Thus far, genes expressed in a periodic pattern in the PSM have been identified essentially in chick and mouse. The Xenopus homologs of these Notch pathway genes, such as Hairy2A or lunatic fringe, appear to be expressed nondynamically in the PSM (Jen et al., 1997; Wu et al., 1996). However, recent analysis of deltaC expression in the zebrafish embryo indicates that it is expressed in a dynamic fashion in the PSM similar to the cycling expression of genes observed in amniotes (Dr. Julian Lewis, personal communication). This observation demonstrates the existence of a segmentation clock in zebrafish. Therefore, the segmentation clock mechanism appears to be conserved among most vertebrates rather than being a specific feature of the amniotes. Remarkably, although different genes are found to oscillate in amniotes and fishes, they are all related to Notch signaling. This would be consistent with a conserved role of the molecular clock in the periodic modulation of Notch signaling. Although the oscillating hairy-like genes of amniotes are related to the Drosophila pair rule-gene hairy, the degree of conservation between different species of the molecular mechanisms involved in segmentation remains a highly controversial topic. In Drosophila all segments are determined simultaneously. However, during vertebrate somitogenesis and segmentation of many different species, including short-germ band insects or annelids, segmentation proceeds by the segmental addition of new segments at the caudal end of the embryo. Moreover, besides the striking expression of the her1 gene in zebrafish, evidence for a pair-rule mechanism acting outside of holometabolous insect has never been provided (Takke and Campos-Ortega, 1999). All the pair-rule homologs analyzed

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in more primitive insects, such as grasshopper, do not exhibit a pair-rule pattern (Davis and Patel, 1999). An additional argument against general conservation of a segmentation mechanism is the lack of any evidence for a role of the Notch signaling pathway during segmentation in Drosophila. It will therefore be particularly interesting to study the expression of genes of the Notch pathway and hairy homologs in short-germ band insects or crustaceans to see whether a segmentation clock exists in more primitive animals that exhibit a progressive mode of segmentation.

Acknowledgments The authors thank Kim Dale, Mike McGrew, and Caroline Jouve for critical reading of this manuscript. Special thanks to Kim Dale for the translation from Spanglish to English. We are also grateful to Dr. Jose Luis de la Pompa for providing the pictures used in Fig. 4 and to Dr. Julian Lewis for sharing unpublished observations. Financial support was provided by the Centre National de la Recherche Scientifique (CNRS), the Association Fran¸caise contre les Myopathies (AFM), the Association pour la Recherche contre le Cancer (ARC), the Fondation pour la Recherche Medicale (FRM), and the Human Frontier Science Programme Organisation (HFSPO). M.M. is currently a recipient of a Marie Curie postdoctoral fellowship from the European Commission.

References Akazawa, C., Sasai, Y., Nakanishi, S., and Kageyama, R. (1992). J. Biol. Chem. 267, 21879–21885. Amakawa, R., Jing, W., Ozawa, K., Matsunami, N., Hamaguchi, Y., Matsuda, F., Kawaichi, M., and Honjo, T. (1993). Genomics 17, 306–315. Aoyama, H., and Asamoto, K. (1988). Development 104, 15–28. Artavanis-Tsakonas, S., Rand, M. D., and Lake, R. J. (1999). Science 284, 770–776. Aulehla, A., and Johnson, R. L. (1999). Dev. Biol. 207, 49–61. Bailey, A. M., and Posakony, J. W. (1995). Genes Dev. 9, 2609–2622. Barnes, G. L., Alexander, P. G., Hsu, C. W., Mariani, B. D., and Tuan, R. S. (1997). Dev. Biol. 189, 95–111. Beatus, P., Lundkvist, J., Oberg, C., and Lendahl, U. (1999). Development 126, 3925–3935. Behrens, J., von Kries, J. P., Kuhl, M., Bruhn, L., Wedlich, D., Grosschedl, R., and Birchmeier, W. (1996). Nature 382, 638–642. Bergemann, A. D., Cheng, H. J., Brambilla, R., Klein, R., and Flanagan, J. G. (1995). Mol. Cell. Biol. 15, 4921–4929. Bettenhausen, B., Hrabe de Angelis, M., Simon, D., Guenet, J. L., and Gossler, A. (1995). Development 121, 2407–2418. Bier, E., Vaessin, H., Younger-Shepherd, S., Jan, L. Y., and Jan, Y. N. (1992). Genes Dev. 6, 2137–2151. Blanar, M. A., Crossley, P. H., Peters, K. G., Steingrimsson, E., Copeland, N. G., Jenkins, N. A., Martin, G. R., and Rutter, W. J. (1995). Proc. Natl. Acad. Sci. USA 92, 5870–5874. Blaumueller, C. M., Qi, H., Zagouras, P., and Artavanis-Tsakonas, S. (1997). Cell 90, 281–291. Bronner-Fraser, M., and Stern, C. (1991). Dev. Biol. 143, 213–217. Brou, C., Logeat, F., Gupta, N., Bessia, C., LeBail, O., Doedens, J. R., Cumano, A., Roux, P., Black, R. A., and Israel, A. (2000). Mol. Cell 5, 207–216. Brunet, J. F., and Ghysen, A. (1999). Bioessays 21, 313–318.

244

Miguel Maroto and Olivier Pourqui´e

Buchberger, A., Seidl, K., Klein, C., Eberhardt, H., and Arnold, H. H. (1998). Dev. Biol. 199, 201–215. Burgess, R., Cserjesi, P., Ligon, K. L., and Olson, E. N. (1995). Dev. Biol. 168, 296–306. Burgess, R., Rawls, A., Brown, D., Bradley, A., and Olson, E. N. (1996). Nature 384, 570–573. Campos-Ortega, J. A. (1995). Mol. Neurobiol. 10, 75–89. Catala, M., Teillet, M. A., and Le Douarin, N. M. (1995). Mech. Dev. 51, 51–65. Chitnis, A., Henrique, D., Lewis, J., Ish-Horowicz, D., and Kintner, C. (1995). Nature 375, 761–766. Christ, B., and Ordahl, C. P. (1995). Anat. Embryol. (Berl.) 191, 381–396. Conlon, R. A., Reaume, A. G., and Rossant, J. (1995). Development 121, 1533–1545. Cooke, J., and Zeeman, E. C. (1976). J. Theor. Biol. 58, 455–476. Davis, G. K., and Patel, N. H. (1999). Trends Cell Biol. 9, M68–M72. de la Pompa, J. L., Wakeham, A., Correia, K. M., Samper, E., Brown, S., Aguilera, R. J., Nakano, T., Honjo, T., Mak, T. W., Rossant, J., and Conlon, R. A. (1997). Development 124, 1139–1148. De Strooper, B., Saftig, P., Craessaerts, K., Vanderstichele, H., Guhde, G., Annaert, W., Von Figura, K., and Van Leuven, F. (1998). Nature 391, 387–390. De Strooper, B., Annaert, W., Cupers, P., Saftig, P., Craessaerts, K., Mumm, J. S., Schroeter, E. H., Schrijvers, V., Wolfe, M. S., Ray, W. J., Goate, A., and Kopan, R. (1999). Nature 398, 518–522. del Amo, F. F., Smith, D. E., Swiatek, P. J., Gendron-Maguire, M., Greenspan, R. J., McMahon, A. P., and Gridley, T. (1992). Development 115, 737–744. del Barco Barrantes, I., Elia, A., Wunnsch, K., Hrabde De Angelis, M., Mak, T., Rossant, J., Conlon, R., Gossler, A., and de la Pompa, J. L. (1999). Curr. Biol. 9, 470–480. Delidakis, C., and Artavanis-Tsakonas, S. (1992). Proc. Natl. Acad. Sci. USA 89, 8731–8735. Dierick, H., and Bejsovec, A. (1999). Curr. Top. Dev. Biol. 43, 153–190. Donoviel, D. B., Hadjantonakis, A. K., Ikeda, M., Zheng, H., Hyslop, P. S., and Bernstein, A. (1999). Genes Dev. 13, 2801–2810. Dornseifer, P., Takke, C., and Campos-Ortega, J. A. (1997). Mech. Dev. 63, 159–171. Dottori, M., Hartley, L., Galea, M., Paxinos, G., Polizzotto, M., Kilpatrick, T., Bartlett, P. F., Murphy, M., Kontgen, F., and Boyd, A. W. (1998). Proc. Natl. Acad. Sci. USA 95, 13248–13253. Dou, S., Zeng, X., Cortes, P., Erdjument-Bromage, H., Tempst, P., Honjo, T., and Vales, L. D. (1994). Mol. Cell. Biol. 14, 3310–3319. Duband, J. L., Dufour, S., Hatta, K., Takeichi, M., Edelman, G. M., and Thiery, J. P. (1987). J. Cell Biol. 104, 1361–1374. Dunwoodie, S. L., Henrique, D., Harrison, S. M., and Beddington, R. S. (1997). Development 124, 3065–3076. Durbin, L., Brennan, C., Shiomi, K., Cooke, J., Barrios, A., Shanmugalingam, S., Guthrie, B., Lindberg, R., and Holder, N. (1998). Genes Dev. 12, 3096–3109. Evrard, Y. A., Lun, Y., Aulehla, A., Gan, L., and Johnson, R. L. (1998). Nature 394, 377–381. Fehon, R. G., Johansen, K., Rebay, I., and Artavanis-Tsakonas, S. (1991). J. Cell Biol. 113, 657–669. Fisher, A., and Caudy, M. (1998). Bioessays 20, 298–306. Fisher, A. L., Ohsako, S., and Caudy, M. (1996). Mol. Cell. Biol. 16, 2670–2677. Fleming, R. J., Gu, Y., and Hukriede, N. A. (1997). Development 124, 2973–2981. Flenniken, A. M., Gale, N. W., Yancopoulos, G. D., and Wilkinson, D. G. (1996). Dev. Biol. 179, 382–401. Forsberg, H., Crozet, F., and Brown, N. A. (1998). Curr. Biol. 8, 1027–1030. Fortini, M. E., and Artavanis-Tsakonas, S. (1994). Cell 79, 273–282. Franklin, J. L., Berechid, B. E., Cutting, F. B., Presente, A., Chambers, C. B., Foltz, D. R., Ferreira, A., and Nye, J. S. (1999). Curr. Biol. 9, 1448–1457. Furukawa, T., Maruyama, S., Kawaichi, M., and Honjo, T. (1992). Cell 69, 1191–1197. Galceran, J., Farinas, I., Depew, M. J., Clevers, H., and Grosschedl, R. (1999). Genes Dev. 13, 709–717. Gale, N. W., Holland, S. J., Valenzuela, D. M., Flenniken, A., Pan, L., Ryan, T. E., Henkemeyer, M., Strebhardt, K., Hirai, H., Wilkinson, D. G., Pawson, T., Davis, S., and Yancopoulos, G. D. (1996). Neuron 17, 9–19.

7. A Molecular Clock Involved in Somite Segmentation

245

George, E. L., Georges-Labouesse, E. N., Patel-King, R. S., Rayburn, H., and Hynes, R. O. (1993). Development 119, 1079–1091. Georges-Labouesse, E. N., George, E. L., Rayburn, H., and Hynes, R. O. (1996). Dev. Dyn. 207, 145–156. Gossler, A., and Hrabe de Angelis, M. (1998). Curr. Top. Dev. Biol. 38, 225–287. Haass, C., and De Strooper, B. (1999). Science 286, 916–919. Haddon, C., Smithers, L., Schneider-Maunoury, S., Coche, T., Henrique, D., and Lewis, J. (1998). Development 125, 359–370. Hamada, Y., Kadokawa, Y., Okabe, M., Ikawa, M., Coleman, J. R., and Tsujimoto, Y. (1999). Development 126, 3415–3424. Hardy, J. (1997). Trends Neurosci. 20, 154–159. Hatada, Y., and Stern, C. D. (1994). Development 120, 2879–2889. Hedgepeth, C. M., Conrad, L. J., Zhang, J., Huang, H. C., Lee, V. M., and Klein, P. S. (1997). Dev. Biol. 185, 82–91. Heitzler, P., and Simpson, P. (1991). Cell 64, 1083–1092. Hing, H. K., Sun, X., and Artavanis-Tsakonas, S. (1994). Mech. Dev. 47, 261–268. Holley, S. A., and Nusslein-Volhard, C. (2000). Curr. Top. Dev. Biol. 47, 247–277. Hrabe de Angelis, M., McIntyre, J., and Gossler, A. (1997). Nature 386, 717–721. Hsu, S. C., Galceran, J., and Grosschedl, R. (1998). Mol. Cell. Biol. 18, 4807–4818. Irvine, K. D., and Wieschaus, E. (1994). Cell 79, 595–606. Ish-Horowicz, D., Howard, K. R., Pinchin, S. M., and Ingham, P. W. (1985). Cold. Spring. Harb. Symp. Quant. Biol. 50, 135–144. Ishibashi, M., Ang, S. L., Shiota, K., Nakanishi, S., Kageyama, R., and Guillemot, F. (1995). Genes Dev. 9, 3136–3148. Jacobsen, T. L., Brennan, K., Arias, A. M., and Muskavitch, M. A. (1998). Development 125, 4531– 4540. Jarriault, S., Brou, C., Logeat, F., Schroeter, E. H., Kopan, R., and Isra¨el, A. (1995). Nature 377, 355–358. Jarriault, S., Le Bail, O., Hirsinger, E., Pourqui´e, O., Logeat, F., Strong, C. F., Brou, C., Seidah, N. G., and Isra¨el, A. (1998). Mol. Cell Biol. 18, 7423–7431. Jen, W. C., Wettstein, D., Chitnis, D., and Kintner, C. (1997). Dev. Suppl. 124, 1169–1178. Jen, W. C., Gawantka, V., Pollet, N., Niehrs, C., and Kintner, C. (1999). Genes Dev. 13, 1486–1499. Johnston, L. A., and Edgar, B. A. (1998). Nature 394, 82–84. Johnston, S. H., Rauskolb, C., Wilson, R., Prabhakaran, B., Irvine, K. D., and Vogt, T. F. (1997). Development 124, 2245–2254. Jouve, C., Palmeirim, I., Henrique, D., Beckers, J., Gossler, A., Ish-Horowicz, D., and Pourquie, O. (2000). Development 127, 1421–1429. Kageyama, R., and Ohtsuka, T. (1999). Cell Res. 9, 179–188. Keller, R. (2000). Curr. Top. Dev. Biol. 47, 183–246. Keynes, R. J., and Stern, C. D. (1984). Nature 310, 786–789. Keynes, R. J., and Stern, C. D. (1988). Development 103, 413–429. Kidd, S., Lieber, T., and Young, M. W. (1998). Genes Dev. 12, 3728–3740. Kimmel, C. B., and Warga, R. M. (1987a). Nature 327, 234–237. Kimmel, C. B., and Warga, R. M. (1987b). Dev. Biol. 124, 269–280. Klein, T., and Arias, A. M. (1998). Dev. Biol. 194, 196–212. Klein, P. S., and Melton, D. A. (1996). Proc. Natl. Acad. Sci. USA 93, 8455–8459. Knust, E., Schrons, H., Grawe, F., and Campos-Ortega, J. A. (1992). Genetics 132, 505–518. Kokubo, H., Lun, Y., and Johnson, R. L. (1999). Biochem. Biophys. Res. Commun. 260, 459–465. Kopan, R., and Turner, D. L. (1996). Curr. Opin. Neurobiol. 6, 594–601. Kopan, R., Schroeter, E. H., Weintraub, H., and Nye, J. S. (1996). Proc. Natl. Acad. Sci. USA 93, 1683–1688.

246

Miguel Maroto and Olivier Pourqui´e

Korinek, V., Barker, N., Willert, K., Molenaar, M., Roose, J., Wagenaar, G., Markman, M., Lamers, W., Destree, O., and Clevers, H. (1998). Mol. Cell. Biol. 18, 1248–1256. Kusumi, K., Sun, E. S., Kerrebrock, A. W., Bronson, R. T., Chi, D. C., Bulotsky, M. S., Spencer, J. B., Birren, B. W., Frankel, W. N., and Lander, E. S. (1998). Nature Genet. 19, 274–278. Lash, J. W., and Yamada, K. (1986). The adhesion signal of fibronectin: a possible trigger mechanism for compaction during somitogenesis. In “Somites in Developing Embryos” (B. R, E. DA and L. JW, eds.), pp. 201–208. Plenum Press, New York and London. Lash, J. W., Seitz, A. W., Cheney, C. M., and Ostrovsky, D. (1984). J. Exp. Zool. 232, 197–206. Lash, J. W., Linask, K. K., and Yamada, K. M. (1987). Dev. Biol. 123, 411–420. Lawson, K. A., Meneses, J. J., and Pedersen, R. A. (1991). Development 113, 891–911. Lecourtois, M., and Schweisguth, F. (1995). Genes Dev. 9, 2598–2608. Lecourtois, M., and Schweisguth, F. (1998). Curr. Biol. 8, 771–774. Leimeister, C., Externbrink, A., Klamt, B., and Gessler, M. (1999). Mech. Dev. 85, 173–177. Linask, K. K., Ludwig, C., Han, M. D., Liu, X., Radice, G. L., and Knudsen, K. A. (1998). Dev. Biol. 202, 85–102. Lindsell, C. E., Shawber, C. J., Boulter, J., and Weinmaster, G. (1995). Cell 80, 909–917. Logeat, F., Bessia, C., Brou, C., LeBail, O., Jarriault, S., Seidah, N. G., and Israel, A. (1998). Proc. Natl. Acad. Sci. USA 95, 8108–8112. Lu, F. M., and Lux, S. E. (1996). Proc. Natl. Acad. Sci. USA 93, 5663–5667. McGrew, M. J., Dale, J. K., Fraboulet, S., and Pourqui´e, O. (1998). Curr. Biol. 8, 979–982. Meier, S. (1984). Cell Differ. 14, 235–243. Meinhardt, H. (1986). Models of segmentation. In “Somites in Developing Embryos” (B. R, E. DA and L. JW, eds.), pp. 179–191. Plenum Press, New York and London. Micchelli, C. A., and Blair, S. S. (1999). Nature 401, 473–476. Micchelli, C. A., Rulifson, E. J., and Blair, S. S. (1997). Development 124, 1485–1495. Molenaar, M., van de Wetering, M., Oosterwegel, M., Peterson-Maduro, J., Godsave, S., Korinek, V., Roose, J., Destree, O., and Clevers, H. (1996). Cell 86, 391–399. Mumm, J. S., Schroeter, E. H., Saxena, M. T., Griesemer, A., Tian, X., Pan, D. J., Ray, W. J., and Kopan, R. (2000). Mol. Cell 5, 197–206. Nakagawa, O., Nakagawa, M., Richardson, J. A., Olson, E. N., and Srivastava, D. (1999). Dev. Biol. 216, 72–84. Nicolas, J. F., Mathis, L., Bonnerot, C., and Saurin, W. (1996). Development 122, 2933–2946. Nieto, M. A., Gilardi-Hebenstreit, P., Charnay, P., and Wilkinson, D. G. (1992). Development 116, 1137–1150. Nye, J. S., and Kopan, R. (1995). Curr. Biol. 5, 966–969. Ohsako, S., Hyer, J., Panganiban, G., Oliver, I., and Caudy, M. (1994). Genes Dev. 8, 2743–2755. Ohtsuka, T., Ishibashi, M., Gradwohl, G., Nakanishi, S., Guillemot, F., and Kageyama, R. (1999). EMBO J. 18, 2196–2207. Oka, C., Nakano, T., Wakeham, A., de la Pompa, J. L., Mori, C., Sakai, T., Okazaki, S., Kawaichi, M., Shiota, K., Mak, T. W., and Honjo, T. (1995). Development 121, 3291–3301. Ostrovsky, D., Cheney, C. M., Seitz, A. W., and Lash, J. W. (1983). Cell Differ. 13, 217–223. Packard, D. S. J., and Meier, S. (1983). Dev. Biol. 97, 191–202. Palmeirim, I., Henrique, D., Ish-Horowicz, D., and Pourqui´e O. (1997). Cell 91, 639–648. Palmeirim, I., Dubrulle, J., Henrique, D., Ish-Horowicz, D., and Pourqui´e, O. (1998). Dev. Genet. 23, 77–85. Pan, D., and Rubin, G. M. (1997). Cell 90, 271–280. Panin, V. M., Papayannopoulos, V., Wilson, R., and Irvine, K. D. (1997). Nature 387, 908–912. Paroush, Z., Finley, R. L. J., Kidd, T., Wainwright, S. M., Ingham, P. W., Brent, R., and Ish-Horowicz, D. (1994). Cell 79, 805–815. Primmett, D. R., Stern, C. D., and Keynes, R. J. (1988). Development 104, 331–339.

7. A Molecular Clock Involved in Somite Segmentation

247

Primmett, D. R., Norris, W. E., Carlson, G. J., Keynes, R. J., and Stern, C. D. (1989). Development 105, 119–130. Psychoyos, D., and Stern, C. D. (1996). Development 122, 1523–1534. Qi, H., Rand, M. D., Wu, X., Sestan, N., Wang, W., Rakic, P., Xu, T., and Artavanis-Tsakonas, S. (1999). Science 283, 91–94. Quertermous, E. E., Hidai, H., Blanar, M. A., and Quertermous, T. (1994). Proc. Natl. Acad. Sci. USA 91, 7066–7070. Rauskolb, C., Correia, T., and Irvine, K. D. (1999). Nature 401, 476–480. Ray, W. J., Yao, M., Mumm, J., Schroeter, E. H., Saftig, P., Wolfe, M., Selkoe, D. J., Kopan, R., and Goate, A. M. (1999). J. Biol. Chem. 274, 36801–36807. Reaume, A. G., Conlon, R. A., Zirngibl, R., Yamaguchi, T. P., and Rossant, J. (1992). Dev. Biol. 154, 377–387. Richardson, M. K., Allen, S. P., Wright, G. M., Raynaud, A., and Hanken, J. (1998). Development 125, 151–160. Rickmann, M., Fawcett, J. W., and Keynes, R. J. (1985). J. Embryol. Exp. Morphol. 90, 437–455. Roy, M. N., Prince, V. E., and Ho, R. K. (1999). Mech. Dev. 85, 27–34. Rushlow, C. A., Hogan, A., Pinchin, S. M., Howe, K. M., Lardelli, M., and Ish-Horowicz, D. (1989). EMBO J. 8, 3095–3103. Saga, Y. (1998). Mech. Dev. 75, 53–66. Saga, Y., Hata, N., Kobayashi, S., Magnuson, T., Seldin, M. F., and Taketo, M. M. (1996). Development 122, 2769–2778. Saga, Y., Hata, N., Koseki, H., and Taketo, M. M. (1997). Genes Dev. 11, 1827–1839. Schroeter, E. H., Kisslinger, J. A., and Kopan, R. (1998). Nature 393, 382–386. Selkoe, D. J. (1994). Curr. Opin. Neurobiol. 4, 708–716. Shen, J., Bronson, R. T., Chen, D. F., Xia, W., Selkoe, D. J., and Tonegawa, S. (1997). Cell 89, 629– 639. Smithers, L., Haddon, C., Jiang, Y., and Lewis, J. (2000). Mech. Dev. 90, 119–123. Sparrow, D. B., Jen, W. C., Kotecha, S., Towers, N., Kintner, C., and Mohun, T. J. (1998). Development 125, 2041–2051. Stambolic, V., Ruel, L., and Woodgett, J. R. (1996). Curr. Biol. 6, 1664–1668. Stern, C. D., Fraser, S. E., Keynes, R. J., and Primmett, D. R. (1988). Development 104, 231–244. Stern, C. D., Hatada, Y., Selleck, M. A., and Storey, K. G. (1992). Dev. Suppl., 151–156. Strom, A., Castella, P., Rockwood, J., Wagner, J., and Caudy, M. (1997). Genes Dev. 11, 3168–3181. Struhl, G., and Adachi, A. (1998). Cell 93, 649–660. Struhl, G., and Greenwald, I. (1999). Nature 398, 522–525. Sun, X., and Artavanis-Tsakonas, S. (1996). Development 122, 2465–2474. Swiatek, P. J., Lindsell, C. E., del Amo, F. F., Weinmaster, G., and Gridley, T. (1994). Genes Dev. 8, 707–719. Takada, S., Stark, K. L., Shea, M. J., Vassileva, G., McMahon, J. A., and McMahon, A. P. (1994) Genes Dev. 8, 174–189. Takebayashi, K., Sasai, Y., Sakai, Y., Watanabe, T., Nakanishi, S., and Kageyama, R. (1994). J. Biol. Chem. 269, 5150–5156. Takke, C., and Campos-Ortega, J. A. (1999). Development 126, 3005–3014. Tam, P. P. L., and Beddington, R. S. P. (1986). The metameric organization of the presomitic mesoderm and somite specification in the mouse embryo. In “Somites in Developing Embroys” (B. R, E. DA and L. JW, eds.), pp. 17–36. Plenum Press, New York and London. Tam, P. P., and Trainor, P. A. (1994). Anat. Embryol. (Berl.) 189, 275–305. Tam, P. P., Goldman, D., Camus, A., and Schoenwolf, G. C. (2000). Curr. Top. Dev. Biol. 47, 1–32. Tamura, K.,Taniguchi, Y., Minoguchi, S., Sakai, T., Tun, T., Furukawa, T., and Honjo, T. (1995). Curr. Biol. 5, 1416–1423.

248

Miguel Maroto and Olivier Pourqui´e

Tomita, K., Hattori, M., Nakamura, E., Nakanishi, S., Minato, N., and Kageyama, R. (1999). Genes Dev. 13, 1203–1210. van de Wetering, M., Cavallo, R., Dooijes, D., van Beest, M., van Es, J., Loureiro, J., Ypma, A., Hursh, D., Jones, T., Bejsovec, A., Peifer, M., Mortin, M., and Clevers, H. (1997). Cell 88, 789–799. Van Doren, M., Ellis, H. M., and Posakony, J. W. (1991). Development 113, 245–255. Wesley, C. S. (1999). Mol. Cell. Biol. 19, 5743–5758. Wheelock, M. J., and Knudsen, K. A. (1991). Differentiation 46, 35–42. Williams, R., Lendahl, U., and Lardelli, M. (1995). Mech. Dev. 53, 357–368. Wolfe, M. S., De Los Angeles, J., Miller, D. D., Xia, W., and Selkoe, D. J. (1999). Biochemistry 38, 11223–30. Wong, P. C., Zheng, H., Chen, H., Becher, M. W., Sirinathsinghji, D. J., Trumbauer. M. E., Chen, H. Y., Price, D. L., Van der Ploeg, L. H., and Sisodia, S. S. (1997. Nature 387, 288–292. Wu, J. Y., Wen, L., Zhang, W. J., and Rao, Y. (1996). Science 273, 355–358. Yang, J. T., Bader, B. L., Kreidberg, J. A., Ullman-Cullere, M., Trevithick, J. E., and Hynes, R. O. (1999). Dev. Biol. 215, 264–277. Ye, Y., Lukinova, N., and Fortini, M. E. (1999). Nature 398, 525–529. Yoshikawa, Y., Fujimori, T., McMahon, A. P., and Takada, S. (1997), Dev. Biol. 183, 234–242. Yuan, Y. P., Schultz, J., Mlodzik, M., and Bork, P. (1997). Cell 88, 9–11. Zernicka-Goetz, M., Pines, J., Ryan, K., Siemering, K. R., Haseloff, J., Evans, M. J., and Gurdon, J. B. (1996). Development 122, 3719–3724. Zhang, N., and Gridley, T. (1998). Nature 394, 374–377. Zhang, Z., Hartmann, H., Do, V. M., Abramowski, D., Sturchler-Pierrat, C., Staufenbiel, M., Sommer, B., van de Wetering, M., Clevers, H., Saftig, P., De Strooper, B., He, X., and Yankner, B. A. (1998). Nature 395, 698–702. Zhou, J., Liyanage, U., Medina, M., Ho, C., Simmons, A. D., Lovett, M., and Kosik, K. S. (1997). Neuroreport 8, 2085–2090.

Subject Index

A Actin cellular functions, 139 hyphal tip growth role binding proteins, 143–144 branch initiation role, 148–149 calcium actin role in gradients, 164 regulation of actin network by calcium, 169–170 cytoplasmic distribution, 139–141 cytoplasmic migration role, 152–153 membrane interactions with actin cytoskeleton, 141–143 membrane protein distribution control, 149–150 microtubule integration of cytoskeletal activity, 153–154 morphogenesis role apical exocytotic gradient regulation by vesicle supply center, 147 turgor pressure effects, 146–147 organelle movement and positioning, 151–152 vesicle transport and exocytosis role, 150–151 myosin interactions in motility, 144 Activin Follistatin binding, 29 mesoderm formation role, 39–41 Adenomatous polyposis coli (APC) mutation in colon cancer, 47 regulation of ␤-catenin levels, 15 Angiotensin II, myofibroblast activation, 94–95 APC, see Adenomatous polyposis coli ASMA, see ␣-Smooth muscle actin ATM knockoout phenotype, 125 meiosis role, 124–125 substrates of kinase, 124–125

ATR localization in synaptonemal complex, 126 meiosis role, 124–126 substrates of kinase, 124, 126 Axin dorsal–ventral specification role, 19 regulation of ␤-catenin levels, 15–16

B BAMBI, signaling, 22 Bix, endoderm differentiation role, 38–39 BLM, meiosis role, 123–124 BMP, see Bone morphogeneic protein Bone morphogeneic protein (BMP) ectoderm formation role, 45–46 embryonic patterning, 28 mesoderm formation role, 43–44 BRCA1, meiosis role, 124 BRCA2, meiosis role, 124

C Caenorhabditis elegans vulval induction, see Vulval induction, Caenorhabditis elegans N-Cadherin, somite formation role, 241 Calcineurin, hyphal extension role, 167–168 Calcium flux apical growth in fungi actin regulation of network by calcium, 169–170 role in gradients, 164 bootstrapping of ions, 164 branching role, 168–169 channels patch clamp, 162–163 spontaneous potassium-permeable channels, 163 stretch-activated channels, 162–163 free calcium detection

249

250 Calcium flux (cont.) fluorescent probe considerations, 155–156 loading of dyes, 156 ratiometric dyes, 157–158 single-wavelength dyes, 157 gradients, 155, 166 homeostasis control, 162 hyphal extension role, 166–168 internal stores, 165–166 membrane-associated calcium measurement with chlortetracycline, 158 total calcium determination electron energy loss spectroscopy, 161 ion microscopy, 162 overview, 158–159 preparative procedures, 159–160 X-ray microanalysis, 161 cellular functions, 154–155 Calmodulin, hyphal extension role, 167 Casein kinase, Dishevelled as substrate, 16–17 ␤-Catenin cell adhesion role, 11–12 dorsal–ventral specification role, 10–12, 14 glycogen synthase kinase-3 regulation and regulators, 14–17 mutation in colon cancer, 47 phosphorylation, 12–13 protein interactions, 13–14 regulation of levels, 14–16 subcellular localization in development, 12–13 Cer anterior endomesoderm expression, 31–32 ectoderm formation role, 45–46 c-hairy1, somite segmentation molecular clock, 225–229 c-hairy2, somite segmentation molecular clock, 227 Chlortetracycline, membrane-associated calcium measurement, 158 Chordin ectoderm formation role, 45–46 endoderm differentiation role, 38 prechordal mesoderm expression, 27 Chromosome, see Meiosis CKI-1, temporal regulation of vulval precursor cell competence and commitment, 198–199 c-Meso1, somite formation role, 238–239

Subject Index COR1(SCP3) knockout phenotype, 118 phosphorylation, 117 synaptonemal complex function, 113 Cortical rotation, Xenopus egg, 8–9 D DAN, tumor suppression, 48 derriere, mesoderm formation role, 42–43 Desmin, myofibroblast expression, 95, 99 DiDeorge syndrome, neural crest development abnormalities, 71–72 Dishevelled (Dsh) dorsal–ventral specification role, 18–19 kinases, 16–17 regulation of glycogen synthase kinase-3, 17 Dkk proteins anterior endomesoderm expression, 32 ectoderm formation role, 46 DMC1 homolog pairing role, 111 meiosis role, 120–121 RAD51 complex, 121–123 Dorsal vegetal region, Spemann organizer gene expression, 24–25 Dorsal–ventral specification, Xenopus laevis cooperation between transforming growth factor-␤ and Wnt signaling, 19–20 cortical rotation, 8–9 cytoplasmic determinants, 5–8 Nieuwkoop center, 9–10 overview, 2–5 transforming growth factor-␤ receptors, Smads, and target genes, 20–22 vegetal cortical cytoplasm/␤-catenin signaling pathway, 10–19 Dsh, see Dishevelled E Ectoderm, formation in Xenopus, 44–46 EELS, see Electron energy loss spectroscopy EGL-17, temporal regulation of vulval precursor cell competence and commitment, 199 Electron energy loss spectroscopy (EELS), total calcium determination in fungi, 161 Endoderm, formation in Xenopus, 35–39 Endodermin, endoderm differentiation role, 36 Endothelin-1 (ET-1), myofibroblast activation, 94, 100

Subject Index eomes, mesoderm formation role, 42–43 EphA4, somite formation role, 239 EphrinB2, somite formation role, 239 ET-1, see Endothelin-1 F FAST-1, signaling, 22 fatvg, RNA localization in frogs, 5–6 FGF, see Fibroblast growth factor Fibroblast growth factor (FGF), mesoderm formation role, 40–41 Fibronectin, somite formation role, 240–241 Follistatin ectoderm formation role, 45 prechordal mesoderm expression, 27, 29 Fringe, Notch signaling regulation, 231, 235 Frizzled, dorsal–ventral specification role, 17 frzb mesoderm formation role, 44 prechordal mesoderm expression, 29 G GATA6, vascular smooth muscle cell phenotype regulation, 79–80 Gax, vascular smooth muscle cell phenotype regulation, 80 Glycogen synthase kinase-3 (GSK-3) binding protein, 16 dorsal–ventral specification role, 19 regulation of ␤-catenin levels, 14–17 regulators, 14–18 Goosecoid (Gsc), prechordal mesoderm expression, 26 Gsc, see Goosecoid GSK-3, see Glycogen synthase kinase-3 H hairy genes, see c-hairy1; c-hairy2 HES-1, somite segmentation molecular clock, 227, 229, 235 Hey/Hesr/HRT proteins, somite segmentation molecular clock, 227, 229 Homologous recombination, see Meiosis Hop1, synaptonemal complex function, 112, 114 Hop2, homolog pairing role, 111

251 Hyphal tip actin binding proteins, 143–144 branch initiation role, 148–149 cytoplasmic distribution, 139–141 cytoplasmic migration role, 152–153 membrane interactions with actin cytoskeleton, 141–143 membrane protein distribution control, 149–150 microtubule integration of cytoskeletal activity, 153–154 morphogenesis role apical exocytotic gradient regulation by vesicle supply center, 147 turgor pressure effects, 146–147 organelle movement and positioning, 151–152 vesicle transport and exocytosis role, 150–151 calcium and apical growth in fungi actin regulation of network by calcium, 169–170 role in gradients, 164 bootstrapping of ions, 164 branching role, 168–169 channels patch clamp, 162–163 spontaneous potassium-permeable channels, 163 stretch-activated channels, 162–163 free calcium detection fluorescent probe considerations, 155–156 loading of dyes, 156 ratiometric dyes, 157–158 single-wavelength dyes, 157 gradients, 155, 166 homeostasis control, 162 hyphal extension role, 166–168 internal stores, 165–166 membrane-associated calcium measurement with chlortetracycline, 158 total calcium determination electron energy loss spectroscopy, 161 ion microscopy, 162 overview, 158–159 preparative procedures, 159–160 X-ray microanalysis, 161

252 Hyphal tip (cont.) cytoskeletal components and inhibitor studies, 137–138, 144 fungal growth characteristics, 136–137, 145–146 growth patterns in yeasts, 145 microtubules in apical growth, 138–139, 150–153 SNARE roles in growth, 148–149 I Id2, myofibroblast cell cycle regulation, 103–104 IFABP, endoderm differentiation role, 36 Ion microscopy, total calcium determination in fungi, 162 L Lef1, ␤-catenin interactions, 13 lef-1, somite formation role, 241–242 Leptin, dorsal–ventral specification role in mouse, 8 LET-23, signaling in vulval precursor cell fate patterning downregulation of ligand-induced activity of LET-23, 209–210 gene expression changes upon RAS activation, 205 inhibition of basal LET-23 activity, 207–209 lateral signaling, 210–211 overview, 192–193 RAS signaling in vulval induction, 203 LIM kinase-1, SRF as substrate, 77–78 LIN-3 lateral signaling, 210–211 RAS signaling in vulval induction, 203 temporal regulation of vulval precursor cell competence and commitment, 196, 199–200, 202 vulval precursor cell fate patterning, 192–193 LIN-4, temporal regulation of vulval precursor cell competence and commitment, 198, 202 LIN-12, signaling in vulval precursor cell fate patterning gene expression changes upon RAS activation, 205 lateral signaling, 210–212 overview, 192–193

Subject Index temporal regulation of vulval precursor cell competence and commitment, 199 LIN-14, temporal regulation of vulval precursor cell competence and commitment, 198, 202 LIN-28, temporal regulation of vulval precursor cell competence and commitment, 198, 202 LIN-39 gene expression changes upon RAS activation, 204–205 spatial regulation of vulval precursor cell competence, 193, 195–196 lunatic fringe, somite formation role, 235–236, 242 M MAB-5, spatial regulation of vulval precursor cell competence, 196 MEF2, see Myocyte enhancer factor 2 Meiosis, see also Synaptonemal complex chromosomes double-stranded breaks, 110–111, 117–119 homologous recombination, 110, 117–119 morphology, 110 proteins associated with cores checkpoint proteins, 124–126 prospects for study, 127 recombination proteins, 119–124 structural characteristics during prophase of meiosis I core formation, 111–115 overview, 111 synapsis, 115–116 synaptonemal complex dissolution, 116–117 phases, 110 Mesoderm anterior endomesoderm, Spemann organizer gene expression genes regulating formation, 30–31 growth factor antagonists, 31–33 mammalian homolog, 33–34 overview, 29–30 formation in Xenopus, 39–44 prechordal mesoderm, Spemann organizer gene expression growth factor antagonists, 27–29 overview, 25 transcription factors, 26–27 timing of induction, 42–43 MesP1, somite formation role, 238

253

Subject Index MesP2, somite formation role, 238 Message transport organizer (METRO) pathway, maternal RNA localization in frogs, 5–6 METRO pathway, see Message transport organizer pathway Microtubules, roles in fungal apical growth, 138–139, 150–153 Milk, endoderm differentiation role, 38–39 Mix.1, endoderm differentiation role, 38–39 Mixer, endoderm differentiation role, 36–38 MLH1, meiosis role, 122–123 MRE11, meiosis role, 119–120 MRFs, see Myogenic regulatory factors Msx1, ectoderm formation role, 46 Myocyte enhancer factor 2 (MEF2) myofibroblast gene regulation, 97–100 vascular smooth muscle cell formation role, 73–74 phenotype regulation, 80 MyoD, myofibroblast gene regulation, 97–98, 101 Myofibroblast cell cycle, 101–104 contractility, 100–101 cytokines and phenotypes, 94–95 muscle structural proteins regulation of expression, 97 types, 95–97 origin and differentiation, 92–94 overview of features and functions, 91–92 platelet-derived growth factor in development, 92–94 prospects for study, 104 Myogenic regulatory factors (MRFs), myofibroblast gene regulation, 97–100 Myogenin, myofibroblast expression, 97, 101–102 Myosin heavy chain, myofibroblast expression and function, 95–96, 99, 100–101 N NBS1, meiosis role, 119–120 NF-␬B, see Nuclear factor-␬B nieuwkoid/dharma, organizer activity, 25 Nieuwkoop center, dorsal–ventral specification in Xenopus, 9–10 Noggin ectoderm formation role, 45–46 endoderm differentiation role, 38 prechordal mesoderm expression, 27

Notch, somite formation signaling pathway Delta ligands, 233 molecular clock interactions, 235–236 mouse studies, 232–234 overview, 229–232 posttranslational processing, 231–232 Xenopus studies, 234–235 zebrafish studies, 234–235 Nuclear factor-␬B (NF-␬B), inflammation mediation in atherosclerosis, 79 P p21, myofibroblast cell cycle regulation, 101–103 p53, myofibroblast cell cycle regulation, 103 paraxis, somite formation role, 239–240 Patch clamp, fungal calcium channels, 162–163 Pax3, cardiac neural crest development role, 72 PDGF, see Platelet-derived growth factor Pem-3, RNA localization, 7 Plasmalemma, actin association, 141–142 Platelet-derived growth factor (PDGF), myofibroblast development role, 92–94 podB, septation role, 145 PP2A, see Protein phosphatase 2A Presomitic mesoderm, see Somite segmentation Protein phosphatase 2A (PP2A), regulation of ␤-catenin levels, 16 R RAD1, meiosis role, 126 RAD50, meiosis role, 119–120 RAD51 chromosome synapsis role, 116 DMC1 complex, 121–123 knockout lethality, 118 meiosis role, 120–121 Red1, synaptonemal complex function, 112, 114 Replication protein A (RP-A), meiosis role, 121–122 Retinoblastoma protein, myofibroblast cell cycle regulation, 103 RP-A, see Replication protein A S SC, see Synaptonemal complex SCP1, see SYN1(SCP1) SCP2, chromosome core component, 114

254 SCP3, see COR1(SCP3) SEL-1, LIN-12 regulation, 212 SEL-10, LIN-12 regulation, 212 sepA, septation role, 145 Siamois dorsal vegetal region expression, 24–25 dorsal–ventral specification role, 10–11, 19 SM22␣, transcriptional regulation, 74–75 Smads classes, 21–22 DPC4 mutation in cancer, 48 Smad2 in anteroposterior patterning and mesoderm formation, 33–34 transforming growth factor-␤ signal transduction, 20–21 SMC proteins, functions, 114–115 ␣-Smooth muscle actin (ASMA), myofibroblast expression, 95, 99 SNARE, roles in hyphal tip growth, 148–149 Somite segmentation N-cadherin role, 241 Drosophila, 242–243 fibronectin role, 240–241 molecular clock c-hairy1 expression and function, 225–229 c-hairy2 expression, 227 conservation in evolution, 242–243 HES-1 expression and function, 227, 229, 235 Hey/Hesr/HRT proteins, 227, 229 Notch signaling pathway Delta ligands, 233 molecular clock interactions, 235–236 mouse studies, 232–234 overview, 229–232 posttranslational processing, 231–232 Xenopus studies, 234–235 zebrafish studies, 234–235 presomitic mesoderm c-Meso1 expression, 238–239 EphA4 expression, 239 EphrinB2 expression, 239 fate mapping of cells, 222–223 Lef-1 expression, 241–242 lunatic fringe expression, 235–236, 242 MesP1 expression, 238 MesP2 expression, 238 paraxis expression, 239–240 structure, 222

Subject Index Tcf-1 expression, 241–242 Thylacine1 expression, 239 somitic border formation, 223–224 somitogenesis models, 224–225 overview, 222–223 SpAN, regulation of bone morphogeneic protein signaling, 28–29 Spectrin, hyphal tip growth role, 142–143 Spemann organizer anterior endomesoderm genes, 29–33 mammalian homolog, 33–34 dorsal vegetal region, 24–25 formation in Xenopus laevis, 23–24, 34 historical perspective of studies, 22–23 prechordal mesoderm region, 25–29 Spo11, meiosis role, 119 SRF phosphorylation, 77–78 protein interactions, 77 regulation of vascular smooth muscle cell growth and differentiation, 75–78, 80–81 STAT3, dorsal–ventral specification role in mouse, 8 SUP-17, LIN-12 regulation, 212 SYN1(SCP1) chromosome synapsis role, 115–116 phosphorylation, 117 Synaptonemal complex (SC) chromosome synapsis proteins, 115–116 dissolution, 116–117 lateral elements, 112 prospects for study, 127 protein composition of cores, 112–113 recombination role, 110 structural function model, 117 T Tcf proteins dorsal–ventral specification role, 14 functions, 13–14 protein interactions, 13 Tcf-1, somite formation role, 241–242 TGF-␤, see Transforming growth factor-␤ Thylacine1, somite formation role, 239 Tolloid, regulation of bone morphogeneic protein signaling, 28 ␤-Transducin repeat-containing protein (␤-TrCP), regulation of ␤-catenin levels, 16

255

Subject Index Transforming growth factor-␤ (TGF-␤) mesoderm differentiation role, 40 myofibroblast activation, 94 receptors, Smads, and target genes, 20–22 vascular smooth muscle cell formation role, 73 Wnt signaling cooperation in dorsal–ventral specification, 19–20 ␤-TrCP, see ␤-Transducin repeat-containing protein Tumorigenesis, developmental pathway overlap in Xenopus laevis, 47–48 twin, dorsal vegetal region expression, 24

patterning genes, table, 194–195 prospects for study, 213–214 RAS signaling gene expression changes upon RAS activation LET-23, 205 LIN-12, 205 LIN-39, 204–205 LET-23 receptor, 203 LIN-3 inductive signal, 203 negative regulation downregulation of ligand-induced activity of LET-23, 209–210 inhibition of basal LET-23 activity, 207–209 overview, 206–207 synthetic multivulva genes, 207–209 specificity, 206 transducing genes and mutation effects, 203–204 vulval precursor cells lineages, 190–191 signaling in fate patterning, 192–193 spatial regulation of cell competence LIN-39, 193, 195–196 MAB-5, 196 temporal regulation of cell competence and commitment cell cycle regulation, 199–202 CKI-1, 198–199 EGL-17, 199 heterochronic pathway, 197–199 identity of vulval precursor cells and daughters, 202–203 LIN-3, 196, 199–200, 202 LIN-4, 198, 202 LIN-12, 199 LIN-14, 198, 202 LIN-28, 198, 202 overview, 196–197

V Vascular smooth muscle cell (VSMC) commitment to lineage, 73–74 coronary vasculature formation, 72 heterogeneity of cells, 72–73 injury and phenotypes, 69–70, 78–79 mesoderm-derived cells and markers, 70–71 neural crest-derived cells, 71–72 phenotype modulation, 78–80 prospects for development and differentiation studies, 80–82 SRF regulation of growth and differentiation, 75–78, 80–81 transcriptional control of differentiation, 74–75 VCC, see Vegetal cortical cytoplasm Vegetal cortical cytoplasm (VCC) dorsal determinant, molecular nature, 18–19 signaling, see ␤-Catenin; Siamois translocation, 2 Wnt molecules in signaling activation, 17–18 VegT endoderm differentiation role, 37, 39 mesoderm differentiation role, 41–43 RNA localization in frogs, 6 Vg1 dorsal–ventral specification role, 19–20 mesoderm formation role, 40–41 RNA localization in frogs, 5–6 VSMC, see Vascular smooth muscle cell Vulval induction, Caenorhabditis elegans evolutionary implications, 212–213 larval stages, 190–191 lateral signaling 2◦ fate specification, 210–211 regulators, 211–212

W Wnt2, anteroposterior patterning and mesoderm formation, 34 WRN, meiosis role, 123–124 X Xanf-1, prechordal mesoderm expression, 26–27 Xblimp1, anterior endomesoderm expression, 31

256 Xbra, mesoderm formation role, 41 Xcat2, RNA localization in frogs, 6 Xdazl, RNA localization in frogs, 6 Xdd1, dorsal–ventral specification role, 19 Xenopus laevis advantages and limitations for embryogenesis studies, 3–4, 48–49 dorsal–ventral specification cooperation between transforming growth factor-␤ and Wnt signaling, 19–20 cortical rotation, 8–9 cytoplasmic determinants, 5–8 Nieuwkoop center, 9–10 overview, 2–5 transforming growth factor-␤ receptors, Smads, and target genes, 20–22 vegetal cortical cytoplasm/␤-catenin signaling pathway, 10–19 generation time, 4 germ layer formation ectoderm, 44–46 endoderm, 35–39 mesoderm, 39–44 modeling, 47 overview, 34–35 prospects for developmental studies, 48–49 Spemann organizer anterior endomesoderm genes, 29–33 mammalian homolog, 33–34

Subject Index dorsal vegetal region, 24–25 formation, 23–24, 34 historical perspective of studies, 22–23 prechordal mesoderm region, 25–29 transgenesis, 4 tumorigenesis and developmental pathways, 47–48 Xfz8, anterior endomesoderm expression, 32–33 Xhex, anterior endomesoderm expression, 31 XIHbox8, endoderm differentiation role, 36 Xlim-1, prechordal mesoderm expression, 26–27 Xnr-1, anterior endomesoderm expression, 30 Xnr-2, anterior endomesoderm expression, 30 Xnr-3, ectoderm formation role, 45 Xotx2, prechordal mesoderm expression, 27 X-ray microanalysis, total calcium determination in fungi, 161 Xsox17, endoderm differentiation role, 36–37 Xwnt-8 dorsal–ventral specification role, 10–11, 17–19 mesoderm formation role, 44 Xwnt-11, RNA localization in frogs, 6 Z Zip1, chromosome synapsis role, 116 Zip2, chromosome synapsis role, 116

Contents of Previous Volumes

Volume 42 Cumulative Subject Index, Volumes 20 through 41

Volume 43 1 Epigenetic Modification and Imprinting of the Mammalian Genome during Development Keith E. Latham

2 A Comparison of Hair Bundle Mechanoreceptors in Sea Anemones and Vertebrate Systems Glen M. Watson and Patricia Mire

3 Developmental of Neural Crest in Xenopus Roberto Mayor, Rodrigo Young, and Alexander Vargas

4 Cell Determination and Transdetermination in Drosophila Imaginal Discs Lisa Maves and Gerold Schubiger

5 Cellular Mechanisms of Wingless/Wnt Signal Transduction Herman Dierick and Amy Bejsovec

6 Seeking Muscle Stem Cells Jeffrey Boone Miller, Laura Schaefer, and Janice A. Dominov

7 Neural Crest Diversification Andrew K. Groves and Marianne Bronner-Fraser

8 Genetic, Molecular, and Morphological Analysis of Compound Leaf Development Tom Goliber, Sharon Kessler, Ju-Jiun Chen, Geeta Bharathan, and Neelima Sinha 257

258

Contents of Previous Volumes

Volume 44 1 Green Fluorescent Protein (GFP) as a Vital Marker in Mammals Masahito Ikawa, Shuichi Yamada, Tomoko Nakanishi, and Masaru Okabe

2 Insights into Development and Genetics from Mouse Chimeras John D. West

3 Molecular Regulation of Pronephric Development Thomas Carroll, John Wallingford, Dan Seufert, and Peter D. Vize

4 Symmetry Breaking in the Zygotes of the Fucoid Algae: Controversies and Recent Progress Kenneth R. Robinson, Michele Wozniak, Rongsun Pu, and Mark Messerli

5 Reevaluating Concepts of Apical Dominance and the Control of Axillary Bud Outgrowth Carolyn A. Napoli, Christine Anne Beveridge, and Kimberley Cathryn Snowden

6 Control of Messenger RNA Stability during Development Aparecida Maria Fontes, Jun-itsu Ito, and Marcelo Jacobs-Lorena

7 EGF Receptor Signaling in Drosophila Oogenesis Laura A. Nilson and Trudi Schupbach

Volume 45 1 Development of the Leaf Epidermis Philip W. Becraft

2 Genes and Their Products in Sea Urchin Development Giovanni Giudice

3 The Organizer of the Gastrulating Mouse Embryo Anne Camus and Patrick P. L. Tam

4 Molecular Genetics of Gynoecium Development in Arabidopsis John L. Bowman, Stuart F. Baum, Yuval Eshed, Joanna Putterill, and John Alvarez

5 Digging Out Roots: Pattern Formation, Cell Division, and Morphogenesis in Plants Ben Scheres and Renze Heidstra

Contents of Previous Volumes

259

Volume 46 1 Maternal Cytoplasmic Factors for Generation of Unique Cleavage Patterns in Animal Embryos Hiroki Nishida, Junji Morokuma, and Takahito Nishikata

2 Multiple Endo-1,4-␤-D-glucanase (Cellulase) Genes in Arabidopsis Elena del Campillo

3 The Anterior Margin of the Mammalian Gastrula: Comparative and Phylogenetic Aspects of its Role in Axis Formation and Head Induction Christoph Viebahn

4 The Other Side of the Embryo: An Appreciation of the Non-D Quadrants in Leech Embryos David A. Weisblat, Franc¸oise Z. Huang, Deborah E. Isaksen, Nai-fia L. Liu, and Paul Chang

5 Sperm Nuclear Activation during Fertilization Shirley J. Wright

6 Fibroblast Growth Factor Signaling Regulates Growth and Morphogenesis at Multiple Steps During Brain Development Flora M. Vaccarino, Michael L. Schwartz, Rossana Raballo, Julianne Rhee, and Richard Lyn-Cook

Volume 47 1 Early Events of Somitogenesis in Higher Vertebrates: Allocation of Precursor Cells during Gastrulation and the Organization of a Moristic Pattern in the Paraxial Mesoderm Patrick P. L. Tam, Devorah Goldman, Anne Camus, and Gary C. Shoenwolf

2 Retrospective Tracing of the Developmental Lineage of the Mouse Myotome Sophie Eloy-Trinquet, Luc Mathis, and Jean-Franc¸ois Nicolas

3 Segmentation of the Paraxial Mesoderm and Vertebrate Somitogenesis Olivier Pourqul´e

4 Segmentation: A View from the Border Claudio D. Stern and Daniel Vasiliauskas

260

Contents of Previous Volumes

5 Genetic Regulation of Somite Formation Alan Rawls, Jeanne Wilson-Rawls, and Eric N. Olsen

6 Hox Genes and the Global Patterning of the Somitic Mesoderm Ann Campbell Burke

7 The Origin and Morphogenesis of Amphibian Somites Ray Keller

8 Somitogenesis in Zebrafish ¨ Scott A. Halley and Christiana Nusslain-Volhard

9 Rostrocaudal Differences within the Somites Confer Segmental Pattern to Trunk Neural Crest Migration Marianne Bronner-Fraser

Volume 48 1 Evolution and Development of Distinct Cell Lineages Derived from Somites Beate Brand-Saberi and Bodo Christ

2 Duality of Molecular Signaling Involved in Vertebral Chondrogenesis Anne-H´el`ene Monsoro-Burq and Nicole Le Douarin

3 Sclerotome Induction and Differentiation Jennifer L. Docker

4 Genetics of Muscle Determination and Development Hans-Henning Arnold and Thomas Braun

5 Multiple Tissue Interactions and Signal Transduction Pathways Control Somite Myogenesis Anne-Ga¨elle Borycki and Charles P. Emerson, Jr.

6 The Birth of Muscle Progenitor Cells in the Mouse: Spatiotemporal Considerations Shahragim Tajbakhsh and Margaret Buckingham

7 Mouse –Chick Chimera: An Experimental System for Study of Somite Development Josiane Fontaine-P´erus

Contents of Previous Volumes

261

8 Transcriptional Regulation during Somitogenesis Dennis Summerbell and Peter W. J. Rigby

9 Determination and Morphogenesis in Myogenic Progenitor Cells: An Experimental Embryological Approach Charles P. Ordahl, Brian A. Williams, and Wilfred Denetclaw

Volume 49 1 The Centrosome and Parthenogenesis ¨ Thomas Kuntziger and Michel Bornens

2 ␥-Tubulin Berl R. Oakley

3 ␥-Tubulin Complexes and Their Role in Microtubule Nucleation Ruwanthi N. Gunawardane, Sofia B. Lizarraga, Christiane Wiese, Andrew Wilde, and Yixian Zheng

4 ␥-Tubulin of Budding Yeast Jackie Vogel and Michael Snyder

5 The Spindle Pole Body of Saccharomyces cerevisiae: Architecture and Assembly of the Core Components Susan E. Francis and Trisha N. Davis

6 The Microtubule Organizing Centers of Schizosaccharomyces pombe Iain M. Hagan and Janni Petersen

7 Comparative Structural, Molecular, and Functional Aspects of the Dictyostelium discoideum Centrosome Ralph Gr¨af, Nicole Brusis, Christine Daunderer, Ursula Euteneuer, Andrea Hestermann, Manfred Schliwa, and Masahiro Ueda

8 Are There Nucleic Acids in the Centrosome? Wallace F. Marshall and Joel L. Rosenbaum

9 Basal Bodies and Centrioles: Their Function and Structure Andrea M. Preble, Thomas M. Giddings, Jr., and Susan K. Dutcher

10 Centriole Duplication and Maturation in Animal Cells B. M. H. Lange, A. J. Faragher, P. March, and K. Gull

262

Contents of Previous Volumes

11 Centrosome Replication in Somatic Cells: The Significance of the G1 Phase Ron Balczon

12 The Coordination of Centrosome Reproduction with Nuclear Events during the Cell Cycle Greenfield Sluder and Edward H. Hinchcliffe

13 Regulating Centrosomes by Protein Phosphorylation Andrew M. Fry, Thibault Mayor, and Erich A. Nigg

14 The Role of the Centrosome in the Development of Malignant Tumors Wilma L. Lingle and Jeffrey L. Salisbury

15 The Centrosome-Associated Aurora/lpl-like Kinase Family T. M. Goepfert and B. R. Brinkley

16 Centrosome Reduction during Mammalian Spermiogenesis G. Manandhar, C. Simerly, and G. Schatten

17 The Centrosome of the Early C. elegans Embryo: Inheritance, Assembly, Replication, and Developmental Roles Kevin F. O ′Connell

18 The Centrosome in Drosophila Oocyte Development Timothy L. Megraw and Thomas C. Kaufman

19 The Centrosome in Early Drosophila Embryogenesis W. F. Rothwell and W. Sullivan

20 Centrosome Maturation Robert E. Palazzo, Jacalyn M. Vogel, Bradley J. Schnackenberg, Dawn R. Hull, and Xingyong Wu

Volume 50 1 Patterning the Early Sea Urchin Embryo Charles A. Ettensohn and Hyla C. Sweet

2 Turning Mesoderm into Blood: The Formation of Hematopoietic Stem Cells during Embryogenesis Alan J. Davidson and Leonard I. Zon

Contents of Previous Volumes

263

3 Mechanisms of Plant Embryo Development Shunong Bai, Lingjing Chen, Mary Alice Yund, and Zinmay Rence Sung

4 Sperm-Mediated Gene Transfer Anthony W. S. Chan, C. Marc Luetjens, and Gerald P. Schatten

5 Gonocyte–Sertoli Cell Interactions during Development of the Neonatal Rodent Testis Joanne M. Orth, William F. Jester, Ling-Hong Li, and Andrew L. Laslett

6 Attributes and Dynamics of the Endoplasmic Reticulum in Mammalian Eggs Douglas Kline

7 Germ Plasm and Molecular Determinants of Germ Cell Fate Douglas W. Houston and Mary Lou King